International Review of Cytology: A Survey of Cell Biology, Volume 239

International Review of A Survey of Cytology Cell Biology VOLUME 239 SERIES EDITORS Geoffrey H. Bourne James F. Dan...

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International Review of

A Survey of

Cytology Cell Biology VOLUME 239

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

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

EDITORIAL ADVISORY BOARD Eve Ida Barak Howard A. Bern Dean Bok William C. Earnshaw Hiroo Fukuda Ray H. Gavin Siamon Gordon Elizabeth D. Hay William R. Jeffry Bruce D. McKee

M. Melkonian Keith E. Mostov Andreas Oksche Vladimir R. Pantic´ Manfred Schliwa Teruo Shimmen Wildred D. Stein Ralph M. Steinman N. Tomilin

International Review of A Survey of

Cytology Cell Biology

Edited by

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

VOLUME 239

Front Cover Photograph: Courtesy of Lynne Cassimeris, Lehigh University.

Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK

This book is printed on acid-free paper. Copyright ß 2004, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2004 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0074-7696/2004 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail: [email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Customer Support’’ and then ‘‘Obtaining Permissions.’’ For all information on all Academic Press publications visit our Web site at www.academicpress.com ISBN: 0-12-364643-X PRINTED IN THE UNITED STATES OF AMERICA 04 05 06 07 08 9 8 7 6 5 4 3 2 1

CONTENTS

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

vii

Multihormonal Control of Vitellogenesis in Lower Vertebrates Alberta Maria Polzonetti-Magni, Gilberto Mosconi, Laura Soverchia, Sakae Kikuyama, and Oliana Carnevali I. II. III. IV. V. VI.

Introduction ............................................................................... History of Vitellogenin................................................................... Vitellogenesis and Reproductive Strategies ......................................... Hormones Regulating Vitellogenin Synthesis ....................................... Vitellogenin Utilization .................................................................. Vitellogenin as Biomarker of the Feminization Process Induced by Environmental Estrogens ............................................................... VII. Concluding Remarks .................................................................... References ................................................................................

1 3 8 17 20 25 32 34

Structure, Evolutionary Conservation, and Functions of Angiotensin- and Endothelin-Converting Enzymes Nathalie Macours, Jeroen Poels, Korneel Hens, Carmen Francis, and Roger Huybrechts I. Introduction ............................................................................... II. Angiotensin-Converting Enzyme Mammals.......................................... III. ACE in Invertebrates..................................................................... v

47 48 55

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CONTENTS

IV. Endothelin-Converting Enzyme: State of the Art in Mammals................... V. Other Roles of ACE and ECE .......................................................... VI. Concluding Remarks .................................................................... References ................................................................................

67 81 84 85

Cell and Molecular Biology of Nucleolar Assembly and Disassembly Patrick J. DiMario I. Introduction............................................................................... II. Nucleolar Organizers .................................................................... III. The Interphase Nucleolus: Compartments, Components, and Functions.................................................................................. IV. Nucleologenesis is Concurrent with the Reinitiation of NOR Transcription.............................................................................. V. Embryonic Nucleologenesis............................................................ VI. Nucleolar Disassembly at Mitosis .................................................... VII. Perspectives .............................................................................. References ................................................................................

100 102 110 121 145 153 158 160

MAPping the Eukaryotic Tree of Life: Structure, Function, and Evolution of the MAP215/Dis1 Family of Microtubule-Associated Proteins David L. Gard, Bret E. Becker, and S. Josh Romney I. II. III. IV. V.

Introduction............................................................................... Structure of MAP215/Dis1-Related Proteins in Protists.......................... Structure and Function of the Dis1-Related Proteins in Fungi .................. Structure and Function of Mor1/Gem1-Related Proteins in Plants............. The Structure and Function of MAP215/Dis1-Related Proteins in Animals................................................................................. VI. Toward Reconciling the Many Functions of MAP215/Dis1 Family Proteins........................................................................... VII. Conclusion ................................................................................ References ................................................................................

180 192 194 208

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

273

213 247 254 265

CONTRIBUTORS

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

Bret E. Becker (179), Department of Biology, University of Utah, Salt Lake City, Utah 84112-0840 Oliana Carnevali (1), Dipartimento di Scienze del Mare, Universita` Politecnica delle Marche, 60131 Ancona, Italy Patrick J. DiMario (99), Department of Biological Sciences, Louisiana State University, Baton Rouge, Louisiana 70803-1715 Carmen Francis (47), Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium David L. Gard (179), Department of Biology, University of Utah, Salt Lake City, Utah 84112-0840 Korneel Hens (47), Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium Roger Huybrechts (47), Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium Sakae Kikuyama (1), Department of Biology, School of Education, Waseda University, Shinjuku-ku, Tokyo 169,8050, Japan Nathalie Macours (47), Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium

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viii

CONTRIBUTORS

Gilberto Mosconi (1), Department of Comparative Morphology and Biochemistry, University of Camerino, V. Camerini 2, 62032 Camerino (MC), Italy Jeroen Poels (47), Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium Alberta Maria Polzonetti-Magni (1), Department of Comparative Morphology and Biochemistry, University of Camerino, V. Camerini 2, 62032 Camerino (MC), Italy S. Josh Romney (179), Department of Biology, University of Utah, Salt Lake City, Utah 84112-0840 Laura Soverchia (1), Department of Comparative Morphology and Biochemistry, University of Camerino, V. Camerini 2, 62032 Camerino (MC), Italy

Multihormonal Control of Vitellogenesis in Lower Vertebrates Alberta Maria Polzonetti-Magni,* Gilberto Mosconi,* Laura Soverchia,* Sakae Kikuyama,{ and Oliana Carnevali{ *Department of Comparative Morphology and Biochemistry, University of Camerino, V. Camerini 2, 62032 Camerino (MC), Italy { Department of Biology, School of Education, Waseda University, Shinjuku-ku, Tokyo 169,8050, Japan { Dipartimento di Scienze del Mare, Universita` Politecnica delle Marche, 60131 Ancona, Italy

The comparative approach on how and when vitellogenesis occurs in the diverse reproductive strategies displayed by aquatic and terrestrial lower vertebrates is presented in this chapter; moreover, attention has been paid to the multihormonal control of hepatic vitellogenin synthesis as it is related to seasonal changes and to vitellogenin use by growing oocytes. The hormonal mechanisms regulating vitellogenin synthesis are also considered, and the effects of environmental estrogens on the feminization process in wildlife and humans have been reported. It is then considered how fundamental nonmammalian models appear to be, for vitellogenesis research, addressed to clarifying the yolkless egg and the evolution of eutherian viviparity. KEY WORDS: Vitellogenin, estradiol-17b, pituitary hormones, reproductive strategies, yolk utilization, xenoestrogens. ß 2004 Elsevier Inc.

I. Introduction ‘‘Omnia ex ovo’’ (Harvey, 1578–1657). Indeed, the egg is a little world containing genetic and metabolic equipment consisting, on the one hand, of nucleic acids and transcriptional factors and, on the other, of proteins, lipids, sugars, enzymes, hormones, vitamins, and growth factors. The egg is, in fact, a complex International Review of Cytology, Vol. 239 0074-7696/04 $35.00

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Copyright 2004, Elsevier Inc. All rights reserved.

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POLZONETTI-MAGNI ET AL.

FIG. 1 Hypothalamus–hypophysial–gonadal axis regulating reproduction through both longand short-loop feedback mechanisms.

and intriguing minilaboratory in which life arises. The term yolk, or vitellus, could theoretically include the vast mixture of materials that are all necessary for embryonic development and that are deposited in the oocytes in storage form; thus, the formation of yolk is one of those physiological processes that was likely to have been studied from the earliest days of biological observation, representing a key event in reproductive success and occurring over a vast range of metazoans from nematodes to insects and vertebrates. Reproduction is the primary function of living organisms by which the evolutionary events create biodiversity in natural populations, and in vertebrates, it is controlled by cellular communications in the hypothalamus– hypophysial–gonadal axis (Fig. 1). In this context, vitellogenesis, a key event in reproductive function, is regulated by gonadal and nongonadal hormones in all oviparous vertebrate species. Therefore, in this review, we attempt to understand how and when vitellogenesis occurs in the diverse reproductive strategies displayed by aquatic and terrestrial lower vertebrates, why vitellogenesis has been chosen by many researchers as a model system to study other biological processes, and last but not least, which hormones play a role in the induction and termination of the vitellogenic process.

MULTIHORMONAL CONTROL OF VITELLOGENESIS

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II. History of Vitellogenin Vitellogenin was first obtained in crude form from an animal when Laskowski diluted the serum of laying hens with water and obtained a precipitate. He called this material serumvitellin, because of the prevailing evidence indicating similarities between yolk and blood constituents in the laying hen. In 1959, Mac Indoe showed that Laskowski’s serumvitellin could be resolved into at least two components noncovalently bound together as a complex. Then, in 1970, a lipophosphoprotein was identified and isolated by Wallace from the blood of vitellogenic females and estrogen-treated males of the amphibian, Xenopus laevis; however, unlike the chicken complex, it could not be dissociated by relatively mild procedures into yolk proteins. The name of vitellogenin, proposed by Pan et al. (1969) as a generic term for female specific blood-borne yolk precursors in insects, was further adopted for X. laevis protein. It was characterized in its native state as a large lipophosphoprotein (460 kDa), with a lipid and protein-phosphorus content of 12% and 1.4%, respectively, with one atom of calcium being associated with every protein-phosphorus group (Wallace, 1970). Xenopus laevis vitellogenin was later shown to be a glycoprotein (Ansari et al., 1971; Redshaw and Follett, 1971) to contain nonstechiometric amounts of biliverdin (Redshaw and Nicholls, 1971). Among vertebrates, vitellogenin is synthesized in an extraovarian site in, at least, all oviparous species so far studied, and the liver has been found to synthesize vitellogenin; the ovarian steroid, estradiol-17b, synthesized under the control of the hypothalamus–hypophysial–gonadal axis, has been considered for a long time to be a most important trigger in many species studied so far. The vitellogenin synthesized in the liver and released in the bloodstream is sequestered by growing oocytes through receptor-mediated endocytosis and enzymatically cleaved into the major yolk protein components, phosvitins, and lipovitellin, which are used as an energy source during embryogenesis. The synthesis and accumulation of yolk proteins, vitellogenesis, is therefore, crucial for oocyte development and for determining spawning success during the seasonally dependent reproductive process (Fig. 2). Many laboratories have extensively studied vitellogenesis in various vertebrate species. Their studies have yielded original and highly important results for understanding how vitellogenesis is strategic in oocyte maturation, and thus in reproductive success in both wild and cultured species (Wallace, 1985). Because a good marker of the vitellogenic process is consistent with the amount of vitellogenin in the blood, a suitable method for measuring plasma vitellogenin was set up by several laboratories; for that purpose, vitellogenins from diVerent species were purified and characterized, with

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FIG. 2 Scheme showing the steps through which vitellogenic process occurs in oviparous vertebrates.

specific antibodies being raised in the rabbit. The first VTG test consisted of the rocket immunoassay on agarose gel (Giorgi et al., 1982). By now, ELISAs have been validated in several diVerent species (Carnevali et al., 2002; Mosconi et al., 1994b, 1998). A. Vitellogenins and Vitellogenin Genes As early as 300 bc, it was realized that the yolk of an egg was not part of the embryo proper but, rather, acted as nourishment for the developing embryo, although the term ‘‘vitellogenin’’ was coined long after the process


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of vitellogenesis had been established as a model for study. Now, however, this term is reserved for yolk protein precursors that have been shown by sequence similarity to belong to an ancient gene family occurring over a vast range of metazoans from nematodes to insects and vertebrates. The vitellogenin genes and structures were widely and extensively reviewed by LaFleur (1999). Multiple vitellogenin genes exist for several vertebrates; in X. laevis, four VTG genes are transcribed, A1, A2, B1, and B2 (Germond et al., 1983; Wahli et al., 1979, 1980) and at least three (I, II, and III) are transcribed in chicken (Byrne et al., 1989; Evans et al., 1988; Wang and Williams, 1983) and two in Fundulus heteroclitus (LaFleur, 1999); VTG genes have been completely sequenced for X. laevis VTGA2 (Gerber-Huber et al., 1987) and chicken VTG II (van het Schip et al., 1987) and were found to be extremely long (17.5–20.3 kb). Each structure has 34 introns and 35 exons, and the interruption of structural sequences (exons) at homologous positions indicates the close relationship of the four genes in Xenopus and indicates that genes A and B arose first from the duplication of an ancestral vitellogenin gene and that the two subgroups of each sequence diverged after genomic duplication. Unlike structural organization, the coding nucleotide sequence varies extensively among genes. Between gene A2 from Xenopus and Chicken II, several of the 35 exons are very similar except in the region codifying for phosvitin, which appears to have evolved rapidly both by point mutations and duplication of serines or short amino acid stretches (Nardelli et al., 1987). Transcription of genes under the steroid hormone control is caused by specific regulatory DNA sequences, termed hormonal responsive elements. The estrogen-responsive element (ERE) present in the 50 flanking region of the X. laevis VTG gene has been characterized in the B1 gene. It is 33 bp long and consists of two 13-bp imperfect palindromic elements, both of which require enhancer activity. A third such element is present further upstream within the 50 flanking region of the gene, but it is unable to confer hormone responsiveness by itself; in contrast, a single 13-bp imperfect palindromic element in the VTG A2 gene is self-suYcient to act as fully active ERE. In the male genome, the VTG gene is silent; in fact, the suppression of VTG genes in the Xenopus male is caused by DNA methylation (Cooper et al., 1987). A sort of hepatic memory has been observed in several species, including amphibians and birds. The yolk protein genes are committed to tissuespecific expression early in development, and a bird given a single injection of estradiol at 15 days from embryogenesis can retain memory for up to 6 months after hatching. The persistance of memory through several rounds of cell division is the result of estrogen-dependent alterations in the chromatine structure of these genes with respect to memory (Edinger et al., 1997). The memory eVect is well evident when one compares the primary and the secondary response; during the latter, the response has a shorter lag and

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reaches a higher magnitude (Beuving and Gruber, 1971). Induction by estrogen results in a linear accumulation of vitellogenin mRNA, with the stability of the messenger being elicited by the presence of estradiol. In cells maintained in the presence of estrogen, VTG mRNA is stable for approximately 3 weeks; when estrogen is removed from the culture within 12–24 hours, it is degraded, indicating an estrogen–mediated stabilization of VTG mRNA against cytoplasmic degradation (Brock and Shapiro, 1983).

B. Biochemical Characterization of Vitellogenins in Fish, Amphibians, and Reptiles All members of the VTG gene family code for proteins that follow a common theme of molecular and cellular characteristics: they are of heterosynthetic origin under hormone induction, are transported by the blood, are taken up by the oocytes through receptor-mediated endocytosis, and are stored in a membrane-bound compartment. Vitellogenin is the precursor of two yolk proteins: the highly acidic phosvitin (PV)/phosvette—about half of all amino acid residues are blocks of polyphosphoserine—and lipovitellin, which is composed of a large and a small subunit (LV1 and LV2, respectively). The A2-deduced amino acid sequence indicated that in VTG A2, PV apparently represents an intervening sequence flanked by LV1 at the N terminus and LV2 near the C terminus. From the parental VTG polypeptide, about 20 kDa are missing; this portion is encoded by exons 30–35, corresponding to the C-terminal end of VTG (Wallace et al., 1990), a cysteine-rich region that has been related to the D domain of the human van Willebrand factor and that is involved in VTG binding to its membrane receptor. In several teleosts in addition to LV and PV, a third class of VTG-derived yolk protein, the b0 component was identified and characterized (Matsubara and Koya, 1997). Regarding the function of vitellogenin and its related yolk proteins, apart from the general hypothesis suggested long ago by Aristotle of their being nutrition for developing embryos, others have recently been put forward. For example, LaFleur (1999) theorized that yolk proteins can be functional as carriers of specific compounds. The yolk also represents in some animals the evolutionary shift occurring during the emergence of viviparous organisms from oviparous ancestors; one component of such a shift in reproductive strategy would be an alteration in hormone regulatory events, as suggested by Tsang and Callard (1987). Even if yolk proteins are nonexistent in mammals, with the exception of egg-laying monotremes, it has been suggested (LaFleur, 1999) that the two human proteins, apo B-100 and the von Willebrand factor, could gain access to the embryo from the mother through the placenta. This scenario seems fascinating, accounting for the


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yolk functions in all vertebrates. However, some evolutionary events occurred in the branch of eutherian mammals, perhaps through an intriguing shift in the steroidogenic pattern inhibiting vitellogenesis or through genetic mutation (Rothchild, 2003). Vitellogenin has been isolated from diVerent phyla of animals including arthropods (Giorgi et al., 1993, 1998; Lui and O’ Connor, 1977; Pan et al., 1969), nematodes (Klass et al., 1979), sea urchins (Amant et al., 1986), fish (Roubal et al., 1997), amphibians (Carnevali et al., 1991a; Ruddack and Wallace, 1968), reptiles (Brown et al., 1997; Carnevali et al., 1991a,b; Janeiro-Cinquini et al., 1999), and birds (Grengard et al., 1964). More recently, it has also been purified and characterized in the amphioxus, or lancelet, by Sun and Zhang (2001) from oocyte-yolk as a homodimer of 320 kDa in native-PAGE. It appears to be composed of two monomeric subunits of 160 kDa in SDS-PAGE and circulates as a homodimer; it is of interest to note that the VTGs from fish, amphibians and birds, nematodes, and arthropods circulate as homodimers. In addition, amino acid composition of the lancelet oocyte–yolk protein was found to be generally similar to that of vitellogenin isolated from diVerent phyla of animals, including both vertebrates and invertebrates. The intriguing aspect of the lancelet is whether it has a liver-like organ, as it is the living invertebrate most closely related to the vertebrates. Vitellogenin has also been isolated and characterized in many teleost species because of its importance in studies applied to wild and aquaculture species of commercial interest (Mosconi et al., 1998; Roubal et al., 1997); it is not only because of the surge of nutritive material used by the embryo during development but also because its large molecular mass enables it to bind and transport calcium and zinc, in addition to delivering other compounds to the oocytes, including hormones, retinols, carotenoids, and riboflavin-binding protein. Vitellogenin is specific to maturing females and is very antigenic, which has facilitated development of immunoassays for its detection in numerous fish species (reviewed by Mommsen and Walsh, 1998; Specker and Sullivan, 1994). Therefore, vitellogenin assessment is considered to be a useful approach in evaluating female maturity related with peripheral gonadal steroid changes, and to assess sex ratio (Heppell and Sullivan, 1999). In addition, because vitellogenin synthesis can also be induced in males and juveniles through exposure to exogenous estradiol or to substances that mimic estrogens, there is increasing interest in using the abnormal production of VTG in male animals as a biological marker for exposure to xenoestrogenic materials (see Section VI). The extensive similarity among cordate vitellogenins can also be recognized through the purification and partial characterization of a reptile, the tuatara (Sphenodon punctatus), VTG (Brown et al., 1997). In that work, it was found that the region located in the N-terminal half of lipovitellin I

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comprises the first 1100 amino acids of intact chicken, frog, and fish VTGs. The lipovitellin I domain appears to function for recognition of VTG by occytic receptors before its internalization. This domain is the most conserved region among the available cordate VTG sequences (11, 45, 52, 58, and 65). Tuatara VTG was also found to have a slightly higher number of residues that were identical to chicken VTG rather than to X. laevis VTG, concordant with the generally accepted notion of a close evolutionary relationship between reptiles and birds, lineages thought to have separated some 240 million years ago. Tuatara is in fact a good model in comparative and evolutionary studies on reproductive biology, as it is sole surviving member of the Sphenodontida, an order of reptiles that otherwise contains only fossils from the Mesozoic. Proteins destined for secretion are known to undergo various posttranslational modification that can be responsible for the spatial configuration and external shape of the molecule, rather than for the primary structure itself. In fact, at protein level, VTGs are not very conserved in the phylogenetic and evolutionary transition from aquatic to terrestrial life (Carnevali and Belvedere, 1991; Carnevali et al., 1991a).

III. Vitellogenesis and Reproductive Strategies Oviparity, ovoviparity, and viviparity are terms used to describe oVspring at the moment of separation from a parent. Viviparity is a form of reproduction in which the embryo begins and also completes its development within or on a part of the parent’s body. Oviparous species all lay eggs with protective covering, from which a larval or juvenile form will later hatch regardless of the state of development at oviposition. The term ovoviviparity has been applied in varying ways and has been well described by Blackburn (1995). Because of the great diversity among vertebrates and the important role of natural selection in reproductive process, it is even more diYcult to generalize about the reproductive pattern. Attainment of sexual maturity occurs at a time characteristic for each species and is followed by a series of reproductive cycles closely related with environmental factors. Bony fishes display the full range of reproductive strategies known for vertebrates. Depending on the species, sexual maturity may be achieved during the first year of life (many teleosts), after more than 15 years of juvenile existence (Atlantic eel, sturgeon), or at some intermediate period. Some animals breed only once after attaining sexual maturity, whereas others exhibit two or more reproductive cycles; a few species may breed as one sex and then change to the opposite sex and breed again.


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Natural environmental factors, such as temperature and photoperiod, the presence of suitable breeding or nesting sites, and food availability, influence the central nervous system and the hypothalamus–hypophysial–gonadal axis and regulate gonadal maturation and secretion of sex hormones. Steroid hormones, pituitary hormones, or both determine development of secondary sexual characters and influence courtship, breeding, and parental behaviors. Like mammals, fishes, amphibians, and reptiles may be either viviparous or oviparous, with the exception of the cyclostomes and birds, which are exclusively oviparous. Ovarian features in lower vertebrates are consistent with oocyte development regulated by hormones through both long-loop feedback and local control mechanisms (Pierantoni et al., 2002; Polzonetti-Magni et al., 1995; Polzonetti-Magni, 1999). Among vertebrates, the intrafollicular processes through which the oocytes reach maturity are thus inextricably bound up with those through which yolk is made, and gets to, is incorporated into, and stored in the oocytes; yolk is the oldest and most common means of maternal investment in reproduction, with only a few exceptions among invertebrates and only the Eutheria among vertebrates; all known metazoans reproduce through a system based on yolk formation, and vitellogenin is the principal constituent of yolk in vertebrates. The follicle, in all known vertebrates, consists of the primary oocytes surrounded by one or more layers of epithelial cells, called granulosa cells. During follicle development, a vascularized layer of cells derived from the ovary’s interstitial tissue surrounds the follicle, a layer called the theca interna, whereas in some species, another layer, the theca externa, may be added. Even though an enormous variation in the size and appearance of the ovary exists, the common organization of the follicle in all the vertebrates can be recognized. Ovarian development and maturation require vast metabolic eVorts made through vitellogenesis, both in the eggs discharged into the aquatic environment and in the cleidoic egg; this is clearly a change that made terrestrial life for vertebrates possible, even if it also occurred in somewhat diVerent form in elasmobranches (sharks, rays, and skates). The ‘‘cleidoic egg,’’ as first described by Needham (1931) is an egg enclosed in a protective shell containing all the ingredients necessary for the accomplishment of embryogenesis outside the mother’s body. The spawner that hatches is usually a free-swimming, independently feeding larva; the cleidoic egg that hatches may be either advanced enough in development to feed itself or be dependent in diVerent ways on parental care. To summarize the relationships between vitellogenesis and reproductive strategies, it should be pointed out that yolk formation is the oldest and most common feature by which the mother supplies the zygote for embryonic development in most of the reproductive strategies, with the exeption of viviparity in eutherian mammals. Because viviparity occurs in all vertebrates

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except birds, the origin of this unique characteristic was found to arouse great interest, and the fascinating aspects of this issue have been very well discussed by Rothchild (2003). The principal constituent of yolk in vertebrates is VTG, made in species-specific varieties. Although the eutherian liver does not make VTG, two lipoproteins involved in cholesterol metabolism, as well as the von Willebrand factor, which is involved in clotting, have a portion homologous to the VTG apolipoprotein (169–172). However, the genetic basis for the eutherian liver’s inability to make VTG is still not well understood.

A. Pattern of Vitellogenesis During Seasonal Reproduction in Fish and Amphibian Models Almost all fish from temperate zones have a seasonal reproductive cycle, and reproductive seasonality is controlled by physiological mechanisms that allow the organism to integrate seasonal environmental variations or cues with a temporal control of reproductive events, so as to optimize reproductive fitness from the Darwinian point of view. The evolutionary causes of reproductive seasonality and its physiological control were extensively reviewed in fish by Mommsen and Walsh (1998) and Nash (1999). Vitellogenesis is a crucial point in the reproductive function and occurs during oogenesis, the major regulatory system of which is in the hypothalamus– pituitary gonadal axis. The gonadotropin-releasing-hormone (GnRH) from the hypothalamus stimulates production and release of gonadotropins from the anterior pituitary; these in turn act on the ovary at the level of the follicle wall to elicit steroid release, which controls a variety of processes, including vitellogenesis. In teleosts, the largest vertebrate group, there are synchronous members that reproduce only once and then perhaps die, and thus all their oocytes develop simultaneously; then there are members showing a type of oogenesis in which at least two populations of oocytes at diVerent stages are present, as well as the asynchronous type characterized by multiple populations of oocytes. The members of the last group often breed several times or continuously throughout the year. The process of atresia or oocyte resumption may occur at any time during oogenesis in many teleost species; this process involves hypertrophy of the granulosa cells and a deterioration of the oocytes, although the purpose of this phenomenon is not yet well understood. The diVerent modes of reproduction, including vitellogenesis, have been extensively studied and reviewed by Connaughton and Aida (1999), Lessman (1999), Reinboth (1999). Nevertheless, it seems of interest to focus on the pattern of vitellogenesis in some teleost models, as for instance the killifish, F. heteroclitus, a cyprinid


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euryhaline oviparous teleost that exhibits semilunar spawning cycles in the wild, coincident with the new and full moon. This semilunar pattern of reproduction is an appropriate laboratory condition allowing daily egg collection. Fundulus, in fact, allowed R. Wallace and coworkers to establish the main and most important concepts and knowledge of fish vitellogenesis (see also Cerda` et al., 1996; LaFleur et al., 1995; Wallace and Selman, 1978, 1981, 1985). The gilthead seabream, Sparus aurata, with its asynchronous type of oogenesis, can be considered a good model for representing the asynchronous spawner; it is widely used in intensive aquaculture, with interesting perspectives for applied researches to improve fish food availability. These fish function as males during the first 2 years of life, and in their third year 80% of them switch sex to become females; females spawn daily, releasing eggs on each occasion. Because spawning may be prolonged over a period of months, vitellogenesis is continuous throughout the breeding season (Barnabe´ and Paris, 1984; Mosconi et al., 1998). As found by Carnevali et al. (1992a), vitellogenin uptake starts in oocytes of 200 m in diameter, and oocytes at diVerent stages of vitellogenesis are present in the ovary. Seabream vitellogenin was characterized as a band with an apparent molecular mass of 180 kDa, corresponding to the VTG monomer (Mosconi et al., 1998); then the specific antibody was provided and an ELISA was validated for measuring VTG during the reproductive season, related with the changes of plasma estradiol-17b. In female broodstock, both VTG and estradiol-17b levels were found to be high in prespawning and spawning females, with the VTG titers being parallel with those of estradiol-17b, whereas in postspawning females, the VTG levels decreased dramatically with even relatively high estradiol being present. In this model, it was found that plasma vitellogenin reflects a balance between that produced by the liver and that sequestered by growing oocytes, which places great demands on vitellogenin from blood. Moreover, it is of interest that the termination of vitellogenin synthesis in the quiescent period occurs notwithstanding relatively high peripheral estradiol (Fig. 3). Even though much research has been done to ascertain the onset of vitellogenesis in fish (Ohkubo et al., 2003; Specker and Sullivan, 1994), little or no evidence exists on vitellogenesis termination. Moreover, experiments have been carried out to study in vitro eVects of estradiol-17b, homologous pituitary homogenate, and recombinant red seabream growth hormone (sbGH) on vitellogenin synthesis and secretion in seabream liver fragments (Mosconi et al., 2002b). GH exerted a direct stimulatory action on hepatic VTG secretion; there appeared to be a specific sensitivity to estrogenic compounds and pituitary hormones, depending on the reproductive phase, as estradiol was more potent than sbGH in the prespawning phase, whereas sbGH was found to be more eVective during the spawning season. Moreover,

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FIG. 3 Trend of plasma vitellogenin and estradiol-17b in female broodstock of the gilthead seabream, Sparus aurata.

a positive correlation between estradiol receptor level and liver responsiveness to estradiol-17b was clearly observed, indicating that estradiol upregulates estrogen receptors that may, in part, be responsible for changes in seasonally dependent sentitivity to circulating estrogen level in the seabream model. In fact, estradiol-17b is one important inductor of vitellogenin synthesis, but even though the plasma estradiol-17b titers of males sometimes match or exceed those of females, normally only the females will synthesize and secrete vitellogenin. This protein is normally never detected in untreated males, although the hepatocytes are capable of synthesizing the yolk precursor when stimulated with unphysiologically high doses of estradiol-17b. This aspect has been pointed out in several oviparous species (Carnevali et al., 1992b, 1995; Gobbetti et al., 1985a; Mosconi et al., 1998, 2002b) including the Mixiniformes Lampetra fluviatilis (Mewes et al., 2002). Regarding chondrichthyan fishes, it has recently been shown that in Torpedo marmorata, oocyte growth is not only caused by the uptake of exogeneous molecules but also by the contributions of the oocytes themselves (Prisco et al., 2002); in oviparous dogfish, Scyliorhinus canicula (Craik, 1978a,b,c,d, 1979), and in the little skate, Raja erinacea (Perez and Callard, 1992), vitellogenesis does not diVer fundamentally from that of other vertebrates in the mechanisms of synthesis and storage of vitellogenin. Like fish, most amphibians are oviparous with external fertilization; however, some viviparous species exist in both anurans (frogs) and urodeles


13

(salamanders). Because many amphibians must return to water to breed, water availability is the limiting factor for their reproductive success. Among all amphibian groups, there are species that typically breed once a year, called seasonal/annual breeders, and others that exhibit biannual breeding. Like cold-blooded vertebrates, amphibians are well-known for their dependence on environmental factors to lay eggs. Although most temperate zone populations are known to have one ovarian cycle per year, populations inhabiting warmer zones may show no seasonality, and females can breed whenever there is suYcient rainfall; many species inhabiting tropical, unpredictable, and aseasonal environments show opportunistic breeding at any time of the year. Temperate zone amphibians primarily breed in early spring, oogenesis occurs in summer, and the ovary is quiescent in winter. Therefore, the cyclical changes of amphibian ovary must be discussed in the frame of their seasonality, bearing in mind the role of environmental cues influencing the reproductive cascade (i.e., the hypothalamus-pituitary-gonadal axis, by which oocyte maturation and vitellogenic process are allowed). The functional units of the ovary are the follicles, each consisting of a single ovum surrounded by granulosa cells and by the theca cells synthesizing the androgen precursor used by granulosa cells for estrogen synthesis. Follicles of diVerent size are present in the frog ovary during the recovery phase. Oocyte yolk proteins (YPs) are present in structures that have a somewhat geometric shape. During oocyte growth, the small nascent platelets increase in size by fusion with one another, and the mature egg contains a heterodisperse size range. In addition to increasing in size, YPs increase in number, and oocytes sampled during the recovery phase show a thin granulosa layer containing follicle cells and a large nucleus provided with numerous nucleoli. The relationships between vitellogenesis and changes in peripheral hormones were studied by Polzonetti-Magni et al. (1998) during the annual reproductive cycle of wild population of Rana esculenta living in a mountain pond at 820 m above sea level (43 030 N, 12 550 E) (Fig. 4). This wild population breeds in late spring (May) and then, particularly in females, a kind of postreproductive refractoriness intervenes, even if environmental conditions are still favorable. Early in autumn, recrudescence begins, characterized by intense vitellogenesis, and it continues until the long winter rest (Polzonetti-Mangi et al., 1990). In that study, a clear correspondence between plasma FSH and estradiol-17b was shown. In females, FSH peak values were found at the beginning of the mating period in parallel with those of plasma vitellogenin and estradiol-17b; in contrast, high LH levels went together with ovarian weight (gonadosomatic index), which is considered a good marker for the plasma sequestration of vitellogenin by growing oocytes. The in vivo results were corroborated by in vitro studies showing the direct eVects of both FSH and LH in inducing hepatic

14


FIG. 4 The relationships between environmental factors and changes of reproductive hormones in the wild population of the frog Rana esculenta.


15

VTG synthesis and release in culture media. In that frog, gonadotropins play a role in the vitellogenic process that consists in the induction of hepatic vitellogenin synthesis and in vitellogenic sequestration by growing follicles. Moreover, the direct eVects of gonadotropins assessed by in vitro experiments, as previously shown using growth hormone and prolactin (PolzonettiMagni et al., 1995), were seasonally dependent; in fact, during recrudescence, only LH was able to induce VTG in the liver. The results obtained on the eVects of pituitary hormones on liver vitellogenin synthesis have been to a large extent accounted for by the contributions of Sakae Kikuyama’s laboratory, in which homologous pituitary hormones were purified and homologous RIAs validated (Kobayashi et al., 1989; Takahashi and Hanaoka, 1981; Yamamoto and Kikuyama, 1981; Yamamoto et al., 1995). Evidence of the involvement of pituitary hormones in vitellogenesis came from experiments performed by Gobbetti et al. (1985a,b), who found that hypophysectomy influenced the vitellogenic process in terms of synthesis and ovarian uptake. Then this knowledge was further sustained by various evidence obtained by both in vitro and in vivo experiments (Carnevali and Mosconi, 1992; Carnevali et al., 1992b, 1993b; Mosconi et al., 1994a,b). When peripheral GH was assessed during the annual reproductive cycle, the two main peaks of GH were found at the breeding period and during the autumn recrudescence, and they were related with high vitellogenin levels in the blood, whereas the highest estradiol-17b plasma levels occurred during the summer refractory period, when vitellogenin concentration is basal or not detectable. Therefore, as pointed out above in S. aurata, the pattern of vitellogenin displays seasonal changes not very related with those of estradiol-17b. In addition, when both types of hormones, estradiol and pituitary, were able to induce in vitro vitellogenin synthesis in the liver, the trends of induction were found to be diVerent, with no synergism between these hormones being present, which indicates diVerent types of hormonal mechanisms operating in the liver (Polzonetti-Magni et al., 1995). Regarding the GH eVects, Carnevali et al. (1995) demonstrated the ability of this hormone to induce, as does estradiol-17b, transcription of the VTG gene both in vivo and in vitro in both sexes; the levels of mRNA were related to those of protein, and seasonal variations were found to occur in the VTG gene transcription under GH and estradiol-17b treatment. Indeed, the more active inducer was GH during the reproductive period and estradiol-17b during the prereproductive phase. Regarding the mechanism of action, the stimulatory eVects of estradiol-17b in the liver have been described in X. laevis, and VTG mRNA transcription seems to be under the control of EREs (Martinez et al., 1987). In the GH mechanism of action, this hormone works through a local production of IGF-I, indicating the involvement of estrogen receptor in VTG induction by IGF-I. Furthermore, the eVects of

16


GH resulted in an increase of cAMP and the appearance of a new phosphotyrosine protein, indicating the occurrence of tyrosine kinase activation (Carnevali et al., 2000). This finding strongly suggests the role played by pituitary hormones in the vitellogenic process, at least in the frog and seabream models; moreover, in the urodele newt, Triturus carnifex, prolactin and GH, as well as estradiol17b, were involved (Mancuso et al., 1996; Zerani et al., 1991) in the vitellogenin process; last, the role of fat bodies should be considered in anurans (Carnevali and Mosconi, 1991).

B. Acquirement of Terrestrial Habitat Through the Reptile Amniotic Egg The amniotic egg that made terrestrial life for vertebrates possible is presumed to have been the reproductive strategy of Mesozoan diapsids, and in fact the fossil eggs of dinosaurs are relatively common in some deposits. The major diVerences between the anamniotic and amniotic egg is consistent with the presence in the latter of three extraembryonic membranes, the chorion, the allantoid, and the amnios. These membranes surround the embryo, which can complete its development and diVerentiative program in an aquatic medium, as fish spawners do, protected by the shell. Reptilian reproduction was reviewed by Lance (1999a,b) and Crews (1999). The primary follicles consist of enlarged oocytes surrounded by a single layer of granulosa cells, with the follicles being generally spherical and arranged in size classes. The developing follicles are elongate rather than spherical and go through a growth stage to reach the previtellogenic phase, when they have a translucent white color; when vitellogenesis proper begins, the follicles take on a yellow color. Apart from being oviparous species, reptiles show marked viviparity, and vitellogenesis in the viviparous lizard was investigated in depth by Gavaud (1986a,b) and Guillette (1985, 1987). In the oviparous model, Podarcis s. sicula Raf., vitellogenin was purified, the native form was present at two polypeptide bands, and the monomeric form in SDS-PAGE was found at about 200 kDa. The specific antibody was raised, and a validated ELISA was applied to measure peripheral VTG together with estradiol-17b during the annual reproductive cycle (Carnevali et al., 1991b). In that work, it was found that plasma vitellogenin levels presented two distinct peaks (in April and June), which coincided with ovarian weight peaks and with those of two ovulatory waves observed over the same period. Plasma estradiol-17b was also measured over the same period to investigate whether the seasonal changes were correlated with the plasma vitellogenin trend (Fig. 5). The


17

FIG. 5 Pattern of vitellogenesis in the oviparous lizard Podarcis s, Sicula Raf.

two peak values recorded for plasma estradiol support the hypothesis that this hormone may regulate hepatic vitellogenin synthesis and, in turn, ovary development. Interestingly, although spring and summer VTG peaks were approximately the same, the corresponding peaks of plasma estradiol-17b were much smaller in summer than in spring. Such a lack of correlation again poses certain questions that can be expressed in several ways: are there ‘‘memory eVects,’’ as suggested by Wallace and Bergink (1974) in Xenopus, and by Ho et al. (1982a,b) in turtle, or do changes in liver sensitivity occur during the season? It is also of particular interest that during summer refractoriness, while VTG titers decline, plasma estrogens remain at a consistent level, as mentioned above for seabream and frog. As previously suggested by Ho et al. (1982b), other hormones than estrogen also can play a role in hepatic vitellogenin production in this oviparous squamate model.

IV. Hormones Regulating Vitellogenin Synthesis On the basis of several data reported here, it is diYcult not to emphasize the high seasonality in the vitellogenesis pattern shown in fish, amphibians and reptiles; in the animal models considered, a kind of refractoriness intervenes

18


in the postreproductive period, when the liver is unresponsive to estradiol17b, whereas pituitary hormones are able to induce vitellogenin synthesis directly. Notwithstanding, there is no doubt that estradiol-17b can be considered a potent hormone in inducing hepatic vitellogenin synthesis, despite the abovementioned evidence for multihormonal control related to the diVerent reproductive strategies, and mainly to the timing of vitellogenic process. Estrogens have diverse eVects on many tissues in both males and females, and it is believed that the majority of these eVects are mediated by estrogen receptors (ERs), which function like growth factors in some way. In addition, estrogens are widely distributed in all vertebrates, and estrogen-like molecules work in several diVerentiative patterns in invertebrates. Regarding pituitary hormones, evidence exists for the role played by them in several fish, amphibians, and reptiles. In fish, even if VTG synthesis in hepatocytes has been found principally to be the direct action of estradiol17b, which is produced in the ovarian follicles under regulation of a gonadotropin (Nagahama, 1994), and administration of estradiol-17b to immature and male fish has induced VTG accumulation in their blood circulation (Mommsen and Walsh, 1998), in vitro experiments using immature eel hepatocytes revealed that growth hormones or prolactin were required for induction of VTG, when physiological doses of estradiol-17b were added to the medium (Know and Mugiya, 1994; Peyon et al., 1996). Moreover, by in vitro studies, the direct eVects of GH in inducing hepatic vitellogenin synthesis were ascertained in seabream liver, where estradiol-17b failed to induce vitellogenin synthesis in the postspawning season (Mosconi et al., 2002b). Pituitary hormones also work directly in the liver of the anuran, R. esculenta, and their eVects with respect to those exerted by estradiol were found to be dependent on sex and season (Polzonetti-Magni et al., 1995). In reptiles, estradiol was synergized by pituitary factors in the iguanid lizard Diploscures dorelis and in the turtle Chrysemys picta (Callard et al., 1972; Ho et al., 1982a,b, 1985). As far as steroid hormones are concerned, it has been found that combined treatment of estradiol-17b and either progesterone or cortisol enforced expression of VTG mRNA in the hepatocyte culture of immature rainbow trout, Oncorynchus mykiss, although cortisol alone failed to induce this expression (Mori et al., 1998). However, it has been suggested that cortisol induced rapid and transient transcription of VTG mRNA in the tilapia, Oreochromis aurens (Ding et al., 1994). Moreover, the involvement of androgens in VTG synthesis has also been reported in certain teleost fish (Hori et al., 1979; Kim and Takemura, 2002; Le Menn et al., 1980; Peyon et al., 1997). In vivo experiments showed that VTG was induced in the blood circulation after 17amethyltestosterone and 5a-dihydrotestosterone treatment in the goldfish Carassius auratus (Hori et al., 1979) and in Gobius niger (Le Menn et al., 1980). Although androgens or testosterone, androsterone, and 17a-methyltestorenone


19

induced in vitro the expression of VTG mRNA in rainbow trout (Mori et al., 1998), its oral administration inhibited both expression of VTG mRNA in the liver and the appearance of VTG in the circulation in Oreochromis niloticus (Lazier et al., 1996). Moreover, biochemical changes that occur in the liver during the induction of vitellogenin synthesis must be taken into account (Duggan and Callard, 2003; Sehgal and Goswami, 2001). For a long time, attention has been addressed, mainly in the laboratory of Ian Callard, to the role played by progesterone in vitellogenesis regulation through the studies carried out in the little skate, R. erinacea and in the painted turtle C. picta. Progesterone is defined as the ‘‘pregnancy hormone’’ of mammals. However, in lower vertebrates, it is a highly versatile hormone that exerts an ample variety of eVects on diversified target organs (Paolucci et al., 1998). In early studies in Callard’s laboratory, it was found that progesterone inhibited seasonal and gonadotropin-induced ovarian growth and estradiol-induced vitellogenin synthesis; moreover, the hypothesis that progesterone may be responsible for the decrease of VTG gene expression that accompanies the evolution of viviparity in vertebrates, as well as the complete inhibition associated with the fully developed mammalian chorioallontoic placenta (Callard et al., 1994), was proposed. Such a role of progesterone has been widely documented by the evidence of the presence of progesterone receptor isoform in the turtle (Custodia-Lora and Callard, 2002) and in the liver of subavian species (Paolucci et al., 1998). The view of Callard et al. (1994) on the role of progesterone as a cause of the loss of vitellogenesis, and to a large extent of the transition to eutherian viviparity, is both very fascinating and well documented in reptiles, in which a postovulatory progesterone peak was also found in the Scincidae lizard (Angelini et al., 1986; Mosconi et al., 1991). On the contrary, Rothchild (2003) suggested that the loss of vitellogenesis in Eutheria was associated with more fundamental causes, probably mutations, and the ability of progesterone to suppress VTG gene expression was considered to be dependent on season and stage of reproductive cycle (Ho, 1991; Polzonetti-Magni et al., 1995). If the origin of viviparity was genetically dependent, as Rothchild suggested (2003), vitellogenin synthesis termination, a process mimicking the loss of vitellogenesis in eutherian mammals, can be perhaps related with a genetically controlled process such as apoptosis. This fundamental process plays a key role in the tissue remodeling occurring during embryonic development and metamorphosis, and in normal tissues it is a key event in the regulation of cell number. This hypothesis is sustained by the work of Assisi et al. (1999), in which it was found that the physiological involution occurring postvitellogenesis in frog liver takes place by programmed cell death, together with the concomitant induction of transglutaminase gene expression, regulated by sex hormones and induced and activated in

20


mammalian apoptotic liver. The role played by sex hormones was also emphasized because circulating levels of estradiol-17b, progesterone, and testosterone increased before the hyperproliferative phase of the liver and their levels dropped in coincidence with the mating season, causing the induction of programmed cell death that leads to involution of the liver mass. This scenario could be convincing even if, as shown above, in the animal models presented here, the levels of sex hormones were high in the postreproductive season, when vitellogenin synthesis was completely terminated. Thus, in addition to long-loop feedback mechanisms, the presence of local regulation of the liver function, perhaps responsible for vitellogenesis termination, can be taken into account. Regarding the possible local regulatory pathways, the GH and IGF system can be considered a good candidate; moreover, interesting results have recently been obtained on the paracrine role of GnRH in the regulation of fish gonadal apoptosis, the eVects of which are dependent on the doses applied and the duration of treatment (Polzonetti-Magni, A.M., Mosconi, G., and Palermo, F., unpublished data). Last other kinds of nonconventional regulatory mechanisms of vitellogenin synthesis have been proposed; Nath and Maitra (2001) found that vitellogenin itself regulates its own synthesis and incorporation in the catfish because heterologous vitellogenin isolated from the Indian major carp, when administered either in semipurified or purified forms, induced complete vitellogenesis in the female catfish. Moreover, a similar regulatory mechanism was also suggested by Reis-Henriques et al. (2000), who found that the in vivo and in vitro vitellogenin concentration inside the oocyte can alter vitellogenin production in the liver by modulating the synthesis of ovarian estradiol-17b; high levels of VTG in the oocyte could in turn suppress the enzymatic chain reactions leading to estradiol-17b biosynthesis.

V. Vitellogenin Utilization The transformation of oogonia into oocytes is commonly described as oogenesis; in many oviparous vertebrates, the prodigious growth of oocytes from microscopic to macroscopic dimensions is mainly the result of one of the most exciting examples of cell regulation, namely vitellogenesis. This process, common to all oviparous vertebrates species, is characterized by hepatic production of a glycolipoprotein, vitellogenin (VTG), under multihormonal control, and its transportation via the bloodstream to the ovary, where it is internalized into the oocytes. The uptake and processing of VTG into developing oocytes was first observed in X. laevis by Wallace and Jared (1969) using in vivo studies with labeled VTG. In fish, some of the first


21

evidence indicating that VTG sequestration represents the major mechanism for oocyte growth was obtained by Tyler et al. (1988). The uptake of VTG into growing oocytes occurs by receptor-mediated endocytosis. Receptors for VTG (VTGR) have been identified at the oocyte surface in several vertebrate groups including birds, amphibians and fish. In fish, for Oreochromis niloticus, L 1766, Chan et al. (1991), using saturation studies and Scatchard analyses, found only a single class of binding sites corresponding to specific receptors; the number and the aYnity (Kd) of these receptors increased from the previtellogenic to vitellogenic stages and remained unchanged at the preovulatory stage, during which aYnity was found highest. However, as observed in other oviparous vertebrates, aYnity was relatively low, which may be accounted for by the high circulating level of vitellogenin in teleosts (Copeland and Thomas, 1988; So et al., 1985). In contrast, no change in Kd was noted in the course of vitellogenesis in the white perch (Tao et al., 1996). Avian and amphibian VTG receptors have been shown to be related both immunologically and at the level of ligand recognition. A combination of genetic and biochemical analyses, including protein purification, demonstrated that chicken oocyte VTGR is also the receptor for other major yolk precursors, very low density lipoproteins (Stifani et al., 1990a,b). Receptors that recognize VTG interact with (a) site(s) located on the lipovitellin portion of VTG, and specifically within the NH2-terminal domain, referred to as lipovitellin I (Stifani et al., 1990a,b; Wahli, 1988). However, by in vitro binding at acidic pH, using the PV region in chicken oocyte, Cutting and Roth (1973) demonstrated that PV was recognized and endocytosed by the developing chicken oocyte. The uptake of VTG into oocytes of the rainbow trout, O. mykiss, is under the control of GtH I. Its injection into maturing vitellogenic females at a dose of 10 mg/kg body weight increased the rate of [3H]VTG uptake more than twofold, eVectively doubling its rate of growth. In vitro, GtH I stimulated VTG uptake in a dose-dependent manner. At a GtH I concentration of 100 ng/mL and above, the rate of VTG uptake was significantly greater than that of the controls, and at 1000 ng/mL it was more than doubled. On the contrary, GtH II did not increase VTG sequestration either in in vivo or in vitro experiments (Prat et al., 1998). Xenopus oocytes exhibited enhanced rates of VTG endocytosis following exposure to insulin in vitro and to human chorionic gonadotropin in vivo. The stimulation of VTG uptake was not caused by an increase in surface VTG receptors. Instead, both hormones acted by stimulating the specific internalization rate of the VTG receptor, although by apparently diVerent mechanisms. Stimulation of the internalization rate by insulin was most obvious at low levels of receptor occupancy. Insulin also increased the aYnity of the VTG receptor for its ligand. The steady-state binding of VTG was biphasic with respect to increasing ligand

22


concentration, primarily because of the use of a nonlinear receptor internalization rate as a function of occupancy, as in models in which there is a cell surface regulatory component that facilitates the internalization of the occupied VTG receptor (Opresko and Wiley, 1987). Once vitellogenin, via a process of receptor-mediated endocytoses, is sequestered by growing oocytes, it is proteolytically cleaved into smaller yolk proteins (Carnevali et al., 1991a, 1992a; Selman and Wallace, 1989; Specker and Sullivan, 1994) and stored as a nutrient required for developing embryos. In amphibians, VTGs give rise to at least two major classes of yolk proteins, an extensively lipidated lipovitellin (LV) and a highly phosphorilated one, PV (Wallace, 1985), and smaller phosphoproteins termed ‘‘phosvettes’’ (Wiley and Wallace, 1981), to form an insoluble matrix packaged in crystals. In the barfin flounder, Verasper moseri, a marine teleost that spawns pelagic eggs, two forms of LV were electrophoretically and immunologically identified in postvitellogenic oocytes; these two forms correspond to the two VTGs (VTGA and VTGB) and are identified in the plasma (Matsubara et al., 1999). One of the catalysts for the intraocytic processing of VTG in yolk proteins is cathepsin D. The involvement of a lysosomal enzyme such as cathepsin D in VTG proteolyses was first demonstrated in tetrapods (Opresko and Karpf, 1987; Yoshizaki and Yonezawa, 1994; Yamamura et al., 1995). In rainbow trout, O. mykiss, an immunocytochemical study of the oocyte showed that cathepsin D is localized in multivesicular bodies that begin to diVerentiate before the phase of vitellogenesis, during which endocytosed VTG is colocalized with cathepsin D in the multivesicular bodies (Sire et al., 1994). The involvement of cathepsin D in the proteolysis of VTG was also demonstrated in seawater fish; in seabream, its enzymatic activity, tested during the diVerent oocyte maturation phases, evidenced the maximum level in early vitellogenesis, when the deposit of yolk proteins was very intense (Carnevali et al., 1999a). In this species, the in vitro approach clearly demonstrated the ability of cathesin D to convert seabream VTG to yolk proteins (Carnevali et al., 1999b). Apart from the initial processing of VTG, the occurrence of additional proteolysis of the yolk proteins during the final oocyte maturation was first observed in F. heteroclitus (Selman and Wallace, 1989; Wallace and Begovac, 1985). This second proteolysis of VTG-derived proteins is unique to teleosts (Byrne et al., 1989) and occurs particularly in marine or brackish-water fishes, which exhibit remarkable hydration of their oocytes during final maturation (Carnevali et al., 1992a; Matsubara and Sawano, 1995; Matsubara and Koya, 1997; Thorsen et al., 1996). Regarding marine pelagic eggs, they contain a high quantity of free amino acids relative to demersal eggs. The increase in free amino acid content is thought to play a significant role in generating the osmotic gradient responsible for water influx during oocyte maturation and hydration (Thorsen et al., 1996). In


23

the barfin flounder, the proteolysis was observed in all three classes of VTGderived yolk proteins during oocyte maturation (Matsubara and Koya, 1997). Similar evidence was obtained in the seabream and sea bass, where the proteolysis was observed both in lipoproteins and phosvitins (Carnevali et al., 1992a, 1993a). In seabream, the hydration process occurring at the end of oocyte maturation is consistent with the reduction of the acidophilic areas of yolk globules and is related to molecular variation in the constituent proteins of yolk (Carnevali et al., 1992a, 1993a). The enzyme responsible for this second proteolytic process was identified in this fish (Fig. 6). From the enzymatic profile of growing oocytes, very high levels of a cystein proteinase, cathepsin L, were revealed in midvitellogenic oocytes. The purification of cathepsin L from the seabream ovary and the assessment of its capability to cleave the LV components by the in vitro system evidenced the ability of this lysosomal enzyme to process the LV components. In fact, after 1 hour of incubation, the highest-molecular-weight LV bands disappeared and digestion was completely abolished by the addition of the cathepsin L specific inhibitor, leupeptin (Carnevali et al., 1999a,b). The involvement of cathepsin D and L in the vitellogenic process was also observed in trout, with a diVerent expression of cathepsin D mRNA being found during oocyte maturation, whereas the maximum expression preceded the period of maximum enzyme activity in rainbow trout (Brooks et al., 1997). The role of cathepsin D in the generation of egg yolk proteins was also demonstrated in chicken by Elkin et al. (1995). Given the large content of lipids in the growing oocyte, high levels of lipoprotein proteinases (LPL) were also well documented in both freshwater and seawater fish (Kwon et al., 2001; Prat et al., 1998). In trout, both LPL and cathepsin B are exclusively expressed during oogenesis (Kwon et al., 2001), and the latter cysteine proteinases seem also to be involved in oocyte maturation both in freshwater (Kwon et al., 2001) and seawater fish (Carnevali et al., 1999a,b). The diVerent pattern of cathepsins and lipase expression during oogenesis indicated temporal roles in yolk protein processing (Kwon et al., 2001). In addition, these data indicate that, in the oocyte, the lysosomal system contains the whole set of hydrolases necessary for yolk formation and degradation, which is in striking contrast to classical lysosomes that rapidly digest any protein down to free amino acids; instead, lysosomes in oocyte cells do not degrade the yolk globule content until a specific developmental stage is reached (Fagotto, 1995). In many marine teleosts, egg buoyancy at spawning is a result of their exceptionally high water content. The acquirement of buoyancy by eggs through the hydration process is accompanied by NaþKþ and free aminoacid accumulation. The inhibition of the vacuolar ATPase-dependent proton pump by bafolomycin A1 in black sea bass suppressed hydration, and Selman et al. (2001) hypothesized a two-step

24


FIG. 6 Role of Cathepsin D and L in vitellogenin cleavage and hydration process occurring in seabream pelagic eggs.

process: Kþ influx (promoting the yolk sphere fusion), and acidification and proteolysis of yolk (promoting the acquirement of water). The elucidation of the mechanism through which oocytes become hydrated is of great interest because it represents a key event of reproduction and, for the genetics of a population, is closely related with the dispersion of eggs. An incorrect hydration process produces sinking eggs unable to develop a viable embryo (Carnevali et al., 2001a,b). Once vitellogenesis is terminated, in Fundulus, the responsiveness of the follicles undergoing oocyte maturation in vitro after gonadotropin and maturation inducing steroid (MIS), 17,20bdihydroxy-4-pregnen-3-one, declines dramatically in oocytes, reaching 1.8– 2.1 mm in the ovary, which indicates that full-grown follicles may be recruited into maturation by diVerent sensitivity to the MIS (Cerda` et al., 1996).


25

The egg contains all the components necessary for the embryo, including all the morphogenetic factors, and the yolk constitutes the main storage compartment and the main food supply for embryonic nutrition. During embryonic development, the materials that accumulated in the oocytes, energy sources that are rapidly degraded to produce free amino acids, fatty acids, phosphorous, calcium, and others, are used by the growing embryo. For yolk degradation, the whole set of enzymes is present in the cytoplasm, and inhibition with cysteine and aspartic proteinase inhibitors results in a blocking of yolk proteolysis and abnormal development or mortality of embryos. In Xenopus, the involvement of cathepsin D and L in yolk utilization was first observed by in vitro experiments (Yoshizaki and Yonezawa, 1996, 1998). These authors found that the limitation to proteolysis of VTG by cathepsin D in amphibians was caused by the insolubility of yolk proteins at physiological salt concentrations. In the sea urchin, during early development, the yolk platelets remain unchanged in terms of phospholipid triglyceride, hexose, sialic acid, total protein, and so forth. However, the platelet is not entirely static because the major yolk proteins undergo limited stepwise proteolytic cleavage during early development. When the yolk platelets become acidified during development, the activation of a cathepsin B-like yolk proteinase occurs because of the degradation of the major yolk protein (Mallay et al., 1992). In the sea bass Dicentrarchus labrax, the degradation of yolk protein, accumulated and stored in the ovaries, is tightly controlled and involves diVerent types of lysosomal enzymes. The serine proteinase, cathepsin B, was found at maximal level during the early stage of development, when no modification of the electrophoretic pattern of yolk proteins occurred. The maximal activities of cathepsin A and L were found in segmentation and are related with yolk protein electrophoretic changes. At the hatching stage, when choriolytic enzymes are produced, cathepsin D reached its maximal level in parallel with changes in the yolk protein electrophoretic pattern (Carnevali et al., 2001a,b). Proteolytic processing of yolk was also well documented in insects, where cysteine proteases were implicated in vitellin processing during embryo development (Cecchettini et al., 2001; Giorgi et al., 1997).

VI. Vitellogenin as Biomarker of the Feminization Process Induced by Environmental Estrogens Environmental estrogens are chemical compounds mimicking estrogen eVects, and are widely considered to be endocrine disrupters (EDs). One major objective in recent ecotoxicological research is to identify anthropogenic chemicals within the aquatic environment that interfere with

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hormonal regulatory pathways of aquatic organisms. Special attention has been addressed to substances that mimic the action of the female sex hormone, estradiol-17b, which has been found to induce morphological and functional alterations of the reproductive tract, primarly in male individuals. Alkylphenolic compounds, such as nonylphenols (NPs), which reach the acquatic environment as a degradation product of nonylphenol ethoxylates, represent one group of industrial chemicals that has been shown to possess estrogenic proprieties in vitro (Jobling and Sumpter, 1993; Polzonetti-Magni et al., 2004; White et al., 1994) and in vivo (Jobling et al., 1996; Mosconi et al., 2002a). Endocrine disrupter chemicals, such as NP, are defined as exogenous substances causing adverse health eVects in an intact organism or its progeny, secondary to changes in endocrine function. The list of chemical compounds behaving as endocrine disrupters has recently been reported in the review by Larkin et al. (2003). Some eVects attributed to these contaminants include reduced fertility and viability of oVspring, as well as impaired hormone activity and altered sexual behavior. EDs are also suspected to aVect human health by increasing the rate of breast cancer (Davis et al., 1993) and other endocrinological diseases. This kind of pollution certainly induces a decrease in fertility, and it would be expected to have catastrophic eVects on a wild population. An assessment of pollutioninduced reproductive dysfunction, however, requires a simple biomarker. Because vitellogenin is present at high concentrations in the plasma of maturing females and is easily measured by radioimmunoassay or ELISA, it may provide a suitable biomarker for environmental pollution. Vitellogenesis involves ovarian estradiol, stimulating the liver to produce vitellogenin, which in turn is incorporated into the developing oocytes. Estrogenic xenobiotics can also act on the hepatic receptors to induce synthesis of vitellogenin, the sex-related protein, because although both male and female adults, as well as immature juveniles, have hepatic estrogen receptors, only the livers of females normally respond to estrogens. Production of vitellogenin by males and juveniles or nonvitellogenic females can therefore provide a bioindicator of exposure to environmental estrogens (Kime et al., 1999) (Fig. 7). In addition, exposure to estradiol-17b results in the synthesis of specific proteins required for reproduction, and several genes that encode proteins induced by this process are, in addition to vitellogenin, the estrogen receptor and choriogenins, which are required for making the egg membrane in fish (reviewed by Larkin et al., 2003). With regard to vitellogenin, exposure to an estrogen mimic increases also the number of estrogen receptors. In fish, the two ERs, a and b, alone or in combination may control diVerent subsets of genes; furthermore, estrogen mimics may bind diVerently to those receptors, acting as agonist and antagonist, giving rise to a very complicated issue in


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FIG. 7 Scheme depicting endocrine disrupters mimicking estrogenic eVects using vitellogenin as biomarker.

researches devoted to the risks of environmental exposure. Recently, various methodologies that can be used to understand at the gene level the consequences of exposure to EDs have been extensively discussed in the review by Larkin et al. (2003); they include Northern blotting, quantitative real-time PCR, diVerential display RT-PCR, and gene arrays. Moreover, two novel biomarkers in the assessment of the eVects of estrogenic and antiestrogenic chemicals in fish have been proposed by Carnevali and Maradonna (2003), who found that cathepsin D (CAT D) gene and its enzymatic active protein could be induced by NP in G. niger and that HSP70 gene expression was also significantly increased.

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Summarizing this issue, biomarkers are valuable tools for estimating the exposure of organisms to environmental pollutants, and the accurate measurement of vitellogenin in male individuals is widely adopted in aquatic vertebrates and invertebrates (RiVeser and Hock, 2002). Therefore, ecotoxicological research needs the help of comparative endocrinology for the development of methods for detecting those substances that indirectly cause adverse eVects in animals by influencing their endocrine system without significant direct eVects (Arnold and McLachlan, 1996; Colborn et al., 1993).

A. Fish and Amphibians as Animal Models Fish represent the largest and most diverse group of vertebrates, and they provide an excellent model for assessing the eVects of contaminants on a biological function such as reproduction. In addition, their intimate association with the aquatic environment makes them an excellent early warning system for environmental health problems that could potentially lead to human health concerns. Most reports of EDs concern feminization phenomena in all classes of vertebrates that may be caused through estrogenic compounds. The growing public interest in the eVects of EDs in aquatic ecosystems was triggered by the work of Sumpter and colleagues that demonstrated that male rainbow trout caged in sewage eZuents were feminized (Purdom et al., 1994). The biomarker used for determination of estrogenicity was VG (Jobling and Sumpter, 1993; Sumpter and Jobling, 1995). In this context, the studies undertaken on vitellogenesis in the last decade were mostly addressed to fish and amphibians as vertebrates representative of the aquatic environment. This issue contributes to this research field, because such eVorts will lead to an increase in current knowledge about the mechanisms regulating vitellogenesis. Hence, methods for purifying vitellogenin from plasma, generating antibodies, and performing assay measurements for vitellogenin were recently validated (Denslow et al., 1999; Shi et al., 2003). In fish, vitellogenin assay was used as a useful biomarker for the detection of estrogenic properties associated with certain compounds, and a doseresponse relationship of the in vivo estrogenicity of such compounds was established in diVerent species. In the liver of rainbow trout, Anderson et al. (1996) investigated the modulation of estradiol-17b-induced vitellogenin synthesis and estrogen receptor by b-Naphthoflavone, which induces the synthesis of the oxidase cytochrome P4501A1. Vitellogenin was induced in rainbow trout also by exposure to 17aethinylestradiol, the main active component of contraceptive pills that is


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more resistant to breakdown than natural estrogen (Verslycke et al., 2002). Again in rainbow trout, NP acted as a weak estrogen in directly exposed adult males, but reproductive success was reduced, as indicated by decreased hatching rates, and a transgenerational eVect mediated by the endocrine system was detected in the oVspring of exposed fish (Schwaiger et al., 2002). The estrogenic response expressed as the induction of vitellogenin synthesis was related in rainbow trout, by Lindholst et al. (2000), to the quantification of internal liver and muscle concentrations of nonmetabolized bisphenol A at the end of the exposure period. This approach seems of great interest because, through this kind of study, a dose–response relationship was established between internal liver concentration of xenoestrogen and the corresponding vitellogenin responses. Moreover, another aspect to be more extensively investigated is an evaluation of the eVects produced by increasing vitellogenin at a physiological level, consisting of a search for the relationship between estrogenic action and homeostasis regulation both in laboratory and wild animals. In this direction, eVorts have been addressed to evaluating the eVect of xenobiotics on reproduction in general, and a biomarker consisting of eggshell zona radiata proteins was found by Celius and Walther (1998) to be a more sensitive parameter, compared to vitellogenesis, for monitoring reproductive eVects of xenoestrogens in Atlantic salmon. To find useful species for monitoring studies on environmental estrogens, estrogen-receptor mRNA (ER mRNA), together with vitellogenin and peripheral sex steroids, was applied to goldfish exposed to 4-nonylphenol (4NP). Treatment with estradiol-17b and 4-NP at concentrations of 107M and 106M induced in the male goldfish a presence in the blood circulation of VTG that was higher compared with that found in animals treated with estradiol-17b. Moreover, 4-NP treatment also induced changes in peripheral sex steroids, and as found in frog and newt, estradiol-17b titers rose in treated male goldfish, perhaps depending on aromatase activation. In those experiments, hepatic estrogen receptor expression was evaluated using RT-PCR methodology, and an increase of Erb1 mRNA expression was found in male goldfish treated with both estradiol-17b and 4-NP (Polzonetti-Magni, A. M., Mosconi, G., and Palermo, F., unpublished data). In fact, the endocrine and reproductive eVects of xenoestrogens are believed to be caused by their ability to mimic (Soto et al., 1991; Sumpter and Jobling, 1995) or inhibit the eVects of endogenous steroid hormones (Toppari et al., 1996; Tyler et al., 1998), to antagonize the eVects, and to disrupt their synthesis and metabolism of hormone receptor (Gillesby and Zacharewski, 1998; Sonnenschein and Soto, 1998; Zacharewski, 1997). Xenoestrogens act at various cellular levels: in binding with nuclear estrogen receptor in target cells (Miodini et al., 1999), in the activation of ER, in nuclear translocation, in the binding of the activated receptor complex to specific DNA motifs-ERE (Weigel, 1996), in the transcription

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of estrogen-dependent genes (Bennetau-Pelissero et al., 1998), and in altering steroid metabolism (Noaksson et al., 2003). As discussed above, the development of tests for detecting endocrinedisrupting chemicals is currently an area of high research priority in all countries in which public institutions have engaged in the task of identifying appropriate experimental goals for evaluation of endocrine-disrupting toxicity to humans and wildlife (Ankley et al., 1997, 1998; Kavlock et al., 1996; Mendes, 2002). Up to now, a well-established marker for exposure of oviparous species to estrogenic chemicals is the induction of VTG levels in males (Harries et al., 1997; Heppell et al., 1995; Mosconi et al., 2002a). Therefore, the VTG ELISA has been applied in both freshwater (Casini et al., 2002; Parks et al., 1999; Rankouhi et al., 2002) and marine fish (Korsgaard and Pedersen, 1998; Madsen et al., 2002). The VTG assays developed to date have mainly focused on fish as the model, but amphibians also need to be represented in studies of toxicity. Amphibians appear to be at high risk from environmental perturbations, as evidenced by a worldwide decline in amphibian populations (Kiesecker et al., 2001). Although the cause of amphibian disappearance is unknown, environmental toxicants, including endocrine disrupters, have been suggested to be among the factors involved (Carey and Bryant, 1995; Fagotti et al., 2004). Moreover, amphibians spend a large part of their life in the water, as spawning and embryonic development are strictly dependent on water availability. Therefore, studies were performed on the exposure of amphibians to estrogenic chemicals and, again in those animal models, VTG assay in the male was used as a biomarker. The interesting findings on this issue were reviewed by Palmer et al. (1998) and by Kloas (2002), and X. laevis was chosen as amphibian model in both in vivo and in vitro experiments. In this frog, disturbances of reproductive pattern have also recently been evidenced (Pickford and Morris, 2003). Because, as explained above, the most important goal of this kind of research is the evaluation of the eVect of estrogenic compounds in the reproductive cascade, wild amphibians must be taken into account. Therefore, a research program was set up using wild populations of frog, R. esculenta, and newt, Triturus carnifex, living in a mountain pond for which previous studies have described the reproductive seasonal cycles (Polzonetti-Magni et al., 2004). Validated ELISAs for VTG were obviously applied to evaluate in a dose–response manner the eVects of the estrogenic compound, 4-NP, in males. The combined in vivo and in vitro experiments allowed us to identify what dose of 4-NP is able to induce not only vitellogenin but also the feminization process in males; in addition, the trend of bioaccumulation and bioavailability of 4-NP were also assessed to link the laboratory experiments with the animal situation in the wild. In fact, as described by Pascolini et al. (2001), the sharp decrease of the amphibian population in that region was noticed because of parasitological diseases,


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perhaps depending on the relationships between endocrine disorders and the immune system. Moreover, 4-NP caused changes in peripheral sex steroids and gonadotropins both in males and females. At pituitary levels, both a remarkable increase in the number of prolactin immunolabeled cells in newt and aromatization in the brain were revealed, as the ability of the brain to aromatize and otherwise transform androgen substrates is phylogenetically ancient (Mosconi et al., 2002a; Polzonetti-Magni et al., 2004). The eVects of estrogenic compounds on aromatase activity may be responsible for the decrease in androgen levels concomitant with high peripheral estradiol-17b levels found not only in adults but also in tadpoles (Fagotti et al., 2004). Aromatase, in fact, is a good biomarker of the exposure to estrogenic compounds because some of them do not interact with estrogen receptors but, through the increasing of aromatase activity, are able to produce a feminization process evidenced by the vitellogenin levels in the males (Polzonetti-Magni et al., 2004).

B. The Effect of the Feminization Process in Wild Populations Fish and amphibians inhabiting polluted water are exposed to complex mixtures of compounds that may have negative consequences on their fitness and reproductive success, with very dramatic changes in animal biodiversity. Some of these compounds are known to exert an estrogenic activity on humans and wildlife, whereas others display antiestrogenic properties (Kirk et al., 2002). Studies on carp have attempted to investigate the combined eVects of estrogenxenobiotic exposure in two biomarkers, VTG and EROD. The latter is cytocrome P450 dependent and is located in the liver endoplasmic reticulum, playing a role in oxidative metabolism of endogenous compounds (steroids), as well as a wide range of xenobiotics (Sole´ et al., 2002). The eVects of exposure to environmental estrogen that can impair the reproductive process in exposed organisms were evaluated in wild carp, Cyprinus carpio L. by Carnevali et al. (2002), who monitored, through a validated ELISA, changes of plasma vitellogenin and estradiol-17b during prespawning, spawning, and postspawning periods, together with those changes of estrogen receptor density in the liver. When a dimer of estrogen receptor complex binds to estrogen-responsive elements in the DNA, the transcription of vitellogenin mRNA (Beato et al., 1995; Perlman et al., 1984; Wahli et al., 1979) and other genes, such as the estrogen receptor (Tata, 1987) and the retinal binding protein (McKearin and Shapiro, 1988; McKearin et al., 1987), are activated. Therefore, the carp can be considered to be a useful sentinel species for biomonitoring studies on environmental estrogens and on the eVects on its reproductive biology. In females, VTG showed high

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seasonality in parallel with plasma changes of estradiol-17b and gonadosomatic index. In addition, in 40% of males VTG was found to be present in the plasma, whereas changes in the liver estrogen receptor were observed in both males and females not parallel with those of vitellogenin and estradiol, even though it is worthy of note that estradiol up-regulates its own receptor. The disturbance of xenoestrogens by competing for estrogen receptor was also reported by Latonnelle et al. (2002) in rainbow trout and Siberian sturgeon, Acipenser baeri, when competitive-binding studies between phytoestrogens and estradiol were performed and after having previously demonstrated the estrogenic eVects of phytoestrogens coming from dietary ingredients of plant origin (Bennetau-Pelissero et al., 1998; Pellissero et al., 1991). Moreover, the evolutionary implications of xenoestrogenic chemicals with reproductive strategies in fish have recently been reported in a review by Arukwe and Goksoyr (2003). In amphibians, the final evidence of a physiological significance of these compounds is consistent with their eVect on sexual diVerentiation, which can be demonstrated by determination of female and male phenotypes of individuals treated with estrogenic compounds during the sensitive phase of larval development (Kloas, 2002) or of those sampled in the wild (Hayes et al., 2002). In the Hayes study, it was found that exposure to waterborne atrazine contamination produced, in male wild leopard frogs, gonadal abnormalities such as retardant development and hermaphroditism. Therefore, the eVect of such compounds must be considered from the standpoint of the reproductive process, which includes also reproductive behavior. Interesting results have been obtained by Cardinali et al. (2004), who found that exposure to estrogenic compounds counteracts reproductive behavior in Poecilia reticulata. Although a widespread incidence of intersex has been reported in estuarine and freshwater fish, little evidence is available on the direct eVect of EDs on fertility. Recently, in Rutilius rutilus, Jobling et al. (2002) evidenced the reduced reproductive capability of intersex fish. It seems that even if VTG is certainly the key biomarker of the feminization process, more eVort must not only be addressed to identifying other good and suitable biomarkers but also to giving more attention to wild population studies in which the knowledge of endocrinology can be integrated with ecotoxicological aspects.

VII. Concluding Remarks According to Darwin, ‘‘it is generally acknowledged that all organic beings have been formed on two great laws: Unity of Type and Conditions of Existence.’’ This sentence also accounts for vitellogenesis, which in general is consistent with VTG synthesis in the liver on multihormonal control, its


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‘‘swimming’’ in the circulation, and VTG uptake by growing oocytes through receptor-mediated endocytosis. Instead, its diVerent patterns are mainly dependent on the adaptive mechanisms allowing reproductive success. The considerations developed in the previous sections indicate that vitellogenin, the key molecule of vitellogenesis, might be the product of a phylogenetic and evolutionary process from invertebrate to vertebrate organisms. Vitellogenin functions as a nutritional source for the developing embryo, rather than as an important functional protein, and its primary sequence has not been conserved among species. Thus, except for short patches of homology along the entire molecule, vitellogenins from diVerent species share little structural similarity. For this reason, an antibody made against VTG from one species is limited in its application as a probe for another, and therefore any studies on vitellogenin need accurate and validated immunological methodologies. Vitellogenin is produced in animals to a large extent; in fact VTG mRNA, synthesized as much as 40,000-fold over background levels (Pakdel et al., 1991), gives rise to as much as 100 mg/mL protein in some species. The patterns of vitellogenesis in reproductive strategies have been studied in terms of measurement of blood vitellogenin related with gonadal and nongonadal hormones; among them, for many years, estradiol-17b has been considered the most important regulator. However, it is precisely from these studies that convincing evidence has arisen on the multihormonal control of liver vitellogenin synthesis corroborated by in vitro experiments conducted in several animal models, representative of both aquatic and terrestrial species. Moreover, in the fish, amphibian, and reptile models here described, the intriguing interplay between pituitary and gonadal steroids related with seasonality can be responsible for modulating vitellogenesis. Unfortunately, less knowledge is available at this time on the hormonal mechanisms regulating vitellogenin uptake by growing oocytes and on those that modulate, perhaps in a paracrine fashion, vitellogenin termination at the end of the seasonal reproductive process. The concepts of how vitellogenin is incorporated, as a yolk, and how and when the yolk is used by the developing embryo have also received due attention. The relationship between vitellogenin and estradiol again plays a crucial role, as vitellogenin assay in male organisms is widely considered a useful biomarker of exposure to environmental chemicals, able to mimic the eVects of endogenous estradiol and, in turn, to determine the feminization process in wildlife and human. Since the paper by Sumpter and Jobling (1995), the research in that field has undergone a marked increase in many laboratories, with diVerent approaches ranging from the hormonal mechanisms of action to ecotoxicological aspects. The great interest now being displayed for VTG, we hope, will throw more light on this fascinating aspect of reproduction. It is

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also clear that nonmammalian (and in particular, lower vertebrate) models appear to be fundamental for vitellogenesis research addressed to clarifying the yolkless egg and the evolution of eutherian viviparity.

Acknowledgments We thank James Burge for his help in revising the English text and this chapter. Financial support was provided by the Italian Ministries of the University (MURST, Cofin, PolzonettiMagni) and of Fisheries and Agriculture.

References Amant, D. R., Carson, D. D., Decker, G. L., Welply, J. K., and Lennarz, W. J. (1986). Characterization of yolk platelets isolated from developing embryos of Arbacia punctulata. Dev. Biol. 113, 342–355. Anderson, M., Olsen, H., Matsumura, F., and Hinton, D. E. (1996). In vivo modulation of 17 b–estradiol-induced vitellogenin synthesis and estrogen receptor in rainbow trout (Oncorhynchus mykiss) liver cells by b-naphthoflavone. Toxicol. Appl. Pharmacol. 98, 422–433. Angelini, F., Ciarcia, G., Picariello, O., Botte, V., and Pagano, M. (1986). Sex steroids and postreproductive refractoriness in the lizard, Podarcis s. sicula. Boll. Zool. 53, 59–62. Ankley, G., Johnson, R., Toth, L., and Detenbeck, N. (1997). Development of a research strategy for assessing the ecological risk of endocrine disruptors. Rev. Toxixol 1, 71–106. Ankley, G., Mihaich, E., Stahl, R., Tillitt, T., Colborn, T., McMaster, S., Miller, R., Bantle, J., Dickwerson, R., Fry, M., Geisy, J., Gray, L. E., Guiney, P., Hutchinson, T., Kramer, V., LeBlanc, G., Mayes, M., Nimrod, A., Peterson, R., Purdy, R., Ringer, R., Thomas, P., Van der Kraak, G., and Zacharewski, T. (1998). Overview of a workshop on screening and testing methods for detecting potential (anti)estrogenic/androgenic chemicals in wildlife. Environ. Toxixol. Chem. 17, 68–87. Ansari, A. Q., Dolphin, P. J., Lazier, C. B., Munday, K. A., and Akhtar, M. (1971). Chemical composition of an oestrogen-induced calcium-binding glycolipophosphoprotein in Xenopus laevis. Biochem J. 122, 107–113. Arnold, S. F., and McLachlan, J. A. (1996). Synergistic signals in the environment. Environ. Health Perspect. 104, 1020–1023. Arukwe, A., and Goksoyr, A. (2003). Eggshell and egg yolk proteins in fish: Hepatic proteins for the next generation: Oogenetic, population, and evolutionary implications of endocrine disruption. Comparative Hepatology 2, 1–21. Assisi, L., Autuori, F., Botte, V., Farrace, M. G., and Piacentini, M. (1999). Hormonal control of ‘‘tissue’’ transglutaminase induction during programmed cell death in frog liver Experimental Cell Research. 247, 339–346. Barnabe´, G., and Paris, J. (1984). Ponte avance´ et ponte normale du loup Dicentrarchus labrax L. a` la station de biologie marine et lagunaire de Se`te. In ‘‘L’Aquaculture du bar et des sparide`s’’ (G. Barnabe´ and R. Billard, Eds.), pp. 63–72. INRA, Paris. Beato, M., Herrlich, P., and Schu¨tz, G. (1995). Steroid hormone receptors: Many actors in searching for a plot. Cell 83, 851–857. Bennetau-Pelissero, C., Flouriot, G., Valotaire, Y., Sumpter, J. P., and Le Menn, F. (1998). Induction of rainbow trout estradiol receptor (rtER)mRNA and vitellogenin (rtVTG) mRNA by phytoestrogens in hepatocyte cultures. Ann. NY Acad. Sci. 839, 600–601.


35

Beuving, G., and Gruber, M. (1971). Induction of phosvitin synthesis in roosters by estradiol injection. Biochim Biophys Acta. 232, 529–536. Blackburn, D. G. (1995). Saltationist and punctuated equilibrium models for the evolution of viviparity and placentation. J. Theor. Biol. 174, 199–216. Brock, M. L., and Shapiro, D. J. (1983). Estrogen stabilizes vitellogenin mRNA against cytoplasmic degradation. Cell 34, 207–214. Brooks, S., Tyler, C. R., Carnevali, O., Coward, K., and Sumpter, J. (1997). Molecular characterization of ovarian cathepsin D in the rainbow trout, Oncorhynchus mykiss. Gene 201, 45–54. Brown, M. A., Carne, A., and Chambers, G. K. (1997). Purification, partial characterization and peptide sequence of vitellogenin from a reptile, the tuatara (Sphenodon punctatus). Comp. Biochem. Physiol. B 117, 159–168. Byrne, B. M., Gruber, M., and Ab, G. (1989). The evolution of egg yolk proteins. Prog. Biophys. Mol. Biol. 53, 33–69. Callard, I. P., Bayne, C. G., and McConnel, W. F. (1972). Hormones and reproduction in the female lizard, Sceloporus cyanogenys. Gen. Comp. Endocrinol. 18, 175–194. Callard, I. P., Giannoukos, G., Charnock-Jones, D. S., Benson, S., and Paolucci, M. (1994). Hormone regulation of vitellogenin genes and the evolution of viviparity. In ‘‘Perspectives in Comparative Endocrinology’’ pp. 325–332. National Research Council of Canada, Canada. Cardinali, M., Maradonna, F., Olivotto, I., Bortoluzzi, G., Polzonetti-Magni, A. M., Mosconi, G., and Carnevali, O. (2004). Temporary impairment of reproduction in freshwater teleost exposed to Nonylphenol. Reproductive Toxicol. 187, 597–604. Carey, C., and Bryant, C. J. (1995). Possible interrelations among environmental toxicants, amphibian development, and declining amphibian populations. Environ. Health Perspect. 103, 13–17. Carnevali, O., and Belvedere, P. (1991). Comparative studies of fish, amphibian, and reptilian vitellogenins. J. Exp. Zool. 259, 18–25. Carnevali, O., and Maradonna, F. (2003). Exposure to xenobiotic compounds: Looking for new biomarkers. Gen. Comp. Endocrinol. 131, 203–209. Carnevali, O., and Mosconi, G. (1991). Fat body involvement in the plasma vitellogenin changes during recovery phase in Rana esculenta. Zool. Sci. 8, 879–884. Carnevali, O., and Mosconi, G. (1992). In vitro induction of vitellogenin synthesis in Rana esculenta: Role of the pituitary. Gen. Comp. Endocrinol. 86, 352–358. Carnevali, O., Belvedere, P., Angelini, F., Colombo, L., and Polzonetti-Magni, A. (1991a). Immunological aspects of fish, amphibian and reptilian vitellogenins. In ‘‘Symposium on the Evolution of Terrestrial Vertebrates’’ (G. Ghiara, F. Angelini, E. Olmo, and L. Varano, Eds.), pp. 439–445. Mucchi, Modena, Italy. Carnevali, O., Carletta, R., Cambi, A., Vita, A., and Bromage, N. (1999a). Yolk formation and degradation during oocyte maturation in seabream Sparus aurata: Involvement of two lysosomal proteinases. Biol. Reprod. 60, 140–146. Carnevali, O., Centonze, F., Brooks, S., Marota, I., and Sumpter, J. P. (1999b). Molecular cloning and expression of ovarian cathepsin D in seabream Sparus aurata. Biol. Reprod. 66, 785–791. Carnevali, O., Mosconi, G., Angelini, F., Limatola, E., Ciarcia, G., and Polzonetti-Magni, A. M. (1991b). Plasma vitellogenin and 17 b-estradiol levels during the annual reproductive cycle of Podarcis sicula Raf. Gen. Comp. Endocrinol. 84, 337–343. Carnevali, O., Mosconi, G., Cambi, A., Ridolfi, S., Zanuy, S., and Polzonetti-Magni, A. M. (2001a). Changes of lysosomal enzyme activities in sea bass Dicentrachus labrax egg and developing embryo. Aquaculture 202, 249–256. Carnevali, O., Mosconi, G., Cardinali, M., Meiri, I., and Polzonetti-Magni, A. M. (2001b). Molecular components related with egg viability in the sea bream Sparus aurata. Mol. Rep. Dev. 58, 330–335.

36


Carnevali, O., Mosconi, G., Habibi, H. R., Elia, A. C., Cardinali, M., and Polzonetti-Magni, A. M. (2002). Validation of an enzyme linked immunosorbent assay (ELISA) for Ciprinus carpio L. vitellogenin, as a biomarker of reproductive disorders. Chem. Ecol. 19, 5–13. Carnevali, O., Mosconi, G., and Polzonetti-Magni, A. M. (2000). Involvement of tyrosine kinase and cAMP in growth hormone-induced vitellogenin synthesis in the anuran, Rana esculenta. Life Sci. 67, 1467–1476. Carnevali, O., Mosconi, G., Roncarati, A., Belvedere, P., Limatola, E., and Polzonetti-Magni, A. M. (1993a). Yolk protein changes during oocyte growth in European sea bass Dicentrarchus labrax L. J. Appl. Icthyol. 9, 175–184. Carnevali, O., Mosconi, G., Roncarati, A., Belvedere, P., Romano, M., and Limatola, E. (1992a). Changes in the electrophoretic pattern of yolk proteins during vitellogenesis in the gilthead sea bream Sparus aurata L. Comp. Biochem. Physiol. Vol. 103B, n. 4, 955–962. Carnevali, O., Mosconi, G., Yamamoto, K., Kobayashi, T., Kikuyama, S., and PolzonettiMagni, A. M. (1992b). Hormonal control of ‘‘in vitro’’ vitellogenin synthesis in Rana esculenta liver: EVects of mammalian and amphibian growth hormone. Gen. Comp. Endocrinol. 88, 406–414. Carnevali, O., Mosconi, G., Yamamoto, K., Kobayashi, T., Kikuyama, S., and PolzonettiMagni, A. M. (1993b). ‘‘In vitro’’ eVects of mammalian and amphibian prolactins on hepatic vitellogenin synthesis in Rana esculenta. J. Endocrinol. 137, 383–389. Carnevali, O., Sabbieti, M. G., Mosconi, G., and Polzonetti-Magni, A. M. (1995). Multihormonal control of vitellogenin mRNA expression in the liver of frog, Rana esculenta. Mol. Cell. Endocrinol. 114, 19–25. Casini, S., Fossi, M. C., Mori, G., and Bjornstad, A. (2002). Vitellogenin induction in Cyprinus carpio treated with 17b-estradiol and 4-nonylphenol. Environ. Monitoring Assessment 75, 235–239. Cecchettini, A., Falleni, A., Gremigni, V., Locci, M. T., Masetti, M., Bradley, J. T., and Giorgi, F. (2001). Yolk utilization in stick insects entails the release of vitellin polypeptides into the perivitelline fluid. Eur. J Cell Biol. 80, 458–465. Celius, T., and Walther, B. T. (1998). DiVerential sensitivity of zonagenesis and vitellogenesis in Atlantic salmon (Salmo salar L) to DDT pesticides. J. Exp. Zool. 281, 259–266. Carda`, J., Calman, B. G., LaFleur, G. J. Jr., and Limesand, S. (1996). Pattern of vitellogenesis and follicle maturational competence during the ovarian follicular cycle of Fundulus heteroclitus. Gen. Comp. Endocrinol. 103, 24–35. Chan, S. L., Tan, C. H., Pang, M. K., and Lam, T. J. (1991). Vitellogenin purification and development of an assay for the vitellogenin receptor in oocyte membranes of tilapia (Oreochromis niloticus. Linnaeus 1766). J. Exp. Zool. 257, 96–109. Colborn, T., Saal, F. S. V., and Soto, A. M. (1993). Developmental eVects of endocrinedisrupting chemicals in wildlife and humans. Environm. Health Perspect 101, 378–384. Connaughton, M. A., and Aida, K. A. M. (1999). Female reproductive system, fish. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 2, pp. 193–204. San Diego: Academic Press. Cooper, D. N., Gerber-Huber, S., Nardelli, D., Schubiger, J. L., and Wahli, W. (1987). The distribution of the ducleotide CpG and Cytosine Methylation in vitellogenin gene family. J. Mol. Evolution. 25, 107–115. Copeland, P. A., and Thomas, P. (1988). The measurement of plasma vitellogenin levels in a marine teleost, the spotted seatrout (Cynoscion nebulosus) by homologous radioimmunoassay. Comp. Biochem. Physiol. B 91, 17–23. Craik, J. C. A. (1978a). Plasma levels of vitellogenin in the elasmobranch Scyliorhinus canicula L. (Lesser spotted dogfish). Comp. Biochem. Physiol. B 60, 9–18. Craik, J. C. A. (1978b). Kinetic studies of vitellogenin metabolism in the elasmobranch Scyliorhinus canicula L. Comp. Biochem. Physiol. A 61, 355–361.


37

Craik, J. C. A. (1978c). An annual cycle of vitellogenesis in the elasmobranch Scyliorhinus canicula. J. Mar. Biol. Ass. UK 58, 719–726. Craik, J. C. A. (1978d). The eVects of oestrogen treatment on certain plasma constituents associated with vitellogenesis in the elasmobranch Scyliorhinus canicula. Gen. Comp. Endocrinol. 35, 455–464. Craik, J. C. A. (1979). Simultaneous measurement of rates of vitellogenin synthesis and plasma levels of oestradiol in an elasmobranch. Gen. Comp. Endocrinol. 38, 264–266. Crews, D. (1999). Reptilian reproduction, Overview. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 4, pp. 254–259. Academic Press, San Diego, CA. Custodia-Lora, N., and Callard, I. P. (2002). Seasonal changes in hepatic progesterone receptor mRNA, estrogen receptor mRNA, and vitellogenin mRNA in the painted turale, Chrysemys picta. Gen. Comp. Endocrinol. 128, 193–204. Cutting, J. A., and Roth, T. F. (1973). Staining of phosphoproteins on acrylamide gel electrophoregrams Anal. Biochem. 54, 386–394. Davis, D. L., Bradlow, H. L., WolV, M., WoodruV, T., Hoel, D. G., and Anton-Culver, H. (1993). Medical hypothesis: Xenoestrogens as preventable causes of breast cancer. Environ. Health Perspect. 101, 372–377. Denslow, N. D., Chow, M. C., Kroll, K. J., and Green, L. (1999). Vitellogenin as a biomarker of exposure for estrogen or estrogen mimics. Ecotoxicology 8, 385–398. Ding, J. L., Lim, E. H., and Lam, T. J. (1994). Cortisol-induced hepatic vitellogenin mRNA in Oreochromis aureus (Steindachner). Gen. Comp. Endocrinol. 96, 276–287. Duggan, A. E., and Callard, I. P. (2003). Lipids and lipid-transporting proteins in Chrysemys picta: Role of gonadal steroids and growth hormone in intact and hypophysectomized turtles. Gen. Comp. Endocrinol. 131, 176–184. Edinger, S. R., Mambo, E., and Evans, M. I. (1997). Estrogen-dependent transcriptional activation and vitellogenin gene memory. Mol. Endocrinol. 11, 1985–1993. Elkin, E. G., Freed, M. B., Denetz, S. A. H., and Bidwell, C. A. (1995). Proteolysis of Japanese quail and chicken plasma apolipoprotein B and vitellogenin by cathepsin D: Similarity of the resulting protein fragments with egg yolk polypeptides. Comp. Biochem. Physiol. 112, 191–196. Evans, M. I., Silva, R., and Burch, J. B. (1988). Isolation of chicken vitellogenin I and III cDNAs and the developmental regulation of five estrogen-responsive genes in the embryonic liver. Genes Dev. 2, 116–124. Fagotti, A., Morosi, L., Di Rosa, I., Clarioni, R., Simoncelli, F., Pascolini, R., Pellegrino, R., Guex, G. D., and Hotz, H. (2004). Bioaccumulation of organochlorine pesticides in frogs of the Rana esculenta complex in central Italy. Amphibia-Reptilia. In press. Fagotto, F. (1995). Regulation of yolk degradation, or how to make sleepy lysosomes. J. Cell Sci. 108, 3645–3647. Gavaud, J. (1986a). Vitellogenesis in the lizard Lacerta vivipara Jacquin. I. Purification and partial characterization of plasma vitellogenin. Gen. Comp. Endocrinol. 63, 1–10. Gavaud, J. (1986b). Vitellogenesis in the lizard Lacerta vivipara Jacquin. II. Vitellogenin synthesis during the reproductive cycle and its control by ovarian steroids. Gen. Comp. Endocrinol. 63, 11–23. Gerber-Huber, S., Nardelli, D., Haefliger, J-A., Cooper, D. N., Givel, F., Germond, J. E., Engel, J., Green, N. M., and Wahli, W. (1987). Precursor-product relationship between vitellogenin and the yolk proteins as derived from the complete sequence of a Xenopus vitellogenin gene. Nucleic Acids Res. 15, 4737–4760. Germond, J. E., ten Heggeler, B., Schubiger, J. L., Walker, P., Westley, B., and Wahli, W. (1983). Vitellogenin B2 gene in Xenopus laevis: Isolation, in vitro transcription and relation to other vitellogenin genes. Nucleic Acids Res. 11, 2979–2997.

38


Gillesby, B. E., and Zacharewski, T. R. (1998). Exoestrogens: Mechanisms of action and strategies for identification and assessment. Environ. Toxixol. Chem. 17, 3–14. Giorgi, F., Cecchettini, A., Falleni, A., Masetti, M., and Gremigni, V. (1998). Vitellogenin is glycosylated in the fat body of the stick insect Carausius morosus and not further modified upon transfer to the ovarian follicle. Micron. 29, 451–460. Giorgi, F., Gobbetti, A., and Polzonetti-Magni, A. (1982). Variations in the vitellogenin titre during the reproductive cycle of Rana esculenta L. Comp. Biochem. Physiol. B 72, 501–506. Giorgi, F., Lucchesi, P., Morelli, A., and Bownes, M. (1993). Ultrastructural analysis of Drosophila ovarian follicles diVering in yolk polypeptide (yps) composition. Development 117, 319–328. Giorgi, F., Yin, L., Cecchettini, A., and Nordin, J. H. (1997). The vitellin-processing protease of Batella germanica is derived from a pro-protease of maternal origin. Tissue Cell 29, 293–303. Gobbetti, A., Polzonetti-Magni, A., Gernai, M., Carnevali, O., and Botte, V. (1985a). Vitellogenin hormonal control in the green frog, Rana esculenta. Interplay between estradiol and pituitary hormones. Comp. Biochem. Physiol. A. 82, 855–885. Gobbetti, A., Polzonetti-Magni, A., and Zerani, M. (1985b). EVect of hypophysectomy on estrogen-induced vitellogenin synthesis in the green frog Rana esculenta complex. Boll. Zool. 52, 343–346. Grengard, O., Gordon, M., Smith, M. A., and Acs, G. (1964). Studies on the mechanism of diethylstilbestrol-induced formation of phosphoprotein in male chickens. J. Biol. Chem. 239, 2079–2082. Guillette, L. J. Jr. (1985). The evolution of egg retention in lizard: A physiological model. In ‘‘The Biology of Australian Frogs and Reptiles’’ (G. Grigg, R. Shine, and H. Ethmann, Eds.), pp. 379–386. Royal Zoological Society of New South Wales, Sydney. Guillette, L. J. Jr. (1987). The evolution of viviparity in fishes, amphibians, reptiles: An endocrine approach. In ‘‘Hormones and Reproduction in Fishes, Amphibians, Reptiles’’ (D. O. Norris and R. E. Jones, Eds.), pp. 523–562. Plenum Press, New York. Harries, J. E., Sheahan, D. A., Jobling, S., Matthiessen, P., Neall, P., Sumpter, J. P., Tylor, T., and Zaman, N. (1997). Estrogenic activity in five United Kingdom rivers detected by measurement of vitellogenesis in caged male trout. Environ. Tox. Chem. 16, 534–542. Hayes, T., Haston, K., Tsui, M., Hoang, A., HaeVele, C., and Vonk, A. (2002). Feminization of male frogs in the wild. Nature 419, 895–896. Heppell, S. A., and Sullivan, C. V. (1999). Gag (Mycteroperca microlepis) vitellogenin: Purification, characterization and use for enzyme-linked immunosorbent assay (ELISA) of female maturity in three species of grouper. Fish Physiol. Biochem. 20, 361–374. Heppell, S. A., Denslow, N. D., Folmar, L. C., and Sullivan, C. V. (1995). Universal assay of vitellogenin as a biomarker for environmental estrogens. Environ Health Perspect. 103, 9–15. Ho, S. M. (1991). Vitellogenesis. In ‘‘Vertebrate Endocrinology: Fundamentals and Biochemical Implications’’ (P. K. T. Pang and M. P. Schreibman, Eds.), Vol. 4, pp. 91–126. Academic Press, New York. Ho, S. M., Danko, D., and Callard, I. P. (1982a). EVect of hypophysectomy and growth hormone on estrogen induced vitellogenesis in the freshwater turtle, Chrysemys picta. Gen. Comp. Endocrinol. 48, 254–260. Ho, S. M., Kleis, S., McPherson, R., Heisermann, C. S., and Callard, I. P. (1982b). Regulation of vitellogenesis in reptiles. Herpetology 38, 40–50. Ho, S. M., Wangh, L. J., and Callard, I. P. (1985). Sexual diVerences in the in vitro induction of vitellogenesis in the turtle: Role of pituitary and growth hormone. Comp. Biochem. Physiol. 81, 467–472. Hori, S. H., Kodama, T., and Tanahashi, K. (1979). Induction of vitellogenin synthesis in goldfish by massive dose of androgens. Gen. Comp. Endocrinol. 37, 306–320.


39

Janeiro-Cinquini, T. R. F., Ribolla, P. E. M., Capurro, M. L., and Winter, C. E. (1999). Vitellogenin and yolk protein processing in Bothrops Jararaca (Wied), a tropical venomous snake. Comp. Biochem. Physiol. B. 122, 189–198. Jobling, S., and Sumpter, J. P. (1993). Detergent components in sewage eZuent are weakly estrogenic to fish: An in vitro study using rainbow trout (Oncorhynchus mykiss) hepatocytes. Envir. Tox. Chem. 27, 361–372. Jobling, S., Coey, S., Whitmore, J. G., Kime, D. E., Van Look, K. J., McAllister, B. G., Beresford, N., Henshaw, A. C., Brighty, G., Tyler, C. R., and Sumpter, J. P. (2002). Wild intersex roach (Rutilus rutilus) have reduced fertility. Biol. Reprod. 67(2), 515–524. Jobling, S., Sheahan, D., Osborn, J. A., Matthiessen, P., and Sumpter, J. P. (1996). Inhibition of testicular growth in rainbow trout (Oncorhynchus mykiss) exposed to estrogenic alkylphenolic chemicals. Envir. Tox. Chem. 15, 194–202. Kavlock, R. J., Daston, G. P., De Rosa, C., Fenner, P., Gray, L. E., Kaattari, S., Lucier, G., Luster, M., Mac, M. J., Maczka, C., Miller, R., Moore, J., Rolland, R., Scott, G., Sheehan, D. M., Sinks, T., and Tilson, H. A. (1996). Research needs for the risk assessment of health and environmental eVects of endocrine disruptors: A report of the US EPA sponsored workshop. Environ. Health Perspect. 104, 715–740. Kiesecker, J. M., Blaustein, A. R., and Belden, L. K. (2001). Complex causes of amphibian population declines. Nature 410, 681–684. Kim, B. H., and Takemura, A. (2002). In vitro vitellogenin synthesis in primary cultures of tilapia hepatocytes. Fisheries Sci. 68, 123–131. Kime, D. E., Nash, J. P., and Scott, A. P. (1999). Vitellogenesis as a biomarker of reproductive disruption by xenobiotics. Aquaculture 177, 345–352. Kirk, L. A., Tyler, C. R., Lye, C. M., and Sumpter, J. P. (2002). Changes in estrogenic and androgenic activities at diVerent stages of treatment in wastewater treatment works. Environ Toxicol. Chem. 21, 972–979. Klass, M. R., Wolf, N., and Hirsh, D. (1979). Further characterization of a temperaturesensitive transformation mutant in Caerorhbditis elegans. Dev. Biol. 69, 329–335. Kloas, W. (2002). Amphibians as a model for the study of endocrine disruptors. Int. Rev. Cytol. 216, 1–57. Know, H. C., and Mugiya, Y. (1994). Involvement of growth hormone and prolactin in the induction of vitellogenin synthesis in primary hepatocyte culture in the eel, Anguilla japonica. Gen. Comp. Endocrinol. 93, 51–60. Kobayashi, T., Kikuyama, S., Yasuda, A., Kawauchi, H., Yamaguchi, K., and Yokoo, Y. (1989). Purification and characterization of bullfrog growth hormone. Gen. Comp. Endocrinol. 73, 417–424. Korsgaard, B., and Pedersen, K. L. (1998). Vitellogenin in Zoarces viviparous: Purification, quantification by ELISA and induction by estradiol-17b and 4-nonylphenol. Comp. Biochem. Physiol. C. 120, 159–166. Kwon, J. Y., Prat, F., Randall, D. J., and Tyler, C. R. (2001). Molecular characterization of putative yolk processing enzymes and their expression during oogenesis and embryogenesis in rainbow trout (Oncorhynchus mykiss). Biol. Rep. 65, 1701–1709. LaFleur, G. J. (1999). Vitellogenins and vitellogenesis. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 4, pp. 985–994. Academic Press, San Diego, CA. LaFleur, G. J. Jr., Byrne, M. B., Kanungo, J., Nelson, L. A., Greenberg, R. M., and Wallace, R. A. (1995). Fundulus heteroclitus vitellogenin: The deduced primary structure of a piscine precursor to noncrystalline, liquid-phase yolk protein. J. Mol. Evol. 41, 505–521. Lance, V. A. (1999a). Female reproductive system, reptiles. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 2, pp. 239–243. Academic Press, San Diego, CA. Lance, V. A. (1999b). Reptilian reproductive cycles. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 4, pp. 260–265. Academic Press, San Diego, CA.

40


Larkin, P., Knoebl, I., and Denslow, N. D. (2003). DiVerential gene expression analysis in fish exposed to endocrine disrupting compounds. Comp. Biochem. Physiol. B 136, 149–161. Latonnelle, K., Fostier, A., Le Menn, F., and Bennetau-Pelisero, C. (2002). Binding aYnities of hepatic nuclear estrogen receptor for phytoestrogens in rainbow trout (Onchorhynchus mykiss) and Siberian sturgeon (Acipenser baeri). Gen. Comp. Endocrinol. 129, 69–79. Lazier, C. B., Langley, S., Ramsey, N. B., and Write, J. M. (1996). Androgen inhibition of vitellogenin gene expression in tilapia (Oreochromis niloticus). Gen. Comp. Endocrinol. 104, 321–329. Le Menn, F., Rochefort, H., and Garcia, M. (1980). EVects of androgen mediated by the estrogen receptor of fish liver: Vitellogenin accumulation. Steroids 35, 315–328. Lessman, C. A. (1999). Oogenesis in nonmammalian vertebrates. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 3, pp. 498–508. Academic Press, San Diego, CA. Lindholst, C., Pedersen, K. L., and Pedersen, S. N. (2000). Estrogenic response of bisphenol A in rainbow trout (Oncorhynchus mykiss). Aquatic Toxicology 48, 87–94. Lui, C. W., and O’Connor, J. D. (1977). Biosynthesis of lipovitellin III. The incorporation of labeled amino acids into the purified lipovitellin of the crab, Pachygrapsus crassipes. J. Exp. Zool. 199, 105–108. Madsen, L. L., Korsgaard, B., and Bjerregaard, P. (2002). 4-tert-octylphenol and 17b-estradiol applied by feeding to flounder Platichthys flesus: Induction of vitellogenin and accumulation in tissues. Marine Environmental Research 54, 729–733. Mallay, S. K., Partin, J. S., Valdizan, M. C., and Lennarz, W. J. (1992). Proteolysis of the major yolk glycoproteins is regulated by acidification of the yolk platelets in sea urchin embryos. J. Cell Biol. 117, 1211–1221. Mancuso, A., Mosconi, G., Colucci, D., Polzonetti, A. M., and Vellano, C. (1996). Hormonal involvement in the production of vitellogenin by Triturus carnifex Laur. Atti Accad. Sci. Torino 130, 123–134. Martinez, E., Givel, F., and Wahli, W. (1987). The estrogen-responsive element as an inducible enhancer: DNA sequence requirements and conversion to a glucocorticoid-responsive element. Embo J. 6, 3719–3727. Matsubara, T., and Koya, Y. (1997). Course of proteolytic cleavage in three classes of yolk proteins during oocyte maturation in barfin flounder Verasper moseri, a marine teleost spawning pelagic eggs. J Exp. Zool. 278, 189–200. Matsubara, T., and Sawano, K. (1995). Proteolytic cleavage of vitellogenin and yolk proteins during vitellogin uptake and oocyte maturation in barfin flounder (Verasper moseri). J. Exp. Zool. 272, 34–45. Matsubara, T., Ohkubo, N., Andoh, T., Sullivan, C. V., and Hara, A. (1999). Two forms of vitellogenin, yielding two distinct lipovitellins, play diVerent roles during oocyte maturation and early development of barfin flounder, Verasper moseri, a marine teleost that spawns pelagic eggs. Dev Biol. 213, 18–32. McKearin, D. M., and Shapiro, D. J. (1988). Persistent estrogen induction of hepatic Xenopus laevis serum retinol binding protein mRNA. J. Biol. Chemistry 263, 3261–3265. McKearin, D. M., Barton, M. C., Keller, M. J., and Shapiro, D. J. (1987). Estrogens induce transcription of the Xenopus laevis serum retinol binding protein gene. J. Biol. Chem. 11, 4939–4942. Mendes, J. J. (2002). The endocrine disrupters: A major medical challenge. Food Chem. Tox. 40, 781–788. Mewes, K. R., Latz, M., Golla, H., and Fisher, A. (2002). Vitellogenin from female and estradiol-stimulated male river lampreys (Lampetra fluviatilis L.). J. Exp. Zool. 292, 52–72.


41

Miodini, P., Fioravanti, L., Di Fronzo, G., and Cappelletti, V. (1999). The two phytoestrogens genistein and quercetin exert diVerent eVects on estrogen receptor function. Br. J. Cancer 80, 1150–1155. Mommsen, T. P., and Walsh, P. J. (1998). Vitellogenin and oocyte assembly. In ‘‘Fish Physiology’’ (W. S. Hoar, D. J. Randal, and E. M. Donaldson, Eds.), pp. 347–406. Academic Press, New York. Mori, T., Matsumoto, H., and Yokota, H. (1998). Androgen-induced vitellogenin gene expression in primary cultures of rainbow trout hepatocytes. J. Steroid Biochem. Mol. Biol. 67, 133–141. Mosconi, G., Carnevali, O., Carletta, R., Nabissi, M., and Polzonetti-Magni, A. M. (1998). Gilthead seabream (Sparus aurata) vitellogenin: Purification, partial characterization, and validation of an enzyme-linked immunosorbent assay (ELISA). Gen, Comp, Endocrinol. 110, 252–261. Mosconi, G., Carnevali, O., Franzoni, M. F., Cottone, E., Lutz, I., Kloas, W., Yamamoto, K., Kikuyama, S., and Polzonetti-Magni, A. M. (2002a). Environmental estrogens and reproductive biology in amphibians. Gen. Comp. Endocrinol. 126, 125–129. Mosconi, G., Carnevali, O., Habibi, H. R., Sanyal, R., and Polzonetti-Magni, A. M. (2002b). Hormonal mechanism regulating hepatic vitellogenin synthesis in the gilthead sea bream, Sparus aurata. Am. J. Physiol.-Cell Physiol. 283, C673–C678. Mosconi, G., Carnevali, O., and Polzonetti-Magni, A. M. (1991). Ovarian development and sex steroid hormones during the reproductive cycle of Podarcis s. sicula Raf. Gynecol Endocrinol. 5, 7–13. Mosconi, G., Carnevali, O., Rorncarati, A., Sabbieti, M. G., Carrera, E., and PolzonettiMagni, A. M. (1992). Modificazioni morfologiche dei globuli di tuorlo durante la maturazione degli ovociti di branzino (Dicentrarchus labrax) e orata (Sparus aurata). Rivista Italiana di Acquacoltura 27, 71–79. Mosconi, G., Yamamoto, K., Carnevali, O., Nabissi, M., Polzonetti-Magni, A., and Kikuyama, S. (1994a). Seasonal changes in plasma growth hormone and prolactin concentrations of the frog Rana esculenta. Gen. Comp. Endocrinol. 93, 380–387. Mosconi, G., Yamamoto, K., Kikuyama, S., Carnevali, O., Mancuso, A., and Vellano, C. (1994b). Seasonal changes of plasma prolactin in the reproduction of the crested newt (Triturus carnifex Laur.). Gen. Comp. Endocrinol. 95, 342–349. Nagahama, Y. (1994). Endocrine regulation of gametogenesis in fish. Int. J. Dev. Biol. 38, 217–229. Nardelli, D., van het Schip, F. D., Gerber-Huber, S., Haefliger, J. A., Gruber, M., Ab, G., and Wahli, W. (1987). Comparison of the organization and fine structure of a chicken and a Xenopus laevis vitellogenin gene. J. Biol. Chem. 262, 15377–15385. Nash, J. P. (1999). Seasonal reproduction, fish. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 4, pp. 329–340. Academic Press, San Diego, CA. Nath, P., and Maitra, S. (2001). Role of two plasma vitellogenins from Indian major carp (Cirrhinus mrigala) in catfish (Clarias batracus) vitellogenesis. Gen. Comp. Endocrinol. 124, 30–44. Needham, J. (1931). ‘‘Chemical Embryology’’ New York: Hafner Publishing, 1963 (reprint of original 1931 edition). Noaksson, E., Linderoth, M., Bosveld, A. T. C., and Balk, L. (2003). Altered steroid metabolism in several teleost species exposed to endocrine disrupting substances in refuse dump leachate. Gen. Comp. Endocrinol. 134, 273–284. Ohkubo, N., Mochida, K., Adachi, S., Hara, A., Hotta, K., Nakamura, Y., and Matsubara, T. (2003). Development of enzyme-linked immunosorbent assay for two forms of vitellogenin in Japanese common goby (Acanthogobius flavimanus). Gen. Comp. Endocrinol. 131, 353–364.

42


Opresko, L. K., and Karpf, R. A. (1987). Specific proteolysis regulates fusion between endocytic compartments in Xenopus oocytes. Cell 51, 557–568. Opresko, L. K., and Wiley, H. S. (1987). Receptor-mediated endocytosis in Xenopus oocytes. II. Evidence for two novel mechanisms of hormonal regulation. J. Biol. Chem. 262, 4116–4123. Pakdel, F., Feon, S., Le Gac, F., Le Menn, F., and Valotaire, Y. (1991). In vivo estrogen induction of hepatic estrogen receptor mRNA and correlation with vitellogenin mRNA in rainbow trout. Mol. Cell Endocrinol. 75, 205–212. Palmer, B. D., Huth, L. K., Pieto, D. L., and Selcer, K. W. (1998). Vitellogenin as a biomarker for xenobiotic estrogens in an amphibian model system. Environmental Toxicology and Chemistry 17, 30–36. Pan, M. L., Bell, W. J., and Telfer, W. H. (1969). Vitellogenin blood protein synthesis by insect fat body. Science 165, 393–394. Paolucci, M., Custodia, N., and Callard, I. P. (1998). Progesterone: EVects and receptors subavian species. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 4, pp. 16–23. Academic Press, San Diego, CA. Parks, L. G., Cheek, A. O., Denslow, N. D., Heppell, S. A., McLachlan, J. A., LeBlanc, G. A., and Sullivan, C. V. (1999). Fathead minnow (Pimephales promelas) vitellogenin: Purification, characterization and quantitative immunoassay for the detection of estrogenic compounds. Comp. Biochem. Physiol. C 123, 113–125. Pascolini, R., Daszak, P., Cunningham, A., Fagotti, A., Tei, S., Vagnetti, D., Bucci, S., and Di Rosa, I. (2001). Parasitism by ‘‘Dermocystidium’’ ranae in a population of Rana esculenta complex in Central Italy, with a taxonomic review and designation of a new genus Amphibiocystidiumn gen. Dis. Aquat. Org. 56, 65–74. Pellissero, C., Le Menn, F., and Kaushik, S. (1991). Estrogenic eVect of dietary soya bean meal on vitellogenesis in cultured Siberian sturgeon Acipenser baeri. Gen. Comp. Endocrinol. 83, 447–457. Perez, L. E., and Callard, I. P. (1992). Identification of vitellogenin in the little skate (Raja erinacea). Comp. Biochem. Physiol. B 103, 699–705. Perlman, A. J., WolVe, A. P., Chapman, J., and Tata, J. R. (1984). Regulation by estrogen receptor of vitellogenin gene transcription in Xenopus hepatocyte culture. Mol. Cell. Endocrinol. 38, 151–161. Peyon, P., Baloche, S., and Burzawa-Ge´rard, E. (1996). Potentiating eVect of growth hormone on vitellogenin synthesis induced by 17b-estradiol in primary culture of female silver eel (Anguilla anguilla L.) hepatocytes. Gen. Comp. Endocrinol. 102, 263–273. Peyon, P., Baloche, S., and Burzawa-Ge´rard, E. (1997). Investigation into the possible role of androgens in the induction of hepatic vitellogenesis in the European eel: In vivo and in vitro studies. Fish Physiol. Biochem. 16, 107–118. Pickford, D. B., and Morris, I. D. (2003). Inhibition of gonadotropin-induced oviposition and ovarian steroidogenesis in the African clawed frog (Xenopus laevis) by the pesticide methoxychlor. Aquatic Toxicology 62, 179–194. Pierantoni, R., Cobellis, G., Meccariello, R., and Fasano, S. (2002). Evolutionary aspects of cellular communication in the vertebrate hypothalamo-hypophysio-gonadal axis. Int. Rev. Cytol. 218, 69–141. Polzonetti-Magni, A. M. (1999). Amphibian ovarian cycles. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 1, pp. 154–160. Academic Press, San Diego, CA. Polzonetti-Magni, A., Carnevali, O., Yamamoto, K., and Kikuyama, S. (1995). Growth hormone and prolactin in amphibian reproduction. Zool. Sci. 12, 683–694.


43

Polzonetti-Magni, A. M., Curini, R., Carnevali, O., Novara, C., Zerani, M., and Gobbetti, A. (1990). Ovarian development and sex steroid hormones during the reproductive cycle of Rana esculenta complex. Zool. Sci. 7, 265–271. Polzonetti-Magni, A. M., Mosconi, G., Carnevali, O., Yamamoto, K., Hanaoka, Y., and Kikuyama, S. (1998). Gonadotropins and reproductive function in the anuran amphibian, Rana esculenta. Biol. Reprod. 58, 88–93. Polzonetti-Magni, A. M., Mosconi, G., Palermo, F., Cottone, E., Forno, S., and Franzoni, M. F. (2004). Amphibians as a model for studying xenoestrogen-dependent neuroendocrine disruption. Ital. J. Zool. In press. Prat, F., Coward, K., Sumpter, J. P., and Tyler, C. R. (1998). Molecular characterization and expression of two ovarian lipoprotein receptors in the rainbow trout, Oncorhynchus mykiss. Biol. Reprod. 58, 1146–1153. Prisco, M., Romano, M., Ricchiari, L., Limatola, E., and Andreuccetti, P. (2002). An ultrastructural study on the vitellogenesis in the spotted ray Torpedo marmorata. Gen. Comp. Endocrinol. 128, 171–179. Purdom, C., Hardiman, P., Bye, V., Eno, N., Tyler, C., and Sumpter, J. (1994). Estrogenic eVects of eZuents from sewage treatment works. Chem. Ecol. 8, 275–285. Rankouhi, R. T, van Holsteijn, I., Letcher, R., Giesy, J. P., and van den Berg, M. (2002). EVects of primary exposure to environmental and natural estrogens on vitellogenin production in carp (Cyprinus carpio) hepatocytes. Toxixological Sciences 67, 75–80. Redshaw, M. R., and Follett, B. K. (1971). The crystalline yolk-platelet proteins and their soluble plasma precursor in an amphibian, Xenopus laevis. Biochem. J. 124, 759–766. Redshaw, M. R., and Nicholls, T. J. (1971). Oestrogen biosynthesis by ovarian tissue of the South African clawed toad, Xenopus laevis Daudin. Gen. Comp. Endocrinol. 16, 85–96. Reinboth, R. (1999). Fish, modes of reproduction. In ‘‘Encyclopedia of Reproduction’’ (E. Knobil and J. D. Neill, Eds.), Vol. 2, pp. 365–372. Academic Press, San Diego, CA. Reis-Henriques, M. A., Ferreira, M., Silva, L., and Dias, A. (2000). Evidence for an involvement of vitellogenin in the steroidogenic activity of rainbow trout (Oncorhynchus mykiss) vitellogenic oocytes. Gen. Comp. Endocrinol. 117, 260–267. RiVeser, M., and Hock, B. (2002). Vitellogenin levels in mussel hemolympha suitable biomarker for the exposure to estrogens? Comp. Biochem. Physiol. C 132, 75–84. Rothchild, I. (2003). The yolkless egg and the evolution of Eutherian viviparity. Biol. Reprod. 68, 337–357. Roubal, W. T., Lomax, D. P., Maryean, L., and Lyndal, L. J. (1997). Purification and partial characterization of English sole (Pleuronectes vetulus) vitellogenin. Comp. Biochem. Physiol. B 118, 613–622. Ruddack, D., and Wallace, R. A. (1968). On the site of phosvitin synthesis in Xenopus laevis. Biochem. Biophys. Acta 155, 299–301. Schwaiger, J., Mallow, U., Ferling, H., Knoerr, S., Braunbeck, Th., Kalbfus, W., and Negele, R. D. (2002). How estrogenic is nonylphenol? A transgenerational study using rainbow trout (Oncorhynchus mykiss) as a test organism Aquatic Toxicol. 59, 177–189. Sehgal, N., and Goswami, S. V. (2001). Biochemical changes in the liver of the Indian freshwater murrel, Channa puncatus (Bloch) during estradiol-induced vitellogenin synthesis. Fish Physiology Biochem. 24, 149–155. Selman, K., and Wallace, R. A. (1989). Cellular aspects of oocyte growth in teleosts. Zool. Sci. 6, 211–231. Selman, K., Wallace, R. A., and Cerda`, J. (2001). Bafilomycin A1 inhibits proteolytic cleavage and hydration but not yolk crystal disassembly or meiosis during maturation of sea bass oocytes. J. Exp. Zool. 290, 265–278.

44


Shi, G., Shao, J., Jiang, G., Wang, Q., Lu, Y., Liu, J., and Liu, J. (2003). Membrane chromatographic method for the rapid purification of vitellogenin from fish plasma. J. Chromatogr. B. 785, 361–368. Sire, M. F., Babin, P. J., and Vernier, J. M. (1994). Involvement of the lysosomal system in yolk protein deposit and degradation during vitellogenesis and embryonic development in trout. J. Exp. Zool. 269, 69–83. So, Y. P., Idler, D. R., and Hwang, S. J. (1985). Plasma vitellogenin in landlocked Atlantic salmon (Salmo solar Ouananiche): Isolation, homologous radioimmunoassay and immunological cross-reactivity with vitellogenin from other teleosts. Comp. Biochem. Physiol. 81B, 63–71. Sole´, M., Barcelo`, D., and Porte, C. (2002). Seasonal variation of plasmatic and hepatic vitellogenin and EROD activity in carp, Cyprinus carpio, in relation to sewage treatment plants. Aqua. Toxicol. 60, 233–248. Sonnenschein, C., and Soto, A. M. (1998). An updated review of environmental estrogen and androgen mimics and antagonists. J. Steroid Biochem. Mol. Biol. 65, 143–150. Soto, A. M., Justicia, H., Wray, J. W., and Sonnenschein, C. (1991). p-nonyl-phenol: An estrogenic xenobiotic released from ‘‘modified’’ polystyrene Environ. Health Perspect. 92, 167–173. Specker, J. L., and Sullivan, C. V. (1994). Vitellogenin in fishes: Status and perspectives. In ‘‘Perspectives in Comparative Endocrinology’’ (K. G. Davey, R. G. Peter, and S. S. Tobe, Eds.), pp. 304–315. National Research Council of Canada, Ottawa. Stifani, S., Barber, D. L., Nimpf, J., and Schneider, W. J. (1990a). A single chicken oocyte plasma membrane protein mediates uptake of very low density lipoprotein and vitellogenin. Proc. Natl. Acad. Sci. USA 87, 1955–1959. Stifani, S., Nimpf, J., and Schneider, W. J. (1990b). Vitellogenesis in Xenopus laevis and chicken: Cognate ligands and oocyte receptors. J. Biol. Chem. 15, 882–888. Sumpter, J. P., and Jobling, S. (1995). Vitellogenesis as a biomarker of estrogenic contamination of the aquatic environment. Environ. Health Prospect. 103, 173–178. Sun, X., and Zhang, S. (2001). Purification and characterization of a putative vitellogenin from the ovary of amphioxus (Branchiostoma belcheri tsingtaunese). Comp. Biochem. Physiol. B 129, 121–127. Takahashi, H., and Hanaoka, Y. (1981). Isolation and characterization of multiple components of basic gonadotropin from bullfrog (Rana catesbeiana) pituitary gland. J. Biochem. 90, 1333–1340. Tao, Y., Berlinsky, D. L., and Sullivan, C. V. (1996). Characterization of a vitellogenin receptor in white perch (Morone Americana). Biol. Rep. 55, 646–656. Tata, J. (1987). Regulation of Xenopus vitellogenin genes by estrogen. In ‘‘Recent Advances in Steroid Hormone Action,’’ pp. 259–283. De Gruyter, Berlin. Thorsen, A., Kjesbu, O. S., Fyhn, H. J., and Solemdal, P. (1996). Physiological mechanisms of buoyancy in eggs from brackish water cod. J. Fish Biol. 48, 457–477. Toppari, J., Larsen, J. C., Christiansen, P., Giwercman, A., Grandjean, P., Guillette, L. J., Jegou, B., Jensen, T. K., Jouannet, P., Keiding, N., LeVers, H., McLachlan, J. A., Meyer, O., Muller, J., Rajpert De Meyts, E., Scheike, T., Sharpe, R., Sumpter, J. P., and Skakebaek, N. E. (1996). Male reproductive health and environmental xenoestrogens. Environ. Health Prospect. 104, 741–803. Tsang, P. C., and Callard, I. P. (1987). Morphological and endocrine correlates of the reproductive cycle of the aplacental viviparous dogfish, Squalus acanthias. Gen. Comp. Endocrinol. 66, 182–189. Tyler, C. R., Jobling, S., and Sumpter, J. P. (1998). Endocrine disruption in wildlife: A critical review of the evidence. Crit. Rev. Toxixol. 28, 319–361.


45

Tyler, C. R., Sumpter, J. P., and Bromage, N. R. (1988). In vivo ovarian uptake and processing of vitellogenin in the rainbow trout Salmo gairdneri. J. Exp. Zool. 246, 171–179. van het Schip, F. D., Samallo, J., Broos, J., Ophuis, J., Mojet, M., Gruber, M., and Ab, G. (1987). Nucleotide sequence of a chicken vitellogenin gene and derived amino acid sequence of the encoded yolk precursor protein. J. Mol. Biol. 20, 196(2): 245–260. Verslycke, T., Vandenbergh, G. F., Versonnen, B., Arijs, K., and Janssen, C. R. (2002). Induction of vitellogenesis in 17a-ethinylestradiol-exposed rainbow trout (Oncorhynchus mykiss): A method comparison. Comp. Biochem. Physiol. C 132, 483–492. Wahli, W. (1988). Evolution and expression of vitellogenin genes. Trends Genet. 4, 227–232. Wahli, W., David, I. B., Wyler, T., Weber, R., and RyVel, G. U. (1980). Comparative analysis of the structural organization of two closely related vitellogenin genes in X. laevis. Cell 20, 107–117. Wahli, W., Wyler, D. I. B., Jaggi, R. B., Weber, R., and RyVel, G. V. (1979). Vitellogenin in Xenopus laevis is encoded in a small family of genes. Cell 16, 535–549. Wallace, R. A. (1970). Studies on amphibian yolk. IX Xenopus vitellogenin. Biochim. Biophys. Acta 215, 176–183. Wallace, R. A. (1985). Vitellogenesis and oocytes growth in nonmammalian vertebrates. In ‘‘Development Biology’’ (L. W. Browder, Ed.), Vol. 1, pp. 127–177. Plenum Press, New York. Wallace, R. A., and Begovac, P. C. (1985). Phosvitins in Fundulus oocytes and eggs. Preliminary chromatographic and electrophoretic analyses together with biological considerations. J. Biol. Chem. 15, 11268–11274. Wallace, R. A., and Bergink, E. W. (1974). Amphibian vitellogenin: Properties, hormonal regulation of hepatic synthesis and ovarian uptake and conversion to yolk proteins. Am. Zool. 14, 1159–1175. Wallace, R. A., and Jared, D. W. (1969). Studies on amphibian yolk. 8. The estrogen-induced hepatic synthesis of a serum lipophosphoprotein and its selective uptake by the ovary and trasformation into yolk platelet proteins in Xenopus laevis. Dev. Biol. 19, 498–526. Wallace, R. A., and Selman, K. (1978). Oogenesis in Fundulus heteroclitus. I. Preliminary observations on oocytes maturation in vivo and in vitro. Dev. Biol. 62, 354–369. Wallace, R. A., and Selman, K. (1981). The reproductive activity of Fundulus heteroclitus females from Woods Hole, Massachusetts, as compared with more southern locations. Copeia 1981, 212–215. Wallace, R. A., and Selman, K. (1985). Major protein changes during vitellogenesis and maturation of Fundulus oocytes. Dev. Biol. 110, 492–498. Wallace, R. A., Hoock, K. L., and Carnevali, O. (1990). Placement of small lipovitellin subunits within the vitellogenin precursor in Xenopus laevis. J. Mol. Biol. 213, 407–409. Wang, S. Y., and Williams, D. L. (1983). Purification of avian vitellogenin III. Comparison with VTG I and II. Biochem. 22, 6206–6210. Weigel, N. L. (1996). Steroid hormone receptors and their regulation by phosphorylation. Biochem. J. 319, 657–667. White, R., Jobling, S., Hoare, S. A., Sumpter, J. P., and Parker, M. G. (1994). Environmentally persistent alkylphenolic compounds are estrogenic. Endocrinology 135, 175–182. Wiley, H. S., and Wallace, R. A. (1981). The structure of vitellogenin. Multiple vitellogenins in Xenopus laevis give rise to multiple forms of the yolk proteins. J. Biol. Chem. 256, 8626–8634. Yamamoto, K., and Kikuyama, S. (1981). Purification and properties of bullfrog prolactin. Endocrinol. Jpn. 28, 59–64. Yamamoto, K., Yamashita, K., Hayakawa, Y., Hanaoka, Y., and Kikuyama, S. (1995). Development and application of a homologous radioimmunoassay for Xenopus prolactin. Gen. Comp. Endocrinol. 99, 28–34.

46


Yamamura, J., Adachi, T., Aoki, N., Nakajima, H., Nakamura, R., and Matsuda, T. (1995). Precursor-product relationship between chicken vitellogenin and the yolk proteins: The 40 kDa yolk plasma glycoprotein is derived from the C-terminal cysteine-rich domain of vitellogenin II. Biochim. Biophys. Acta 1244, 384–394. Yoshizaki, N., and Yonezawa, S. (1994). Cathepsin D activity in the vitellogenesi of Xenopus laevis. Dev. Growth DiVer. 36, 299–306. Yoshizaki, N., and Yonezawa, S. (1996). Salt concentration dependency of vitellogeni processing by cathepsin D in Xenopus laevis. Dev. Growth DiVer. 38, 549–556. Yoshizaki, N., and Yonezawa, S. (1998). Cystein proteinase plays a key role for the initiation of yolk digestion during development of Xenopus laevis. Dev. Growth DiVer. 25, 659–667. Zacharewski, T. (1997). In vitro bioassay for assessing estrogenic substances. Environ. Sci. Technol. 31, 613–623. Zerani, M., Vellano, C., Amabili, F., Carnevali, O., Andreoletti, G. E., and Polzonetti-Magni, A. M. (1991). Sex steroid profile and plasma vitellogenin during the annual reproductive cycle of the crested newt (Triturus carnifex Laur.). Gen. Comp. Endocrinol. 82, 337–344.

Structure, Evolutionary Conservation, and Functions of Angiotensin- and Endothelin-Converting Enzymes Nathalie Macours, Jeroen Poels, Korneel Hens, Carmen Francis, and Roger Huybrechts Laboratory for Developmental Physiology, Genomics and Proteomics, Katholieke Universteit Leuven, Naamsestraat 59, B-3000 Leuven, Belgium

Angiotensin-converting enzyme, a member of the M2 metalloprotease family, and endothelin-converting enzyme, a member of the M13 family, are key components in the regulation of blood pressure and electrolyte balance in mammals. From this point of view, they serve as important drug targets. Recently, the involvement of these enzymes in the development of Alzheimer’s disease was discovered. The existence of homologs of these enzymes in invertebrates indicates that these enzyme systems are highly conserved during evolution. Most invertebrates lack a closed circulatory system, which excludes the need for blood pressure regulators. Therefore, these organisms represent excellent targets for gaining new insights and revealing additional physiological roles of these important enzymes. This chapter reviews the structural and functional aspects of ACE and ECE and will particularly focus on these enzyme homologues in invertebrates. KEY WORDS: Endothelin-converting enzyme, Invertebrates, angiotensinconverting enzyme, Metalloproteases, ACE, ECE. ß 2004 Elsevier Inc.

I. Introduction Protease (peptidase, proteinase) is the general name for an enzyme that cleaves peptide bonds in peptides and proteins causing an irreversible modification or destruction to their substrate. Proteases are, based on their nature International Review of Cytology, Vol. 239 0074-7696/04 $35.00

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of catalysis, classified into five major groups: serine proteases, cysteine proteases, aspartic proteases, metalloproteases, and proteases with an unknown mechanism (Barret et al., 1998). From these groups, the metalloproteases, with more than 30 families identified to date, are most diverse. In mammalian physiology, the high interest in metalloproteases originates from the fact that they are attractive targets for drug design. In insects, regulatory peptides are the major and most diverse class of signaling molecules. They act as transmitters and modulators in the nervous system and as hormones controlling key physiological processes, behavior, and development. Peptidases are recognized as the key components of this peptidergic system, functioning in biosynthesis of active peptides from their precursors and in peptide inactivation. Structural analysis of metalloproteases from invertebrate and vertebrate species indicates that these enzymes are very well conserved and arose early during evolution. The evolutionary conservation of universal peptidesignaling mechanisms has led to the general acceptance of the idea that invertebrates provide excellent models for the study of genetics and physiology. The information and knowledge gained by studying invertebrate peptidases will provide novel insights in the process of evolution and may reveal conserved functions of vertebrate peptidases that have remained unknown until now.

II. Angiotensin-Converting Enzyme Mammals Angiotensin-converting enzyme (ACE, dipeptidyl carboxypeptidase I [DCP I], peptidase P, carboxycathepsin, peptidyl-dipeptidase A, peptidyl-dipeptidase I, kininase II, EC 3.4.15.1) was originally isolated in 1956 as a ‘‘hypertensin-converting enzyme’’ (Skeggs et al., 1956). Classified as a member of the M2 gluzincin family, ACE acts as a zinc-metalloprotease. The general reaction mechanism is identical for all zinc-metalloproteases. Coordination of the zinc ion within the catalytic site of metalloproteases is typically aVected by an HEXXH motif, a third zinc ligand (E) located carboxyl to this motif, and a water molecule. The histidine residues within the active site motif are known to directly coordinate the active site zinc ion, whereas the HExxH glutamate has been shown to coordinate weakly to zinc via the activated molecule, thus facilitating the acid–base catalytic mechanism. The renin-angiotensin system (RAS) plays a key role in the regulation of blood pressure and of fluid and electrolyte balance in mammals (Inagami, 1994). Being part of the renin–angiotensin system, ACE converts the decapeptide angiotensin I (AngI) into the potent vasopressor angiotensin II (AngII). ACE also inactivates the vasodilatory peptide bradykinin

ANGIOTENSIN- AND ENDOTHELIN-CONVERTING ENZYMES

49

(BK). Hence, inhibition of ACE results in lowering the blood pressure by preventing formation of the hypertensive AngII and by inactivation of the hypotensive BK (Fig. 1). ACE is one of the most studied mammalian peptidases, and in particular for medical applications. ACE inhibitors are widely used in treatment of hypertension, diabetic nephropathy, and heart failure (Dell’Italia et al., 2002). The production of captopril, the first clinically used ACE inhibitor, represents a classical example of drug design (Menard and Patchett, 2001).

A. Structure and Isoforms Vertebrate ACE is classified as a type I ecto-enzyme, with the major part of the protein, including the active site, located on the external surface of cells. ACE is anchored in the plasma membrane by the C-terminal part of the protein. There is an N-terminal signal peptide present, required for the passage through the endoplasmatic reticulum. Two distinct forms of ACE are known: a somatic form (sACE), which is found in many tissues, and a

FIG. 1 Components of the renin–angiotensin system with the principal enzymes ACE and ACE2 involved in the generation of biologically active peptides. m; vasoconstrincting eVect; d; vasodilating eVect.

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smaller iso-enzyme, germinal ACE (gACE, also called testicular ACE [tACE]), which is found exclusively in testes, where it has a crucial role in fertility (Fig. 2). The soluble, circulating form of the sACE protein is produced by a ‘‘secretase,’’ probably a zinc-metalloprotease, which releases ACE from the membrane by cleaving oV the extracellular part (Turner and Hooper, 2002). The two forms diVer in that the somatic isoform contains two active sites, whereas the germinal form contains only one. sACE is composed of two very similar domains (C and N domain), which are both catalytically active (Wei et al., 1991). tACE is, apart from its N-terminal sequence, identical to the C-domain of sACE. The close similarity of the N and C domains of mammalian sACE and the fact that exons 4–11 and 17–24 of the human ACE gene, which code for the N and C domain, respectively, are very similar in size and have similar codon phases at exon–intron boundaries, strongly indicates that the coding gene arose from duplication of an ancestral ACE gene coding for a single domain enzyme (Cornell et al., 1995). The ancestral copy is predicted to be the C-domain, equivalent to the germinal isoform (Lattion et al., 1989). The

FIG. 2 Schematic representation of human and Drosophila ACE protein and gene structure. Exons are indicated by boxes. Exons and corresponding amino acid regions in the protein are colored correspondingly. Zinc-binding sites (HExxH) are indicated. Human somatic and testicular promoters are indicated by arrows. One-kilobase intronic sequence was omitted at the // signs. (See also color insert.)


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two isoforms are transcribed from a single gene, under the control of two distinct promoters. sACE is transcribed from a promoter region upstream of the duplication, and tACE from a promoter within intron 12 (Langford et al., 1991). The two domains of sACE have relatively broad substrate specificities and share some enzymatic features, but there are several biochemical properties that diVerentiate the two active sites (Corvol et al., 1995). For example, the hematopoietic stem cell regulator, N-acetyl-Ser-Asp-Lys-Pro (AcSDKP), is an in vivo substrate for the N-domain, but not for the C domain of human ACE (Azizi et al., 1996). The N and C domain can also be diVerentiated by their response to chloride ions, with the C domain’s specific activity being dependent on chloride ion concentration (Wei et al., 1991). Their inhibitory aYnity profiles also diVer, illustrated by the existence of an N-domain-specific inhibitor (RXP 407) (Dive et al., 1999).

B. Biological Role ACE acts as a peptidyl-dipeptidase, removing the C-terminal dipeptide from its substrate, but it can also act as an aminopeptidase and as an endopeptidase on peptides that are amidated at the C terminus. To date, only angiotensin I, bradykinin (Yang, 1970), and the haemoregulatory peptide Ac-DSKP (Azizi et al., 2001) have been confirmed as in vivo substrates for mammalian ACE. The conversion of AngI into the vasoconstrictor AngII and the degradation of the vasodilator bradykinin reflect the key role of ACE as a component of the mammalian RAS. In recent years, the RAS turned out to be a far more complex regulatory system that involves more angiotensin-derived mediators (AngI, AngII, AngIII, AngIV, Ang(1–9)), Ang(1–7)) then initially recognized (Fig. 1). To this day, it includes four identified receptors (Turner and Hooper, 2002). In vivo models using ACE-deficient mice have greatly advanced the knowledge of the RAS and confirmed the role of ACE in blood pressure regulation and fertility. Knock-out mice all show reduced blood pressure, altered kidney morphology, impaired somatic growth, and male sterility (Esther et al., 1996; Eriksson et al., 2002). These mice produce normal numbers of sperm that are indistinguishable from wild-type sperm in assays of viability, motility, capacitation, and induction of the acrosome reaction. Absence of ACE, however, does cause defects in sperm transport in the oviducts and in binding to zonae pellucidae (Hagaman et al., 1998). Knockout of tACE, leaving the sACE intact, has proven that tACE expression is both necessary and suYcient for fulfilling the role of ACE in male fertility (Ramaraj et al., 1998). Recently, immunocytochemical studies have shown that tACE is completely shed from

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the sperm membrane before ejaculation. This would imply that the defects in sperm transport in the oviducts and in binding to zonae pellucidae after tACE knockout are not directly related with tACE activity (Metayer et al. 2002), so the exact role of tACE remains to be uncovered. The fact that to date no in vivo substrate has been found for tACE is the main bottleneck in tACE research. Expression of sACE, either in renal proximal tubes or in vasculature, is suYcient for maintaining normal kidney functions. However, for maintaining blood pressure, sACE must be expressed in vascular endothelial cells (Kessler et al., 2003). Expression of tACE alone is suYcient to restore male fertility in mutant mice without curing other problems of Ace / mice (Ramaraj et al., 1998). However, sACE cannot substitute for tACE, demonstrating that the two isozymes are not interchangeable for fertility functions (Kessler et al., 2000). Yet tACE expression in serum of Ace/ mice can substitute for sACE in maintaining normal renal structure and functions without restoring blood pressure (Kessler et al., 2002). All this indicates that the two isoforms of ACE are indispensable, as suggested by their high degree of evolutionary conservation. Also, the specific physiological function of ACE requires its expression in the correct tissue. The peptide AcSDKP reversibly prevents the recruitment of pluripotent haematopoietic stem cells into the S phase of the cellular cycle by maintaining them into the Go phase. Because this peptide has been found to be an in vivo substrate specific for the N domain of sACE, it is suggested that ACE is implicated in the process of hematopoietic stem cell regulation by permanently degrading this natural circulating inhibitor (Azizi et al., 2001; Rousseau et al., 1995). However, in a very recent study, mutation of the N domain zinc-binding site in mice, rendering this N domain inactive indeed, resulted in accumulation of AcDSKp, but no physiological eVect was seen (Fuchs et al., 2004). ACE acts as an endopeptidase on the multifunctional neuropeptides substance P and cholecystokinin and may degrade the luteinizing hormone releasing hormone (LH–RH) (Skidgel and Erdos, 1985). Its biological significance in this context is not known, and the possibility of functional cross talk between the systems has not been excluded (Turner and Hooper, 2002). The broad in vitro specificity (Table I) and the presence of ACE in species that lack an identifiable RAS shows that the relevance of ACE to the animal must be more complex than just its role in the RAS.

C. ACE2 Up until now, only one other member of the M2 family other than ACE has been identified. Almost 50 years after the discovery of ACE itself, two groups (Donoghue et al., 2000; Tipnis et al., 2000) introduced the first homolog of

TABLE I Primary Sequences of Peptides Hydrolyzed by ACE and ACE2 ACE2

ACE Substrate

Sequence

Substrate

Sequence

Ang I

DRVYIHPF#HL

Ang I

DRVYIHPFH#L

Ang (1-9)

DRVYIHP#FH

Ang II

DRVYIHP#F

Ang (1-7)

DRVYI#HP

Apelin 18 (36)

(. . .)QRPRLSHKGPMP#F

Bradykinin

RPPGF#SP#FR

Des-Arg9 bradykinin

RPPGFSP#F

Enkephalins

YGG#FM(L)-NH2

b-Casamorphin

YPFVEP#I

Substance P

RPLPQQFF#G#LM-NH2

Neocasamorphin

YPVEP#I

LH-RH

EHW#SGLRPG

Dynorphin A 1-13

YGGFLRRIRPKL#K

Haemoregulatory peptide

NAcSD#KP

Neurotensin 1-8

Pe-LYENKP#R

CCK-8

NY#(SO3H)MGWM#DF-NH2

Cleavage sites are indicated by an arrow.

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mammalian ACE. This protein and its homolog were named ACE2 and ACEH, respectively, and were shown to be an essential regulator of heart function (Crackower et al., 2002). Human ACE2 is an 805–amino acid–long type I integral membrane protein, including a 17–amino acid N-terminal signal peptide and a putative C-terminal membrane anchor. The extracellular domain shares a 42% sequence identity with the catalytic domain of sACE. It is 33% identical to human testicular (germinal) ACE and also contains a single catalytic site. ACE2 contains six putative N-linked glycosylation sites on the extracellular part (Tipnis et al., 2000). The C-terminal domain of ACE2 shares a 48% sequence identity with collectrin (Vickers et al., 2002; Zhang et al., 2001), a developmentally regulated renal transmembrane glycoprotein. The many similarities in the genomic sequence of ACE2 and ACE indicate that there is an evolutionary relationship between these two genes (Turner and Hooper, 2002). ACE2 transcripts are detected in heart, lung, kidney, the gastrointestinal tract, and testis (Harmer et al., 2002; Komatsu et al., 2002). ACE2 is located at the endothelium of coronary and intrarenal vessels and in the renal tubular epithelium. Like ACE, ACE2 also appears to be susceptible to cleavage and secretion from the cell surface (Donoghue et al., 2000). However, the sequence identity of the juxtamembrane regions of ACE and ACE2 is diVerent. The cleavage site present in ACE (–Ala-Arg Ser-Glu–) is absent in ACE2. Hence, a diVerent secretase may be involved in the shedding of ACE2. ACE2 does not have the same substrate specificity as ACE, nor is it inhibited by the typical ACE inhibitors such as enalapril, lisinopril, and captropril. ACE2 hydrolyzes AngI and AngII but cleaves neither bradykinin nor the typical ACE substrate Hip-His-Leu (Fig. 1). To hydrolyze AngI and AngII, ACE2 acts as a carboxypeptidase, removing one amino acid from the C terminus of the substrate. This is also the case for kinetensin, des-Arg9 bradykinin, and neurotensin (Donoghue et al., 2000; Turner et al., 2002). Several other regulatory peptides, including the enkephalins, are not substrates of ACE2. It seems that ACE2 prefers to cleave a single C-terminal hydrophobic residue from its substrates (Table I) and consequently does not hydrolyze bradykinin that contains an arginine at its C terminus. ACE2 is the first mammalian carboxypeptidase identified that contains the HExxH active site. ACE2 also acts on apelin-13 and apelin-36 peptides, of which the role is not yet fully understood but involves an eVect on hypertension and vasoconstriction (Lee et al., 2000). Two opioid peptides, dynorphin A(1–13) and b-casamorphin, are also ACE2 substrates (Eriksson et al., 2002; Vickers et al., 2002). These peptides activate G protein-coupled opioid receptors that regulate pain perception and are suggested to negatively aVect cardiomyocyte contractility (Pugsley, 2002; Ventura et al., 1992; WenzlaV et al., 1998). The diVerences in substrate specificity between ACE and ACE2 have important physiological consequences. Whereas ACE activity leads to the


55

formation of the vasoconstrictor AngII, ACE2 is involved in the formation of vasodilator mediators (Danilczyk et al., 2003). The targeted disruption of ACE2 in mice results in increased AngII levels, impaired cardiac contractility, and up-regulation of hypoxia-induced genes in the heart (Crackower et al., 2002). Renal ACE is implicated in diabetic nephropathy. In the diabetic kidney, levels of ACE2 transcription and protein expression are downregulated. It is suggested that this regulation occurs through the actions of ACE2 in the RAS (Tikellis et al., 2003). Another, rather unexpected function of ACE2 was recently discovered. It was found to act as a functional receptor for the Severe Acute Respiratory Syndrome (SARS) coronavirus (CoV) (Dimitrov, 2003; Li et al., 2003; Xiao et al., 2003). SARS is the abbreviation for the CoV-induced disease that causes SARS, with a mortality rate of approximately 10% (Ksiazek et al., 2003; Kuiken et al., 2003). The virus associates with cellular receptors to mediate infection of the target cells (Gallagher and Buchmeier, 2001; Holmes, 2003). ACE2, acting as a cellular receptor, eYciently binds to the S1 domain of SARS-CoV, eYciently supporting the replication of SARSCoV. Adding a soluble form of ACE2, but not that of ACE, blocks the association of SARS-CoV and its target cell. Replication of the virus in cells and the formation of syncitia can be blocked by antibodies against ACE2, which indicates the importance of ACE2 in the replication of SARS-CoV (Li et al., 2003). Several attempts to reveal the three-dimensional structure of ACE2 have been made very recently. This provides an insight into the structural variations underlying the diVerences in substrate specificity between ACE and ACE2. It seems that these diVerences occur in the ligand binding cavity, influencing the binding of the peptide carboxyterminus (Guy et al., 2003). The crystal structure of ACE2, native and inhibitor bound, revealed a large inhibitordependent bending movement of the two enzyme domains to one another, positioning important catalytic residues (Towler et al., 2004). It has been suggested that ridges surrounding the cavity at the top of the molecule serve as a binding region for SARS (Prabakaran et al., 2004). The discovery of this SARS-related function of ACE2, together with the knowledge of its structure, may contribute substantially to the possible treatment of SARS patients.

III. ACE in Invertebrates Lamango and Isaac (1993) reported the metabolization of [D-Ala2, Leu5] enkephalin by head membranes of the housefly, Musca domestica. This hydrolysis was only partially inhibited by phosphoramidon, an inhibitor of endopeptidase-24.11. The combination of phosphoramidon and captopril,

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however, fully inhibited the cleavage of enkephalin, suggesting the presence in this fly of a second enkephalin-metabolizing enzyme in addition to endopeptidase-24.11. It turned out to be zn 87-kDa-enzyme that was activated by ZnCl2 and inhibited by EDTA as well as the mammalian ACE inhibitors captopril, fosinoprilat, and enalaprilat. This represented the discovery of the first invertebrate ACE homolog (Lamango and Isaac, 1994). Following the report on this ACE-like activity, the first results of putative Drosophila melanogaster ACE cDNA clones were published (Cornell et al., 1993). Since then, ACE homologs have been detected in many invertebrate species. Cloning or purification of ACE from the fruit fly D. melanogaster (Cornell et al., 1995; Tatei et al., 1995), the cattle tick Boophilus microplus (Jarmey et al., 1995), the leech Theromyzon tessalutum (Laurent and Salzet, 1996), the housefly M. domestica (Lamango et al., 1996), the buValo fly Haematobia irritans exigua (WijVels et al., 1996), the mosquito Anopheles stephensi (Ekbote et al., 1999), the silk moth Bombyx mori (Quan et al., 2001), the grey fleshfly Neobellieria bullata (Vandingenen et al., 2002b), and the locust Locusta migratoria (Macours et al., 2003a) have contributed to the molecular, biochemical, and evolutionary characterization of invertebrate ACEs. The first D. melanogaster ACE homolog that was discovered is abbreviated as Ance, because ace was already used for acetylcholine esterase in the fruit fly. Of the invertebrate ACEs known today, the two D. melanogaster ACE homolog (Ance and Acer) have been studied in most detail. Ance is also known under the name Race, as the enzyme was discovered, sequenced, and named by two groups simultaneously (Cornell et al., 1995; Tatei et al., 1995). Nowadays, the focus on ACE research is beginning to shift from purification-characterization to a more physiological point of view. Considering the fact that most invertebrates have an open circulatory system, there is probably no need for a renin–angiotensin system analogous to that found in vertebrates, at least not for the regulation of blood pressure. Hence, understanding the biology of ACE homologs in invertebrates may, in due time, provide substantial information on additional roles for ACE.

A. Structure In contrast to the occurrence of both a double and a single domain ACE in vertebrates, characterization of ACE homologs from diVerent invertebrates showed that the overall size of these proteins was comparable to the nonglycosylated testicular form of mammalian ACE. Cloning of the corresponding cDNAs confirmed the existence of this single-domain form in insects. Until now, there has been no evidence for a functional two-domain form of insect ACE. In addition to the absence of a two-domain form, a structural diVerence exists between insect ACE and the mammalian enzyme,


57

located in the N- and C-terminal ends. Cloning of the D. melanogaster ACE homologue (Ance) revealed that the insect form of ACE lacks both the heavily glycosylated N-terminal extension and the C-terminal transmembrane and cytoplasmic regions that are present in mammalian testicular ACE. The enzyme contains an N-terminal secretory signal peptide, which indicates that the mature enzyme is being secreted. Hence, Ance was predicted to code for a soluble secreted protein (Fig. 2). Confirmation of this prediction came when 90% of recombinantly expressed Ance activity was found in the cell culture medium (Cornell et al., 1995). Although insect ACE activity was primarily detected in a membrane preparation from heads of the housefly, M. domestica (Lamango and Isaac, 1994), this ACE protein was later found not to behave as an integral membrane protein. The membrane-associated activity was probably the result of a contamination of the membrane fraction with soluble ACE (Lamango et al., 1996). Therefore, the general form of insect ACE seems to be a secreted single-domain protein with no recognizable C-terminal membrane anchor. The molecular identification of the ACE enzyme in B. microplus, however (Jarmey et al., 1995), which does posses a C-terminal hydrophobic region, showed that the membrane-bound form of ACE is not restricted to vertebrate organisms. Also, in the cockroach Leucophaea maderae and the noctoid moth Lacanobia oleracea, a substantial amount of ACE activity was detected in head membranes (Isaac et al., 2002). The nature of this membrane-associated ACE activity in these two organisms is, as yet, unidentified. 1. Sequence Conservation Sequence comparison between human testicular ACE and several invertebrate ACE homologs showed that the overall sequence identity is approximately 40% and exceeds 70% in the regions surrounding the active site. In human ACE, 17 and 8 potential N-linked glycosylation sites (Asn-Xaa-Ser/ Thr motifs) are present in the somatic isoform and in the germinal isoform, respectively. This contrasts to the situation in invertebrates, where ACE homologs have only three or fewer potential glycosylation sites. In D. melanogaster, it was shown that glycosylation of ACE is not required for the enzymatic properties. Expression of an unglycosylated mutant of Ance resulted in the secretion of a catalytically active enzyme, with the same substrate and inhibitor aYnity as the wild-type enzyme (Williams et al., 1996). 2. Three-Dimensional Structure Kim et al. (2003) determined the crystal structure of the D. melanogaster ACE homolog Ance. Ance is composed of 21 a-helices and three antiparallel b-strands. The substrate-binding channel exists as a large continuous internal

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path that comprises the entire protein molecule. The composition of this substrate-binding site is unusual in that it is composed of two unequal-sized ‘‘chambers’’ analogous to a peanut shell. The large chamber, referred to as the N chamber, contains the binding site of the N-terminal sequence of the substrate angiotensin I. The smaller chamber, called the C chamber, will bind the carboxy-dipeptide of the substrate. The size of these chambers is suYciently large for binding short peptide substrates. The bottleneck region connecting these two chambers encloses the zinc ion critical for catalysis and locates the inhibitor binding sites. The opening of these chambers to the exterior appears to be relatively small for the passage of peptide substrates, so flexibility and a so-called ‘‘breathing motion’’ are probably required for eYcient catalysis.

B. Evolutionary Aspects The presence of significant ACE activity was reported in extracts of the horse anemone, Actinia equine, a member of the cnidarians (Coates et al., 2000b). To this day, no ACE peptidase activity has been detected in nematode tissues. However, there is one copy of an ACE-like gene (CD42D8.5) with a recorded expressed sequence tag (EST) present in the genome of the nematode Caenorhabditis elegans. Yet taking a closer look at this ACE-like gene reveals that the amino acids crucial for enzyme activity are absent. The sequence lacks all three zinc coordinating His residues, which means that the C. elegans ACE homolog does not have a functional zinc-binding domain. Therefore, it cannot be classified as a metallopeptidase homolog. Studies on this ACE-like nonmetallopeptidase, termed ACN-1, demonstrated that the protein plays an essential role in larval molting and adult morphogenesis. If ACN-1 is indeed evolutionarily related to the ancestral ACE, it means that during evolution it lost its peptidase activity but acquired sites for new molecular interactions that have a direct or indirect eVect on the molting process in nematodes (Brooks et al., 2003). This phenomenon can be further expanded to other organisms and enzymes, as in the genome of D. melanogaster and Anopheles gambiae, 15–25% of the protease-like genes lack one or more catalytic residues. Some of these proteins have probably lost the peptidase activity of their ancestor while acquiring new functions. The analysis of the human ACE gene at the structural level forwarded the view that the vertebrate somatic form was transcribed from a tandem duplication (Lattion et al., 1989). In all vertebrate species examined, from humans at the top down to the electric fish Torpedo marmorata (Turner et al., 1987), the somatic ACE enzyme exists as the two-domain form. The analysis of the ACE gene of the housefly M. domestica revealed that this organism comprises a smaller form of the enzyme, indicating that this species does not


59

contain the duplicated form of ACE. The existence of this single-domain isoform was confirmed when the Ance cDNA of D. melanogaster was sequenced (Cornell et al., 1995). All other invertebrate ACE enzymes that were discovered later on also consisted of the single-domain isoform. This leads to the view that invertebrate ACE represents the ancestral copy of the vertebrate two-domain enzyme and that the duplication event occurred before vertebrate radiation. However, this course of ACE evolution started to be questioned when the characterization of D. melanogaster ACE homologs began. The high level of homology between Ance and Acer points toward the existence of an ancestral ACE gene. In addition, studying the relationship of Ance and Acer to the N and C domains of human somatic ACE showed that Ance was more enzymatically related to the C domain than to the N domain of human ACE (Williams et al., 1996). Acer, however, has much less in common with this C domain. The active sites of the N domain of sACE and Acer share structural features that permit the binding of the RXP407 inhibitor, which does not bind the C domain of sACE. An analysis of the genomic region around the Ance gene in D. melanogaster has led to the identification of four additional ACE-like genes (Isaac et al., 2000). From these four, Ance 3, 4, and 5 seem to be transcribed, because ESTs were reported for these genes. With the information available today, no EST seems to be present for Ance 2. The predicted protein sequence corresponding the Ance 3 gene includes a Cterminal transmembrane anchor, as is seen in vertebrate ACE. Ance 2 codes for an ACE-like protein with a putative signal peptide, but without a Cterminal membrane region. Both genes lack crucial active site residues. These two gene predictions are interesting because alternative splicing of the predicted transcripts would lead to a two-domain, membrane- bound ACE-like product, as is seen in the vertebrate somatic form (Coates et al., 2000a). Taken together, this information indicates that the duplication event that gave rise to the vertebrate tandem protein predates the divergence of the Ecdysozoa and deuterostomes, and that the diVering activities on the N and C domains have been maintained over a long period of time.

C. Substrate Specificity Insect ACE is capable of hydrolyzing the known in vivo substrates of mammalian ACE, angiotensin I, bradykinin, and the hemoregulatory peptide AcDSKP. However, there are no reports of insect homologs of these known in vivo substrates, and none of the many insect peptides isolated to date shows any structural resemblance to them. The search for angiotensin I–like peptides in the brain of several insect species could not prove otherwise (Isaac et al., 1998b; Schoofs et al., 1998). In addition, a search in the D. melanogaster

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MACOURS ET AL.

genome did not reveal any potential homologs of the precursor proteins of angiotensin I and bradykinin (Isaac et al., 2000). This is in contrast to the situation in leeches, where several reports on the presence of members of the renin–angiotensin system have been published (Laurent and Salzet, 1996; Salzet and Verger-Bocquet, 2001). Cleavage experiments with ACE from M. domestica showed that insect ACE exhibits a broad in vitro substrate specificity. The in vitro substrates of mammalian ACE, cholecystokinin, enkephalins, substance P, and LH-RH, are successfully cleaved by the endopeptidase activity of Musca ACE. This shows that insect ACE can hydrolyze C-terminally amidated peptides in vitro, functioning as an endopeptidase (Isaac et al., 1997a; Lamango et al., 1997). M. domestica ACE is unable to hydrolyze either CCAP (crustacean cardioactive peptide) or proctolin. Apparently, the penultimate C-terminal prolyl residue of proctolin makes this peptide resistant to ACE hydrolysis. The lack of hydrolysis of CCAP is likely a result of the presence of a disulfide bridge, which results in a secondary structure that prevents access to the active site. A novel activity of insect ACE, namely, prohormone processing, was suggested when it was found to be capable of successfully removing basic dipeptides from locustamyotropin oligopeptides possessing a C-terminal Gly-Lys-Arg and Gly-Arg-Arg extension (Isaac et al., 1998b). Many insect peptides are synthesized as inactive prohormones and need posttranslational processing to acquire their biological activity. In several prohormones, the sequence -Gly-Arg/Lys-Arg- is an internal consensus recognition sequence for endoproteolysis by prohomone convertases, generating peptide intermediates with a C-terminal –Gly-Arg/Lys-Arg– extension. These are substrates for carboxypeptidase E (CPE) that sequentially removes the two basic amino acids from the C terminus, which is followed by amidation of the C terminus (Isaac et al., 1997b). The fact that insect ACE is capable of hydrolyzing these partially processed prohormone-like peptides, together with the intracellular colocalization of the enzyme with locustamyotropin peptides, implies a role for ACE in the biosynthesis of peptide hormones and neuropeptides (Schoofs et al., 1998). Zhu et al. (2001) and coworkers demonstrated that the hexapeptide trypsin modulating oostatic factor from N. bullata (Neb-TMOF) (Bylemans et al., 1994) is selectively hydrolyzed by ACE that is present in substantial amounts in the fly’s haemolymph (Zhu et al., 2001). Neb-TMOF is capable of regulating vitellogenesis and is thought to be released by the ovaries and transported to the midgut, where it terminates the protein meal–induced trypsin biosynthesis. Captopril feeding experiments suggest that NebTMOF represents a prime candidate for being an in vivo substrate for circulating insect ACE (Vandingenen et al., 2002a). TMOF was also detected in Aedes aegypti, probably synthesized by the brain (Borovsky and Meola,


61

2004). However, this Aea-TMOF, with six prolines at its C terminus, is not cleaved by Neb-ACE (K. Hens, personal communication). Recently, new substrates were purified from the ovaries of N. bullata. Their primary structures were identified as NKLKPSWQWISL (Neb-ODAIF), NKLKPSQWISLSD (Neb-ODAIF-11–13), NKLKPSQWI (Neb-ODAIF- 11–9), NKLKPSQ (Neb-ODAIF- 11–7), SLKPSNWLTPSE (Neb-ODAIF-2), and LEQIYHL. These peptides show significant homology to the N-terminal part of fly yolk proteins (Hens et al., 2002; Vandingenen et al., 2002b). Kinetic analysis of these peptides with insect ACE and human ACE shows that the N. bullata ACE shares enzymatical properties with the C-domain sACE, in addition to features that seem unique to invertebrate ACE. Characterization of the Ance protein demonstrated that this enzyme eYciently hydrolyzes angiotensin I and bradykinin (Cornell et al., 1995; Williams et al., 1996). Ance can operate as an endopeptidase, for instance on tachykinins, substance P, and leucokinins. Still, the nonamidated form of a peptide is always preferentially cleaved over the amidated form. Furthermore, insect ACE preferentially acts as a dipeptidase and shows reduced tripeptidase activity. Of the insect peptides tested, locustatachykinin (Lom TK-1) proved to be the best substrate for Ance. However, the D. melanogaster tachykinins turned out to be poor Ance substrates (Siviter et al., 2002). Acer is not capable of cleaving angiotensin I (Houard et al., 1998), and most peptides tested were hardly degraded by Acer (Siviter et al., 2002). To this day, there are no clues concerning the identity of the endogenous Acer substrates.

D. Biological Role The structural and biochemical similarities between mammalian and invertebrate ACE do not necessarily imply a functional conservation as well. Although the presence of an invertebrate RAS has been reported in a leech that feeds on ducks (Laurent et al., 1995; Laurent and Salzet, 1995a,b, 1996), no other components of the RAS other than ACE have been described in insects or other invertebrates. It is therefore highly improbable that this invertebrate enzyme would physiologically act as a converting enzyme for an angiotensin-like peptide. As already mentioned, functionality of mammalian ACE is, however, not restricted to the RAS. Both its broad substrate specificity and widespread tissue distribution indicate its involvement in several other (but, unfortunately, still poorly described) physiological functions. To fully unravel the in vivo role of ACE in insects, information about substrate specificity and tissue and cellular localization of the enzyme, as well as its putative colocalization with its substrates, has to be combined. These

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data, together with the studies of ACE mutants and the eVects of ACE inhibitors on insect physiology, will eventually lead to the elucidation of the roles of insect ACE. Today’s concept on insect ACE functionality focuses on its possible role in reproduction, development, and defense. 1. Reproduction The discovery of ACE in testes of several insect species, including H. irritans exigua (WijVels et al., 1996), Lymantria dispar (Loeb et al., 1998), N. bullata, the Colorado potato beetle Leptinotarsa decemlineata, and the locust L. migratoria (Isaac et al., 1999; Schoofs et al., 1998), implies that the function of tACE may have been conserved during the course of evolution. In L. decemlineata, ACE was detected in the germ cells, more particularly in developing spermatids and mature spermatozoa, whereas in L. migratoria, ACE was found in peripheral somatic cell bodies of the apical compartment of the testicular follicles and not in the germ cells. In the haematophagus fly H. irritans, expression of testicular ACE is induced after a blood meal, indicating a specific role in the maturation of spermatozoa (WijVels et al., 1996). The possible importance of testicular ACE in reproduction became apparent by the observation that male D. melanogaster transheterozygotes of two mutant Ance alleles were found to be sterile (Tatei et al., 1995), with the infertility linked to the failure of the spermatocytes to develop into active mature spermatozoa. Spermatogenesis proceeds normally in the mutants up to the spermatid stage, but these spermatids do not succeed in diVerentiating into spermatozoa. This change is accompanied by nuclear scattering and an abnormal morphology of the spermatids. Together with the observed expression pattern of Ance, the enzyme is believed to be required for spermatid diVerentiation through the processing of a regulatory peptide synthesized within the developing cyst (Hurst et al., 2003). The involvement of ACE in the control of spermatogenesis was also demonstrated by studies of the eVects of AII and bovine ACE on the ability of the testes of L. dispar to synthesize and release ecdysteroids in response to testis ecdysiotropin (TE). The eVect of TE on the initiation and up-regulation of ecdysteroid synthesis could be imitated by both AII and bovine ACE, raising the possibility that ACE is involved in the production of an AII-like peptide that modulates ecdysteroid synthesis in the testes of insects (Loeb et al., 1998). In female adults of the mosquito A. stephensi, ACE activity increases by a factor 2.5 after a blood meal. It is not known where this induced ACE is synthesized, but it accumulates in developing ovaries and in mosquito eggs (Ekbote et al., 1999). Adding ACE inhibitors (captopril and lisinopril) to the blood meal of the mosquito leads to a size reduction of the batch of eggs laid in


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a dose-dependent manner. The ACE inhibitors reduce fecundity by interfering with the transfer of the oocytes along the oviducts (Ekbote et al., 2003a). In the reproductive tissue of the tomato moth Lacanobia aleracea, almost all of the ACE activity is concentrated in the accessory glands of the male and in the spermatheca and bursa copulatrix of the (virgin) female. These high activities may imply the importance of ACE for peptide metabolism in the male and female reproductive tract. Changes in the levels of ACE activity in the reproductive tissues of males and females during mating, point toward a transfer of ACE from male to female during time of copulation (Ekbote et al., 2003b). This male ACE donated in the spermatophore might form a hitherto unknown male component of an energy-producing metabolic pathway, as is seen in the serine endopeptidase-dependent proteolytic cascade in the spermatophore of B. mori (Aigaki et al., 1987, 1988). In the fleshfly N. bullata, feeding of captopril results in an increase of vitellogenin titers in the fly hemolymph. It has been suggested that this eVect is mediated through the trypsin-modulating oostatic factor TMOF. As already discussed, Neb-TMOF is an in vitro substrate of insect ACE. However, if TMOF were to be inactivated by ACE, inhibition of ACE would lead to an increase of TMOF levels in the hemolymph, resulting in a reduced vitellogenin synthesis. Therefore, an activating eVect of ACE on TMOF is postulated (Vandingenen et al., 2001). The discovery of ACE interactive peptides in the ovaries of the fleshfly (Vandingenen et al., 2002a), and the discovery that these peptide substrates originate from yolk gene products, strengthen the idea that ACE is a regulator of vitellogenic and developmental processes (Hens et al., 2002; Vandingenen et al., 2002b). These physiological observations, in combination with the discovery of ACE protein in the gonads of numerous insects, undoubtedly show that ACE is a vital enzyme in insect reproduction. The further elucidation of the full role of ACE in this process necessitates including the description of all endogenous substrates. 2. Development and Metamorphosis The observation that some mutations in the D. melanogaster Ance gene result in death during larval/pupal stages (Tatei et al., 1995) indicates that insect ACE is important for normal growth and development. Many reports from diVerent and independent laboratories have confirmed the role of insect ACE in development and metamorphosis. Along with the cloning of the first D. melanogaster ACE homolog (Race, Ance), the gene coding for this protein was shown to be a target of the homeobox regulatory gene zerknullt (zen) and decapentaplegic (dpp) (Tatei et al., 1995). A 533-bp-long enhancer located in the Ance promotor region covers three zen protein binding sites and was shown to mediate selective expression in the amnioserosa. During early

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development, Ance is activated by zen and becomes associated with the diVerentiating amnioserosa and with the anterior and posterior midgut, where it persists throughout embryogenesis. In late embryos, Ance expression is detected in epithelial cells of the midgut and in the pericardial cells of the heart. Ance expression is lost from the presumptive amnioserosa in zen and dpp mutants; the expression in the gut stays unaVected. In D. melanogaster larvae, high expression of Ance is found before and peaks during pupal development, indicating a physiological role for the enzyme during metamorphosis (Wilson et al., 2002). Also, in the tomato moth, L. oleracea, ACE activity increases approximately fourfold in the last larval instar and in the early pupal stage (Ekbote et al., 2003b). In insects, each larval molting is preceded by high ecdysteroid titers, and for pupal and adult development, multiple pulses of ecdysteroid hormones are essential. In wing discs of B. mori, the ACE gene (BmAcer) was found to be directly 30-hydroxyecdysone (20E) inducible in a dose-dependent manner. Because this ecdysteroid-dependency was not found in other tissues examined, transcription of BmAcer seems to be organ specific (Quan et al., 2001). During the transition from larva to pupa in D. melanogaster, the expression of Ance in imaginal disc cells was found to be induced by ecdysteroids. These data point toward a fundamental role for ACE during metamorphosis of holometabolous insects. Several relationships between ACE and metamorphosis are possible: the conversion of precursors into biologically active peptides necessary for metamorphosis, inactivation of signaling peptides, or recycling of amino acids from larval proteins for the synthesis of pupal and adult cuticle/tissues. In A. stephensi, ACE expression is induced by a blood meal and ACE accumulates in developing ovaries, from which it passes into the mosquito eggs (Ekbote et al., 1999). This indicates a role for ACE-generating peptides that are essential for oocyte development and embryogenesis. The characterization of ACN-1 in the nematode C. elegans shows that it is not necessarily the proteolytic function of ACE that is involved in developmental processes. The hypodermal expression of acn-1 is under the control of nuclear hormone receptors that regulate molting in C. elegans. Functional acn-1 knockout by RNAi causes morphological defects in larvae and adults. More specifically, ACN-1 was found to be required for larval molting, male tail development, and formation of adult alae (Brooks et al., 2003). 3. ACE in the Central Nervous System: A Role in Prohormone Processing? In N. bullata, L. migratoria, L. decemlineata, L. maderae, the walking stick insect Carausius morosus, B. mori, and the caterpillar Mamestra brassica, ACE immunoreactivity is seen in neuropil regions of the brain. In L. Migratoria, L. maderae, B. mori, and M. brassica, ACE immunoreactivity is also present in


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neurosecretory cells of the brain (Schoofs et al., 1998). In L. migratoria, immunoreactive staining can also be traced in the controlateral nervus corporis cardiaca I (NCC I), corpora cardiaca (CC), and suboesophageal ganglion (SOG), and in C. morosus, L. decemlineata, and L. maderae, ACE immunoreactivity was also detected in the corpora cardiaca (Veelaert et al., 1999). The presence of ACE in neurosecretory cells and neuropil regions of the insect brain points toward a dual role for ACE in the insect nervous system (Schoofs et al., 1998). The localization of ACE in the neuropil indicates the metabolic inactivation of peptidic neurotransmitters. The presence of peptides from the adipokinetic hormone (AKH), kinin, and locustatachykinin family in the central nervous system (CNS), as well as the knowledge that these peptides are hydrolyzed by ACE (Lamango et al., 1997), indicates that they might be potential in vivo substrates for insect neuronal ACE. In neurosecretory cells, ACE is likely to be involved in the conversion of prohormones to active peptide hormones that are secreted into the hemolymph by the corpora cardiaca or corpora allata (Isaac et al., 1998b). Because insect ACE has been shown to exert in vitro prohormone processing activities against the locustamyotropins, and locustamyotropin-containing cells of the locust brain and suboesophageal ganglion also contain ACE, this insect enzyme most probably plays a role in the biosynthesis of these hormones (Veelaert et al., 1999). 4. ACE and the Defense System In contrast to the situation in vertebrates, which have a dual system of immune reaction, namely, an innate and an acquired one, invertebrates only rely on their innate immune responses. This system comprises both humoral and cellular defense responses. Well-known humoral responses include the synthesis of a broad spectrum of antimicrobial peptides/proteins (Bulet et al., 1999), as well as two proteolytic cascades: the blood coagulation system identified in the horseshoe crab (Iwanaga, 1993, 2002) and the prophenoloxidase-activating system (pro-PO-AS) in insects and crustaceans (So¨derha¨ll and Cerenius, 1998). Cellular defenses cover hemocyte-mediated responses like phagocytosis, nodulation, and encapsulation. In insects, a large number of peptides is released into the hemolymph, where they are known to regulate a large diversity of physiological functions. The half-life of these peptides can be regulated by proteases and peptidases present in the hemolymph. The discovery of ACE activity in the hemolymph of insects (Isaac and Lamango, 1994), together with the knowledge of putative prohormone converting activity of ACE, indicated the possible involvement of ACE in the proteolytic processing of insect antibacterial peptides from their precursors (Isaac et al., 1998a). Supportive data concerning this hypothesis came from expression studies in larvae from the sheep blowfly Lucilia cuprina and the

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old screwworm Chrysomya bezziana, which are known to secrete or excrete chymotrypsins and trypsins in larval cultures and wound sites. The knowledge that an ACE homolog is also secreted or excreted from these larvae indicated a dual role in wound formation. It may degrade chymotryptic and tryptic digestion products from host tissue proteins and may influence the host vascular and inflammatory response by processing or degrading host regulatory peptides (WijVels et al., 1997). In the endoparasitic wasp, Pimpla hypochondriaca, the female envenomates and oviposits into pupae of some lepidopteran species. This venom was shown to suppress the hemocyte-mediated immune responses of the host by impairing encapsulation and phagocytosis (Parkinson et al., 2002; Richards and Parkinson, 2000). The presence of ACE in this venom, together with the presence of antibacterial activity, implies that ACE could be one of the enzymes involved in the processing of antibacterial peptides from the venom sac (Dani et al., 2003). All the mentioned data indeed point toward an involvement of ACE in the defense system, but they are still only hypothetical. The first physiological evidence for the existence of a connection between ACE and defense came from studies in the locust L. migratoria. Locusts that are subjected to an immune challenge through injection of lipopolysaccharides (LPS) into the hemocoel showed an increase in ACE mRNA-expression. Comparing the level of ACE transcription from LPS-treated animals with control animals revealed an approximately 10-fold increase in ACE transcription 6 hours after the LPS-injection (Macours et al., 2003b). The exact nature of the ACE-defense connection remains to be clarified. In mammals, all blood cells derive from hematopoietic stem cells that diVerentiate into diVerent lineages. The involvement of ACE here resides in the inactivation of Ac-SDKP (N-acetyl-seryl-aspartyl-lysyl-proline), a peptide that prevents the recruitment of hematopoietic stem cells (Azizi et al., 1996; Baudin, 2002). Insects also continue to produce hemocytes via a division of stem cells or by continued division of hemocytes in circulation (Jones, 1970; RatcliVe et al., 1985). On the basis of this information, a possible connection between ACE and the immune system can be suggested in the replenishment of the hemocytes that are involved in the insect immune response. The possibility of ACE being involved in the processing of antimicrobial peptides in locust hemolymph cannot be excluded, but so far, except for lysozyme and coagulogen, no major antibacterial activity could be detected in locust hemolymph (Van Sambeek and Weisner, 1999). The rise in ACE expression that follows LPS injection is not likely to result from an involvement of ACE in the prophenoloxidase cascade, as this pathway is not activated by injection of LPS. However, the immune system of the locust responds to LPS injection through the formation of nodules (Goldsworthy and Chandrakant, 2002; HoVman et al., 1974). Therefore ACE may be required in the process of nodule formation. This speculation is in agreement with the recent findings


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of Goldsworthy and colleagues (2003), who investigated the eVect of LPS on the prophenoloxidase cascade and nodule formation in the locust. Activation of prophenoloxidase by LPS could only be achieved by coinjection with AKH, not by the injection of LPS alone. When captopril was coinjected, the activation of the PPO cascade remained unaVected (with/without AKH). However, captopril did inhibit the nodule formation induced by LPS (Goldsworthy et al., 2003). Considering these data, a possible role for ACE in the immune system of insects may reside in the regulation of nodulation. E. ACE in Leeches To date, leeches are the only group of invertebrates in which the existence of a renin–angiotensin system has been reported. In Theromyzon tessulatum, a parasite of ducks, an angiotensin II (AII)-like peptide was isolated (Salzet et al., 1993). The full sequencing of such AII-like peptide was realized in another leech species, namely Erpobdella octoculata (Salzet et al., 1995). This leech AII peptide diVers from the vertebrate AII by a c-terminal amidation. Injection of this AII into the leech results in a decrease in mass, reflicting a diuretic eVect. Also in T. tessulatum, a renin-like peptide was isolated (Laurent and Salzet, 1995a) and partial peptidic sequences were identified. This enzyme was found capable of releasing AI from an AI-like precursor, Angiotensinogen (Laurent et al., 1995), by cleavage of the Leu10–Leu11 bond. The mass of this invertebrate renin (ca. 32 kDa) was lower than that of vertebrate active renin (ca. 44 kDa) but displayed a comparable activity. The leech renin is localized in the excretory and nervous system, which implies the involvement of the leech RAS in osmoregulation. The angiotensin-converting enzyme of this leech-RAS is a glycosylated membrane-bound enzyme of 120 kDa that is released into the hemolymph in a hydrophilic form (100 kDa) (Laurent and Salzet, 1996; Vandenbulcke et al., 1997). The presence of this RAS in coelomocytes of leeches and the involvement of each part of this system in the immune response inhibition indicates the implication of the RAS in host–parasite cross-talk (Salzet and Verger-Bocquet, 2001).

IV. Endothelin-Converting Enzyme: State of the Art in Mammals A. Introduction Endothelin-converting enzymes (ECEs) belong to the class of type II integral membrane zinc metalloproteases named for their ability to hydrolyze big endothelins (big ETs) into the smaller vasoactive endothelins. Yanagisawa

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and colleagues (1988) were the first to predict the existence of ECEs based on the cloning of the prepro-endothelin gene. ECE is classified into the neutral endopeptidase or M13 group of proteins, which contains type II membrane glycoproteins with zinc peptidase catalytic activity. At present mammalian M13 family of zinc proteases consists of seven known members: neutral endopeptidase (NEP); the endothelin-converting enzymes ECE-1, ECE-2, and ECE-3; the Kell blood group antigen (Kell); the phosphate regulating gene (PEX); X-converting enzyme (XCE); and secreted endopeptidase (SEP). Each member of this family shows homology to the others, mainly in the C-terminal part of the sequence that is responsible for catalytic activity. In addition, there are structural similarities in that all M13 members have short intracellular domains. Where identified, these enzymes have roles in the processing or metabolism of regulatory peptides and therefore represent potential therapeutic targets.

B. Structure and Isoforms ECE-1 is a membrane-bound protein with a short N-terminal cytoplasmatic tail, a transmembrane hydrophobic domain, and a large extracellular domain, which shares high sequence homology to NEP and other members of the metallopeptidase family. Ten cysteine residues in the ECE sequence are conserved among the other members of the family and are probably involved in disulfide bridges. The ECE protein has 10 predicted sites for N glycosylation in the extracellular domain. This is consistent with the observation that ECE is a highly glycosylated protein and explains the diVerence between the predicted molecular mass and the mass shown on SDS-PAGE. Purification and cloning studies of rat ECE-1 have shown that it consists as a disulphidelinked dimer, in which Cys412 is solely responsible for the formation of the intermolecular disulphide bond (Shimada et al., 1996). The large extracellular domain contains the zinc-binding consensus sequence HEXXH, in which the glutamic acid (E) is the most important residue for catalytic activity. The two flanking histidine (H) residues act as zinc coordinating amino acids. The third ligand for the zinc atom is E651, whereas H716 is involved in the stabilization of the tetrahedral intermediate during the transition state (Shimada et al., 1995). Four ECE-1 isoforms have been identified in humans and were named ECE-1a (758 amino acids), ECE-1b (770 amino acids), ECE-1c (754 amino acids) and ECE-1d (767 amino acids) (Jafri and Ergul, 2003; Valdenaire et al., 1995, 1999a). The four isoforms are transcribed from a single gene and result from the use of alternate promoters located upstream of specific exons. They only diVer in the first half (approximately) of their cytoplasmatic tail and are identical over their remaining


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sequence, including the enzymatic catalytic site. As a logical consequence, the isoforms display comparable converting activity. The specificity of the isoforms thus lies in their cytosolic tails, which results in a diVerent subcellular distribution. Whereas ECE-1a and ECE-1c are present in the plasma membrane, ECE-1b and ECE-1d are retained inside the cell. Recently it was shown that ECE-1 isoforms can heterodimerize. It was suggested that heterodimerization of ECE-1 isoforms could constitute a means for regulating their subcellular distribution and, subsequently, their extracellular activity (Muller et al., 2003). Considering the importance of the biosynthesis of both the intracellular and extracellular endothelins, the subcellular distribution of ECE isoforms may play a central role in the regulation of the endothelin system.

C. Biological Role The endothelin system is composed of endothelin (ET) peptides and their receptors (Fig. 3a), which together aVect a wide variety of physiological functions and pathological conditions. The discovery of this system started in the mid-1980s, when an increasing awareness of the role of endothelial cells as active components of the vascular system led to the description of an endothelial cell–derived constricting factor (Hickey et al., 1985). In 1988, a 21–amino acid vasoconstricting factor termed ‘‘endothelin’’ was isolated from cultured porcine aortic endothelial cells (Yanagisawa et al., 1988). The expression of ET is associated with many pathological processes and with tumor growth. To fulfill their wide spectrum of physiological functions, ETs act through autocrine and paracrine mechanisms. Their biosynthesis thus requires tight local control. Endothelin-1 (ET-1) is a 21–amino acid peptide with a hydrophobic C terminus. Within 1 year of its discovery, two structurally related peptides were identified and termed ET-2 and ET-3, respectively. The three isoforms are encoded by three diVerent genes and show highly conserved sequences in several species. All isoforms are composed of 21 amino acids with two intrachain disulfide bridges. ET-2 exhibits the closest structural similarity to ET-1, diVering by only three amino acid residues, whereas ET-3 diVers by six amino acids. ET isoforms are widely distributed among various cells and tissues. Of the three isoforms, the most widely distributed, best studied, and most potent is ET-1. ET-1 shares great structural homology to sarafotoxin, a snake (Atractapsis engaddensis) venom that induces myocardial infarction by exaggerated contraction of the cardiac vessels and interruption of the blood supply to the heart. ETs are produced by a variety of organ and body tissues such as the lung and kidney as well as the brain, pituitary, peripheral endocrine tissues, and

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FIG. 3 (a) The endothelin pathway. (b) Human big ET1–3. Cleavage by ECE leads to the generation of ET1–3 at the N-terminal and an inactive C-terminal fragment. Nonconserved amino acids are in grey boxes.

placenta. Most abundantly, endothelial cells of the vascular endothelium produce ET. ET binds to two types of receptors (ETA and ETB) (Arai et al., 1990; Sakurai et al., 1990). Both contain seven transmembrane domains and activate an overlapping set of G proteins, producing an array of diVerent physiological responses (Douglas and Ohlstein, 1997). Although structurally highly similar, these receptors possess diVerent aYnities for the three


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isoforms of ET. Like many other regulatory peptides, the ETs are maturated through proteolytic processing from larger precursor polypeptides, termed ‘‘prepro-ETs.’’ This proteolytic cleavage generates an essentially inactive intermediate referred to as big ET. The subsequent hydrolysis of big ET into the final active product ET, and a presumed inactive C-terminal fragment, is catalyzed by ECE (Fig. 3a). In the case of big ET-1 and ET-2, proteolysis occurs at the Trp21-Val22 bond. ET-3 is formed through cleavage of the Trp21-Ile22 bond of big ET-3 (Inoue et al., 1989) (Fig. 3b). The conversion of big ET to ET can occur in the extracellular medium and in the secretory pathway; cells thus secrete big ETs either alone or together with endothelins (Parnot et al., 1997). The physiological importance of the cleavage of big ET is indicated by a 140-fold increase in vasoconstrictor potency on cleavage to ET. This makes ECE an important activating protease in the biosynthetic pathway of ET. ECE-1 null mice exhibit a phenotype similar to that of ET-1- or ETA-deficient mice, demonstrating the significance of ECE-1 in generating bio-available ET-1 (Yanagisawa et al., 1998). The endothelin system plays a variety of roles in both normal physiology and pathological conditions in a number of tissues/organs of the body. In blood vessels, ET maintains a basal level of vasoconstriction and is involved in the development of hypertension, atherosclerosis, and vasospasm after subarachnoid haemorrhage. In the heart, the endothelin system aVects force and rate of contraction and mediates hypertrophy and remodeling in congestive heart failure. In lungs, ET regulates the tone of both airways and blood vessels and is involved in the development of pulmonary hypertension. Kidney-ET regulates water and sodium excretion and acid–base balance, and it participates in the pathophysiology of acute and chronic renal failure. In the brain, the ET system modulates cardiorespiratory centers and hormone release (Kedzierski and Yanagisawa, 2001). In ovaries, ET is shown to play a role in regulating the female reproductive cycle, where it functions as an important component of the luteolytic cascade (Levy et al., 2003). Egidy and colleagues (2000) suggested that ET, as a powerful vasoconstrictor and mitogenic peptide that is produced by many cell lines, might play a role in cancer progression of the colon. Studies by Johnson and colleagues (1999) revealed that ECE-1 possesses a broad in vitro substrate specificity and is potentially involved in the metabolism of peptides distinct from ET. They found that neurotensin, substance P, bradykinin, and the oxidized insulin b chain are in vitro substrates for ECE-1. Given this in vitro substrate specificity, it is likely that ECE-1 is involved in the degradation and processing of many biologically active peptides, both at the cell surface and in the secretory pathway (Johnson et al., 1999).

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D. ECE-2 and ECE-3 A second form of ECE, ECE-2, was cloned from bovine adrenal cortex and was reported to possess a 59% identity with ECE-1. The main structural features of ECE-1 are conserved in ECE-2 (Emoto and Yanagisawa 1995). Both are type II integral membrane proteins with the HEXXH consensus sequence. The 10 cysteine residues conserved between ECE-1, NEP, and Kell, are also conserved in ECE-2, as is the cysteine residue responsible for dimerization of ECE-1. ECE-2 is also heavily glycosylated, with 10 possible glycosylation signals. Although ECE-2 shows specific activity to produce mature ET-1 from big ET-1, like ECE-1, the striking diVerence between ECE-2 and ECE-1 is the acidic pH optimum of 5.5 of ECE-2, in contrast to the neutral pH optimum of ECE-1. This indicates that ECE-2 is involved in ET synthesis in the intracellular compartments in which the pH is acidic (Emoto and Yanagisawa, 1995). ECE-2 has been reported to be expressed in various organs and tissues including cultured human vascular endothelial cells, uterus, ovary, heart, lung, and liver, indicating various functions for the enzyme. Recent targeting studies with mouse ECE-2 revealed that it plays crucial roles in the formation of cardiac outflow structures (Yanagisawa et al., 2000). Four subisoforms of ECE-2 that diVer only in their N-terminal cytoplasmatic tails were termed ECE-2a-1, ECE-2a-2, ECE-2b-1 and ECE-2b-2, ECE-2a-1 and ECE-2a-2 are expressed in a variety of tissues including liver, kidney, adrenal gland, testis, and endothelial cells. ECE-2b-1 and ECE-2b-2 are abundantly expressed in the brain and adrenal gland (Ikeda et al., 2002). ECE-3 was purified from bovine microsomes. This enzyme was found to be specific for the conversion of big ET-3 to ET-3 and was inhibited by phosphoramidon (Hasegawa et al., 1998). To the best of our knowledge, this is the only report on an ECE-3 enzyme.

E. Other Members of the M13 Metallopeptidase Family 1. Neutral Endopeptidase Neutral endopeptidase (neprilysin, NEP), a 90–110-kDa plasma membrane protein, is the prototype and best characterized member of the M13 zinc metallopeptidase family. NEP is identical to the neutrophil and clusterdiVerentiation antigen CD10 and is also known as the common acute lymphoblastic leukemia antigen (CALLA), which is mainly associated with the precursors of B lymphocytes (Letarte et al., 1988). Purification and subsequent cloning of NEP revealed it to be a type II integral membrane protein of approximately 700 residues. Neprylisin’s extracellular domain,


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which includes the active site, also contains 12 cysteine residues, 10 of which are involved in highly conserved disulfide bridges that are important in the maintenance of structure and activity of the enzyme (Oefner et al., 2000). NEP comprises three subisoforms that result from alternative splicing in the 50 untranslated region, indicating that these spliceforms do not exhibit functional diVerences (Ishimaru and Shipp, 1995). Regarding its catalytic activity, NEP shares some similarity with the bacterial zinc metalloproteinase thermolysin, and in particular in its substrate specificity (Malfroy et al., 1988). NEP exists as an ectoenzyme that preferentially hydrolyzes extracellular oligopeptides ( human hybrids faithfully localized within the mouse nucleoli (an important point considered in more detail later). Mouse UBF associated with the human rDNA, although the human genes remained silent, but the observation indicates that UBF is necessary but not suYcient for transcription. The authors concluded that species diVerences in the critical

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TIF-IB/SL1 factor leads to silencing of the human rDNA in mouse > human cell hybrids. Taken together, the ChIP assays and the cell hybrid assays support the hypothesis that UBF plays a structural role in nucleolar chromatin within interphase FCs and mitotic NORs. This structural role of prepositioned UBF is concomitant with its functional role in recruiting TIF-IB/SL1 (Reeder et al., 1995). Topoisomerase I is another key regulator in rDNA transcription. Most topoisomerase I in the interphase nucleus enriches in nucleoli (Fleischmann et al., 1984; Muller et al., 1985), but it readily exchanges with the nucleoplasm (Christensen et al., 2002a). Rose et al. (1988) showed that topoisomerase I activity associates tightly with Pol I purified from rat hepatoma cells. In their study, antibodies directed against the topoisomerase blocked the relaxation of supercoiled DNA in vitro. Several reports described the requirement of topoisomerase I (Topo I) activity for Pol I transcription (Brill et al., 1987; Garg et al., 1987; Zhang et al., 1988). Previous studies showed the presence of Topo I in mitotic NORs (Guldner et al., 1986; Rose et al., 1988). Christensen et al. (2002b) showed that the nonconserved amino terminus of topoisomerase I functions as an adaptor to anchor a subfraction of the protein to the FCs and the mitotic NORs. Weisenberger et al. (1993) showed that the topoisomerase I inhibitor, camptothecin, blocks postmitotic nucleologenesis in mammalian PtK2 cells. The author’s results are further described below. Antibody localizations clearly show enrichments of Pol I, UBF, and Topo I within the FCs. The fraction of Pol I actually engaged in rRNA transcription may be small; most Pol I is thought to reside in preinitiation complexes (Grummt, 1999) within the FC. Whether the majority of transcription factors within the FCs actively engages in rDNA transcription, or instead assembles into quiescent preinitiation complexes remains a debatable issue. Some argue that transcription occurs within the FCs, and others support the model in which rDNA loops out from the FC into the DFCs, where it transcribes pre-rRNA (Hoza´k et al., 1994; Raska et al., 1995; Schwarzacher and Wachtler, 1993; Thiry and Goessens, 1996; Wachtler et al., 1989). Taking the middle ground, we cite more recent studies that indicate that FCs are repositories for silent rDNA with bound but quiescent transcription complexes. The active rDNA genes loop out toward the FC cortex and perhaps on into the DFC, where early pre-rRNA processing occurs (Biggiogera et al., 2001; Cheutin et al., 2002; Cmarko et al., 2000; Huang, 2002; Koberna et al., 2002; Malıńsky et al., 2002). 3. The Dense Fibrillar Component: Pre-rRNA Processing and Early Ribosome Assembly The DFC can assume various morphologies that reflect the intensity of ribosome biosynthesis. For example, the DFC forms a reticulated nucleolonema in cells producing ribosomes at maximal rates. In most cells, the DFC

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forms a more compact concentric sphere around the FC in cells in which ribosome synthesis is less than maximal. DFCs appear dark in electron micrographs in part because of the aYnity between metal stains and resident DFC components, and in part because of the high concentration of these components. The DFCs also appear dark by phase-contrast microscopy. Although the DFC may support (arguably) pre-rRNA transcription, quick pulse-labeling experiments have clearly established the DFC as the site of initial pre-rRNA processing and early ribosome assembly. Ribosomal RNA is synthesized as a precursor of 47S in mammals, 38S in Drosophila, and 35S in yeast. The nascent rRNA is about 7000 nucleotides long. The linear organization of these precursors is well conserved despite the evolutionary divergence of the species. From its 50 end, the pre-rRNA consists of an external transcribed spacer (ETS), the 18S rRNA, internal transcribed spacer 1 (ITS1), the 5.8S rRNA, ITS2, the 28S rRNA, and then a short 30 tail. Processing of the pre-rRNA includes endonucleolytic cleavage, chemical modifications, RNA folding, and the association of ribosomal proteins. Processing begins cotranscriptionally. The bulk of ribosomal subunit assembly is thought to occur in the DFC, continue in the GC, and linger to a certain extent while the large and small ribosomal subunits are in transit through the nucleoplasm to the nuclear envelope. We know that the final processing steps actually occur to the cytoplasm. Yeast has thus far provided the most detailed information on subunit assembly (reviewed by Tschochner and Hurt, 2003). The yeast 35S prerRNA associates with nonribosomal processing factors, ribosomal proteins, and the 5S rRNA to form an initial 90S preribosomal particle (Fig. 1 of Tschochner and Hurt, 2003). Chemical modifications within the 90S particle include 20 -O-methylation of the ribose moieties of specific nucleotides, nitrogenous base methylations, and site-specific uridine to pseudouridine conversions (pseudouridylation). These chemical modifications occur within pre-rRNA regions that will eventually generate mature 18S, 5.8S, and 25S RNAs found in intact ribosomes. The pre-90S particle bifurcates into a pre40S particle (small ribosomal subunit) and a pre-60S particle (the large ribosomal subunit) that then follow their respective processing and assembly pathways. Endonucleolytic cleavage and exonucleolytic trimming within the pre-90S particle remove external and internal spacer sequences, thus liberating a 20S intermediate rRNA. This yields the 18S mature rRNA within the 40S pre-ribosomal subunit. Concomitantly, the 5S rRNA synthesized outside the nucleolus joins the 5.8S and 25S rRNAs to form the 60S preribosomal subunit. Large and small subunit assembly occurs as ribosomal proteins enter the nucleolus from the cytoplasm and associate with the maturing rRNAs. Our future research challenge is to decipher the order of these processing and assembly events (Tschochner and Hurt, 2003).


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Relatively few snoRNAs (U3, U8, U22, and MRP in vertebrates, and U3, U14, MRP, and snR30 in yeast) play associative roles in the multiple cleavage events of pre-rRNA (Gerbi, 1995; Maxwell and Fournier, 1995; Yu et al., 1999). These various snoRNAs exist as small nucleolar ribonucleoprotein particles (snoRNPs). U3 is the most abundant snoRNA. It is well conserved from yeast to mammals, and it is a founding member of the box C/D family of snoRNAs (see below). The U3 snoRNP makes the first cleavages within the pre-rRNA, beginning at A0 in the 50 ETS (Borovjagin and Gerbi, 1999; Mougey et al., 1993a,b) to liberate the 18S region. In Saccharomyces cerevisiae, U3 associates with 28 proteins ranging in size from 13 to 200 kDa to form a processome of 2.2 MDa (Dragon et al., 2002; Grandi et al., 2002) that initiates assembly of the small ribosomal subunit. The U8 snoRNP, however liberates the 5.8S and 28S regions (Peculis and Steitz, 1993, 1994). Specificity in cleavage is determined by base pairing between the nascent rRNA and the specific snoRNAs (Hughes, 1996; Sharma and Tollervey, 1999). Far more snoRNAs (200) participate in chemically modifying the prerRNA (reviewed by Bachellerie et al., 2002; Kiss, 2002; Tollervey and Kiss, 1997; Yu et al., 1999). These latter snoRNAs fall into two families. The first is the box C/D (short conserved sequences) snoRNAs that serve as guides for site-specific methylation of the pre-rRNA. There are 55 20 -O-methylated sites in S. cerevisiae rRNA, 86 sites in A. thaliana, and 105–107 sites in mammalian rRNA. Specific base pairing between the C/D box guide RNAs and the pre-rRNA determines the site of methylation. Proteins associated with the box C/D snoRNAs include fibrillarin (Nop1p in yeast), Nop56p, Nop58p, and the 15.5-kDa protein (also known as Snu13p in yeast). Fibrillarin (34 kDa) is the putative methyltransferase (Tollervey et al., 1993; Wang et al., 2000); it resides exclusively within the DFC. Because cleavage and methylation are considered early pre-RNA processing events, the most centric localization of fibrillarin within the standard ring nucleolus should indicate the earliest accumulations of nascent pre-rRNA. In fact, several labs have shown enrichments of fibrillarin at the FC-DFC borders (Shah et al., 1996) or to distinct foci within the DFC (Beven et al., 1996; Mosgoeller et al., 1998), thus indicating that the DFC is functionally subcompartmentalized. The second family of snoRNAs shares a conserved hinge sequence and an ACA motif three nucleotides from their 30 end. These box H/ACA snoRNAs serve as guides for site-specific pseudouridylation (Ganot et al., 1997; Ni et al., 1997). There are 44 pseudouridines in yeast rRNA and 91–93 in mammalian rRNA. Again, base pairing between the guide RNA and prerRNA determines the actual site of pseudouridylation (Bachellerie et al., 2002). Proteins associated with the box H/ACA snoRNAs are Dyskerin (Cbf5p in yeast), Gar1p, Nop10p, and Nhp2p. Dyskerin is the putative pseudouridylase (Hoang and Ferre´-D’Amare´, 2001; Lafontaine et al., 1998;

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Zebarjadian et al., 1999). The sites for methylation and pseudouridylation are conserved in the mature rRNAs, and interestingly, these modifications lie within or near the functional domains of the ribosome (e.g., the peptidyl transferase domain of the large ribosomal subunit). Although the precise role for these numerous methylations and pseudouridylations remain elusive, they are thought to help fold the rRNA and allow for proper interactions with ribosomal proteins (Bachellerie et al., 2002). Interestingly, the 15.5-kDa/ Snu13p protein of the box C/D snoRNPs associates with the pre-mRNA spliceosomal U4/U6-U5 snRNPs (Peculis, 2000; Watkins et al., 2000), and 15.5-kDa/Snu13p contains a central region that is conserved in Nhp2p. Both 15.5-kDa/Snu13p and Nhp2p share similarities with ribosomal protein L30. The association of 15.5-kDa/Snu13p with snoRNPs and spliceosomal RNPs and its sequence similarities with Nhp2p and L30 indicate common ancestral origins for these RNP proteins (Watkins et al., 2000; reviewed by Peculis, 2000). In addition to the few snoRNP proteins just mentioned, an estimated 350 proteins participate in ribosome assembly (Andersen et al., 2002; Leung et al., 2003; Scherl et al., 2002). These proteins can be functionally divided into rDNA helicases and topoisomerases, endo- and exoribonucleases, RNA helicases, molecular chaperones, intranuclear transport proteins, and putative GTPases that are involved in nuclear export (for detailed reviews on function, see Olson et al., 2002; Tschochner and Hurt, 2003). A few of these proteins are relatively abundant within nucleoli and they likely play important roles in nucleolar assembly and disassembly. The most abundant and intensely studied of the vertebrate proteins include nucleolin, B23, and Nopp140. Nucleolin (110 kDa) interacts with the nascent pre-rRNA primarily within the DFC, but also somewhat in the GC (Ghisolfi-Nieto et al., 1996; Herrera and Olson, 1986) to perhaps establish proper RNA folding for rRNA processing events and ribosome assembly (Bouvet et al., 1998; Ginisty et al., 1998, 1999; Olson, 1990). Nucleolin is a modular protein (Lapeyre et al., 1987); its amino terminal third contains alternating acidic and basic domains, its central domain contains four RNA-binding domains, and its carboxy terminus is rich in glycine and dimethylarginine residues that form several Arg-Gly-Gly (RGG) motifs that are common to many other RNA-binding proteins (Burd and Dreyfuss, 1994; Draper, 1995). Nucleolin’s amino terminal basic domains have several Cdk1/cyclin B phosphorylation sites. As such, it is tempting to speculate that the phosphorylation of an abundant nucleolar protein during prophase may help disassemble the nucleolus, and its dephosphorylation may drive nucleolar reassembly during telophase (see below). Olson et al. (2002) provided a recent and thorough review on nucleolar protein B23 (38 kDa). In brief, B23 resides in the DFC and GC and is


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thought to interact with nucleolin (Li et al., 1996; Pellar and DiMario, 2003; Pinõl-Roma, 1999), nucleolar p120 (Valdez et al., 1994), and the HIV Rev protein (Fankhauser et al., 1991; Szebeni et al., 1997). B23 displays nucleic acid binding activity, nuclease activity, in vitro chaperone activity (Olson et al., 2002; Szebeni and Olson, 1999; Szebeni et al., 2003), and possible centrosome functions (Zatsepina et al., 1999). B23 likely acts as a chaperone in the protein-rich nucleolus. In its dephosphorylated form, B23 binds other nucleolar proteins or ribosomal proteins to prevent their erroneous aggregation and to ensure their proper assembly into ribosomal subunits. On phosphorylation by a nucleolar CKII, B23 releases its protein substrate (Szebeni et al., 2003). Like nucleolin (Lischwe et al., 1982, 1985a), fibrillarin contains RGG motifs (Christensen and Fuxa, 1988; Lischwe et al., 1985b), thus implicating its interactions with RNA (Ghisolfi et al., 1992a,b), perhaps in a methylation-dependent manner (Henry and Silver, 1996; Liu and Dreyfuss, 1995; Najbauer et al., 1993; Pellar and DiMario, 2003; Xu et al., 2003). Nopp140 (140–180 kDa in vertebrates) resides in the DFCs. Except for its amino and carboxy tails, vertebrate Nopp140 consists of alternating acidic and basic domains (Cairns and McStay, 1995; Meier and Blobel, 1992). The acidic domains are rich in serine residues that are phosphorylated by a nucleolar casein kinase type II during interphase (Meier, 1996), whereas the basic domains contain a few Cdk1/cyclin B phosphorylation sites. Rat Nopp140 shuttles between the nucleus and the cytoplasm (Meier and Blobel, 1992), as do nucleolin and B23. The physiological significance of nucleolar protein shuttling still remains uncertain. In vitro, Nopp140 binds nuclear localization signals (NLSs) in other nucleophilic proteins when its many serine residues are phosphorylated by CKII (Meier and Blobel, 1990). This is consistent with the hypothesis that Nopp140 and other nucleolar shuttling proteins serve as chaperones in the transport of ribosomal proteins into the nucleus, or perhaps ribosomal subunits out of the nucleus. Nopp140 interacts with NAP57 (Meier and Blobel, 1994), one of the four basal proteins subunits of snoRNPs required for pseudouridylation, and NAP65, a constituent protein of box C/D snoRNPs (Yang et al., 2000). Nopp140 interacts with both classes of snoRNPs in a phosphorylation-dependent manner (Wang et al., 2002) to perhaps act as a chaperone in snoRNP assembly or transport. Interestingly, Nopp140 is not required for the RNA-modifying activities for either of these two classes of snoRNPs (Wang et al., 2002). Although Nopp140 is not known to bind nucleic acids, Nopp140 may participate indirectly in pre-rRNA processing and modification by serving as a scaVold for processing events. Isoforms of Nopp140 have been described in humans (Pai and Yeh, 1996), in Drosophila (Waggener and DiMario, 2002), and in Chironomus (Sun et al., 2002). In the latter study, the authors treated cells with ribonuclease that normally redistributes nucleolar

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proteins such as nucleolin to the nucleoplasm, but showed that the Chironomus Nopp140-like p100 remains in place. The result indicates that Nopp140like proteins may serve as a structural framework for nucleolar activity. If so, their dynamic interactions in the cell cycle may regulate nucleolar assembly and disassembly. Mostly because of their high relative abundance, studies of nucleolar assembly and disassembly use these four proteins as cytological markers. Many other less abundant nucleolar proteins exist (Andersen et al., 2002; Olson et al., 2002; Tschochner and Hurt, 2003), and their localizations during nucleolar assembly and disassembly are forthcoming or have yet to be determined. 4. The Granular Component and Nuclear Export The granular component is the most peripheral zone of the nucleolus. Ribosomal subunit assembly continues in this compartment as the 40S and 60S particles prepare to make their transit from the DFC toward the nuclear envelope. No fibrillarin resides in the GCs, and smaller amounts of nucleolin reside in the GCs than in the DFCs. Conversely, greater amounts of B23 reside in the GCs (Olson et al., 2002; Spector et al., 1984). The supposition is that rRNA processing and subunit assembly are in their final stages before subunit release and export. Subunit release from the GC may depend on the final removal of processing machinery and restructuring of the subunits. Various chaperones, helicases, ATPases (Rix7p) and GTPases (Nog1p, Nog2p/Nug2p, and Nug1p), and other accessory proteins are implicated in these final events, some of which would presumably occur within the GCs (Johnson et al., 2002). In yeast, subunit maturation and intranuclear transport requires the Noc protein complexes: Noc1p-Noc2p associates with the pre-90S particle and the pre-60S subunits within nucleoli, and the Noc2pNoc3p complex is thought to play a role in transporting the pre-60S subunit through the nucleoplasm to the nuclear envelope. In contrast, the Noc4Nop14p complex associates with the pre-90S particle and the 40S subunit at least up to the nuclear envelope (Milkereit et al., 2001, 2003). Transport of the large 60S subunit across the nuclear envelope was reviewed by Johnson et al. (2002). Briefly, the 60S and 40S subunit export pathways in yeast and Xenopus are independent of each other, but both are dependent on CRM1, which serves as receptor for cargos bearing nuclear export signals (NESs). CRM1 links export cargos to the RAN-GTP export process. In addition to CRM1, Nmd3p is necessary for 60S transport out of the nucleus. Nmd3p contains both a NES and a NLS for nuclear import; thus, Nmd3p shuttles between the nucleus and the cytoplasm. Nmd3p is believed to be one of the last proteins to bind the 60S subunit in the nucleus, perhaps by binding ribosomal protein Rp110p. Thus, Nmd3p is the


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NES-containing adaptor that links the 60S subunit to CRM1 and the export machinery. Factors involved with 40S export (Kre33p and Krr1p) have been suggested (Grandi et al., 2002), but the export mechanism for the 40S subunit is less established than that for the 60S subunit.

B. Nucleolar Multitasking Novel nucleolar functions have been described in the last several years (Cockell and Gasser, 1999; Olson et al., 2002; Pederson, 1998a, 1998b; Pederson and Politz, 2000; Visintin and Amon, 2000). In yeast and metazoans, these functions include the processing of small RNAs not usually associated with nucleoli or ribosomes. The functional characterization of box C/D snoRNAs (methylation) and box H/ACA snoRNAs (pseudouridylation) represented a major breakthrough in our understanding of nucleolar function, not only with regard to chemical modifications of ribosomal RNA but also in the modifications of nonribosomal RNAs that transit through the nucleolus. For instance, chemical modification of vertebrate spliceosomal snRNAs U2 (Yu et al., 2001), U4, U5, and U6 (Gerbi and Lange, 2002; Gerbi et al., 2003; Lange and Gerbi, 2000) occurs in the nucleolus. In addition to the spliceosomal snRNAs passing through the nucleolus for modification, other small RNAs transit through the nucleolus either for chemical modification or for partial assembly into their respective RNP complexes. The telomerase RNA contains a box H/ACA domain at its 30 end, which targets this RNA to nucleoli (Mitchell et al., 1999; Narayanan et al., 1999). The RNase P RNA is another small RNA that transits the nucleolus. RNase P normally cleaves the 50 leader sequence of pre-tRNAs, and its greatest steady state concentration is dispersed throughout the nucleus. However, rhodamine-tagged RNase P RNA, when injected into rat kidney cell nuclei, rapidly accumulates within the DFC of nucleoli and localizes coincidently with endogenous fibrillarin before dispersing to the rest of the nucleus (Jacobson et al., 1997). RNase P proteins, Rpp14 and Rpp29, localize to the DFC, and Rpp38 is dispersed throughout the nucleolus. Both Rpp29 and Rpp38 also localize to Cajal bodies (Jarrous et al., 1999). Transfer RNAs also transit through the nucleolus (Bertrand et al., 1998; Pederson and Politz, 2000). The SRP RNA along with three out of four proteins normally associated with the intact cytoplasmic SRP transit through the nucleolus (Jacobson and Pederson, 1998; Politz et al., 1998, 2000). Specifically, the Alu domain and internal helix 8 (within the S domain) of the SRP RNA are required for its nucleolar localization. SRP proteins 19, 68, and 72, but not 54, also transit through the nucleolus, indicating that partial SRP assembly occurs within the nucleolus. Interestingly, the SRP RNA localizes to subregions of

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the GCs that do not contain significant amounts of 28S rRNA (Politz et al., 2002). This observation indicates that certain territories within the nucleolus are reserved for SRP assembly rather than ribosome assembly. The transit of SRP components through the nucleolus indicates a possible coordination with ribosome assembly (Pederson and Politz, 2000; Politz et al., 2002). The nucleolus plays a fundamental role in budding yeast cell cycle progression, and this is mediated by a complex of proteins known as RENT (for regulator of nucleolar silencing and telophase exit) (Cockell and Gasser, 1999). Yeast proteins Net1, Cdc14, Sir2, and Nan1 constitute the complex. Net1 binds rDNA throughout the cell cycle, promotes rDNA expression during interphase, and is required for nucleolar integrity (Straight et al., 1999). Cdc14 is a cell cycle regulatory phosphatase, and Sir2 (silent information regulator) is a NADþ-dependent chromatin histone deacetylase. Nan1 is the least characterized of the four. The RENT complex is intact from G1 through late anaphase. Its association with rDNA is dependent on interactions with nucleolar protein, Fob1, and subunits of Pol I (Huang and Moazed, 2003). In late anaphase, Cdc14 is released from its negative regulator, Net1 (also known as Cfi1, Visintin et al., 1999), that has tethered Cdc14 within the nucleolus (Traverso et al., 2001). Once released, Cdc14 diVuses through the nucleus and cytoplasm in a Tem1-dependent fashion (Shou et al., 1999) to activate Cdh1/Hct1, a regulator of the anaphase-promoting complex/cyclosome, which in turn inactivates mitotic cyclins Clb2 and Clb3. The released Cdc14 also promotes the accumulation of Sic1, the inhibitor of Clb/ Cdc28. In a positive feedback loop, Cdc14 also activates Swi5, the transcription factor that promotes SIC1 gene expression. All three Cdc14 activities are necessary to exit mitosis in budding yeast. Finally, dephosphorylation of nucleolar Net1 by Cdc14 may restore Cdc14 to the nucleolus as the cell proceeds into interphase (Shou et al., 1999). The RENT complex in budding yeast also functions in rDNA silencing; specifically, Sir2 represses rDNA expression (Bryk et al., 1997; Smith and Boeke, 1997) and recombination (Gottlieb and Esposito, 1989; Sinclair and Guarente, 1997). In the study by Smith and Boeke (1997), the Ty1 retrotransposon marked with the URA3 gene integrated eYciently into the rDNA, but expression of the ectopic URA3 gene was silenced in a Sir2-dependent manner, and loss of Sir2 increased psoralen accessibility to the rDNA as measured by DNA cross linking. Loss of Sir2 also causes a 10-fold increase in the rate of recombination of rDNA as measured by the accumulation of extra-chromosomal rDNA circles that accumulate in the yeast mother cell because of aberrant recombination within the rDNA locus. Accumulation of these circles is toxic and is believed to trigger senescence (Guarente, 2000; Sinclair and Guarente, 1997). Although novel nucleolar roles in small, nonribosomal RNA processing is well established and still expanding for yeasts


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and metazoans (Gerbi et al., 2003), the possible roles that Sir2-related proteins and the nucleolus may have in metazoan cell aging remain a forefront for investigation. Thus far, we have described the intricate and dynamic choreographies necessary to assemble ribosomal subunits within the interphase nucleolus (its traditional function), and we have briefly surveyed novel nucleolar functions. Determining the precise interactions of nucleolar macromolecules is our current and future challenge. Knowledge of how these interactions are regulated will explain nucleologenesis, nucleolar maintenance during interphase, and eventually nucleolar disassembly in mitosis.

IV. Nucleologenesis is Concurrent with the Reinitiation of NOR Transcription Although rRNA transcription ceases in prophase, the transcription factors UBF, TIF-IB/SL1, RNA Pol I, TTF-1, and DNA topoisomerase I remain bound to the mitotic NORs. Reinitiation of rDNA transcription in late anaphase/early telophase is dependent on the (re)activation of the transcription complexes. Transcription reinitiation is in general responsible for the formation of intact nucleoli in the subsequent interphase. The question addressed in this first section is what factors are involved with the reinitiation of Pol I transcription? In part B we ask whether or not transcription is necessary for PNB formation, and in part C we look at the details of nucleolar reassembly beginning in telophase.

A. Reinitiation of rDNA Transcription in Late Anaphase/Early Telophase Bell et al. (1997) added demembranated Xenopus sperm nuclei to activated egg extracts to assemble male pronuclei. Antibodies directed against Xenopus UBF clearly labeled either a single focus (there is only one NOR in Xenopus on chromosome 12) or a series of small necklace-like beads that the authors interpret to be individual rDNA repeats within the single NOR. UBF associated with the rDNA within the paternal NOR was maternally inherited from the egg cytoplasm; no UBF was packaged in the sperm. Interestingly, although UBF is present on the NORs from the pronuclear stage and onward throughout embryogenesis, no TIF-IB/SL1 or Pol I could be detected until the midblastula stage, when rDNA transcription initiates. This observation supports the view that UBF acts as a structural chromatin

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protein that associates with the rDNA throughout the cell cycle regardless of transcriptional activity, and that it is UBF that recruits the other Pol I factors to the rDNA promoters. Formation of intact, active nucleoli after mitosis appears to be a function of rDNA transcription. Benavente et al. (1987) injected antibodies directed against Pol I into prometaphase and metaphase PtK2 cells and followed the injected cells as they divided into postmitotic daughter cells. Anti-Pol I clearly blocked the reformation of typical nucleoli in the daughter cells, perhaps by redistributing Pol I from the NORs/fibrillar centers and thus blocking rDNA transcription. Despite the lack of intact nucleoli, numerous prenucleolar bodies containing fibrillarin were readily apparent by immunofluorescence microscopy. Transmission electron microscopy showed that these PNBs consisted of densely packed fibrillar material typical of DFCs and that, occasionally, a PNB could be found associated with putative fibrillar centers that were identified by their morphology and heavy metal staining characteristics. As described earlier, topoisomerase I is implicated in the reactivation of NOR transcription. Weisenberger et al. (1993) used the Topo I inhibitor, camptothecin, to block reinitiation of NOR transcription in telophase, and thus the reformation of nucleoli. Metaphase cells treated with the drug completed mitosis, but their daughter cells lacked nucleoli in the next interphase. Antibodies directed against fibrillarin again stained prenucleolar bodies in the interphase nucleoplasm of the camptothecin-treated cells, whereas antibodies against Topo I and RNA Pol I continued to label inactive NORs. The observations again support the requirement for reactivating rDNA transcription to initiate reformation of intact, functional nucleoli. Instead of blocking rDNA transcription, Sirri et al. (2000) confirmed the hypothesis by prematurely reinitiating rDNA transcription in metaphase and anaphase HeLa cells. Maintaining transcription repression during mitosis is a function of Cdk1-cyclin B kinase. Sirri et al. (2000), therefore, used the drug roscovitin, a powerful (IC50 < 7 mM) inhibitor of Cdk1-cyclin/B kinase, to block continued phosphorylation of Pol I transcription factors. Transcription factors that are likely substrates for Cdk1-cyclin B are TBP (its phosphorylation would down-regulate all transcription in the nucleus), TAF1110, and TTF-1. Their phosphorylation during prophase is thought to shut down rDNA transcription (see later discussion). Roscovitin inhibition relieves rDNA transcription repression, but in a protein phosphatase 1 (PP1)-dependent manner. On blocking Cdk1-cyclin B kinase, dephosphorylation of the transcription factors by PP1 would reinitiate their activity, and thus their rDNA transcription. Sirri et al. (2000) observed transcription reinitiation only at the NORs and not at other loci, most likely because the NORs maintain a preinitiated Pol I transcription machinery (i.e., UBF, TIF-IB/SL1, and Pol I) during mitosis. Okadaic acid, a powerful PPI


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inhibitor, blocked transcription reinitiation in the presence of roscovitin. Interestingly, the reinitiation of rDNA transcription in metaphase and anaphase produced the expected 47S pre-rRNA, but the precursor RNA was not processed to mature 18S, 5.8S, or 28S rRNAs. Apparently, the roscovitintreated mitotic cells still lacked the appropriate mechanism(s) to reassemble the preexisting processing machinery back into functional nucleoli that can process pre-rRNA.

B. Formation of Nucleolus-Like Bodies in the Absence of rRNA Synthesis The anucleolate (0-nu) mutation in Xenopus laevis was originally described by Elsdale et al. (1958) and was characterized in molecular terms by Brown and Gurdon (1964), Wallace and Birnstiel (1966), Brown (1967), and Brown and Weber (1968). Esper and Barr (1964) and Hay and Gurdon (1967) described the prenucleolar bodies and pseudonucleoli that form in anucleolate embryos. Prenucleolar bodies form in the 0-nu mutants in the early cleavage stages of embryogenesis, just as they do in heterozygous embryos that contain one functional NOR (1-nu) or in homozygous wild-type embryos that contain the normal number of two NORs (2-nu). No diVerences in fine structure were noted between the PNBs in the three genotypes during the early cleavage stages and up to gastrulation. Later in development, however, definitive nucleoli with DFCs and GCs formed in the 1- and 2-nu embryos as PNBs fused, and PNBs persisted in the 0-nu embryos. By the tail bud stage, many cell types within 0-nu embryos (e.g., presumptive myoblasts) managed to form pseudonucleoli that segregated a large fibrillar region from a smaller granular (capped) region. Without a normal complement of ribosomal genes, fibrillarin and its associated U3, U15, and U17 remained diVuse and localized within a certain class of PNBs. Nucleolin was also diVuse within the nuclei, and it localized to a second class of PNBs (Crosio et al., 1997). A few normal and abnormal rDNA genes may still remain in the 0-nu genome (Steele et al., 1984; Tashiro et al., 1986), and these few genes could generate suYcient rRNA to account for the small pseudonucleoli. Any ribosomes that may be produced, however, are insuYcient to sustain the organism; 0-nu embryos survive only until the swimming tadpole stage because of large pools of maternal ribosomes produced by the oocyte before the meiotic divisions. NORs exist on both the X and Y chromosomes in Drosophila melanogaster, and deletions of rDNA in D. melanogaster are referred to as bobbed mutations (Ritossa, 1976). According to Ritossa, the first bobbed mutation was discovered by Bridges (Morgan et al., 1925), and although many diVerent bobbed alleles have been described since then, they have been little used in characterizing embryonic nucleologenesis in Drosophila (see later discussion).

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Instead of genetic rDNA deletions, other approaches have used colcemid (Philips and Philips, 1979) or colchicine (Hernandez-Verdun et al., 1979, 1991) to disrupt the mitotic spindle and thus to perturb normal segregation of the replicated chromosomes. In the following interphase, micronuclei arise that contain aneuploid (reduced) complements of chromosomes. Micronuclei could contain one or a few chromosomes. The technique eVectively separates chromosomes that either contain or lack NORs. Phillips and Phillips (1979) used newly initiated X. laevis cell cultures to generate micronuclei. Because X. laevis has only two NOR-bearing (homologous) chromosomes, most micronuclei lacked NOR chromosomes. Micronuclei containing the NOR chromosomes (confirmed by in situ hybridization) were easily scored by the formation of large, fully functional nucleoli with typical ultrastructural morphologies. Conversely, micronuclei lacking NOR chromosomes formed nucleolus-like bodies that were smaller than true nucleoli, appeared electron-dense at the ultrastructural level, and consisted of a fibrous material (DFC), but with a granular cap—much like that produced by Act D treatment. FCs were not apparent. Phillips and Phillips (1979) noted that the nucleolus-like bodies contained RNA. We now know that this is partially processed pre-rRNA, presumably from the previous interphase (Dundr and Olson, 1998; see later discussion). The Xenopus micronuclei that contained true nucleoli did not contain any of the nucleolus-like bodies, presumably because their nucleolar material incorporated into intact, functional nucleoli. DiVerences in the ultrastructure of these nucleolus-like bodies may occur between diVerent cell types: Hernandez-Verdun et al. (1979, 1991) noted that similar nucleolus-like bodies (‘‘blobs’’ or ‘‘dots’’) formed in micronuclei of colchicine-treated rat kangaroo PTO cells or PtK1 cells, but that these bodies lacked the granular caps observed by Phillips and Phillips (1979) in the Xenopus bodies. These nucleolus-like bodies (dots) in micronuclei of PtK1 cells contained antigens specific for all three compartments of interphase nucleoli—the FCs, DFCs, and GCs—although their ultrastructure closely resembled the DFCs of interphase nucleoli (Hernandez-Verdun et al., 1991). To summarize thus far, in the absence of rDNA transcription, prenucleolar bodies and nucleolus-like bodies can form, but intact, functional nucleoli fail to assemble. To better examine the formation of these bodies, Bell et al. (1992) used bacteriophage lambda phage DNA to generate pseudonuclei in activated Xenopus egg extracts. The use of lambda DNA ensured that there were no ribosomal genes present that could potentially give rise to micronucleoli. Pseudonuclei formed 90 minutes after adding purified lambda DNA to the extract. These nuclei contained electron-dense spheres that ranged from 0.15 to 0.4 mm in diameter. The bodies were first detectable at 45 minutes but grew in size over time. Immuno-fluorescence localization of specific antibodies


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showed that these dense bodies contained fibrillarin, a 180-kDa protein that was presumably the Nopp140 homologue in Xenopus (Cairns and McStay, 1995), nucleolin, and at least some B23. The bodies did not contain Pol I, Topo I, or the 7-2-RNP. In situ hybridizations failed to detect the U3 snoRNA or the 18S and 28S ribosomal RNAs (see below, however). By adding wheat germ agglutinin (WGA) to the extract before the addition of DNA, Bell et al. (1992) were able to form pseudonuclei that lacked nuclear pore complexes. No dense bodies formed in these pseudonuclei, indicating that their de novo formation is dependent on soluble, depolymerized components that must traverse the nuclear envelope. Thus, the first step in nucleologenesis is the assembly of prenucleolar bodies. Without rDNA transcription, however, fully functional nucleoli fail to form. Detailed molecular studies of de novo nucleologenesis in Xenopus embryos (Verheggen et al., 1998, 2000) further support the notion that initial nucleologenesis occurs in the absence of rRNA synthesis. Xenopus embryos remain transcriptionally silent through their first 12 rapid cell divisions. Transcription of Pol II genes begins at the midblastula transition (MBT is 7 hours after fertilization at 23 C) as cell cycles increase in duration. Pol III genes followed by Pol I genes begin their transcription in a sequential order over the next several division cycles (for details, see Verheggen et al., 1998, and references therein). In 6-hour embryos, the bulk of UBF localizes to the cytoplasm, whereas after the MBT, the majority of UBF localizes to the nuclei. A similar cytoplasmic-to-nuclear redistribution occurs for nucleolin (Meßmer and Dreyer, 1993). Nuclear UBF consistently associated with the NORs from the 6-hour time point and onward throughout embryogenesis. Before MBT, fibrillarin localized to typical PNBs, and nucleolin localized to smaller, more diVusely distributed particles. At MBT, nucleolin, fibrillarin, and partially processed maternal pre-rRNA began to regroup into new nucleolar domains surrounding the rDNA. These nucleolar domains resembled DFC-like structures. Using in situ run-on incorporation of Br-UTP in isolated embryonic nuclei as a sensitive assay for Pol I transcription, coupled with immuno-localizations of fibrillarin in the same nuclei, the authors showed that the earliest Pol I transcription started at 9 hours postfertilization (at the end of MBT) in a very low percentage of embryonic nuclei. The vast majority of embryonic nucleoli activated their Pol I transcription after the end of MBT. This was well after the first regroupments of fibrillarin into the nucleolar domains that formed around the rDNA (NORs). The significant finding is that nucleolar-like structures do form before (in the absence of) rDNA transcription. Several earlier studies used Act D in concentrations ranging from 0.04 to 0.2 mg/mL to block Pol I transcription in telophase and early interphase (Benavente et al., 1987; Dousset et al., 2000; Gimeńez-Martin et al., 1977; Ochs et al., 1985; Scheer et al., 1975; Semeshin et al., 1975). Act D

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intercalates into DNA with a preference for GC-rich sequences (e.g., ribosomal DNA). Thus, very low concentrations of Act D selectively inhibit RNA Pol I versus Pol II and Pol III. In the presence of Act D, UBF and Pol I remain bound to the NORs (Dousset et al., 2000), and PNBs still form in telophase (Benavente et al., 1987; Dousset et al., 2000; Ochs et al., 1985). Act D (0.08 to 0.2 mg/mL) prevents PNBs fusion at the NORs, presumably by blocking Pol I transcription, but at lower concentrations (0.05 mg/mL) small nucleolar-like bodies still form at the NORs (Semeshin et al., 1975). In a detailed study using slightly lower concentrations of Act D (0.04 mg/m), Dousset et al. (2000) showed that nucleolin and fibrillarin initially associate with UBF-positive NORs in Act D–treated telophase cells. As the cells progressed into G1, two types of nucleolar-like bodies were evident by transmission electron microscopy. The first were typical electron-dense PNBs that failed to fuse, but the second appeared as small segregated mininucleoli that had separated into light (fibrillar) and dark (granular) regions as they appear by transmission electron microscopy. The light regions contained UBF, nucleolin, and fibrillarin, whereas the dark regions contained partially processed pre-rRNA (see later discussion). These segregated mininucleoli strongly resembled interphase nucleoli that segregate into light and dark regions (capped nucleoli) because of Act D inhibition of rRNA synthesis (see Fig. 1.7 of Busch and Smetana, 1970). C. Nucleolar Assembly in Telophase/Early G1 1. A Primer for Nucleologenesis The observations described above support a two-step model for nucleologenesis (Fig. 8 of Dousset et al., 2000): first, in early telophase, competent NORs with the bound transcription factors UBF, TIF-IB/SL1, and Pol I directly recruit pre-rRNA processing components (e.g., nucleolin and fibrillarin) along with residual nonprocessed pre-rRNA synthesized in late G2/early M phase. Some initial recruitment may occur in the absence of new pre-rRNA synthesis in very early telophase. The second aspect of nucleologenesis is the transfer of PNB material to nascent nucleoli; this transfer is dependent on reinitiating rRNA transcription. However, questions remain: Where do PNBs come from? What is the molecular composition of PNBs? What kinetic relationships exist between cytoplasmic NDF and the PNBs, and between the PNBs and the reactivated NORs? We describe what we know about nucleolar disassembly later in the cyclical pathway, but to set the stage for a discussion of reassembly, let me remind the reader that on disassembly, most nucleolar processing components scatter to the cytoplasm and the perichromosomal regions (sheaths). It is from this point that we begin our description of nucleologenesis.


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2. Perichromosomal Regions As nucleoli disassemble in prophase of mitosis, their transcriptional components remain associated with the mitotic NORs, and several of their prerRNA processing and ribosome assembly components redistribute first to a nucleoplasmic network and then to the peripheral regions of the mitotic chromosomes. Several early studies described silver-staining nucleolar proteins coating the mitotic chromosomes (Paweletz and Risuenõ, 1982). In particular, the ultrastructural images provided by Paweletz and Risuenõ (1982) clearly demonstrated argyrophilic material first permeating the chromatin of prophase HeLa cells as nucleoli disassembled. By metaphase, the material formed a complete peripheral sheath around the individual chromosomes. These sheaths remained intact and associated with the anaphase chromosomes as the chromosomes moved to opposite poles. As the chromosomes began to decondense at telophase, the peripheral sheaths began to break down, forming silver-stained accumulations within interchromosomal spaces. These accumulations then gave rise to prenucleolar bodies that then fused at the NORs. Paweletz and Risuenõ (1982) convincingly demonstrated that the silver-stained material, tracked from prophase to telophase at the ultrastructural level using a sensitive silver impregnation technique, had to be nucleolar in origin. Specifically, the addition of 2 mM adenosine to the culture medium prevented the perichromosomal sheaths from forming around the chromosomes in prophase cells, whereas the exogenous adenosine prevented small, silver-stained bodies (presumably PNBs) from fusing with the NORs in telophase cells. Exogenous adenosine at 2 mM disrupts interphase nucleoli. Yasuda and Maul (1990) carefully tracked fibrillarin through mitosis and then provided some provocative ideas concerning the function of the perichromosomal sheath. In early prophase, fibrillarin is released from the nucleolus. At first, fibrillarin accumulated in between the condensing chromosomes. Soon after, however, it associated with chromosome surfaces before the nucleolus and nuclear envelope had completed disassembled. Yasuda and Maul (1990) speculated that mitotic phosphorylation may regulate the quick transfer of free fibrillarin to the chromosome surfaces. Fibrillarin associated with the chromosome surfaces either as relatively large RNP granules or as fine, densely packed material, which they interpreted to be fibrillarin free of any RNA. Except for centromeric regions, fibrillarin localized along the sides of the chromosomes that faced the cytoplasm. This localization was coincident with lamin B. Gautier et al. (1992a,b,c, 1994) used several autoimmune sera to localize nucleolar antigens to the perichromosomal sheath during mitosis. Table I summarizes those known nucleolar components that associate with the perichromosomal regions during mitosis in animal and plant cells. Gautier

TABLE I Distribution of Nucleolar Components during Anaphase and Early Telophase Cyto

PCS

NORs

þ

þ

NDF

PNBs

References

snoRNAs U3

þ

Jimeńez-Garcıá et al., 1994

þ

Azum-Ge´lade et al., 1994

þ þ

þ

Gautier et al., 1992a,b, 1994

þ

Beven et al., 1996 Dundr et al., 1997

U8

128

U14

þ

þ

þ

þ

þ

Dundr and Olson, 1998 þ

Beven et al., 1996

þ

Ochs et al., 1985

Nucleolar/ribosomal proteins from the DFCs and GCs Fibrillarin

þ

Jimeńez-Garcıá et al., 1989 þ

Yasuda and Maul, 1990

þ

Gautier et al., 1992, 1994 þ

Azum-Ge´lade et al., 1994 Hernandez-Verdun and Gautier, 1994

þ

þ

þ

Medina et al., 1995 Weisenberger and Scheer, 1995 Beven et al., 1996 Dundr et al., 1996, 1997

þ B23

þ

þ

þ

Fomproix et al., 1998

þ

Ochs and Busch, 1984; Ochs et al., 1985

Schmidt-Zachmann et al., 1987 Dundr et al., 1996, 1997 Dundr and Olson, 1998 Ribocharin

þ

Perichromonucleolin

þ

Nucleolin

þ þ

Hu¨gle et al., 1985b Shi et al., 1987 þ þ

þ

Ochs and Busch, 1984; Ochs et al., 1985

þ

Medina et al., 1995 Weisenberger and Scheer, 1995 Dundr et al., 1996, 1997 Pinõl-Roma, 1999

Ki-67 antigen

þ

Gerdes et al., 1984

þ

Verheijen et al., 1989

129

Dundr and Olson, 1998

hPop1 þ

p52 (GC)

Gautier et al., 1994

þ p66 (GC)

þ

p103 (DFC and GC)

þ

No55

þ

þ

þ

Dundr et al., 1996, 1997 Gautier et al., 1992a Gautier et al., 1992a

þ

þ

Ochs et al., 1996 Dundr et al., 1997

SSBI

þ

þ

Nopp140 (DFC)

þ

þ

Beven et al., 1996

/þ

Pai et al., 1995 Dundr et al., 1997

þ

Ribosomal Protein S1

Hu¨gle et al., 1985a

Proteins of the FCs RNA Pol I (FC)

þ

Scheer and Rose, 1984 (continued)

TABLE I (continued) Cyto

PCS

NORs

NDF

References

PNBs

Haaf et al., 1988 Gilbert et al., 1995 UBF (FC)

þ

Chan et al., 1991 Rendon et al., 1992 Roussel et al., 1993

Topo I (FC)

þ

Guldner et al., 1986

pp135 (FC)

þ

Pfeifle et al., 1986

Nucleolar RNA

þ

þ

18S-ITS1-5.8 S -ITS2-28 S

þ

þ

Preribosomal RNA Lepoint and Goessens, 1978

130

þ

þ*

32 S pre-rRNA

Pinõl-Roma, 1999

0

Blum et al., 1986

5 ETS leader 50 ETS core

Medina et al., 1995

þ

þ

{


Pierron and Puvion-Dutilleul, 1996

þ

Weisenberger and Scheer, 1995

þ

{

Pierron and Puvion-Dutilleul, 1996

Beven et al., 1996

þ

Dundr and Olson, 1998 Blum et al., 1986

þ*

Pinõl-Roma, 1999

þ 18 S

þ

þ

þ

Dundr et al., 2000 Dundr and Olson, 1998

þ*

Pinõl-Roma, 1999

þ ITS1

þ

þ

þ

28 S

Dundr et al., 2000

þ

Beven et al., 1996

þ


þ


þ*

30 ETS

þ

Pinõl-Roma, 1999 þ


Cyto, cytoplasm; PCS, perichromosomal sheath; NDF, nucleolar derived foci; PNBs, prenucleolar bodies; NOR, nucleolar organizer regions; FC, fibrillar center; DFC, dense fibrillar component; GC, granular component; ETS, external transcribed spnur. * Pinõl-Roma (1999) used metaphase arrested cells. Presumably, the detected ribosomal RNAs were associated with the peri-chromosomal sheath. { Physarum polycephalum has a closed mitosis in its plasmodium stage, and its rDNA segregates into many small nucleolar remnants during mitosis (see text) that are categorized here as PNBs.

131

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PATRICK J. DIMARIO

et al. (1992b) showed that nucleolar antigens first move away from the nucleoli in prophase toward the nuclear envelope along defined paths within the nucleoplasm. By metaphase, the antigens have reaccumulated into the perichromosomal sheath. Association of nucleolar proteins to the chromosome periphery is concurrent with chromosome condensation, and Gautier et al. (1992a) suggested that there may be a functional correlation between these two events, although no direct correlation has yet been established. Once associated with the periphery, the nucleolar antigens may act as passengers on chromatids that separate at anaphase (Gautier et al., 1992a, 1994). The chromosomal sheath may simply serve as an eYcient mechanism to distribute nucleolar proteins equally to daughter cells. One autoimmune serum, D09, labeled not only the periphery of HeLa metaphase chromosomes but also an interconnected network between chromosomes. Thus, a simplistic passenger model may not be suYcient for all nucleolar antigens. What directs nucleolar proteins to the chromosome periphery remains an open question. Gautier et al. (1992a, 1994) also used immunoprecipitation and fluorescence in situ hybridization to establish that snoRNA U3 resides within the perichromosomal sheath; the authors suggested that the U3 snoRNP particle may be kept intact during mitosis. Further extending these studies, Gautier et al. (1992b) showed that the perichromosomal sheath was fairly uniform over the length of the chromosomes, including the telomeres. The exception was at centromeric regions, where the sheath was not visible. Centromeric regions appeared as dark zones in their immunolocalization of nucleolar antigens, perhaps because of the exclusion of nucleolar antigens by centromeric proteins (CENPs). Finally, the perichromosomal sheath resisted extraction when isolated chromosomes were treated with hypotonic solutions known to disrupt interphase nucleoli (Gautier et al., 1992b). This observation indicates that interactions holding the nucleolar antigens to the sheath are not necessarily the same interactions encountered by these antigens in interphase nucleoli. Gautier et al. (1992c) also used cryosectioning and freeze-substitution (using acetone to substitute, followed by 1% OsO4 in acetone, and then Epon embedding) to examine the perichromosomal sheath at the ultrastructural level. In cryosections, the perichromosomal material consisted of closely packed fibrils unevenly distributed around the chromosomes. The sheath was thin but dense in some regions, and it was thick and loosely packed in other regions. In freeze-substituted cells, the sheath formed a continuous fibrous layer surrounding each chromosome and often between the sister chromatids of a given mitotic chromosome. The thickness of the sheath varied, however. In addition to the fibrous nature of the sheath, the freeze-substituted material displayed granules with 11–16-nm diameters that were interconnected by thinner fibrils to form chainlike structures. These granules appeared similar to interchromatin granules of interphase cells prepared in similar manners.


133

Several other studies demonstrated the localization of nucleolar processing components to the perichromosomal sheath (see Table I). Medina et al. (1995) suggested that nucleolar material within the perichromosomal sheath gives rise to interchromosomal material that then forms PNBs. In addition to nucleolar components, the perichromosomal sheath also contains nonnucleolar proteins such as perichromonucleolin (PCN) (Shi et al., 1987), the phosphorylated form of nucleoplasmin (Dilworth, 1991), nuclear envelope proteins, and proteins that appear diVuse in the interphase nucleus (Rattner, 1992). What role might nucleolar proteins play in coating the mitotic chromosomes in higher eukaryotes? Yasuda and Maul (1990) suggested that the fibrillarin-containing RNP material works in concert with other perichromosomal proteins (perichromin) to coat and insulate the chromosomes from the surrounding cytoplasm. This perichromosomal sheath could also help maintain the compacted state of the chromosomes, whereas selective loss of sheath proteins in telophase and early G1 could result in nonrandom distributions of chromosomal domains within the reforming nuclei. Recall that McClintock (1934) put forth a similar hypothesis. Perhaps the sheath plays a structural role in chromatin condensation or a functional role by sequestering cell cycle regulatory proteins in addition to the nucleolar proteins, as suggested by Savino et al. (2001). The sheath may protect the condensed DNA, or it may serve to equally divide nucleolar components in cells that normally disassemble their nucleoli (Hernandez-Verdun, 2003; Hernandez-Verdun et al., 1993). Clearly, the perichromosomal sheath is an aspect of nuclear cell biology that warrants further investigations. 3. PNBs In Depth a. Early Work Lafontaine (1958) provided a detailed historical account of the early work on prenucleolar material in anaphase and telophase cells. According to Lafontaine (1958), fusion of prenucleolar bodies to form nucleoli in telophase was first noted by Van Camp (1924) well before Heitz and McClintock established the secondary constrictions as the NORs. Some of the early ultrastructural observations on prenucleolar material in prenucleolar bodies and perichromosomal sheaths were made in Allium cepa (onion) and Vicia faba (broad bean) by Lafontaine (1958) and Lafontaine and Chouinard (1963). Jacob and Sirlin (1963) described PNBs in the larval salivary glands of Bradysia mycorum (a sciarid), and Stevens (1965) examined embryonic neuroblasts of Chortophaga viridifasciata (grasshopper). In her report, Stevens noted the reappearance of nucleolar material in anaphase as small granules (0.2–0.3 mm in diameter) on the surfaces of the chromosomes. Next to appear were bodies of 0.3–0.7 mm in diameter that had lost their attachment to the chromosomes, but that still maintained a composition

134

PATRICK J. DIMARIO

quite similar to the smaller granules still attached to the chromosome surfaces. Similar to studies that preceded hers, Stevens suggested a sequential flow of nucleolar material from the chromosome surfaces to PNBs, and then to the growing telophase nucleoli. As telophase nucleoli grew in size, the number of PNBs diminished. Morcillo et al. (1976) emphasized that PNBs in A cepa did not result from a synthesis of pre-rRNA because these bodies appeared before the reinitiation of rRNA synthesis. b. Composition Table I summarizes the nucleolar components that localize to PNBs. Modern molecular techniques and confocal microscopy have allowed in-depth analyses of the PNBs. For example, Jimeńez-Garcıá et al. (1994) localized the U3 snRNP to PNBs in HeLa and NRK (normal rat kidney) epithelial cells during telophase. These PNBs did not apparently contain rRNA or ribosomal protein S6. In HeLa cells, ribosomal RNA was excluded from the chromosome arms in the HeLa cells from prophase to anaphase but reappeared in reforming nucleoli during telophase. In the NRK cells, however, rRNA localized between the chromosome arms from prophase to telophase but was not directly associated with the condensed chromatin in the form of a sheath. Bell and Scheer (1996) depleted Xenopus egg extracts of individual nucleolar proteins to determine which component is necessary for PNB formation. Interestingly, egg extracts immuno-depleted separately of fibrillarin, nucleolin, Nopp180 (the Xenopus version of Nopp140), or B23 could still assemble PNBs. Only the fibrillarin-depleted extract seemed to have fewer PNBs. Extracts lacking U3 by RNase digestion were also able to assemble PNBs. Of note among the various nuclear components within the egg extract is p80 coilin, the marker protein for the RNP-rich nuclear organelles called Cajal bodies (CBs; Gall, 2000, 2003; Ogg and Lamond, 2002). CBs were originally described by Ramoń y Cajal (1903) as accessory bodies to nucleoli. In addition to coilin, CBs contain the nucleolar proteins fibrillarin, Nopp140, NAP57, Gar1, the large subunit of RNA Pol I, TOPO I, ribosomal protein S6, and snoRNAs U3 and U8. CBs also contain several pre-mRNA splicing components (SM-containing snRNPs, splicing snRNAs). CBs do not contain nucleolin, B23, pre-rRNAs, and pre-mRNAs. Thus, CBs are not directly involved in pre-rRNA processing or in pre-mRNA splicing. The primary function of CBs may be to preassemble and chemically modify the processing components before they are shipped oV to the nucleolus or to sites of premRNA splicing. With respect to PNBs, Bell and Scheer (1996) demonstrated ultrastructural diVerences between CBs and PNBs that formed in Xenopus egg extracts: PNBs appeared electron dense and quite similar to the DFCs of interphase nucleoli. CBs, in contrast, appeared less electron dense, with their RNPs loosely packed. Bell and Scheer (1996) showed that although nucleolin


135

and B23 (NO38 in Xenopus) did not localize to CBs, p80 coilin localized to PNBs along with fibrillarin. Coilin, however, did not localize to reforming nucleoli during telophase. c. PNBs Are Dynamic Savino et al. (1999, 2001) characterized the flow of nucleolar material between PNBs, and from PNBs to the reforming nucleoli. They used three-dimensional deconvolution imaging and time-lapse (fourth dimension) microscopy to localize GFP-tagged fibrillarin and Nop52 in stably transfected HeLa cells from metaphase to G1. As described above, fibrillarin normally localizes in the DFC during interphase to function early in 50 ETS processing and site-specific methylation of pre-rRNA. Conversely, Nop56 localizes to the GC and assists in the later removal of the second ITS. Fibrillarin-GFP and Nop56-GFP localized normally at all points of the cell cycle; fibrillarin-GFP localized to the peri-chromosomal sheath, PNBs, CBs during metaphase and anaphase, and to the reforming nucleolus during telophase. Nop56-GFP colocalized with fibrillarin to the sheath during metaphase and anaphase. Most interestingly, however, the authors showed diVerent kinetics of redistribution and relocalization for the two proteins. As soon as the chromosomes reached the poles, fibrillarin-GFP was the first to dissociate from the chromosomal sheath as it entered PNBs and reactivating NORs. Nop56-GFP, in contract, was much slower to dissociate from the sheath. Nop56 redistribute to PNBs, but these so-called ‘‘late PNBs’’ were distinct from the ‘‘early PNBs’’ that carried fibrillarin. Nop56 finally reached the nucleoli in early G1. Using time-lapse video microscopy, Savino et al. (2001) confirmed the existence of at least early and late PNBs. The fibrillarin-containing PNBs were relatively short lived, lasting only 15 5 minutes, whereas the Nop56containing PNBs were stable, lasting about 80 20 minutes. Using fourdimensional imaging, the authors observed PNBs oscillating (without directional movement) toward reforming nucleoli. They also observed bridges linking one PNB to another. As the GFP signal diminished in one interconnected PNB, it increased in the other, presumably by the unidirectional flow of nucleolar material via connecting bridges. Likewise, those PNBs containing Nop-56 that made contact with nucleoli did not completely fuse with reforming nucleoli. Instead, material transferred in a unidirectional manner from the PNB to the nucleolus via fibrillar bridges that remain compositionally and mechanistically unexplored. The model of PNB function put forth by Savino et al. (2001), therefore, describes a directional flow of nucleolar material between PNBs, and then from PNBs to more stable and rapidly expanding NORs. Photobleaching (FRAP) assays by Dundr et al. (2000) and Phair and Mistelli (2000) (see later discussion) indicate that rapid diVusion through the nucleoplasm also plays an important role in moving nucleolar components between PNBs, and from PNBs to reforming nucleoli.

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PATRICK J. DIMARIO

The production of pre-rRNA in the NORs is likely the molecular sink ensuring the unidirectional flow of processing complexes toward the nucleolus. 4. NDF Dundr et al. (1996) first described NDF in mitotic CMT3 monkey cells stably transfected to express the HIV Rev protein; specifically, they examined the distribution of the HIV Rev protein as it compared with several endogenous nonribosomal nucleolar proteins. Wild-type Rev colocalized with the DFC proteins fibrillarin, B23, and nucleolin, and with the GC protein, p52, as they moved from disassembling nucleoli in prophase to the perichromosomal regions in prometaphase and metaphase. Beyond this point, Rev leaves the general paradigm for nucleolar proteins. At telophase, when PNBs formed inside the reforming nuclear envelope, the Rev protein remained in the cytoplasm, within discrete foci, now referred to as the NDF. Rev rejoins the nucleolus in early G1, once rRNA transcription had been initiated and subsequently maintained (Dundr et al., 1995). NDF in CMT3 cells are quite numerous during anaphase (typically 25–50 per cell, with some cells containing over 100). The NDF distribute throughout the cytoplasm but are excluded from the spindle. NDF were not a consequence of Rev expression, as similar foci were observed in nontransfected COS-7 cells and HeLa cells (see below). Although the NDF in the Rev-expressing CMT3 cells were variable in size, they are typically 1–2 mm in other cell types. NDF are composed of RNP particles that contain several nonribosomal nucleolar components (see Table I). Dundr et al. (1997) carefully described the molecular composition of NDF at the various stages of mitosis. Nucleolar proteins fibrillarin, B23, nucleolin, and p52 associate with NDF during anaphase and telophase. The U3 snoRNA is also present in B23-containing NDF during anaphase, and a subsequent paper showed the presence of U8 in NDF (Dundr and Olson, 1998). These various nucleolar components likely track together, first within the chromosomal periphery (sheath) from the time nucleoli disassemble in prophase, and then to the NDF in anaphase, to the PNBs that form in late anaphase and telophase as the nuclear envelope reforms, and finally to the reforming nucleoli in telophase and early G1. Interestingly, Dundr et al. (1997) showed that Nopp140 localizes to the cytoplasm as nucleoli disassemble, but later in anaphase and telophase, Nopp140 remains independent of NDF and PNBs before its direct association with reforming nucleoli in telophase. NDF first appear outside the mitotic spindle in early anaphase. In telophase, however, the number of cytoplasmic NDF declines significantly as the number of nucleoplasmic PNBs increases and as nucleoli begin their reassembly at the NORs. Most nucleolar components so far examined follow this


137

pathway, although Medina et al. (1995) suggested that the perichromosomal sheath gives rise to irregular interchromosomal material, which in turn gives rise to PNBs. 5. Partially Processed Pre-rRNA in the NDF A few early studies indicated the persistence of partially processed pre-rRNA through mitosis and on into the subsequent interphase (Abramova and Neyfakh, 1973; Fan and Penman, 1971). Fan and Penman (1971) isolated RNA from interphase and metaphase-arrested cells and showed that persistent 45 S and 32 S pre-rRNA in the metaphase-arrested cells was stable during mitosis and identical to that isolated from the interphase cells. They also showed that on release of metaphase block, precursor rRNAs were once again unstable (i.e., processed), and that the resumed processing mechanism was insensitive to the block of rDNA transcription by Act D. Processing was also largely insensitive to the block in new protein synthesis by cycloheximide. Fan and Penman further showed that the partially processed rRNA associated with the mitotic chromatin (presumably the perichromosomal sheath) after fractionation of the metaphase arrested cells. More recently, Dundr and Olson (1998) used fluorescence in situ hybridizations and Northern blot analyses to show the existence of partially processed pre-rRNA in NDF. Selective biotinylated antisense riboprobes complementary to the 50 ETS core, the 18 S region, ITS1, the tail end of the 28 S regions, and the 30 ETS, all labeled NDF in anaphase CMT3 (monkey) cells. Specifically, the 50 ETS core riboprobe labeled NDF and the perichromosomal sheath, both of which contained B23. The 18S- and 28S-specific probes labeled the cytoplasm in a general manner as expected, but both probes also labeled discrete B23-containing foci within the cytoplasm of anaphase and telophase cells. The 30 ETS-specific probe labeled NDF, the cytoplasm, and reactivated nucleoli during telophase. This 30 ETS signal, however, was significantly weaker than the 50 ETS core signal, indicating that a large proportion of the transcripts were completed and terminated by the time they were released on mitotic arrest of rDNA transcription. Although the riboprobe for the 50 ETS leader sequence labeled intranucleolar regions from interphase through prophase, the same riboprobe failed to label any structure from prometaphase through late telophase, at which time it labeled only the reactivated nucleoli. This probe for the 50 leader completely failed to label B23-containing cytoplasmic NDF and nucleoplasmic PNBs. The data are consistent with the rapid cleavage of the 50 ETS leader from the 47 S precursor to yield the 46 S intermediate with its 30 ETS still intact, as compared to the 45 S that lacks the 30 ETS. None of the riboprobes labeled the NORs from prometaphase through anaphase, indicating a complete release of pre-rRNA from the mitotically silenced rDNA. As mentioned above, U8

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occurs in NDF. Because U8 functions relatively late in the removal of 5.8 S and 28 S rRNAs, NDF apparently contain early- and late-acting processing components, perhaps still associated as complexes. According to Dundr and Olson (1998), the NDF may preserve the integrity of these processing complexes until nucleoli reform in telophase. 6. NDF and PNB Dynamics In a study similar to the dynamic studies of Savino et al. (2001), Dundr et al. (2000) used time-lapse studies of green fluorescent protein (GFP)-tagged B23, fibrillarin, and nucleolin in transfected CMT3 cells to show random movements of NDF (1.8–15 mm/min) during telophase, and in rare cases fusions between NDF. The number of NDF declined during telophase as the amount of GFP fluorescence increased at the reforming nuclear envelope. The interpretation here is that as cytoplasmic NDF disassembled in telophase, their component macromolecules entered the reforming telophase nucleus. From immunofluorescence microscopy, fluorescent in situ hybridization (FISH), fluorescence recovery after photobleaching (FRAP), and ultrastructural examinations, Dundr et al. (2000) concluded that NDF and PNBs contain the same nucleolar components (proteins and pre-rRNA), and they have similar fibrogranular structures. Thus as NDF disassemble, their components enter the nucleus and contribute to the formation of PNBs. In their studies, Dundr et al. showed that the PNBs in early telophase appeared irregular in shape and displayed rapid movement (2.4 mm/min), although many PNBs were interconnected. Some PNBs fused to form larger ones, and other PNBs made contact with reforming nucleoli via thin fluorescent threads. Time-lapse microscopy showed that as the size and fluorescence intensity of reemerging nucleoli both increased, the number, size, and fluorescence intensities of the PNBs concomitantly decreased. Dundr et al. (2000) further showed that the timing of nucleolar component association with reforming nucleoli was not identical for diVerent components. For example, B23 and fibrillarin colocalized to NDF, perichromosomal regions, and the cytoplasm in general in early anaphase. In late anaphase, however, localization of the two proteins began to diVer in their localizations. Both proteins continued their colocalization to NDF and perichromosomal regions, but fibrillarin also appeared in the small nucleoli that were just beginning to reform, whereas B23 did not associate with nucleoli until late telophase. Some B23 even remained associated with PNBs that persisted into early G1. 7. Summary Model for Postmitotic Mammalian Nucleologenesis Dundr et al. (2000) put forth the following model for mammalian postmitotic nucleologenesis (Fig. 2). Transcription comes to a halt in early mitosis, but those transcripts that were under construction at the very onset of mitosis are


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FIG. 2 Models for nucleolar disassembly, the perichromosomal regions, and initial postmitotic nucleologenesis. (A) At the onset of mitosis (prophase), the nuclear envelope is still present, but chromosomes become visible as thin threads. The nucleolus begins to disassemble by the movement of processing components (white arrows) out toward the nuclear envelope. (B) By metaphase, much of the nucleolar processing components line the periphery of the condensed chromosomes to form the perichromosomal compartment (black arrows). More processing components occupy spaces between the chromosomes (white arrow). (C) Nucleologenesis begins in telophase. The nuclear envelope begins its reassembly (oval-shaped vesicles), and the chromosomes begin to decondense. Competent NORs (the darkened secondary constriction denoting silver-staining transcription factors) are the first loci to reactivate transcription. Prenucleolar bodies (PNBs, black and white arrows) form from nucleolar RNP material that previously associated with the perichromosomal and interchromosomal spaces. The black PNBs contain early processing components such as partially processed pre-rRNA, nucleolin, and fibrillarin. The flow of these components back to reactivated nucleoli is by diVusion. The picture shows a unidirectional flow of these materials toward the NOR, but diVusion may be in all directions. The white PNBs contain late processing components such as B23 and Nop52. Either the late components are slower to diVuse from the PNBs as compared to the early processing components, or the late processing components constitute their own unique PNBs. Nucleus derived foci (NDF, white arrow head in the cytoplasm) are closely related to PNBs in RNP composition and dynamic assembly/disassembly. NDF disassemble by diVusion, and their components transport across the reforming nuclear envelope.

completed and released from the transcription machinery. As the DFC breaks down in prophase (Fig. 2A), these partially processed pre-rRNAs (lacking their 50 ETS leader sequences) remain associated with the DFC components (proteins and snoRNAs). This DFC material forms the perichromosomal nucleolar material by metaphase (Fig. 2B). At anaphase, some of this material remains within the perichromosomal region, while other portions form cytoplasmic NDF. As the nuclear envelope reforms around decondensing chromosomes in telophase, nucleolar material translocates from the NDF to the nuclear envelope and then across the envelope into the nucleoplasm, where it then incorporates into PNBs (Fig. 2C). Reactivation of rDNA transcription in late anaphase/early telophase initiates

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the return of PNBs (DFC material) back to the NORs (FCs). The staggered return of individual processing components (fibrillarin and nucleolin before B23) may reflect the order in which these respective proteins are needed for the sequential processing of pre-rRNA. Dundr et al. (2000) suggested that this return of DFC material to the reactivated NORs was by diVusion rather than by an active transport mechanism.

D. Other Model Systems 1. Closed Mitosis of Physarum The plasmodium of Physarum polycephalum displays a slightly diVerent program for nucleologenesis. The plasmodium contains multiple diploid nuclei that synchronously undergo a closed mitosis (Guttes et al., 1968, for an ultrastructural analysis of mitosis in Physarum). A diploid nucleus in Physarum contains approximately 200 linear minichromosomes; each chromosome is 60 kbp in length, with two divergent rDNA genes transcribed from the center toward the ends (Pierron and Puvion-Dutilleul, 1993, and references therein). At prophase, a large central nucleolus disassembles into multiple nucleolar remnants that retain a minichromosome with attached DFC and GC regions. Disassembly of the large central nucleolus is concomitant with the cessation of rDNA transcription in prophase. Guttes et al. (1968) reported that by mid- to late prophase the nucleolar remnants move farther apart from each other and continue to disassemble. In addition to these nucleolar remnants, Guttes et al. (1968) noted a thin layer of ‘‘seemingly amorphous material’’ on one side of the chromosomes. Pierron and Puvion-Dutilleul (1993) used high-resolution in situ hybridizations with rDNA probes to show that by late prophase, the nucleolar remnants were scattered between the chromosomes, but the authors made no mention of perichromosomal labeling with biotinylated rDNA probes. By metaphase the remnants were located at the nuclear border outside the metaphase plate area. Guttes et al. (1968) showed that the nucleolar remnants moved with the daughter plates toward the poles during anaphase; most of the remnants were attached to the side of daughter plates adjacent to the interzone separating the daughter plates. Pierron and Puvion-Dutilleul (1993) showed that these remnants still consisted of DFC and GC components. By telophase, the remnants collected at the internal surface of the daughter chromatids that had moved to the poles. The nuclear region between the chromosome masses also contained remnants, but the chromosomes themselves remained unlabeled by a biotinylated rDNA probe. At the end of telophase, the remnants (now referred to as PNBs) were dispersed among the mass of still-condensed chromatin. These PNBs still retained DFC- and GC-like regions, but with a


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translucent central core. The PNBs progressively fused into from one to three nucleoli by G1, and these small nucleoli further coalesced into the one central nucleolus by G2. In their 1993 paper, Pierron and Puvion-Dutilleul demonstrated persistence of rRNA within the mitotic nucleolar remnants, and in their 1996 report, they clearly demonstrated that the rRNA contains the 50 ETS core, but not the 50 ETS leader. Apparently, the 50 ETS leader is cleaved and rapidly degraded within the DFC of the late G2 nucleolus. 2. Amphibian Oogenesis The amphibian oocyte amplifies its rDNA to generate multiple extrachromosomal nucleoli. In Xenopus, for example, ribosomal DNA is selectively amplified during late zygotene and early pachytene of prophase I to produce 30 pg of extrachromosomal rDNA per oocyte nucleus (Gall, 1969; Macgregor, 1972; Scheer and Dabauvalle, 1985). This rDNA amplification is remarkable considering the tetraploid oocyte at this stage of meiosis contains 12.8 pg of chromosomal DNA. At first, the amplified rDNA exists as a large aggregate cap within the nucleus. By early to middiplotene, this rDNA disperses into approximately 1000 extrachromosomal nucleoli (Van Gansen and Schram, 1972). The molecular mechanisms for this unique form of nucleologenesis have remained poorly understood. Toward this end, Spring et al. (1996) and Mais et al. (2002) examined the earliest formation of the multiple nucleoli in Xenopus oocytes. Spring et al. (1996) showed that 2–4 small rDNA clusters derived from the aggregate cap fused to the outside perimeter of prenucleolar protein bodies in late stage I/early stage II oocytes. From S1 transcript analyses, the authors suggested that this fusion occurs largely in the absence of rDNA transcription. With rDNA patches on their periphery, several prenucleolar bodies fused to form ribbon-like complexes. In stage II, the external rDNA translocates into the prenucleolar bodies. Translocation is concomitant with decondensation of the rDNA and transcription initiation. Mais et al. (2002) showed that early diplotene oocytes with diameters of 70 mm contain three kinds of morphologically distinct nuclear structures. The first two were granular bodies of irregular shape that contained nucleolin and spheroidal bodies of fibrillar texture that contained nucleolin and fibrillarin. On the basis of their morphologies and protein compositions, Mais et al. (2002) interpreted the granular bodies to be prenucleolar GCs and the fibrillar bodies to represent pre-DFCs. The third morphological entity was the most interesting: These were rodlike structures that intermingled with, and were adjacent to, the granular and fibrillar granules. In some cases, the rods actually made contact with the other prenucleolar granules. These rods failed to stain with fibrillarin or nucleolin antibodies. Anti-UBF, however, intensely stained the central axes of the rods, leaving the more peripheral

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filamentous regions unlabeled. The sensitive TdT-immunogold method for detecting DNA at the ultrastructural level (Thiry, 1992; Thiry et al., 1993) showed that the more peripheral filamentous regions of the rods did contain DNA, and presumably amplified rDNA. The rods, therefore, represent extrachromosomal NORs that originate from the aggregated cap of amplified rDNA. Interestingly, the rods have similar dimensions and silver-staining properties as found for NORs in somatic mitotic chromosomes. As diplotene progresses, the rods fuse with the filamentous granules to establish the multiple extrachromosomal nucleoli. Future work is necessary to determine whether this fusion is dependent on initiation of rDNA transcription within the rods, or whether other factors such as RNA-independent mechanisms (i.e., protein–protein interactions) may contribute to fusion, and thus nucleologenesis. Although amplified rDNA and multiple extrachromosomal nucleoli have been particularly well studied in amphibian oocytes (e.g., Gall, 1978; Miller, 1981), the phenomenon also exists in other organisms. As examples, amplified rDNA exists in some insects represented in the orders Orthoptera, Neuroptera, Coleoptera, Diptera, and Siphonaptera; in the mollusk Spisula solidissima; in the brine shrimp Artemia salina; and in several bony fish (Kloc et al., 1995; Kubrakiewicz and Bili’nski, 1995; Tobler, 1975). In addition, the unicellular alga Euglena gracilis carries between 400 and 8000 extrachromosomal rDNA circles, each circle containing one rDNA unit (Ravel-Chapuis, 1988). Naegleria gruberi and four other species from the order Schizopyrenida also contain extrachromosomal rDNA plasmids (Clark and Cross, 1988). Finally, hypotrichous ciliates amplify their rDNA during macronuclear diVerentiation (Maercker et al., 1997). Future research on nucleologenesis should be greatly enhanced by the continued characterization of rDNA amplification in these various organisms. 3. Nucleolar ‘‘Assembly’’ in Yeast The yeast nucleolus does not disassemble during mitosis, so postmitotic nucleologenesis as observed in higher eukaryotes may not strictly apply. Yet molecular-genetic studies in yeast have been invaluable in determining what molecular events maintain nucleolar structure (Nomura, 2001). The nucleolus in wild-type Saccharomyces cerevisiae is a relatively large organelle taking up about one-third the nuclear volume. It is crescent-shaped and positioned on one side of the nucleus, closely adjacent to the nuclear envelope. Approximately 100–150 tandemly repeated copies of the rRNA genes reside in one nucleolar organizer on chromosome XII in S. cerevisiae (Petes, 1979). A general impression is that synthesis of the 35S pre-rRNA is the nucleating factor/event driving nucleolar assembly (Me´le`se and Xue, 1995; Warner, 1990).


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Exogenous extrachromosomal plasmids that express 35S pre-rRNA were used to assess the apparent requirements for the tandem arrays of rDNA genes within NORs and for Pol I transcription of this DNA in the process of building a nucleolus (Nierras et al., 1997; Nomura, 2001; Oakes et al., 1993, 1998, 1999; Trumtel et al., 2000). Nomura’s lab use a strain of S. cerevisiae that was completely deficient for the NOR genes on chromosome XII (rdn), but carried one of two multicopy plasmids. The first plasmid (the Pol I helper plasmid) carried a single rDNA repeat with its intact promoter elements. Many small nucleoli formed when multiple copies of this plasmid were transcribed by normal Pol I within the cells. These mininucleoli maintained the same uniform electron density normally seen in the large, crescentshaped nucleolus in wild-type strains, and these multiple nucleoli continued to localize at the nuclear periphery (Oakes et al., 1998). The second plasmid (the Pol II helper plasmid) carried the identical rRNA coding sequence, but without the normal Pol I promoter. Instead it used the GAL7 promoter to drive RNA Pol II transcription of the rRNA, again in the same rdn cells. In this latter case, the cells contained a single, rounded nucleolus that had little or no contact with the nuclear periphery. Unlike the normal crescent-shaped nucleolus in wild-type cells or the mininucleoli formed with the Pol I helper plasmid, the lone nucleolus formed by the Pol II helper plasmids displayed distinct intranucleolar subcompartments. The observed diVerences in nuclear position (peripheral versus central) and intranucleolar substructures were not caused by plasmid copy numbers, but were thought to be caused by the two diVerent transcription machineries (Pol I vs. Pol II). Further, these diVerences in nucleolar organization do not seem to be dependent on the two types of cis-acting promoter sequences: The rdn strain displayed a centralized round nucleolus when it also carried a temperature-sensitive Pol I mutation (rpa12) and expressed rRNA from the Pol I helper plasmid at the growth-permissive temperature. In other words, an intact Pol I with its properly assembled transcription complex may be the primary determinant in forming an intact yeast nucleolus and for placing that nucleolus in close apposition to the nuclear envelope. When mininucleoli formed in rdn cells with the Pol I helper plasmid and a normal Pol I, the nucleoli localized to the nuclear envelope, but their association with the envelope may prevent the individual mininucleoli from fusing into a single larger nucleolus. Nierras et al. (1997) performed similar studies with strains deficient for the NOR, but expressing 35 S pre-rRNA from plasmid-borne genes. They specifically examined pre-rRNA synthesis and processing in the absence of a true nucleolus. Although the plasmid-borne genes were 10% as active as normal tandem rRNA genes in a strain with an intact NOR, methylation and processing of the 35 S transcripts produced on the plasmids occurred normally in the absence of an organized nucleolus. Nop1p (fibrillarin) spread diVusely throughout the nuclei of these strains carrying the plasmid-borne rRNA

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genes. Their results indicated that pre-rRNA synthesis and its processing were distributed to the dispersed plasmids in the absence of a true nucleolus. Trumtel et al. (2000) also examined the tandem organization of rRNA genes and their transcripts as requirements for nucleologenesis in S. cerevisiae. They too used strains deficient in chromosomal rRNA gene transcription: Strain L1494 largely lacked the NOR rRNA genes (ChernoV et al., 1994), and strain NOY 558 lacked Rrnp7p, a component of core factor (CF) that is instrumental in recruiting Pol I to the core promoter element (Keys et al., 1994; Nomura, 2001). Strain L1494 expressed the 35 S pre-rRNA using normal Pol I and a plasmid-borne rRNA gene with its normal Po I promoter. Strain NOY 558 expressed 35 S using Pol II and a plasmid-borne rRNA gene driven by the GAL7 promoter. Using both cryofixation and chemical fixation, Trumtel et al. (2000) recognized the three standard morphological regions (FCs, DFCs, and GCs) within crescent-shaped nucleoli of the respective wild-type parental strains. Nucleolar regions within the two mutant strains that expressed 35 S from the plasmids were morphologically ‘‘peculiar,’’ to use the authors’ description. Strain L1494 contained a well-defined electron-dense region (a DFC) and a lighter granular component, but no FC. The two substructures stretched out over more internal surface area of the nuclear envelope than does a typical crescent-shaped nucleolus. Localization assays identified the 35 S pre-rRNA, Pol I, and processing components Ssb1, Nop1p, and Gar1p within the elongated DFCs of the L1494 strain. Conversely, strain NOY 558 contained one to three spherical mininucleoli scattered throughout the nucleoplasm. These mininucleoli contained a central compact region and a peripheral fibrillar region with interspersed granules. Localization studies showed that the majority of 35 S pre-rRNA and its processing components resided within the peripheral region, but not within the compact core region. By growing NOY 558 cells in glucose, Trumtel et al. (2000) shut down the GAL7 promoter, and thus 35 S transcription. With the loss of pre-rRNA, processing components of the peripheral fibrillar regions dispersed to the nucleoplasm, and the average number of mininucleoli declined. Interestingly, the central compact regions (that remain enigmatic) were left largely unaVected. This dispersal eVect of the peripheral regions could be reversed on returning the cells to a galactose-containing medium. The primary conclusion from this work is that rDNA transcription is the primary determinant for nucleologenesis in yeast. 4. Ectopic Nucleoli in Drosophila: A Model for Nucleologenesis Karpen et al. (1988) used P element-mediated transformation to insert a single Drosophila rDNA gene into several ectopic (i.e., non-NOR) sites within the Drosophila genome. The insertion gene contained the entire rDNA transcription unit along with 50 and 30 flanking intergenic spacer


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sequences (IGSs contain several tandem 240-bp repeats with homology to the rDNA promoter). Karpen et al. (1988) characterized four transformed fly lines in which the rDNA transgene inserted within euchromatic regions 1A, 23E, 68BC, or 94B (the endogenous NORs lie within centromeric heterochromatin of the X and Y chromosomes). Mininucleoli formed at the four ectopic sites on salivary gland polytene chromosomes as determined by RNA–RNA in situ hybridizations that employed either complementary or noncomplementary (control) ETS RNA probes, and by immuno-cytochemistry that used an antibody specific for a 47-kDa nucleolar antigen. In a functional genetic assay, Karpen et al. (1988) showed that the single rDNA insert in region 1A was able to partially rescue the three phenotypes (lethality, a delay in adult eclosion from the pupal case, and perturbations in abdominal cuticle formation in adult flies) typically associated with bobbed (bb) deficiencies that lack endogenous rDNA genes at the normal NORs. Thus, a single rDNA repeat is suYcient to induce mininucleolus formation. McKee et al. (1992) also inserted a full-length rDNA gene along with upstream and downstream IGSs into one ectopic site near the distal tip of the Drosophila X-chromosome. Similar to Karpen et al. (1988), McKee et al. observed a mininucleolus at the ectopic site. McKee et al. (1992) carried the analysis further, however, by creating a series of targeted deletions within the single rDNA gene. These deletions were created by introducing the P element transposase by genetic cross. The transposase then mobilized the P-element harboring the rDNA gene, but imprecise excision of the P element resulted in deletions that eliminated all or part of the single rDNA transcription unit. These deletions prevented formation of the mininucleolus at the site. Some deletions retained IGSs, but they too failed to form mininucleoli, thus suggesting that the transcribed portions of the rDNA gene are necessary for nucleolus formation. One deletion eliminated parts of the 5.8 S region, ITS2, the entire 28 S region, and interestingly, one of the 240-bp IGS repeat sequences at the 50 end of the gene. This deletion also failed to form a mininucleolus, presumably because of the elimination of the downstream coding portions (5.8 S–28 S regions) of the transcription unit rather than the missing single 50 IGS. Thus, production of the entire pre-rRNA may be required for mininucleolus formation.

V. Embryonic Nucleologenesis A. Drosophila Embryogenesis In the past, Drosophila has served as an excellent model organism to characterize rDNA within NORs (Ritossa, 1976), the Minute genes that encode ribosomal proteins (Kay and Jacobs-Lorena, 1987), and as described above,

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the ectopic expression of individual rDNA transcription units that result in the formation of mininucleoli (Karpen et al., 1988; McKee et al., 1992). More recently, several laboratories have initiated studies in Drosophila to characterize nucleolar structure and function. Although not an exhaustive list, recent publications regarding Drosophila nucleoli include those describing small nucleolar RNAs (Tycowski and Steitz, 2001), the Nop60B protein (called minifly) that is the pseudouridylase homologue of human dyskerin (Giordano et al., 1999, 2002; KauVman et al., 2003; Phillips et al., 1998), a DEAD-box RNA helicase encoded by pitchoune (ZaVran et al., 1998), a KH domain protein necessary for pre-rRNA processing encoded by dribble (Chan et al., 2001), and two isoforms of Nopp140 (Waggener and DiMario, 2002). One of the isoforms (Nopp140-True) contains a carboxy terminus that is well conserved with mammalian versions of Nopp140 and is considered a true homolog. The other isoform, Nopp140-RGG, contains a RGG-rich carboxy domain that is common to various RNA-associated proteins but is not observed previously in vertebrate Nopp140 proteins. Both isoforms localize to nucleoli. Surprisingly, Drosophila has not been extensively used to characterize the cell biology of nucleologenesis, so I present some of our work on nucleolar dynamics during Drosophila embryogenesis. We separately transformed Drosophila to express GFP-Nopp140-True or GFP-Nopp140-RGG. Both proteins serve as cytological markers for nucleoli in nearly all cell types during embryogenesis, larval development, and in the adult. We used two separate GFP-Nopp140-RGG transgenic strains to study nucleologenesis in Drosophila embryos. After fertilization, but before cellularization, the Drosophila embryo is a syncytium of rapidly dividing nuclei (Foe et al., 1993). Most of these nuclei migrate out toward the periphery of the embryo to be enclosed by forming blastoderm cells. We examined embryos shortly after fertilization and on through to the late blastoderm stage. GFPNopp140-RGG was not apparent at the earliest precellular stages; it was not detected in any of the syncytial nuclei, nor was it detected in any prenucleolar-like bodies, the existence of which has yet to be established in the Drosophila embryo. We first observed GFP-Nopp140-RGG in the peripheral nuclei during the cellularization process (Fig. 3). Specifically, the tagged protein first appeared within a single large spot within individual peripheral nuclei when descending cell furrows were half complete. These spots should be the nucleoli that are known to form at this stage. We did not see nucleoli form in the ‘‘yolk’’ nuclei that remain behind in the syncytial cytoplasm. The most interesting observation is that nucleoli did not form in the posterior pole cells. Pole cells are the germ-line progenitors that are actually the first cells to form in the embryo (Mahowald, 2001; St. Johnston, 1993). GFP-Nopp140-RGG is clearly present within the pole cell nuclei, but it appears diVuse, and at this resolution, we see no indication of


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FIG. 3 At least two distinct programs for nucleologenesis in the Drosophila embryo. Females homozygous for P[Hsp70-gfp-Nopp140-RGG] were heat shocked for 1 hour and left overnight at room temperature to produce maternal product (mRNA and protein) within their developing egg chambers. Fertilized eggs were collected the next day and allowed to develop at room temperatures while viewing with a confocal microscope. (A) Posterior of a blastoderm stage embryo. Blastoderm cells line the periphery of the yolk-filled syncitial cytoplasm. The black arrow points to the most posterior pole cells, the germ-line progenitors. (B) GFP-Nopp140RGG first appeared in newly formed nucleoli when blastoderm cell furrows were half complete. This is the stage at which nucleoli first become evident (Foe et al., 1993; Mahowald and Hardy, 1985). Although each peripheral blastula cell contained a nucleolus, the germ line pole cells did not display nucleoli. GFP-Nopp140-RGG localized to pole cell nuclei (white arrow), but the protein remained somewhat diVuse within these nuclei. Pole cell nucleoli form after gastrulation begins (Mahowald, 1963). Bar = 40 mm. (See also color insert.)

prenucleolar-like bodies containing GFP-Nopp140-RGG (Fig. 3B), perhaps in keeping with the findings of Dundr et al. (1997) for Nopp140 during nucleologenesis. It is clear from this preliminary observation that there are at least two programs for nucleologenesis in the Drosophila embryo, one for the peripheral blastoderm cells and another for the posterior pole cells. We assume that nucleologenesis in the pole cells is delayed because of a programmed lag in pre-rRNA synthesis at the nucleolar organizers. If we assume that rDNA expression initiates nucleologenesis, what regulates the rapid onset of rDNA transcription in the blastoderm cells but a delay in the pole cells? Could cytoplasmic determinants in the posterior egg cytoplasm (e.g., the germ plasm or polar granules; Mahowald, 2001) directly or indirectly block rDNA transcription within the pole cells during the blastoderm stage? Further, how are nucleolar components stored in the pole cells before rDNA transcription? Drosophila should be amenable to decipher the dynamics of nucleologenesis during embryonic development.

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B. Xenopus Embryogenesis Meßmer and Dreyer (1993) followed the two isoforms of Xenopus laevis nucleolin from oocyte maturation on through to gastrulation. Progesterone induces maturation of the prophase I–arrested oocyte. On induction, Maturation Promoting Factor (MPF or Cdk1/cyclin B) activity rises sharply leading to vesiculation of the nuclear envelope and the disassembly of the extrachromosomal multiple nucleoli. The nucleoli disassemble down to their fibrillar centers (Shah et al., 1996; P. DiMario, unpublished observations). Parenthetically, the rDNA within these fibrillar centers persists within unfertilized eggs (Brown, 1966) and remains detectable until gastrulation (Brown and Weber, 1968). The hyperphosphorylation of several T/SPAKK motifs within the amino-terminal third of nucleolin is closely correlated with nucleolar disassembly. MPF-phosphorylated nucleolin redistributes to the egg cytoplasm as the nucleoli disassemble, but this hyperphosphorylation by MPF may not be the driving mechanism for the observed redistribution (Zhu et al., 1999). The fertilized Xenopus egg undergoes 12 synchronous rounds of mitosis to reach the midblastula transition (MBT). These early cleavages are very rapid; the cell cycles lack G1 and G2 phases, thus precluding transcription by the three classes of RNA polymerase (Brown and Littna, 1964; Hay and Gurdon, 1967; Wallace, 1963). During the rapid cleavages, nucleolin remains cytoplasmic. G1 and G2 phases appear at the midblastula transition—the point at which cell cycles slow down and transcription begins. Nucleolin begins to translocate from the cytoplasm to the nucleolus at MBT. Because nucleoli do not appear until the late blastula/early gastrula stage (De Capoa et al., 1983), maternal nucleolin that enters the nucleus at MBT accumulates within prenucleolar bodies (Meßmer and Dreyer, 1993). Bell et al. (1997) used immuno-fluorescence microscopy to localize UBF, TBP, and Pol I during the assembly of pronuclei from sperm chromatin added to Xenopus egg extracts and during embryogenesis following fertilization. UBF consistently associated with the rDNA (NORs) in the extract-assembled nuclei and in the embryonic nuclei. The observations indicate that UBF associates with the NORs from the earliest pronuclear stage and onward through embryogenesis. Interestingly, TBP and Pol I failed to associate with UBF in the silent NORs of the pronuclei and the rapidly cleaving embryonic blastomere nuclei. TBP and Pol I first associated with the NORs at the MBT. Bell et al. (1997) suggest that UBF forms an architecturally open chromatin complex with the rDNA that is then recognized by TBP and Pol I only at the MBT, thus creating a transcriptional-competent complex. Verheggen et al. (1998, 2000) argued that partial nucleolar formation in Xenopus embryos can occur in the absence of Pol I activity. In their 1998


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paper, Verheggen et al. used run-on transcription assays with Br-UTP followed by immuno-fluorescence localization of the newly synthesized RNA. Using Act D (inhibition RNA Pol I), a-amanitin (inhibition of RNA Pol II and III inhibition), and aphidocolin (inhibition of DNA Pols a, d for chromosome replication, and for DNA repair) to determine what classes of polymerases were active at particular times in development, the authors showed that RNA Pol II and III initiated transcription in a small percentage of embryonic nuclei at 7 hours postfertilization (the beginning of midblastula transition, MBT), but that 75% of the nuclei had initiated Pol II and III transcription by 9 hours postfertilization (end of MBT). Conversely, RNA Pol I first started transcription at 9 hours postfertilization in 1–2% of embryonic nuclei. Pol I transcription was detectable in 40% of the nuclei by 11 hours, and in 90% of the nuclei by 12 hours postfertilization. The striking observation made by Verheggen et al. (1998) was that fibrillarin and nucleolin colocalized to ‘‘nucleolar domains’’ that formed during MBT (7–9 hours postfertilization). These nucleolar domains were detected by immunofluorescence microscopy in the absence of pre-rRNA synthesis (e.g., a complete lack of Br-UTP labeling). Their observation suggests that in Xenopus embryos, preliminary nucleolar assembly at NORs can occur in the absence of pre-rRNA synthesis (Me´le`se and Xue, 1995). The nucleolar domains that formed during MBT, however, did contained unprocessed pre-rRNA of maternal origin, as detected by fluorescent in situ hybridization using 50 ETS and ITS1 probes. Thus, nucleologenesis may be dependent on the synthesis of nascent pre-rRNA at the NOR or the delivery of maternal pre-rRNA to the NORs by previously assembled PNBs. Verheggen et al. (2000) then established the existence of two classes of PNBs; the first class (PNBs I) contains the early pre-rRNA processing components fibrillarin, U3, and U8, whereas PNBs II contain the later processing component B23 (called NO38 in Xenopus). In all likelihood, PNBs II may derive from PNBs I as early processing components transfer to the nascent nucleoli, leaving late processing components behind. Verheggen et al. (2000) showed that fibrillarin (PNBs I) and NO38 (PNBs II) began to fuse to nucleolar domains, but with diVerent rates. By 9 hours, PNBs I had apparently fused to the NORs, and a large number of NO38containing PNBs II remained in the nucleoplasm until 11 hours postfertilization. An important observation made by Verheggen et al. (2000) was that PNBs could fuse to nucleolar domains in the absence of rDNA transcription. In 9- or 12-hour embryos treated with Act D (0.5 mg/mL), fibrillarin-containing PNBs I could still fuse to nucleolar domains, but these accumulations did not correspond to NORs that were silenced by the Act D. The only transcription sites in these Act D–treated cells (as detected by Br-UTP incorporation/anti-Br-UTP staining) were inhibited by a-amanitin; thus, only Pol II or Pol III were active. No Pol I transcription was detected in

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the Act D–treated cells, despite the fact that fibrillarin-containing PNBs I could still fuse to nucleolar domains. The possibility that any pre-rRNA synthesis before Act D treatment may initiate PNBs I recruitment was reduced by the observation that most Pol I remains scattered within nuclear speckles at 9 hours postfertilization. In a related Xenopus erythrocyte model for embryonic nucleologenesis, Verheggen et al. (2001) addressed the hypothesis that rDNA transcription is required for full nucleolar formation. The Xenopus erythrocyte nucleus is transcriptionally silent; its chromatin is condensed, and the nucleolus is reduced to a persistent remnant. At the ultrastructural level, this nucleolar remnant maintains a ring-shaped fibrillar region with a central clear zone. A few RNP granules lie on the periphery of the fibrillar region. Processing components fibrillarin, nucleolin, B23/NO38, U3, and U8 were all represented within the remnants, along with some partially processed pre-rRNA containing ITS1, but apparently not the 50 ETS. An electrophoretic isoform of UBF (64–67 kDa instead of 82–85 kDa) localized to a single spot within the remnant. Pol I, however, was not detected within the remnant nucleoli by immuno-fluorescence microscopy, and the erythrocyte nucleoli failed to incorporate Br-UTP during in situ run-on assays. These observations indicate that portions of the rDNA transcriptional machinery (UBF and the rDNA), and at least some rRNA processing components, can maintain their association in the absence of rDNA transcription. Although the precise mechanism or mechanisms for maintaining the associations remain unknown, Verheggen et al. (2001) suggest that the partially processed pre-rRNA could be a critical link in maintaining these associations. In eVorts to mimic the onset of nucleologenesis in embryonic cells, Verheggen et al. placed erythrocyte nuclei into Xenopus egg extracts. After 1 hour at 23 C, 80% of the nuclei could partially decondense their chromatin. Only 10% of the nuclei could fully decondense their chromatin. In nuclei exhibiting fully decondensed chromatin, UBF maintained its close proximity to rDNA, but it redistributed from the one spot to several smaller foci. Even though chromatin had greatly decondensed, these nuclei were incapable of transcription. Extra amounts of the processing components fibrillarin, nucleolin, U3, and U8 within the egg extract colocalized to large, dense PNBs within these nuclei, whereas imported B23/NO38 localized to distinct smaller nuclear dots. All examined processing components failed to associate with the rDNA. The observations indicate that although quiescent rDNA could remodel as determined by the multiple UBF-containing foci, and nascent PNBs could form within the erythrocyte nuclei, intact nucleoli failed to form in the absence of rDNA transcription. With UBF present, the authors reasoned that other important transcription factors were either missing or incapable of supporting transcription, and as described later, their reasoning seems to be correct.


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Bell and Scheer (1999) showed that Xenopus eggs contain high levels of maternally derived Pol I. This Pol I could be easily detected within in vitro– assembled pronuclei that form in egg extracts on the addition of sperm chromatin. Failure to detect Pol I in early blastula nuclei, however, may be a result of the failure rapidly cleaving nuclei to adequately form nuclear pore complexes that are necessary for the translocation of Pol I from the cytoplasm. As a result Pol I may never reach a detectable concentration within rapidly cleaving blastula nuclei. With the appearance of G1 and G2, however, Pol I localizes to nuclear speckles that exist from the midblastula stage to the neurula stage. These speckles are distinct from prenucleolar bodies because the processing proteins nucleolin, fibrillarin, and Nopp180 fail to colocalize with Pol I in these speckles. Portions of this nuclear Pol I begin to associate with the forming nucleoli in the late blastula/early gastrula stages. TBF (a component of SL1) was hardly detectable in the oocyte, but its abundance increased steadily to the gastrula stage—the point at which nucleoli first appear. Before the onset of rDNA transcription, TBP colocalized with Pol I in the nuclear speckles. The low abundance of TBP and the mostly compact nature of chromatin during rapid blastomere cleavages may prevent the formation of productive transcription complexes on rDNA promoters. The pulse of TBP production at late blastula/early gastrula may reflect a need to activate virgin rDNA genes that never before transcribed. Later, at the tail bud stage, TBP returned to low concentrations typical of somatic cells. UBF, however, maintained its NOR localization at all stages of embryogenesis. Ribosomal DNA transcription activates at late blastula/gastrula, perhaps because of the lengthened cell cycles that in turn allow NOR chromatin to decondense, and because of the association of transiently abundant TBP with the UBF-bound rDNA promoters. At this point, Pol I gradually redistributes form the nuclear speckles to the forming nucleoli.

C. Nucleologenesis in Mammalian Embryos Nucleologenesis in mammalian embryos does not initiate immediately after fertilization, but rather begins at diVerent points of early cleavage in a speciesspecific manner (Baran et al., 1996, and references therein). Much of the cytochemical (silver staining) and immuno-cytochemical work on mammalian embryonic nucleologenesis has been conducted in cow (Baran et al., 1996; Kopec˘ny´ et al., 2000; Laurincik et al., 2002), pig (Hyttel et al., 2000; Kopec˘ny´ et al., 1996a; Laurincik et al., 1995), mouse (Baran et al., 1995; Biggiogera et al., 1994; Zatsepina et al., 2003), and rabbit embryos (Baran et al., 1997; Kan˘ka et al., 1996) where nucleolus precursor bodies (NPBs, see Flećhon and Kopec˘ny´, 1998) exist in the preimplantation embryos. On the basis of their

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molecular composition, NPBs have been categorized into two classes, the bovine-type and mouse-type. NPBs in pronuclei, two-cell, and four-cell and bovine (and goat) embryos are small, electron-dense spheres containing tightly packed fibrils. An important distinguishing feature for this class of mammalian NPBs (as opposed to others, see below) is the presence of silver-staining proteins throughout the bodies. Kopec˘ny´ et al. (1996b, 2000) described these NPBs in two-cell bovine embryos produced either in vivo or by in vitro techniques. Ethidium bromide– phosphotungstic acid staining indicated the presence of nucleic acids (RNA) within these bodies. In addition to NPBs in two-cell embryos, another class of nuclear body (the ‘‘loose bodies’’) contained snRNAs and is closely related to somatic cell interchromatin granules. A third class of nuclear bodies in these embryos, the dense bodies, appeared similar to NPBs. Although dense bodies contained Sm-antigens, which are absent in NPBs, the dense bodies lacked B23, which is present in NPBs. All three types of nuclear bodies could exist within the same nucleus, often in close proximity to each other. Nucleologenesis in bovine embryos begins primarily in the eight-cell stage (the fourth cell cycle), with the preformed NPBs gradually interacting with rDNA as it resumes Pol I transcription. By the eight-cell stage, the smaller NPBs observed in the two-cell stage have fused into a larger NPB. Bovine embryonic nucleologenesis during the eight-cell stage has been divided into several steps based on the NPB morphology (Kopec˘ny´, 1990; Kopec˘ny´ et al., 1989). Baran et al. (1996) showed that in the first step, compact NPBs make contact with closely residing rDNA-containing chromatin, after which the rDNA penetrates into the NPB (Kopec˘ny´ et al., 1989). Apparently, the rDNA infiltrates deeply into the NPBs to initiate rRNA transcription. In the second step, NPBs form a large central vacuole and accumulate nucleolin into compact fibrillar networks. Similar to delayed associations of B23 during somatic postmitotic nucleologenesis, B23 joins the fibrillar component of the NPB in the third step, after the initial localization of nucleolin. Also in the third step, bovine NPBs acquire secondary vacuoles that are surrounded by the fibrillar component, and the NPBs display rRNA for the first time in clusters in the dense fibrillar regions. In the fourth and final step, fully reticulated nucleoli contain typical DFCs with rRNA and processing proteins (Baran et al., 1996) encircling a fibrillar center, along with a peripheral granular component that expanded significantly in the 16-cell stage (Kopec˘ny´ et al., 1989). Mouse-type NPBs include those found also in rat, hamster, rabbit, pig, and human embryos. These NPBs appear early in the pronucleus as very prominent nuclear bodies (unlike the bovine-type NPBs, which are rather inconspicuous). Like the bovine-type NPBs, mouse-type NPBs consist of a dense fibrillar material, but argyrophilic proteins localize only to the periphery of the mouse-type NPB, and not within it (Flećhon and Kopec˘ny´, 1998).


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Mouse-type NPBs do not contain rDNA, but they do contain processing components U3, fibrillarin, nucleolin, and B23. Embryonic nucleologenesis in mouse begins in the two-cell stage (instead of the eight-cell stage, as in bovine embryos). Interestingly, Zatsepina et al. (2000) showed that UBF and Pol I dissociate from the chromosomes during meiosis I of mouse oogenesis, and that the two proteins are undetectable by metaphase II, perhaps having degraded. New protein synthesis begins 4–5 hours before initiation of rDNA transcription. Zatsepina et al. (2003) showed that new UBF and Pol I first transit through Cajal bodies (Gall, 2000, 2003) before their assembly on the NORs. Transcription begins 4–5 hours later, during the second half of the two-cell stage. Reactivated NOR chromatin does not infiltrate deeply into core of the mouse-type NPBs; thus, nucleolar fibrillar centers and transcription sites form initially at the periphery of some NPBs. The NPB core remains enigmatic. Although this core does not stain with silver, a few nucleolar proteins may localize to it. A few reports suggested that the core may contain maternal spliceosomal snRNPs (Flećhon and Kopec˘ny´, 1998), suggesting the core may be similar to Cajal bodies (Gall et al., 1995) or sphere organelles of amphibian oocytes (Gall and Callan, 1989). Nucleolonema containing nucleolin and B23 eventually form on the NPB periphery in four-cell mouse embryos (Baran et al., 1995). Typical postmitotic mechanisms for nucleologenesis are restored by the eight-cell stage of mouse embryogenesis (Zatsepina et al., 2003).

VI. Nucleolar Disassembly at Mitosis A careful review of the entire nucleolar cycle is necessary to best understand nucleologenesis. Here we review nucleolar disassembly during prophase. This aspect of the nucleolar cycle has received relatively little experimental attention beyond the level of rDNA transcription. Although most would agree that nucleolar disassembly is primarily caused by the loss of pre-rRNA production, important questions remain concerning the role that mitotic phosphorylation of several processing components may have in the disassembly process (Belenguer et al., 1990; Gulli et al., 1997; Peter et al., 1990a). We know that Act D inhibition of Pol I transcription will cause a redistribution of processing component to the nucleoplasmin of interphase cells (Scheer et al., 1975), but what function does Cdk1/cyclin B phosphorylation of nucleolin, B23, and perhaps other pre-rRNA processing proteins have in their redistribution to the mitotic cytoplasm? One can easily imagine that M-phase phosphorylation may facilitate nucleolar disassembly and that reversing this phosphorylation during anaphase and telophase will play a role in PNB and NDF assembly, and ultimately in the reassembly of the nucleolus. Such a paradigm is found

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for the nuclear lamina (Heald and McKeon, 1990), but we need to be cautious with this apparently simplistic assumption.

A. Mitotic Phosphorylation of Nucleolar Transcription Factors The onset of mitosis is marked by the rapid rise in Cdk1/cyclin B kinase activity and the M-phase phosphorylation of many diVerent substrates (Basi and Draetta, 1995). M-phase phosphorylation of transcription factors is believed to be the driving mechanism for mitotic repression of chromosome transcription. For example, Kuhn et al. (1998) suggested that mitotic repression of transcription could result from the loss of transcription factors from gene promoters, modification of the chromatin template, or chromosome condensation. We already described above the molecular architecture of the mitotic NOR, and how UBF, Pol I, SL1, and Topo I remain associated with the mitotic NOR. To address chromatin condensation, Hartl et al. (1993) showed the in vitro repression of 5S and tRNA genes (Pol III transcription) using mitotic cell extracts, and they were able to maintain an open chromatin configuration by using VM-26, an inhibitor of topoisomerase II that blocks chromatin condensation. Their results suggest that chromatin compaction is not required for mitotic repression of at least the 5S gene, at least in vitro. Gottesfeld et al. (1994) then supplemented extracts from asynchronous cells with Cdk1/cyclin B, and again showed a repression in Pol III transcription, but they carried the work farther to show that TFIIIB was phosphorylated by the kinase. Later studies verified the findings that mitotic extracts can silence Pol II and III transcription by phosphorylating the TBP, a component of TFIID and TFIIIB (Leresche et al., 1996; White et al., 1995). With respect to Pol I transcription, Kuhn et al. (1998) used an elaborate add-back transcription rescue assay to show that only exogenous, unphosphorylated TIF-IB/SL1 (a complex containing TBP and TAF1 110) could rescue rDNA transcription in a mitotic extract in which endogenous factors had been previously inactivated by thio-phosphorylation and normal kinase activities were blocked by 6-dimethyl-aminopurine (DMAP) and the nonhydrolyzable ATP analogue, adenylyl-imidodiphosphate (AMP-PNP). None of the other unphosphorylated transcription factors (Pol I itself, UBF, TIF-IA, or TIF-IC) could rescue transcription when added to the extract. Heix et al. (1998) also showed that the phosphorylation of SL1 repressed in vitro Pol I transcription, and that Cdk1/cyclin B phosphorylated both TBP and hTAFI110. They showed, however, that it was the phosphorylation of TAFI110 rather than TBP that blocked mitotic transcription. Heix et al. (1998) further showed that normal protein–protein interactions between the SL1 complex and UBF


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were impaired by mitotic phosphorylation of SL1. Their results indicate that mitotic phosphorylation disrupts the preinitiation complex (i.e., SL1 interacting with UBF), thus blocking Pol I transcription of rDNA at the onset of mitosis. Blocking this interaction occurs even though SL1 and UBF remain associated with the NOR. Sirri et al. (2000) actually restored rDNA transcription in colchicine-arrested or taxol-treated mitotic HeLa cells. Pol I transcription was restored in these arrested cells by blocking Cdk1/cyclin B activity with the drug roscovitine, a highly selective inhibitor of cyclin-dependent kinases (e.g., Sirri et al., 2000). Pol I transcription resumed at the NOR during prometaphase, metaphase, and anaphase on Cdk1/cyclin B inhibition by roscovitine: In situ BrUTP labeling during these stages colocalized with anti-UBF labeling. Further, unprocessed 47S pre-rRNA accumulated in the roscovitine-treated mitotic cells, and its synthesis was sensitive to the low doses (0.05 mg/mL) of Act D typically used to block Pol I transcription. Sirri et al. (2000) then showed that the mitotic phosphorylation of TBP, TAFI110, and TTF-1 was largely reversed on roscovitine treatment, indicating that phosphatases are constantly at work during mitosis, attempting to dephosphorylate the Pol I transcription factors. Indeed, roscovitine-induced resumption in Pol I transcription could be blocked by okadaic acid, an inhibitor of protein phosphatase 1 (PP1). Sirri et al. (2000) suggested that isoform PP1-g, which remains associated with mitotic chromosomes, may be responsible for the reactivation of SL1. Interestingly, the newly synthesized 47S pre-rRNA in these roscovitine-treated cells was not processed—it only accumulated. This observation indicates that restoration of rDNA transcription and pre-rRNA processing in telophase/early G1 may be under separate controls (see following).

B. Mitotic Phosphorylation of Pre-rRNA Processing Components In addition to the transcription factor SL1, pre-rRNA processing components are also physiological substrates for Cdk1/cyclin B. These include nucleolin and B23 (Belenguer et al., 1990; Peter et al., 1990a). Perhaps their mitosis-specific phosphorylation contributes to the dispersion of the nucleolus during prophase, analogous to what happens to the nuclear lamina when its lamin proteins are similarly phosphorylated (Peter et al., 1990b). To test this, we prepared a modified form of Chinese hamster nucleolin in which eight of nine Cdk1/cyclin B phosphorylation sites (–TPXKK–) within the amino terminus were substituted (–NTXKK–) by site-directed mutagenesis (the remaining site was simply lost during DNA manipulations). The mutated protein was not a substrate for in vitro phosphorylation by cdk1/cyclin B, but we observed no adverse dominant-negative eVects on nucleolar disassembly

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or reassembly when we overexpressed the mutant in diVerent mammalian cell types. The mutant protein redistributed normally from nucleoli to the cytoplasm during prophase and metaphase, and it localized to PNBs and the reforming nucleoli during telophase. Overexpression prevented us from adequately investigating the perichromosomal regions. Because the mutant protein localized properly, we argued that mitotic phosphorylation of nucleolin is not necessary for its correct mitotic redistribution, but rather for its normal interphase function in pre-rRNA processing. Further eVorts should define the eVects of M-phase phosphorylation of pre-rRNA processing components. We described above how Sirri et al. (2000) reactivated rDNA transcription in colchicine-arrested mitotic HeLa cells by blocking cdk1/cyclin B with roscovitine. The newly synthesized 47S transcript, however, accumulated within these cells without being processed. This observation indicated that block in cdk1/cyclin B activity at the end of mitosis is not suYcient to reassemble pre-rRNA processing components in nucleologenesis, and that separate mechanisms must regulate reactivation of rDNA transcription versus pre-rRNA processing. To further address this hypothesis, Sirri et al. (2002) again used roscovitine to block general cdk activity, but now at the M/G1 transition. Control experiments showed that metaphase-synchronized cells treated with roscovitine could proceed through the M/G1 transition, but these cells failed to form nucleoli. Fibrillarin could still relocalize to the newly activated sites of rDNA transcription in roscovitine-treated cells, as in nontreated control cells, but another processing protein, Nop52, could not. Again, the 46S pre-rRNA accumulated without being processed in these roscovitine-treated, M/G1-transitioning cells. Not surprisingly, roscovitine (and other cdk inhibitors) dramatically disrupted the structural integrity of nucleoli in interphase cells, and the compounds partially blocked both rDNA transcription and rRNA processing (46S accumulated while 45S matured) during interphase. From their observations and those of Dousset et al. (2000), Sirri et al. (2002) argued that the formation of nucleoli at the end of mitosis is not governed solely by the reactivation of rDNA transcription, and that normal nucleolar structures and functions are maintained by interphase-specific cdk activities.

C. Persistent Nucleoli Nucleoli that persist well into mitosis have been noted in several cell types (Sheldon et al., 1981). Examples include Chinese hamster ovary and Dede cells (Brinkley, 1965; Hsu et al., 1965; Noel et al., 1971) and various transformed cell lines including HeLa and Erlich ascites (Guttes et al., 1968). Why these nucleolar remnants persist later into prophase remains unknown.


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Sheldon et al. (1981) demonstrated persistent nucleoli in five mouse embryonal carcinoma cell lines. Sixty to 88% of the cells maintained nucleolar remnants into metaphase and anaphase. (Nucleoli remaining in prometaphase were not considered persistent.) True persistent nucleoli maintained contact with the metaphase and anaphase chromosomes in the 99% of the cells that displayed persistent nucleoli. When these cells diVerentiated, the percentage of cells displaying persistent nucleoli dropped to below 30%. Pulse labeling with tritiated uridine followed by autoradiography showed that persistent nucleoli in metaphase had half the number of silver grains as interphase nucleoli and twice as many grains as metaphase cells without persistent nucleoli. The incorporation of tritiated uridine in the persistent nucleoli was sensitive to Act D (0.1–1.0 mg/mL). The result strongly indicate that continued (albeit reduced) rRNA synthesis is responsible for persistent nucleoli in metaphase. The question remains, How do these cells maintain active Pol I transcription so deep into mitosis? Clearly, the inhibitory eVects of Cdk1/cyclin B phosphorylation on Pol I transcription factors should be the first focus of investigation.

D. Coming Full Circle: Reactivation of Pol I Transcription Klein and Grummt (1999) showed that UBF is also phosphorylated by cdk1/ cyclin B during mitosis, and that its progressive reactivation (dephosphorylation) during early G1 is tightly linked to the progressive reactivation of Pol I transcription during this same period. The postmitotic reactivation of UBF was slower than that for SL1, and a number of possibilities could explain this diVerence. Because UBF reactivation proved to be sensitive to okadaic acid, Klein and Grummt (1999) favored the possibility that two separate phosphatases may remove M-phase phosphorylation at diVerent rates to reactivate SL1 and UBF. They suggested that protein phosphatase 2A (PP2A) may be responsible for this postmitotic reactivation of UBF. Grummt’s laboratory (Voit et al., 1999) further showed that the upregulation of UBF in serum-stimulated NIH 3T3 cells was dependent on its phosphorylation by G1-specific cdk4/cyclin D and cdk2/cyclin E complexes. This phosphorylation occurred on Ser484, a conserved residue. Using an in vitro UBF-responsive reconstitution transcription assay, Voit et al. (1999) showed that transcription was strongly reduced in the presence of the UBF1 mutant, S484A. Their observations indicate that increasing rates of Pol I transcription in G1 closely correlate with the phosphorylation of UBF at S484, and that this phosphorylation is the result of the appearance of G1-specific cdk-cyclin complexes. UBF is phosphorylated at multiple sites, and the timing of these phosphorylation events is closely coupled to cell proliferation and rDNA transcription. Hannan et al. (2000a,b) emphasized

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this link between UBF function and cell proliferation by showing hypophosphorylated retinoblastoma protein redistributed to nucleoli as culture cells reached confluence and their rDNA transcription diminished significantly. Hannan et al. (2000a) showed that the hypophosphorylated form of Rb inhibited rDNA transcription by directly binding UBF. The interaction between Rb (or p130) and UBF does not dislodge UBF from the rDNA but, rather, blocks the normal interaction between UBF and SL1 (Hannan et al., 2000b). Unfortunately, we do not have the space to elaborate on these important (and very interesting) mechanisms of interphase-specific regulation of rDNA transcription. The reader is referred to Grummt (1999) and Hannan et al. (2000a,b) for a complete description of this topic.

VII. Perspectives In 1970, Busch and Smetana stated that the mechanisms of nucleolar disassembly and reassembly were largely unknown. Since then, we have made tremendous strides in our understanding of nucleologenesis as described in the preceding sections. Many significant questions still remain regarding nucleologenesis, but an equally fascinating path of discovery lies before us in understanding the disassembly process in prophase and how these events lead to assembly in telophase. For example, it is reasonably well established that mitotic phosphorylation of Pol I transcription factors block their function, resulting in the shut-down of rDNA transcription. A loss of newly produced pre-rRNA may then disband processing components to the nucleoplasm and the cytoplasm. However, that leaves us with the question of why are processing components (e.g., nucleolin, B23) phosphorylated during mitosis? We can easily speculate that mitotic phosphorylation disrupts interphase-specific associations, and perhaps establishes mitosis-specific associations. But what are these associations? Could reorganization of processing components from interphase- to mitosis-specific associations be linked to the formation of the perichromosomal sheath? Alternatively, could reversing mitotic phosphorylation of processing components trigger the disassembly of the perichromosomal sheath and initiate assembly of NDF and PNBs? The perichromosomal sheath remains poorly understood in terms of its formation, overall composition, and possible functions. What attracts nucleolar processing components to the surface of mitotic chromosomes? Are the mechanisms that condense chromosomes directly or indirectly related to accumulating nucleolar components as a surface coat? In terms of function, perhaps the sheath insulates the chromosomes from the cytoplasm after the nuclear envelope disassembles. Perhaps it serves simply to


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carry nucleolar components into daughter nuclei. It is interesting to note that a perichromosomal sheath is lacking in yeast, which display a closed mitosis. Thus, an insulator function seems more plausible than a carrier function. In her 1934 paper, McClintock speculated that the coating on mitotic chromosomes may play a functional role. After all these years, we have yet to assign a clear function to the perichromosomal sheath. What mitotic events block the further processing of pre-rRNA and ribosome assembly during nucleolar breakdown? Is it possible that partially processed residual pre-rRNA functions to maintain the integrity of the perichromosomal sheath, PNBs, and NDF, as suggested by the work of Pinõl-Roma (1999)? How NDF and PNBs release their components to the reactivated NORs is also incompletely understood. For example, what initiates NDF and PNB disassembly? Could reversing a mitotic phosphorylation allow processing proteins (e.g., nucleolin and B23) and their associated snoRNAs to rapidly diVuse from the PNBs to the NORs? What regulatory signals or events link resumption of rDNA transcription (the initiation of nucleologenesis) to the return of pre-rRNA processing components to the NORs? Perhaps as one possibility, postmitotic dephosphorylation of UBF and SL1 triggers the recruitment of processing components. Alternatively, the reappearance of the 50 ETS leader at the NORs may signal an initial coalescence of processing components, with a full-fledged recruitment taking place once more downstream regions of the pre-rRNA become available. The two possibilities are not exclusive of one another. In past years, molecular techniques have identified the many snoRNAs. More recently, proteomic techniques (mass spectroscopy) have identified the vast array of nucleolar proteins (Andersen et al., 2002; Scherl et al., 2002). These many snoRNAs and proteins are the pieces of a very complicated structural and functional puzzle. Simply having the pieces before us has been a monumental step forward in deciphering the dynamics of the nucleolus. Our future challenge, however, is to put the pieces together in terms of functional associations that in their entirety describe conventional and nonconventional functions of the nucleolus (Olson et al., 2002). As daunting as this challenge may seem, knowing how these nucleolar macromolecules reassociate during nucleologenesis will certainly shed light on the many nucleolar functions during interphase.

Acknowledgments I thank Mark Olson at the University of Mississippi Medical Center, Jackson, Mississippi, for his insightful comments on the manuscript. National Science Foundation grant MCB-0234245 supported our work on Drosophila nucleologenesis, first presented in this review.

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References Abramova, N. B., and Neyfakh, A. A. (1973). Migration of newly synthesized RNA during mitosis. III. Nuclear RNA in the cytoplasm of metaphase cells. Exp. Cell Res. 77, 136–142. Andersen, J. S., Lyon, C. E., Fox, A. H., Leung, A. K. L., Lam, Y. W., Steen, H., Mann, M., and Lamond, A. I. (2002). Directed proteomic analysis of the human nucleolus. Curr. Biol. 12, 1–11. Azum-Ge´lade, M. C., Noaillac-Depeyre, J., Caizergues-Ferrer, M., and Gas, N. (1994). Cell cycle redistribution of U3 snRNA and fibrillarin. Presence in the cytoplasmic nucleolus remnant and in the prenucleolar bodies at telophase. J. Cell Sci. 107, 463–475. Bachellerie, J. P., Cavaille, J., and Huttenhofer, A. (2002). The expanding snoRNA world. Biochemie 84, 775–790. Baran, V., Vesela´, J., Reha´k, P., Koppel, J., and Flećhon, J.-E. (1995). Localization of fibrillarin and nucleolin in nucleoli of mouse preimplantation embryos. Mol. Reprod. Dev. 40, 305–310. Baran, V., Flećhon, J.-E., and Pivko, J. (1996). Nucleologenesis in the cleaving bovine embryo: Immunocytochemical aspects. Mol. Reprod. Dev. 44, 63–70. Baran, V., Mercier, Y., Renard, J.-P., and Flećhon, J.-E. (1997). Nucleolar substructures of rabbit cleaving embryos: An immunocytochemical study. Mol. Reprod. Dev. 48, 34–44. Basi, G., and Draetta, G. (1995). The cdc2 kinase: Structure, activation, and its role at mitosis in vertebrate cells. In ‘‘The Cell Cycle’’ (C. Hutchison and D. M. Glover, Eds.), pp. 106–143. IRL Press/Oxford University Press, Oxford. Belenguer, P., Caizergues-Ferrer, M., Labbe, J.-C., Doreé, M., and Amalric, F. (1990). Mitosisspecific phosphorylation of nucleolin by p34cdc2 protein kinase. Mol. Cell. Biol. 10, 3607–3618. Bell, P., and Scheer, U. (1996). Prenucleolar bodies contain coilin and are assembled in Xenopus egg extract depleted of specific nucleolar proteins and U3 RNA. J. Cell Sci. 109, 43–54. Bell, P., and Scheer, U. (1999). Developmental changes in RNA polymerase I and TATA boxbinding protein during early Xenopus embryogenesis. Exp. Cell Res. 248, 122–135. Bell, P., Dabauvalle, M.-C., and Scheer, U. (1992). In vitro assembly of prenucleolar bodies in Xenopus egg extract. J. Cell Biol. 118, 1297–1304. Bell, P., Mais, C., McStay, B., and Scheer, U. (1997). Association of the nucleolar transcription factor UBF with the transcriptionally inactive rRNA genes of pronuclei and early Xenopus embryos. J. Cell Sci. 110, 2053–2063. Benavente, R., Rose, K. M., Reimer, G., Hu¨gle-Do¨rr, B., and Scheer, U. (1987). Inhibition of nucleolar reformation after microinjection of antibodies to RNA polymerase I into mitotic cells. J. Cell Biol. 105, 1483–1491. Bernhard, W., and Granboulan, N. (1968). Electron microscopy of the nucleolus in vertebrate cells. In ‘‘The Nucleus’’ (A. J. Dalton and F. Haguenau, Eds.), Vol. 3, pp. 81–149. Academic Press, New York. Bertrand, E., Houser-Scott, F., Kendall, A., Singer, R. H., and Engelke, D. R. (1998). Nucleolar localization of early tRNA processing. Genes Dev. 12, 2463–2468. Beven, A. F., Lee, R., Razaz, M., Leader, D. J., Brown, J. W. S., and Shaw, P. J. (1996). The organization of ribosomal RNA processing correlates with the distribution of nucleolar snRNAs. J. Cell Sci. 109, 1241–1251. Biggiogera, M., Malatesta, M., Abolhassani-Dadras, S., Amalric, F., Rothblum, L. I., and Fakan, S. (2001). Revealing the unseen: The organizer region of the nucleolus. J. Cell Sci. 114, 3199–3205. Biggiogera, M., Martin, T. E., Gordon, J., Amalric, F., and Fakan, S. (1994). Physiological inactive nucleoli contain nucleoplasmic ribonucleoproteins: Immunoelectron microscopy of mouse spermatids and early embryos. Exp. Cell Res. 213, 55–63.


161

Blum, B., Pierron, G., Seebeck, T., and Braun, R. (1986). Processing in the external transcribed spacer of ribosomal RNA from Physarum polycephalum. Nucl. Acids. Res. 14, 3153–3166. Borovjagin, A. V., and Gerbi, S. A. (1999). U3 small nucleolar RNA is essential for cleavage at sites 1, 2 and 3 in pre-rRNA and determines which rRNA processing pathway is taken in Xenopus oocytes. J. Mol. Biol. 286, 1347–1363. Bouvet, P., Diaz, J. J., Kleinbeiter, K., Madjar, J. J., and Amalric, F. (1998). Nucleolin interacts with several ribosomal proteins through its RGG domain. J. Biol. Chem. 273, 19025–19029. Brill, S. J., DiNardo, S., Voelker-Meiman, K., and Sternglanz, R. (1987). Need for DNA topoisomerase activity as swivel for DNA replication and for transcription of ribosomal genes. Nature 326, 414–416. Brinkley, B. R. (1965). The fine structure of the nucleolus in mitotic divisions of Chinese hamster cells in vitro. J. Cell Biol. 27, 411–422. Brown, D. D. (1966). The nucleolus and synthesis of ribosomal RNA during oogenesis and embryogenesis of Xenopus laevis. Nat. Cancer Inst. Mono. 23, 297–309. Brown, D. D. (1967). The genes for ribosomal RNA and their transcription during amphibian development. Curr. Top. Dev. Biol. 2, 47–73. Brown, D. D., and Gurdon, J. B. (1964). Absence of ribosomal RNA synthesis in the anucleolate mutant of Xenopus laevis. Proc. Natl. Acad. Sci. USA 51, 139–146. Brown, D. D., and Littna, E. (1964). RNA synthesis during the development of Xenopus laevis, the South African clawed toad. J. Mol. Biol. 8, 669–687. Brown, D. D., and Weber, C. S. (1968). Gene linkage by RNA-DNA hybridization. II. Arrangement of the redundant gene sequences for 28S and 18S ribosomal RNA. J. Mol. Biol. 34, 681–697. Bryk, M., Banerjee, M., Murphy, M., Knudsen, K. E., Garfinkel, D. J., and Curcio, M. J. (1997). Transcriptional silencing of Ty1 elements in the RD1 locus of yeast. Genes Dev. 11, 255–269. Burd, C. G., and Dreyfuss, G. (1994). Conserved structures and diversity of functions of RNAbinding proteins. Science 265, 615–621. Busch, H., and Smetana, K. (1970). ‘‘The Nucleolus.’’ Academic Press, New York. Cairns, C., and McStay, B. (1995). Identification and cDNA cloning of a Xenopus nucleolar phosphoprotein, xNopp180, that is the homolog of the rat nucleolar protein Nopp140. J. Cell Sci. 108, 3339–3347. Cavanaugh, A. H., Hempel, W. M., Taylor, L. J., Rogalsky, V., Todorov, G., and Rothblum, L. I. (1995). Activity of RNA polymerase I transcription factor UBF blocked by Rb gene product. Nature 374, 177–180. Chan, E. K. L., Imai, H., Hamel, J. C., and Tan, E. M. (1991). Human autoantibody to RNA polymerase I transcription factor hUBF. Molecular identity of nucleolus organizer region autoantigen NOR-90 and ribosomal RNA transcription upstream binding factor. J. Exp. Med. 174, 1239–1244. Chan, H. Y. E., Brogna, S., and O’Kane, C. J. (2001). Dribble, the Drosophila KRR1p homologue, is involved in rRNA processing. Mol. Biol. Cell. 12, 1409–1419. ChernoV, Y. O., Vincent, A., and Liebman, S. W. (1994). Mutations in eukaryotic 18S ribosomal RNA aVect translational fidelity and resistance to aminoglycoside antibiotics. EMBO J. 16, 4770–4776. Cheutin, T., O’Donohue, M.-F., Beorchia, A., Vandelaer, M., Kaplan, H., Defe´ver, B., Ploton, D., and Thiry, M. (2002). Three-dimensional organization of active rRNA genes within the nucleolus. J. Cell Sci. 115, 3297–3307. Christensen, M. E., and Fuxa, K. P. (1988). The nucleolar protein, B-36, contains a glycine and dimethylarginine-rich sequence conserved in several other RNA-binding proteins. Biochem. Biophys. Res. Commun. 155, 1278–1283.

162

PATRICK J. DIMARIO

Christensen, M., Barthelmes, H., Feineis, S., Knudson, B., Boege, F., and Mielke, C. (2002a). Changes in mobility account for camptothecin-induced subnuclear relocation of topoisomerase I. J. Biol. Chem. 277, 15661–15665. Christensen, M. O., Barthelmes, H. U., Boege, F., and Mielke, C. (2002b). The N-terminal domain anchors human topoisomerase I at fibrillar centers of nucleoli and nucleolar organizer regions of mitotic chromosomes. J. Biol. Chem. 277, 35932–35938. Clark, C. G., and Cross, G. A. (1988). Circular ribosomal RNA genes are a general feature of schizopyrenid amoebae. J. Protozool. 35, 326–329. Cmarko, D., Verschure, P. V., Rothblum, L. I., Hernandez-Verdun, D., Amalric, F., van Driel, R., and Fakan, S. (2000). Ultrastructural analysis of nucleolar transcription in cells microinjected with 5-bromo-UTP. Histochem. Cell Biol. 113, 181–187. Cockell, M. M., and Gasser, S. M. (1999). The nucleolus: Nucleolar space for RENT. Curr. Biol. 9, R575–R576. Conconi, A., Widmer, R. M., Koller, T., and Sogo, J. M. (1989). Two diVerent chromatin structures coexist in ribosomal RNA genes throughout the cell cycle. Cell 57, 753–761. Crosio, C., Campioni, N., Cardinali, B., Amaldi, F., and Pierandrei-Amaldi, P. (1997). Small nucleolar RNAs and nucleolar proteins in Xenopus anucleolate embryos. Chromosoma 105, 452–458. Daskal, Y., Smetana, K., and Busch, H. (1980). Evidence from studies on segregated nucleoli that nucleolar silver staining proteins C23 and B23 are in the fibrillar component. Exp. Cell Res. 127, 285–291. De Capoa, A., Pelliccia, F., Marlekaj, P., Ciofi-Luzzatto, A. R., and Buongiorno Nardelli, M. (1983). Silver positivity of the NORs during embryonic development of Xenopus laevis. Exp. Cell Res. 147, 472–478. Derenzini, M. (2000). The AgNORs. Micron 31, 117–120. Dilworth, S. M. (1991). A perichromosomal region contains proteins phosphorylated during mitosis in Xenopus laevis cells. J. Cell Sci. 98, 309–315. Dobigny, G., Ozouf-Costaz, C., Bonilla, C., and Volobouev, V. (2002). ‘‘Ag-NORS’’ are not always true NORS: New evidence in mammals. Cytogen. Genome Res. 98, 75–77. Dousset, T., Wang, C., Verheggen, C., Chen, D., Hernandez-Verdun, D., and Huang, S. (2000). Initiation of nucleolar assembly is independent of RNA polymerase I transcription. Mol. Biol. Cell. 11, 2705–2717. Draper, D. E. (1995). Protein-RNA recognition. Annu. Rev. Biochem. 64, 593–620. Dragon, F., Gallagher, J. E. G., Compagnone-Post, P. A., Mitchell, B. M., Porwancher, K. A., Wehner, K. A., Wormsley, S., Settlage, R. E., Shabanowitz, J., Osheim, Y., Beyer, A. L., Hunt, D. F., and Baserga, S. J. (2002). A large nucleolar U3 ribonucleoprotein required for 18S ribosomal RNA biogenesis. Nature 417, 967–970. Dundr, M., and Olson, M. O. J. (1998). Partially processed pre-rRNA is preserved in association with processing components in nucleolus-derived foci during mitosis. Mol. Biol. Cell. 9, 2407–2422. Dundr, M., Leno, G. H., Hammarskjo¨ld, M.-L., Rekosh, D., Helga-Maria, C., and Olson, M. O. J. (1995). The roles of nucleolar structure and function in the subcellular location of the HIV-1 Rev protein. J. Cell Sci. 108, 2811–2823. Dundr, M., Leno, G. H., Lewis, N., Rekosh, D., Hammarskjo¨ld, M.-L., and Olson, M. O. J. (1996). Location of the HIV-1 Rev protein during mitosis: Inactivation of the nuclear export signal alters the pathway for postmitotic recentry into nucleoli. J. Cell Sci. 109, 2239–2251. Dundr, M., Meier, U. T., Lewis, N., Rekosh, D., Hammarskjo¨ld, M.-L., and Olson, M. O. J. (1997). A class of nonribosomal nucleolar components is located in chromosome periphery and in nucleolus-derived foci during anaphase and telophase. Chromosoma 105, 407–417.


163

Dundr, M., Misteli, T., and Olson, M. O. J. (2000). The dynamics of postmitotic reassembly of the nucleolus. J. Cell Biol. 150, 433–446. Elsdale, T. R., Fischberg, M., and Smith, S. (1958). A mutation that reduces nucleolar number in Xenopus laevis. Exp. Cell Res. 14, 642–643. Esper, H., and Barr, H. J. (1964). A study of the developmental cytology of a mutation aVecting nucleoli in Xenopus embryos. Dev. Biol. 10, 105–121. Fakan, S., and Puvion, E. (1980). Ultrastructural visualization of nucleolar and extranucleolar RNA synthesis and distribution. Int. Rev. Cyt. 65, 255–299. Fan, H., and Penman, S. (1971). Regulation of synthesis and processing of nucleolar components in metaphase-arrested cells. J. Mol. Biol. 59, 27–42. Fankhauser, C., Izaurralde, E., Adachi, Y., Wingfield, P., and Laemmli, U. K. (1991). Specific complex of human immunodeficiency virus type 1 Rev and nucleolar B23 proteins: Dissociation by the Rev response element. Mol. Cell. Biol. 11, 2567–2575. Flećhon, J.-E., and Kopec˘ny´, V. (1998). The nature of the ‘‘nucleolus precursor body’’ in early preimplantation embryos: A review of fine-structure cytochemical, immunocytochemical and autoradiographic data related to nucleolar function Zygote 6, 183–191. Fleischmann, G., Pflugfelder, G., Steiner, E. K., Javaherian, K., Howard, G. C., Wang, J. C., and Elgin, S. (1984). Drosophila DNA topoisomerase I is associated with transcriptionally active regions of the genome. Proc. Natl. Acad. Sci. USA 81, 6958–6962. Foe, V. E., Odell, G. M., and Edgar, B. A. (1993). Mitosis and morphogenesis in the Drosophila embryo: Point and counterpoint. In ‘‘The Development of Drosophila melanogaster’’ (M. Bate and A. Marinez-Aries, Eds.), Vol. 1, pp. 149–300. Cold Spring Harbor Press, Cold Spring Harbor, NY. Fomproix, N., Ge´brane-Youne`s, J., and Hernandez-Verdun, D. (1998). EVects of antifibrillarin on the building of functional nucleoli at the end of mitosis. J. Cell Sci. 111, 359–372. Gall, J. G. (1969). The genes for ribosomal RNA during oogenesis. Genet. Suppl. 61, 121–132. Gall, J. G. (1978). Early studies on gene amplification. Harvey Lect. 71, 55–70. Gall, J. G. (2000). Cajal bodies: the first 100 years. Annu. Rev. Cell Dev. Biol. 16, 273–300. Gall, J. G. (2003). The centennial of the Cajal body. Nat. Rev. Mol. Cell Biol. 4, 975–980. Gall, J. G., and Callan, H. G. (1989). The sphere organelle contains small nuclear ribonucleoproteins. Proc. Natl. Acad. Sci. USA 86, 6635–6639. Gall, J. G., Tsvetkov, A., Wu, Z., and Murphy, C. (1995). Is the sphere organelle/coiled body a universal nuclear component? Dev. Genet. 16, 25–35. Ganot, P., Bortolin, M.-L., and Kiss, T. (1997). Site-specific pseudouridine formation in preribosomal RNA is guided by small nucleolar RNAs. Cell 89, 799–809. Garg, L. C., DiAngelo, S., and Jacob, S. T. (1987). Role of DNA topoisomerase I in the transcription of supercoiled rDNA gene. Proc. Natl. Acad. Sci. USA 84, 3185–3188. Gates, R. R. (1942). Nucleoli and related nuclear structures. Botan. Rev. 8, 337–409. Gautier, T., Dauphin-Villemant, C., Andre´, C., Masson, C., Arnoult, J., and HernandezVerdun, D. (1992a). Identification and characterization of a new set of nucleolar ribonucleoproteins which line the chromosomes during mitosis. Exp. Cell Res. 200, 5–15. Gautier, T., Fomproix, N., Masson, C., Azum-Ge´lade, M. C., Gas, N., and Hernandez-Verdun, D. (1994). Fate of specific nucleolar perichromosomal proteins during mitosis: Cellular distribution and association with U3 snoRNA. Biol. Cell 82, 81–93. Gautier, T., Robert-Nicoud, M., Guilly, M.-N., and Hernandez-Verdun, D. (1992b). Relocation of nucleolar proteins around chromosomes at mitosis. A study by confocal laser scanning microscopy. J. Cell Sci. 102, 729–737. Gautier, T., Masson, C., Quintana, C., Arnoult, J., and Hernandez-Verdun, D. (1992c). The ultrastructure of the chromosome periphery in human cell lines. An in situ study using cryomethods in electron microscopy. Chromosoma 101, 502–510.

164

PATRICK J. DIMARIO

Ge´brane-Youne`s, J., Fomproix, N., and Hernandez-Verdun, D. (1997). When rDNA transcription is arrested during mitosis, UBF is still associated with non-condensed rDNA. J. Cell Sci. 110, 2429–2440. Gerbi, S. A. (1995). Small nucleolar RNA. Biochem. Cell. Biol. 73, 845–858. Gerbi, S. A., and Lange, T. S. (2002). All small nuclear RNAs (snRNAs) of the [U4/U6.U5] trisnRNP localize to nucleoli; Identification of the nucleolar localization element of U6 snRNA Mol. Biol. Cell. 13, 3123–3127. Gerbi, S. A., Borovjagin, A. V., and Lange, T. S. (2003). The nucleolus: A site of ribonucleoprotein maturation. Curr. Opin. Cell Biol. 15, 318–325. Gerdes, J., Lemke, H., Baisch, H., Wacker, H. H., Schwab, U., and Stein, H. (1984). Cell cycle analysis of a cell proliferation-associated human nuclear antigen defined by the monoclonal antibody Ki-67. J. Immunol. 133, 1710–1715. Ghisolfi, L., Gerard, J., Amalric, F., and Erard, M. (1992a). The glycine-rich domain of nucleolin has an unusual supersecondary structure responsible for its RNA-helix-destabilizing properties. J. Biol. Chem. 267, 2955–2959. Ghisolfi-Nieto, L., Joseph, G., Puvion-Dutilleul, F., Amalric, F., and Bouvet, P. (1996). Nucleolin is a sequence-specific RNA-binding protein: Characterization of targets on preribosomal RNA. J. Mol. Biol. 260, 34–53. Ghisolfi, L., Kharrat, A., Joseph, G., Amalric, F., and Erard, M. (1992b). Concerted activities of the RNA recognition and the glycine-rich C-terminal domains of nucleolin are required for eYcient complex formation with pre-ribosomal RNA. Eur. J. Biochem. 209, 541–548. Gilbert, N., Lucas, L., Klein, C., Menager, M., Bonnet, N., and Ploton, D. (1995). Threedimensional co-location of RNA polymerase I and DNA during interphase and mitosis by confocal microscopy. J. Cell Sci. 108, 115–125. Gimeńez-Martin, G., De la Torre, C., Lo´pez-Saéz, J. F., and Esponda, P. (1977). Plant nucleolus; structure and physiology. Cytobiologie 14, 421–462. Ginisty, H., Amalric, F., and Bouvet, P. (1998). Nucleolin functions in the first step of ribosomal RNA processing. EMBO J. 17, 1476–1486. Ginisty, H., He´le`ne, S., Roger, B., and Bouvet, P. (1999). Structure and functions of nucleolin. J. Cell Sci. 112, 761–772. Giordano, E., Peluso, I., Senger, S., and Furia, M. (1999). minifly, a Drosophila gene required for ribosome biogenesis. J. Cell Biol. 144, 1123–1133. Giordano, E., Rendina, R., Peluso, I., and Furia, M. (2002). RNAi triggered by symmetrically transcribed transgenes in Drosophila melanogaster. Genetics 160, 637–648. Goessens, G., and Lepoint, A. (1974). The fine structure of the nucleolus during interphase and mitosis in Ehrlich tumor cells cultivated in vitro. Exp. Cell Res. 87, 63–72. Goessens, G. (1984). Nucleolar structure. Int. Rev. Cytol. 87, 107–158. Goodpasture, C., and Bloom, S. E. (1975). Visualization of nucleolar organizer regions in mammalian chromosomes using silver staining. Chromosoma 53, 37–50. Gottesfeld, J. M., Wolf, V. J., Dang, T., Forbes, D. J., and Hartl, P. (1994). Mitotic repression of RNA polymerase III transcription in vitro mediated by phosphorylation of a TFIIIB component. Science 263, 81–84. Gottlieb, S., and Esposito, R. E. (1989). A new role for a yeast transcriptional silencer gene, SIR2, in regulation of recombination in ribosomal DNA. Cell 56, 771–776. Grandi, P., Rybin, V., Bassler, J., Petfalski, E., Strauss, D., Marzioch, M., Schafer, T., Kuster, B., Tschochner, H., Tollervey, D., Gavin, A. C., and Hurt, E. (2002). 90S preribosomes include the 35S-pre-rRNA, the U3 snoRNP, and 40S-subunit processing factors but predominantly lack 60S synthesis factors. Mol. Cell 10, 105–115. Grummt, I. (1999). Regulation of mammalian ribosomal gene transcription by RNA polymerase I. Prog. Nucleic Acid Res. Mol. Biol. 62, 109–154.


165

Guarente, L. (2000). Sir2 links chromatin silencing, metabolism, and aging. Genes Dev. 14, 1021–1026. Guldner, H.-H., Szostecki, C., Vosber, H.-P., Lakomek, H.-J., Penner, E., and Bautz, F. A. (1986). Scl 70 autoantibodies from scleroderma patients recognize a 95 kDa protein identified as DNA topoisomerase I. Chromosoma 94, 132–138. Gulli, M-P., Faubladier, M., Sicard, H., and Caizergues-Ferrer, M. (1997). Mitosis-specific phosphorylation of gar2, a fission yeast nucleolar protein structurally related to nucleolin. Chromosoma 105, 532–541. Guttes, S., Guttes, E., and Ellis, R. (1968). Electron microscope study of mitosis in Physarum polycephalum. J. Ultrast. Res. 22, 508–529. Haaf, T., Reimer, G., and Schmid, M. (1988). Immunocytogenetics: Localization of trenscriptionally active rRNA genes in nucleoli and nucleolus organizer regions by use of human autoantibodies to RNA polymerase I. Cytogenet. Cell. Genet. 48, 35–42. Hadjiolov, A. A. (1985). The nucleolus and ribosome biogenesis. Springer, New York. Hannan, K. M., Kennedy, B. K., Cavanaugh, A. H., Hannan, R. D., Hirschler-Laszkiewicz, I., JeVerson, L. S., and Rothblum, L. I. (2000a). RNA polymerase I transcription in confluent cells: RB downregulates rDNA transcription during confluence-induced cell cycle arrest. Oncogene 19, 3487–3497. Hannan, K. M., Hannan, R. D., Smith, S. D., JeVerson, L. S., Lun, M., and Rothblum, L. I. (2000b). Rb and p130 regulate RNA polymerase I transcription: Rb disrupts the interaction between UBF and SL1. Oncogene 19, 4988–4999. Hartl, P., Gottesfeld, J., and Forbes, D. J. (1993). Mitotic repression of transcription in vitro. J. Cell Biol. 120, 613–624. Hay, E. D. (1968). Structure and function of the nucleolus in developing cells. In ‘‘The Nucleus’’ (A. J. Dalton and F. Haguenau, Eds.), Vol. 3, pp. 1–79. Academic Press, New York. Hay, E. D., and Gurdon, J. B. (1967). Fine structure of the nucleolus in normal and mutant Xenopus embryos. J. Cell Sci. 2, 151–162. Heald, R., and McKeon, F. (1990). Mutations of phosphorylation sites in lamin A that prevent nuclear lamina disassembly in mitosis. Cell 61, 579–589. Heitz, E. (1931). Die ursache der gesetzma¨ssigen zahl, lage, form und gro¨sse pflanzlicher nukleolen. Planta (Berlin) 12, 775–844. Heix, J., Vente, A., Voit, R., Budde, A., Michaelidis, T. M., and Grummt, I. (1998). Mitotic silencing of human rRNA synthesis: Inactivation of the promoter selectivity factor SL1 by cdc2/cyclin B-mediated phosphorylation. EMBO J. 17, 7373–7381. Heliot, L., Kaplan, H., Lucas, L., Klein, C., Beorchia, A., Doco-Fenzy, M., Menager, M., Thiry, M., O’Donohue, M.-F., and Ploton, D. (1997). Electron tomography of the metaphase nucleolar organizer regions: Evidence for a twisted-loop organization. Mol. Biol. Cell. 8, 2199–2216. Henry, M. F., and Silver, P. A. (1996). A novel methyltransferase (Hmt1p) modifies poly(A)þRNA-binding proteins. Mol. Cell. Biol. 16, 3668–3678. Hernandez-Verdun, D. (2003). Behavior of the nucleolus during mitosis. In ‘‘The Nucleolus’’ (M.O.J. Olson, Ed.), Chapter 5. Kluwer Academic/Plenum Publishing, New York. Hernandez-Verdun, D., and Derenzini, M. (1983). Non-nucleosomal configuration of chromatin in nucleolar organizer regions of metaphase chromosomes in situ. Eur. J. Cell Biol. 31, 360–365. Hernandez-Verdun, D., and Gautier, T. (1994). The chromosome periphery during mitosis. Bioessays 16, 179–185. Hernandez-Verdun, D., and Roussel, P. (2003). Regulators of nucleolar functions. Prog. Cell Cycle Res. 5, 301–308. Hernandez-Verdun, D., Bouteille, M., Ege, T., and Ringertz, N. R. (1979). Fine structure of nucleoli in micronucleated cells. Exp. Cell Res. 124, 223–235.

166

PATRICK J. DIMARIO

Hernandez-Verdun, D., Robert-Nicoud, M., Geraud, G., and Masson, C. (1991). Behaviour of nucleolar proteins in nuclei lacking ribosomal genes. A study by confocal laser scanning microscopy. J. Cell Sci. 98, 99–105. Hernandez-Verdun, D., Roussel, P., and Gautier, T. (1993). Nucleolar proteins during mitosis. Chromosomes Today 11, 79–90. Hernandez-Verdun, D., Roussel, P., and Ge´brane-Youne`s, G. (2002). Emerging concepts of nucleolar assembly. J. Cell Sci. 115, 2265–2270. Herrera, A. H., and Olson, M. O. J. (1986). Association of protein C23 with rapidly labeled nucleolar RNA. Biochemistry 25, 6258–6264. Hisatake, K., Nishimura, T., Maeda, Y., Hanada, K., Song, C. Z., and Muramatsu, M. (1991). Cloning and structural analysis of cDNA and the gene for mouse transcription factor UBF. Nucl. Acids Res. 19, 4631–4637. Hoang, C., and Ferre´-D’Amare´, A. R. (2001). Cocrystal structure of a tRNA Psi55 pseudouridine synthase: Nucleotide flipping by an RNA-modifying enzyme. Cell 107, 929–939. Howell, W. M. (1982). Selective staining of nucleolus organizer regions (NORs). In ‘‘The Cell Nucleus’’ (H. Busch and L. Rothblum, Eds.), Vol 11, pp. 89–142. Academic Press, New York. Hoza´k, P., Cook, P. R., Scho¨fer, C., Mosgo¨ller, W., and Wachtler, F. (1994). Site of transcription of ribosomal RNA and intranucleolar structure in HeLa cells. J. Cell Sci. 107, 639–648. Hsu, T. C., Arrighi, F. E., Klevecz, R. R., and Brinkley, B. R. (1965). The nucleoli in mitotic divisions of mammalian cells in vitro. J. Cell Biol. 26, 539–553. Huang, S. (2002). Building an eYcient factory: Where is pre-rRNA synthesized in the nucleolus? J. Cell Biol. 157, 739–741. Huang, J., and Moazed, D. (2003). Association of the RENT complex with nontranscribed and coding regions of rDNA and a regional requirement for the replication fork block protein Fob1 in rDNA silencing. Genes Dev. 17, 2162–2172. Hughes, J. M. (1996). Functional base-pairing interaction between highly conserved elements of U3 small nucleolar RNA and the small ribosomal subunit RNA. J. Mol. Biol. 259, 645–654. Hu¨gle, B., Hazan, R., Scheer, U., and Franke, W. W. (1985a). Localization of ribosomal protein S1 in the granular component of the interphase nucleolus and its distribution during mitosis. J. Cell Biol. 100, 873–886. Hu¨gle, B., Scheer, U., and Franke, W. W. (1985b). Ribocharin: A nuclear Mr 40000 protein specific to particles of the large ribosomal subunit. Cell 41, 615–627. Hyttel, P., Laurincik, J., Rosenkranz, C., Rath, D., Niemann, H., Ochs, R. L., and Schellander, K. (2000). Nucleolar proteins and ultrastructure in preimplantation porcine embryos developed in vivo. Biol. Reprod. 63, 1848–1856. Jacob, J., and Sirlin, J. L. (1963). Electron microscopic studies on salivary gland cells. J. Cell Biol. 17, 153–165. Jacobson, M. R., and Pederson, T. (1998). Localization of signal recognition particle RNA in the nucleolus of mammalian cells. Proc. Natl. Acad. Sci. USA 95, 7981–7986. Jacobson, M. R., Cao, L.-G., Taneja, K., Singer, R. H., Wang, Y.-L., and Pederson, T. (1997). Nuclear domains of the RNA subunit of RNase P. J. Cell Sci. 110, 829–837. Jarrous, N., Wolenski, J. S., Wesolowski, D., Lee, C., and Altman, S. (1999). Localization in the nucleolus and coiled bodies of protein subunits of the ribonucleoprotein ribonuclease P. J. Cell Biol. 146, 559–571. Jimeńez-Garcia, L. F., Rothblum, L. I., Busch, H., and Ochs, R. L. (1989). Nucleologenesis: Use of non-isotopic in situ hybridization and immunocytochemistry to compare the localization of rDNA and nucleolar proteins during mitosis. Biol. Cell 65, 239–246.


167

Jimeńez-Garcıá, L. F., Segura-Valdez, M. de L., Ochs, R. L., Rothblum, L. I., Hannan, R., and Spector, D. L. (1994). Nucleologenesis: U3 snRNA-containing prenucleolar bodies move to sites of active pre-rRNA transcription after mitosis. Mol. Biol. Cell 5, 955–966. Johnson, A., Lund, E., and Dahlberg, J. (2002). Nuclear export of ribosomal subunits. Trends Biochem. Sci. 27, 580–585. Jordan, E. G., and Cullis, C. A. (1982). ‘‘The Nucleolus.’’ Cambridge University Press, Cambridge. Jordan, P., Mannervik, M., Tora, L., and Carmo-Fonseca, M. (1996). In vivo evidence that TATA-binding protein/SL1 colocalizes with UBF and RNA polymerase I when rRNA synthesis is either active or inactive. J. Cell Biol. 133, 225–234. June´ra, H. R., Masson, C., Ge´raud, G., Suja, J., and Hernandez-Verdun, D. (1997). Involvement of in situ conformation of ribosomal genes and selective distribution of UBF in rRNA transcription. Mol. Biol. Cell 8, 145–156. Kan˘ka, J., Hoza´k, P., Heyman, Y., Chesne´, P., Degrolard, J., Renard, J. P., and Flećhon, J. E. (1996). Transcriptional activity and nucleolar ultrastructure of embryonic rabbit nuclei after transplantation to enucleated oocytes. Mol. Reprod. Dev. 43, 135–144. Karpen, G. H., Schaefer, J. E., and Laird, C. D. (1988). A Drosophila rRNA gene located in euchromatin is active in transcription and nucleolus formation. Genes Dev. 2, 1745–1763. KauVman, T., Tran, J., and Dinardo, S. (2003). Mutations in Nop60B, the Drosophila homolog of human Dyskeratosis congenita 1, aVect the maintenance of the germ-line lineage during spermatogenesis. Dev. Biol. 253, 189–199. Kay, M. A., and Jacobs-Lorena, M. (1987). Developmental genetics of ribosome synthesis in Drosophila. Trends Genet. 3, 347–351. Keys, D. A., Vu, L., SteVan, J. S., Dodd, J. A., Yamamoto, R. T., Nogi, Y., and Nomura, M. (1994). RRN6 and RRN7 encode subunits of a multiprotein complex essential for the initiation of rDNA transcription by RNA polymerase I in Saccharomyces cerevisiae. Genes Dev. 8, 2349–2362. Kiss, T. (2002). Small nucleolar RNAs: An abundant group of noncoding RNAs with diverse cellular functions. Cell 109, 145–148. Klein, J., and Grummt, I. (1999). Cell cycle-dependent regulation of RNA polymerase I transcription: The nucleolar transcription factor UBF is inactive in mitosis and early G1. Proc. Natl. Acad. Sci. USA 96, 6096–6101. Kloc, M., Matuszewski, B., and Nurkowska, J. (1995). Ribosomal gene amplification in the oocytes of Creophilus maxillosus (Staphylinidae, Coleoptera-polyphaga)—an insect with telotrophic ovaries. Folia Histochem. Cytobiol. 33, 267–276. Koberna, K., Malıńsky, J., Pliss, A., Masata, M., Vecerova´, J., Fialova´, M., Bedna´r, J., and Raska, I. (2002). Ribosomal genes in focus: new transcripts label the dense fibrillar components and form clusters indicative of ‘‘Christmas trees’’ in situ J. Cell Biol. 157, 743–748. Kopec˘ny´, V. (1990). Nucleologenesis in cow embryo: Relation between onset of transcription and penetration of DNA into nucleolar precursor body. In ‘‘Nuclear Structure and Function’’ (J. R. Harris and I. B. Zbarsky, Eds.), pp. 187–191. Academic Press, New York. Kopec˘ny´, V., Biggiogera, M., Laurincik, J., Pivko, J., Grafenau, P., Martin, T. E., Fu, X. D., and Fakan, S. (1996a). Fine structural cytochemical and immunocytochemical analysis of nucleic acids and ribonucleoprotein distribution in nuclei of pig oocytes and early preimplantation embryos. Chromosoma 104, 561–574. Kopec˘ny´, V., Biggiogera, M., Pivko, J., Grafenau, P., Pavlock, A., Malatesta, M., Martin, T. E., and Fakan, S. (1996b). The cell nucleus in early bovine and caprine preimplantation embryos: Fine structural cytochemistry and immunoelectron microscopy. Eur. J. Cell Biol. 70, 361–372.

168

PATRICK J. DIMARIO

Kopec˘ny´, V., Biggiogera, M., Pivko, J., Pavlok, A., Martin, T. E., Kaufmann, S. H., Shaper, J. H., and Fakan, S. (2000). Fine-structural cytochemical and immunocytochemical observations on nuclear bodies in the bovine 2-cell embryo. Zygote 8, 315–328. Kopec˘ny´, V., Flećhon, J.-E., Camous, S., and Fulka, J., Jr. (1989). Nucleologenesis and the onset of transcription in the eight-cell bovine embryo: Fine-structural autoradiographic study. Mol. Reprod. Dev. 1, 79–90. Kubrakiewicz, J., and Bili’nski, S. M. (1995). Extrachromosomal amplification of rDNA in oocytes of Hemerobius spp. (Insecta, Neuroptera). Chromosoma 103, 606–612. Kuhn, A., Vente, A., Doree, M., and Grummt, I. (1998). Mitotic phosphorylation of the TBPcontaining SL1 represses ribosomal gene transcription. J. Mol. Biol. 284, 1–5. Kuhn, A., Voit, R., Stefanovsky, V., Evers, R., Bianchi, M., and Grummt, I. (1994). Functional diVerences between the two splice variants of the nucleolar transcription factor UBF: The second HMG box determines specificity of DNA binding and transcriptional activity. EMBO J. 13, 416–424. Lafontaine, J. G. (1958). Structure and mode of formation of the nucleolus in meristematic cells of Vicia faba and Allium cepa. J. Biophysic. Biochem. Cytol. 4, 777–784 with plates 392–398. Lafontaine, J. G., and Chouinard, L. A. (1963). A correlated light and electron microscopic study of the nucleolar material during mitosis in Vicia faba. J. Cell. Biol. 17, 167–201. Lafontaine, D. L. J., Bousquet-Antonelli, C., Henry, Y., Caizergues-Ferrer, M., and Tollervey, D. (1998). The box H + ACA snoRNAs carry Cbf5p, the putative rRNA pseudouridine synthase. Genes Dev. 12, 527–537. Lange, T. S., and Gerbi, S. A. (2000). Transient nucleolar localization of U6 small nuclear RNA. Mol. Biol. Cell 11, 2419–2428. Lapeyre, B., Bourbon, H., and Amalric, F. (1987). Nucleolin, the major nucleolar protein of growing eukaryotic cells: An unusual protein structure revealed by the nucleotide sequence. Proc. Natl. Acad. Sci. USA 84, 1472–1476. Laurincik, J., Hyttel, P., and Kopec˘ny´, V. (1995). DNA synthesis and pronucleus development in pig zygotes obtained in vivo: An autoradiographic and ultrastructural study. Mol. Reprod. Dev. 40, 325–332. Laurincik, J., Zakhartchenko, V., Stojkovic, M., Brem, G., Wolf, E., Muller, M., Ochs, R. L., and Maddox-Hyttel, P. (2002). Nucleolar protein allocation and ultrastructure in bovine embryos produced by nuclear transfer from granulose cells. Mol. Reprod. Dev. 61, 477–487. Lepoint, A., and Goessens, G. (1978). Nucleologenesis in Ehrlich tumour cells. Exp. Cell Res. 117, 89–94. Leresche, A., Wolf, V. J., and Gottesfeld, J. M. (1996). Repression of RNA polymerase II and III transcription during M phase of the cell cycle. Exp. Cell Res. 229, 282–288. Leung, A. K. L., and Lamond, A. I. (2003). The dynamics of the nucleolus. Crit. Rev. Eukaryot. Gene Expr. 13, 39–54. Leung, A. K., Andersen, J. S., Mann, M., and Lamond, A. I. (2003). Bioinformatic analysis of the nucleolus. Biochem. J. 376, 553–569. Li, Y.-P., Busch, R. K., Valdez, B. C., and Busch, H. (1996). C23 interacts with B23, a putative nucleolar-localization-signal-binding protein. Eur. J. Biochem. 237, 153–158. Lin, M. (1955). Chromosomal control of nuclear composition in maize. Chromosoma 7, 340–370. Lischwe, M. A., Cook, R. G., Ahn, Y. S., Yeoman, L. C., and Busch, H. (1985a). Clustering of glycine and NG,NG-dimethylarginine in nucleolar protein C23. Biochemistry 24, 6025–6028. Lischwe, M. A., Ochs, R. L., Reddy, R., Cook, R. G., Yeoman, L. C., Tan, E. M., Reichlin, M., and Busch, H. (1985b). Purification and partial characterization of nucleolar scleroderma antigen (Mr 43,000; pI 8.5) rich in NG,NG -dimethylarginine. J. Biol. Chem. 260, 14304 –14310.


169

Lischwe, M. A., Roberts, K. D., Yeoman, L. C., and Busch, H. (1982). Nucleolar specific acidic phosphoprotein C23 is highly methylated. J. Biol. Chem. 25, 14600–14602. Liu, Q., and Dreyfuss, G. (1995). In vivo and in vitro arginine methylation of RNA-binding proteins. Mol. Cell. Biol. 15, 2800–2808. Macgregor, H. C. (1972). The nucleolus and its genes in amphibian oogenesis. Biol. Rev. 47, 177–210. Maercker, C., Harjes, P., Neben, M., Niemann, H., Sianidis, G., and Lipps, H. J. (1997). The formation of new nucleoli during macronuclear development of the hypotrichous ciliate Stylonychia lemnae visualize by in situ hybridization. Chromosome Res. 5, 333–335. Mahowald, A. P. (1963). Ultrastructural diVerentiations during formation of the blastoderm in the Drosophila melanogaster embryo. Dev. Biol. 8, 186–204. Mahowald, A. P. (2001). Assembly of the Drosophila germ plasm. Int. Rev. Cytol. 203, 187–213. Mahowald, A. P., and Hardy, P. A. (1985). Genetics of Drosophila embryogenesis. Ann. Rev. Genet. 19, 149–177. Mais, C., McStay, B., and Scheer, U. (2002). On the formation of amplified nucleoli during early Xenopus oogenesis. J. Struct. Biol. 140, 214–226. Malıńsky, J., Kobrena, K., Bedna´r, J., Stulı´k, J., and Raska, I. (2002). Searching for active ribosomal genes in situ: Light microscopy in light of the electron beam. J. Struct. Biol. 140, 227–231. Matsui, S., and Sandberg, A. A. (1985). Intranuclear compartmentalization of DNA-dependent RNA polymerases: Association of RNA polymerase I with nucleolar organizing chromosomes. Chromosoma 92, 1–6. Maxwell, E. S., and Fournier, M. J. (1995). The small nucleolar RNAs. Ann. Rev. Biochem. 35, 897–933. McClintock, B. (1934). The relationship of a particular chromosomal element to the development of the nucleoli in Zea mays. Zeitschr. Zellf. Mikr. Anat. 21, 294–328. McKee, B. D., Habera, L., and Vrana, J. A. (1992). Evidence that intergenic spacer repeats of Drosophila melanogaster rRNA genes function as X -Y pairing sites in male meiosis, and a general model for achiasmatic pairing. Genetics 132, 529–544. McStay, B., Hu, C. H., Pikaard, C. S., and Reeder, R. H. (1991). xUBF and Rib 1 are both required for formation of a stable polymerase I promoter complex in X. laevis. EMBO J. 10, 2297–2303. Medina, F. J., Cerdido, A., and Fernańdez-Go´mez, M. E. (1995). Components of the nucleolar processing complex (pre-rRNA, fibrillarin, and nucleolin) colocalize during mitosis and are incorporated to daughter cell nucleoli. Exp. Cell Res. 221, 111–125. Meier, U. T. (1996). Comparison of the rat nucleolar protein Nopp140 with its yeast homolog SRP40. J. Biol. Chem. 271, 19376–19384. Meier, U. T., and Blobel, G. (1990). A nuclear localization signal binding protein in the nucleolus. J. Cell Biol. 111, 2235–2245. Meier, U. T., and Blobel, G. (1992). Nopp140 shuttles on tracks between nucleolus and cytoplasm. Cell 70, 127–138. Meier, U. T., and Blobel, G. (1994). NAP57, a mammalian nucleolar protein with a putative homolog in yeast and bacteria. J. Cell Biol. 127, 1505–1514. Me´le`se, T., and Xue, Z. (1995). The nucleolus: An organelle formed by the act of building a ribosome. Curr. Opin. Cell Biol. 7, 319–324. Meßmer, B., and Dreyer, C. (1993). Requirements for nuclear translocation and nucleolar accumulation of nucleolin of Xenopus laevis. Eur. J. Cell Biol. 61, 369–382. Milkereit, P., Gadal, O., Podtelejnikov, A., Trumtel, S., Gas, N., Petfalski, E., Tollervey, D., Mann, M., Hurt, E., and Tschochner, H. (2001). Maturation and intranuclear transport of pre-ribosomes requires noc proteins. Cell 105, 499–509.

170

PATRICK J. DIMARIO

Milkereit, P., Strauss, D., Bassler, J., Gadal, O., Ku¨hn, H., Schu¨tz, S., Gas, N., Lechner, J., Hurt, E., and Tschochner, H. (2003). A Noc complex specifically involved in the formation and nuclear export of ribosomal 40 S subunits. J. Biol Chem. 278, 4072–4081. Miller, O. L., Jr. (1981). The nucleolus, chromosomes, and visualization of genetic activity. J. Cell Biol. 91, 15s–27s. Mitchell, J. R., Cheng, J., and Collins, K. (1999). A box H/ACA small nucleolar RNA-like domain at the human telomerase 30 -end. Mol. Biol. Cell 19, 567–576. Montgomery, T. H. (1898). Comparative cytological studies, with especial regard to the morphology of the nucleolus. J. Morph. 15, 265–582. Morcillo, G., De La Torre, C., and Gimeńez-Martin, G. (1976). Nucleolar transcription during plant mitosis. Exp. Cell Res. 102, 311–316. Morgan, T. H., Bridges, C. B., and Sturtevant, A. H. (1925). The genetics of Drosophila melanogaster. Biblphia. Genet. 2, 1–262. Mosgoeller, W., Scho¨fer, C., Wesierska-Gadek, J., Steiner, M., Mu¨ller, M., and Wachtler, F. (1998). Ribosomal gene transcription is organized in foci within nucleolar components. Histochem. Cell Biol. 109, 111–118. Moss, T., and Stefanovsky, V. Y. (1995). Promotion and regulation of ribosomal transcription in eukaryotes by RNA polymerase I. Prog. Nucleic Acid Res. Mol. Biol. 50, 25–65. Moss, T., and Stefanovsky, V. Y. (2002). At the center of eukaryotic life. Cell 109, 545–548. Mougey, E. B., Pape, L. K., and Sollner-Webb, B. (1993a). A U3 small nuclear ribonucleoprotein-requiring processing event in the 50 external transcribed spacer of Xenopus precursor rRNA. Mol. Cell. Biol. 13, 5990–5998. Mougey, E. B., O’Reilly, M., Osheim, Y., Miller, O. L., Jr., Beyer, A., and Sollner-Webb, B. (1993b). The terminal balls characteristic of eukaryotic rRNA transcription units in chromatin spreads are rRNA processing complexes. Genes Dev. 7, 1609–1619. Muller, M. T., Pfund, W. P., Mehta, V. B., and Trask, D. R. (1985). Eukaryotic type topoisomerase is enriched in the nucleous and catalytically active on ribosomal DNA. EMBO J. 4, 1237–1243. Najbauer, J., Johnson, B. A., Young, A. L., and Aswad, D. W. (1993). Peptides with sequences similar to glycine, arginine-rich motifs in proteins interacting with RNA are eYciently recognized by methyltransferase(s) modifying arginine in numerous proteins. J. Biol. Chem. 268, 10501–10509. Narayanan, A., Lukowiak, A., Jady, B. E., Dragon, F., Kiss, T., Terns, R. M., and Terns, M. P. (1999). Nucleolar localization signals of box H/ACA small nucleolar RNAs. EMBO J. 18, 5120–5130. Ni, J., Tien, A. L., and Fournier, M. J. (1997). Small nucleolar RNAs direct site-specific synthesis of pseudouridine in ribosomal RNA. Cell 89, 565–573. Nierras, C. R., Liebman, S. W., and Warner, J. R. (1997). Does Saccharomyces need an organized nucleolus? Chromosoma 105, 444–451. Noel, J. S., Dewey, W. C., Abel, J. H., Jr., and Thompson, R. P. (1971). Ultrastructure of the nucleolus during the Chinese hamster cell cycle. J. Cell Biol. 49, 830–847. Nomura, M. (2001). Ribosomal RNA genes, RNA polymerases, nucleolar structures, and synthesis of rRNA in the yeast Saccharomyces cerevisiae. Cold Spring Harbor Symp. Quant. Biol. 66, 555–565. Oakes, M., Aris, J. P., Brockenbrough, J. S., Wai, H., Vu, L., and Nomura, M. (1998). Mutational analysis of the structure and localization of the nucleolus in the yeast Saccharomyces cerevisiae. J. Cell Biol. 143, 23–34. Oakes, M., Nogi, Y., Clark, M. W., and Nomura, M. (1993). Structural alterations of the nucleolus in mutants of Saccharomyces cerevisiae defective in RNA polymerase I. Mol. Cell. Biol. 13, 2441–2455.


171

Oakes, M., Siddiqi, I., Vu, L., Aris, J., and Nomura, M. (1999). Transcription factor UAF, expansion and contraction of ribosomal DNA (rDNA) repeats, and RNA polymerase switch in transcription of yeast rDNA. Mol. Cell. Biol. 19, 8559–8569. Ochs, R. L., and Busch, H. (1984). Further evidence that phosphoprotein C23 (110 kD/pI 5.1) is the nucleolar silver staining protein. Exp. Cell Res. 152, 260–265. Ochs, R. L., Lischwe, M. A., Shen, E., Carroll, R. E., and Busch, H. (1985). Nucleologenesis: Composition and fate of prenucleolar bodies. Chromosoma 92, 330–336. Ochs, R. L., Stein, T. W., Jr., Chan, E. K., Ruutu, M., and Tan, E. M. (1996). cDNA cloning and characterization of a novel nucleolar protein. Mol. Biol. Cell. 7, 1015–1024. Ogata, M. (1883). Vera¨nderung der Pancreaszellen bei der Sekretion. Arch. Anat. Physiol. Abt. 405–437. Ogg, S. C., and Lamond, A. I. (2002). Cajal bodies and coilin—moving towards function. J. Cell Biol. 159, 17–21. Olson, M. O. J. (1990). The role of proteins in nucleolar structure and function. In ‘‘The Eukaryotic Nucleus: Molecular Biochemistry and Macromolecular Assemblies’’ (P. R. Straus and S. H. Wilson, Eds.), Vol. 2, pp. 519–559. Telford Press, West Caldwell, NJ. Olson, M. O. J., Hingorani, K., and Szebeni, A. (2002). Conventional and nonconventional roles of the nucleolus. Int. Rev. Cyt. 219, 199–266. O’Mahony, D. J., and Rothblum, L. I. (1991). Identification of two forms of the RNA polymerase I transcription factor UBF. Proc. Natl. Acad. Sci. USA 88, 3180–3184. O’Mahony, D. J., Xie, W. Q., Smith, S. D., Singer, H. A., and Rothblum, L. I. (1992). DiVerential phosphorylation and localization of the transcription factor UBF in vivo in response to serum deprivation. J. Biol. Chem. 267, 35–38. O’Sullivan, A. C., Sullivan, G. J., and McStay, B. (2002). UBF binding in vivo is not restricted to regulatory sequences within the vertebrate ribosomal DNA repeat. Mol. Cell. Biol. 22, 657–668. Paweletz, N., and Risuenõ, M. C. (1982). Transmission electron microscopic studies on the mitotic cycle of nucleolar proteins impregnated with silver. Chromosoma 85, 261–273. Pai, C.-Y., Chen, H.-K., Sheu, H.-L., and Yeh, N.-H. (1995). Cell cycle-dependent alterations of a highly phosphorylated nucleolar protein p130 are associated with nucleologenesis. J. Cell Sci. 108, 1911–1920. Pai, C.-Y., and Yeh, N.-H. (1996). Cell proliferation-dependent expression of two isoforms of the nucleolar phosphoprotein p130. Biochem. Biophys. Res. Commun. 221, 581–587. Paule, M. R. (1998). Transcription of ribosomal genes by eukaryotic polymerase I. Berlin, Springer Verlag. Peculis, B. A., and Steitz, J. A. (1993). Distribution of U8 nucleolar snRNA inhibits 5.8S and 28S rRNA processing in the Xenopus oocyte. Cell 73, 1233–1245. Peculis, B. A., and Steitz, J. A. (1994). Sequence and structural elements critical for U8 snRNP function in Xenopus oocytes are evolutionarily conserved. Genes Dev. 8, 2241–2255. Peculis, B. A. (2000). RNA-binding proteins: If it looks like a sn(o)RNA . . . . Curr. Biol. 10, R916–R918. Pederson, T. (1998a). The plurifunctional nucleolus. Nucleic Acids Res. 26, 3871–3876. Pederson, T. (1998b). Growth factors in the nucleolus? J. Cell Biol. 143, 279–281. Pederson, T., and Politz, J. C. (2000). The nucleolus and the four ribonucleoproteins of translation. J. Cell Biol. 148, 1091–1095. Pellar, G. J., and DiMario, P. J. (2003). Deletion and site-specific mutagenesis of nucleolin’s carboxy GAR domain. Chromosoma 111, 461–469. Peter, M., Nakagawa, J., Doreé, M., Labbe´, J. C., and Nigg, E. A. (1990a). Identification of major nucleolar proteins as candidate mitotic substrates of cdc2 kinase. Cell 60, 791–801.

172

PATRICK J. DIMARIO

Peter, M., Nakagawa, J., Doreé, M., Labbe´, J. C., and Nigg, E. A. (1990b). In vitro disassembly of the nuclear lamina and M phase-specific phosphorylation of lamins by cdc2 kinase. Cell 61, 591–602. Petes, T. (1979). Yeast ribosomal DNA genes are located in chromosome XII. Proc. Natl. Acad. Sci. USA 76, 410–414. Pfeifle, J., Anderer, F. A., and Franke, M. (1986). Characterisation of nucleolar proteins as autoantigens using human autoimmune sera. Ann. Rheum. Dis. 45, 978–986. Phair, R. D., and Mistelli, T. (2000). High mobility of proteins in the mammalian cell nucleus. Nature 404, 604–609. Phillips, B., Billin, A. N., Cadwell, C., Buchholz, R., Erickson, C., Merriam, J. R., Carbon, J., and Poole, S. J. (1998). The Nop60B gene of Drosophila encodes an essential nucleolar protein that functions in yeast. Mol. Gen. Genet. 260, 20–29. Philips, S. G., and Phillips, D. M. (1979). Nucleolus-like bodies in micronuclei of cultured Xenopus cells. Exp. Cell Res. 120, 295–306. Pierron, G., and Puvion-Dutilleul, F. (1993). Mitotic segregation of the nucleolar RNA in Physarum polycephalum. Exp. Cell Res. 208, 509–517. Pierron, G., and Puvion-Dutilleul, F. (1996). Localization of the newly initiated and processed ribosomal primary transcripts during the mitotic cycle in Physarum polycephalum. Exp. Cell Res. 229, 407–420. Pinõl-Roma, S. (1999). Association of nonribosomal nucleolar proteins in ribonucleoprotein complexes during interphase and mitosis. Mol. Biol. Cell 10, 77–90. Ploton, D., Gilbert, N., Menager, M., Kaplan, H., and Adnet, J. J. (1994). Three-dimensional co-localization of nucleolar argyrophilic components and DNA in cell nuclei by confocal microscopy. J. Histochem. Cytochem. 42, 137–148. Ploton, D., Thiry, M., Menager, M., Lepoint, A., Adnet, J.-J., and Goessens, G. (1987). Behaviour of nucleolus during mitosis. A comparative ultrastructural study of various cancerous cell lines using the Ag-NOR staining procedure. Chromosoma 95, 95–107. Politz, J. C., Kilroy, S. M., Cohen, H. R., and Pederson, T. (1998). Endogenous signal recognition particle RNA is present in the nucleolus as well as the peri-nuclear regions of normal rat kidney cells. Mol. Biol. Cell 9, 190a. Politz, J. C., Lewandowski, L. B., and Pederson, T. (2002). Signal recognition particle RNA localization within the nucleolus diVers from the classical sites of ribosome synthesis. J. Cell Biol. 159, 411–418. Politz, J. C., Yarovoi, S., Kilroy, S. M., Gowda, K., Zwieb, C., and Pederson, T. (2000). Signal recognition particle components in the nucleolus. Proc. Natl. Acad. Sci. USA 97, 55–60. Ramoń y Cajal, S. R. (1903). Un sencillo metodo de coloacion seletiva del reticulo protoplasmico y sus efectos en los diversos organos nerviosos de vertebrados y invertebrados. Trab. Lab. Invest. Biol. 2, 129–221. Raska, I., Dundr, M., Koberna, K., Melca´k, I., Riuenõ, M.-C., and To¨ro¨k, I. (1995). Does the synthesis of ribosomal RNA take place within nuclear fibrillar centers or dense fibrillar components? A critical appraisal. J. Struct. Biol. 114, 1–22. Ravel-Chapuis, P. (1988). Nuclear rDNA in Euglena gracilis: Paucity of chromosomal units and replication of extrachromosomal units. Nucleic Acids Res. 16, 4801–4810. Rattner, J. B. (1992). Integrating chromosome structure with function. Chromosoma 101, 259–264. Reeder, R. H., Pikaard, C. S., and McStay, B. (1995). UBF, an architectural element for RNA polymerase I promoters. In ‘‘Nucleic Acids and Molecular Biology’’ (F. Eckstein and D. M. J. Lilley, Eds.), Vol. 9, pp. 251–263. Springer, Berlin. Rendon, M. C., Rodrigo, R. M., Goenechea, L. G., Garcia-Herdugo, G., Valdivia, M. M., and Moreno, F. J. (1992). Characterization and immunolocalization of a nucleolar antigen with anti-NOR serum in HeLa cells. Exp. Cell Res. 200, 393–403.


173

Reichel, R., Monstein, H.-J., Jansen, H. W., Philipson, L., and Benecke, B.-J. (1982). Small nuclear RNAs are encoded on the nontranscribed region of ribosomal spacer DNA. Proc. Natl. Acad. Sci. USA 79, 3106–3110. Ritossa, F. (1976). The bobbed locus. In ‘‘The Genetics and Biology of Drosophila’’ (M. Ashburner and E. Novitski, Eds.), Vol. 1b, pp. 801–846. Academic Press, London. Robert-Fortel, I., June´ra, H. R., Ge´raud, G., and Hernandez-Verdun, D. (1993). Threedimensional organization of the ribosomal genes and Ag-NOR proteins during interphase and mitosis in PtK1 cells studied by confocal microscopy. Chromosoma 102, 146–157. Rose, K. M., Szopa, J., Han, F., Cheng, Y.-C., Richter, A., and Scheer, U. (1988). Association of DNA topoisomerase I and RNA polymerase I: A possible role for topoisomerase I in ribosomal gene transcription. Chromosoma 96, 411–416. Roussel, P., and Hernanduz-Verdun, D. (1994). Identification of Ag-NOR proteins, markers of proliferation related to ribosomal gene activity. Exp. Cell Res. 214, 465–472. Roussel, P., Andre´, C., Comai, L., and Hernandez-Verdun, D. (1996). The rDNA transcription machinery is assembled during mitosis in active NORs and absent in inactive NORs. J. Cell Biol. 133, 235–246. Roussel, P., Andre, C., Masson, C., Geraud, G., and Hernandez-Verdun, D. (1993). Localization of the RNA polymerase I transcription factor hUBF during the cell cycle. J. Cell Sci. 104, 327–337. Savino, T. M., Bastos, R., Jansen, E., and Hernandez-Verdun, D. (1999). The nucleolar antigen Nop52, the human homologue of the yeast ribosomal RNA processing RRP1, is recruited at late stages of nucleologenesis. J. Cell Sci. 112, 1889–1900. Savino, T. M., Ge´brane-Youne`s, J., De May, J., Sibarita, J.-B., and Hernandez-Verdun, D. (2001). Nucleolar assembly of the rRNA processing machinery in living cells. J. Cell Biol. 153, 1097–1110. Scheer, U., and Dabauvalle, M.-C. (1985). Functional organization of the amphibian oocyte nucleus. ‘‘Developmental Biology, A Comprehensive Synthesis’’ Vol. 1, pp. 385–430. Plenum, New York. Scheer, U., and Rose, K. M. (1984). Localization of RNA polymerase I in interphase cells and mitotic chromosomes by light and electron microscopic immunocytochemistry. Proc. Natl. Acad. Sci. USA 81, 1431–1435. Scheer, U., Thiry, M., and Goessens, G. (1993). Structure, function and assembly of the nucleolus. Trends Cell Biol. 3, 236–241. Scheer, U., Trendelenburg, M. F., and Franke, W. W. (1975). EVects of actinomycin D on the association of newly formed ribonucleoproteins with the cistrons of ribosomal RNA in Triturus oocytes. J. Cell Biol. 65, 163–179. Scheer, U., and Weisenberger, D. (1994). The nucleolus. Curr. Opin. Cell Biol. 6, 354–359. Scherl, A., Coute´, Y., Deón, C., Calle´, A., Kindbeiter, K., Sanchez, J.-C., Greco, A., Hochstrasser, D., and Diaz, J.-J. (2002). Functional proteomic analysis of human nucleolus. Mol. Biol. Cell 13, 4100–4109. Schmidt-Zachmann, M. S., Hugle-Dorr, B., and Franke, W. W. (1987). A constitutive nucleolar protein identified as a member of the nucleoplasmin family. EMBO J. 6, 1881–1890. Schwarzacher, H. G., Mikelsaar, A. V., and Schnedl, W. (1978). The nature of the Ag-staining of nucleolus organizer regions. Electron and light-microscopic studies on human cells in interphase, mitosis, and meiosis. Cytogen. Cell Gen. 20, 24–39. Schwarzacher, H. G., and Wachtler, F. (1993). The nucleolus. Anat. Embryol. 188, 515–536. Semeshin, V. F., Sherudilo, A. I., and Belyaeva, E. S. (1975). Nucleoli formation under inhibited RNA synthesis. Exp. Cell Res. 93, 458–467.

174

PATRICK J. DIMARIO

Shah, S. B., Terry, C. D., Wells, D. A., and DiMario, P. J. (1996). Structural changes in oocyte nucleoli of Xenopus laevis during oogenesis and meiotic maturation. Chromosoma 105, 111–121. Sharma, K., and Tollervey, D. (1999). Base pairing between U3 small nucleolar RNA and the 50 end of 18S is required for pre-rRNA processing. Mol. Cell Biol. 19, 6012–6019. Shaw, P. J., and Jordan, E. G. (1995). The nucleolus. Annu. Rev. Cell Dev. Biol. 11, 93–121. Sheldon, S., Speers, W. C., and Lehman, J. M. (1981). Nucleolar persistence in embryonal carcinoma cells. Exp. Cell Res. 132, 185–192. Shi, L. J., Ni, Z. M., Zhao, S., Wang, G., and Yang, Y. (1987). Involvement of a nucleolar component, perichromonucleolin, in the condensation and decondensation of chromosomes. Proc. Natl. Acad. Sci. USA 84, 7953–7956. Shou, W., Seol, J. H., Shevchenko, A., Baskerville, C., Moazed, D., Chen, Z. W., Jang, J., Shevchenko, A., Charbonneau, H., and Deshaies, R. J. (1999). Exit from mitosis is triggered by Tem1-dependent release of the protein phosphatase Cdc14 from nucleolar RENT complex. Cell 97, 233–244. Sinclair, D. A., and Guarente, L. (1997). Extrachromosomal rDNA circles—A cause of aging in yeast. Cell 91, 1–20. Sirri, V., Hernandez-Verdun, D., and Roussel, P. (2002). Cyclin-dependent kinases govern formation maintenance of the nucleolus. J. Cell Biol. 156, 969–981. Sirri, V., Roussel, P., and Hernadez-Werdun, D. (1999). The mitotically phosphorylated form of the transcription termination factor TTF-1 is associated with the repressed rDNA transcription machinery. J. Cell Sci. 112, 3259–3268. Sirri, V., Roussel, P., and Hernandez-Verdun, D. (2000). In vivo release of mitotic silencing of ribosomal gene transcription does not give rise to precursor ribosomal RNA processing. J. Cell Biol. 148, 259–270. Smetana, K., and Busch, H. (1974). The nucleolus and nucleolar DNA. In ‘‘The Cell Nucleus’’ (H. Busch, Ed.), Vol. 1, pp. 73–147. Academic Press, New York. Smith, J. S., and Boeke, J. D. (1997). An unusual form of transcriptional silencing in yeast ribosomal DNA. Genes Dev. 11, 241–254. Spector, D. L., Ochs, R. L., and Busch, H. (1984). Silver staining, immunofluorescence, and immunoelectron microscopic localization of nucleolar phosphoproteins B23 and C23. Chromosoma 90, 13–148. Spring, H., Meissner, B., Fischer, R., Mouzaki, D., and Trendelenburg, M. F. (1996). Special arrangement of intra-nucleolar rDNA chromatin in amplified Xenopus oocyte nucleoli: Structural changes precede the onset of rDNA transcription. Int. J. Dev. Biol. 40, 263–272. St. Johnston, D. (1993). Pole plasm and the posterior group genes. In ‘‘The Development of Drosophila melanogaster’’ (M. Bate and A. Marinez-Aries, Eds.), Vol. 1, pp. 325–363. Cold Spring Harbor Press, Cold Spring Harbor, NY. Steele, R. E., Thomas, P. S., and Reeder, R. H. (1984). Anucleolate frog embryos contain ribosomal DNA sequences and a nucleolar antigen. Dev. Biol. 102, 409–416. Stevens, B. J. (1965). The fine structure of the nucleolus during mitosis in the grasshopper neuroblast cell. J. Cell Biol. 24, 349–368. Straight, A. F., Shou, W., Dowd, G. J., Turck, C. W., Deshaies, R. J., Johnson, A. D., and Moazed, D. (1999). Net1, a Sir2-associated nucleolar protein required for rDNA silencing and nucleolar integrity. Cell 97, 245–256. Strasburger, E. (1882). Ueber den Theilungsvorgang der Zellkerne und das Verha¨ltniss der Kerntheilung zur Zelltheilung. Arch. Mikroskop. Anat. 21, 476–588. Strasburger, E. (1893). Zu dem jetzigen Stande der Kern- und Zelltheilngsfragen. Anat. Anz. 8, 177–191.


175

¨ ber cytoplasmastrukturen, Kern- und Zelltheilung. Jahrb. f. wiss. Bot. Strasburger, E. (1897). U 30, 221–251. Suja, J., and Hernandez-Verdun, D. (1996). The Ag-NOR proteins present a crescent-shaped distribution at the secondary constriction of metaphase PtK1 chromosomes. Cytogenet. Cell Genet. 75, 155–161. Suja, J. A., Ge´brane-Youne`s, J., Ge´raud, G., and Hernandez-Verdun, D. (1997). Relative distribution of rDNA and proteins of the RNA polymerase I transcription machinery at chromosomal NORs. Chromosoma 105, 459–469. Sullivan, G. J., Bridger, J. M., Cuthbert, A. P., Newbold, R. F., Bickmore, W. A., and McStay, B. (2001). Human acrocentric chromosomes with transcriptionally silent nucleolar organizer regions associate with nucleoli. EMBO J. 20, 2867–2877. Sumner, A. T. (1990). ‘‘Chromosome Banding.’’ Unwin Hyman, London. Sumner, A. T. (2003). ‘‘Chromosomes: Organization and Function.’’ Blackwell, Malden, MA. Sun, X., Zhao, J., Jin, S., Palka, K., Visa, N., Aissouni, Y., Daneholt, B., and AlzhanovaEricsson, A. T. (2002). A novel protein localized to the fibrillar compartment of the nucleolus and to the brush border of a secretory cell. Eur. J. Cell Biol. 81, 125–137. Szebeni, A., and Olson, M. O. J. (1999). Nucleolar protein B23 has molecular chaperone activities. Protein Sci. 8, 905–912. Szebeni, A., Hingorani, K., Negi, S., and Olson, M. O. J. (2003). Role of protein kinase CK2 phosphorylation in the molecular chaperone activity of nucleolar protein B23. J. Biol. Chem. 278, 9107–9115. Szebeni, A., Mehrotra, B., Baumann, A., Adam, S. A., Wingfield, P. T., and Olson, M. O. J. (1997). Nucleolar protein B23 stimulates nuclear import of the HIV-1 Rev protein and NLSconjugated albumin. Biochemistry 36, 3941–3949. Tashiro, K., Shiokawa, K., Yamana, K., and Sakaki, Y. (1986). Structural analysis of ribosomal DNA homologous in nucleolus-less mutant of Xenopus laevis. Gene 44, 299–306. Thiry, M. (1992). Highly sensitive immunodetection of DNA on sections with exogenous terminal deoxynucleotidyl transferase and nonisotopic nucleotide analogues. J. Histochem. Cytochem. 40, 411–419. Thiry, M., and Goessens, G. (1996). ‘‘The Nucleolus During the Cell Cycle.’’ Springer, Heidelberg. Thiry, M., Ploton, D., Menager, M., and Goessens, G. (1993). Ultrastructural distribution of DNA within the nucleolus of various animal cell lines or tissues revealed by terminal deoxynucleotidyl transferase. Cell Tissue Res. 271, 33–45. Tobler, H. (1975). Occurrence and developmental significance of gene amplification. In ‘‘Biochemistry of Animal Development’’ (R. Weber, Ed.), Vol. 3, pp. 91–143. Academic Press, New York. Tollervey, D., and Kiss, T. (1997). Function and synthesis of small nucleolar RNAs. Curr. Opin. Cell Biol. 9, 337–342. Tollervey, D., Lehtonen, H., Jansen, R., Kern, H., and Hurt, E. C. (1993). Temperaturesensitive mutations demonstrate roles for yeast fibrillarin in pre-rRNA processing, pre-rRNA methylation, and ribosome assembly. Cell 72, 443–457. Traverso, E. E., Baskerville, C., Liu, Y., Shou, W., James, P., Deshaies, R. J., and Charbonneau, H. (2001). Characterization of the Netl cell cycle-dependent regulator of the Cdc14 phosphatase from budding yeast. J. Biol. Chem. 276, 21924–21931. Trere`, D., Pession, A., and Derenzini, M. (1989). The silver-stained proteins of interphasic nucleolar organizer regions as a parameter of cell duplication rate. Exp. Cell Res. 184, 131–137. Trumtel, S., Le´ger-Silvestre, I., Gleizes, P.-W., Teulie`res, F., and Gas, N. (2000). Assembly and functional organization of the nucleolus: ultrastructural analyses of Saccharomyces cerevisiae mutants. Mol. Biol. Cell 11, 2175–2189.

176

PATRICK J. DIMARIO

Tschochner, H., and Hurt, E. (2003). Pre-ribosomes on the road from the nucleus to the cytoplasm. Trends Cell Biol. 13, 255–263. Tycowski, K. T., and Steitz, J. A. (2001). Non-coding snoRNA host genes in Drosophila: Expression strategies for modification guide snoRNAs. Eur. J. Cell Biol. 80, 119–125. Valdez, B. C., Henning, D., Le, T. V., and Busch, H. (1995). Specific aspartic acid-rich sequences are responsible for silver staining nucleolar proteins. Biochem. Biophys. Res. Comm. 207, 485–491. Valdez, B. C., Perlaky, L., Henning, D., Saijo, Y., Chan, P. K., and Busch, H. (1994). Identification of the nuclear and nucleolar localization signals of the protein p120. Interaction with translocation protein B23. J. Biol. Chem. 269, 23776–23783. Van Camp, G. M. (1924). Le roˆle du nucleóle dans la caryocine`se somatique. La Cellule 34, 5–49. Van Gansen, P., and Schram, A. (1972). Evolution of the nucleoli during oogenesis in Xenopus laevis studied by electron microscopy. J. Cell Sci. 10, 339–367. Verheggen, C., Almouzni, G., and Hernandez-Verdun, D. (2000). The ribosomal RNA processing machinery is recruited to the nucleolar domain before RNA polymerase I during Xenopus laevis development. J. Cell Biol. 149, 293–305. Verheggen, C., Le Panse, S., Almouzni, G., and Hernandez-Verdun, D. (1998). Presence of prerRNAs before activation of polymerase I transcription in the building process of nucleoli during early development of Xenopus laevis. J. Cell Biol. 142, 1167–1180. Verheggen, C., Le Panse, S., Almouzni, G., and Hernandez-Verdun, D. (2001). Maintenance of nucleolar machineries and pre-rRNAs in remnant nucleolus of erythrocyte nuclei and remodeling in Xenopus egg extracts. Exp. Cell Res. 269, 23–34. Verheijen, R., Kuijpers, H. J. H., van Driel, R., Beck, J. L. M., van Dierendonck, J. H., BrakenhoV, G. J., and Ramaekers, F. C. S. (1989). Ki-67 detects a nuclear matrix-associated proliferation-related antigen. II. Localization in mitotic cells and association with chromosomes. J. Cell Sci. 92, 531–540. Vincent, W. S. (1955). Structure and chemistry of nucleoli. Int. Rev. Cytol. 4, 269–298. Visintin, R., and Amon, A. (2000). The nucleolus: The magician’s hat for cell cycle tricks. Curr. Opin. Cell Biol. 12, 372–377. Visintin, R., Hwang, E. S., and Amon, A. (1999). Cfi1 prevents premature exit from mitosis by anchoring Cdc14 phosphatase in the nucleolus. Nature 398, 818–823. Voit, R., HoVmann, M., and Grummt, I. (1999). Phosphorylation by G1-specific cdk-cyclin complexes activates the nucleolar transcription factor UBF. EMBO J. 18, 1891–1899. Voit, R., Kuhn, A., Sander, E. E., and Grummt, I. (1995). Activation of mammalian ribosomal gene transcription requires phosphorylation of the nucleolar transcription factor UBF. Nucleic Acids Res. 23, 2593–2599. Voit, R., Scha¨fer, K., and Grummt, I. (1997). Mechanism of repression of RNA polymerase I transcription by retinoblastoma protein. Mol. Cell. Biol. 17, 4230–4237. Voit, R., Schnapp, A., Kuhn, A., Rosenbauer, H., Hirschmann, P., Stunnenberg, H. G., and Grummt, I. (1992). The nucleolar transcription factor mUBF is phosphorylated by casein kinase in the C-terminal hyperacidic tail which is essential for transactivation. EMBO J. 11, 2211–2218. Wachtler, F., and Stahl, A. (1993). The nucleolus—A structural and functional interpretation. Micron 24, 473–505. Wachtler, F., Hartung, M., Devictor, M., Wiegant, J., and Stahl, A. (1989). Ribosomal DNA is located and transcribed in the dense fibrillar component of human Sertoli-cell nucleoli. Exp. Cell Res. 184, 61–71. Waggener, J. M., and DiMario, P. J. (2002). Two splice variants of Nopp140 in Drosophila melanogaster. Mol. Biol. Cell 13, 362–381.


177

Wallace, H. (1963). Nucleolar growth and fusion during cellular diVerentiation. J. Morphol. 112, 261–278. Wallace, H., and Birnstiel, M. L. (1966). Ribosomal cistrons and the nucleolar organizer. Biochim. Biophys. Acta 114, 296–310. Wang, C., Query, C. C., and Meier, U. T. (2002). Immunopurified small nucleolar ribonucleoprotein particles pseudouridylate rRNA independently of their association with phosphorylated Nopp140. Mol. Cell. Biol. 22, 8457–8466. Wang, H., Boisvert, D., Kim, K. K., Kim, R., and Kim, S. H. (2000). Crystal structure of a ˚ fibrillarin homologue from Methanococcus jannaschii, a hyperthermophile, at 1.6 A resolution. EMBO J. 19, 317–323. Warner, J. R. (1990). The nucleolus and ribosome formation. Curr. Opin. Cell Biol. 2, 521–527. Watkins, N. J., Segault, V., Charpentier, B., Nottrott, S., Fabrizio, P., Bachi, A., Wilm, M., Roshbash, M., Branlant, C., and Luhrmann, R. (2000). A common core RNP structure shared between the small nucleolar Box C/D RNPs and the spliceosomal U4 snRNP. Cell 103, 457–466. Weisenberger, D., and Scheer, U. (1995). A possible mechanism for the inhibition of ribosomal RNA gene transcription during mitosis. J. Cell Biol. 129, 561–575. Weisenberger, D., Scheer, U., and Benavente, R. (1993). The DNA topoisomerase I inhibitor camptothecin blocks postmitotic reformation of nucleoli in mammalian cells. Eur. J. Cell Biol. 61, 189–192. White, R. J., Gottlieb, T. M., Downes, C. S., and Jackson, S. P. (1995). Mitotic regulation of a TATA-binding-protein-containing complex. Mol. Cell. Biol. 15, 1983–1992. Wilson, E. B. (1925). ‘‘The Cell in Development and Heredity.’’ Macmillan, New York. Woolford, J. L., Jr., and Warner, J. R. (1991). The ribosome and its synthesis. In ‘‘The Molecular and Cellular Biology of the Yeast Saccharomyces: Genome Dynamics, Protein Synthesis, and Energetics’’ (J. R. Broach, J. R. Pringle, and E. W. Jones, Eds.), Vol. 1, pp. 587–626. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Xu, C., Henry, P. A., Setya, A., and Henry, M. F. (2003). In vivo analysis of nucleolar proteins modified by the yeast arginine methyltransferase Hmt1/Rmt1p. RNA 9, 746–759. Yang, Y., Isaac, C., Wang, C., Dragon, F., Pogacic, V., and Meier, U. T. (2000). Conserved composition of mammalian BoxH/ACA and Box C/D small nucleolar ribonucleoprotein particles and their interaction with the common factor Nopp140. Mol. Biol. Cell 11, 567–577. Yasuda, Y., and Maul, G. G. (1990). A nucleolar auto-antigen is part of a major chromosomal surface component. Chromosoma 99, 152–160. Yu, Y.-T., Scharl, E. C., Smith, C. M., and Steitz, J. A. (1999). The growing world of small nuclear ribonucleoproteins. In ‘‘The RNA World’’ (R. F. Gesteland, T. R. Cech, and J. F. Atkins, Eds.), 2nd ed., pp. 487–524. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Yu, Y.-T., Shu, M. D., Narayanan, A., Terns, R. M., Terns, M. P., and Steitz, J. A. (2001). Internal modification of U2 small nuclear (sn)RNA occurs in nucleoli of Xenopus oocytes. J. Cell Biol. 152, 1279–1288. ZaVran, S., Chartier, A., Gallant, P., Astier, M., Arquier, N., Doherty, D., Gratecos, D., and Se´me´riva, M. (1998). A Drosophila RNA helicase gene, pitchoune, is required for cell growth and proliferation and is a potential target of d-Myc. Development 125, 3571–3584. Zatsepina, O., Baly, C., Chebrout, M., and Debey, P. (2003). The step-wise assembly of a functional nucleolus in preimplantation mouse embryos involves the Cajal (coiled) body. Dev. Biol. 253, 66–83.

178

PATRICK J. DIMARIO

Zatsepina, O. V., Bounio-Baly, C., and Amirand, C. (2000). Functional and molecular reorganization of the nucleolar apparatus in maturing mouse occytes. Dev. Biol. 223, 354–370. Zatsepina, O. V., Rousselet, A., Chan, P. K., Olson, M. O. J., Jordan, E. G., and Bornens, M. (1999). The nucleolar phosphoprotein B23 redistributes in part to the spindle poles during mitosis. J. Cell Sci. 112, 455–466. Zatsepina, O. V., Voit, R., Grummt, I., Spring, H., Semenov, M. V., and Trendelenburg, M. F. (1993). The RNA polymerase I-specific transcription initiation factor UBF is associated with transcriptionally active and inactive ribosomal genes. Chromosoma 102, 599–611. Zebarjadian, Y., King, T., Fournier, M. J., Clarke, L., and Carbon, J. (1999). Point mutations in yeast CBF5 can abolish in vivo pseudouridylation of rRNA. Mol. Cell. Biol. 19, 7461–7472. Zhang, H., Wang, J. C., and Liu, L. (1988). Involvement of DNA topoisomerase I in transcription of human ribosomal RNA genes. Proc. Natl. Acad. Sci. USA 85, 1060–1064. Zhu, Y., Lu, D., and DiMario, P. J. (1999). Nucleolin, defective for MPF phosphorylation, localizes normally during mitosis and nucleologenesis. Histochem. Cell Biol. 111, 477–487.

MAPping the Eukaryotic Tree of Life: Structure, Function, and Evolution of the MAP215/Dis1 Family of Microtubule-Associated Proteins David L. Gard, Bret E. Becker, and S. Josh Romney Department of Biology, University of Utah, Salt Lake City, Utah 84112-0840

The MAP215/Dis1 family of proteins is an evolutionarily ancient family of microtubule-associated proteins, with characterized members in all major kingdoms of eukaryotes, including fungi (Stu2 in S. cerevisiae, Dis1 and Alp14 in S. pombe), Dictyostelium (DdCP224), plants (Mor1 in A. thaliana and TMBP200 in N. tabaccum), and animals (Zyg9 in C. elegans, Msps in Drosophila, XMAP215 in Xenopus, and ch-TOG in humans). All MAP215/Dis1 proteins (with the exception of those in plants) localize to microtubule-organizing centers (MTOCs), including spindle pole bodies in yeast and centrosomes in animals, and all bind to microtubules in vitro and/or in vivo. Diverse roles in regulating microtubule assembly and organization have been proposed for individual family members, and a substantial body of evidence suggests that MAP215/Dis1-related proteins play critical roles in the assembly and function of the meiotic/mitotic spindles and/ or cell division. An extensive search of public databases (including both EST and genome databases) identified partial sequences predicted to encode more than three dozen new members of the MAP215/Dis1 family, including putative MAP215/Dis1-related proteins in Giardia lamblia and four other protists, sixteen additional species of fungi, six plants, and twelve animals. The structure and function of MAP215/Dis1 proteins are discussed in relation to the evolution of this ancient family of microtubule-associated proteins. KEY WORDS: Microtubules, Microtubule-associated proteins, MAPs, Dis1, Alp14, Stu2, DdCP224, Zyg9, Minispindles, Msps, XMAP215, ch-TOG, Mor1, Gem1, TACC proteins. ß 2004 Elsevier Inc.

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I. Introduction Microtubules (MTs), composed of a- and b-tubulin heterodimers, play a number of roles in processes that are critical for the structure, function, and survival of eukaryotic cells. Perhaps foremost among these processes is the assembly and function of the spindle, a complex MT-based machine that is responsible for the segregation of chromosomes during the meiotic and mitotic divisions of eukaryotes. Individual MTs are highly dynamic structures, undergoing alternating phases of elongation and rapid shortening, with transitions between these phases termed catastrophe (elongation to rapid shortening) and rescue (rapid shortening to elongation) (Desai and Mitchison, 1997; Mitchison and Kirschner, 1984). A third state, in which a MT neither grows nor shrinks, but remains constant in length, termed a ‘‘pause,’’ has also been described (Tran et al., 1997). This behavior, termed ‘‘dynamic instability,’’ has been observed both with purified tubulin subunits in vitro (Horio and Hotani, 1986; Walker et al., 1988) and in living cells (Cassimeris et al., 1988; Joshi, 1998; Sammak and Borisy, 1988; Shelden and Wadsworth, 1993). However, the dynamic behavior of individual MTs in living cells diVers markedly from that of MTs assembled from pure subunits in vitro (Desai and Mitchison, 1997). Although the basic structure and dynamics of MTs are inherently derived from the tubulin subunits themselves, it has become increasingly clear that the dynamics, organization, and function of MTs are modulated by a diverse array of proteins collectively referred to as microtubule-associated proteins, or MAPs (Cassimeris and Spittle, 2001). The first MAPs to be identified and characterized, including the Tau, MAP2, and MAP1 families from vertebrate brain, dampened or suppressed the inherent dynamic instability of MTs (Drechsel et al., 1992; Kowalski and Williams, 1993; Pryer et al., 1992), consistent with the long, stable, MTs found in neuronal axons and dendrites. However, the stabilizing functions of these neuronal MAPs were not suYcient to explain the complex organization, rapid reorganization, and function of MT arrays during the division cycle of nonneuronal cells. Subsequent studies have since identified several distinct classes of MAPs that have diverse roles in regulating the assembly, organization, and disassembly of MT arrays, including nucleation factors (g-tubulin and its associated proteins in the g-TuRC; Moritz and Agard, 2001); monomer binding proteins such as OP18/stathmin that inhibit MT assembly or induce catastrophe (Cassimeris, 2002; Lawler, 1998); MAPs that promote MT assembly and stability in a manner similar to neuronal MAPs (MAP4 is an example) (Cassimeris and Spittle, 2001); end-binding proteins that stabilize or destabilize MTs (Schroer, 2001; Schuyler and Pellman, 2001); catastrophe factors including the kin I family of kinesin-related proteins (reviewed in Desai et al.,

THE MAP215/Dis1 FAMILY

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FIG. 1 XMAP215 dramatically promotes assembly of tubulin from centrosomes in vitro. (A) Small asters, consisting of a only a few, short MTs, are assembled from 0.9 mg/mL phosphocellulose-purified bovine brain tubulin (5 min at 37 C). (B) At the same tubulin concentration, addition of 0.02 mg/mL XMAP215 purified from activated Xenopus eggs (corresponding to a 1:90 molar ratio of XMAP215:TB) dramatically stimulates MT elongation. Bar is 10 mm. Reproduced from Gard and Kirschner (1987) by permission.

1999); severing proteins, such as katanin (McNally and Vale, 1993; Quarmby, 2000); and the MAP215/Dis11 family of MT regulators: an evolutionarily conserved family of proteins that regulate MT assembly, dynamics, or function, related to XMAP215 in Xenopus laevis and Dis1 in Schizosaccaromyces pombe (Kinoshita et al., 2002; Ohkura et al., 2001). All of these classes of proteins are thought to act in concert to orchestrate the complex organization and function of MTs during the life of eukaryotic cells. However, a growing body of evidence indicates that the MAP215/Dis1 family of MAPs plays one or more critical roles in the assembly, organization, and function of MTs during the division of eukaryotic cells. In 1987, Gard and Kirschner (1987) reported the isolation and initial characterization of a novel MAP from the eggs of the African frog X. laevis. This MAP, initially named XMAP (Xenopus Microtubule-Assembly Protein) (Gard and Kirschner, 1987) exhibited the unique property of dramatically and preferentially promoting the elongation of MT plus-ends (see Fig. 1). On the basis of their initial characterization of XMAP function, the authors 1

This family of proteins has previously been referred to as the Dis1/TOG or XMAP215/TOG family. MAP215/Dis1 will be used throughout this discussion, recognizing the initial two family members to be identified and characterized, but dropping the organism specifying ‘‘X.’’ XMAP215 will hereinafter refer specifically to the protein family member in Xenopus (X. laevis and X. tropicalis).

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proposed that this novel MAP might play a role in the rapid assembly and organization of MTs during the processes of fertilization and the rapid division cycles of the early frog embryo (Gard and Kirschner, 1987). Since the identification of additional MAPs in Xenopus eggs (particularly those now known as XMAP230 and XMAP310) (Andersen and Karsenti, 1997; Andersen et al., 1994), XMAP has been referred to as XMAP215, denoting its species of origin and apparent molecular mass determined by SDS-PAGE. Nearly coincident with the initial description of XMAP215, Ohkura et al. (1988) reported the identification and initial genetic characterization of coldsensitive mutations of the DIS1 gene in the fission yeast S. pombe. Yeast cells containing mutations in the DIS1 gene exhibit defects in sister chromatid segregation and arrest in mitosis (Ohkura et al., 1988). Subsequent studies revealed that the DIS1 gene encodes a 93-kDa protein (Dis1) associated with spindle MTs and spindle pole bodies (SPBs), which was implicated in several aspects of spindle function (Nabeshima et al., 1995; Nakaseko et al., 1996; see Section III.A). The serendipitous cloning of ch-TOG (colonic-hepatic Tumor Overexpressed Gene), a cDNA encoding a human protein overexpressed in tumors, during a screen for cytochrome p450 (Charrasse et al., 1995) forged the link that bound together these heretofore distinct families of MAPs in such highly diverse eukaryotic taxa as yeast and humans. Preliminary characterization revealed that ch-TOG protein was expressed at low levels in all cells but was abundant in neurons and (as its name implies) tumor cells (Charrasse et al., 1995, 1996). On the basis of peptide sequences and immunological cross reaction, Charrasse et al. (1998) proposed that ch-TOG was the human homolog of XMAP215, and subsequent reports revealed that Dis1 and Alp14, Stu2, DdCP224, Zyg9,2 and minispindles (Msps) were XMAP215/ TOG homologs in S. pombe (Garcia et al., 2001; Nakaseko et al., 2001), Saccharomyces cerevisiae (Wang and HuVaker, 1997), Dictyostelium discoideum (Graf et al., 2000), Caenorhabditis elegans (Matthews et al., 1998), and Drosophila (Matthews et al., 1998), respectively. Since those initial reports, it has become clear that the MAP215/Dis1 family represents an ancient family of MAPs, perhaps extending to the very origin of eukaryotes. The 10 MAP215/Dis1-related proteins that have been characterized to date (Dis1 and Alp14 in S. pombe, Stu2 in S. cerevisiae, DdCP224 in Dictyostelium, Zyg9 in C. elegans, Msps in Drosophila, XMAP215 in Xenopus, 2 Identification of the Zyg9 locus in Caenorhabditis elegans was first reported in 1986 (Kemphues, 1986). Thus, Zyg9 might be considered one of the founding members of the MAP215/Dis1 family. However, characterization of the Zyg9 gene product was not reported until 1998 (Matthews et al., 1998), eleven years after the identification and characterization of the XMAP215 in Xenopus (Gard and Kirschner, 1987), and three years after characterization of p93Dis1 protein in S. pombe (Nabeshima et al., 1995).


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ch-TOG in humans, Mor1 in A. thaliana, and TMBP200 in Nicotiana tabaccum) share commonality of both structure and function: All are composed of a core consisting of two or four TOG domains (Kinoshita et al., 2002; Ohkura et al., 2001), with each domain composed of as many as five HEAT repeats (Neuwald and Hirano, 2000), and all have been implicated in regulating the assembly, dynamics, organization, or function of MTs, particularly those of the meiotic and mitotic spindles. Several recent reviews have discussed the unifying structural features and diverse functions of the MAP215/Dis1 family of proteins (Kinoshita et al., 2002; Ohkura et al., 2001). The following discussion will reconsider the structure and function of the MAP215/Dis1-related proteins in light of the identification of more than 35 new family members encoded by sequences identified in the partial or complete genomes from protists, fungi, animals, and plants. The purpose of this discussion is twofold: to review and consolidate what is known about the structures and functions of MAP215/Dis1-related proteins in all taxa in which they have been identified to date, and to highlight what is not yet known, propose models for structure or function based on evolutionarily conserved features of MAP215/Dis1-related proteins, and thereby stimulate new avenues of study.

A. MAP215/Dis1 Family: Deeply Rooted in the Eukaryote Tree of Life To date, 10 members of the MAP215/Dis1 family of proteins, spanning taxa from yeast to vertebrates, have been identified and functionally characterized using a combination of biochemical, cell biological, or genetic approaches. However, more than three dozen additional related proteins were identified by searching databases from the numerous genome sequencing and expressed sequence tag (EST) projects being carried out worldwide (Appendix I). SuYcient sequence information is available to map more than 30 of these predicted proteins, along with the more well-characterized family members, within the MAP215/Dis1 family tree (see Fig. 2). The MAP215/Dis1 family is deeply rooted in the eukaryotic branch of the tree of life. Sequence alignment and comparison of the first 750 amino acids of 46 proteins reveals deeply rooted branches corresponding to fungal proteins, including Stu2 (S. cerevisiae), Dis1 (S. pombe), and Alp14 (S. pombe); animals, including the protist D. discoideum (DdCP224), nematodes (Zyg9 in C. elegans), insects (Msps in Drosophila meldnogaster), and chordates (XMAP215 and ch-TOG in X. laevis and Homo sapiens, respectively); and plants (Mor1/Gem1 in A. thaliana and TMBP200 in N. tabacum). Each of these major branches may contain two or more monophyletic subfamilies (Fig. 2) (Sections III and V), which will be discussed in more detail later.

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FIG. 2 The MAP215/Dis1 family of MT-associated proteins is deeply rooted in the eukaryotic branch of the tree of life. About 700 amino acids (including TOG domains 1A and 1B) from 46 characterized (bold) and putative members of the MAP215/Dis1 family were aligned with ClustalW (using the PAM215 weight table; DNAStar, Lasergene, Inc.). Phylogenetic trees were


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In addition to the fungal, animal, and plant branches, five putative family members have been identified in protists, including Giardia lamblia, Entamoeba histolytica, trypanosomes (T. brucei and T. cruzzi), and a microsporidian (Encephilitazooan cuniculi). Although the proteins predicted from the genomes of these more primitive eukaryotes are the most divergent members of the MAP215/Dis1 family of the proteins and have yet to be ascribed a function, they share many of the structural features characteristic of bona fide members of the MAP215/Dis1 family.

B. MAP215/Dis1 Proteins: A Common Structural Organization 1. N-Terminal HEAT Repeats Clustered into TOG Domains MAP215/Dis1-related proteins range from fewer than 900 amino acids to more than 2000 amino acids in length. Most are enriched in basic amino acids. The calculated isoelectric points for the characterized family members range from 7.5 (Mor1 in A. thaliana and Zyg9 in C. elegans) to 8.6 (Stu2 in S. cerevisiae). Sequence analysis and structural predictions indicate that all members of the MAP215/Dis1 family share a common core structure consisting of 10–25 HEAT repeats (Fig. 3) (Neuwald and Hirano, 2000). HEAT (HuntingtinElongation A-TOR) repeats consist of poorly conserved 35–50–amino acid domains characterized by tandemly arranged a-helices with conserved hydrophobic or charged residues found in a number of eukaryotic proteins (reviewed in Andrade and Bork, 1995; Andrade et al., 2001a,b), where they are thought to provide a flexible protein scaVold for protein–protein interactions (Andrade et al., 2001a,b; Groves et al., 1999; Kobe et al., 1999; Neuwald and Hirano, 2000). HEAT repeats in MAP215/Dis1-related proteins are clustered into TOG domains: a higher-order sequence repeat approximately 240 amino acids in length that includes up to five individual HEAT repeats (Kinoshita et al., 2002; Ohkura et al., 2001). The N-terminal region of MAP215/Dis1 family members consist of two or four TOG domains organized as tandem repeats. Pairwise sequence comparisons of all four TOG domains (200 amino acids) from 15 species (three basidiomycote fungi, six animals, three plants, and three protists/ others; see Table I) revealed that the TOG domains fall into four distinct classes corresponding to their positions in the amino acid sequence (from N ! C): 1A,

constructed in PAUP (V4.0) based on parsimony. Shown is the single most parsimonious tree resulting from a heuristic search of more than 7 106 rearrangements (100 randomly seeded replicates). Branch lengths are shown above and bootstrap values (500 replicates) over 50% are shown below the branches (bootstrap values R substitution), Physcomitrella, and six species of higher plants, the target serine has been replaced by either glutamate or aspartate in all of these proteins, mimicking phosphorylation. Phosphomimetic substitutions are also present in the more highly diverged sequences identified in the MAP215/Dis1-related proteins of Drosophila (Msps in D. melanogaster and a Msps-related protein in D. pseudoobscura)

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and E. histolytica. All of the divergent Tau repeats in which the conserved serine has been substituted with a nonphosphorylatable amino acid (UMMAP215, DdCP224, Msps, and all plants), with the exception of the more divergent sequence in E. histolytica, contain one or more nearby serine or threonine residues. The potential phosphorylation of the Tau repeat in MAP215/Dis1 proteins by MARK/PAR1 kinases, and its role in the regulation of MAP215/Dis1 protein function, begs further investigation.

II. Structure of MAP215/Dis1-Related Proteins in Protists Five predicted proteins identified by genome BLAST searches fall outside of the major branches of the XMAP215/Dis1 family: one protein each in the diplomonad Giardia lamblia, the microsporidian E. cuniculi, and E. histolytica, and two family members (one each) in the kinetoplastids T. cruzzi and T. brucei. Although only 11–13% identical in amino acid sequence to Dis1 in S. pombe and 11–16% identical to XMAP215 in Xenopus, each of these gene products is predicted to include two or more TOG domains and one or more other sequence motifs or features found in bone fide members of the MAP215/Dis1 family of proteins (see Fig. 3). The putative MAP215/Dis1 family member in the diplomonad G. lamblia (GL-MAP215) is just 12% identical to Dis1 in S. pombe and 11% to XMAP215 in X. laevis overall. However, GL-MAP215 is predicted to include two TOG domains, each with multiple HEAT repeats. Sequence comparisons revealed that the first TOG domain of GL-MAP215 (200 amino acids) is 16–20% identical to that of the MAP215/Dis1-related proteins in euascomycote fungi, including Aspergillus and Neurospora, and that the two predicted TOG domains in GL-MAP215 correspond to A-type (signature sequence: A K Q R L) and B-type (signature sequence: W K E R K) domains characteristic of MAP215/Dis1 proteins. With a calculated isoelectric point (pI) of 7.64, the predicted GL-MAP215 protein is mildly basic, similar to other members of the MAP215/Dis1 family. However, a stretch of 85 amino acids (734–818) is enriched in basic amino acids (15% lysine and arginine), with a calculated pI of 9.1. The predicted GL-MAP215 protein lacks other sequence features or motifs found in MAP215/Dis1 family members, such as predicted coiled–coil regions or CDK1 phosphorylation targets. Both parsimony (Fig. 2) and sequence distance trees (not shown) place GL-MAP215 at the root of the MAP215/Dis1 family tree, consistent with current views that Giardia is a member of an early-branching eukaryotic lineage (Baldauf et al., 2000). The MAP215/Dis1 family member predicted from the genome of the microsporidian E. cuniculi (EC-MAP215) is just 11% identical in amino


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acid sequence to GL-MAP215, 11% identical to Dis1 in S. pombe, and 13% identical to XMAP215 in Xenopus overall. However, EC-MAP215 is predicted to include two TOG domains corresponding to types A and B, with the N terminal signature sequences W K D R Q and W K T R L, respectively. The amino acid sequence of the first and second TOG domains of ECMAP215 are 20–23% and 13–19% to the 1A and 1B domains of MAP215/ Dis1-related proteins in euascomycote fungi, respectively (EC-MAP215 TOG 1B is 20% identical to TOG 1B of Dis1). With a calculated pI of 5.73, EC-MAP215 is uncharacteristically acidic for a MAP215/Dis1 family member. However, like MAP215/Dis1-related proteins in yeast, EC-MAP215 includes a domain (amino acids 551-637) that is enriched in basic amino acids (23% lysine and arginine), with a predicted pI of 9.4. Interestingly, the predicted sequence of EC-MAP215 also includes a single copy of an 18–amino acid sequence (amino acids 587–604) that is 44% identical to the consensus sequence of the single Tau repeat found in MAP215/Dis1-related proteins in animals and plant (see Fig. 5). This putative Tau repeat in EC-MAP215 includes conserved leucines at the N- and C-terminal ends of the repeat, the core KIG sequence, and a conservative substitution of threonine for the conserved serine (position 596). Although the functional significance of this sequence has yet to be established, evidence from fungi and animals suggests that it may serve to target MAP215/ Dis1-related proteins to MTOCs (Sections IV.D.4e and IV.D.5). The incomplete genome sequences of T. cruzzi and T. brucei each encode a putative member of the MAP215/Dis1 family (TC-MAP215 and TBMAP215). As would be expected, pairwise comparison of their amino acid sequences indicates that TC-MAP215 and TB-MAP215 are more similar to each other (53% identical) than they are to other Dis1/MAP215 family members: 11–12% identical to GL-MAP215, 12% identical to Dis1 in S. pombe, and 14–15% identical to XMAP215 in Xenopus overall. However, the amino acids sequences of the first TOG domains of the two putative trypanosome MAP215/Dis1-related proteins are 20–24% identical to those of MAP215/Dis1-related proteins in euascomycote fungi and are 23% and 25% identical to a similar region in Alp14 (S. pombe). Phylogenetic analysis places the Trypanosome MAP215 proteins in a subfamily that also includes EC-MAP215. However, in contrast to ECMAP215, which contains just two TOG domains (Fig. 3), putative MAP215/Dis1 proteins in kinetoplastids are predicted to include four TOG domains, corresponding to 1A, 1B, 2A, and 2B (see Fig. 3). TOG domains 2A and 2B in TC-MAP215 and TB-MAP215 are 18–21% and 25% identical to corresponding domains in Mor1/Gem1 and TMBP200 in plants, respectively. The putative trypanosome MAP215/Dis1-related proteins also include canonical target sequences (S/T P X K/R) for the cyclin-dependent kinase

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CDK1, a master regulator of the eukaryotic cell cycle (two in TC-MAP215 and three in TB-MAP215). Finally, a single MAP215/Dis1 family member is predicted from the incomplete genome of the enteric pathogen E. histolytica (EH-MAP215). On the basis of alignment with other MAP215/Dis1 proteins, the 762 amino acids obtained for EH-MAP215 appear to be missing 10–15 amino acids from their N terminus, and an additional sequence might be missing from the C terminus. EH-MAP215 is 11% identical in amino acid sequence to GL-MAP215, 13% to Dis1 in S. pombe, and 16% to XMAP215 in X. laevis. However, The TOG domains (1A and 1B) of EH-MAP215 are 19–23% identical to TOG 1A in MAP215/Dis1 proteins of animals and plants and 21% identical to TOG 1B of puVerfish, respectively. With a calculated pI of 9.3, EH-MAP215 is characteristically basic. Moreover, a 136–amino acid sequence (513–638) is enriched in basic amino acids (23% lysine and arginine) with a calculated pI of 10.21. EH-MAP215 lacks predicted CDK targets and coiled–coil regions. Finally, a 16–amino acid sequence 602-LINELKNKIGDPKRGL-617 near the C terminus of EH-MAP215 is similar (50% sequence identity with three appropriately spaced insertions and one deletion) to the 18–amino acid Tau repeat found in other MAP215/Dis1 family members. At the core of this putative Tau repeat is the four–amino acid sequence KIGD, which is identical to the core sequence found in all MAP215/Dis1 family members in plants (Section IV). The five putative MAP215/Dis1 proteins discussed above have been identified solely from BLAST searches of incomplete genome sequences. Further investigation will be required to confirm their identities and determine whether they, in keeping with other members of the MAP215/Dis1 protein family, regulate the assembly, organization, and function of MTs or spindles. If they indeed represent bona fide MAP215/Dis1 family members, their identification in these early branching taxa of eukaryotes, in combination with the identification of MAP215/Dis1 proteins in each of the three major kingdoms of eukaryotes, indicates that the evolution of the MAP215/Dis1 family can be traced to the very origin of eukaryotes.

III. Structure and Function of the Dis1-Related Proteins in Fungi Nearly half (20 of 50) of the MAP215/Dis1-related proteins currently known or identified for this report are found in fungi, no doubt a reflection of the number of fungal genome sequencing projects and the relatively small size of fungal genomes. Three family members were originally identified in genetic screens: Dis1 and Alp14 in the archaeascomycote fission yeast


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S. pombe (Nakaseko et al., 2001; Ohkura et al., 1988; RadcliVe et al., 1998), and Stu2 in the hemiascomycote budding yeast S. cerevisiae (Wang and HuVaker, 1997). The remainder of the proteins were identified/predicted by BLAST of complete or incomplete fungal genome database using these three yeast sequences as probes. In the phylogenetic tree shown in Figure 2, Dis1 in the archaeascomycote S. pombe falls out as the most basal member of the MAP215/Dis1 family in fungi. The remaining family members all fall into one of three groups: the Alp14/Stu2 group, which includes Alp14 in S. pombe, Stu2 in the hemiascomycote S. cerevisiae, and related proteins in other hemiascomycotes (Candida albicans and six additional species of Saccharomyces); a group of MAP215/ Dis1-related proteins found in the archaeascomycote P. carinii and five genera of euascomycotes, including Aspergillus sp. and Neurospora; and a group of MAP215/Dis1-related proteins identified/predicted from the incomplete genome sequences of four basidiomycotes, including the causative agent of corn smut, Ustilago maydis (Section V.D).

A. Dis1-Related Proteins in Ascomycotes Despite substantial sequence divergence between subfamilies (Dis1 in S. pombe and Stu2 in S. cerevisiae are only 16% identical at the amino acid level), MAP215/Dis1-related proteins in ascomycotes (including the archaeascomycotes, hemisascomycotes, and euascomycotes) share a common structural organization. All ascomycote family members are 900 amino acids in length and include two TOG domains, with conserved signature sequences and up to five HEAT repeats each, within their N-terminal 500 amino acids. Sequence alignments and phylogenetic analysis reveal that the TOG domains in the Dis1-related proteins of ascomycote fungi correspond to the 1A and 1B classes. The second TOG domain (1B) is followed by a basic domain (100–150 amino acids) that is highly enriched in lysine (K) and arginine (R): the frequency of these two amino acids ranges from 12% to nearly 20%, and isoelectric points (pI) from 9.3 to 12. This basic domain (KR domain, see Fig. 7) overlaps regions required for MT binding in Dis1, Alp14, and Stu2 (Nabeshima et al., 1995; Nakaseko et al., 1996, 2001; Wang and HuVaker, 1997; see below). The KR domain precedes a 75–100–amino acid domain near the carboxy terminus (between amino acids 625–800) that is predicted to form a coiled–coil (PAIRCOIL probability scores range from a low of 0.5 [S. kluyveri] to >0.9 [most other species]; http://paircoil.lcs.mit. edu/cgi-bin/paircoil). All of the proteins are predicted to include one or more target sites for phosphorylation by cyclin-dependent kinases. Although the absolute positioning of these predicted phosphorylation sites within the amino acid sequence is not highly conserved, most are located in the

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C-terminal third of the protein sequence, and often within the KR domain. Finally, all MAP215/Dis1-related proteins in ascomycotes include a conserved sequence of 28 amino acids near their C terminus that may function to target them to SPBs (the fungal MTOC), kinetochores, or the plus-ends of MTs (see Section V.C).

B. Functional Analysis of Dis1-Related Proteins in Yeast Genetic analysis of Dis1, Alp14, and Stu2 indicates that the homology between these proteins extends beyond structure to function. The phenotype of loss-of-function mutations or deletions includes defects in the assembly or function of the mitotic spindle and defects in the assembly, organization, or dynamics of interphase MTs (Garcia et al., 2001, 2002; Kosco et al., 2001; Nabeshima et al., 1998; Nakaseko et al., 2001; Ohkura et al., 1988; Severin et al., 2001; Wang and HuVaker, 1997). All three proteins localize to spindle or astral MTs and SPBs during M phase and to cytoplasmic MTs during interphase (Garcia et al., 2001; Kosco et al., 2001; Nabeshima et al., 1995; Nakaseko et al., 1996, 2001; Usui et al., 2003; Wang and HuVaker, 1997). The fission yeast S. pombe occupies a unique position as the sole organism in which two distinct MAP215/Dis1 family members, Dis1 and Alp14, have been identified (Garcia et al., 2001; Nabeshima et al., 1995; Nakaseko et al., 2001; Ohkura et al., 1988; RadcliVe et al., 1998). Dis1 was originally identified in a genetic screen for mutations that disrupted spindle function, resulting in defects in sister chromatid segregation (Ohkura et al., 1988). Cells bearing loss of function mutations in Dis1 or Alp14 display overlapping, but not identical, defects in spindle function. Prometaphase congression movements of centromeres are suppressed, and chromosomes are maloriented on the metaphase spindles in cells bearing mutations in DIS1 (Nabeshima et al., 1998). Despite arresting in M phase with activated CDK (Kinoshita et al., 1991), dis1 cells exhibit premature and inappropriate anaphase-like spindle elongation (Nabeshima et al., 1998; Ohkura et al., 1988). On the basis of these observations, Nabeshima et al. (1998) proposed that Dis1 links kinetochore MTs to sister chromatids in the mitotic spindle. In dis1 cells, that link is uncoupled or fails to form, resulting in the observed segregation defects. The authors argue that the premature spindle elongation observed in dis1 mutants results directly from the loss of kinetochore forces that normally balance and oppose spindle elongation (Nabeshima et al., 1998). Alp14 was independently identified as a Dis1-related protein (Mtc1) in a genome search (Nakaseko et al., 2001) and in a genetic screen that identified mutations disrupting the characteristic polar growth of S. pombe cells (Alp ¼ Altered polarity) (Garcia et al., 2001; RadcliVe et al., 1998). alp14 cells


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exhibit defects in interphase MTs and assemble long, thin spindles that often buckle or break, phenotypes consistent with defects in the assembly or stabilization of cytoplasmic and spindle MTs (Garcia et al., 2001; Nakaseko et al., 2001). Although highly divergent in amino acid sequence (only 24% identical over their full length), Dis1 and Alp14 include all the structural features characteristic of MAP215/Dis1-related proteins in ascomycotes. Each includes two TOG domains (1A and 1B), a KR domain enriched in the basic amino acids lysine (K) and arginine (R), a predicted coiled–coil domain near their C terminus, multiple predicted CDK phosphorylation targets, and a conserved C-terminal domain (see Fig. 3). Results from genetic analyses suggest that Dis1 and Alp14 function in overlapping, but nonidentical, pathways. Neither gene is essential for viability (Garcia et al., 2001; Nakaseko et al., 2001; Ohkura et al., 1988), but loss of either results in mitotic defects and slowed growth. Combinations of dis1 and alp14 loss of function mutations are synthetically lethal (Garcia et al., 2001; Nakaseko et al., 2001), suggesting that they function in the same pathway. Although defects in alp14 mutants can be suppressed by overexpression of Dis1, defects in Dis1 mutants are only partially suppressed by overexpression of Alp14 (Garcia et al., 2001; Nakaseko et al., 2001). Interestingly, lossof-function dis1 mutants are cold sensitive (Nabeshima et al., 1998; Ohkura et al., 1988), whereas loss-of-function alp14 mutants are heat sensitive (Nakaseko et al., 2001), indicating that they may perform similar functions during growth under diVerent environmental conditions. Using antibodies and expression of green fluorescent protein (GFP)tagged protein, Nabeshima et al. (1995) observed Dis1 associated with cytoplasmic MTs during interphase, weakly associated with spindle MTs during M phase, and strongly localized to SPBs during anaphase. Alp14 was observed as punctae along interphase MTs, associated with kinetochore MTs of the mitotic spindle, and bound to SPBs (Garcia et al., 2001; Nakaseko et al., 2001). More recently, higher-resolution imaging in live cells revealed that Dis1 undergoes a cell cycle–dependent relocalization from interphase MTs to the kinetochores (and weakly along spindle MTs) during mid-M phase and to the SPBs during anaphase (Nakaseko et al., 2001). Both Dis1 and Alp14 coimmunoprecipitate markers for centromeric DNA (Garcia et al., 2001; Nakaseko et al., 2001), providing additional evidence that Dis1 and Alp14 might function at the kinetochore as a link or adapter between kinetochore MTs and chromatids. The centromere-binding region of Dis1 has been mapped to the C-terminal 225 amino acids, overlapping the SPB-targeting domain (Nakaseko et al., 2001) (see following). Finally, centromere mobility and chromosome attachment to the spindle are disrupted in cells bearing mutations in Dis1 or Alp14, consistent with a model in which Dis1 and Alp14

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function as adaptors between MTs and kinetochores/chromosomes (Garcia et al., 2001; Nabeshima et al., 1998; Nakaseko et al., 2001). Both Dis1 and Alp14 have also been proposed to regulate the dynamics of cytoplasmic or spindle MTs (Garcia et al., 2001; Nakaseko et al., 2001). In contrast to results indicating that Stu2, the MAP215/Dis1 family member in budding yeast S. cerevisiae, antagonizes the eVects of Kin I catastrophe factors in vivo (Severin et al., 2001; however, see Kosco et al., 2001; van Breugel et al., 2003), genetic interactions suggest that Dis1 and Alp14 act cooperatively with the destabilizing kinesin-related proteins Klp5 and Klp6 (Garcia et al., 2002). This observation is more in keeping with recent results indicating that Stu2 promotes MT dynamics in vivo (Kosco et al., 2001) and destabilizes MTs in vitro (van Breugel et al., 2003). Confirmation of the dual roles of Dis1 and Alp14 as MT linkers and regulators awaits direct examination of the eVects of these two proteins on MT dynamics. Two alleles of STU2 (Suppressor of tubulin, stu2-1 and stu2-2) were identified as dominant extragenic suppressors of the cold-sensitive tub2-423 allele of b-tubulin in the budding yeast S. cerevisiae (Wang and HuVaker, 1997). Cloning and sequence comparisons revealed that Stu2 (the product of the STU2 gene) was structurally similar to Dis1 (16% identical) and Alp14 (21% identical) in S. pombe. Stu2 protein localizes to cytoplasmic MTs during interphase and to astral and spindle MTs during mitosis and is highly enriched at SPBs, where it is bound to both the cytoplasmic and nuclear faces (Kosco et al., 2001; Wang and HuVaker, 1997) (see Fig. 6). Kosco et al. (2001) and Usui et al. (2003) reported that Stu2 is enriched at the plus ends of cytoplasmic and astral MTs, although localization of Stu2 to kinetochores has not been reported. Unlike Dis or Alp14 in S. pombe, which are not individually essential for viability (however, they are synthetically lethal) (Garcia et al., 2001; Nakaseko et al., 2001), genetic analysis revealed that Stu2 is essential for mitotic cell growth (Wang and HuVaker, 1997). Depletion or disruption of Stu2 function arrests cells in metaphase of mitosis before sister chromatid segregation and anaphase spindle elongation (Kosco et al., 2001; Wang and HuVaker, 1997). A fraction of Stu2-depleted cells exhibit elongated spindles that appeared thinner and that often are bent or broken (Kosco et al., 2001), similar to the spindle phenotype associated with Alp14 defects in S. pombe (Garcia et al., 2001; Nakaseko et al., 2001). Severin et al. (2001) demonstrated that mad2/stu2 cells, bearing mutations in both the MAD2 component of the spindle assembly checkpoint and Stu2, failed to elongate anaphase spindles despite release from the MAD-dependent mitotic arrest, suggesting that Stu2 cells were unable to assemble or stabilize spindle MTs. Moreover, deletions of the KIP3 kinesin-related protein (in a triple mutant stu2-10/kip3/mad2) partially suppressed the stu2 phenotype


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FIG. 6 Stu2 protein is associated with interphase MTs, spindles, and spindle pole bodies. Images of GFP fluorescence in live yeast cells expressing Stu2-GFP from the endogenous promoter. In unbudded cells (A–B), Stu2 is localized to the spindle pole bodies (arrowheads) and to foci corresponding to MT plus-ends (arrows), often at the cell cortex. In pre-anaphase cells (C and D), Stu2 is localized to the poles of short mitotic spindles (arrowheads), and more faintly to the central spindle in a region where MT plus ends are found (arrows). In elongating anaphase spindles (E–F), Stu2 was localized to spindle poles (arrowheads), along spindle MTs, and in the overlap zone where MT plus-ends are concentrated (arrows). A–D are single focal planes. E and F are projections from optical Z-series (bar is 1 mm). Figure courtesy of Dr. HuVaker. Reprinted from Molecular Biology of the Cell (Mol. Biol. Cell 2001 12, 2870–2880) with permission by the American Society of Cell Biology.

(Severin et al., 2001), leading the authors to conclude that Stu2 stabilizes MTs by counteracting the destabilizing eVects of the kinesin-related protein KIP3, analogous to the antagonistic roles of XMAP215 and XKCM1 in Xenopus egg extracts (Tournebize et al., 2000). However, subsequent studies revealed that depletion of Stu2 suppressed the dynamics of both cytoplasmic and spindle MTs in vivo, indicating that Stu2 destabilizes MTs (Kosco et al., 2001). By monitoring the dynamics of individual cytoplasmic MTs in interphase and preanaphase cells, Kosco et al. (2001) demonstrated that depletion of Stu2 suppresses both catastrophes and rescues and increases the frequency of pauses with little eVect on the rates of MT elongation or rapid shortening. By inference, Stu2 promotes MT

FIG. 7 The amino acid sequence of the MT binding domain of Stu2 mapped in S. cerevisiae is not well conserved among other Saccharomycetaciae. Alignment of an 110–amino acid sequence following the second TOG domains from six species of Saccharomyces, corresponding to the MT binding domain of Stu2 mapped by Wang and HuVaker (1997). Black shading represents amino acids identical to the 50% consensus (shown above); grey shading represents conservative substitutions. f represents a nonpolar amino acid. The MT binding domain of Stu2, identified by Wang and HuVaker (1997) is shown in a grey box below the sequences, with the bipartate repeats (1A-1B:2A-2B) indicated. Although well conserved in four species (S. cerevisiae, S. paradoxus, S. mikatae, and S. bayanus), the sequences corresponding to the MT binding domain in S. castelli and S. kluyveri are more highly diverged (see text). Asterisks denote the CDK1 phosphorylation targets predicted by Wang and HuVaker (1997). S. castelli includes just one of the two predicted CDK1 targets and S. kluyveri includes neither. However, corresponding region in these and all other MAP215/Dis1-related proteins in ascomycotes are rich in basic amino acids (see text).


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dynamics by promoting catastrophe and rescue and decreasing the frequency of pauses (Kosco et al., 2001). Suppression of MT dynamics in Stu2-depleted cells resulted in a number of defects in MT organization and function. Although the length of cytoplasmic MTs was not significantly diVerent in Stu2-depleted cells, the number of cytoplasmic MTs was substantially reduced after Stu2 depletion, which presumably accounts for the observed defects in spindle orientation and positioning relative to the bud neck (Kosco et al., 2001). Suppression of the dynamics of kinetochore MTs resulted in secondary disruption of chromatid attachment and suppression of the centromere motility and dynamics in preanaphase spindles (Kosco et al., 2001). However, the molecular basis for the observed defects in anaphase spindle elongation in Stu2-depleted cells (Kosco et al., 2001; Severin et al., 2001), even in the absence of a MAD2-dependent spindle checkpoint (Severin et al., 2001), is less easily reconciled with a primary role in promoting MT destabilization and disassembly. The rather surprising conclusion that Stu2 destabilizes MTs in vivo (Kosco et al., 2001) (in light of previous indications that Stu2 acted antagonistically to Kin I-related catastrophe factors; Severin et al., 2001) has recently been confirmed in vitro by examining the eVect of recombinantly produced Stu2 on the dynamics of individual MTs (van Breugel et al., 2003). Consistent with results obtained by depleting Stu2 in vivo (Kosco et al., 2001), addition of Stu2 to growing MTs in vitro significantly decreased the rate of elongation and increased the frequency of catastrophe (van Breugel et al., 2003).

C. Correlating the Structure and Function of Dis1-Related Proteins in Yeast Dis1, Alp14, and Stu2 proteins share (at least) three functional characteristics: all bind to MTs in vivo and in vitro, they influence the dynamics function of MT plus-ends, and all bind to the SPB (Garcia et al., 2001; Kosco et al., 2001; Nabeshima et al., 1995; Nakaseko et al., 2001; Wang and HuVaker, 1997). The substantial divergence in amino acid sequence among Dis1, Alp14, and Stu2 made the initial correlation of protein function with structure diYcult. The identification of MAP215/Dis1-related proteins in a wide variety of fungi, including archaeascomycotes, hemiascomycotes, and euascomycotes, as well as six nearly complete sequences and a seventh partial sequence from within the hemiascomycote genus Saccharomyces, thus provides an unprecedented opportunity to correlate the known protein functions with conserved sequence elements and protein structure. The N terminus of all Dis1-related proteins in ascomycotes includes two TOG domains, each composed of as many as five HEAT repeats. No specific

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function has been ascribed to the TOG domains. However, in pairwise sequence comparisons between taxa, the TOG domains exhibit greater sequence conservation than the proteins as a whole, suggesting that sequence divergence is limited by functional constraints. The two single amino acid substitutions responsible for the Stu2 phenotype in S. cerevisiae (D513Y and T514A) (Wang and HuVaker, 1997) map to a seven–amino acid sequence (IVNDTQP, amino acids 510–516 in S. cerevisiae) that is strictly conserved among all seven species of Saccharomyces for which sequence is available, and the core of this sequence is highly conserved across all species of ascomycotes (the aspartate is conserved in Dis1-related proteins through all fungi, including four basidiomycotes). This sequence is found near the middle of the last HEAT repeat in the second TOG domain (1B), underscoring the functional importance of the conserved HEAT repeats and TOG domains. The regions of Dis1, Alp14, and Stu2 responsible for MT binding in vitro and in vivo have all been mapped to 150 amino acids adjacent to the second TOG domain (amino acids 518–641 in Dis1, 559–709 in Alp14, and 557–658 in Stu2) (Nabeshima et al., 1995; Nakaseko et al., 1996, 2001; Wang and HuVaker, 1997). However, comparison of the three MT binding domains reveals little, if any, sequence conservation. This apparent lack of sequence conservation is perhaps not surprising, given the overall divergence in the amino acid sequences of these three proteins. Although no obvious protein motifs were recognized within the MT binding domains of Dis1 and Alp14, Wang and HuVaker (1997) reported that the MT binding domain of Stu2 comprised a tandem duplication of a short, bipartite sequence. Deletion analysis revealed that both copies of this repeat contributed to MT binding in vitro. Moreover, each repeat included a predicted target for phosphorylation by CDK1 (one strongly predicted target in the first repeat, and a less conserved target in the second) (Wang and HuVaker, 1997), suggesting a mechanism by which MT binding might be regulated during the cell cycle. Somewhat surprisingly, the 92–amino acid MT binding domain in S. cerevisiae Stu2 identified by Wang and HuVaker (1997) is not well conserved among Stu2-related proteins identified within the genus Saccharomyces. Pairwise comparisons of the MT binding domain of Stu2 with homologous regions of Stu2-related proteins in five additional species of Saccharomyces (S. kluyveri, S. castellii, S. bayanus, S. mikitae, and S. paradoxus) reveals that amino acid sequence conservation ranges from a low of 18% identity (comparing S. kluyveri to S. paradoxus) to 90% identity (comparing S. paradoxus to S. cerevisiae). In particular, only 10 of 75 amino acids in the two imperfect repeats (1A/B and 2A/B) observed in the S. cerevisiae MT binding domain are conserved in the Stu2 homolog in S. kluyveri. Moreover, the Stu2 homolog in S. castellii is predicted to have deletions of six amino acids in both the first and second domains of repeat 1 (referred to by Wang and HuVaker as 1A and 1B) (Wang and HuVaker, 1997), with only 15 of 75


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amino acids conserved overall. For comparison, sequence conservation of the N-terminal region containing both TOG domains ranges from 40% (comparing S. kluyveri to S. paradoxus) to 93% identity (comparing S. paradoxus to S. cerevisiae). The considerable divergence in amino acid sequence observed in comparisons of the MT binding domains of Dis1, Alp14, Stu2, and the corresponding regions in Dis1-related proteins in all ascomycotes, suggests that the ability to bind MTs is not sequence dependent. Instead, MT binding might require other chemical or physical properties of the MT binding domains. Indeed, the MT binding domains of all three proteins, Dis1, Alp14, and Stu2, overlap with the highly basic KR domain shared by all members of the MAP215/Dis1 family identified in ascomycotes, indicating that MT binding might be dependent on electrostatic interactions between the KR domain of yeast MAP215/Dis1 proteins and tubulins. This hypothesis is supported by observations that the binding of Stu2 to MTs is disrupted by high salt (Wang and HuVaker, 1997), which disrupts electrostatic interactions, and is consistent with proposed interactions between vertebrate brain MAPs (including Tau, MAP1, and MAP2) with the acidic C terminus of b-tubulin (Littauer et al., 1986; Rodionov et al., 1990; Serrano et al., 1984, 1985). However, unlike vertebrate brain MAPs, both Stu2 and XMAP215 (the MAP215/Dis1 family member in Xenopus) bind to MTs assembled from tubulin in which the acidic C terminus of b-tubulin has been removed by mild proteolysis with subtilisin (Spittle et al., 2000; van Breugel et al., 2003). This observation suggests that the binding interaction of MAP215/Dis1-related proteins with tubulin is distinct from that of vertebrate brain MAPs. All members of the Dis1 family in ascomycotes include a 75–100–amino acid domain near the C terminus that is predicted with high probability (PAIRCOIL probability scores >0.50) to form a coiled–coil. Stu2 has recently been shown to be a stable dimer in solution (van Breugel et al., 2003), raising the possibility that the coiled–coil regions of MAP215/Dis1-related proteins in yeast function as dimerization domains, and suggesting that other MAP215/Dis1-related proteins in yeast also function as dimers. It is not clear whether Dis1 and Alp14, which interact genetically and colocalize to SPBs, kinetochores, and spindles, might also physically interact to form heterodimers via their coiled–coil domains. Dis1, Alp14, and Stu2 all localize to the SPB in a cell cycle–dependent manner. The regions of Dis1, Alp14, and Stu2 required for targeting these proteins to the SPB have all been mapped to the C termini (Nakaseko et al., 1996; Sato et al., 2004; Usui et al., 2003), corresponding to amino acids 645– 882 in Dis1 (Nakaseko et al., 1996), 853–888 in Stu2 (Usui et al., 2003), and 697–809 in Alp14 (Sato et al., 2004). Several recent studies have demonstrated that Stu2 is targeted to the SPBs via specific interactions with a 72kDa coiled–coil component of SPBs known as SPC72 (Chen et al., 1998;

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Soues and Adams, 1998). SPC72 was independently identified as a component of SPBs (Soues and Adams, 1998) and as a Stu2 binding partner in two-hybrid screens (Chen et al., 1998). Disruption of SPC72 function phenocopies many of the defects observed in Stu2 mutants (Chen et al., 1998; Soues and Adams, 1998). SPC72 is localized to the nuclear face of the SPB, through interactions of its C-terminal domain with KAR1 and NUD1 (Gruneberg et al., 2000; Pereira et al., 1999), and contains additional binding sites near its amino terminus for the TUB4 g-tubulin complex (Knop and Schiebel, 1998; Pereira et al., 1998), and Stu2 (Usui et al., 2003). Usui et al. (2003) demonstrated that SPC72, TUB4 complex proteins, and Stu2 can be coimmunoprecipitated, indicating that they form a ternary protein complex. Because SPC72 self associates (Chen et al., 1998), it was unclear whether TUB4 and Stu2 bind to the same SPC72 molecule. Usui et al. (2003) mapped the SPC72/SPB targeting sequence to the Cterminal 35 amino acids of Stu2. Interestingly, sequence alignment of all Dis1 family members identified in ascomycotes revealed a highly conserved 28– amino acid sequence motif that falls within the putative SPC72 binding/SPB targeting domain of Stu2 and the SPB targeting domain of Dis1 (see Fig. 8). The sequence of the 23–amino acid core of this conserved domain is more than 55% similar in Dis1-related proteins of ALL ascomycotes and is >60% identical and >85% similar in Dis-related proteins from hemiascomycotes (Saccharomyces and Candida) and euascomycotes (Aspergillus, Coccidioides, Magneporthe, and Neurospora). The high degree of amino acid sequence conservation apparent in this C-terminal SPB targeting domain, extending across all ascomycote yeast species (and even to basidiomycotes; section V.D), suggests that this protein domain is under considerable selective pressure and perhaps functions as the binding site for interacting proteins (i.e., SPC72) that anchor Dis1 family members to the SPB. Although BLAST searches of all available ascomycote genomes (both complete and incomplete) identified SPC72 protein homologs (ranging from 66 to 81% sequence identity) in only four Saccharomyces species. (S. kudriavzevii [AACI01000739.1], S. mikitae [AACH01000231.1], S. paradoxus [AABY01000017.1], and S. bayanus [AACA01000020.1]). other SPC72 homologs may yet be identified as the genome sequences of additional yeast genomes are completed (see following). The predicted coiled–coil structure of SPC72 and its role in targeting Stu2 to the SPB of budding yeast have lead to suggestions that SPC72 is the functional (though perhaps not evolutionarily related) homolog of the TACC proteins that target MAP215/Dis1 family members to the centrosomes of ‘‘higher’’ eukaryotes (Section V.D.5). However, SPC72 has only been observed at the cytoplasmic face of the SPB (Rout and Kilmartin, 1991; Soues and Adams, 1998), whereas Stu2 has been localized to both cytoplasmic


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FIG. 8 A conserved sequence in the C-termini of MAP215/Dis1 proteins in fungi may represent a SPB targeting/SPC72 binding domain. The SPB targeting domain of Stu2 has been mapped to a 35–amino acid sequence at the C terminus of Stu2, which includes a conserved 23 amino acid sequence found at or near the C terminus of all MAP215/Dis1-related proteins in ascomycote fungi, including archaeascomycotes (Dis1 and Alp14 in S. pombe), hemiascomycotes (C. albicans and all Saccharomycetaciae, incl. Stu2 in S. cerevisiae), and several euascomycotes (incl. A. nidulans and N. crassa). Black shading represents amino acids identical to the 50% consensus (shown above; f represents a nonpolar amino acid, þ to basic amino acids, to acidic amino acids); grey shading represents conservative substitutions. A related sequence is also found in UM-MAP250, the MAP215/Dis1 homolog in the basidiomycote fungus Ustilago maydis, where it overlaps the Tau repeat (bolded; see text).

and nuclear faces (Kosco et al., 2001). Thus, additional factors might play a role in targeting Stu2 and other Dis1-related proteins to the spindle poles bodies in yeast. Recently, Sato et al. (2004) demonstrated that targeting of Alp14 to the spindles and SPBs of S. pombe is dependent on the product of the Alp7 gene. Mutations in Alp7 result in altered cell polarity and mitotic defects similar to those observed in alp14 cells. Alp7 encodes a coiled–coil protein that is associated with MTs during interphase and to the spindle and SPBs during M-phase (Sato et al., 2004), where it colocalizes with Alp14. The localization of these two proteins is interdependent: Localization of Alp14 to SPBs requires Alp7, and localization of Alp7 to spindles and kinetochores requires Alp14. Moreover, Alp7 and Alp14 form a complex mediated by the C-terminal coiled–coil domain of Alp7 and the C-terminal 113 amino acids of Alp14 (697–809). On the basis of these results, Sato et al. (2004) concluded

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that Alp7 is the functional homolog of the TACC family of coiled–coil proteins required for targeting of MAP215/Dis1 family members to the centrosomes of higher eukaryotes (Sections V.D.5 and V.E). Moreover, they suggest that Slk9, the S. cerevisiae protein most similar to Alp7, might also play a role in Stu2 localization and function in budding yeast (Sato et al., 2004). Although the evolutionary relationships among SPC72, Alp7, and TACC proteins are unclear, these proteins appear to perform homologous functions in targeting MAP215/Dis1-related proteins to MTOCs in yeast and animals. Interestingly, all are coiled–coil proteins that interact with the C-termini of their MAP215/Dis1-related binding partners. We propose that the mechanisms for targeting yeast MAP215/Dis1-related proteins to SPBs are evolutionarily conserved, relying on specific interactions with coiled–coil components of the SPB with the conserved sequence motif identified near the C termini of these proteins (Section V.E). Stu2 has also been observed to be associated with MT plus ends (Kosco et al., 2001; Usui et al., 2003). Although the MT binding domain of Stu2 maps to amino acids 557–658, overlapping the KR domain, Usui et al. (2003) found that targeting to the plus-ends of cytoplasmic and astral MTs was dependent on the same C-terminal 35 amino acids that are required for targeting to the SPBs. Similar results were obtained for Alp14 (Sato et al., 2004), in which targeting of Alp14 to spindle MTs was dependent on its SPB targeting domain, rather than its MT binding domain. Moreover, targeting of Alp7 to kinetochores and spindle MTs was dependent on interactions with Alp14 (Sato et al., 2004). Similarly, C-terminal sequences suYcient for targeting Dis1 to the centromeres of mitotic chromosomes overlap with those required for targeting to the SPB, and centromere targeting was at least partially dependent on MTs (Nakaseko et al., 2001). These apparently contradictory observations can be reconciled by a model in which Stu2, Alp14 and Alp7, and Dis1 are targeted to the SPBs, where they can then be loaded onto the plus ends of elongating MTs for translocation to plus ends or kinetochores (Section V.E).

D. Structure of Dis1-Related Proteins in Basidiomycotes Genes encoding MAP215/Dis1-related proteins were identified by BLAST in the genomes of four species of basidiomycotes: Ustilago maydis, the causative agent of corn smut disease; Phanerochaete chrysosporium, white rot; Cryptococcus neoformans, the causative agent of cryptococcosis commonly observed in patients with AIDs; and Coprinopsis cineria (Coprinus cinereus, the Grey inkcap). The predicted protein from U. maydis (UM-MAP215) is encoded by a single open reading frame of 2238 amino acids and is likely to


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represent the complete protein sequence. However, based on size and sequence comparisons with UM-MAP215 (2238 amino acids), the MAP215/ Dis1-related proteins predicted from P. chrysosporium (1970 amino acids), C. neoformans (1727 amino acids), and C. cineria (1864 amino acids) genomes are likely to be incomplete, missing 250–500 amino acids from their C termini. The available sequences of the four MAP215/Dis1-related proteins in basidiomycotes are 37–63% identical, and they constitute a monophyletic group by both parsimony (Fig. 2) and distance analysis (not shown). The basidiomycote MAP215/Dis1-related proteins are 16–19% identical to Dis1 in S. pombe and 23–29% identical to related proteins in euascomycotes (including Aspergillus and Neurospora). Although exhibiting only limited sequence similarity to MAP215/Dis1-related proteins in higher eukaryotes (13–17% identity to plants and 18–21% to vertebrates), the predicted sequence of UM-MAP215 exhibits several features characteristic of MAP215/Dis1 family members in animals and plants. First, the predicted sequence of UM-MAP215, at 2238 amino acids, is substantially longer than the related proteins in Ustilago’s ascomycote cousins, which range from 816 (S. castellii) to 940 amino acids (N. crassa). The size of UM-MAP215 is comparable to that of MAP215/Dis1 family members in insects (Msps in Drosophila is 2050 amino acids), vertebrates (the zygotic isoform of XMAP215 in X. laevis is 2030 amino acids), and plants (MOR1 in Arabidopsis is 1978 amino acids). Although the predicted sequences in other basidiomycotes appear incomplete, each is larger than 1970 amino acids. In keeping with its predicted size, the sequence of UM-MAP215 includes four strongly predicted TOG domains, corresponding to 1A (four predicted HEAT repeats; 24–44% identical to TOG 1A in fungi, plants, and animals), 1B (five predicted HEAT repeats; 25–42% identical to TOG 1B in fungi, plants, and animals), 2A (two predicted HEAT repeats; 18–30% identical to TOG 2A in fungi, plants, and animals), and 2B (five predicted HEAT repeats; 14–34% identical to TOG 2B in fungi, plants, and animals). The N-terminal signature sequences of each of the four TOG domains of UMMAP215 are very similar (80–100% identical) to those in ch-TOG and XMAP215. Like XMAP215 and ch-TOG (Charrasse et al., 1998), the predicted sequence of UM-MAP215 includes a domain rich in alanine, proline, and serine following the fourth TOG domain (amino acids 1130–1350; Section V.D). This potential S,P domain overlaps a fifth cluster of HEAT repeats. Like the Dis1-related proteins in ascomycotes, UM-MAP215 includes a domain (amino acids 1862–1897) that is predicted to form a coiled–coil (with a PAIRCOIL probability score >0.5). However, in UM-MAP215, the predicted coiled–coil is much shorter (just 36 amino acids, compared to 100

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amino acids in ascomycotes) and may not support stable protein–protein interactions. Finally, the predicted UM-MAP215 sequence includes 18 amino acids near the C terminus (amino acids 2094–2111) that are 50% identical to the Tau repeat in the human MAP215/Dis1-related protein ch-TOG and 22% identical to the first MT binding repeat of human Tau. The KIGS sequence of ch-TOG has been replaced with RISD in UM-MAP215, in which the phosphorylatable serine has been replaced with the phosphomimic aspartate. Interestingly, 26 amino acids (2082–2107) overlapping the putative Tau repeat of UM-MAP215 are 42% identical and 50% similar to the consensus of a conserved sequence found near the C termini of MAP215/Dis1-related proteins identified in ascomycotes. In S. cerevisiae, this conserved sequence maps within the 35–amino acid region required for targeting Stu2 to SPBs and is proposed to function as a binding site for SPC72 (see earlier). On the basis of these observations, and on additional evidence from studies of the localization and targeting of XMAP215 to centrosomes in frog cells (Popov et al., 2001) (Section V.D.4), the Tau repeat-related sequences in UMMAP215 and other MAP215/Dis1-related proteins may function to target these proteins to MTOCs.

IV. Structure and Function of Mor1/Gem1-Related Proteins in Plants Two temperature-sensitive alleles of MOR1 (mor1-1 and mor1-2) (Microtubule Organization 1), the first MAP215/Dis1-related protein to be discovered in plants, were identified in a visual screen for mutations that disrupted the organization of cortical MTs in the epidermal cells of Arabidopsis seedlings (Whittington et al., 2001). When grown at the permissive temperature, homozygous mor1 seedlings are indistinguishable from wild-type plants. mor1 seedlings grown at the restrictive temperature are stunted and exhibit radial swelling of organs consistent with defects in polar cell elongation caused by disorganization of the cortical MTs and resulting disruption of cellulose deposition. Examination of the MT organization in mor1 plants after shifting them to the restrictive temperature revealed a rapid disruption of the cortical MT array, including shortening or fragmentation of the MTs and loss of their characteristic circumferential organization (Whittington et al., 2001). Cortical MT organization was restored on returning the plants to the permissive temperature, indicating that the changes in MT organization result from a reversible temperature-dependent inactivation of the MOR1 product.


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Although neither of the two original alleles of MOR1 caused mitotic or cell division defects, MOR1 was subsequently shown to be allelic to GEM1 (GEMini pollen 1) (Twell et al., 2002). The gem1-1 and gem1-2 mutations are homozygous lethal, indicating that MOR1/GEM1 is required for some essential cellular process. GEM1 mutations disrupt cell polarity and division during pollen development, resulting in a twin-celled pollen phenotype (Twell et al., 2002). Immunofluorescence confirmed that Mor1/Gem1 localizes to all four MT arrays during the plant cell cycle (Twell et al., 2002) (Fig. 9). Antibodies to Mor1/Gem1 stain the interphase array of cortical MTs, consistent with the disruption of these MTs by mutations in Mor1/Gem1. Mor1/Gem1 also is localized to the preprophase band of MTs that marks the future division site. During M phase, Mor1/Gem1 antibodies stain the mitotic spindle, consistent with the localization of MAP215/Dis1-related proteins in animal cells. Mor1/ Gem1 also localizes to the phragmoplast, a unique MT array required for the assembly of the nascent cell plate during cytokinesis in plant cells. Despite the observed localization of Mor1/Gem1 to spindle MTs during M-phase, none of the four MOR1/GEM1 alleles disrupts nuclear division during pollen development (Twell et al., 2002). This is somewhat surprising in light of the essential role of MAP215/Dis1-related proteins in spindle assembly and function in yeast (Garcia et al., 2001; Kosco et al., 2001; Rockmill and Fogel, 1988; Wang and HuVaker, 1997) and animals (Becker et al., 2003; Cassimeris and Morabito, 2004; Cullen et al., 1999; Holmfeldt et al., 2004; Tournebize et al., 2000). Although the existing MOR1/GEM1 alleles may not represent the null phenotype (mor1-1 and mor1-2 are point mutations near the N terminus and gem1-1 and gem1-2 both produce truncated products, see following), the lack of spindle defects might result from basic diVerences in the organization and function of spindles in animals and plants. In animals, MAP215/Dis1-related proteins are localized to centrosomes and are required for organizing the poles of mitotic spindles (Section V.D). Spindles in higher plants lack classic centrosomes (Vaughn and Harper, 1998), and Mor1/Gem1 is not significantly concentrated at the spindle poles (Twell et al., 2002). It is possible that the lack of mitotic defects reflects this basic diVerence in the organization of the spindle poles in plant cells. However, MAP215/Dis1-related proteins are required for assembly and organization of acentrosomal meiotic spindles in Drosophila (Msps) (Cullen and Ohkura, 2001) and Xenopus oocytes (XMAP215; Becker et al., 2003), and many other aspects of spindle structure and function are conserved between animals and plants. Thus, the role of Mor1/Gem1 in plant mitosis, or lack thereof, remains uncertain. Building on these observations, and the known eVects of XMAP215 on MT dynamics in animal cells (Section V.D.2), Twell et al. (2002) proposed that Mor1/Gem1 might promote the assembly or dynamics of MT plus ends

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FIG. 9 Mutations in MOR1/GEM1 cause temperature-dependent disruption of cortical MTs in leaf epidermal cells of Arabidopsis. (A) MTs in wild-type cells are organized circumferentially. (B) At the restrictive temperature, cortical MTs in cells from mor1 homozygotes appear shorter and disorganized (scale bar is 25 mm). Immunofluorescence microscopy reveals that the MOR1/ GEM1 product is associated with MTs of (C) interphase cells, (D) the preprophase band, (E) mitotic spindles, and (F) phragmoplasts of cultured Arabidopsis cells (C, D, E, and F stained with anti-tubulin; C0 , D0 , E0 , and F0 stained with anti-MOR1/GEM1). A and B are reprinted from Whittington et al. (2001) courtesy of Dr. Wasteneys and reproduced with the permission C–F are reprinted from Twell et al. (2002) courtesy of Dr. Hussey, and reproduced with permission.

required for assembly of interphase MT arrays and the phragmoplast during M phase. However, identification of the Mor1/Gem1 homolog TMBP200 as a MT bundling protein in cultured tobacco cells (Yasuhara et al., 2002) suggests that Mor1/Gem1 family members might also function in the


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bundling and organization of phragmoplast MTs, a function that is also consistent with the observed localization of Mor1/Gem1 to the midzone of phragmoplast in dividing cells and of the cytokinetic defects observed in cells bearing the gem1-1 and gem1-2 alleles (Twell et al., 2002). Five complete or nearly complete sequences for Mor1/Gem1 family members are available (see Appendix I): cDNAs encoding Mor1/Gem1 in A. thaliana (Whittington et al., 2001) and TMBP200 in N. tabacum (Yasuhara et al., 2002) have been cloned and sequenced, nearly complete sequences of Mor1/Gem1 homologs in O. sativa (rice) and P. trichocarpa (Poplar) were predicted from genomic DNA sequence, and a partial sequence for a Mor1/ Gem1 homolog in Z. mays was compiled from multiple EST sequences. Partial sequences for Mor1/Gem1 family members were identified in ESTs from Medicago truncatula, Glycine max (soybean), and Physcomitrella patens (a moss). Finally, a long, but apparently incomplete, sequence for a Mor1/ Gem1 family member was identified in the partial genomic sequence of the green algae, C. reinhardtii. The structure of Mor1/Gem1 homologs is highly conserved within the plant kingdom. Pairwise comparison of their amino acid sequences of Mor1/Gem1 homologs in higher plants reveals sequence identities ranging from 70% to 80%. Mor1/Gem1 homologs in algae and the lower plants are more divergent: The 71–amino acid fragment of the Mor1/Gem1 homolog predicted from an EST from P. patens is 44 –53% identical to the corresponding region in higher plant proteins (which are 70% identical over the same region), and the incomplete predicted amino acid sequence of the Mor1/Gem1 homolog in the green algae C. reinhardtii is 32–33% identical to that of higher plants over 800 amino acids. For comparison, sequences of the plant Mor1/Gem1 homologs are 18–28% identical to MAP215/Dis1-related proteins in vertebrates and 11–24% identical to those in fungi. Within the higher plants, sequence conservation extends throughout the entire length of the predicted proteins, with three short variable regions in the C-terminal half of the proteins (amino acids 1070–1120, 1420–1460, and 1860–1950). The amino acid sequences of all of the plant homologs (for which suYcient sequence is available) are predicted to include four TOG domains corresponding to 1A, 1B, 2A, and 2B. The single–amino acid substitutions identified in the mor1-1 (L174F) and mor1-2 (E195K) alleles in Arabidopsis fall within the fourth predicted HEAT repeat of TOG domain 1A (the first repeat predicted by Whittington et al., 2001), underscoring the importance of the HEAT repeats in the structure and function of MAP215/Dis1 family members. These single–amino acid substitutions result in a temperaturedependent disorganization of cortical MTs but do not impair either nuclear or cytoplasmic division (Whittington et al., 2001). Presumably, at the restrictive temperature, the function of the Mor1/Gem1 protein is altered in a

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manner that specifically compromises the assembly and organization of the cortical MT array. The MT binding domain of Mor1/Gem1 in A. thaliana has been roughly mapped to the C-terminal 856 amino acids (amino acids 1123–1978) (Twell et al., 2002). This region overlaps the high-aYnity MT binding domain mapped in XMAP215 (Section V.D.4) and also includes the 18–amino acid sequence related to the Tau repeat, which is postulated to function as a centrosomal targeting domain in MAP215/Dis1 family members in animal cells (Section V.D.5). Although cells of higher plants lack centrosomes (Vaughn and Harper, 1998), this sequence might perform a homologous function, allowing Mor1/Gem1-related proteins to bind MT nucleation factors in plant cells. There, as proposed in Section VI.D, Mor1/Gem1-related proteins in plant cells might also be loaded to the plus ends of MTs for transport or targeting to their destination (e.g., to the plus ends of phragmoplast MTs). Interestingly, as discussed in section I.B.3, the conserved serine at the core of the Tau repeat in MAP215/Dis1-related proteins in animal cells (which is also conserved in the Tau repeats of the Tau, MAP2, and MAP4 families of vertebrate MAPs) has been substituted with nonphosphorylatable glutamate (E) or aspartate (D) in all Mor1/Gem1-related proteins in the plants, from Chlamydomonas through Arabidopsis. Although these amino acids are not subject to phosphorylation, their negative charge might mimic a phosphorylated state. The more severe alleles gem1-1 and gem1-2 produce truncated protein products retaining 1326 and 411 amino acids of the 1978–amino acid wildtype Mor1/Gem1 protein, respectively (Twell et al., 2002). The gem1-1 product, which exhibits a phenotype of intermediate severity, would include all four TOG domains but would be missing part of the mapped MT binding region (notably, that which overlaps the higher-aYnity MT binding domain in XMAP215) and the Tau repeat. The most severe allele, gem1-2, produces a protein of just 411 amino acids, truncating the wild-type protein in the fourth predicted HEAT repeat of TOG domain 1B. This allele’s pollen division phenotype is nearly 100% penetrant, and the gem1-2 phenotype thus may represent the null phenotype (Twell et al., 2002). Interestingly, although pollen carrying the gem1-2 mutation fails to undergo normal cytokinesis, it exhibits no defects in nuclear division (Twell et al., 2002). Three alternative explanations for this puzzling observation cannot yet be distinguished: gem12 does not represent a true null phenotype, and the product retaining only 411 amino acids of the wild-type protein provides suYcient function to support mitosis; suYcient maternally produced GEM1/MOR1 protein is carried forward from the microspore mother cell to support subsequent pollen mitoses; and Mor1/Gem1 is not required for assembly and function of spindles during pollen division. Further research into the mitotic functions


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of Mor1/Gem1 proteins will be required to discriminate between these hypotheses.

V. The Structure and Function of MAP215/Dis1-Related Proteins in Animals The animal branch of the MAP215/Dis1 family tree currently includes 15 family members, 14 of which fall into three subfamilies: the Zyg9 subfamily in nematodes, the Minispindles (Msps) subfamily in arthropods (insects), and the XMAP215/TOG subfamily in vertebrates (and the invertebrate chordates Ciona intestinalis and Ciona savignyi). In the phylogenetic tree presented in Figure 2, DdCP224, the MAP215/Dis1-related protein in the ramicristate protist D. discoideum, also maps to the animal branch of the MAP215/Dis1 family tree, where it falls as the most basal member, outside of the three well-defined subgroups. However, bootstrap values placing DdCP224 with animals are low, making its inclusion in the MAP215/TOG subfamily uncertain. MAP215/Dis1-related proteins in animals share a common structural organization. With the exception of the Zyg9 subfamily in nematodes (discussed further later), MAP215/Dis1-related proteins of animals (including DdCP224 in Dictyostelium) are large (1900–2100 amino acids) and include four complete TOG domains [(1A:1B)(2A:2B)] within their first 1100 amino acids. The N-terminal TOG domains represent the largest block of sequence conservation between MAP215/Dis1-related proteins of the diVerent subfamilies: Sequence identities range from 37–42% (comparing insects to vertebrates) to over 70% (within the vertebrate subfamily). Following the fourth TOG domain, most MAP215/Dis1-related proteins in animals include additional clusters of HEAT repeats. However, these domains lack the signature sequences identifiable near the amino end of the first four TOG domains. All members of the animal branch of the MAP215/Dis1 family include one or more predicted targets for the cell cycle kinase CDK1. Finally, again with the exception of the Zyg9 subfamily in nematodes, all MAP215/ Dis1 family members in animals (for which complete amino sequences can be predicted) include a single 18–amino acid sequence related to the Tau repeats found in the vertebrate MAPs of the Tau, MAP2, and MAP4 families. In addition to their structural similarities, MAP215/Dis1-related proteins in animals whose function has been characterized, either through genetic mutations or using biochemical or cell biological approaches, share several functional characteristics: all associate with centrosomes (the central MTOC of animal cells) and spindle poles, all bind MTs in vitro and in vivo and

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regulate MT assembly and or dynamics, and their function is required for normal assembly and organization of meiotic and/or mitotic spindles.

A. Structure and Function of DdCP224 in D. discoideum DdCP224 was identified in a monoclonal antibodies screen for components of the uniquely-organized centrosome of the cellular slime mold D. discoideum (Graf et al., 2000). Analysis of cDNAs encoding DdCP224 predicted a 2015–amino acid member of the MAP215/Dis1 family of MAPs (Graf et al., 2000). Pairwise sequence comparisons reveal that DdCP224 is 13–24% identical to MAP215/Dis1-related proteins in fungi, 17–25% to those of higher plants, and 15–29% to those of animals. Immunolocalization (Graf et al., 2000) revealed that DdCP224 is associated with the centrosomes of interphase cells. Comparison of the localization of DdCP224 with that of g-tubulin indicated that DdCP224 was localized to the periphery of the centrosome corona, corresponding to the pericentriolar material of centrosomes in animal cells. Weaker staining was also apparent along some interphase MTs. The centrosomes were also strongly stained by DdCP224 antibodies throughout M phase, along with weaker staining of spindle MTs. In metaphase cells, DdCP224 was also localized to the kinetochore region of the central spindle. Examination of DdCP224 localization in nocodazole-treated cells revealed that the association of DdCP224 with centrosomes is independent of MTs. However, kinetochore staining was substantially reduced or eliminated in nocodazole-treated cells, indicating that targeting or anchoring DdCP224 to the kinetochore requires intact MTs. During late M phase and cytokinesis, DdCP224 also localized to the region of the midbody. Cells underexpressing a DdCP224-GFP fusion in place of the endogenous protein exhibit a reduction in the number and length of interphase MTs and exhibit hypersensitivity of the remaining MTs to nocodazole (Graf et al., 2003). In addition, regrowth of the cytoplasmic MT array during recovery from nocodazole was severely delayed or inhibited. Finally, partially purified DdCP224 was observed to copellet with MTs in vitro (Graf et al., 2000). Together, these observations indicate that DdCP224 binds MTs, where it may function to promote nucleation or elongation. Interestingly, overexpression of full-length DdCP224 fused to GFP or a truncated form corresponding to amino acids 809–2015 induced the formation of supernumerary centrosomes (Graf et al., 2003). These additional centrosomes included g-tubulin, were ultrastructurally similar to endogenous centrosomes, and were observed to separate and duplicate during the cell cycle (albeit somewhat delayed with regard to the endogenous centrosomes). Excess DdCP224 thus appears to be capable of inducing the assembly of


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functional, bona fide centrosomes, recruiting other centrosomal components and organizing them into higher-order structures unique to Dictyostelium centrosomes. This observation indicates that, in addition to its roles in the regulation of MT nucleation or dynamics, DdCP224 may play key roles in centrosome organization and reproduction (Graf et al., 2003). The sequence and structural organization of DdCP224 has many features in common with the MAP215/Dis1-related proteins in higher eukaryotes (animals and plants), but it also shares features with fungal family members and includes several unique features. The size of DdCP224 (2015 amino acids) is comparable to MAP215/Dis1 family members in higher eukaryotes (1900–2100 amino acids). DdCP224 includes four TOG domains [(1A:1B)(2A:2B)] in its N-terminal half (amino acids 11100), each predicted to include three to five HEAT repeats. As with many of the larger MAP215/ Dis1 family members, additional clusters of HEAT repeats are found in the second half of the protein sequence. However, these clusters of HEAT repeats lack the signature sequences characteristic of the first four TOG domains. When expressed as a GFP fusion, a fragment of DdCP224 corresponding to amino acids 1-813 was diVusely distributed in the cytoplasm during interphase (Graf et al., 2000). However, a GFP fusion with amino acids 809–2015 of DdCP224 also failed to localize to interphase MTs, but it did localize to spindle MTs and kinetochores during M phase (Graf et al., 2000). These results suggest that binding to interphase MTs may require cooperative interactions with the N-terminal TOG domains and undefined sequences in the C-terminal half of DdCP224, and that binding to spindle MTs might rely on distinct interactions requiring sequences downstream of the TOG domains. The distribution of DdCP224, similar to that of other MAP215/Dis1related proteins, varies with the cell cycle (Graf et al., 2000). However, unlike MAP215/Dis1-related proteins in all fungi and animals, DdCP224 lacks S/T P X K/R sequences that are predicted to serve as targets for the cell cycle kinase CDK1. A single SPK sequence is located near the N terminus of DdCP224 (amino acids 136–138). The distribution and function of DdCP224 during the cell cycle may thus rely on downstream cell cycle regulators. DdCP224 includes two short stretches (amino acids 1211–1234 and 1989– 2015) that are predicted with low probability to form a coiled–coil (PAIRCOIL scores less than 0.5). Coiled–coil domains are characteristic features of MAP215/Dis1-related proteins in fungi. However, the low PAIRCOIL scores and limited size of the predicted coiled–coil domains of DdCP224 make it unlikely that these domains are suYcient to support stable protein– protein interactions.

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When expressed as a fusion with GFP, a fragment of DdCP224 corresponding to amino acids 809–2015 is targeted to both interphase and M-phase centrosomes (Graf et al., 2000), indicating that the centrosomal targeting function resides in the C-terminal half of DdCP224. This region includes an 18–amino acid domain near its C terminus (amino acids 1819– 1836) that is 61% identical and 83% similar to the consensus sequence of the Tau repeat found in MAP215/Dis1-related proteins in E. cuniculi, U. maydis, all plants, and all animals. Notably, the serine in the highly conserved KIGS sequence that forms the core of Tau repeat in vertebrate MAPs (Tau, MAP2, and MAP4) and in most MAP215/Dis1 family members in animals has been replaced by the polar amino acid asparagine. The lack of finer functional mapping leaves any conclusions regarding the functional relationship between the Tau repeat in DdCP224 and targeting to centrosomes uncertain. However, inclusion of the putative Tau repeat from DdCP224 within a sequence that, when expressed as a GFP fusion, is targeted to centrosomes, further supports speculation that the Tau repeat of MAP215/Dis1-related proteins function as centrosomal targeting domains (Sections V.D.5 and V.E). Finally, DdCP224 includes several short stretches of sequence that are highly enriched in single amino acids including serine (21 amino acids from 1057 to 1077, includes 14 serines), threonine (a run of six amino acids from 1750 to 1755, and another run of 11, from 1778 to 1788), asparagine (a run of 21 from 1372 to 1392, and 19 amino acids from 1704 to 1722 includes 11 asparagines; and a run of 26 asparagines interrupted by a single I from 1914 to 1941), and glutamine (21 amino acids from 1083 to 1103 include 18 glutamines, a run of 14 amino acids from 1394 to 1407). The significance of these sequence motifs is currently unknown.

B. Structure and Function of Zyg9-Related Proteins in Nematodes Zyg9 was originally identified as a maternal eVect mutation that causes embryonic lethality during early development of the nematode worm C. elegans (Kemphues et al., 1986). Embryos lacking functional Zyg9 exhibit disorganized meiotic spindles and subsequently fail to undergo pronuclear migration. The first mitotic spindle that is assembled in zyg9 embryos lacks the characteristic long astral MTs characteristic of wild-type spindles (Albertson, 1984; Kemphues et al., 1986). Disruption of the astral MTs accounts for the observed failure of the migration and rotation of the nuclear-centrosomal complex. Embryos lacking Zyg9 function undergo several aberrant divisions before further development arrests (Kemphues et al., 1986).


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Immunolocalization revealed that Zyg9 is associated with the spindles of both meiotic and mitotic divisions (Matthews et al., 1998). Zyg9 is distributed throughout both meiotic spindles and is enriched in the spindle poles. During mitosis, Zyg9 is largely restricted to the centrosomal regions that constitute the poles of the mitotic spindles, although weak associations with the central spindle are observed from late prometaphase through early anaphase (Matthews et al., 1998). Localization of Zyg9 to the central spindle (Matthews et al., 1998) further suggests that Zyg9 is enriched at MT plus ends. During interphase, Zyg9 appears to be diVusely distributed throughout the cytoplasm (Matthews et al., 1998), indicating that localization of Zyg9 to centrosomes is regulated during the cell cycle. Alternatively, the association of Zyg9 with interphase MTs or centrosomes might be masked by other cellular components. Recently, independent reports from three laboratories have demonstrated that targeting of Zyg9 to the centrosomes of mitotic spindles is dependent on TAC-1 (Bellanger and Gonczy, 2003; Le Bot et al., 2003; Srayko et al., 2003), the single member of the TACC (Transforming Acidic Coiled–Coil) protein family predicted from the C. elegans genome, and that Zyg9 and TAC-1 form a physical complex (Bellanger and Gonczy, 2003; Srayko et al., 2003). TACC family proteins have also been implicated in the binding and targeting of MAP215/Dis1-related proteins to centrosomes in Drosophila (Gergely et al., 2000b; Giet et al., 2002; Lee et al., 2001. Sections IV.D-3 and IV.D-5) and mammals (Conte et al., 2003; Gergely et al., 2000a,b; LauVart et al., 2002) (Sections IV.D-4e and IV.D-5). The cellular distribution of TAC-1 was similar to that of Zyg9, including localization to centrosomes and spindle poles (Bellanger and Gonczy, 2003; Le Bot et al., 2003; Srayko et al., 2003). In addition, TAC-1 localized to the interface between MTs and kinetochores (Srayko et al., 2003). Targeting of TAC-1 (and by inference, Zyg9) to centrosomes is not dependent on MTs but requires activity of a C. elegans homolog of Aurora-kinase (AIR1) (Bellanger and Gonczy, 2003; Le Bot et al., 2003). Interestingly, Bellanger and Gonczy (2003) observed that depletion of TAC-1 resulted in a concomitant reduction of Zyg9 protein levels, and vice versa, suggesting that formation of the TAC-1/Zyg9 complex regulates the stability of each of the protein components (see also results from Srayko et al., 2003). Depletion of either Zyg9 or TAC-1 results in similar phenotypes, including a reduction in the length of astral MTs and spindles (Bellanger and Gonczy, 2003; Kemphues et al., 1986; Le Bot et al., 2003; Srayko et al., 2003). Depletion of Zyg9 or TAC-1 does not appear to limit MT nucleation (Srayko et al., 2003), but TAC-1 depletion does diminish MT dynamics (Bellanger and Gonczy, 2003). Together, these observations suggest that the Zyg9/TAC-1 complex is required for the assembly or

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stabilization of long MTs (Bellanger and Gonczy, 2003; Le Bot et al., 2003; Srayko et al., 2003). Cloning and sequencing of Zyg9 revealed that it was structurally similar to members of the MAP215/Dis1 family of MAPs identified in vertebrates and yeast (Matthews et al., 1998). Database searches of the incomplete genomes of C. briggsae and Brugia malaya identified/predicted similar proteins in these nematode species that are 86% and 46% identical in amino acid sequence to CeZyg9, respectively. However, with the exception of DdCP224 in Dictyostelium (whose association with family members in animals is uncertain), the Zyg9 subfamily in nematodes is the most structurally divergent subfamily of MAP215/Dis1-related proteins in animals. Zyg9-related proteins are shorter (only 1400 amino acids) than those of other animals (which are typically 1900–2100 amino acids in length), and pairwise comparisons reveal sequence identities of 11–17% between Zyg9-related proteins and DdCP224 in Dictyostelium, 17–28% between Zyg9-related proteins and Msps-related proteins in insects, and 19–30% between Zyg9-related proteins and members of the XMAP215/TOG subfamily in vertebrates (for comparison, Msps-related proteins are 30–45% identical to their vertebrate counterparts). Unlike their cousins in insects and vertebrates (and even DdCP224 in Dictyostelium), Zyg9-related proteins include only two well-conserved TOG domains, each with five predicted HEAT repeats. A cluster of three additional HEAT repeats predicted to follow the second TOG domain lacks a conserved signature sequence. Interestingly, the signature sequences of the two TOG domains in Zyg9 from both C. elegans and C. briggsae are K W Q E R K, and those of the B. malaya Zyg9-related protein are K W T E R R and K W V E R R. The fourth position of both TOG domain signature sequences in all three Zyg9-related proteins identified to date is occupied by an E, which is more similar to the B-type signature consensus W K/Q D/E R K in other MAP215/ Dis1 family members. Indeed, sequence distance and parsimony analysis (not shown) place both Zyg9 TOG domains in the 1B class of TOG domains. Moreover, pairwise alignment of the two TOG domains of Zyg9 (amino acids 1–237 vs. 290–522) with each other reveal that they are 51% identical, and comparable levels of sequence identity are also observed between the first and second TOG domains of Zyg9-related proteins in C. briggsae (50%) and B. malaya (46%; for comparison, conservation of the amino acid sequences of TOG domains in XMAP215 ranges from 14% to 20%). MAP215/Dis1 family members from trypanosomes, a fungus (U. maydis), all plants, and all animals except nematodes are predicted to include four TOG domains, suggesting that the duplication event that gave rise to the second pair of TOG domains occurred early in eukaryotic evolution,


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before the divergence of the major eukaryotic lineages. The exceptional organization of the Zyg9-related proteins in three species of nematodes, including the lack of identifiable A-type TOG domains and the presence of two highly similar B-type TOG domains, suggests that substantial rearrangement of the Zyg9 gene occurred in a common ancestor of these three nematode species. The observed sequence organization of modern Zyg9-related proteins is consistent with an evolutionary scenario in which three of the four TOG domains of the ancestral Zyg9 protein, corresponding to both A-type domains and one of the B-type domains, were lost by deletion of the corresponding gene segments. The gene seqment encoding the remaining B-type TOG domain then duplicated to give rise to the (B:B) organization observed in Zyg9-related proteins from these three modern nematodes. Sequencing and comparison of Zyg9-related proteins from additional species of nematodes and more distantly related taxa may provide additional insight into the evolution of the Zyg9 subfamily of MAP215/Dis1-related proteins. The amino acid sequences of both Zyg9 in C. elegans and the Zyg9-related protein in C. briggsae include a single predicted target for phosphorylation by CDK1 (802-SPLK-805), consistent with the observed cell-cycle regulation of Zyg9 localization in C. elegans embryos (Matthews et al., 1998; Srayko et al., 2003). This conserved CDK1 target is absent from the sequence of the Zyg9-related protein in B. malaya, which includes two predicted CDK1 targets (680-TPVR-683 and 694-SPPK-697). Three overlapping regions responsible for TAC-1 binding have been mapped to the C-terminal two-thirds of Zyg9, corresponding to amino acids 654–937, 908–1167, and 1162–1415, consistent with mapping of MTOC targeting domains to the C termini of MAP215/Dis1-related proteins in fungi and animals (Sections V.D.5 and V.F). However, the three Zyg9related proteins identified in nematodes to date lack sequences similar to the Tau repeats, which are found in MAP215/Dis1-related proteins from fungi (U. maydis; Section V.D), protists (Dictyostelium; Section V.A), and all animals (Section V.D.5 and V.E).

C. Structure and Function of Minispindles (Msps)-Related Proteins in Insects Minispindles (Msps protein) was identified in a cytological screen for mutants with mitotic defects in the central nervous systems of larvae (Cullen et al., 1999). Although normal in many respects, larvae homozygous for msps mutations grow more slowly and die before pupation. Dissection revealed that msps larvae lacked imaginal discs. Cytological examination of the developing CNS revealed an increased mitotic index and paucity

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of anaphase cells, indicating that msps arrested the cell cycle in M phase. Immunostaining with antitubulin subsequently revealed that the msps mutation disrupted the organization of mitotic spindles, with a high incidence (more than 50%) of multipolar and disorganized spindles (Cullen et al., 1999). Cloning and sequencing of the Msps gene from D. melanogaster predicted that it encodes a protein product of 2050 amino acids that is related to the MAP215/Dis1 family of MAPs (Cullen et al., 1999). In addition to Msps in D. melanogaster, an apparently complete Msps-related protein has been predicted from genomic DNA sequences of Drosophila pseudoobscura (DpMsps) and partial sequences from Anopheles gambia (mosquito; AgMsps: 1843 amino acids) and honey bee Apis mellifera (AmMsps: 1090 amino acids) (Appendix I). The four Msps-related proteins in insects share 56–76% amino acid identity over their entire sequence and are 28–34% identical to XMAP215 in Xenopus. Immunolocalization revealed that Msps protein is associated with centrosomes throughout the cell cycle in syncytial embryos and throughout the spindle (including the poles) during M phase (Cullen et al., 1999). The latter observation suggests that Msps associates with MTs of the central spindle, rather than specifically with kinetochores or kinetochore MTs, and is consistent with results demonstrating that Msps protein binds to MT in vitro (Cullen et al., 1999). A similar distribution of Msps protein is observed in cellularized embryos during M phase. However, Msps does not localize to centrosomes of interphase cells postcellularization (Cullen et al., 1999). Msps also is observed at the acentrosomal poles of female meiotic spindles in Drosophila (Cullen and Ohkura, 2001), and msps mutations disrupt the maintenance of meiotic spindle organization, leading to a high incidence of tripolar spindles (Cullen and Ohkura, 2001). On the basis of the observed defects in spindle assembly in msps mutants, Msps has been proposed to play roles in the anchoring and bundling of spindle MTs required for establishment and maintenance of spindle organization (Cullen and Ohkura, 2001; Cullen et al., 1999). Recent studies demonstrate that the targeting of Msps to centrosomes and spindles poles requires D-TACC (Gergely et al., 2000b; Giet et al., 2002; Lee et al., 2001), a member of the conserved family of TACC proteins identified in diverse taxa including C. elegans, D. melanogaster, and vertebrates (Bellanger and Gonczy, 2003; Conte et al., 2003; Gergely, 2002; Gergely et al., 2000a,b; Le Bot et al., 2003; Lee et al., 2001; RaV, 2002; Srayko et al., 2003; Theurkauf, 2001). D-TACC and Msps form a physical complex, and disruption or depletion of D-TACC phenocopies many of the eVects observed with mutations in Msps, resulting in the formation of disordered, multipolar spindles (Gergely et al., 2000b; Giet et al., 2002; Lee et al., 2001). As in C. elegans, targeting of D-TACC to centrosomes and spindle poles is


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dependent on Aurora-kinase (Giet et al., 2002). Interestingly, D-TACC is also required for the polar localization of Msps in the acentrosomal spindles of female meiosis in Drosophila (Cullen and Ohkura, 2001), suggesting that the interplay between these proteins is not restricted to conventional centrosomes. All Msps-related proteins are predicted to include four TOG domains [(1A:1B)(2A:2B)], each with two to five predicted HEAT repeats and sequences similar to the TOG domain signature sequences. Two additional clusters of HEAT repeats are predicted between amino acids 1210–1385 (four HEAT repeats) and 1580–1780 (four HEAT repeats). However, these clusters lack the conserved TOG domain signature sequences. Conservation between the insect proteins and vertebrate MAP215/Dis1-related proteins is highest in the N-terminal TOG domains: The amino acid sequences of the first TOG domain of the insect proteins are 50–52% identical to the corresponding region of XMAP215, and amino acid sequence identities over the first 1150 amino acids range from 41 to 42%. All of the Msps-related proteins for which suYcient sequence is available are predicted to include from two to five predicted targets for the CDK1 cell cycle kinase. Although no such target sequences are found in the available sequence from Apis mellifera (the N-terminal 1090 amino acids), the majority of the predicted CDK1 targets in the other Msps-related proteins are found in the C-terminal half of the protein. Both of the Msps proteins from Drosophilids include an 18–amino acid sequence near their C termini that exhibits limited similarity (33% identity and 56% similarity) to the consensus sequence of the single Tau repeat observed in other members of the MAP215/Dis1 family. These putative Tau repeats are the most divergent of any identified in the MAP215/Disrelated proteins in animals. Interestingly, they are most similar to the putative Tau repeat in the basidiomycote fungus U. maydis, with which they share 50% amino acid sequence identity and 61% similarity. Like the U. maydis Tau repeat (and those from plants), the conserved serine of the core KIGS sequence has been replaced with the phosphomimic aspartate (D) in both Drosophilid sequences.

D. Structure and Function of XMAP215/ch-TOG-Related Proteins in Chordates XMAP215, one of the founding members of the MAP215/Dis1 family of MAPs, was first identified and isolated from Xenopus eggs by virtue of its ability to promote centrosome-nucleated MT assembly in vitro (Gard and Kirschner, 1987). The subsequent cloning and sequencing of cDNAs encoding human ch-TOG (Charrasse et al., 1995), the human homolog of

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XMAP215 (Charrasse et al., 1998), forged the link that bound related proteins from yeast, nematodes, flies, and vertebrates into the evolutionarily ancient MAP215/Dis1 family of MAPs. As might be expected of homologous proteins from two species of vertebrates, XMAP215 and ch-TOG share a number of functional properties: both proteins bind to MTs in vitro (Charrasse et al., 1998; Gard and Kirschner, 1987), both proteins promote MT assembly in vitro (Charrasse et al., 1998; Gard and Kirschner, 1987; Vasquez et al., 1994; however, the eVect of ch-TOG on MT assembly is much less dramatic than that of XMAP215), both are localized to spindle MTs in vivo (Becker et al., 2003; Tournebize et al., 2000) (Fig. 10), and both XMAP215 and ch-TOG are associated with centrosomes or spindle poles in vivo (Charrasse et al., 1998; Gard et al., 1995; Popov et al., 2001; Tournebize et al., 2000; see Fig. 10 and Fig. 11). In recognition of the extensive structural and functional similarities between these two MAP215/Dis1-related proteins, discussion of the structures and functions of XMAP215 and ch-TOG will be combined. However, it should be remembered that these proteins are not identical, and significant divergence in structure and function might have occurred during the separate evolution of frogs and humans. 1. Structure of XMAP215/ch-TOG The XMAP215/TOG subfamily of MAP215/Dis1-related proteins in chordates and hemichordates currently includes 12 members (see Appendix I), two of which (XMAP215 in X. laevis and ch-TOG in humans) have been studied in detail at both the structural and functional levels. The remaining family members are proteins predicted from genome sequences or ESTs and include complete or nearly complete protein sequences for Xenopus tropicalis, mouse (Mus musculus), rat (Rattus norvegicus), zebrafish (Danio rerio), and puVerfish (Takefugu rubrio), and partial sequences (ranging from a few hundred to more than 1000 amino acids) predicted from incomplete genome sequences or ESTs from chicken (Gallus gallus), lizard (Anolis sagrei), and two species of tunicates (C. intestinalis and C. savignyi). Members of the XMAP215/TOG subfamily share 44–94% amino acid identity (from tunicates through humans) and are more distantly related to the Msps (28–47% identity) and Zyg9 (15–30% identity) subfamilies in animals, the Mor1/Gem1-related proteins in plants (17–27% identity), and Dis1-related proteins in fungi (11–25% identity). In addition to sequence similarities, members of the XMAP215/TOG subfamily share structural features characteristic of MAP215/Dis1-related proteins in animals, including at least four TOG domains, each with up to five HEAT repeats, one or more predicted CDK1 phosphorylation targets, and a single C-terminal Tau repeat.


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FIG. 10 XMAP215 is associated with interphase MTs and mitotic spindles in Xenopus blastomeres, and with meiotic spindles in maturing Xenopus oocytes. Confocal immunofluorescence microscopy reveals that XMAP215 is associated with MTs in interphase blastomeres (A–A00 ), spindle MTs in mitotic blastomeres (B–B00 ), and meiotic spindles in maturing oocytes (C–C00 ). XMAP215 appears to be distributed as punctae along MTs in all stages of the cell cycle, and is concentrated at the plus ends of some MTs in interphase cells (small arrowheads in A– A0 ). Note the prominent staining of the centrosomes (larger arrowheads in A–A0 ) and midbodies (arrows in A–A0 ) in interphase, and spindle poles in mitosis (arrowheads in B–B0 ). In contrast, XMAP215 is not substantially concentrated at the poles of meiotic spindles (arrowheads in C–C0 ). A–C were stained with monoclonal anti-a-tubulin, A0 –C0 were stained with a cocktail of three XMAP215 anti-peptide antibodies, and A00 –C00 were stained with Yo-Pro 3 (Becker et al., 2003). C–C 0 are reproduced from Becker et al. (2003) with permission.

The sequences of XMAP215, ch-TOG, and other family members, are predicted to include four TOG domains (amino acid numbering for XMAP215): 1A (amino acids 1–263), 1B (amino acids 264–543), 2A (amino acids 580–839), and 2B (amino acids 840–1115). Each TOG domain

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GARD ET AL.

FIG. 11 Depletion of ch-TOG or XMAP215 in vivo disrupts centrosome structure and spindle assembly during M-phase. (A) A bipolar spindle with focused poles in a normal (untreated) HeLa cell. (B) a spindle in ch-TOG-depleted HeLa cells exhibits multiple poles and detachment of spindle microtubules from the spindle poles (see Cassimeris and Morabito, 2004). (C–C0 ) a normal bipolar spindle in a 10 hour Xenopus blastomere. (D–D0 ) a multipolar spindle in a blastomere from a Xenopus embryo injected with a morpholino antisense oligonucleotide specific for XMAP215 (C and D stained with antitubulin; C0 –D0 stained with YoPro-3). Figures 11A and B are courtesy of Dr. Cassimeris.

includes sequences similar to the conserved signature sequence, is predicted to include up to five closely spaced HEAT repeats, and includes a C-terminal trailing sequence of variable length. Neuwald and Hirano (2000) predicted that ch-TOG (and, by inference, XMAP215, with which it shares 80% sequence identity) (Becker and Gard, 2000; Tournebize et al., 2000) included 20 HEAT repeats: 16 HEAT repeats organized into the four TOG domains, and four additional repeats located between amino acids 1243 and 1396 (corresponding to amino acids 1277–1430 in XMAP215) which may represent a fifth, less well conserved, TOG domain. However, this potential fifth TOG domain, similar to those found in other taxa, lacks a conserved signature sequence. On the basis of alignment of the four TOG domains, Cassimeris et al. (2001) predicted that each TOG domain in XMAP215 might include five HEAT repeats, bringing the total number of HEAT repeats in XMAP215 and ch-TOG to at least 24. Purified XMAP215 behaves like an elongate monomer in solution (Cassimeris et al., 2001; Gard and Kirschner, 1987), and electron microscopy revealed a semiflexible rod-shaped molecule


225

FIG. 12 XMAP215 is a long, thin molecule. A gallery of unidirectionally shadowed images of XMAP215 from two diVerent XMAP215 preparations. The mean XMAP215 length was 60 nm. Most molecules were straight, but a significant number were bent. Reproduced from Cassimeris et al. (2001) with permission.

with a contour length of 60 nm (Cassimeris et al., 2001; see Fig. 12). From comparison of the calculated mass density of XMAP215 with that of other HEAT repeat proteins, Cassimeris et al. (2001) concluded that XMAP215 might consist almost entirely of HEAT repeats. XMAP215 and ch-TOG also include a serine- and proline-rich (S,P) domain (amino acids 10001200) similar to that found in MAP4 (West et al., 1991), and a single Tau repeat. The functional significance of the S,P domain in XMAP215 and ch-TOG is unclear. The Tau repeat is proposed to function in targeting XMAP215 and ch-TOG to centrosomes (Sections I.B.3 and V.D.5). 2. XMAP215 Promotes Microtubule Assembly and Dynamics In Vitro XMAP215 and ch-TOG are arguably the most well characterized members of the MAP215/Dis1 family of MAPs in terms of both structure and function. Initial characterization of XMAP215 activity revealed two striking features: first, XMAP215 dramatically promoted MT assembly to a degree not seen with more conventional MAPs, and second, XMAP215 preferentially promoted assembly at MT plus ends, with little eVect on minus-end assembly

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GARD ET AL.

(Gard and Kirschner, 1987). Subsequent studies revealed that, unlike previously characterized MAPs that suppress the dynamic behavior of MTs (Drechsel et al., 1992; Kowalski and Williams, 1993; Pryer et al., 1992), XMAP215 promotes both the assembly and dynamics of MT plus ends (Vasquez et al., 1994). Specifically, addition of XMAP215 to purified tubulin promoted a substantial (as much as 10-fold at a molar ratio of 1:20) increase in the rate of plus-end elongation, promoted a two- to three-fold increase in the rate of rapid shortening, eliminated rescue, and did not suppress catastrophe (Vasquez et al., 1994). In all aspects, XMAP215 preferentially aVected MT plus ends, with little or no eVects noted on minus-end assembly or dynamics. Thus, in a system of reconstituted components, XMAP215 promotes the assembly of long, highly dynamic MTs (Gard and Kirschner, 1987; Vasquez et al., 1994). Depletion of XMAP215 substantially reduced MT assembly and increased MT dynamics in Xenopus egg extracts (Tournebize et al., 2000). Immunodepletion of XMAP215 from interphase extracts, to 30–50% of its normal abundance in eggs, resulted in a corresponding decrease in the rates of elongation and rapid shortening (Tournebize et al., 2000), consistent with the observed promotion of elongation and rapid shortening of MTs assembled from purified components (Vasquez et al., 1994). In contrast, immunodepletion of XMAP215 from M-phase arrested egg extracts had no apparent eVect on the rates of either MT elongation or rapid shortening (Tournebize et al., 2000). For comparison, nearly complete immunodepletion of the putative MAP4 homolog XMAP230, which is also abundant in Xenopus eggs and early embryos, had no significant eVect on MT elongation rates in either interphase or M-phase egg extracts (Cha et al., 1999). These observations indicate that XMAP215 plays a significant role in regulating the rates of MT elongation and rapid shortening during interphase in Xenopus eggs and are consistent with the proposed role of XMAP215 in promoting the rapid assembly of the sperm aster and cortical MT arrays of fertilized Xenopus eggs (Gard and Kirschner, 1987; Gard et al., 1995). In contrast to results obtained from purified components, depletion of XMAP215 from either interphase or mitotic egg extracts strongly promotes catastrophe (Tournebize et al., 2000): observed catastrophe frequencies in XMAP215-depleted interphase extracts were sevenfold higher than those of mock-depleted interphase extracts, whereas those of XMAP215-depleted M-phase extracts are more than twofold greater than similarly prepared control extracts. The promotion of MT catastrophe induced by depletion of XMAP215 in both interphase and M-phase extracts was nearly completely suppressed by addition of antibodies that inhibit XKCM1 (Tournebize et al., 2000), a catastrophe-promoting member of the KinI family of kinesin-related proteins (Desai et al., 1999; Kline-Smith and Walczak, 2002; Walczak et al., 1996). These results indicate that XMAP215 and XKCM1


227

act antagonistically to regulate MT dynamics by modulating the frequency of catastrophe during both M phase and interphase (Tournebize et al., 2000). The subsequent reconstitution of many of the dynamic parameters of MT assembly observed in extracts with a three-component system consisting of XMAP215, XKCM1, and tubulin (Kinoshita et al., 2001) adds further support to the antagonistic roles of XMAP215 and XKCM1 and their importance in regulating MT dynamics. However, the contribution of other cytoplasmic factors, including XMAP230, XMAP310, stathmin, EB1, and others, and the role of XMAP215 in modulating the rates of MT elongation and rapid shortening during interphase, should not be discounted. The observed diVerences between XMAP215-depleted extracts in interphase and M phase may be reflections of cell cycle–dependent regulation of XMAP215 function. XMAP215 is hyperphosphorylated in M phase (Gard and Kirschner, 1987), and phosphorylation by CDK1 in vitro reduces assembly promotion without aVecting MT binding (Vasquez et al., 1999; Section V.D.6). Depletion of XMAP215, whose elongation- and disassembly promoting functions have already been ‘‘turned oV’’ by CDK1-dependent phosphorylation, might not be expected to have substantial eVects on MT elongation or rapid shortening rates in M-phase extracts. However, phosphorylation by CDK1 had little eVect on XMAP215 binding (Vasquez et al., 1999), and phosphorylated XMAP215 might still be suYcient to suppress XKCM1-induced catastrophes, as observed in M-phase extracts (Tournebize et al., 2000). The biochemical/molecular mechanism or mechanisms by which XMAP215 promotes both plus-end elongation and rapid shortening remain unclear. The observed enhancement of plus-end elongation rates exceeds that which could be obtained by reducing or eliminating subunit loss through reductions in the oV-rate constant (KoV) (Gard and Kirschner, 1987; Vasquez et al., 1994). The novel eVects of XMAP215 on MT dynamics indicate that the interactions between XMAP215 and MTs are distinct from those of other well-characterized MAPs (Tau, MAP2, and MAP4), a conclusion that is supported by results from studies of ch-TOG binding to MTs in vitro (Spittle et al., 2000; discussed in more detail later). Models for promotion of elongation have included facilitating orientation or addition of tubulin dimers, facilitating addition of tubulin oligomers, or increasing the number of protofilaments available for subunit addition (Gard and Kirschner, 1987; Spittle et al., 2000; Vasquez et al., 1994). Proposed models for promotion of rapid shortening by XMAP215 have included XMAP215-induced weakening/stress of lateral interactions between protofilaments, introduction of defects into the MT lattice, or modulation of tubulin’s intrinsic GTPase activity (Shirasu-Hiza et al., 2003; Vasquez et al., 1994, 1999). Recently, Shirasu-Hiza et al. (2003) reported that full-length and N-terminal fragments of XMAP215 promote disassembly

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of GMPPCP-stabilized MTs, both in the context of egg extracts and using purified components. The ends of GMPPCP MTs are blunt (Shirasu-Hiza et al., 2003). In contrast, MTs undergoing elongation or rapid shortening exhibit sheet-like extensions, or splayed protofilaments, respectively (Chretien et al., 1995; Mandelkow et al., 1991; Simon and Salmon, 1990). On the basis of these diVerences in ultrastructure, Shirasu-Hiza et al. (2003) suggested that the ends of GMPPCP MTs represent ‘‘paused’’ MTs (those that are neither growing nor shrinking) and proposed a model in which XMAP215 inhibits the paused state, favoring either growth or rapid shortening. In their model, XMAP215-dependent reduction of shortduration ‘‘micropauses’’ could contribute to the modest (two- to threefold) enhancement of rapid shortening produced by XMAP215 (Shirasu-Hiza et al., 2003). However, it is less clear how this model accounts for XMAP215’s dramatic promotion of MT elongation rates (six- to 10-fold at subsaturating stoichiometries). Characterization of the functional eVects of ch-TOG on the assembly of MTs in vitro has been much less extensive, largely because of technical and other limitations of purifying the required amount of material from mammalian sources. However, partially purified fractions enriched in ch-TOG from porcine brain enhanced the rate and extent of tubulin polymerization in turbidometric assays and promoted a twofold increase in the elongation rate of individual MTs viewed by video-enhanced DIC microscopy (Charrasse et al., 1998). In contrast to the marked plus-end preference of XMAP215, porcine ch-TOG aVected the plus and minus ends of MTs equally (Charrasse et al., 1998). Whether ch-TOG’s lack of plus-end preference results from technical limitations, or rather reflects an inherent diVerence in the functions of ch-TOG and XMAP215, remains unclear. In addition to its eVect on the dynamics of individual MTs, Popov et al. (2002) demonstrated that XMAP215 is required for MT nucleation by centrosomes in Xenopus eggs extracts. In their studies, incubation in XMAP215-depleted Xenopus egg extracts was unable to restore MT nucleation from salt-stripped centrosome scaVolds, in contrast to untreated or mock-depleted extracts. Moreover, XMAP215-coated beads nucleated MT asters both in egg extracts and from purified tubulin. Nucleation of MT asters by XMAP215-beads in egg extracts was dependent on g-tubulin (Popov et al., 2002), suggesting that XMAP215 cooperates with g-tubulin to nucleate MTs in cytoplasm. However, XMAP215-loaded beads were able to nucleate MTs and organize asters from purified tubulin in the absence of any other proteins. The authors concluded that XMAP215 is able to both nucleate MT assembly from purified tubulin, perhaps by stabilizing protofilament interactions, and anchor MTs and organize them into asters (Popov et al., 2002). Together with the observed localization of XMAP215 and


229

other MAP215/Dis1 family members to centrosomes and other MTOCs (Charrasse et al., 1998; Gard et al., 1995; Tournebize et al., 2000), these results suggest that XMAP215 may play a role in MT nucleation or anchoring in vivo (for additional discussion of the role of XMAP215 at MTOCs, see Section VI.D). 3. Role of XMAP215/ch-TOG in the Organization of Spindles and Spindle Poles XMAP215 is associated with interphase MTs in cultured Xenopus cells (Tournebize et al., 2000) and early Xenopus embryos (Fig. 10A,B) (B. Becker and D. Gard, unpublished observation), where it appears to be enriched at the plus-ends of some MTs (arrows in figure 10A–B). In addition, both XMAP215 and ch-TOG are conspicuously associated with spindle poles and spindle MTs during M phase (see Fig. 10) (Becker et al., 2003; Charrasse et al., 1998; Gard et al., 1995; Popov et al., 2001; Tournebize et al., 2000). Several recent studies have addressed the role or roles of TOG or XMAP215 in spindle assembly and MT dynamics in vivo by using antibody injections (Becker et al., 2003) or siRNA to deplete the proteins in living cells (Cassimeris and Morabito, 2004; Conte et al., 2003; Gergely et al., 2003; Holmfeldt et al., 2004). Becker et al. (2003) reported that XMAP215 is associated with MTs of the first meiotic spindle, and the transient MT array from which it is derived, during the progesterone-induced maturation of Xenopus oocytes (Becker et al., 2003). In contrast to the observed localization of XMAP215 to the poles of mitotic spindles in Xenopus embryos (Gard et al., 1995; see Fig. 9), and cultured cells (Popov et al., 2001; Tournebize et al., 2000) and Msps to the acentrosomal poles of meiotic spindles in Drosophila oocytes (Cullen and Ohkura, 2001), XMAP215 appeared uniformly distributed throughout the first meiotic spindles of Xenopus oocytes, with no apparent concentration at the poles. Microinjection of anti-XMAP215 antibodies dramatically disrupted spindle assembly and organization (Becker et al., 2003). Most antibody-injected oocytes assembled numerous small asters of short MTs that appeared to be nucleated from aggregates of XMAP215 and the injected antibody, consistent with MT nucleation from beads coated with XMAP215 or XMAP215 antibodies reported by Popov et al. (2002), whereas the animal hemispheres of the remaining injected oocytes were largely devoid of MTs. Microinjection of anti-XKCM1, a member of the KinI family of kinesinrelated catastrophe factors (Desai et al., 1999; Walczak et al., 1996) also disrupted spindle assembly (Becker et al., 2003) but appeared to promote MT assembly or stabilization. The observed defects in spindle assembly and organization in oocytes injected with anti-XMAP215 or anti-XKCM1 are

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GARD ET AL.

consistent with the antagonistic eVects of these two proteins on MT assembly and dynamics in Xenopus egg extracts in vitro (Kinoshita et al., 2001; Tournebize et al., 2000). In contrast, results from recent studies indicate that siRNA-induced depletion of ch-TOG in cultured human cells has little eVect on overall MT assembly or dynamics, but profoundly aVects the organization of mitotic spindles by dramatically disrupting the organization and function of the spindle poles (Cassimeris and Morabito, 2004; Conte et al., 2003; Gergely et al., 2003; Holmfeldt et al., 2004). Depletion of ch-TOG had no apparent aVect on the assembly and organization of interphase MTs (Cassimeris and Morabito, 2004; Holmfeldt et al., 2004) and little eVect (1727

AACO01000063.1

Causitive agent of cryptococcosis. Predicted protein from whole genome shotgun sequence. Cryptococcus neoformans var. grubii H99 cont1.63_scaVold6.

Ustilago maydis

UmMAP215

2238

AACP01000242.1

Corn smut. Predicted protein from whole genome shotgun sequence. Ustilago maydis 521 cont1.242_scaVold23.

PcMAP215

>1970

ScaVold_3|351753|358780

White rot. Predicted protein from draft 1 of the P. chrysosporium genome. Sequenced by the D.O.E. Joint Genome Institute: http://genome.jgi-psf.org/whiterot/

Arabidopsis thaliana

Mor1 Gem1

1978

AF367246.1

Thale crest. Protein predicted from mRNA, complete cds. Whittington et al., 2001.

Chlamydomonas reinhardtii

CrMor1

?

ScaVold_376|4879|15895

Green alga. 800 amino acids predicted from incomplete genome sequence using AtMor1 as template. Sequenced by the D.O.E. Joint Genome Institute: http://genome.jgipsf.org/chlamy/

Glycine max

GmMor1

?

Gm_002_283485

Soybean. Three ESTs predict 435 amino acids, including the Tau repeat: amino acids 1574–1778, 1736–1900, and 1926–1978 (based on AtMOr1). EST sequences from The National Center of Genomic Resources Legume Information Center: http://www.comparativelegumes.org/

Phanerochaete chrysosporium Plants and algae

Gm_001_358471 Gm_001_300967

259 Medicago truncatula

MtMor1

?

Mt_002_66354 Mt_001_94112 Mt_001_106339 Mt_001_109182

Four ESTs predict 680 amino acids including the Tau repeat: amino acids 1269–1394, 1424–1650, 1706–1950, and 1894–1978 (based on AtMOR1). EST sequences from The National Center of Genomic Resources Legume Information Center: http://www.comparativelegumes.org/

Nicotiana Tabaccum

TMBP200

2029

BAB88648.1

Tobacco. Predicted from mRNA. Yasuhara et al., 2002.

Oryza sativa

OrMor1

1920

BAB64822

Rice. Predicted from Oryza sativa nipponbare (GA3) genomic DNA, chromosome 1, BAC clone:B1148D12.

Physcomitrella patens

PpMor1

?

BJ582083.1

Moss. 71 amino acids, including the Tau repeat, predicted from an EST. http://www.cosmoss.org/ (continued )

APPENDIX I (continued) Organism Populus trichocarpa

Gene/Protein PtMor1

Amino Acids >1800

Accession Number (or other identification) Individual sequence chromatograms.

Notes Poplar. Amino acids 1–890 and 906–1842 (based on AtMor1) predicted from incomplete genome sequenced by the D.O.E. Joint Genome Institute: http://genome.jgi-psf.org/ poplar/

260

Individual sequence chromatograms: POPS11988; TREE539817.b1; TREE688230.g1; TREE730147.y1; TREE877788.b1; POPS16017.b1; TREE289774.b2; TREE843658.g1 TREE90555.b2; VAO12782.g2; XX1299199; YFS7464.g1; YFS130568.x1; YFS797727; YFS 418233 Zea Mays

ZmMor1

CD052294 BM428946 BM418349 BM895204 CF037203 BM277032 Al649650 BM428770 CD986907 BQ048821 CD991588 CA828389

Corn. Assembled from EST sequences from the NCBI/ Genbank (http://www.ncbi.nlm.nih.gov/BLAST/) Additional ESTs available from the Maize Genetics and Genome Database (http://www.maizegdb.org/blast.php).

Animals: nematodes Brugia malayi

BrnZyg9

Caenorhabditis elegans

Zyg9

Caenorhabditis briggsae

CbZyg9

1415

TIGR_6279_1132919

Causitive agent of filiarisis. Amino acids 9–867 predicted from incomplete genome sequence. The Sequencing eVort is part of the International Brugia Genome Sequencing Project and is supported by an award from the National Institute of Allergy and Infectious Diseases, National Institutes of Health.

AF03519

Predicted protein from mRNA, complete CDS. Matthews et al., 1998.

CAE57635.1

Hypothetical protein CBG00620; Nematode Sequencing Project, Wellcome Trust Sanger Institute and Genome Sequencing Center, Washington University, St. Louis, MO 63110.

Animals: Insects

261

Anopheles gambia

AgMsps

>1850

XP_309487.1 EAA05144

Mosquito. 1851 N-terminal amino acids predicted from genome sequence. The Anopheles Genome Sequencing Consortium.

Apis metlifera

AmMsps

>1125

AADG01012959.1

Honey bee. 1125 N-terminal amino acids predicted from whole genome shotgun sequence, Apis mellifera strain DH4 Contig 12959. Splicing predicted from predicted. D. melanogaster msps.

Drosophila melanogaster

Msps

2050

AJ249115

Drosophila melanogaster msps gene for microtubule associated protein. Cullen et al., 1999.

Drosophila pseudoobscura

DpMsps

2175

AADE01000185.1

Protein predicted from whole genome sequence, chromosome 2 strain MV2–25 Cont7464. Splicing predicted from predicted D. melanogaster msps. (continued )

APPENDIX I (continued) Organism

Amino Acids

Gene/Protein

Accession Number (or other identification)

Notes

Animals: chordates Ciona intestinales

CiMAP215

?

AABS01000174.1

Tunicate. Predicted protein from whole genome shotgun sequence. Similarity with the predicted Xenopus, Fugu, and C. savignyii proteins were used to predict the first 1375 amino acids of the C. intestinalis protein.

Ciona savignyi

CsMAP215

?

AACT01008099.1

Tunicate. Predicted protein from overlapping contigs of whole genome shotgun sequence. Similarity with the predicted Xenopus, Fugu, and C. intestinalis proteins were used to predict the first 1375 amino acids of the C. intestinalis protein.

AACT01010629.1 AACT01041397.1 AACT01032920.1

262

AACT01010626.1 AACT01008098.1 AACT01010625.1 Takifugu rubripes

TrMAP215

Danio rerio

DrMAP215

2032

CAAB01004128.

PuVerfish. Predicted protein from whole genome sequence. Shotgun assembly SCAFFOLD_412.

ENSDARG00000005446

Zebrafish. Ensembl Danio rerio (http://www.ensembl.org/ Danio_rerio/).

AL927739.1

cDNAs encoding the C-terminus of the D. rerio MAP215 are available through NCBI (http://www.ncbi.nim.nih.gov/)

AW175221.1 CK362884.1 CK361751. 1BM861374.1 CB354558.1 BQ479631.1 BQ263962.1

Ictalurus Punctata

lpMAP215

?

IpuJx13_39_A02_23.ab1 IpuJx14_27_CO4_23_13May02_0 34.ab1

Xenopus laevis

XMAP215

2065

CAB61894

2066

AAG34914

2030

AAG34915

Channel catfish. Two ESTs encoding amino acids 1446–1712 (numbering based upon human KIAA0097). http:// morag.umsmed.edu/ African clawed frog. Predicted proteins from mRNA, complete CDS. The predicted products of CAB61894 and AAG34914 diVer by 72 single amino acid substitutions and a single amino acid insertion. It is not clear whether these reflect natural polymorphism or cloning errors. The predicted products of AAG34914 and AAG34915 diVer by 24 single amino acid substitutions and the presence of a 36 amino acid insert (in AAG34914) that represents developmentally-regulated alternative splicing (see text). Tournebize et al., 2000; Becker and Gard, 2000.

Xenopus tropicalis

XtMAP215

263 Anolis sagrei

AsMAP215

>2000

?

scaVold_26739 13753

Protein predicted from incomplete genome sequence using XMAP215 as template.

scaVold_40867 8424

Sequenced by the D.O.E. Joint Genome Institute: http:// genome.jgi-psf.org/xenopus/

scaVold_8041 29765

The predicted sequence includes Ins1 and Ins2 (see text).

CB035631

Anolis lizard. Overlapping partial cDNA sequences from limb bud. Together, these two sequences encode 162 amino acids with similarity to MAP215. Library was constructed by Jeremy Gibson-Brown. DNA sequencing by: Washington University Genome Sequencing Center.

CB280918

Gallus gallus

GgMAP215

>2000

ChEST1004b23 (aa 1-200) ChEST808m11 (aa 105–373) ChEST591c12 (aa 262–471)

Chicken. A nearly complete sequence can be compiled from ESTs encoding MAP215-related proteins included in the BBSRC ChickEST Database: http://www.chick.umist.ac.uk/

ChEST174b17 (aa 580–793) ChEST65e9 (aa 987–1206) ChEST853f15 (aa 1146–1366) (continued )

APPENDIX I (continued) Organism

Gene/Protein

Amino Acids

Accession Number (or other identification) ChEST394g14 (aa 1398–1618) ChEST407k2 (aa 1575–1684) ChEST373h4 (aa 1690–1869)

Notes Ten clones provided as examples (amino acid numbering corresponding to ch-TOG); Clone ChEST892n2 (amino acid 1806–2032) includes the Tau repeat.

ChEST892n2 (aa 1806–2032)

264

Mus musculus

MmMAP215

2025

AL691489

House mouse. The predicted sequence includes Ins1 (see text).

Rattus norvegicus

RNMAP215

>1900

XP_230281 (N-term 870 aa)

Norwegian rat. These reference sequences were predicted from NCBI contig NW_043641 and NW_047647 by automated computational analysis. Additional ESTs are also identified by BLAST of NCBI databases. XP 230281 appears to be missing one or more internal exons.

XP_230282 (C-term 1915 aa)

Homo sapiens

ch-TOG

1972

NP_055571

KIAA0097

2032

BAA07892

Human. ch-TOG (NP 055571) lacks both Ins1 and Ins2; KIAA00097 includes Ins1 but lacks Ins2. Charrasse et al., 1995; Nagase et al., 1995.


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References Albertson, D. G. (1984). Formation of the first cleavage spindle in nematode embryos. Dev. Biol. 101, 61–72. Amos, L. A., and Baker, T. S. (1979). The three-dimensional structure of tubulin protofilaments. Nature 279, 607–612. Andersen, S. S., and Karsenti, E. (1997). XMAP310: A Xenopus rescue-promoting factor localized to the mitotic spindle. J. Cell. Biol. 139, 975–983. Andersen, S. S., Buendia, B., Dominguez, J. E., Sawyer, A., and Karsenti, E. (1994). EVect on microtubule dynamics of XMAP230, a microtubule-associated protein present in Xenopus laevis eggs and dividing cells. J. Cell. Biol. 127, 1289–1299. Andrade, M. A., and Bork, P. (1995). HEAT repeats in the Huntington’s disease protein. Nat. Genet. 11, 115–116. Andrade, M. A., Perez-Iratxeta, C., and Ponting, C. P. (2001a). Protein repeats: structures, functions, and evolution. J. Struct. Biol. 134, 117–131. Andrade, M. A., Petosa, C., O’Donoghue, S. I., Muller, C. W., and Bork, P. (2001b). Comparison of ARM and HEAT protein repeats. J. Mol. Biol. 309, 1–18. Baker, T. S., and Amos, L. A. (1978). Structure of the tubulin dimer in zinc-induced sheets. J. Mol. Biol. 123, 89–106. Baldauf, S. L., Roger, A. J., Wenk-Siefert, I., and Doolittle, W. F. (2000). A kingdom-level phylogeny of eukaryotes based on combined protein data. Science 290, 972–977. Becker, B. E., and Gard, D. L. (2000). Multiple isoforms of the high molecular weight microtubule associated protein XMAP215 are expressed during development in Xenopus. Cell Motil. Cytoskeleton 47, 282–295. Becker, B. E., Romney, S. J., and Gard, D. L. (2003). XMAP215, XKCM1, NuMA, and cytoplasmic dynein are required for the assembly and organization of the transient microtubule array during the maturation of Xenopus oocytes. Dev. Biol. 261, 488–505. Bellanger, J. M., and Gonczy, P. (2003). TAC-1 and ZYG-9 form a complex that promotes microtubule assembly in C. elegans embryos. Curr. Biol. 13, 1488–1498. Belmont, L. D., Hyman, A. A., Sawin, K. E., and Mitchison, T. J. (1990). Real-time visualization of cell cycle-dependent changes in microtubule dynamics in cytoplasmic extracts. Cell 62, 579–589. Biernat, J., Gustke, N., Drewes, G., Mandelkow, E. M., and Mandelkow, E. (1993). Phosphorylation of Ser262 strongly reduces binding of tau to microtubules: Distinction between PHF-like immunoreactivity and microtubule binding. Neuron 11, 153–163. Cassimeris, L. (2002). The oncoprotein 18/stathmin family of microtubule destabilizers. Curr. Opin. Cell Biol. 14, 18–24. Cassimeris, L., Gard, D., Tran, P. T., and Erickson, H. P. (2001). XMAP215 is a long thin molecule that does not increase microtubule stiVness. J. Cell Sci. 114, 3025–3033. Cassimeris, L., and Morabito, J. (2004). TOGp, the human homolog of XMAP215/Dis1, is required for centrosome integrity, spindle pole organization, and bipolar spindle assembly. Mol. Biol. Cell 15, 1580–1590. Cassimeris, L., Pryer, N. K., and Salmon, E. D. (1988). Real-time observations of microtubule dynamic instability in living cells. J. Cell Biol. 107, 2223–2231. Cassimeris, L., and Spittle, C. (2001). Regulation of microtubule-associated proteins. Int. Rev. Cytol. 210, 163–226. Cha, B., Cassimeris, L., and Gard, D. L. (1999). XMAP230 is required for normal spindle assembly in vivo and in vitro. J. Cell Sci. 112, 4337–4346. Chapin, S. J., and Bulinski, J. C. (1991). Non-neuronal 210 10(3) Mr microtubule-associated protein (MAP4) contains a domain homologous to the microtubule-binding domains of neuronal MAP2 and tau. J. Cell Sci. 98, 27–36.

266

GARD ET AL.

Chapin, S. J., and Bulinski, J. C. (1992). Microtubule stabilization by assembly-promoting microtubule-associated proteins: A repeat performance. Cell Motil. Cytoskeleton 23, 236–243. Charrasse, S., Coubes, P., Arrancibia, S., and Larroque, C. (1996). Expression of the tumor over-expressed ch-TOG gene in human and baboon brain. Neurosci. Lett. 212, 119–122. Charrasse, S., Mazel, M., Taviaux, S., Berta, P., Chow, T., and Larroque, C. (1995). Characterization of the cDNA and pattern of expression of a new gene over-expressed in human hepatomas and colonic tumors. Eur. J. Biochem. 234, 406–413. Charrasse, S., Schroeder, M., Gauthier-Rouviere, C., Ango, F., Cassimeris, L., Gard, D. L., and Larroque, C. (1998). The TOGp protein is a new human microtubule-associated protein homologous to the Xenopus XMAP215. J. Cell Sci. 111, 1371–1383. Chen, H. M., Schmeichel, K. L., Mian, I. S., Lelievre, S., Petersen, O. W., and Bissell, M. J. (2000). AZU-1: A candidate breast tumor suppressor and biomarker for tumor progression. Mol. Biol. Cell 11, 1357–1367. Chen, X. P., Yin, H., and HuVaker, T. C. (1998). The yeast spindle pole body component Spc72p interacts with Stu2p and is required for proper microtubule assembly. J. Cell Biol. 141, 1169–1179. Chretien, D., Fuller, S. D., and Karsenti, E. (1995). Structure of growing microtubule ends: Two-dimensional sheets close into tubes at variable rates. J. Cell Biol. 129, 1311–1328. Conte, N., Delaval, B., Ginestier, C., Ferrand, A., Isnardon, D., Larroque, C., Prigent, C., Seraphin, B., Jacquemier, J., and Birnbaum, D. (2003). TACC1-chTOG-Aurora A protein complex in breast cancer. Oncogene 22, 8102–8116. Cullen, C. F., and Ohkura, H. (2001). Msps protein is localized to acentrosomal poles to ensure bipolarity of Drosophila meiotic spindles. Nat. Cell Biol. 3, 637–642. Cullen, C. F., Deak, P., Glover, D. M., and Ohkura, H. (1999). Mini spindles: A gene encoding a conserved microtubule-associated protein required for the integrity of the mitotic spindle in Drosophila. J. Cell Biol. 146, 1005–1018. Desai, A., and Mitchison, T. J. (1997). Microtubule polymerization dynamics. Annu. Rev. Cell. Dev. Biol. 13, 83–117. Desai, A., Verma, S., Mitchison, T. J., and Walczak, C. E. (1999). Kin I kinesins are microtubule-destabilizing enzymes. Cell 96, 69–78. Dionne, M. A., Sanchez, A., and Compton, D. A. (2000). ch-TOGp is required for microtubule aster formation in a mammalian mitotic extract. J. Biol. Chem. 275, 12346–12352. Doxsey, S. (2001). Re-evaluating centrosome function. Nat. Rev. Mol. Cell. Biol. 2, 688–698. Drechsel, D. N., Hyman, A. A., Cobb, M. H., and Kirschner, M. W. (1992). Modulation of the dynamic instability of tubulin assembly by the microtubule-associated protein tau. Mol. Biol. Cell 3, 1141–1154. Drewes, G., Ebneth, A., and Mandelkow, E. M. (1998). MAPs, MARKs and microtubule dynamics. Trends Biochem. Sci. 23, 307–311. Drewes, G., Trinczek, B., Illenberger, S., Biernat, J., Schmitt-Ulms, G., Meyer, H. E., Mandelkow, E. M., and Mandelkow, E. (1995). Microtubule-associated protein/microtubule aYnity-regulating kinase (p110mark). A novel protein kinase that regulates tau-microtubule interactions and dynamic instability by phosphorylation at the Alzheimer-specific site serine 262. J. Biol. Chem. 270, 7679–7688. Duensing, S., and Munger, K. (2001). Centrosome abnormalities, genomic instability and carcinogenic progression. Biochim. Biophys. Acta 2, M81–M88. Francis, S. E., and Davis, T. N. (2000). The spindle pole body of Saccharomyces cerevisiae: Architecture and assembly of the core components. Curr. Top. Dev. Biol. 49, 105–132. Galagan, J. E., Calvo, S. E., Borkovich, K. A., Selker, E. U., Read, N. D., et al. (2003). The genome sequence of the filamentous fungus Neurospora crassa. Nature 422, 859–868.


267

Garcia, M. A., Koonrugsa, N., and Toda, T. (2002). Spindle-kinetochore attachment requires the combined action of Kin I-like Klp5/6 and Alp14/Dis1-MAPs in fission yeast. EMBO J. 21, 6015–6024. Garcia, M. A., Vardy, L., Koonrugsa, N., and Toda, T. (2001). Fission yeast ch-TOG/ XMAP215 homologue Alp14 connects mitotic spindles with the kinetochore and is a component of the Mad2-dependent spindle checkpoint. EMBO J. 20, 3389–3401. Gard, D. L., and Kirschner, M. W. (1987). A microtubule-associated protein from Xenopus eggs that specifically promotes assembly at the plus-end. J. Cell Biol. 105, 2203–2215. Gard, D. L., Cha, B. J., and Schroeder, M. M. (1995). Confocal immunofluorescence microscopy of microtubules, microtubule-associated proteins, and microtubule-organizing centers during amphibian oogenesis and early development. Curr. Top. Dev. Biol. 31, 383–431. Gergely, F. (2002). Centrosomal TACCtics. Bioessays 24, 915–925. Gergely, F., Draviam, V. M., and RaV, J. W. (2003). The ch-TOG/XMAP215 protein is essential for spindle pole organization in human somatic cells. Genes Dev. 17, 336–341. Gergely, F., Karlsson, C., Still, I., Cowell, J., Kilmartin, J., and RaV, J. W. (2000a). The TACC domain identifies a family of centrosomal proteins that can interact with microtubules. Proc. Natl. Acad. Sci. USA 97, 14352–14357. Gergely, F., Kidd, D., JeVers, K., Wakefield, J. G., and RaV, J. W. (2000b). D-TACC: A novel centrosomal protein required for normal spindle function in the early Drosophila embryo. EMBO J. 19, 241–252. Giet, R., McLean, D., Descamps, S., Lee, M. J., RaV, J. W., Prigent, C., and Glover, D. M. (2002). ‘‘Drosophila aurora A kinase is required to localize D-TACC to centrosomes and to regulate astral microtubules. J. Cell Biol. 156, 437–451. Gliksman, N. R., Parsons, S. F., and Salmon, E. D. (1992). Okadaic acid induces interphase to mitotic-like microtubule dynamic instability by inactivating rescue. J. Cell Biol. 119, 1271–1276. Graf, R., Daunderer, C., and Schliwa, M. (2000). Dictyostelium DdCP224 is a microtubuleassociated protein and a permanent centrosomal resident involved in centrosome duplication. J. Cell Sci. 113, 1747–1758. Graf, R., Euteneuer, U., Ho, T. H., and Rehberg, M. (2003). Regulated expression of the centrosomal protein DdCP224 aVects microtubule dynamics and reveals mechanisms for the control of supernumerary centrosome number. Mol. Biol. Cell 14, 4067–4074. Groisman, I., Huang, Y. S., Mendez, R., Cao, Q., Theurkauf, W., and Richter, J. D. (2000). CPEB, maskin, and cyclin B1 mRNA at the mitotic apparatus: Implications for local translational control of cell division. Cell 103, 435–447. Groves, M. R., Hanlon, N., Turowski, P., Hemmings, B. A., and Barford, D. (1999). The structure of the protein phosphatase 2A PR65/A subunit reveals the conformation of its 15 tandemly repeated HEAT motifs. Cell 96, 99–110. Gruneberg, U., Campbell, K., Simpson, C., Grindlay, J., and Schiebel, E. (2000). Nud1p links astral microtubule organization and the control of exit from mitosis. EMBO J. 19, 6475–6488. Hasegawa, M., Morishima-Kawashima, M., Takio, K., Suzuki, M., Titani, K., and Ihara, Y. (1992). Protein sequence and mass spectrometric analyses of tau in the Alzheimer’s disease brain. J. Biol. Chem. 267, 17047–17054. Hayden, J. H., Bowser, S. S., and Rieder, C. L. (1990). Kinetochores capture astral microtubules during chromosome attachment to the mitotic spindle: Direct visualization in live newt lung cells. J. Cell Biol. 111, 1039–1045. Holmfeldt, P., Stenmark, S., and Gullberg, M. (2004). DiVerential functional interplay of TOGp/XMAP215 and the KinI kinesin MCAK during interphase and mitosis. EMBO J. 23, 627–637. Horio, T., and Hotani, H. (1986). Visualization of the dynamic instability of individual microtubules by dark-field microscopy. Nature 321, 605–607.

268

GARD ET AL.

Hoshi, M., Ohta, K., Gotoh, Y., Mori, A., Murofushi, H., Sakai, H., and Nishida, E. (1992). Mitogen-activated-protein-kinase-catalyzed phosphorylation of microtubule-associated proteins, microtubule-associated protein 2 and microtubule-associated protein 4, induces an alteration in their function. Eur. J. Biochem. 203, 43–52. Illenberger, S., Drewes, G., Trinczek, B., Biernat, J., Meyer, H. E., Olmsted, J. B., Mandelkow, E. M., and Mandelkow, E. (1996). Phosphorylation of microtubule-associated proteins MAP2 and MAP4 by the protein kinase p110mark. Phosphorylation sites and regulation of microtubule dynamics. J. Biol. Chem. 271, 10834–10843. Jameson, L., and Caplow, M. (1981). Modification of microtubule steady-state dynamics by phosphorylation of the microtubule-associated proteins. Proc. Natl. Acad. Sci. USA 78, 3413–3417. Joshi, H. C. (1998). Microtubule dynamics in living cells. Curr. Opin. Cell Biol. 10, 35–44. Katinka, M. D., Duprat, S., Cornillot, E., Metenier, G., Thomarat, F., Prensier, G., Barbe, V., et al. (2001). Genome sequence and gene compaction of the eukaryotic parasite Encephalitozoon cuniculi. Nature 414, 450–453. Kemphues, K. J., Wolf, N., Wood, W. B., and Hirsh, D. (1986). Two loci required for cytoplasmic organization in early embryos of Caenorhabditis elegans. Dev. Biol. 113, 449–460. Kinoshita, K., Arnal, I., Desai, A., Drechsel, D. N., and Hyman, A. A. (2001). Reconstitution of physiological microtubule dynamics using purified components. Science 294, 1340–1343. Kinoshita, N., Yamano, H., Le BouVant-Sladeczek, F., Kurooka, H., Ohkura, H., Stone, E. M., Takeuchi, M., Toda, T., Yoshida, T., and Yanagida, M. (1991). Sister-chromatid separation and protein dephosphorylation in mitosis. Cold Spring Harb. Symp. Quant. Biol. 56, 621–628. Kinoshita, K., Habermann, B., and Hyman, A. A. (2002). XMAP215: A key component of the dynamic microtubule cytoskeleton. Trends Cell Biol. 12, 267–273. Kline-Smith, S. L., and Walczak, C. E. (2002). The microtubule-destabilizing kinesin XKCM1 regulates microtubule dynamic instability in cells. Mol. Biol. Cell 13, 2718–2731. Knop, M., and Schiebel, E. (1998). Receptors determine the cellular localization of a gammatubulin complex and thereby the site of microtubule formation. EMBO J. 17, 3952–3967. Kobe, B., Gleichmann, T., Horne, J., Jennings, I. G., Scotney, P. D., and Teh, T. (1999). Turn up the HEAT. Structure Fold. Des. 7, R91–R97. Kosco, K. A., Pearson, C. G., Maddox, P. S., Wang, P. J., Adams, I. R., Salmon, E. D., Bloom, K., and HuVaker, T. C. (2001). Control of microtubule dynamics by Stu2p is essential for spindle orientation and metaphase chromosome alignment in yeast. Mol. Biol. Cell 12, 2870–2880. Kowalski, R. J., and Williams, R. C., Jr. (1993). Microtubule-associated protein 2 alters the dynamic properties of microtubule assembly and disassembly. J. Biol. Chem. 268, 9847–9855. Kramer, A., and Ho, A. D. (2001). Centrosome aberrations and cancer. Onkologie 24, 538–544. LauVart, B., Howell, S. J., Tasch, J. E., Cowell, J. K., and Still, I. H. (2002). Interaction of the transforming acidic coiled-coil 1 (TACC1) protein with ch-TOG and GAS41/NuBI1 suggests multiple TACC1-containing protein complexes in human cells. Biochem. J. 363, 195–200. Lawler, S. (1998). Microtubule dynamics: If you need a shrink try stathmin/Op18. Curr. Biol. 8, R212–R214. Le Bot, N., Tsai, M. C., Andrews, R. K., and Ahringer, J. (2003). TAC-1 a regulator of microtubule length in the C. elegans embryo. Curr. Biol. 13, 1499–1505. Lee, G., Cowan, N., and Kirschner, M. (1988). The primary structure and heterogeneity of tau protein from mouse brain. Science 239, 285–288. Lee, M. J., Gergely, F., JeVers, K., Peak-Chew, S. Y., and RaV, J. W. (2001). Msps/XMAP215 interacts with the centrosomal protein D-TACC to regulate microtubule behaviour. Nat. Cell Biol. 3, 643–649.


269

Lindwall, G., and Cole, R. D. (1984). Phosphorylation aVects the ability of tau protein to promote microtubule assembly. J. Biol. Chem. 259, 5301–5305. Littauer, U. Z., Giveon, D., Thierauf, M., Ginzburg, I., and Ponstingl, H. (1986). Common and distinct tubulin binding sites for microtubule-associated proteins. Proc. Natl. Acad. Sci. USA 83, 7162–7166. Mandelkow, E. M., Mandelkow, E., and Milligan, R. A. (1991). Microtubule dynamics and microtubule caps: A time-resolved cryo-electron microscopy study. J. Cell Biol. 114, 977–991. Matthews, L. R., Carter, P., Thierry-Mieg, D., and Kemphues, K. (1998). ZYG-9, a Caenorhabditis elegans protein required for microtubule organization and function, is a component of meiotic and mitotic spindle poles. J. Cell Biol. 141, 1159–1168. McNally, F. J., and Vale, R. D. (1993). Identification of katanin, an ATPase that severs and disassembles stable microtubules. Cell 75, 419–429. Mitchison, T., and Kirschner, M. (1984). Dynamic instability of microtubule growth. Nature 312, 237–242. Morgan, D. O. (1997). Cyclin-dependent kinases: Engines, clocks, and microprocessors. Annu Rev. Cell Dev. Biol. 13, 261–291. Moritz, M., and Agard, D. A. (2001). Gamma-tubulin complexes and microtubule nucleation. Curr. Opin. Struct. Biol. 11, 174–181. Nabeshima, K., Kurooka, H., Takeuchi, M., Kinoshita, K., Nakaseko, Y., and Yanagida, M. (1995). p93dis1, which is required for sister chromatid separation, is a novel microtubule and spindle pole body-associating protein phosphorylated at the Cdc2 target sites. Genes Dev. 9, 1572–1585. Nabeshima, K., Nakagawa, T., Straight, A. F., Murray, A., Chikashige, Y., Yamashita, Y. M., Hiraoka, Y., and Yanagida, M. (1998). Dynamics of centromeres during metaphaseanaphase transition in fission yeast: Dis1 is implicated in force balance in metaphase bipolar spindle. Mol. Biol. Cell 9, 3211–3225. Nagase, T., Miyajima, N., Tanaka, A., Sazuka, T., Seki, N., Sato, S., Tabata, S., Ishikawa, K., Kawarabayasi, Y., Kotani, H., and Nomura, N. (1995). Prediction of the coding sequences of unidentified human genes. III. The coding sequences of 40 new genes (KIAA0081KIAA0120) deduced by analysis of cDNA clones from human cell line KG-1. DNA Res. 2, 51–59. Nakaseko, Y., Goshima, G., Morishita, J., and Yanagida, M. (2001). M phase-specific kinetochore proteins in fission yeast: Microtubule-associating Dis1 and Mtc1 display rapid separation and segregation during anaphase. Curr. Biol. 11, 537–549. Nakaseko, Y., Nabeshima, K., Kinoshita, K., and Yanagida, M. (1996). Dissection of fission yeast microtubule associating protein p93Dis1: Regions implicated in regulated localization and microtubule interaction. Genes Cells 1, 633–644. Neuwald, A. F., and Hirano, T. (2000). HEAT repeats associated with condensins, cohesins, and other complexes involved in chromosome-related functions. Genome Res. 10, 1445–1452. Ohkura, H., Adachi, Y., Kinoshita, N., Niwa, O., Toda, T., and Yanagida, M. (1988). Coldsensitive and caVeine-supersensitive mutants of the Schizosaccharomyces pombe dis genes implicated in sister chromatid separation during mitosis. EMBO. J. 7, 1465–1473. Ohkura, H., Garcia, M. A., and Toda, T. (2001). Dis1/TOG universal microtubule adaptors— one MAP for all? J. Cell Sci. 114, 3805–3812. Parsons, S. F., and Salmon, E. D. (1997). Microtubule assembly in clarified Xenopus egg extracts. Cell. Motil. Cytoskeleton 36, 1–11. Pereira, G., Grueneberg, U., Knop, M., and Schiebel, E. (1999). Interaction of the yeast gamma-tubulin complex-binding protein Spc72p with Kar1p is essential for microtubule function during karyogamy. EMBO J. 18, 4180–4195.

270

GARD ET AL.

Pereira, G., Knop, M., and Schiebel, E. (1998). Spc98p directs the yeast gamma-tubulin complex into the nucleus and is subject to cell cycle-dependent phosphorylation on the nuclear side of the spindle pole body. Mol. Biol. Cell 9, 775–793. Pihan, G. A., Purohit, A., Wallace, J., Knecht, H., Woda, B., Quesenberry, P., and Doxsey, S. J. (1998). Centrosome defects and genetic instability in malignant tumors. Cancer Res. 58, 3974–3985. Pihan, G. A., Purohit, A., Wallace, J., Malhotra, R., Liotta, L., and Doxsey, S. J. (2001). Centrosome defects can account for cellular and genetic changes that characterize prostate cancer progression. Cancer Res. 61, 2212–2219. Popov, A. V., Pozniakovsky, A., Arnal, I., Antony, C., Ashford, A. J., Kinoshita, K., Tournebize, R., Hyman, A. A., and Karsenti, E. (2001). XMAP215 regulates microtubule dynamics through two distinct domains. EMBO J. 20, 397–410. Popov, A. V., Severin, F., and Karsenti, E. (2002). XMAP215 is required for the microtubulenucleating activity of centrosomes. Curr. Biol. 12, 1326–1330. Pryer, N. K., Walker, R. A., Skeen, V. P., Bourns, B. D., Soboeiro, M. F., and Salmon, E. D. (1992). Brain microtubule-associated proteins modulate microtubule dynamic instability in vitro. Real-time observations using video microscopy. J. Cell Sci. 103, 965–976. Quarmby, L. (2000). Cellular samurai: Katanin and the severing of microtubules. J. Cell Sci. 113, 2821–2827. RadcliVe, P., Hirata, D., Childs, D., Vardy, L., and Toda, T. (1998). Identification of novel temperature-sensitive lethal alleles in essential beta-tubulin and nonessential alpha 2-tubulin genes as fission yeast polarity mutants. Mol. Biol. Cell 9, 1757–1771. RaV, J. W. (2002). Centrosomes and cancer: Lessons from a TACC. Trends Cell Biol. 12, 222–225. Rehberg, M., and Graf, R. (2002). Dictyostelium EB1 is a genuine centrosomal component required for proper spindle formation. Mol. Biol. Cell. 13, 2301–2310. Rockmill, B., and Fogel, S. (1988). DIS1: A yeast gene required for proper meiotic chromosome disjunction. Genetics 119, 261–272. Rodionov, V. I., Gyoeva, F. K., Kashina, A. S., Kuznetsov, S. A., and Gelfand, V. I. (1990). Microtubule-associated proteins and microtubule-based translocators have diVerent binding sites on tubulin molecule. J. Biol. Chem. 265, 5702–5707. Roos, U. P. (1975). Fine structure of an organelle associated with the nucleus and cytoplasmic microtubules in the cellular slime mould Polysphondylium violaceum. J. Cell Sci. 18(2), 315–326. Rout, M. P., and Kilmartin, J. V. (1991). Yeast spindle pole body components. Cold Spring Harb. Symp. Quant. Biol. 56, 687–692. Sammak, P. J., and Borisy, G. G. (1988). Direct observation of microtubule dynamics in living cells. Nature 332, 724–726. Sato, M., Vardy, L., Angel Garcia, M., Koonrugsa, N., and Toda, T. (2004). Interdependency of fission yeast Alp14/TOG and coiled coil protein Alp7 in microtubule localization and bipolar spindle formation. Mol. Biol. Cell 15, 1609–1622. Saxton, W. M., Stemple, D. L., Leslie, R. J., Salmon, E. D., Zavortink, M., and McIntosh, J. R. (1984). Tubulin dynamics in cultured mammalian cells. J. Cell Biol. 99(6), 2175–2186. Schroer, T. A. (2001). Microtubules don and doV their caps: Dynamic attachments at plus and minus ends. Curr. Opin. Cell Biol. 13, 92–96. Schuyler, S. C., and Pellman, D. (2001). Microtubule ‘‘plus-end-tracking proteins’’: The end is just the beginning Cell 105, 421–424. Serrano, L., Avila, J., and Maccioni, R. B. (1984). Controlled proteolysis of tubulin by subtilisin: localization of the site for MAP2 interaction. Biochemistry 23, 4675–4681. Serrano, L., Montejo de Garcini, E., Hernandez, M. A., and Avila, J. (1985). Localization of the tubulin binding site for tau protein. Eur. J. Biochem. 153, 595–600.


271

Severin, F., Hebermann, B., HuVaker, T., and Hyman, T. (2001). Stu2 promotes mitotic spindle elongation in anaphase. J. Cell Biol. 153, 435–442. Shelden, E., and Wadsworth, P. (1993). Observation and quantification of individual microtubule behavior in vivo: Microtubule dynamics are cell-type specific. J. Cell Biol. 120, 935–945. Shiina, N., Moriguchi, T., Ohta, K., Gotoh, Y., and Nishida, S. (1992). Regulation of a major microtubule-associated protein by MPF and MAP kinase. EMBO J. 11, 3977–3984. Shirasu-Hiza, M., Coughlin, P., and Mitchison, T. (2003). Identification of XMAP215 as a microtubule-destabilizing factor in Xenopus egg extract by biochemical purification. J. Cell Biol. 161, 349–358. Simon, J. R., and Salmon, E. D. (1990). The structure of microtubule ends during the elongation and shortening phases of dynamic instability examined by negative-stain electron microscopy. J. Cell Sci. 96, 571–582. Snyder, M. (1994). The spindle pole body of yeast. Chromosoma 103, 369–380. Soues, S., and Adams, I. R. (1998). SPC72: A spindle pole component required for spindle orientation in the yeast Saccharomyces cerevisiae. J. Cell Sci. 111(Pt 18), 2809–2818. Spittle, C., Charrasse, S., Larroque, C., and Cassimeris, L. (2000). The interaction of TOGp with microtubules and tubulin. J. Biol. Chem. 275, 20748–20753. Srayko, M., Quintin, S., Schwager, A., and Hyman, A. A. (2003). Caenorhabditis elegans TAC-1 and ZYG-9 form a complex that is essential for long astral and spindle microtubules. Curr. Biol. 13, 1506–1511. Stebbins-Boaz, B., Cao, Q., de Moor, C. H., Mendez, R., and Richter, J. D. (1999). Maskin is a CPEB-associated factor that transiently interacts with elF-4E. Mol. Cell 4, 1017–1027. Still, I. H., Hamilton, M., Vince, P., Wolfman, A., and Cowell, J. K. (1999a). Cloning of TACC1, an embryonically expressed, potentially transforming coiled coil containing gene, from the 8p11 breast cancer amplicon. Oncogene 18, 4032–4038. Still, I. H., Vince, P., and Cowell, J. K. (1999b). The third member of the transforming acidic coiled coil-containing gene family, TACC3, maps in 4p16, close to translocation breakpoints in multiple myeloma, and is upregulated in various cancer cell lines. Genomics 58, 165–170. Theurkauf, W. E. (2001). TACCing down the spindle poles. Nat. Cell Biol. 3, E159–E161. Tournebize, R., Popov, A., Kinoshita, K., Ashford, A. J., Rybina, S., Pozniakovsky, A., Mayer, T. U., Walczak, C. E., Karsenti, E., and Hyman, A. A. (2000). Control of microtubule dynamics by the antagonistic activities of XMAP215 and XKCM1 in Xenopus egg extracts. Nat. Cell Biol. 2, 13–19. Tran, P. T., Walker, R. A., and Salmon, E. D. (1997). A metastable intermediate state of microtubule dynamic instability that diVers significantly between plus and minus ends. J. Cell Biol. 138, 105–117. Twell, D., Park, S. K., Hawkins, T. J., Schubert, D., Schmidt, R., Smertenko, A., and Hussey, P. J. (2002). MOR1/GEM1 has an essential role in the plant-specific cytokinetic phragmoplast. Nat. Cell Biol. 4, 711–714. Usui, T., Maekawa, H., Pereira, G., and Schiebel, E. (2003). The XMAP215 homologue Stu2 at yeast spindle pole bodies regulates microtubule dynamics and anchorage. EMBO J. 22, 4779–4793. van Breugel, M., Drechsel, D., and Hyman, A. (2003). Stu2p, the budding yeast member of the conserved Dis1/XMAP215 family of microtubule-associated proteins is a plus end-binding microtubule destabilizer. J. Cell Biol. 161, 359–369. Vasquez, R. J., Gard, D. L., and Cassimeris, L. (1994). XMAP from Xenopus eggs promotes rapid plus end assembly of microtubules and rapid microtubule polymer turnover. J. Cell Biol. 127, 985–993. Vasquez, R. J., Gard, D. L., and Cassimeris, L. (1999). Phosphorylation by CDK1 regulates XMAP215 function in vitro. Cell Motil. Cytoskeleton 43, 310–321.

272

GARD ET AL.

Vaughn, K. C., and Harper, J. D. (1998). Microtubule-organizing centers and nucleating sites in land plants. Int. Rev. Cytol. 181, 75–149. Verde, F., Labbe, J. C., Doree, M., and Karsenti, E. (1990). Regulation of microtubule dynamics by cdc2 protein kinase in cell-free extracts of Xenopus eggs. Nature 343, 233–238. Walczak, C. E., Mitchison, T. J., and Desai, A. (1996). XKCM1: A Xenopus kinesin-related protein that regulates microtubule dynamics during mitotic spindle assembly. Cell 84, 37–47. Walker, R. A., O’Brien, E. T., Pryer, N. K., Soboeiro, M. F., Voter, W. A., Erickson, H. P., and Salmon, E. D. (1988). Dynamic instability of individual microtubules analyzed by video light microscopy: Rate constants and transition frequencies. J. Cell Biol. 107, 1437–1448. Wang, D., Lewis, S. A., and Cowan, N. J. (1988). Complete sequence of a cDNA encoding mouse MAP2. Nucleic Acids Res. 16, 11369–11370. Wang, P. J., and HuVaker, T. C. (1997). Stu2p: A microtubule-binding protein that is an essential component of the yeast spindle pole body. J. Cell Biol. 139, 1271–1280. West, R. R., Tenbarge, K. M., and Olmsted, J. B. (1991). A model for microtubule-associated protein 4 structure. Domains defined by comparisons of human, mouse, and bovine sequences. J. Biol. Chem. 266, 21886–21896. Whittington, A. T., Vugrek, O., Wei, K. J., Hasenbein, N. G., Sugimoto, K., Rashbrooke, M. C., and Wasteneys, G. O. (2001). MOR1 is essential for organizing cortical microtubules in plants. Nature 411, 610–613. Wordeman, L., and Mitchison, T. J. (1995). Identification and partial characterization of mitotic centromere-associated kinesin, a kinesin-related protein that associates with centromeres during mitosis. J. Cell Biol. 128, 95–104. Yasuhara, H., Muraoka, M., Shogaki, H., Mori, H., and Sonobe, S. (2002). TMBP200, a microtubule bundling polypeptide isolated from telophase tobacco BY-2 cells is a MOR1 homologue. Plant Cell Physiol. 43, 595–603.

INDEX

A A. californica. See Aplysia californica A. stephensi. See Anopheles stephensi A. thaliana. See Arabidopsis thaliana ACE. See Angiotensin-converting enzyme Actinia equine, ACE with, 58 Adipokinetic hormone (AKH-1), ECE with, 77 AKH-1. See Adipokinetic hormone Allium cepa, perichromosomal sheath in, 133 Alp14 fungi with, 194–198, 201–203, 205–206 MAP215/Dis1 family with, 179, 182, 183, 189, 193, 194–198, 201–203, 205–206, 247–251 reconciling functions of, 247–251 yeast with, 201–203, 205–206 Alzheimer’s disease, NEP with, 73, 83–84 Amphibians environmental estrogens influencing, 28–31 oogenesis in, 141–142 reproductive strategies with, 10–16 VTG in, 6–8, 28–31 AngI. See Angiotension I AngII. See Angiotension II Angiotensin-converting enzyme (ACE) ACE2 and, 52–55 Alzheimer’s disease with, 83–84 biological role of, 51–53 blood pressure regulation with, 51–52 concluding remarks on, 84–85 invertebrates with, 55–67 ACE biological role in, 61–67 ACE sequence conservation in, 57 ACE structure in, 56–58

273

ACE substrate specificity in, 59–61 central nervous system in, 64–65 defense system in, 65–67 development in, 63–64 evolutionary aspects of, 58–59 leeches in, 67 metamorphosis in, 63–64 prohormone processing in, 64–65 reproduction in, 62–63 isoforms of, 49–51 mammals with, 48–55 other roles for, 81–84 prokaryotes with, 81–83 RAS with, 48, 49, 51, 52, 55, 56, 60, 61, 67 structure of, 49–51 Angiotension I (AngI), 48, 51, 53, 54, 82 Angiotension II (AngII), 48, 49, 51, 53, 54, 55, 82 Anopheles gambiae, ACE with, 58 Anopheles stephensi, ACE with, 56, 62, 64 Aplysia californica, ECE with, 77, 79, 81 Arabidopsis thaliana, MAP215/Dis1 family in, 179, 183, 185, 186, 187, 211, 212 Artemia salina, nucleologenesis with, 142 Ascomycotes, MAP215/Dis1 family with, 195–196, 200 Atractapsis engaddensis, ECE with, 69

B Basidiomycotes, MAP215/Dis1 family in, 184, 206–208 Bombyx mori ACE with, 56, 63, 64 ECE with, 79, 81

274

INDEX

Boophilus microplus, ACE with, 56, 57 Bradykinin ACE with, 48, 51, 53, 54, 59–60, 61 ECE with, 71, 75, 76 Bradysia mycorum, perichromosomal sheath in, 133 Brugia malayi, MAP215/Dis1 family in, 189, 211

C C. cineria. See Coprinopsis cineria C. elegans. See Caenortibditis elegans C. intestinalis. See Ciona intestinalis C. morosus. See Carausius morosus C. neoformans. See Cryptococcus neoformans C. picta. See Chrysemys picta C. reinhardtii. See Chlamydamonas reinhardtii C. savignyi. See Ciona savignyi Caenortibditis elegans ACE with, 58, 64 ECE with, 78–80 MAP215/Dis1 family in, 179, 182, 183, 185, 186, 189, 216–220, 240–243, 245, 249, 252, 255 Cancer MAP215/Dis1 family’s role in, 245–247 mitotic NOR in, 104 NEP and, 73 Carassius auratus, vitellogenesis with, 18 Carausius morosus, ACE with, 64, 65 CAT D. See Cathepsin D Cathepsin D (CAT D), vitellogenesis with, 22–23, 24, 25, 27 CCK-8, ACE with, 53 CENP. See Centromeric proteins Centromeric proteins (CENP), 132 Centrosomes, MAP215/Dis1 family in, 239–247 ch-TOG. See Colonic-hepatic Tumor Overexpressed Gene ChIP. See Chromatin immunoprecipitations Chlamydamonas, MAP215/Dis1 family in, 190, 191 Chlamydamonas reinhardtii, MAP215/Dis1 family in, 189, 211 Chordates, XMAP215 in, 221–239 C-terminal domain with, 233, 235–236

CDK1-dependent phosphorylation regulation of, 236–237 centrosome targeting of, 235–236 microtubule assembly promoted by, 225–229 multiple MT binding domains with, 232–235 spindles organized with, 190, 210, 223, 229–232 structure/function correlation with, 237–239 structure of, 222–225 Chortophaga viridifasciata, perichromosomal sheath in, 133 Chromatin immunoprecipitations (ChIP), 112–113 Chrysemys picta, vitellogenesis with, 18, 19 Chrysomya bezziana, ACE with, 66 Ciona intestinalis, MAP215/Dis1 family in, 187, 213, 222, 236 Ciona savignyi, MAP215/Dis1 family in, 213, 222, 236 Colonic-hepatic Tumor Overexpressed Gene (ch-TOG) chordates with, 221–239 MAP215/Dis1 family with, 182–183, 189–191, 197, 207–208, 221–239, 240, 242, 243, 244, 245, 247–254 multiple MT binding domains with, 232–235 spindles organized with, 190, 210, 223, 229–232 structure/function correlation with, 237–239 structure of, 222–225 Coprinopsis cineria, MAP215/Dis1 family in, 206–207 Cryptococcus neoformans, MAP215/Dis1 family in, 206–207 Cyprinus carpio, vitellogenesis with, 31

D DdCP224 D. discoideum, 179, 182, 183, 213, 214–216, 218, 253 MAP215/Dis1 family with, 179, 182, 183, 185, 191, 192, 213–216, 218, 247, 251, 253, 254 reconciling functions of, 251, 253, 254

275

INDEX Dense fibrillar component (DFC), nucleolar assembly/disassembly with, 104, 110–111, 113–119, 122, 123, 124, 125, 128–131, 134, 135, 136, 139–141, 144, 152 DFC. See Dense fibrillar component Dicentrarchus labrax, vitellogenesis with, 25 Dictyostelium discoideum, MAP215/Dis1 family in, 182, 183, 187, 189, 191, 213–216, 247 Dis1. See MAP215/Dis1 family Drosophila ectopic nucleoli in, 144–145 embryogenesis in, 145–148 MAP215/Dis1 family in, 179, 182–183, 190, 191, 207, 209, 217, 220, 221, 229–231, 240–242, 245, 252, 254, 255 Msps with, 179, 182, 183, 186, 189, 191, 207, 220, 221, 242, 245, 252, 254 nucleolar assembly/disassembly in, 111, 114, 117, 123, 144–148 nucleologenesis model with, 144–145 Drosophila melanogaster ACE protein with, 50, 56, 79 MAP215/Dis1 family in, 183, 190 nucleolus-like bodies in, 123

E E. cuniculi. See Encephilitazooan cuniculi E. histolytica. See Entamoeba histolytica ED. See Endocrine disrupters Encephilitazooan cuniculi, MAP215/Dis1 family in, 185, 186, 187, 188, 189, 190, 191, 192, 216, 236, 255 Endocrine disrupters (ED), environmental estrogens as, 25–28, 32 Endothelin-converting enzymes (ECE) Alzheimer’s disease with, 83–84 biological role of, 69–71 concluding remarks on, 84–85 EC2 and, 69, 71 EC3 and, 69, 71 introduction to, 67–68 invertebrates with, 76–81 C. elegans, 78–80 endopeptidase activity in, 77–78 Hydra vulgaris, 78–80 insect, 80–81 NEP in, 81 isoforms of, 68–69

Kell blood group with, 68, 73–74 mammals with, 67–81 neutral endopeptidase with, 68, 72–73 other roles for, 81–84 PEX with, 68, 74–75 prokaryotes with, 81–83 secreted endopeptidase with, 68, 75 structure of, 68–69 XCE with, 68, 75–76 Endothelin (ET) ECE with, 69–81, 82 pathway of, 70, 71 Enkephalin ACE with, 53, 54, 55–56 ECE with, 73, 76, 77 Entamoeba histolytica, MAP215/Dis1 family in, 185, 186, 187, 188, 189, 190, 192, 194, 255 Environmental estrogen, feminization process from, 25–32 ER. See Estrogen receptor ERE. See Estrogen-responsive element Erpobdella octoculata, ACE with, 67 Estradiol-17b feminization process with, 26, 29, 31, 32 hypothalamus-hypophysial gonadal axis in, 3 reproductive strategies and, 11–13, 15, 16 seasonal reproduction and, 11–13 vitellogenesis with, 1, 3, 5–6, 7, 11–13, 15–20, 26, 29, 31–33 VTG synthesis and, 17–20 Estrogen receptor (ER), vitellogenesis with, 12, 15, 18, 26, 28, 29, 31–32 Estrogen-responsive element (ERE), vitellogenesis with, 5, 15, 29 ETS. See External transcribed spacer Euglena gracilis, nucleologenesis with, 142 Expressed sequence tag (EST), MAP215/Dis1 family with, 179, 183, 211, 222 External transcribed spacer (ETS), nucleolar assembly/disassembly in, 109, 114, 115, 130, 135, 137, 139, 141, 145, 149, 150, 159

F F. heteroclitus. See Fundulus heteroclitus Feminization process environmental estrogen in, 25–32 VTG as biomarker for, 25–32

276

INDEX

Feminization process (continued ) animal models of, 28–31 wild populations and, 31–32 Fish environmental estrogens influencing, 28–31 reproductive strategies with, 10–16 VTG in, 6–8, 28–31 Fundulus heteroclitus, vitellogenesis with, 5, 10, 22 Fungi ascomycotes, 195–196, 200 basidiomycote, 184, 206–208 functional analysis of yeast, 186, 196–201 MAP215/Dis1 family in, 186, 194–208 structure/function of yeast, 201–206

G G. lamblia. See Giardia lamblia G. max. See Glycine max G. niger. See Gobius niger Gem1 MAP215/Dis1 family with, 179, 183, 186, 189, 208–213, 222, 248 plants with, 208–213 Giardia lamblia, MAP215/Dis1 family in, 179, 185, 186, 187, 188, 192, 255 Gloeobacter violaceus, prokaryotes with, 81 Glycine max, MAP215/Dis1 family in, 190, 211 Gobius niger, vitellogenesis with, 18, 27

H H. vulgaris. See Hydra vulgaris Haematobia irritans exigua, ACE with, 56, 62 Haemoregulatory peptide, ACE with, 51, 53 HIV Rev protein, 117, 136 Hydra vulgaris, ECE with, 78–80, 79

I Insects ECE with, 80–81 MAP215/Dis1 family in, 219–221 Internal transcribed spacer (ITS), nucleolar assembly/disassembly in, 109, 114, 130, 135, 137, 145, 149, 150

Invertebrates ACE in, 55–67 biological role in, 61–67 central nervous system with, 64–65 defense system with, 65–67 development with, 63–64 evolutionary aspects of, 58–59 leeches with, 67 metamorphosis and, 63–64 prohormone processing of, 64–65 reproduction with, 62–63 sequence conservation in, 57 structure of, 56–58 substrate specificity of, 59–61 ECE in, 76–81 C. elegans, 78–80 endopeptidase activity of, 77–78 Hydra vulgaris, 78–80 insect, 80–81 NEP in, 81 ITS. See Internal transcribed spacer

K Kell blood group, ECE with, 68, 73–74

L L. decemlineata. See Leptinotarsa decemlineata L. dispar. See Lymantria dispar L. oleracea. See Lacanobia oleracea Lacanobia aleracea, ACE with, 63 Lacanobia oleracea, ACE with, 57, 64 Lampetra fluviatilis, reproductive strategies with, 12 Leeches, ACE with, 67 Leptinotarsa decemlineata, ACE with, 62, 64, 65 Leucophaea maderae ACE with, 57, 64, 65 ECE with, 77 Locust migratoria ACE with, 56, 62, 64, 65, 66 ECE with, 77, 78, 79, 80 Luteinizing hormone releasing hormone (LH-RH), ACE with, 53, 60 Lymantria dispar ACE with, 62 ECE with, 77

277

INDEX

M M13 EC2 and, 69, 71 EC3 and, 69, 71 invertebrates with, 76–81 C. elegans, 78–80 endopeptidase activity in, 77–78 Hydra vulgaris, 78–80 insect, 80–81 NEP in, 81 Kell blood group with, 68, 73–74 neutral endopeptidase with, 68, 72–73 PEX with, 68, 74–75 secreted endopeptidase with, 68, 75 XCE with, 68, 75–76 M. brassica. See Mamestra brassica M. domestica. See Musca domestica Mamestra brassica, ACE with, 64 Manduca sexta, ECE with, 79 MAP215/Dis1 family animals with, 213–247 centrosomes in, 239–247 D. discoideum, 214–216 DdCP224 in, 214–216 genome stability in, 245–247 insect, 219–221 Msps related protein in, 219–221 nematode, 216–219 other development in, 243–245, 246 TACC proteins in, 239–243 Zyg9-related proteins in, 216–219 cancer role with, 245–247 ch-TOG related proteins in, 221–239 multiple MT binding domains with, 232–235 spindles organized with, 190, 210, 223, 229–232 structure/function correlation with, 237–239 structure of, 222–225 chordates with, 221–239 common structural organization of, 185–192 cyclin-dependent kinases targets in, 187–189 N-terminal HEAT repeats in, 185–187, 188 single Tau repeat in, 189–192 TOG domains in, 185–187, 188 conclusion to, 254–255

eukaryote tree of life with, 183–185 eukaryotes with, 179–255 fungi with, 186, 194–208 ascomycotes, 195–196, 200 basidiomycote, 184, 206–208 functional analysis of yeast, 186, 196–201 structure/function of yeast, 201–206 introduction to, 180–192 MT loading zones in, 254 MTOC associated with, 250–252 MTOC function with, 252–254 nucleation/stabilization/anchoring of MTs in, 252–254 peripheral functions of, 249–250 plants with, 208–213 protists with, 184, 186, 190, 192–194 reconciling functions of, 238, 247–254 XMAP215 in, 221–239 C-terminal domain with, 233, 235–236 CDK1-dependent phosphorylation regulation of, 236–237 centrosome targeting of, 233, 235–236 microtubule assembly promoted by, 225–229 multiple MT binding domains with, 232–235 spindles organized with, 190, 210, 223, 229–232 structure/function correlation with, 237–239 structure of, 222–225 Medicago truncatula, MAP215/Dis1 family in, 190, 211 Microtubule-associated proteins (MAP), 179–181. See also MAP215/Dis1 family Microtubule-organizing centers (MTOC) evolutionarily ancient association of, 250–252 MAP215/Dis1 family with, 179, 191, 193, 196, 206, 208, 213, 219, 228, 238, 239, 248–254 MT loading zones with, 254 nucleation/stabilization/anchoring of MTs with, 252–254 Microtubules (MT) MAP215/Dis1 family with, 179–255 animals and, 213–247 ascomycotes and, 195–196, 200 basidiomycotes and, 184, 206–208 fungi and, 186, 194–208

278

INDEX

Microtubules (MT) (continued ) insects and, 219–221 introduction to, 180–192 nematodes and, 216–219 plants and, 208–213 protists and, 184, 186, 190, 192–194 yeast and, 186, 196–206 nucleation/stabilization/anchoring of, 252–254 Minispindles (Msps) Drosophila with, 179, 182, 183, 186, 189, 191, 207, 220, 221, 242, 245, 252, 254 insects with, 219–221 introduction to, 179, 182, 183, 186, 189, 191–192 MAP215/Dis1 family, 179, 182, 183, 186, 189, 191–192, 207, 209, 213, 218, 219–221, 222, 229–231, 241, 242, 245, 251, 252, 254 Mor1 A. thaliana with, 179, 183, 185, 186 MAP215/Dis1 family with, 179, 183, 185, 186, 189, 193, 207, 208–213, 222, 248, 249, 250, 251 plants with, 208–213 Msps. See Minispindles MT. See Microtubules MTOC. See Microtubule-organizing centers Musca domestica ACE with, 55, 56, 57, 58, 60 ECE with, 77

N N. tabaccum. See Nicotiana tabaccum Naegleria gruberi, nucleologenesis with, 142 Nematodes, MAP215/Dis1 family in, 216–219 NEP. See Neutral endopeptidase NES. See Nuclear export signals Neutral endopeptidase (NEP) Alzheimer’s disease with, 73, 83–84 cancer with, 73 ECE with, 67, 72–73, 74, 75, 76, 77, 78, 79, 80, 81, 82, 83, 84, 85 Nicotiana tabaccum, MAP215/Dis1 family in, 179, 183, 187, 249 NOR. See Nucleolar organizers Nuclear export signals (NES), nucleolar assembly/disassembly with, 118–119

Nucleolar assembly/disassembly disassembly at mitosis in, 153–158 persistent nucleoli with, 156–157 Pol I transcription reactivation with, 157–158 pre-RNA processing component phosphorylation with, 155–156 transcription factors phosphorylation with, 153–155 embryonic nucleologenesis with, 145–153 Drosophila in, 145–148 mammals in, 151–153 Xenopus in, 148–151 interphase nucleolus with, 106, 110–121 DFC in, 113–118 early ribosome assembly in, 113–118 fibrillar center in, 110–111 granular component in, 118–119 nuclear export in, 118–119 nucleolar multitasking in, 119–121 pre-rDNA processing in, 113–118 rDNA transcription in, 111–113 traditional nuclear function and, 106, 110–119 introduction to, 100–102 nucleolar organizers with, 102–110 composition of, 104–110 early pioneers of, 102–104 modern studies of, 104–110 molecular structure of, 104–110 SAT-chromosomes and, 102–103 UBF in, 104–110 nucleologenesis concurrent with NOR transcription in, 121–145 amphibian oogenesis model and, 141–142 closed mitosis model and, 140–141 Drosophila ectopic nucleoli model and, 144–145 late anaphase/early telophase reinitiation of, 121–123 NDF in, 128–131, 136–138 nucleolar assembly in, 126–140 nucleologenesis and, 126 nucleolus-like body formation in, 123–126 partially processed pre-rRNA in, 137–138 perichromosomal region and, 127–133 Physarum model and, 140–141

279

INDEX PNB/NDF dynamics in, 138 PNBs in, 133–136, 138 postmitotic mammalian nucleologenesis model and, 138–140 yeast nucleolar assembly model and, 142–144 perspectives on, 158 Nucleolar organizers (NOR) composition of, 104–110 early pioneers of, 102–104 mitosis with, 104–110, 111, 113, 121, 127, 154–155 modern studies of, 104–110 molecular structure of, 104–110 nucleolar assembly/disassembly, 100, 102–110, 111, 113, 121–145, 148–149, 151, 153, 154, 155, 159 nucleolar perspectives with, 159 SAT-chromosomes and, 102–103 three-dimensional model of, 106 transcription with, 121–145 amphibian oogenesis model and, 141–142 closed mitosis model and, 140–141 Drosophila ectopic nucleoli model and, 144–145 late anaphase/early telophase reinitiation of, 121–123 NDF in, 128–131, 136–138 nucleolar assembly in, 126–140 nucleologenesis and, 126 nucleolus-like body formation in, 123–126 partially processed pre-rRNA in, 137–138 perichromosomal region and, 127–133 Physarum model and, 140–141 PNB/NDF dynamics in, 138 PNBs in, 133–136, 138 postmitotic mammalian nucleologenesis model and, 138–140 yeast nucleolar assembly model and, 142–144 UBF in, 104–110, 121–122, 154–155 Nucleologenesis amphibian oogenesis model and, 141–142 closed mitosis model and, 140–141 Drosophila ectopic nucleoli model and, 144–145 embryos with, 145–153 Drosophila in, 145–148

mammals in, 151–153 Xenopus in, 148–151 late anaphase/early telophase reinitiation of, 121–123 NDF in, 128–131, 136–138 nucleolar assembly/disassembly with, 99–101, 103, 110, 111, 113, 121–153, 156, 158–159 nucleolar assembly in, 126–140 nucleolus-like body formation in, 123–126 partially processed pre-rRNA in, 137–138 perichromosomal region and, 127–133 perspectives on, 158–159 Physarum model and, 140–141 PNB/NDF dynamics in, 138 PNBs in, 133–136, 138 postmitotic mammalian nucleologenesis model and, 138–140 primer on, 126 yeast nucleolar assembly model and, 142–144

O Oncorynchus mykiss, vitellogenesis with, 18, 21, 22 Oreochromis inlohins, vitellogenesis with, 19 Oreochromis niloticus, vitellogenesis with, 21

P P. chrysosporium. See Phanerochaete chrysosporium P. gingivalis. See Porphyromonas gingivalis P. patens. See Physcomitrella patens Perichromosomal sheath introduction to, 99–101 NDF with, 136–138 nucleolar assembly/disassembly with, 99–101, 104, 126, 127–133, 136, 137, 138, 139, 158–159 nucleolar component distribution with, 128–131 nucleologenesis with, 126, 127–133, 136–139 perspectives on, 158–159 region of, 127–133 PEX. See Phosphate regulating gene Phanerochaete chrysosporium, MAP215/Dis1 family in, 187, 206–207

280

INDEX

Phosphate regulating gene (PEX), ECE with, 68, 74–75 Physarum polycephalum, nucleologenesis with, 140–141 Physcomitrella patens, MAP215/Dis1 family in, 189, 211 Pimpla hypochondriaca, ACE with, 66 Pituitary hormones reproductive strategies with, 9, 11, 15–16 vitellogenesis with, 1, 9, 11, 15–16, 18, 31, 33 Plants, MAP215/Dis1 family in, 208–213 Poecilia reticulata, vitellogenesis with, 32 Porphyromonas gingivalis, prokaryotes with, 82 Prenucleolar bodies (PNB) composition of, 134–135 in depth look at, 133–136 dynamism of, 135–136 early work on, 133–134 introduction to, 99–100 NDF dynamics with, 136, 138 nucleolar assembly/disassembly with, 99–100, 121–125, 126, 127, 128–131, 133–136–140, 141, 149–150, 153, 156, 158–159 nucleolar component distribution with, 128–131 nucleologenesis with, 121–125, 126, 127, 128–131, 133–141, 139 perspectives on, 158–159 Prohormone processing, ACE with, 64–65 Prokaryotes, ECE/ACE with, 81–83 Protease, five major groups of, 47–48 Protists, MAP215/Dis1 family in, 184, 186, 190, 192–194

R R. erinacea. See Raja erinacea R. esculenta. See Rana esculenta Raja erinacea, reproductive strategies with, 12, 19 Rana esculenta reproductive strategies with, 13, 14 vitellogenesis with, 13, 14, 18, 30 Renin-angiotensin system (RAS) ACE in, 48, 49, 51, 52, 55, 56, 60, 61, 67 biological role of, 51, 52 blood pressure regulation with, 48, 51, 56

components of, 49 leeches with, 60, 67 Reproductive strategies, vitellogenesis with, 8–17 amniotic egg and, 16–17 amphibian models of, 10–16 fish models of, 10–16 habitat acquirement in, 16–17 hypothalamus-hypophysial gonadal axis in, 2 seasonal patterns in, 10–16 Sparus aurata in, 11, 12, 15 Reptiles, VTG in, 6–8 Rutilius rutilus, vitellogenesis with, 32

S S. cerevisiae. See Saccharomyces cerevisiae S. pombe. See Schizosaccaromyces pombe Saccharomyces cerevisiae nucleologenesis with, 142 Stu2 in, 179, 182, 183, 185, 189, 195, 198, 200, 202–203, 205, 206, 208, 249, 251, 254 SARS. See Severe Acute Respiratory Syndrome SAT-chromosomes (Sine acido thymonucleinico), NOR with, 102–103 Schistocerca gregaria, ECE with, 77 Schizosaccaromyces pombe, MAP215/Dis1 family in, 179, 181–183, 186, 189, 192–196, 198, 205, 207, 249, 251 Scyliorhinus canicula, reproductive strategies with, 12 Secreted endopeptidase (SEP), ECE with, 68, 75 Severe Acute Respiratory Syndrome (SARS), ACE2 with, 55 Shewanella oneidensis, prokaryotes with, 81 Sine acido thymonucleinico. See SAT-chromosomes Sparus aurata, reproductive strategies with, 11, 12, 15 Sphenodon punctatus, vitellogenesis with, 7 Spisula solidissima, nucleologenesis with, 142 Stu2 fungi with, 195–196, 198–206, 208 MAP215/Dis1 family with, 179, 182, 183, 185, 189, 191, 195–196, 198–206, 208, 231, 247–251, 254 reconciling functions of, 247–251, 254

281

INDEX S. cerevisiae with, 179, 182, 183, 185, 189, 195, 198, 200, 202–203, 205, 206, 208, 249, 251, 254 Substance P ACE with, 52, 53, 60, 61, 82 ECE with, 71, 73, 75, 82

rDNA transcription with, 111–113, 121–122 Ustilago maydis, MAP215/Dis1 family in, 186, 187, 189, 190, 191, 195, 205, 206, 216, 218, 219, 221, 236

V T T. tessalutum. See Theromyzon tessalutum TACC proteins. See Transforming Acidic Coiled-Coil proteins TATA-binding protein (TBP), nucleolar assembly/disassembly with, 107, 111, 122, 148, 151, 154, 155 TE. See Testis ecdysiotropin Testis ecdysiotropin (TE), ACE with, 62 Theromyzon tessalutum, ACE with, 56, 67 TOG domains, MAP215/Dis1 family structure with, 183, 184, 185–187, 188, 192–194, 196–197, 200, 201–203, 207, 211–213, 215, 218–219, 221–225, 232, 235 Torpedo marmorata ACE with, 58 reproductive strategies with, 12 Transforming Acidic Coiled-Coil (TACC) proteins centrosomes targeted by, 239–247 MAP215/Dis1 family with, 197, 204, 206, 217, 220–221, 236, 237, 238, 239–247, 252, 254 MTOC and, 254 Triturus carnifex, vitellogenesis with, 16, 30

U U. maydis. See Ustilago maydis Upstream binding factor (UBF) interphase nucleoli with, 121–122 mitotic NOR with, 104–110, 121–122, 154–155 mitotic phosphorylation with, 154–155 nucleolar perspectives with, 159 nucleologenesis with, 121–122, 125–126, 128, 141, 148, 150–151, 153 Pol I transcription with, 157–158

Verasper moseri, vitellogenesis with, 22 Vertebrates, vitellogenesis in, 1–34 amphibian, 6–8, 28–31 fish, 6–8, 28–31 hormone regulation with, 17–20 reproductive strategies with, 8–17 reptile, 6–8 utilization of, 20–25 Vicia faba, perichromosomal sheath in, 133 Vitellogenesis concluding remarks on, 32–34 history of, 3–8 VTG in, 4–6 Xenopus laevis in, 3, 5 hormone regulation for, 17–20 introduction to, 1–3 multihormonal control of, 1–34 reproductive strategies with, 8–17 amniotic egg and, 16–17 amphibian models of, 10–16 fish models of, 10–16 habitat acquirement in, 16–17 hypothalamus-hypophysial gonadal axis in, 2 seasonal patterns in, 10–16 Sparus aurata in, 11, 12, 15 VTG utilization and, 20–25 Vitellogenin (VTG) amphibians with, 6–8, 28–31 biochemical characterization of, 6–8 feminization process marked by, 25–32 animal models of, 28–31 wild populations and, 31–32 fish with, 6–8, 28–31 history of, 4–6 hormone regulation with, 17–20 reproductive strategies with, 8–17 reptiles with, 6–8 structure of, 5 utilization of, 20–25 VTG. See Vitellogenin

282

INDEX

X X. laevis. See Xenopus laevis X-converting enzyme (XCE), ECE with, 68, 75–76 Xanthomonas axonopodis, prokaryotes with, 82 XCE. See X-converting enzyme Xenoestrogen reproductive effects of, 29 vitellogenesis and, 1, 7, 29, 32 Xenopus embryogenesis in, 148–151 MAP215/Dis1 family in, 179, 181–182, 186, 188, 190, 192, 193, 199, 203, 209, 220, 221, 222, 223, 226, 228–232, 234, 236–237, 240, 241, 243, 245, 246, 249, 252 nucleolar assembly/disassembly in, 111, 112, 118, 121, 123–125, 134–135, 141, 148–151 XMAP215 in, 179, 181, 182, 186, 192–194, 199, 207, 220, 222, 228, 231, 232, 234, 241 Xenopus laevis embryogenesis in, 148 MAP215/Dis1 family in, 181, 186, 190, 241, 243, 245, 246, 249, 252 nucleolus-like bodies in, 123–124 vitellogenesis with, 3, 5, 8, 15, 20, 30 Xenopus tropicalis, MAP215/Dis1 family in, 190, 222 XMAP215 animals with, 213, 218, 220, 221–239, 241–245, 246, 247 C-terminal domain with, 233, 235–236 CDK1-dependent phosphorylation regulation of, 236–237 centrosome targeting of, 233, 235–236 chordates with, 221–239

introduction to, 179, 181–183 MAP215/Dis1 family with, 179, 181–183, 186, 187–188, 192–194, 199, 203, 207–209, 212, 213, 218, 220, 221–239, 241–245, 246, 247, 248–253 microtubule assembly promoted by, 225–229 multiple MT binding domains with, 232–235 protists with, 192–194 reconciling functions of, 248–253 spindles organized with, 190, 210, 223, 229–232 structure/function correlation with, 237–239 structure of, 222–225 tubulin assembly with, 181 Xenopus with, 179, 181, 182, 186, 192–194, 199, 207, 220, 222, 228, 231, 232, 234, 241

Y Yeast, MAP215/Dis1 family in functional analysis of, 186, 196–201 structure/function of, 201–206

Z Zyg9 C. elegans with, 179, 182, 183, 185, 186, 189, 216–219, 241, 242–243, 245, 249, 252 MAP215/Dis1 family with, 179, 182, 183, 185, 186, 188, 189, 213, 216–219, 222, 241, 242–243, 245, 249, 251, 252 nematodes with, 216–219

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