International Review of Cytology, Vol. 198 [1st ed.] 0123646022, 9780123646026, 9780080548630

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International Review of

A Survey of

Cytology Cell Biology VOLUME 198

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 Rosa Beddington Howard A. Bern Robert A. Bloodgood Dean Bok Rene Couteaux Marie A. DiBerardino Laurence Etkin Hiroo Fukuda Elizabeth D. Hay P. Mark Hogarth William R. Jeffrey Anthony P. Mahowald

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

International Review of A Survey of

Cytology Cell Biology

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

VOLUME 198

ACADEMIC PRESS San Diego London Boston New York Sydney Tokyo Toronto

Acaedemic Press is an imprint of Elsevier, Inc. 84 Theobald’s Road, London WC1X 8RR, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands Linacre House, Jordan Hill, Oxford OX2 8DP, UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA Copyright © 2002 Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material

International Review of Cytology: A Survey of Cell Biology, Volume 198 Edited by Kwang W. Jeon ISBN 978-0-12-364602-6

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CONTENTS

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

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Somite Formation and Patterning Estelle Hirsinger, Caroline Jouve, Julien Dubrulle, and Oliver Pourquie´ I. II. III. IV. V. VI. VII.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Origin and Commitment of the Somitic Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Somite Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antero-posterior Compartmentalization of the Somites . . . . . . . . . . . . . . . . . . . . . . . . . . Dorso-ventral Patterning of the Avian Somite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Medio-lateral Polarity of the Avian Somite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 5 8 14 18 37 47 49

Morphogenesis of the Eggshell in Drosophila Gail L. Waring I. II. III. IV. V. VI. VII.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Morphology of the Eggshell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Identification of Eggshell Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eggshell Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Posttranslational Processing of Eggshell Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v

67 69 73 76 85 89 102 104

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Cell Biology of the Cytochrome P-450 in the Liver Shinsuke Kanamura and Jun Watanabe I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulatory Mechanisms for P-450 Expression in Mammalian Hepatocytes . . . . . . . . . . . Membrane Topology of P-450 Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sublobular Expression of P-450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

109 110 126 134 140 140

Alternative Protein Sorting Pathways John Kim, Sidney V. Scott, and Daniel J. Klionsky I. II. III. IV. V. VI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Retrograde Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Degradative Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nondegradative Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Channel-Mediated Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

153 154 165 179 188 191 191

Structural Correlates of the Transepithelial Water Transport Ekaterina S. Snigirevskaya and Yan Yu. Komissarchik I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Features of Epithelia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Role of Tight Junctions in Transepithelial Transport . . . . . . . . . . . . . . . . . . . . . . . . . Ultrastructure of Apical Membranes of Epitheliocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . Participation of the Vacuolar System in the Cell Response on Induced Water Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. The Role of the Cytoskeleton in Transcellular Water Transport . . . . . . . . . . . . . . . . . . . . VII. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

203 205 215 224 240 250 255 256

Membrane Trafficking and Processing in Paramecium Richard D. Allen and Agnes K. Fok I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Membrane Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

277 279

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III. Membrane Modulation and Membrane Transport Systems . . . . . . . . . . . . . . . . . . . . . . . IV. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

304 311 312

Role of Natural Benzoxazinones in the Survival Strategy of Plants Dieter Sicker, Monika Frey, Margot Schulz, and Alfons Gierl I. II. III. IV. V. VI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . General Characteristics of Benzoxazinones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biosynthesis of Benzoxazinoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mode of Action Benzoxazinoid Aglucones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Molecular Allelopathy of Benzoxazinones and Benzoxazolinones . . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

319 320 324 332 335 341 341

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

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CONTRIBUTORS

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

Richard D. Allen (277), Pacific Biomedical Research Center, Department of Microbiology, University of Hawaii, Honolulu, Hawaii 96822 Julien Dubrulle (1), Developmental Biology Institute of Marseille, Campus de Luminy, Case 907, 13288 Marseille Cedex 09, France Agnes K. Fok (277), Biology Program, University of Hawaii at Manoa, Honolulu, Hawaii, 96822 Monika Frey (319), Institut fu¨r Genetik, Technische Universita¨t Mu¨nchen, D-85747 Garching, Germany Alfons Gierl (319), Institut fu¨r Genetik, Technische Universita¨t Mu¨nchen, D-85747 Garching, Germany Estelle Hirsinger (1), Institute of Neuroscience, University of Oregon, Eugene, Oregon 97402 Caroline Jouve (1), Developmental Biology Institute of Marseille, Campus de Luminy, Case 907, 13288 Marseille Cedex 09, France Shinsuke Kanamura (109), Department of Anatomy, Kansai Medical University, Osaka 570-8506, Japan John Kim (153), Section of Microbiology, Division of Biological Sciences, University of California, Davis, Davis, California 95616 Daniel J. Klionsky (153), Section of Microbiology, Division of Biological Sciences, University of California, Davis, Davis, California 95616 Yan Yu. Komissarchik (203), Institute for Cytology, Russian Academy of Sciences, St. Petersburg 194064, Russia ix

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CONTRIBUTORS

Oliver Pourquie´ (1), Developmental Biology Institute of Marseille, Campus de Luminy, Case 907, 13288 Marseille Cedex 09, France Margot Schulz (319), Institute of Agricultural Botany, University of Bonn, D-531115 Bonn, Germany Sidney V. Scott (153), Section of Microbiology, Division of Biological Sciences, University of California, Davis, Davis, California 95616 Dieter Sicker (319), Institute of Organic Chemistry, University of Leipzig, D-04103 Leipzig, Germany Ekaterina S. Snigirevskaya (203), Institute for Cytology, Russian Academy of Sciences, St. Petersburg 194064, Russia Gail L. Waring (67), Department of Biology, Marquette University, Milwaukee, Wisconsin 53201-1881 Jun Watanabe (109), Department of Anatomy, Kansai Medical University, Osaka 5708506, Japan

Somite Formation and Patterning Estelle Hirsinger, Caroline Jouve, Julien Dubrulle, and Olivier Pourquie´ Laboratoire de Ge´ne´tique et de Physiologie du De´veloppement (LGPD), Developmental Biology Institute of Marseille (IBDM), CNRS-INSERMUniversite´ de la Me´diterrane´e-AP de Marseille, Campus de Luminy, Case 907, 13288 Marseille Cedex 09, France

As a consequence of their segmented arrangement and the diversity of their tissue derivatives, somites are key elements in the establishment of the metameric body plan in vertebrates. This article aims to largely review what is known about somite development, from the initial stages of somite formation through the process of somite regionalization along the three major body axes. The role of both cell intrinsic mechanisms and environmental cues are evaluated. The periodic and bilaterally synchronous nature of somite formation is proposed to rely on the existence of a developmental clock. Molecular mechanisms underlying these events are reported. The importance of an antero-posterior somitic polarity with respect to somite formation on one hand and body segmentation on the other hand is discussed. Finally, the mechanisms leading to the regionalization of somites along the dorso-ventral and medio-lateral axes are reviewed. This somitic compartmentalization is believed to underlie the segregation of dermis, skeleton, and dorsal and appendicular musculature. KEY WORDS: Somite, Chick embryo, Muscle, Vertebra, Dermis, Segmentation, Somitogenesis, Signaling: 䊚 2000 Academic Press.

I. Introduction In vertebrates, all skeletal muscles of the trunk arise from the transient embryonic somites. In amniotes, these structures first appear as epithelial spheres, which provide the early framework on which the segmental pattern of the body is established. Somite formation occurs sequentially and synchronously on both sides of the neural tube with each epithelial sphere pinching off from the anterior extremity of the caudal unsegmented paraxial mesoderm (Fig. 1). Somites form with a constant periodicity until the International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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Copyright 䉷 2000 by Academic Press All rights of reproduction in any form reserved.

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FIG. 1 Schematic representation of paraxial mesoderm differentiation. Somitic material is produced by ingression of material from the epiblast through the anterior primitive streak and later on from the tail bud. These cells become organized as two stripes of mesenchymal PSM which subsequently form epithelial spheres called somites. This process called somite segmentation occurs in a coordinated fashion on both sides of the midline organs (neural tube and notochord). Somites become subsequently polarized into a ventral mesenchymal compartment called the sclerotome which will give rise to the axial skeleton and a dorsal epithelial dermomyotome. This latter compartment will yield all the skeletal muscles of the body and the dermis of the back. It will later subdivide into a myotome which comes to lie

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definitive number, characteristic of the species, is reached (Richardson et al., 1998). In amniotes, somites give rise to the vertebral column, ribs, all the skeletal muscles of the trunk and limbs, and the dermis of the back; they also contribute endothelial cells, smooth muscles, and connective tissues such as the meninges. In this review, we will summarize our current knowledge of the development of somites in the chick embryo and compare this to what is known in the mouse embryo. Due to the major advantages of using avian embryos as a model system to study embryonic development, such as their accessibility at early stages, much of our understanding of somitogenesis has come from analyses of this organism. Genetic studies in mice greatly contributed to the analysis of this mechanism at the molecular level. We will discuss our current comprehension of somitogenesis from the establishment of the somitic territory during gastrulation to the specification of the various somitic lineages. Somite regionalization along the antero-posterior body axis and the role of the homeotic genes in that process is not discussed here; refer to Gossler and Hrabe de Angelis (1998) for a recent review on the subject. In the chick embryo, the somitic material becomes specified during gastrulation when a lateral territory of the epiblast ingresses into the primitive streak. This invaginated material forms the presumptive territory of the paraxial mesoderm, which thereafter resides in the rostral primitive streak. During regression of the primitive streak and tail bud elongation, paraxial mesoderm migrates through the node and laterally and comes to lie on both sides of the neuraxis. The first somite forms from the anterior end of the presomitic mesoderm (PSM) after 24 hr of incubation at stage HH 7 (Hamburger, 1992). During chick somitogenesis, which lasts up to embryonic day 5, approximately 52 somites are formed. Somites follow a rostrocaudal gradient of differentiation, the anterior-most somites being the oldest ones. Therefore, at different embryonic ages, the differentiation stage of a somite can be measured with respect to the time elapsed since its segmentation. This led Ordahl (1993) to propose a somite staging system reflecting this intrinsic state of differentiation, independent of the embryonic stage. According to this system, the most recently formed somite is called somite I, the second most recently formed is somite II, and so on. This nomenclature will be extensively used in this review.

between the sclerotome and the dermatome. The myotome provides all the epaxial muscles. dm, dermomyotome; m, myotome; nc, notochord; np, neural plate; nt, neural tube; psm, PSM; sc, somitocoele; scl, sclerotome; wd, wolffian duct. The level of somite I and V (SI and SV) is indicated.

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The primary segmentation of the vertebrate embryo displayed by somitic organization underlies much of the segmental organization of the body. The primary segments, the somites, can be further subdivided into anterior and posterior compartments, which exhibit different properties with respect to neural crest cell and motor axon migration through the somite. This somitic subdivision is thus responsible for peripheral nervous system segmentation. The embryonic vasculatory system initiates development during somitogenesis. While the dorsal aorta elongates caudally, branches arise laterally and bifurcate dorsally each passing through an intersomitic cleft. Thus, the segmental pattern of blood vessels is also largely imposed by the somites (Christ et al., 1998). Recently, new insights have been gained about the molecular mechanisms involved in somite segmentation and the formation of rostral and caudal somite compartments. Implications of the Notch signaling pathway in these processes and the role of a newly identified molecular clock linked to segmentation will be discussed. In the chick and mouse embryo, the first somite lies immediately caudal to the otic vesicle (Huang et al., 1997) and participates in the formation of the occipital bone of the skull. Anterior to this somite, the paraxial mesoderm is referred to as head or cephalic mesoderm. This anterior tissue will contribute to skeletal muscles and bones of the head. The issue of whether this tissue is segmented is still a matter of controversy. In this review, we will focus strictly on somites. Somites bud off from the anterior segmental plate as epithelial balls surrounding a cavity, the somitocoele, containing mesenchymal cells. By somite stage III–IV, the dorsal portion of the somite remains epithelial and constitutes the dermomyotome while its ventral moiety undergoes an epithelio-mesenchymal transition leading to the formation of the sclerotome. The sclerotome, which also contains the somitocoele cells, gives rise to the skeletal elements. By stage X, these cells migrate either ventrally toward the notochord to form vertebral bodies and intervertebral discs, or dorsally to constitute neural arches while at the thoracic level they also form the proximal ribs. The distal ribs have been shown to derive from the dermomyotome (Kato and Aoyama, 1998). The dermomyotome predominantly contains dermal precursors, which contribute to the dermis of the back, and the skeletal muscle precursors. By stage VII, the myotome, from which paraspinal muscles arise, starts to migrate from the dermomyotomal medial edge and comes to lie between the dermomyotome and the sclerotome. Laterally at the interlimb level, a lateral myotome forms in mirror image to the medial edge and gives rise to abdominal and intercostal muscles. At the wing level from a stage VII somite and at the leg level from a stage I somite, cells disperse from the dermomyotome lateral edge, migrate through lateral mesoderm and settle in limb mesenchyme where they coalesce to form premuscular masses and

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further differentiate as appendicular muscles. Connective tissues of the trunk muscles also derive from somites, whereas those of the limb muscles along with their skeletal elements arise from the lateral plate. Endothelial cells arise from all the compartments of the somite to contribute to vessel formation throughout the trunk region (Wilting et al., 1995). By stage XV, the dermomyotome undergoes a global epithelio-mesenchymal transition, releasing muscle precursors ventrally and precursors of the back dermis dorsally, which migrate farther dorsally to reach their final position under the epidermis. The other regions of dermis derive from lateral mesoderm. Thus, the epithelial somite, which initially appears as a homogenous structure, undergoes regionalized morphogenetic transformations leading to the formation of a great variety of tissues. How is this diversity of cell types generated within the epithelial somite? After discussing the means by which paraxial mesoderm is specified and subsequently segments into somites, we will review the studies of how the somites are molecularly patterned along the three classical axes (i.e., the antero-posterior, the dorsoventral, and the medio-lateral axes). The intersections of these axes define compartments from which different cell types arise. Cell autonomous mechanisms controlling cell fate decisions will also be discussed.

II. Origin and Commitment of the Somitic Cells A. Origin of Somitic Cells in the Gastrulating Embryo Various lineage tracing techniques have been used to follow cell movement and to fate map presumptive territories in the avian blastoderm (Nicolet, 1971; Ooi et al., 1986; Psychoyos and Stern, 1996a; Spratt, 1955). These include labeling cells with vital dyes or with fluorescent or histochemical markers or constructing quail–chick chimeras. Using such techniques in the chick embryo, the progenitors of the paraxial mesoderm have been localized in mid and definitive primitive streak stages (stages 2–4 HH) to the epiblast lateral to the midline and subsequently to the anterior primitive streak. When the primitive streak regresses, it lays down in its wake, on both sides of the axis, a stream of mesenchymal PSM cells which will subsequently segment into somites (Hatada and Stern, 1994; Psychoyos and Stern, 1996a; Schoenwolf et al., 1992a; Selleck and Stern, 1991). When the primitive streak has totally regressed, its remnant occupies a small zone at the caudal end of the embryo which forms the tail bud. This structure will provide the remainder of the paraxial mesoderm cells (Catala et al., 1995). A similar distribution of the somite presumptive territory is also found in the mouse embryo where progenitors of the somites have been

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localized to the rostral primitive streak and then to the tail bud (Tam and Beddington, 1987, 1992). After complete ingression of the epiblastic material around stage 4 in chick, the presumptive cells of the somites are found in the anterior part of the primitive streak (Psychoyos and Stern, 1996a; Schoenwolf et al., 1992a; Selleck and Stern, 1991). They reside in this structure and later in the tail bud as a population of stem cells giving rise to the progenitor cells from which all the somites arise (Nicolas and Bonnerot, 1988; Nicolas et al., 1996; Stern, 1992). The antero-posterior organization of the embryonic mesodermal territories of the primitive streak corresponds to the future medio-lateral organization of the embryo. Thus the most rostral territory corresponds to the notochord, followed more caudally by the somites and then by the intermediate mesoderm, lateral plate and extra-embryonic mesoderm (Psychoyos and Stern, 1996a; Schoenwolf et al., 1992b). This order also seems to apply within a defined territory because, in the case of somites, the most rostral part of the streak contains the medial somitic progenitors whereas lateral somitic cells are localized in more caudal aspects of the rostral streak (Selleck and Stern, 1991). In the primitive streak, there is no evident prepattern reflecting the future distribution of the somitic cells along the antero-posterior axis, at any defined time point. However, comparing cells at equivalent levels of the primitive streak at different ages shows that they are fated to populate progressively more caudal somites as the age of the embryo increases. It is therefore unlikely that the antero-posterior distribution of the somites reflects by an early spatial arrangement of progenitor cells in the primitive streak.

B. Specification of Precursors Cells to Form Paraxial Mesoderm Paraxial mesoderm precursors in the epiblast and the primitive streak show extensive developmental plasticity and are not stably committed to a definitive fate. In chick, the prospective paraxial mesoderm territory in the epiblast can differentiate into neural tissue when ectopically grafted into the prospective neurectoderm (Garcia-Martinez et al., 1997). Similarly, when anterior streak tissue is transplanted to the posterior streak level, it differentiates into extra-embryonic or lateral plate mesoderm indicating that somitic precursor cells in the anterior primitive streak are also not committed (Beddington, 1982; Garcia-Martinez and Schoenwolf, 1992). These anterior primitive streak cells can also contribute to the neural plate when transplanted to the prospective neuro-ectoderm region of the epiblast (GarciaMartinez et al., 1997) and will differentiate into notochord when grafted

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into Hensen’s node of the mid-streak state embryo (Selleck and Stern, 1992). Even more surprisingly, complete ablation of Hensen’s node and rostral primitive streak including the whole presumptive somitic territory leads to the development of a normal embryo when performed at stage 3⫹/4⫺ in the chick embryo (Psychoyos and Stern, 1996b). These experiments demonstrate the lack of commitment of both the epiblast and rostral primitive streak to form paraxial mesoderm. Even in the PSM and caudal somites, cells are not stably committed to a somitic fate. Quail caudal somites, when grafted under the ectoderm of a primitive streak stage embryo, form mainly mesoderm but also, to a lesser extent, endoderm, lateral plate, and endothelium (Veini and Bellairs, 1991). In addition, cell lineage studies using injection of a fluorescent tracer into cells of the caudal part of the PSM show that these cells can contribute both to somites and to intermediate or lateral mesoderm (Stern et al., 1988). It has also been shown that PSM can be converted to lateral plate by treatment with BMP4 (Tonegawa et al., 1997; Tonegawa and Takahashi, 1998). At the PSM level, paraxial mesoderm identity seems to correspond to an unstable state which is maintained by a balance between opposing signals such as BMP4 and its antagonists Noggin or Chordin, expressed on either side of the PSM. Whereas the fate of cells ingressing at different levels of the primitive streak are well described in mouse and chick embryos, the molecular mechanisms that specify these fates, and in particular those responsible for the commitment of these cells to form paraxial mesoderm, are poorly understood. In chick, Hensen’s node appears to be dispensable for paraxial mesoderm specification because its ablation, at least up to stage 4⫺, does not affect somitogenesis (Psychoyos and Stern, 1996b). In addition, mouse embryos lacking the node and notochord do form somites (Ang and Rossant, 1994; Weinstein et al., 1994). Mutational analyses in mice have begun to elucidate the role of signalling molecules during paraxial mesoderm specification. The phenotype of FGFR1-deficient mice indicates a potential role for FGF signaling in specifying paraxial mesoderm cell fates. FGFR1-deficient embryos exhibit an expansion of the axial mesoderm, primitive streak defects, and an absence of somites (Yamaguchi et al., 1994). The loss of somites seems to be a secondary consequence of the expansion of the axial mesoderm at the expense of paraxial mesoderm. In embryos lacking another signaling molecule, Wnt3a, somitogenesis is disrupted after the formation of the first 5–6 somites (Takada et al., 1994). A similar phenotype has been observed in mutant embryos lacking Brachyury (Wilson et al., 1995). In the Tbx6 mutant, trunk somites are transformed into ectopic neural tubes flanking the axis, resulting in a mouse with three neural tubes (Chapman and Papaioannou, 1998). These molecules therefore appear to be involved in segregating

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the presumptive embryonic territories corresponding to axial mesoderm, paraxial mesoderm, and neural plate. In all these mutants, it seems that the first few somites that are formed are spared from the effects of the mutation. A similar observation has also been made in most of the mutants of the Notch-delta signaling pathway in which only the first few somites form. This suggests that the activity of these genes is required specifically for the formation and segmentation of the more posterior somitic mesoderm, indicating that formation of the anterior somites could have different requirements. Although a few genes have been characterized as potential regulators of paraxial mesoderm identity, thus far, no gene has been found to be required specifically for the specification of the paraxial mesoderm along the entire body axis.

III. Somite Formation A. A Prepattern of Segmentation in the Paraxial Mesoderm? Once paraxial mesoderm has been formed, the next issue is to understand how the mesenchymal PSM tissue becomes transformed into epithelial spheres that form iteratively and synchronously on both sides of the embryo. Meier has proposed that the PSM displays a metameric arrangement of groups of cells called somitomeres. These transient structures are visible only by stereo scanning electron microscopy (Meier, 1984), and their existence remains controversial (Wachtler et al., 1982b). According to Meier and colleagues, these groups of mesenchymal cells are organized around a central point and are formed in a strictly anterior to posterior order. The number of somitomeres in the PSM varies between species but remains constant within a given species (12 in the chicken and 6 in the mouse), even though the length of the PSM changes during development (Tam and Beddington, 1986). The number of somitomeres was shown to correspond to the number of prospective somites of the PSM, thus reflecting an early spatial allocation of these cells along the antero-posterior axis ( Jacobson, 1988; Tam and Beddington, 1986). Lineage analysis of chick or mouse PSM cells suggests that somitomeres are not acting as clonal compartments like the rhombomeres (Fraser et al., 1990). Clonal descendants of single cells or groups of cells are often found to cross the boundary between somitomeres (Bagnall, 1992; Bagnall et al., 1992; Stern et al., 1988; Tam, 1988; Tam and Beddington, 1987). Furthermore, when a portion of chick segmental plate is replaced by a piece of quail segmental plate of an equivalent size, which is known to contain more

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prospective somites, the number of somites formed by the quail transplant is equal to the number of chick prospective somites that have been removed and not the same as the number of quail prospective somites that have been transplanted (Packard et al., 1993). Therefore, if a prepattern of the PSM exists, it is not entirely fixed and is subject to regulating processes until it undergoes segmentation. Moreover, genetic expression patterns offer little support for the somitomere concept since no gene has been observed to be expressed in a somitomeric fashion except in the most rostral PSM. Therefore it seems that apart from the original arguments based on scanning electron microscope images, little evidence has been provided to confirm the existence of somitomeres in the PSM. Nonetheless, because little cell movement is observed in the PSM, the distribution of cells in this tissue, to some extent, reflects a prepattern corresponding to their future relative position in the somites along the antero-posterior axis.

B. Tissue Interactions Involved in Somite Formation Interactions between the paraxial mesoderm and surrounding tissues, in particular the axial structures, were postulated to be linked directly to the process of somite formation (Fraser, 1960; Keynes and Stern, 1988). It was observed that the PSM can segment in isolation of the neural tube, the notochord, or the endoderm (Packard and Jacobson, 1976; Sandor and Fazakas-Todea, 1980; Tam and Beddington, 1986). However, these explants failed to form somites in the absence of ectoderm (Ladher et al., 1996; Lash and Yamada, 1986; Palmeirim et al., 1998), although surprisingly, they displayed segmental properties as evidenced by deltal mRNA expression (Palmeirim et al., 1998). In addition, ablation of the ectoderm in vivo prevents somite formation in the operated region (Sosic et al., 1997). Therefore, establishment of the segmental pattern in the PSM appears to be an intrinsic property of this tissue, although formation of the epithelial somite requires signaling from the ectoderm.

C. Cell–Cell and Cell–Matrix Interactions During Somitogenesis Cells destined to form somites undergo profound changes in their adhesive properties during somitogenesis. These cell–cell and cell–matrix interactions are essential to somite formation, as was demonstrated by disrupting these interactions. Specific cell–cell adhesion is dependent not only on the presence of tissue-specific cell adhesion molecules (CAMs) but also on the

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spatio-temporal regulation of CAM types. In the chick paraxial mesoderm, N-cadherin, cadherin-11, and NCAM expression are up-regulated during differentiation (Duband et al., 1987; Kimura et al., 1995). Enhanced cadherin expression may be related to the increase in cell adhesivity observed during PSM maturation and somite formation (Bellairs et al., 1978). Accordingly, the epithelial structure of the newly formed somites can be disrupted by in vitro treatment with antibodies against N-cadherin (Duband et al., 1987). A null mutation of N-cadherin led to small and irregular somites whose epithelial structure was partially disrupted (Radice et al., 1997). In contrast, the loss of function mutation in the NCAM gene had no effect on somite formation (Cremer et al., 1994; Duband et al., 1987). In mouse and chick embryos, differentiation of the PSM is accompanied by the elaboration of matrix materials (Duband et al., 1987; Newgreen, 1984; Ostrovsky et al., 1988). In the mouse, the amount of fibronectin in the mesenchyme and at the interface with adjacent epithelia increases during the maturation of the PSM. Interestingly, somites do not form in mouse embryos mutant for the fibronectin gene (George et al., 1993). Similar phenotypes are observed in mice mutant for focal adhesion kinase (FAK), a nonreceptor tyrosine kinase implicated in transducing signals generated by cell–matrix interactions (Furuta et al., 1995). Thus, the dynamic expression patterns of adhesion molecules, along with experiments disrupting the interactions they mediate, suggest that cell–cell and cell– matrix interactions play an important role during somitogenesis.

D. Molecular Aspects of Somitogenesis 1. A Molecular Clock Linked to Somitogenesis Several theoretical models have been proposed to account for somitogenesis, a process characterized by its rythmicity and bilateral synchrony. Some of these models are based on cell communication and implicate adhesion molecules or Notch-delta mediated lateral inhibition as the driving force of somite formation (Conlon et al., 1995). But these models do not account for the periodicity of this process. Others such as the ‘‘clock and wave front’’ model (Cooke and Zeeman, 1976), Meinhardt’s model (Meinhardt, 1986), or the cell cycle model (Stern et al., 1988) proposed the existence of an oscillator or clock in the presomitic cells. The purpose of such an oscillator was to generate a temporal periodicity, which would be translated into the spatial periodicity of the somites. Recently, the identification of c-hairy1, an avian homolog of the fly pair rule gene, has provided molecular support for the existence of such a clock linked to segmentation (Palmeirim et al., 1997). This gene is strongly expressed in the PSM where its mRNA

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exhibits cyclic waves of expression whose temporal periodicity corresponds to the time required for the formation of one somite (Palmeirim et al., 1997). In vitro studies have demonstrated that these waves result from an intrinsic property of the PSM and do not rely on cell migration or extrinsic signal propagation. Prospective somitic cells express pulses of c-hairy1 mRNA as soon as they leave the site of gastrulation and enter the PSM. Because the PSM contains 12 prospective somites in the chick, cells will undergo 12 c-hairy1 expression cycles before their incorporation into a somite. In zebrafish, a homolog of hairy, her1, has been identified (Muller et al., 1996). This gene presents no dynamic PSM expression as observed in chick, but it is expressed in alternate primordia of the presumptive somites and thus follows a pair-rule expression in the PSM. her-1 expression constitutes the only evidence for a pair-rule type of mechanism in vertebrates. This result raises the possibility that part of the machinery involved in the segmentation process has been conserved between Drosophila and zebrafish. Interestingly, lunatic fringe has been recently identified in the chick and mouse as another gene expressed in a cyclical fashion in the PSM (Forsberg et al., 1998; McGrew et al., 1998). lunatic fringe mRNA is expressed in a rhythmic fashion in the PSM, with a periodicity corresponding to the formation of one somite. When protein synthesis is blocked, the lunatic fringe wave is abolished, in the contrast to c-hairy1, suggesting that lunatic fringe could act dowstream of c-hairy1. It appears that c-hairy1 and lunatic fringe are two genes regulated by the clock linked to somitogenesis. The role of the lunatic fringe gene has been investigated in the mouse by mutational analysis. Mutant embryos exhibit defects in somite formation, demonstrating a crucial role for this gene during somitogenesis (Evrard et al., 1998; Zhang and Gridley, 1998). In Drosophila, fringe has been shown to play a role in modulating Notch-delta signaling, resulting in the establishment of the dorso-ventral boundary of the wing margin (Fleming et al., 1997; Panin et al., 1997). As described later, the Notch-delta signaling pathway is indeed involved in defining somite boundaries. 2. Role of the Notch Signaling Pathway Some major contributions to the understanding of the molecular mechanisms underlying somitogenesis have come from the use of gene knockout experiments in the mouse. Surprisingly enough, the study of the homologs of fly neurogenic genes in vertebrates revealed that homologs of these genes implicated in the Notch-delta pathway are expressed in the PSM, including the receptors Notch1 (Conlon et al., 1995; Swiatek et al., 1994), and Notch2 (Lindsell et al., 1996; Weinmaster et al., 1992); the ligands

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Delta1 (Bettenhausen et al., 1995), Delta3 (Dunwoodie et al., 1997), and Serrate/Jagged1 (Lindsell et al., 1996; Mitsiadis et al., 1998); downstream effectors such as the suppressor of hairless homolog RBP-jK (Oka et al., 1995) and the enhancer of split homolog HES5 (de la Pompa et al., 1997), as well as other genes involved in this pathway such as Presenilin1 (Wong et al., 1997) and lunatic fringe (Evrard et al., 1998; Forsberg et al., 1998; Johnston et al., 1997; McGrew et al., 1998; Zhang and Gridley, 1998). Inactivation of most of these genes leads to a strong disruption of somitic segmentation. In Notch1 mutant mice, somite formation occurs but its bilateral coordination is affected (Conlon et al., 1995). A similar phenotype is observed in mutants for Delta1 and Delta3, where somites are present but not fully epithelialized (Hrabe de Angelis et al., 1997; Kusumi et al., 1998). Mutation of the mouse homologs of other genes involved in Notch signaling such as RBP-jK or Presenilin1 also generates somitogenesis defects (Oka et al., 1995; Wong et al., 1997). RBP-jK is the vertebrate homolog of the fly suppressor of hairless gene which codes for a transcription factor acting downstream of Notch (Fortini and Artavanis-Tsakonas, 1994; Jarriault et al., 1995). Mice embryos mutant for RBP-jK have irregularly shaped somites, as in the Notch1 and Delta1 mutants, however the phenotype is more severe than in the Notch1 mutant, suggesting some degree of Notch redundancy during embryogenesis. This might be expected because other Notch genes are similarly expressed in the PSM. The Presenilin1 gene codes for a multipass transmembrane protein whose C. elegans homolog has been shown to act in the Notch-delta pathway (Rohan de Silva and Patel, 1997). Mice mutant for Presenilin1 exhibit somitogenesis defects similar to those observed for the other members of the Notch pathway (Wong et al., 1997). The importance of the Notch-delta pathway during somitogenesis is further supported by the analysis of Xenopus X-Delta2. Injection of RNA coding for dominant negative form of X-Delta2, as well as ectopic expression of X-Delta2 in Xenopus embryos leads to perturbations of segmentation and, in the more severe cases, abolishes the segmental pattern ( Jen et al., 1997). It is apparent that any disruption of the activity of the Notch signaling pathway leads to abnormal somitogenesis without the loss of segments. Therefore, the Notch-delta pathway does not seem to play a crucial role in the establishment of the basic metameric pattern, but rather seems to coordinate and fine-tune somite formation. In Drosophila, this pathway is implicated, along with other systems, in the process of lateral inhibition, which controls cell fate choice in the nervous system and in the process of boundary definition in the establishment of the wing margin (Blair, 1997; Kimble and Simpson, 1997). In vertebrates the role of this pathway during somitogenesis might be more related to this latter function, as evidenced by the phenotype of lunatic fringe mutant mice (Evrard et al., 1998; Zhang and Gridley, 1998). In homozygous null mice, somites form but exhibit

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irregularities in size and shape. Loss of lunatic fringe function also leads, in the rostral PSM, to loss of the sharply demarcated domains of expression of several genes encoding members of the Notch-Delta signaling. Thus, a possible explanation for the lunatic fringe phenotype is that the loss of Notch-delta signaling may alter the positioning of the somite boundaries and the allocation of cells to the nascent somite. From these observations, it is likely that the Notch-delta signaling pathway acts downstream of the clock linked to somitogenesis and that lunatic fringe is the molecular link between this clock and the Notch-delta pathway. Thus, clock control of the local modulation of the Notch signaling pathway could confer the periodic arrangement of boundaries that underly the segmental body plan.

3. Other Players in Somitogenesis A significant role for bHLH transcription factors other than hairy during somitogenesis has been demonstrated. The related transcription factors Mesp1 and 2 in the mouse, Thylacine1 in the frog, and c-meso1 in the chick are expressed in the PSM as a stripe in its rostral part (Buchberger et al., 1998; Saga et al., 1996, 1997; Sparrow et al., 1998). A mutation in the Mesp2 gene leads to defects in somitogenesis, particularly at the level of the caudal somites (Saga et al., 1997). In these mutants, expression of Notch1 and 2 as well as FGFR1 is strongly down-regulated in the PSM, suggesting that Mesp2 acts upstream of these factors. Accordingly, overexpression studies of the frog Mesp homolog, Thylacine1, indicate that it is part of the Notch signaling cascade important for somite segmentation. Another bHLH transcription factor, paraxis, is expressed in the rostral PSM and in the newly formed epithelial somite (Blanar et al., 1995; Burgess et al., 1995). Mutation of this gene in the mouse leads to a loss of somite formation, whereas the segmented arrangement of the paraxial mesoderm derivatives is maintained (Burgess et al., 1996). Furthermore, blocking expression of paraxis, using antisense oligonucleotides in the chick, leads to an impairment of somite epithelialization (Barnes et al., 1997). In the chick embryo, paraxis expression correlates with the epithelialization of the PSM, and it was shown that its expression is under the control of factors derived from the ectoderm, which is necessary for somite epithelialization (Gallera, 1966; Palmeirim et al., 1998; Sosic et al., 1997). However, paraxis expression is not sufficient for epithelization because its expression is maintained in cultured isolated PSM explants, which fail to form epithelial somites (Palmeirim et al., 1998). These results show that the segmentation process of mesodermal derivatives can be dissociated from the formation of the epithelial somite.

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IV. Antero-posterior Compartmentalization of the Somites As discussed in the next section, somite formation might also necessitate the juxtaposition of anterior and posterior compartments within the future somite. Later on, this compartmentalization along the antero-posterior axis is clearly visible in the sclerotome whose anterior and posterior halves exhibit distinct cellular and molecular properties.

A. Anatomical and Biochemical Compartmentalization of the Sclerotome During somite maturation, the ventral part of the somite de-epithelializes to form the sclerotome, a mesenchymal tissue, which contains the precursors of the axial skeleton. The polarization of the sclerotome into rostral and caudal compartments, which differ in their permissivity to neural crest cell and motor axon migration, has been well studied and has given rise to the notion of antero-posterior (AP) compartmentalization of the somite. Histological analyses have shown that the two sclerotome halves are morphologically separated by a cleft called the Von Ebner’s fissure. This fissure separates the rostral from the caudal sclerotome. Both compartments differ in their cell density which is higher in the caudal half (Stern and Keynes, 1987; Von Ebner, 1888). Thus, the paraxial mesoderm becomes histologically segmented by two kinds of boundaries—the intersomitic clefts and the intrasomitic clefts. Rostral and caudal sclerotome halves also differ in the biochemical composition of their extracellular matrix. The caudal moiety contains versican, a chondroitin sulfate proteoglycan (Landolt et al., 1995), as well as collagen IX (Ring et al., 1996), and reacts with fluorescent Peanut agglutinin (PNA), whereas the rostral moiety is devoid of these molecules (Bagnall and Sanders, 1989). Several cell adhesion molecules are also differentially expressed in the sclerotome. For example, T-cadherin is specifically present in the caudal half of the somite (Ranscht and Bronner-Fraser, 1991). Thus, these morphological, biochemical, and molecular differences between the two parts of the sclerotome are thought to play an important role in the control of migration of both neural crest cells and motor axons. In vitro studies have implicated adhesion molecules, such as T-cadherin or collagen IX, in inhibiting both axon and neural crest cell migration (Fredette et al., 1996; Ring et al., 1996). More recently, Eph family ligands and receptors have been shown to display differential AP sclerotomal expression and are implicated in the

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control of axon guidance in the paraxial mesoderm. EphB3, an Eph receptor tyrosine kinase, localizes to the rostral half of the sclerotome and is also expressed by neural crest cells migrating through this territory, whereas the ligand ephrin B1 is restricted to the caudal half (Krull et al., 1997), where neural crest cells do not migrate. Similarly, Wang and Anderson (1997) have demonstrated the presence of the Eph-related receptors TUJ1 in both migrating neural crest cells and motor axons, whereas Lerk-2, an Eph ligand, is detected in regions devoid of peripheral nervous system derivatives such as the caudal half of the sclerotome and the dermomyotome. Neural crest cells are unable to migrate on substrates containing Eph family ligands such as HtkL or Lerk-2, suggesting that these molecules provide repulsive signals to the migrating neural crest cells (Krull et al., 1997; Wang and Anderson, 1997). Thus, neural crest cell migration and motor axon outgrowth in the rostral part of the sclerotome is likely to be due to a repulsive activity of molecules specifically expressed in the caudal half of the sclerotome. These Eph-related molecules are also likely to play a role in defining or maintaining the integrity of compartments or compartment boundaries. In the hindbrain, cek8, an Eph-related receptor, is expressed in rhombomeres 3 and 5. In retinoic acid-treated embryos, the posterior morphological boundaries of rhombomeres are lost, and this phenomenon is preceded by the down-regulation of cek8 in rhombomere 5 (Nittenberg et al., 1997). A similar role for the Eph-related family of molecules in the segregation of the rostral and caudal somitic compartments and in maintainance of the intrasomitic boundary was recently demonstrated in zebrafish (Durbin et al., 1998).

B. Sclerotomal Fate and the Resegmentation Theory Subdivision of the sclerotome into rostral and caudal compartments preempts the future contribution of the sclerotomal cells to the vertebral column (see for a review Christ and Wilting, 1992). Cell lineage analyses of various parts of the somite, including the caudal and rostral halves have been performed using the quail–chick chimaera system. By orthotopical replacement of one cervical somite, it has been possible to follow the contribution of a single embryonic somite to the vertebral column (Bagnall et al., 1998). Cells derived from a single somite colonize a delimited region which comprises one half of each of two adjacent vertebrae as well as the intervening disc (Bagnall et al., 1988; Huang et al., 1996). Moreover, grafting trains of multiple caudal or rostral halves has shown that each part of the sclerotome contributes to defined vertebral structures. In multiple caudal half-somite grafts, the pedicle of the vertebral arch is abnormally extended

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and the intervertebral disc almost disappears. In contrast, in multiple rostral half-somite replacements, the pedicle is almost absent and the intervertebral disc is present (Goldstein and Kalcheim, 1992). These results suggest that a vertebra is formed from the caudal parts of one somite pair and the rostral parts of the next pair. This phenomenon, which was first described by Remak (1850) and later reinforced by von Ebner (1888) during the last century, led to the theory of resegmentation: the boundaries of the definitive adult segments do not correspond to the embryonic metameric units because they are shifted by half a segment. Thus, the initial segments, the somites could represent ‘‘parasegments,’’ a situation which is similar to that described in the fly for body segments (Lawrence, 1992). This concept is however still controversial because conflicting results have been obtained (Baur, 1969; Blechschmidt, 1957; Christ et al., 1979; Tajbakhsh and Sporle, 1998; Verbout, 1985).

C. Establishment of Antero-posterior Compartmentalization 1. Intrinsic Determination: Cellular and Molecular Aspects In contrast to the dorso-ventral or medio-lateral polarity of the somite, AP polarity is already established when the somite forms (Aoyama and Asamoto, 1988). Experimental evidence has demonstrated that this polarity is not determined by surrounding cues provided by the environment of the somite. For instance, inversion of the AP axis of the PSM leads to an inversion of the progression of segmentation from caudal to rostral. The somites formed from the inverted PSM present a reversed AP polarity as evidenced by neural crest cells which now migrate through the caudal part of the somite (former rostral) or by the inversion of the c-delta1 expression pattern (Bronner-Fraser and Stern, 1991; Palmeirim et al., 1998). All these data indicate that AP compartmentalization is determined within the PSM. Several genes have been identified; they are restricted to either the caudal or rostral part of the newly formed somite (i.e., before the sclerotome has formed, such that this rostrocaudal compartmentalization is already apparent by the epithelial stage). Most of the genes belonging to the Notchdelta pathway are expressed in such a restricted pattern (Bettenhausen et al., 1995; Dunwoodie et al., 1997; Henrique et al., 1995; Reaume et al., 1992). In the mouse, both Notch1 and Notch2, as well as Delta1 and Delta3, are expressed in the PSM and subsequently in either the rostral or the caudal compartments of the newly formed somites (Bettenhausen et al., 1995; del Amo et al., 1992; Dunwoodie et al., 1997; Reaume et al., 1992; Saga et al., 1996; Williams et al., 1995). In the Delta1 mutant, no epithelial somite forms in the caudal region of the embryo. The caudal identity of

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the sclerotome is lost, but the primary metameric pattern is established as indicated by the segmental arrangement of myotomes. A similar phenotype altering rostro-caudal compartmentalization is also observed in the mouse lunatic fringe mutant (Evrard et al., 1998; Zhang and Gridley, 1998). All these data suggest that the Notch-delta pathway is involved in specifying the rostro-caudal identity of the somites, as well as in maintaining segments borders, as mentioned earlier (Hrabe de Angelis et al., 1997). Similar to the mouse, chick c-delta1 is expressed in the PSM and is then restricted to the caudal part of the epithelial somite. PSM cultured in vitro without any surrounding tissue has been shown to display a segmental expression of this gene (Palmeirim et al., 1998). Alternate stripes of cdelta1 appear sequentially whereas no epithelialization is observed. Thus, epithelialization and segmental patterning defined as the rostro-caudal compartmentalization of gene expression in the rostral PSM are two independent events that can be uncoupled in vitro.

2. Relationship Between Antero-posterior Polarity and Somite Formation The following results suggest that somitic AP compartmentalization might be a prerequisite for somite formation. When multiple caudal or rostral halves of epithelial somites are placed adjacent to each other, sclerotomal cells derived from like half-somites mix, whereas juxtaposition of cells from unlike half-somites leads to a strict segregation of each population (i.e., rostral vs caudal) (Stern and Keynes, 1987). This suggests that formation and maintenance of segment borders could depend on the juxtaposition of two cell-states, such as A and P as proposed in the Meinhardt model (Meinhardt, 1986). In this model, cells of the PSM would oscillate between two different states and would eventually undergo stable acquisition of one of these states. Juxtaposition of cells in one state with cells in the other creates a border, similar to the mechanism for formation of parasegmental boundaries in Drosophila. However, a system of two-segment periodicity must be postulated and superimposed on this two-state model to enable border formation only between P and A of different somites and not between A and P of the same somite. Such a system could require the activity of genes similar to the pair-rule genes in Drosophila. In summary, the role of AP compartmentalization of the somites in the establishment of segmental pattern within the body, such as for the peripheral nervous system and vertebrae, is beginning to be well understood. Within somites, it is still unclear whether this compartmentalization is restricted to the sclerotome or whether it also impinges upon the development of the dermomyotome.

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At the molecular level, Eph molecules have been shown to have a restricted expression in the paraxial mesoderm and are thought to play a major role in maintaining the boundaries of each half somite domain. Cell–cell interactions mediated by the Notch pathway are implicated in the specification of rostro-caudal polarity of the somite as well as in the establishment of segment borders. Nevertheless, further experiments should be designed to address the molecular and cellular events that mediate alternation of anterior and posterior compartments in the paraxial mesoderm. Besides, the exact relationships between the existence of these compartments and somite formation are still unclear.

V. Dorso-ventral Patterning of the Avian Somite A. Three Different Lineages Segregate along the Dorso-ventral Axis 1. Somites Give Rise to Skeleton, Muscle, and Dermis Once formed, the epithelial somite then undergoes regionalized morphogenetic transformations along a second axis, the dorso-ventral axis. This process leads to the segregation of different somitic lineages, dermis, muscle, and skeleton. The somitic contribution to the formation of the axial skeleton has been mapped using quail–chick chimeras (Christ and Wilting, 1992). Although the six most rostral somites give rise to the basi-occipital bone of the skull (Couly et al., 1993), all the axial body skeleton derives from the remaining somites (Christ and Ordahl, 1995). The vertebrae and intervertebral disks, the ribs, and part of the scapula derive from the somites, whereas the limb and girdle skeleton derive from the lateral plate. It has now been well established that the vertebral body arises from cells of the ventro-medial sclerotome which migrate ventrally toward the notochord (Christ and Wilting, 1992; Ordahl and Le Douarin, 1992). The vertebral pedicles of the neural arches have been shown to derive from the caudal part of the somitic compartment, whereas the rostral somitic half yields the intervertebral disk (Goldstein and Kalcheim, 1992). The dorsal spinous process of the vertebra also derives from the somite, and its formation requires a specific set of interactions with the surrounding structures like the ectoderm and neural tube (Monsoro-Burq et al., 1994; Takahashi et al., 1992). The origin of the ribs is more complex because their vertebral part derives from the somitocoele and caudal somite cells (Huang et al., 1994) whereas the sternal part derives from the lateral dermomyotome (Kato and Aoyama, 1998). Therefore, although the axial skeleton arises primarily

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from the sclerotome, the dermomyotome is also able to give rise to some skeletal derivatives. The anatomical origin of the myotome has been and still is a subject of controversy. Three major models have been proposed. Williams, among others, proposed in 1910 that the myotome originates from the medial and, to a lesser extent, from the lateral edges of the dermomyotome. Later on, Langman and Nelson (1968) concluded, from labeling experiments with tritiated thymidine, that cells from the whole surface of the dermomyotome delaminate and form the underlying myotome. Later still, Mestres and Hinrichsen (1976) were led to believe that sclerotomal cells in contact with the overlying dermatome reaggregate to form the myotome. These different models have been put to the test using progressively more refined techniques. Christ and colleagues (1978), using quail–chick grafting techniques, demonstrated that the myotome derives exclusively from the dermomyotome, refuting Mestres and Hinrichsen’s model. By studying the onset of desmin expression in the somite, Kaehn and collaborators (1988) reached the conclusion that myogenic precursors arise strictly from the cranio-medial corner of the dermomyotome, contradicting the results obtained by Langmann and Nelson. More recently, DiI injection mapping of the dermomyotome (Denetclaw et al., 1997) demonstrated that myogenic precursors emerge not only from the cranio-medial edge of the dermomyotome but also from the entire extent of the medial lip. The existence of a subset of myotomal cells which become postmitotic very early at the epithelial somite stage, has been reported in the chick by Kahane and co-workers (1998b). These very early muscle precursors were termed pioneer cells and arise from the dorso-medial epithelial somite. They are the first cells to migrate beneath the dermomyotome and elongate along the antero-posterior axis to form the primary myotome. By analogy with Drosophila and zebrafish muscle pioneers (Ho et al., 1983; see Jellies, 1990 for a review), this primary myotome composed of pioneer cells could serve as a longitudinal scaffold, providing guidance cues for the migration of a secondary wave of myotomal precursors, which arise from the anterior and posterior lips of the dermomyotome (Kahane et al., 1998a). Due to the lack of morphological criteria to follow the differentiation of dermal cells, dermis differentiation is far less well understood than myotome or sclerotome differentiation. Only the dermis of the back arises from the somite (Christ and Ordahl, 1995). The dermis of the limbs is derived from the somatopleura, and the dermis of the face arises from the neural crest (Couly and Le Douarin, 1988). It is generally assumed that dermis arises from the dermatome (i.e., the epithelial plate located above the myotome). Beginning at E3 in the chick embryo, the dermatomes lose their epithelial structures, and their cells migrate beneath the epidermis (Brill et al., 1995).

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2. External Cues Are Involved in the Segregation of the Three Somitic Lineages How is the dorso-ventral axis established and thereby how are the different lineages specified? Two major mechanisms are classically considered to regulate cell diversification—either cell lineage relationships or environmental cues. Cell lineage analyses have been conducted in the PSM (Stern et al., 1988). When a single cell is labeled at any level of the unsegmented mesoderm, its progeny is not restricted to dermatome, myotome, or sclerotome. Progeny of the injected cell can be found in two or all three of these derivatives. Therefore it appears that, in the PSM at least, the segregation of the three fates does not rely on cell lineage mechanisms. To ask whether the establishment of the dorso-ventral axis is controlled by external cues, dorso-ventral inversion of the last three somites (Aoyama and Asamoto, 1988), or heterotopic grafts of dorsal or ventral somite pieces (Aoyama, 1993; Christ et al., 1992) were performed. The manipulated somites I and II differentiate according to their new local environment (i.e., the former dorsal half forms the sclerotome, whereas the former ventral side generates the dermomyotome). In contrast, somite III exhibits an inverted polarity suggesting that, from this level forward, the dorso-ventral axis may be irreversibly determined (Aoyama and Asamoto, 1988). Along the same lines, when quail ventral somitic halves are grafted in place of chick dorsal halves at the level of the two last formed somites, the quail cells form dermomyotome and subsequently differentiate as dermis and muscle (Christ et al., 1992). The lability of the dorso-ventral axis is progressively lost as one moves toward the anterior extremity. Taken together these results suggest that the establishment of dorsoventral polarity is under the control of environmental cues. What is their nature? Are they inductive, in the sense that they instruct a naive tissue to adopt particular cell fates, or are they permissive, in that they promote an intrinsic and preexisting potential of differentiation?

B. Self Differentiation or Induction? 1. Historical Aspects Whether somitic differentiation is an example of self-differentiation or whether it is dependent upon extrinsic cues has been a long-standing debate. Theoretical models have varied along with experimental strategies. Since the beginning of the century, numerous studies focused on chondrogenesis, mainly because manifestations of on-going chondrocyte differentiation were readily observable under the microscope. Based on the observations that isolated chick somites grafted on the chorio-allantoidian membrane

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underwent chondrogenesis and that this phenomenon was amplified if axial structures were present, the concept of self differentiation prevailed until the early 1940s (Hall, 1977). Extirpation experiments performed in the 1940s in amphibian and avian embryos established that, in vivo, the notochord and neural tube influence cartilage differentiation (Holtzer and Detwiler, 1953; Horstadius, 1944; Strudel, 1955; Watterson et al., 1954). In the 1950s, in vitro analyses of mice explant combinations clarified that cartilage forms from somitic mesoderm only if an inducer is present (Grobstein and Holtzer, 1955). Finally in the 1960s attention focused on the somite as a responding tissue, and induction was consequently considered as enhancing a preexisting potential rather than as imposing new potential on a naive tissue (Hall, 1977). 2. The Commitment State of Somitic Cells Evolves along Temporal and Spatial Axes With the characterization of molecular markers for sclerotogenic and myogenic lineages in the 1980s, the controversy of ‘‘self-differentiation versus induction’’ is now being revisited in detail with respect to chondrogenesis and myogenesis. In vivo, the paraxial mesoderm at the unsegmented level does not exhibit molecular differentiation along a particular lineage. It is only in the segmented region that somites start to express differentiation markers in a rostro-caudal gradient. To assay the state of commitment of paraxial mesoderm, this tissue was explanted and cultured in isolation. Observations of in vitro cultures of chick or mice explants led to the conclusion that, at a given stage, PSM and the last two or three formed somites are uncommited toward any lineage. In contrast, more rostral somites differentiate autonomously, suggesting that somites progress along an anteroposterior gradient of commitment (Buffinger and Stockdale, 1994; Cossu et al., 1996; Ebensperger et al., 1995; Fan and Tessier-Lavigne, 1994; Kos et al., 1998; Mu¨ller et al., 1996; Mu¨nsterberg and Lassar, 1995; Reshef et al., 1998; Rong et al., 1992; Spence et al., 1996). In addition to the antero-posterior variation of the commitment state, somites of a given level explanted at different embryonic ages exhibit increasing myogenic capacities with age (Kenny-Mobbs and Thorogood, 1987; Rong et al., 1992; Stern and Hauschka, 1995; Vivarelli and Cossu, 1986). This occurs because the rate of myogenic onset is approximately 1.5 times faster than that of somite formation (Borman and Yorde, 1994a, 1994b). These data suggest that myogenesis is controlled by both somitic competence and developmentally regulated differentiation signals and not solely by the temporal sequence of somite formation (Borycki et al., 1997; Rong et al., 1992). In contrast, activation of sclerotome markers always occurs at the same somitic level, suggesting that it is strictly controlled by

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somite formation and differentiation competence, rather than by timing of the differentiation signals (Borycki et al., 1997). Consequently, it appears that the state of somite commitment develops along both temporal and spatial axes and that specification of muscle and skeleton lineages exhibit different modalities. Another means by which one can determine when cells are stably committed is to place them in a challenging environment, that is an environment known to drive cells towards an alternative fate. By grafting tissue fragments into the limb bud, different groups have shown that myogenic potential arises in the primitive streak during gastrulation (Chen and Solursh, 1991; Krenn et al., 1988; von Kirschhofer et al., 1994) and that muscular and chondrogenic determination occurs as early as the 8-somite stage (Wachtler et al., 1982a). Williams and Ordahl (1997) used a different challenge assay in which the somitic dorso-medial quadrant was exposed in vivo to supernumerary notochords that act as sclerotome inducers. They demonstrated that a number of muscle cells manage to differentiate in this ventralizing environment, indicating their irreversible commitment to the myogenic lineage. This number increases with the age of the donor somite. In a parallel study, Dokter and Ordahl (1998) demonstrated that it is not until somite stage XII that the first cartilage progenitors cells are determined and not until E4 that the entire sclerotome is stably committed to cartilage differentiation. Therefore myogenic and sclerotogenic determination occur progressively and asynchronously during development. This suggests that, at a given time point, the somite is composed of a mosaic of cellular territories, each having reached a different state of commitment toward a specific lineage. 3. Environmental Signals Are Required for Somite Differentiation As previously mentioned, culturing PSM or epithelial somites in isolation or in association with adjacent structures revealed that this explanted tissue does not undergo differentiation unless an adjacent structure is present in the culture (Buffinger and Stockdale, 1994; Cossu et al., 1996; Ebensperger et al., 1995; Fan and Tessier-Lavigne, 1994; Kos et al., 1998; Mu¨ller et al., 1996; Mu¨nsterberg and Lassar, 1995; Reshef et al., 1998; Rong et al., 1992; Spence et al., 1996). These data demonstrate that adjacent structures are necessary for correct development of the somite, but they do not determine the exact nature of their influences. In addition to controlling the competence of the paraxial mesoderm, which will be discussed later, intercellular communications within paraxial mesoderm have been shown to play a role in a ‘‘community effect.’’ First described in Xenopus mesoderm induction (Gurdon, 1988), the ability of a cell to respond to inducing signals is enhanced when it is surrounded by

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neighboring cells following the same differentiation pathway at the same time. Indeed, such a mechanism is observed in mammals (Cossu et al., 1995) and in cultures of chick dissociated epiblasts, in which myogenesis occurs to a greater extent when cells are plated at high density than at low density (George-Weinstein et al., 1996). This ‘‘community effect’’ might be mediated by homophilic cell–cell adhesion molecules, the cadherins, as evidenced by their expression patterns and blocking experiments (Duband et al., 1987; George-Weinstein et al., 1997; Hatta and Takeichi, 1986; Holt et al., 1994). 4. But Cell–Autonomous Mechanisms May Also Function Despite the data described earlier, one cannot rule out the possibility of a cell autonomous component contributing to cell fate when one considers in vivo or tissue dissociation studies such as those described later. When the neural tube/notochord complex, which is thought to be essential for myogenic and sclerotogenic differentiation, is extirpated in vivo, a few myogenic or sclerogenic cells nevertheless differentiate from somites which have not been exposed to axial organ influence after segmentation (Bober et al., 1994; Teillet et al., 1998). Similar results have been obtained when the presumptive territory of notochord and floor plate is ablated (Catala et al., 1996). Moreover, fragments or dissociated cultures of segmental plate or of epithelial somites unlike intact explanted tissues, can undergo myogenesis and chondrogenesis (Gamel et al., 1995; George-Weinstein et al., 1994). One interpretation of these data is that cells of the PSM are already endowed with myogenic potential and that cellular interactions, possibly mediated by the Notch signaling pathway, prevent precocious expression of this potential. Signals derived from surrounding structures might therefore be permissive rather than instructive thereby imposing temporal control on the onset of chondrogenic and myogenic differentiation in the paraxial mesoderm. George-Weinstein and co-workers (1998) proposed an explanation to reconcile the autonomous versus inductive views. Populations of stably committed cells are randomly distributed in the mesoderm and cohabit with uncommitted cells. Differentiation of the latter depends upon communication between these two populations, regulated both by cell–cell interactions within the mesoderm and by influences from surrounding tissues. C. Combinatorial and Antagonistic Signals Emanating from the Overlying Ectoderm, the Neural Tube, and the Notochord Establish the Dorso-ventral Polarity Somites are surrounded by the ectoderm dorsally, the lateral plate laterally, the neural tube and notochord medially, and the endoderm ventrally. Ex-

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cept for the endoderm, which as yet has not been studied in this context in great detail, the putative influence of each of these structures has been examined. 1. The Notochord and Floor Plate Exert a Ventralizing Influence on the Somite Following on from the pioneering work of many laboratories (Hall, 1977), several groups have provided evidence that the notochord and floor plate are structures which are both necessary and sufficient for directing the differentiation of the adjacent paraxial mesoderm tissue toward the sclerotomal lineage. The patterning activities of these two structures on the ventral neural tube are strikingly similar. Notochord has been shown to specify the ventral part of the neural tube and more precisely to promote directly the differentiation of the floor plate and the bilaterally positioned motoneurons. The floor plate subsequently exhibits a patterning activity similar to that of the notochord on motoneurons (Placzek, 1995). In the case of the paraxial mesoderm, when a supernumerary notochord is grafted dorsal to the PSM, the dorsal compartment appears to be transformed into a ventral one: dorsal cells undergo an epithelio-mesenchymal transition such that no dermomyotome is formed (Pourquie´ et al., 1993); the expression domains of sclerotomal markers such as Pax-1 and Pax-9 are dorsally expanded, whereas dorsal markers (Pax-3 and Pax-7) are downregulated (Brand-Saberi et al., 1993; Dietrich et al., 1998, 1997; Goulding et al., 1994). Subsequently, extra cartilage is produced at the expense of axial musculature and dermis (Goulding et al., 1994; Pourquie´ et al., 1993). A floor plate can also convert the dermomyotome towards a sclerotomal fate (Brand-Saberi et al., 1993; Pourquie´ et al., 1993). In the loop-tail mouse mutant, as a consequence of overdifferentiation of the notochord and floor plate, the neural tube does not close dorsally and somites appear ventralized as evidenced by up-regulation of Pax-1 expression and down-regulation of Pax-3. This somite phenotype is likely to be due to increased signaling from the expanded ventralizing structures (Greene et al., 1998). In vitro experiments in which naive presomitic cells were cocultured with notochord or floor plate, led to similar conclusions about the ventralizing activities of these structures (Ebensperger et al., 1995; Fan and Tessier-Lavigne, 1994; Mu¨ller et al., 1996). Conversely, early ablation of the notochord along its whole length, such that floor plate is not induced, or ablation of both floor plate and notochord leads to an absence of Pax-1, Pax-9, and MyoD and to a ventral extension of the Pax-3 and Pax-7 expression domains in most of the region of the ablation (Dietrich et al., 1997; Goulding et al., 1994; Pownall et al., 1996; Teillet et al., 1998). If these embryos are allowed to develop further, no

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ventral derivatives like cartilage are observed, whereas dorsal ones like muscle and dermis are present (Van Straaten and Hekking, 1991). Analyses of mouse mutants in which notochord and floor plate development are affected support these results. In the truncate and Brachyury curtailed mutant embryos, the notochord does not develop: Pax-1 expression is never activated, whereas the whole somite expresses Pax-3 (Dietrich et al., 1993). In the Danforth’s short-tail and pintail mutant embryos, the notochord degenerates secondarily: Pax-1 expression is induced but is subsequently lost, whereas Pax-3 expression invades the whole somite (Asakura and Tapscott, 1998; Dietrich et al., 1993; Koseki et al., 1993). These data therefore suggest that notochord signaling is required for sclerotome development. If the ablation is performed after induction of the floor plate, then somites develop normally, indicating that the notochord and floor plate have redundant patterning activities (Dietrich et al., 1997; Ebensperger et al., 1995; see also Watterson et al., 1954). Thus, as in the case of dorso-ventral patterning of the neural tube, these two structures seem to exhibit strikingly similar dorso-ventral patterning activities on the somite. It is not yet clear whether they act synergistically or in a cascade in which the floor plate acts as a functional relay of the notochord. These observations have led to the proposal that dorsal differentiation in the somite reflects the existence of a default pathway for the somitic lineages (Pourquie´ et al., 1993). This hypothesis provides an alternative to the myogenic induction models, which will be discussed later. In the absence of notochord and floor plate signaling, somitic cells would differentiate toward the muscle and dermal lineages. This hypothesis implies that dorsal lineages require only permissive signals for their differentiation, whereas ventral lineages require instructive signaling provided by the floor plate and notochord. This default model is further supported by results from in vitro culture of dissociated chick epiblast in which 99% of the cells undergo myogenesis after 2 hr, suggesting that muscle differentiation could occur autonomously as early as epiblast stages before gastrulation. Such a precocious event would be prevented in vivo by tissular integrity and presumably cell communication (George-Weinstein et al., 1996). 2. The Dorsal Ectoderm and Neural Tube Deliver Dorsalizing Signals That Antagonize the Notochord/Floor Plate Influence Whether they are of an inductive or of a permissive nature, signals emanating from surrounding structures play a role in defining a dorsal identity. In vivo manipulations consisting of ablation or ectopic grafts and in vitro experiments using chick embryos have shown that the ectoderm and/or the dorsal neural tube are necessary and sufficient for the epithelialization of the

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somite and subsequently of the dermomyotome (Gallera, 1966; Palmeirim et al., 1998; Sosic et al., 1997; Spence et al., 1996). The individual roles of these structures have been investigated in vitro. When naive, non-dorso-ventrally determined murine presomitic explants are cultured with notochord or floor plate, these cells express a sclerotomal marker (Pax-1). When similar explants are cultured with dorsal ectoderm or dorsal neural tube, they express dermomyotomal markers such as Pax-3, Pax-7, and Sim-1 (Fan and Tessier-Lavigne, 1994; Maroto et al., 1997; Reshef et al., 1998). Moreover, if such coculture is performed with more mature somites that already have sclerotogenic properties, cartilage differentiation is strongly impeded (Kenny-Mobbs and Thorogood, 1987). In vivo experiments have aided our understanding of the complex network of interactions between the different structures. Independent ablation of ectoderm or dorsal neural tube does not affect the dorso-ventral polarity of somites (Dietrich et al., 1998, 1997; Hirano et al., 1995; Kuratani et al., 1994). In contrast, ectopic grafts of these tissues individually leads to ectopic activation of the dorsal marker Pax-3 and down-regulation of the ventral marker Pax-1 (Dietrich et al., 1997). Both structures have to be removed at the same time in order to obtain fully ventralized somites (Dietrich et al., 1997). In the open brain mutant mice, the neural tube lacks its dorsal domain leading to abnormal expression of Pax-3 and dermomyotome defects. Whether the ectoderm is affected in these mice is not known (Spo¨rle et al., 1996). Taken together, these results indicate that ectoderm and dorsal neural tube have necessary and redundant dorsalising activities that antagonize the ventralizing signals from notochord and floor plate. However, these dorsal signals can be overridden by ventral signals. For example, the graft of an ectopic notochord dorsal to the PSM leads to complete absence of muscles and dermis on the grafted side (Pourquie´ et al., 1993). Accordingly, the dorso-ventral inversion of both neural tube-notochord complex leads to a concomitant dorso-ventral inversion of this axis in the juxtaposed somites (Dietrich et al., 1997; Spence et al., 1996). A role for the dorsal neural tube in patterning the dorsal sclerotome, fated to form the neural arches, is also suggested by the analysis of the open brain mouse mutant (Spo¨rle et al., 1996). Furthermore, the notochord mutant, Danforth’s short-tail, exhibits defects solely in the ventral sclerotome resulting in abnormal vertebrae formation (Koseki et al., 1993). Therefore, antagonistic influences from the dorsal neural tube and notochord/ floor plate might also act to pattern the sclerotome itself along the dorsoventral axis. As differentiation proceeds, the ectoderm is also necessary for dermatome-specific gene expression such as gMHox (Kuratani et al., 1994). The dorsal neural tube is required for a subsequent differentiation step of this compartment, the epithelial-mesenchymal conversion of the dermatome precursors (Brill et al., 1995).

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In summary, it seems that notochord/floor plate, dorsal ectoderm, and neural tube exert antagonistic influences to pattern the somite along the dorso-ventral axis. It is also noteworthy that correct dorso-ventral patterning of the neural tube is necessary for proper dorso-ventral patterning of somites. Interestingly, the nature of the notochord, floor plate and neural tube signals seem to be diffusible, whereas the ectoderm effect requires cell–cell contact (Cooper, 1965; Fan and Tessier-Lavigne, 1994; Flower and Grobstein, 1967; Lash et al., 1957). 3. The Neural Tube/Ectoderm and Notochord/Floor Plate Are Involved in Myogenic Differentiation The precise means by which myogenesis is initiated and is then maintained in the somite has been and still is highly controversial, although it is now widely accepted that the neural tube, surface ectoderm, lateral plate, notochord, and floor plate are all involved to some extent in these processes. The discrepancies, which exist between the various studies addressing this issue, might be accounted for by the different strategies used and by variability in the age of the embryos and in the time of operation. The observation that myogenesis in the embryo is first initiated in the somitic territory apposed to the dorsal neural tube prompted several laboratories to test the myogenic-inducing role of the neural tube. Using in vitro approaches, it was demonstrated that the paraxial mesoderm and the newly formed somites require axial signaling to differentiate into muscle (KennyMobbs and Thorogood, 1987; Packard and Jacobson, 1976; Rong et al., 1992; Vivarelli and Cossu, 1986). More recently, this question was revisited by a number of labs and yielded a series of conflicting results. Buffinger and Stockdale (1994) reported that the neural tube but not the notochord could elicit myogenesis from segmental plate explants. In contrast, Stern and collaborators (1995), showed that, if the complex neural tube/notochord induces a 100% myogenic response in the paraxial mesoderm, the dorsal neural tube induces an 80% response, whereas the ventral neural tube with or without notochord triggers a 30% and a 10% myogenic response, respectively. A role for the dorsal neural tube is further confirmed by the phenotype of the open brain mutant mice in which muscle differentiation is impaired. These mice exhibit altered expression patterns of Pax-3 and myf-5 followed by severe defects in muscle formation (Spo¨rle et al., 1996). Mu¨nsterberg and Lassar (1995), for their part have shown that, in vitro, myogenic induction from segmental plate requires coculture with both notochord and neural tube, whereas more rostral somites require only neural tube signaling to differentiate. They thus propose a model involving a two step mechanism whereby notochord signaling would confer upon the somite the competence to respond to myogenic signals from the neural

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tube. In vivo manipulations have provided further evidence in favor of such a model. Caudal ablation of the notochord leads to failure of MyoD induction, but when the notochord is removed more anteriorly, the presence of the neural tube and ectoderm are sufficient to maintain MyoD expression (Pownall et al., 1996). A parallel situation is found in the case of the Danforth’s short-tail mutant mice: in anterior regions, a notochord develops but degenerates secondarily, whereas the somites at this level exhibit normal myogenesis raising the possibility that initial exposure to the notochord in the presence of the neural tube/floor plate and the ectoderm was sufficient to initiate myogenesis (Asakura and Tapscott, 1998). Conversely, in neural tube-ablated embryos, myogenic markers are induced but not maintained (Bober et al., 1994). The role of the notochord and floor plate in myogenic specification has been further addressed in vivo by performing ectopic grafts of these structures. In contrast to sclerotome induction by the notochord, no tissue adjacent to the paraxial mesoderm has been shown to exhibit such a clear inducing capacity in the recruitment of paraxial mesodermal cells toward the myogenic pathway. However, ectopic notochord grafts have been reported to induce short-term ectopic MyoD activation in the region of the graft in in vitro cultures of quail embryos (Pownall et al., 1996). In the majority of cases, notochord implantation leads to an inhibition of the myogenic program in nearby cells, although it promotes the process in cells located at a distance from the graft (Bober et al., 1994; Brand-Saberi et al., 1993; Dietrich et al., 1997; Pourquie´ et al., 1993). In most of the cases, these grafting experiments result in abnormal MyoD expression in the dermomyotome. This could result either from premature activation of the myogenic factors in territories which are normally fated to give rise to muscle, like the lateral somite, or from disrupted architecture of the myotome itself resulting in local concentrations of MyoD expressing cells. The only evidence for instructive signaling in myogenesis comes from dorsoventral inversion of the neural tube which can elicit formation of an ectopic dermomyotome from the ventral somite (Dietrich et al., 1997; Spence et al., 1996). In conclusion, both axial structures seem important for myogenesis, but the permissive or inductive nature of the signal emanating from these axial structures is still unclear. In vivo, if the notochord and neural tube are both removed, no epaxial myogenesis occurs because all medial somitic cells die within 24 hours (Teillet and Le Douarin, 1983). These structures thus provide at least a trophic support to somitic cells. Backgraft of either the neural tube or the notochord alone rescues myogenesis (Rong et al., 1992). Backgraft of the dorsal neural tube is far less efficient in rescuing MyoD expression indicating that most of this trophic (i.e., permissive) effect can be accounted for by the notochord and, to a lesser extent, by the ventral neural tube (Teillet et al., 1998). The role of the notochord in initializing the myogenic program

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is further questioned by the fact that ablation of the presumptive territory of the notochord and floor plate or simulatneous ablation of the neural tube and notochord does not prevent the subsequent differentiation of a few muscle cells (Teillet et al., 1998; Bober et al., 1994; Catala et al., 1996). Moreover, in the caudal region of a Danforth’s short-tail mutant, where neither notochord nor floor plate develop, weak myf-5 is nevertheless expressed in somites (Asakura and Tapscott, 1998). In summary, axial structures play an important role in myogenic differentiation, but this role seems to be more of a permissive (or trophic) nature. A third tissue, the dorsal ectoderm, may also regulate myogenesis. Culture of isolated lateral PSM (Cossu et al., 1995) or caudal somites (Reshef et al., 1998) with the overlying ectoderm leads to the activation of MyoD expression. Moreover, ablation and ectopic graft experiments led Dietrich and collaborators (1998; 1997; see also Hirano et al., 1995) to conclude that, as for sclerotome induction, ectoderm and dorsal neural tubes have redundant myogenic activities (see also Spence et al., 1996). Controversy also exists concerning the mechanisms involved in this process. Buffinger and Stockdale (1995), using transfilter cultures, proposed that the signals emanating from both the neural tube and the notochord are diffusible, whereas other groups have shown that neural tube and ectoderm require close contact with the somitic tissue to exert their effects (Cossu et al., 1996; Stern and Hauschka, 1995). To conclude, these conflicting results reflect the complexity of the regulation of the onset of myogenesis. Developmental time windows seem to be a crucial parameter; various surrounding structures send combinatorial and/or antagonizing signals to the paraxial mesoderm, which subsequently interprets these signals according to its own state of maturation. In addition, because some of these signals are diffusible, it is possible that different signaling threshold levels elicit different cellular responses. This morphogen-type response of somitic cells could account for the observation that the myotome forms at an intermediate position relative to dorsalizing (the ectoderm and the dorsal neural tube) and ventralizing signals (the notochord and the floor plate) (Dietrich et al., 1997). Alternatively, myogenesis could reflect a default pathway of the somitic cells and the role of these signals would then be to provide trophic support to somitic cells and to control precisely the timing of the onset of terminal differentiation (Pourquie´ et al., 1995, 1993; Teillet et al., 1998).

D. Shh and Wnt Proteins Are Key Players in the Segregation of the Different Somitic Lineages The previously described complexity of surrounding tissue interactions that pattern the somite is also reflected at the molecular level. The predominant

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role of two genes, both encoding secreted molecules, has been extensively examined in vivo and in vitro. Sonic hedgehog (Shh) is expressed in the notochord and floor plate along their entire length (Echelard et al., 1993; Krauss et al., 1993; Marti et al., 1995b; Riddle et al., 1993), whereas several Wnt proteins are expressed either in the dorsal neural tube (Wnt1, 3a, 4) or in the dorsal ectoderm (Wnt4, 6, 7a) (Hollyday et al., 1995; Marcelle et al., 1997; McMahon and Bradley, 1990; Parr et al., 1993; Tajbakhsh et al., 1998). Moreover, these factors have been shown to exhibit patterning activities in diverse systems (reviewed in Cadigan and Nusse, 1997; Tanabe and Jessell, 1996). Therefore, because these molecules are present at the right time and place, they appear to be good candidates for mediating somite patterning. 1. Shh Promotes Ventral Cell Fates whereas Wnt Proteins Specify Dorsal Lineages In vitro and in vivo experiments have provided evidence of a role for Shh in promoting sclerotome differentiation. Coculture of uncommitted paraxial mesoderm explants either with cells secreting Shh (Fan and TessierLavigne, 1994) or with purified Shh protein (Fan et al., 1995; Maroto et al., 1997; Mu¨nsterberg et al., 1995; Reshef et al., 1998) leads to activation of Pax-1 expression, as was obtained in similar cocultures with notochord. Accordingly, replacement of notochord and floor plate by a Shh secreting bead or by secreting cells or, conversely, antisense inhibition of quail Shh expression together demonstrated, in vivo, that Shh is important for maintaining and/or inducing Pax-1 expression (Borycki et al., 1998; Teillet et al., 1998). Furthermore, in an in vitro assay, Shh protein was shown to be capable of antagonizing the induction of dermomyotomal markers elicited by dorsal neural tube (Fan and Tessier-Lavigne, 1994; Maroto et al., 1997). In accordance with these studies, ectopic expression of Shh in the chick paraxial mesoderm using a retroviral expression system, revealed that Pax-1 expression expanded dorsally while Pax-3 was down-regulated and the dermatome was anatomically perturbed ( Johnson et al., 1994). An equivalent situation is found in the loop-tail mutant embryo in which Shh is expressed in an abnormally broad expression domain. The Pax-1 expression domain is expanded dorsally while Pax-3 expression diminishes (Greene et al., 1998). These experiments suggest that exposure to Shh can convert the dorsal somitic compartment toward ventral cell fates, and therefore can mimick the effect of the graft of a supernumary notochord in a dorsal position. However, several lines of evidence indicate that Shh does not account for all the ventralizing properties of the notochord. First, in the Shh retroviral overexpression studies in the chick, a strong MyoD up-regulation is ob-

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served in the overexpression domain. In contrast, notochord grafts lead to MyoD down-regulation in the close vicinity. Second, such a direct role for Shh in sclerotome induction appears unlikely because of the phenotype of Shh homozygous null mice. Sclerotome derivatives are, as expected, severly reduced, and the Pax-3 expression domain extends ventrally. Surprisingly, however, Pax-1 expression is transiently induced although, in agreement with the results obtained in vitro and in vivo, this expression is not maintained (Chiang et al., 1996). A similar situation is found in the case of the Danforth’s short-tail mutant (Asakura and Tapscott, 1998) or after ablation of the notochord/neural tube complex, both scenarios being that the embryos lose the source of Shh protein (Teillet et al., 1998): the somites which develop in the absence of Shh signaling activity exhibit weak Pax-1 expression. These data suggest that Shh is involved in the maintenance of Pax-1 expression rather than in its induction in the epithelial somite. In a reciprocal manner, retroviral ectopic expression of Wnt-1 in somites elicits the opposite response: Pax-1 and Pax-9 expression are lost and at E10, the axial skeleton is almost completely absent, suggesting that the development of the sclerotome has been severely impaired. Nevertheless, unlike the capacity of Shh to extend Pax-1 expression dorsally, Wnt-1 overexpression does not expand the Pax-3 expression domain ventrally (Capdevila et al., 1998). In vitro experiments support the model that different Wnt proteins mediate the dorsalizing role of the neural tube and ectoderm (Fan et al., 1997; Maroto et al., 1997). Uncommitted mesoderm cultured with cells expressing Wnt-1, 3a, 4, or 6 expresses dermomyotomal markers. Interposition of a nucleopore filter leads to a decrease in the effect of Wnt-4 and 6 activities, whereas the signaling potential of Wnt-1 and 3a is unaffected (Fab et al., 1997). Considering their expression patterns and their diffusibility, Wnt-1 and 3a appear to be good candidates for the secreted dorsalizing factor derived from the neural tube, whereas Wnt-4 and 6 meet all the necessary characteristics of the contact-dependent dorsalizing derived factor from the ectoderm. The phenotype of the compound mutant Wnt-1/Wnt-3a highlights the redundant influences of these two sets of Wnt genes in this process: the dorso-ventral polarity of their somites is normal, the Wnt proteins from the ectoderm possibly compensating for the loss of those from the neural tube (Ikeya and Takada, 1998). The antagonistic activities of Shh and Wnts has been demonstrated in sandwich cultures where a naive mesoderm explant is positioned between Shh-expressing cells on one side and Wnt-1-expressing cells on the other. This results in the induction of Pax-1 and Pax-3 expression in domains close to the ventralizing and dorsalizing signal sources, respectively. When Shh signaling is increased, the Pax-1 expression domain extends toward

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the Wnt-1 protein source whereas the Pax-3 expression domain is reduced (Fan et al., 1997). These data indicate that Wnt-1 and Shh signaling pattern the paraxial mesoderm in an antagonistic manner suggestive of a concentrationdependent action. Such a concentration-dependent action has been previously documented for patterning of the neural tube (Roelink et al., 1995). However, a strong caveat to this model arises from experiments performed by Teillet et al. (1998). When they replaced the neural tube/notochord complex with Shh expressing cells, a fairly normal dorso-ventral patterning of the somites was observed. Accordingly, surprisingly normal vertebrallike structures and associated muscles formed. These experiments suggest that Shh essentially rescues the death of somitic cells which normally occurs after ablation of the neural tube and notochord (Teillet and Le Douarin, 1983). If the choice to differentiate into muscle or into cartilage results from exposure to different levels of Shh and Wnt signals, then it seems very unlikely that grafting a pellet of cells producing an uncontrolled quantity of Shh would rescue an exact pattern of cartilage and muscles. Moreover in the rescue experiments, two different cell lines producing Shh were used—a stable transfectant and a cell line infected by a retrovirus-producing Shh. It is very unlikely that both lines produce the same amount of Shh, yet they elicit the same phenotype. These experiments therefore strongly argue in favor of a prepattern of the paraxial mesoderm along the dorso-ventral axis. The role of Shh and Wnt in this case would thus be permissive rather than instructive. Furthermore, studies aimed at addressing the diffusibility of these molecules demonstrate the complexity of the biochemistry of the two molecules. In vivo, Shh is autocatalytically processed into a soluble. C-terminus form and an N-terminus form (Shh-N), which becomes anchored in the membrane via a cholesterol bond (Lee et al., 1994). However, the N-terminus form alone accounts for all the signaling activity of the molecule both in patterning the mesoderm and the neural tube (Fan et al., 1995; Marti et al., 1995a; Mu¨nsterberg et al., 1995; Roelink et al., 1995). If the effects of Shh-N are direct, then the protein needs to be released from the membrane and to subsequently diffuse from the inducing tissue. Surprisingly however, cells transfected with full-length Shh exhibit a Pax-1-inducing activity, whereas the N-terminal form is not detected in the culture medium. In Drosophila, a gene necessary for Hedgehog diffusion has been characterized: Tout-velu is a fly homolog of a human tumor suppressor gene, Ext-1, that is also implicated in Hedgehog protein diffusion, thereby suggesting the existence of a conserved mechanism implicated in this process (Bellaiche et al., 1998). On the other hand, analysis of Shh signaling in limb development favors the idea that its long-range signaling activity is indirect

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and mediated by the release of a diffusible secondary signal (Yang et al., 1997). Similar to Shh, Wnt proteins have been shown to be poorly soluble and to bind tightly to proteoglycans at the cell surface (Smolich et al., 1993). In Drosophila, the Wg long-range activity appears to be mediated by active intracellular transport of the protein from one cell to the next (Gumbiner, 1998). Thus, whether Wnt-1 and Shh proteins act directly by diffusion or cellular transport or whether their range of activity is mediated in a relay fashion by triggering a second diffusible messenger is still unclear. 2. Combinatorial Signalling by Shh and Wnt Proteins Promote Myogenesis Although these molecules seem to have antagonistic effects on the specification of the dorsal-most and ventral-most derivatives of somites, they seem to specify cooperatively the intermediate compartment, the myotome. Coculture of unsegmented mesoderm or caudal somites in the presence of Shh alone does not result in muscle marker activation (Fan and TessierLavigne, 1994; Kos et al., 1998; Mu¨nsterberg et al., 1995; Tajbakhsh et al., 1998). In contrast, PSM culture in the presence of Wnt proteins (Wnt-1 or 4 or 5a or 6 or 7a) results in a low-level induction of myogenic precursors (Stern et al., 1995; Tajbakhsh et al., 1998). Therefore, it appears that Wnt proteins and Shh, when individually applied to naive mesoderm, are not fully sufficient to promote muscle development fully. Furthermore, in the case of the Shh knockout (Chiang et al., 1996) or as seen in the caudal region of the Danforth’s short-tail embryos (Asakura and Tapscott, 1998), expression of myf-5 is activated despite the lack of Shh protein. The phenotype of embryos lacking both Wnt-1 and Wnt-3a genes suggests that these genes are essential for the early myf-5 expression (Ikeya and Takada, 1998). Taken together, these results suggest that, as for the specification of the dorsal somitic compartment, the Wnt proteins produced by dorsal neural tube (Wnt-1, 3a, 4) and dorsal ectoderm (Wnt-4, 6, 7a) are both involved in myogenic specification. The situation is different when both signaling pathways are activated simultaneously. When PSM is cocultured in the presence of both Shh and Wnt proteins, MyoD and myf-5 expression are robustly up-regulated (Maroto et al., 1997; Mu¨nsterberg et al., 1995; Reshef et al., 1998; Tajbakhsh et al., 1998). When caudal somites or PSM from older embryos that have had a longer exposure to the influence of the notochord in situ are cocultured with Wnt proteins alone, myogenic precursors differentiate. Conversely, in experiments using antisense inhibition of Shh expression, MyoD is no longer induced in caudal somites (Borycki et al., 1998). There appears to be a difference in the requirement of continued Shh signaling in vivo and in

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vitro because the continuous presence of Shh in vitro is not required for the maintenance of MyoD expression in explants (Mu¨nsterberg et al., 1995), whereas antisense inhibition experiments suggest that in vivo maintenance of previously induced MyoD expression requires continuous Shh signaling (Borycki et al., 1998). The results of these experiments suggest that Shh and Wnt fulfill the criteria for the combinatorial signals from the notochord and the dorsal neural tube of the model proposed by Munsterberg and Lassar (1995). They also provide molecular evidence in favor of the model described earlier in which notochord signaling, mediated by Shh, and neural tube signaling, mediated by Wnt proteins, acts synergistically to promote somitic myogenesis. 3. How Is the Response to Axial Signaling Spatially and Temporally Regulated? Considering the previous data, two major comments can be made. First, it appears that, although Wnt and Shh signals are permanently present along the whole length of the antero-posterior axis, sclerotome formation at all embryonic stages is initiated only after somite segmentation; myogenesis is activated still later in somites from stage 11 HH onward (Borycki et al., 1998; Buffinger and Stockdale, 1994; Dietrich et al., 1997). Secondly, Shh signaling can induce both myotomal and sclerotomal lineages. So one might wonder how cells can interpret and respond to the same signal in such distinct ways. With respect to the regulation of the initation of myogenesis and chondrogenesis, different mechanisms may be involved: a. The Notch Signaling Pathway The Notch signaling pathway, which is well established as a regulator of cell competence (Artavanis-Tsakonas et al., 1999), is a candidate for regulating the state of responsiveness of the paraxial mesoderm. The Notch1 and Notch2 receptors are expressed in the mouse PSM and are down-regulated, along with their ligands Dll1 and Dll3, as somites form (Bettenhausen et al., 1995; del Amo et al., 1992; Dunwoodie et al., 1997; Hayashi et al., 1996; Henrique et al., 1995; Myat et al., 1996; Palmeirim et al., 1998; Reaume et al., 1992; Swiatek et al., 1994; Williams et al., 1995). Besides its role in the segmentation process discussed previously, forced activation of Notch signaling in Xenopus embryos or in mammalian cultures leads to a blockade of myogenesis ( Jarriault et al., 1998; Kato et al., 1997; Kopan et al., 1994; Lindsell et al., 1995; Luo et al., 1997; Nye et al., 1994; Shawber et al., 1996), possibly by directly interfering with the activity of MyoD (Kopan et al., 1994; Nye et al., 1994). Accordingly, in Drosophila, Notch has been implicated in regulation of myogenic differentiation (Baylies et al., 1998). As proposed by George-Weinstein et al.,

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(1998), Notch signaling pathway could therefore prevent precocious expression of the myogenic potential present from these early stages in the paraxial mesoderm. b. Controlling Expression of the Shh Receptor, Patched In avian embryos, regulation of the expression of Patched, the Shh receptor, could provide another molecular mechanism for temporally restricting PSM responsiveness. Patched is not expressed in the PSM, which may explain why neither sclerotome nor myotome specification occurs at this level despite the close vicinity of Shh-expressing tissues. Furthermore, in avian embryos younger than stage HH11, Patched is expressed exclusively in ventral somites, thereby explaining why myogenesis, unlike sclerotome formation, is not induced before stage 11 HH (Borycki et al., 1998). Similar mechanisms may occur in mammals because the two mouse Patched genes isolated so far exhibit expression patterns similar to their avian homologs during early development (Goodrich et al., 1996; Hahn et al., 1996; Motoyama et al., 1998). c. Presence of Inhibitors of the Wnt Pathway A third mechanism possibly involved in the regulation of mesodermal competence is based on Wnt signaling inhibitors. In Xenopus, one such molecule, Frzb, which is a soluble form of the Wnt receptor, Frizzled, has been shown to bind some Wnt proteins in vitro thereby inhibiting their activities (Leyns et al., 1997; Wang et al., 1997a, 1997b). Murine homologs of this gene have been cloned, and these are expressed in the PSM and the somites (Hoang et al., 1998). Cerberus, a member of the DAN family, has also been shown, in Xenopus, to inhibit Wnt signaling (Bouwmeester et al., 1996; Glinka et al., 1997). Its murine homolog is expressed in the PSM and the two last formed somites (Belo et al., 1997; Biben et al., 1998; Shawlot et al., 1998). The presence of these inhibitors of the Wnt pathway in the unsegmented mesoderm might provide a molecular mechanism to prevent precocious myogenic induction. Therefore activation of Notch signaling, restricted expression of the Shh receptor, or expression of Wnt signaling inhibitors, in the PSM, could account for the restricted responsiveness of the paraxial mesoderm to differentiation signals. With respect to Shh possibly eliciting myogenic or sclerotogenic differential responses, two mechanisms can be considered: controling expression of the effectors genes of the Shh pathway and cis-regulation of the myogenic factors activity. d. Controling Expression of the Effectors Genes of the Shh Pathway Gli and Gli2/4 are the vertebrate homologs of the fly transcription factor Cubitus interruptus (Ci) which is known to mediate the nuclear transcriptional

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activity of Hedgehog signaling in Drosophila. The Gli genes are differentially expressed in somites (Borycki et al., 1998), and, as was described earlier for Patched, the time of their activation is correlated with somite segmentation. The Gli expression domain is confined to the ventro-medial domain of somites and its activation is regulated by the notochord and Shh itself. Gli2/4 is initially expressed throughout the whole somite but is later restricted to its medio-dorsal domain from where MyoD expressing cells arise; its activation is regulated by the surface ectoderm (Borycki et al., 1998). The differential expression patterns of genes downstream of the Shh pathway might thus provide a mechanism by which Shh signaling may be differentially interpreted by different parts of the somite: Gli would mediate sclerotome specification while Gli2/4 may transduce the myogenic effect of Shh signaling. Obviously, to validate such a model, one needs to show that the different Gli genes transactivate different target genes and thereby evoke different cellular responses. Similar mechanisms may occur in mammals because the three Gli genes isolated so far exhibit expression patterns similar to their quail homologs during early development. Moreover, simple or double mutations of these genes lead to abnormal skeletal differentiation in the mutant mice (Hui et al., 1994; Mo et al., 1997; Platt et al., 1997). e. Cis-Regulation of the Myogenic Factors Activity The onset of myogenesis is controled by the bHLH transcription factors, MyoD and myf-5, which, at the trunk level, act downstream of the Pax-3 transcription factor (Maroto et al., 1997; Tajbakhsh et al., 1997). Both MyoD and myf-5 bind to the ubiquitous bHLH E12 protein, and subsequently this heterodimer transactivates muscle specific markers (reviewed in Weintraub et al., 1991). Two related murine bHLH genes, MTwist and Dermo-1, along with members of the Id family, encode HLH proteins that lack the basic domain essential for specific DNA binding (Benezra et al., 1990; Christy et al., 1991; Li et al., 1995; Riechmann et al., 1994; Sun et al., 1991; Wolf et al., 1991). These proteins exhibit similar but distinct HLH sequences, expression patterns, and activities. Unlike other bHLH proteins, they seem to inhibit myogenesis in a competitive manner by binding to E12 and/or the DNA binding site of myogenic factors but without transactivating muscle gene transcription (Benezra et al., 1990; Hebrok et al., 1994; Jen et al., 1992; Li et al., 1995; Melnikova and Christy, 1996; Spicer et al., 1996). Their transcripts are detected in the PSM, epithelial somites, and are later excluded from the myotome, but they persist in the sclerotome and dermomyotome (Evans and O’Brien, 1993; Fuchtbauer, 1995; Li et al., 1995; Stoetzel et al., 1995; Wang et al., 1992; Wolf et al., 1991). Therefore, they appear to be good candidates, first, to block precocious myogenic induction in the PSM and, second, to restrict myogenic induction to the myotome. Nevertheless,

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targeted disruption of the Mtwist gene does not lead to ectopic myogenesis (Chen and Behringer, 1995), although this may be due to functional redundancy. Another controversial aspect of a putative early inhibitory role for Mtwist is that, although its transcripts are present in the PSM and in epithelial somites, no protein could be detected in these tissues until somites differentiate (Gitelman, 1997). Another protein that is likely to play a role in the spatial restriction of myogenesis to the myotome compartment is I-mf, a mouse protein of unknown structure which is strongly expressed in the sclerotome (Chen et al., 1996). I-mf is capable of repressing myogenesis. This is likely to be due either to retaining the MyoD family members in the cytoplasm by binding to them and masking their nuclear localization signal or by interfering with their DNA binding activities (Chen et al., 1996). The differential expression domains of Dermo-1, Mtwist, Id proteins, and I-mf in the sclerotome and/ or dermomyotome may explain in part how somitic compartments respond differently to axial signaling mediated by Shh and Wnt. In summary, extrinsic cues emanating from structures surrounding the somites pattern paraxial mesoderm along the dorso-ventral axis, thereby segregating the different somitic lineages. Establishment of the mediolateral axis results in creating, within each of the three lineages, medial and lateral compartments, which also differ in terms of differentiation modalities and cell fate.

VI. Medio-lateral Polarity of the Avian Somite The third axis set up in the developing somite is the medio-lateral axis. It is discussed here in a separate section although it is apparent that the establishment of the dorso-ventral and medio-lateral axes are interconnected because these processes implicate similar molecular pathways and tissular interactions. Whereas the medio-lateral polarity of the dermomyotome compartments has been extensively examined, data are not yet clear about medio-lateral polarity within the sclerotome compartment.

A. Two Different Myogenic Lineages Arise from Somites 1. Two Myogenic Lineages Emerge from Different Somitic Compartments In 1895, Fischel described that muscles of the back arise from the myotome, and that the ventral edge of the dermomyotome generates abdominal mus-

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cles while migrating cells from the lateral somite colonize the limb bud to form appendicular muscles. This has been demonstrated by replacing medial chick halves with their quail counterparts and vice versa (Ordahl and Le Douarin, 1992; Ordahl and Williams, 1998). The axial skeleton and epaxial myogenic lineage, which will give rise to the paravertebral musculature, arise from the medial half of the somite, whereas the hypaxial myogenic lineage, which will generate muscles of the body wall (abdominal and intercostal muscles) and limbs, emerges from the lateral somitic moiety.

2. Muscular Development Is Different in the Epaxial and Hypaxial Lineages at the Brachial Level Cells of these two lineages undergo muscular differentiation according to different modalities. Epaxial precursors start differentiating in situ in the somite and activate myogenic genes as early as myotome formation (Buckingham, 1992; Pownall and Emerson, 1992). In contrast, differentiation of hypaxial precursors is delayed until they have migrated from the somite to their final location and activation of myogenic genes occurs 48 hours later than in the epaxial lineage, when muscle masses start to condense. Moreover, the epaxial muscles initially differentiate as mononuclear myocytes (Keynes and Stern, 1988), unlike the multinucleate hypaxial muscles (Rutz et al., 1982). Last, each somitic half seems to be differentially dependent upon trophic factors released from axial structures. Neural tube and notochord ablation are followed by necrosis within the myotome and sclerotome and the subsequent absence of axial muscles and skeleton. In contrast, the hypaxial musculature develops normally and therefore appears to be trophically independent of the axial structures (Asakura and Tapscott, 1998; Rong et al., 1992; Teillet et al., 1998; Teillet and Le Douarin, 1983). Another important difference between the two lineages is that epaxial muscles are innervated by the dorsal ramus of the spinal nerve, whereas the hypaxial muscles are innervated by the ventral ramus (Ordahl, 1993). Because hypaxial cells differentiate 48 hr later than their epaxial neighbors and only after having reached their final destination, the cellular readout of a lateral identity might consist of delaying overt differentiation of committed cells (Cossu et al., 1996; Pourquie´ et al., 1995, 1996) until they have reached their final destination. While these different developmental modalities occur at the limb level, the same is not strictly true at the thoracic and abdominal levels. In these regions, the hypaxial musculature forms from a ventral myotome, this being a mirror-image of the dorsal myotome (Christ et al., 1983) and differentiation modalities are similar, although delayed, to that of the epaxial lineage (Christ and Ordahl, 1995). Therefore the distinction between epaxial and hypaxial lineages is much less obvious at the thoracic

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and abdominal levels compared to the brachial ones (Ordahl and Williams, 1998). In conclusion, the two somitic lineages exhibit a different developmental history in terms of responsivity, origin, and fate. One might wonder whether these two compartments are specified differently because the cells that populate each of them are initially different or because an initially homogenous paraxial mesoderm becomes patterned along the medio-lateral axis by extrinsic cues. 3. Epaxial and Hypaxial Muscle Cells Have a Distinct Origin during Gastrulation To explore the first hypothesis, cell lineage analyses have been conducted. Marking cells in the Hensen’s node region with various techniques first led Pasteels (1937) and then Selleck and Stern (1991) to the conclusion that medial and lateral compartments in the somite originate from distinct populations of gastrulating cells. The medial compartment has been shown to derive from cells ingressing in the node itself while the progeny of cells ingressing at the level of the anterior primitive streak, just caudal to the node, populate the lateral compartment. This view was challenged by Schoenwolf and collaborators (1992b) who found that somitic cells derive exclusively from progenitors ingressing at the level of the primitive streak. Nevertheless, a more recent study confirms that medial and lateral compartments derive from cell populations emerging from different regions of the primitive streak which subsequently migrate along distinct routes (Psychoyos and Stern, 1996a). 4. Somitic Medio-lateral Patterning Is Under the Control of External Cues To investigate somite plasticity along the medio-lateral axis, switch-graft experiments have been conducted in which medial halves of the last-formed somites are replaced with lateral halves and vice versa (Ordahl and Le Douarin, 1992). The manipulated somites subsequently exhibit normal development. Even more dramatically, when PSM is grafted inside the lateral plate, it adopts a lateral plate fate (Tonegawa et al., 1997). Therefore, similar to what has been shown for the establishment of dorso-ventral polarity, external cues are involved in the establishment of somitic mediolateral polarity. Likewise, this polarity is not yet determined at the level of the PSM or the last-formed somites. Nevertheless, unlike the dorsoventral polarity, both intrinsic and extrinsic mechanisms seem to be involved in establishing the medio-lateral axis. Indeed, even though cells of the PSM are not yet allocated to a dorsal or a ventral compartment, they are already

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specified, albeit not fully committed, to a medial or a lateral fate. The apparent requirement for intrinsic and extrinsic cues could provide a proofreading process in which a medio-lateral axis would initially be outlined in a reversible manner according to cell lineage criteria and then confirmed according to environmental cues.

B. The Lateral Plate Exerts a Lateralizing Signal on the Somite which Is Antagonized by Medial Structures 1. Axial Structures and Lateral Plate Have Anatagonistic Patterning Influences Considering their positions relative to the somite, the influences of the lateral plate, neural tube, ectoderm, and notochord have been assessed in vivo by microsurgery (Dietrich et al., 1998; Pourquie´ et al., 1995; 1996; Pownall et al., 1996; Tonegawa et al., 1997). When the somite is isolated from the lateral plate by a slit, lateral dermomyotomal markers such as Pax-3 and Sim-1 are lost; concommitantly, MyoD expression is up-regulated as if the lateral identity was converted to a medial identity. Conversely, grafting lateral plate tissue medially to the somite converts the adjacent compartment to a lateral identity as evidenced by down-regulation of the medial markers MyoD and myf-5 and up-regulation of lateral markers (Dietrich et al., 1998; Pourquie´ et al., 1995; 1996). In vitro recombination experiments have led to similar conclusions (Cossu et al., 1996; Gamel et al., 1995; Mu¨ller et al., 1996). In addition, when paraxial mesoderm is grafted inside the lateral plate, it exhibits lateral plate characteristics: the paraxial marker Pax-3 is lost and the lateral plate marker cytokeratin is up-regulated (Tonegawa et al., 1997). These experiments demonstrated that the lateral plate produces a signal, which is required for the specification of lateral somitic identity. Further experiments have demonstrated that the axial structures could antagonize this lateralizing signal (Dietrich et al., 1998; Pourquie´ et al., 1996; Pownall et al., 1996). Ablation of the axial structures results in lateralization of the whole somite, whereas lateral grafts of these tissues lead to the conversion of the lateral somitic half toward a medial identity. Moreover, the presence of notochord and/or neural tube tissue in cultures of caudal somite explants promotes the expression of the medial marker Nkx3.1 (Kos et al., 1998). In addition, open brain mutant embryos which lack a differentiated dorsal neural tube exhibit a lack of epaxial muscles whereas hypaxial muscles develop normally (Spo¨rle et al., 1996). These results imply that medio-lateral polarity is established through the antagonistic influences of the lateral plate and axial structures. If the par-

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axial mesoderm is exposed to only one of these structures, the developing somites are either totally medial or totally lateral in character, depending on the identity of this structure. 2. Ectoderm Plays a Pivotal Role in Linking Dorso-ventral and Medio-lateral Axes The role of the ectoderm in the specification of the somite medio-lateral axis has been studied in vitro in the mouse embryo. In mouse explant cultures, the ectoderm can promote myogenesis in the lateral somite by first inducing expression of the MyoD gene. This induction appears to involve the Wnt7b gene (Tajbakhsh et al., 1998). In the medial domain, myogenesis requires the neural tube and involves myf-5 activation first probably in response to Wnt1 signalling (Cossu et al., 1996; Tajbakhsh et al., 1998). In the chick embryo, ectoderm ablation in a lateral region leads to conversion of the dorso-lateral quadrant toward a ventral fate, unlike the medial compartment, which maintains its dorsal fate. This is most likely due to the presence of the neural tube. Conversely, ventral grafts of ectoderm induce ectopic dorsal markers in both cases. Moreover, these markers are of a lateral nature if the lateral ectoderm is placed ventro-laterally (Dietrich et al., 1998). These results highlight the fact that ectoderm plays a pivotal role linking the dorso-ventral and medio-lateral axes: by itself, it specifies the dorsal compartment and, in combination with lateral plate, the dorsolateral identity. Consequently, because ablation of the notochord/floor plate complex does not lead to the ventral expansion of dorso-lateral markers (Dietrich et al., 1998), one might wonder what structure specifies the ventral aspect of the lateral quadrant. Endoderm would be a suitable candidate although such an issue is difficult to address because of the lack of specific ventro-lateral markers. Further experiments in which permeable or impermeable obstacles were implanted between the somite and either of the adjacent structures demonstrated the diffusible nature of the signals emitted by both the neural tube and the lateral plate (Pourquie´ et al., 1996). In addition, in vivo and in vitro studies (Mu¨ller et al., 1996; Tonegawa et al., 1997) suggest that the lateralizing signal acts in a concentration-dependent manner. In contrast, the influence of the ectoderm seems to require cell–cell contact (Dietrich et al., 1998). 3. Specification of Trunk Versus Limb Hypaxial Lineages As mentioned earlier, in addition to medio-lateral patterning, the lateral somite is also regionalized in terms of thoracic and abdominal levels versus

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brachial regions. Depending on the axial level, the hypaxial lineage exhibits various differentiation modalities and cell fates. At the thoracic and abdominal levels, hypaxial cells forming the trunk (abdominal and intercostal) muscles undergo an ‘‘epaxial-like’’ differentiation program unlike those at the limb level. Characterization of the chick homeobox gene Lbx1 provided a specific marker for the migrating hypaxial lineage (Dietrich et al., 1998; Mennerich et al., 1998) (i.e., for the cells that disperse and migrate out from somites to form the limb musculature). The limb-level identity of the hypaxial cells has been shown to be conferred by the lateral plate (Kieny, 1960). Indeed, grafts of limb-level lateral plate to another axial level result in the outgrowth of an extra-limb at this level (Kieny, 1960) and to the ectopic induction of Lbx1 (Dietrich et al., 1998). Whether Lbx1 behaves as a marker or as a key regulator in specifying this limb subpopulation has still to be determined. In mice, an analogous mechanism might operate. It has been proposed from analyses of homozygous null mutant mice that the myogenic factors MyoD and myf-5 differentially regulate the development of limb versus trunk skeletal muscle respectively (Kablar et al., 1997; Ordahl and Williams, 1998). Cells that first activate myf-5, whose expression depends upon neural tube signals, would differentiate into ‘‘epaxial-like’’ muscles (paraspinal, abdominal, and intercostal muscles), whereas those that first activate MyoD expression, which is regulated by the ectoderm, form limb muscles (Cossu et al., 1996; Kablar et al., 1997; Ordahl and Williams, 1998). From these data, it appears that three muscle populations arise from the combination of medio-lateral polarity and antero-posterior regionalization: the epaxial paraspinal muscles, the abdominal and intercostal muscles that have a hypaxial origin but an epaxial-like development and the hypaxial limb muscles.

C. BMPs, Wnt, Shh, and Noggin Interact to Pattern Somites along the Medio-lateral Axis 1. BMP4 Mediates the Lateral Plate Effect Via a Concentration-Dependent Mechanism The first molecule shown to play a role in medio-lateral patterning was BMP4, a member of the TGF-웁 family (Pourquie´ et al., 1996). This secreted molecule is highly expressed in the lateral plate and has been previously shown to play crucial roles in inductive interactions during embryogenesis in both vertebrates and invertebrates (Hogan, 1996). It therefore exhibits all the characteristics expected of a candidate molecule that would mediate lateral plate signaling. To test its role, BMP4-expressing cells were grafted

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medially between the neural tube and the somite. Expression of the dorsomedial marker MyoD and the ventro-medial marker Pax-1 were lost, whereas that of the lateral marker Sim-1 was extended medially as if the whole somite was lateralized (Dietrich et al., 1998; Hirsinger et al., 1997; Pourquie´ et al., 1996). Furthermore, in vitro experiments show that BMP4 blocks the medializing signal delivered by axial structures (Reshef et al., 1998). Unfortunately, the study of the BMP4 null mouse is not very informative because most mutant mice arrest development at the egg cylinder stage and therefore show little or no mesoderm formation (Winnier et al., 1995). Therefore BMP4 activity seems to account for most of the lateral plate effects described earlier. Whereas BMP4 can induce Sim1 expression in the lateral somite, it is unable to mimick the Lbx1 gene inducing activity of the limb-level lateral plate (Dietrich et al., 1998). This is, however, not surprising considering its homogenous expression pattern along the rostro-caudal axis. Candidate molecules such as the SGF/HGF also failed to induce ectopic Lbx1 expression in the paraxial mesoderm (Mennerich et al., 1998). To define further the mechanistic details of lateralization, a quantitative analysis of BMP4 effects has been conducted. When COS cells, strongly expressing BMP4, are grafted into the unsegmented mesoderm, this tissue adopts a lateral plate identity (i.e., Pax-3 and Sim-1 expression are lost) and cytokeratin expression, a marker of the lateral plate, is activated; when BMP4 cells are more and more diluted with control cells, the paraxial mesoderm is less and less lateralized. The minimal perturbation corresponds to a somite having two lateral halves (Tonegawa et al., 1997). These results show that BMP4 acts in a concentration-dependent manner. To decide whether BMP4 acts as a true morphogen, it must in addition exhibit a direct long-range effect. Xenopus studies suggest that BMP4 acts directly but diffuses poorly (Dosch et al., 1997; Jones et al., 1996, 1998). Its Drosophila countepart, Decapentaplegic, is however highly diffusible (Lecuit et al., 1996; Nellen et al., 1996). This issue is therefore still open to debate. 2. Noggin Antagonizes the BMP4 Lateralizing Effect In addition to a BMP4 concentration-dependent effect, previous studies have shown the necessity of a medializing signal, antagonizing BMP4 lateralizing signal, in order to specify a medial and a lateral identity within the somite. A molecular candidate to mediate this medializing signal is Noggin. Noggin is a secreted protein (Smith and Harland, 1992), which has been shown to bind to BMP4 and thereby prevent it from binding to its receptor (Zimmerman et al., 1996). In this way, it acts as a BMP4 antagonist. Among other sites, it is dynamically expressed in the paraxial mesoderm and its presumptive territory in locations compatible with a putative role as a

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BMP4 antagonist (Capdevila and Johnson, 1998; Hirsinger et al., 1997; Marcelle et al., 1997; Tonegawa and Takahashi, 1998). To examine whether Noggin blocks the BMP4 lateralizing signal in somites, the protein has been ectopically expressed in the paraxial mesoderm either by grafting Noggin expressing cells (Hirsinger et al., 1997; Marcelle et al., 1997; Reshef et al., 1998; Tonegawa and Takahashi, 1998) or by retroviral-mediated overexpression (Capdevila and Johnson, 1998). In all cases, ectopic Noggin expression converts the lateral compartment into a medial one, as evidenced by the down-regulation of Sim-1 and the lateral expansion of MyoD and Pax-1 expression domains. At an earlier developmental stage, when Noggin-expressing cells are implanted in the presumptive region of the lateral plate, ectopic somites are formed, suggesting that the lateral plate fate has been converted to a paraxial mesoderm fate by blocking the BMP4-lateralizing signal (Tonegawa and Takahashi, 1998). Conversely, when Noggin expression is abolished in homozygous null mice, the somitic phenotype shows a medial extension of the Sim-1 expression domain (McMahon et al., 1998). These data indicate that the whole somite becomes lateralized. These results taken together with in vitro studies (Reshef et al., 1998) are consistent with the hypothesis that Noggin acts as an antagonist of BMP4 signaling in somite differentiation. Initially, the antagonistic interactions between Noggin and BMP4 lead to the definition of the boundary between paraxial and lateral mesoderm; as development proceeds, this mechanism is progressively involved in specifying lateral and medial compartments within the somite. Unlike the case with BMP4, it is well-established that in Xenopus Noggin diffusion is long range and its effect is direct (Dosch et al., 1997; Jones and Smith, 1998). Therefore the BMP4 activity gradient is more likely to be achieved by the long range effects of its antagonists than by diffusion of the BMP4 protein itself ( Jones and Smith, 1998). To date, it is unclear whether Noggin has activities independent of this antagonistic role. No Noggin receptor has been identified so far. Results obtained in vitro are contradictory since, Reshef and colleagues (1998) find that naive somites cultured with Noggin are not driven toward any particular differentiation pathways, whereas McMahon and collaborators (1998) argue that Noggin can induce Pax-1 expression in presomitic explants. Different experimental procedures could account for these discrepancies. As described earlier, PSM is temporally delayed in its response to the prior presence of differentiation signals. As previously mentioned, activation of Notch signaling, regulation of the Shh pathway by its downstream genes, or presence of Wnt antagonists are strong candidates for the possible mechanisms that could account for this phenomenon. Blockade of BMP4 signal by Noggin might be another factor. Indeed, at the level of unseg-

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mented mesoderm, Noggin is expressed in the lateral edge of the paraxial mesoderm, in apposition to the BMP4 expression domain (Hirsinger et al., unpublished data; Capdevila and Johnson, 1998; Marcelle et al., 1997; Tonegawa and Takahashi, 1998). Therefore, Noggin activity in this location could prevent the PSM from responding to signals emanating from the lateral plate and additionally act to maintain the boundary between paraxial mesoderm and lateral mesoderm (Tonegawa and Takahashi, 1998). In addition, it has been shown in Xenopus and mouse that the DAN family member Cerberus, in addition to antagonising Wnt signaling, blocks BMP4 activity, by binding to it (Biben et al., 1998; Hsu et al., 1998). Murine cerberus is found in the PSM and the two last formed somites (Belo et al., 1997; Biben et al., 1998; Shawlot et al., 1998). Therefore, presence of Noggin and Cerberus in these locations could provide an additional molecular mechanism for the delayed response of the PSM toward environmental signals. 3. Wnt and Shh Pathways Antagonize BMP4 Signaling Via Noggin The previous results suggest that Noggin plays a role in differentiation events, such as myogenesis and sclerogenesis, that are controlled by the Wnt and Shh pathways. Therefore the possibility that these two factors might also antagonize BMP4 has been investigated. Furthermore, these genes are medially expressed, in the dorsal neural tube/ectoderm and in the notochord/floor plate, respectively. Retroviral-mediated ectopic expression of Shh (Capdevila et al., 1998; Johnson et al., 1994) and Wnt-1 (Capdevila et al., 1998; Johnson et al., 1994) or grafts of Shh or Wnt-1 expressing cells (Hirsinger et al., 1997; Marcelle et al., 1997) alter the normal patterning along the medio-lateral axis. In all cases, medial markers (MyoD, Wnt-11) are laterally expanded while the lateral marker Sim-1 is lost. Accordingly, the phenotypes of embryos mutant either for Shh or for Wnt-1/Wnt-3a support these data (Chiang et al., 1996; Ikeya and Takada, 1998): the lateral somitic compartment is expanded at the expense of the medial compartment. Therefore, Shh and Wnt-1 appear to exhibit the same antagonistic activities as the structures in which they are expressed and as Noggin itself. As a consequence, one might wonder whether these factors exert their effect independently of Noggin or if they are part of the same pathway. Grafts of Shh expressing cells was shown to induce Noggin expression ectopically whereas cells producing Wnt-1 mimick the influence of the dorsal neural tube on Noggin expression (Hirsinger et al., 1997). Furthermore, Wnt-1 protein is capable of inducing Noggin expression in naive somites in vitro (Reshef et al., 1998). Accordingly, the major site of Noggin

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expression in the somite is the medial dermomyotome, in close vicinity to the Wnt-1 expressing domain in the dorsal neural tube; Noggin is also transiently found colocalized with Shh in the notochord and floor plate. Therefore Noggin may act downstream of both the Wnt and Shh pathways where it functions to antagonize the BMP4 lateralizing signal. In parallel, other lines of evidence suggest that Noggin and Shh might also act synergistically in the specification of the sclerotomal lineage (McMahon et al., 1998). Shh is able to activate Pax-1 expression in cultured PSM; however, in conjunction with Noggin, lower levels of Shh are required to obtain the same effect. Some aspects of the Noggin mutant mouse phenotype provide further support for this hypothesis (McMahon et al., 1998). In conclusion, Noggin, present in the medial dermomyotome, seems to be downstream of both the axial Shh and Wnt-1 pathways and appears to act synergistically with Shh to block the effect of lateral plate-derived BMP4 signaling. However, Noggin is not the only BMP4 antagonist expressed in somites. A Follistatin-related gene, Flik, has been cloned in chick (Patel et al., 1996) and is expressed in the medial dermomyotome (Amthor et al., 1996). Similar to Noggin, Flik expression is regulated by the neural tube (Amthor et al., 1996). Because Follistatin has been shown to bind to BMP4 and thereby block its activity (Fainsod et al., 1997), a similar role might be expected for Flik. Another likely candidate is the secreted protein Chordin, which is expressed in the notochord (Sasai et al., 1994) and has been shown to diffuse long distances ( Jones and Smith, 1998). Like Noggin and Follistatin, it acts to block BMP4 activity by binding to it (Piccolo et al., 1996). These secreted molecules thus exhibit features compatible with a role in defining a medial somitic identity by blocking a BMP4 lateralizing signal. 4. Dorsal Neural Tube-Derived BMP4 Can Indirectly Promote Medial Somitic Identity BMP4 expression in the dorsal neural tube has been shown to play a role in the dorso-ventral patterning of the neural tube (Liem et al., 1995), but one might question how the presence of BMP4 in this location can be reconciled with its lateralizing activity on the somite. Firstly, the somite and the neural tube are each surrounded by a heparin-rich basal lamina, which is likely to inhibit movement of heparin binding factors like BMP4. Secondly, Noggin is expressed in the adjacent medial dermomyotome, adjacent to the dorsal neural tube, and is thus correctly spatially located for it to block any BMP4 effect on the paraxial mesoderm. Third, Marcelle and collaborators (1997) have shown in vivo that injecting cells, that express BMP4 into the neural tube, can activate the ectopic expression of Wnt-1 in the dorsal neural tube. This activation then positively regulates the

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expression of a medial dermomyotomal marker, Wnt-11 (Marcelle et al., 1997) and also promotes Noggin expression in that region (Hirsinger et al., 1997). Thus, in an indirect manner, BMP4 could promote the specification of a medial compartment while simultaneously being prevented from directly lateralizing the somite by the expression of Noggin in this region. 5. The Origin of Ribs and the Lateral Sclerotome It is noteworthy that the medio-lateral polarity of the dermomyotome has been extensively studied but little is known about the existence of such a regionalization within the sclerotome. For example, the existence of a lateral sclerotome is not yet well established, although this compartment appears to be regionalized in terms of developmental fate and gene expression patterns. Its medial region, expressing high levels of Pax-1, gives rise to the vertebral body and intervertebral discs, whereas the neural arches and proximal ribs probably derive from the lateral sclerotome, which expresses Sim-1 and high levels of Pax-9 (Balling et al., 1996; Pourquie´ et al., 1996). However, whereas Sim-1 expression is promoted by lateralizing cues and is down-regulated by medializing signals (Pourquie´ et al., 1996), as would be expected for a marker of the lateral compartment, both Pax genes are positively regulated by medializing signals and negatively regulated by lateralizing ones (Hirsinger et al., 1997; Mu¨ller et al., 1996), thereby suggesting that they both are markers of the medial compartment. Fate mapping of half somite derivatives in the chick have led to the suggestion that ribs are derived from the lateral somitic half (Christ and Ordahl, 1995). Surprisingly, recent fate mapping experiments have demonstrated that in chick the distal rib originates in the dermomyotome (Kato and Aoyama, 1998). Ablation of progressively more lateral regions of the dermomyotome affects progressively more distal parts of the ribs. Therefore ribs and intercostal muscles that interact in the adult animal derive from the same embryological structure, the dermomyotome, whereas the rest of the axial skeleton derives from the sclerotome. Accordingly, myf-5 knockout mice exhibit defects both in distal rib formation and in the development of intercostal muscles (Braun et al., 1992), suggesting that, in addition to their closely related anatomical origin, their subsequent development might be regulated by the same molecular cascades.

VII. Conclusion Taking into account all these results, it appears that tissues surrounding the somites build a complex network of synergistic and antagonistic interactions

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leading to the definition of eight major somitic compartments (Fig. 2). Each compartment gives rise to distinct sublineages such as epaxial and hypaxial muscles, dermis, anterior and posterior part of the vertebra, and so on. Therefore, a cell reads its position by assessing and integrating the different patterning signals. Interpreting these external cues in the context of its intrinsic characteristics, this cell will adopt a particular pathway of differentiation. Surrounding structures and their signals are crucial for somitic differentiation. However, the exact cellular mechanisms employed are still controversial: are these signals inductive or only permissive, allowing the expansion of previously committed cells as suggested by George-Weinstein and collaborators (1998)? Alternatively, both types of mechanisms may be in operation because they are not theoretically mutually exclusive.

FIG. 2 Schematic representation of the interactions between signaling molecules involved in somite medio-lateral and dorso-ventral patterning. Because the antero-posterior axis is not taken into account on this scheme, only four of the eight somitic compartments are shown. See text. Ec, ectoderm; En, endoderm; LP, lateral plate; LS, lateral somite; MS, medial somite.

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One form of permissive signal may be, for example, to provide trophic support. It is known that the neural tube/notochord complex delivers trophic signals necessary for the survival of the medial somite and for the maintenance and amplification of myotomal and sclerotomal cells (Asakura and Tapscott, 1998; Rong et al., 1992; Strudel, 1955; Teillet et al., 1998; Teillet and Le Douarin, 1983). When the complex is ablated, Pax-1 and MyoD expression seems nevertheless transiently induced, questioning any role for these structures in the initiation of differentiation markers (Teillet et al., 1998). Ectoderm can compensate for the loss of axial tissue in the induction of MyoD (Dietrich et al., 1997), but the case of Pax-1 remains unresolved. It has recently been shown that the neural tube/notochord trophic effect is mediated by Shh (Teillet et al., 1998). In addition, Shh has been shown to induce cell proliferation in the retina and lung, in the PSM, and in committed skeletal muscles (Bellusci et al., 1997; Duprez et al., 1998; Fan et al., 1995; Jensen and Wallace, 1997). However, the Shh null mice which show transient expression of Pax-1 suggest that this signal, while mediating some notochordal properties such as trophic and maintenance activities, is not able to substitute for all the activities. The question is now to decipher whether the activity of this molecule in the notochord and the floor plate can be separated from that of other signals in these tissues and whether it can exert, in addition, any specific inductive properties.

Acknowledgments We thank Drs Kim Dale, Mike McGrew, Miguel Maroto, and Pr. Monte Westerfield for their helpful comments and their critical reading of the manuscript. Financial support for our own work on somitogenesis was provided by the Centre National de la Recherche Scientifique (CNRS), the Association Franc¸aise contre les myopathies (AFM), the Association pour la Recherche contre le Cancer (ARC), and the Fondation pour la Recherche Me´dicale (FRM).

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Morphogenesis of the Eggshell in Drosophila Gail L. Waring Department of Biology, Marquette University, Milwaukee, WI 53201-1881

The Drosophila eggshell is a specialized extracellular matrix that forms between the oocyte and overlaying somatic follicle cells during the latter stages of oogenesis. Largely proteinaceous, the eggshell is a highly organized multilayered structure with regional specializations designed to perform a variety of functions. Production of a functional eggshell features: (1) the differentiation of subsets of follicle cells in response to ovarian signals, (2) directed migrations of the follicle cells within the developing egg chamber, (3) expression of eggshell structural genes by the follicle cells in a defined temporal and spatial order, (4) postdepositional modifications of the eggshell proteins including several temporally regulated proteolytic cleavage events, and (5) regulated trafficking of several eggshell proteins in the assembling structure. By exploiting the genetic advantages of Drosophila and using evolution as a guide, the eggshell provides an excellent experimental system to study, in vivo, molecular mechanisms used to regulate protein–protein interactions throughout the assembly of a complex extracellular architecture in a developing organism. KEY WORDS: Oogenesis, Drosophila eggshell, Supramolecular assembly, Extracellular trafficking, Follicle cells. 䊚 2000 Academic Press.

I. Introduction The formation of functional three-dimensional architectures is a fundamental process that occurs in all cells and developing organisms. The Drosophila eggshell is a large extracellular architecture which forms during late oogenesis between the oocyte and overlying follicle cells. Over the course of approximately 30 hr, the follicle cells secrete eggshell proteins into the extracellular milieu, where they assemble into a highly organized structure International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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featuring both radial and regional complexity. Although the ultrastructure of the assembling and mature eggshell has been extensively studied and reviewed (Margaritis, 1985; Margaritis and Mazzini 1998), little is known about the molecular mechanisms that underlie the assembly process. Over the past 20 years, several eggshell genes have been cloned and characterized. It is clear from these studies that eggshell genes are activated and repressed in a well-defined temporal order. However, it is not clear whether, or to what extent, the timing of the appearances of the eggshell proteins controls their interactions extracellularly. Although it has been appreciated for some time that regional specializations in the eggshell are related to different subsets of follicle cells, only recently have the molecular signals that lead to subdivisions within the follicle cell epithelial begun to be deciphered. Although eggshell morphology is distinct in regions that underlie different follicle cell populations, molecular differentiation in these regions has proven elusive. With few exceptions all follicle cell subpopulations appear to express the same array of major eggshell genes. Until recently, assembly of the multilayered body of the eggshell was limited to the realm of electron microscopic autoradiography. The recent acquisition of specific antibodies has allowed the fates of individual eggshell proteins to be monitored and localized in the assembling and completed structure. These studies have revealed complex extracellular processing events and unanticipated dynamics and molecular trafficking between the different eggshell layers. This review begins with a brief overview of eggshell morphology to convey the structural complexity that emerges from the assembly process. The morphology section is followed by a section considering the biochemical complexity of the eggshell as revealed by one- and two-dimensional gel analyses of purified eggshells from wild-type females and mutant females exhibiting aberrant eggshell morphology. Evolutionarily conserved structural features and amino acid sequence motifs of the major eggshell proteins are highlighted in a section on eggshell genes. A discussion of postdepositional processing and proteolytic cleavage of several eggshell proteins follows. The last section on assembly begins with the elaboration of the specialized regions with an emphasis on the molecular signaling that occurs between the oocyte and follicle cells that leads to the differentiation and proper positioning of the follicle cells that produce specializations in the anterior region of the eggshell. Production and stabilization of the multilayered eggshell concludes the review. Highlighted in this section are the intricate interlayer interactions and regulated molecular trafficking that appear to be a central feature of eggshell assembly in Drosophila.

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II. Morphology of the Eggshell A. Egg Chamber Development The eggshell is produced by epithelial follicle cells during the latter stages of oogenesis. Egg chambers, consisting of three cell types (nurse cells, oocyte, and follicle cells) are formed in the germarium at the tip of the ovary. In the germarium, 16 interconnected germ-line cells (15 nurse cells and 1 oocyte) become enveloped by a layer of follicle cells. Egg chambers leave the germarium as stage 2 egg chambers with the oocyte occupying the most posterior position in the germ cell cluster. Fourteen stages of oogenesis have been described based on morphological criteria, including egg chamber size, the proportion of the egg chamber occupied by the oocyte, the position of the follicle cells, and the appearance of the eggshell coverings (King, 1970; Spradling, 1993). During stages 2–7, the egg chamber increases in size as the nurse cell nuclei endoreplicate and the follicle cells divide. During the vitellogenic stages, 8–10, the follicular epithelium rearranges, and the oocyte enlarges via the stage-specific uptake of yolk proteins produced by the fat body and follicle cells. By stage 10, the yolkfilled oocyte occupies one-half of the volume of the egg chamber. At the end of stage 10, the cytoplasmic contents of the nurse cells are squeezed into the oocyte through the interconnecting cytoplasmic bridges; hence, in stage 11 egg chambers the oocyte is larger than the nurse cell complex. Stages 12–14 are distinguished by the appearances of the egg chambers at the anterior ends. In stage 12 egg chambers, nurse cell nuclei are present, and the elaboration of specialized respiratory structures, the dorsal appendages, begins. The appendages become prominent in stage 13 egg chambers; by stage 14, remnants of the nurse cell nuclei have disappeared and the dorsal appendages are complete. Figure 1 shows morphological changes associated with egg chamber development.

B. Radial Complexity The eggshell is a complex multilayered structure which exhibits regional and radial complexity. The specialized regions include the micropyle, dorsal appendages, operculum, and collar at the anterior end (Fig. 2B), the main body, and the posterior pole. Radial complexity is most conspicuous in the main body region where five morphologically distinct layers have been identified. The main body contains three major proteinaceous layers, an oocyte proximal vitelline membrane, an inner chorionic layer (ICL), and

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FIG. 1 Stages of egg chamber development. The egg chamber consists of 15 nurse cells (nc) and the oocyte (oo) surrounded by somatic follicle cells (fc). The small, interconnected egg chambers shown in panel (a) are bright field images of stages 2 and 3 egg chambers emerging from the germarium (G). Panel (b) shows the tip of a feulgen-stained ovary highlighting the polyploid nurse cell nuclei in stages 2–6 egg chambers. Yolk deposition begins in stage 8, continues through stage 9 (c), and terminates at the end of stage 10. Panel (d) shows a stage 10 egg chamber with the nurse cells occupying the anterior half of the egg chamber and the yolk-filled oocyte surrounded by a layer of columnar follicle cells occupying the posterior half. During stages 12–14, the dorsal appendages (da) are synthesized at the anterior end of the oocyte as the nurse cell nuclei degrade. In late stage 14, the follicle cells die and are sloughed off, the dorsal appendages separate as the paddle-like appearance of their tips becomes visible. (e) stage 12, (f ) stage 13, (g) early stage 14, and (h) a mature stage 14 egg chamber.

an outer endochorion (Fig. 2A). A lipid wax layer forms between the vitelline membrane and ICL and an ill-defined outermost nonproteinaceous exochorion layer completes the eggshell (Fig. 2A). The ICL, endochorion, and exochorion are collectively termed the chorion. The vitelline membrane is the first layer of the eggshell produced by the follicular epithelium. Its deposition begins in stage 9 and is completed by late stage 10. The vitelline membrane is deposited as a discontinuous layer that is traversed by micropillar projections from both the follicle cells and oocyte. During late stage 10, the micropillar projections are withdrawn, and the ‘‘vitelline bodies’’ fuse to form a continuous, morphologically homogeneous layer around the oocyte. The vitelline membrane is formed as a

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FIG. 2 Eggshell morphology. (A) Transmission electron micrograph showing the multilayered eggshell as seen in the central main body region: ex, exochorion; oe, outer endochorion; ie, inner endochorion; icl, inner chorionic layer; vm, vitelline membrane. (B) Scanning electron micrograph of the anterior region of the eggshell: op, operculum; c, collar; m, micropyle; da, dorsal appendage; ms, mainshell.

1.7 애m thick layer, but thins as the oocyte grows during the terminal stages of oogenesis to a uniform 0.3 애m thick layer (Margaritis, 1985). Deposition of the wax layer begins in late stage 10 with the accumulation of lipid-filled vesicles between the follicle cells and vitelline membrane. Synthesis and secretion of the lipid vesicles continues through stage 12. As the lipid vesicles accumulate on the surface of the vitelline membrane, they

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take on a flat appearance. During stage 14, the vesicles are compressed into three to four planes of overlapping plaques which create a waterimpermeable layer between the vitelline membrane and chorion. Compression is a consequence of opposing forces created by: (1) the late swelling of the oocyte due to water uptake and (2) the mechanical resistance provided by the formation of the ICL and endochorion layers that surround the wax layer. The compressed wax layer, less than 5 nm in cross section, protects the embryo from desiccation (Margaritis et al., 1980; Papassideri et al., 1993). Fine structural analysis of the ICL shows it first appears as a 10-nm electron-dense layer complexed with the endochorion of stage 12 egg chambers (Papassideri and Margaritis, 1996). As the egg chambers mature, the ICL gradually thickens to a 30-nm bilamellar complex which remains associated with the inner endochorion through stage 13. At the end of choriogenesis (stage 14), the ICL detaches from the endochorion and assumes its final thickness (40 nm) and unique crystalline substructure. Three-dimensional reconstructions from negatively stained EM images indicate that the structural units within the cystalline ICL consist of two types of subunits, each 3 nm in diameter. The subunits appear to be grouped into an octamer as four dimer pairs. Margaritis et al. (1984) suggest each dimer pair consists of polypeptides approximately 40 kDa in size, whereas Akey et al. (1987) suggest a dimer of 150 kDa and an octamer of 600 kDa. It has been postulated that the ICL aids in morphogenesis of the wax layer (Papassideri and Margaritis, 1996) or may serve as a bridge between the vitelline membrane/wax layer and the endochorion floor which allows the larva to shed the inner vitelline membrane/wax layer during hatching (Akey et al. 1987). Ultrastructural analyses of stage 14 egg chambers and isolated endochorions reveal the tripartite nature of this layer (Margaritis et al., 1980). A thin, fenestrated inner layer or floor is separated from a thick outer roof layer by vertical pillars creating cavities that facilitate gas exchange. The continuous outer roof network displays ridges. It defines the borders of the follicle cell imprints. The imprints in the main body of the eggshell are fairly uniform, with those on the dorsal side being more elongate than those found ventrally. Freeze-fractured views of endochorion components reveal globular structures interconnected via fine fibrils (Margaritis and Mazzini, 1998).

C. Regional Complexity Like most insect eggs, several regional specializations are apparent on the surface of Drosophila eggs. The most prominent specialized structures include the micropyle, operculum, and dorsal respiratory appendages at the anterior end of the egg (Fig. 2B). The micropyle is a specialized structure

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consisting of chorion and vitelline membrane that protrudes from the eggshell and allows sperm to gain access to the oocyte plasma membrane. The physical dimensions of the micropyle (20 애m in length, 10 애m in width at its base, with a central pore of 0.8 애m) are believed to provide a block to polyspermy by restricting passage to a single sperm (Perotti, 1974). The operculum is a specialized region that has evolved to facilitate hatching of the larva at the end of embryogenesis. The lateral and anterior margins of the operculum, the collar region, mark an abrupt transition to the follicle cell imprints found on the ventral side of the main shell. By serving as gills, the dorsal respiratory appendages on either side of the midline allow the eggs to respire when they are submerged under water. Long (300-애m) cylindrical structures with paddle-shaped tips, the appendages consist largely of a network of modified pillars with air spaces. The ventral surfaces of the appendages contain a fine network akin to the roof network in the main body, whereas the dorsal surface consists of isolated, porous plaques. Extensive reviews of the ultrastructure of the eggshell layers and specialized regions have been published by Margaritis (1985) and, more recently, by Margaritis and Mazzini (1998).

III. Identification of Eggshell Proteins A. Biochemical Analysis Although insoluble in the mature eggshell, eggshell proteins can be solubilized with denaturing SDS containing buffers well into stage 14 of egg chamber development. Several proteins have been electrophoretically resolved from eggshells purified from stage 10 and stage 14 egg chambers. Egg chamber proteins that become selectively enriched in purified eggshells are considered putative eggshell proteins. Other identifying features include stage-specific synthesis by the follicle cells during the period of eggshell deposition and ready labeling with radioactive proline, a major constituent of both the vitelline membrane (18%) and the chorion (11%) (Kafatos et al., 1977). By differential sieving and centrifugation, Petri et al. (1976) mass-isolated stage 14 egg chambers from adult flies. After vigorous shaking in distilled water, the outer chorion layers were separated from the remaining vitelline membrane-bound oocyte. The chorion fraction was solubilized in strong denaturing reagents and six major proteins, detected by Coomassie blue stain, were resolved on SDS-urea gels. To facilitate detection and provide greater resolution, Waring and Mahowald (1979) used two-dimensional NEPGHE gels to fractionate eggshell

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proteins which were radiolabeled in vivo during the choriogenic stages (11–14). Stage 14 egg chambers were disrupted by sonication in a tricine based buffer containing 1% Triton X-100 and 5% 웁-mercaptoethanol. Eggshell fragments were separated from the other egg chamber components by low-speed centrifugation. Scanning and transmission electron microscopy of the pelleted fraction indicated that eggshells purified in this manner consisted of endochorion and exochorion with no evidence of either the vitelline membrane or ICL layers. Approximately 20 SDS-soluble proteins in the purified eggshells were resolved by two-dimensional gel electrophoresis, including the six major chorion proteins previously identified by Petri et al. (1976). Choriogenic stage-specific synthesis patterns were established for most of these proteins by electrophoretic analysis of pulse-labeled egg chamber extracts from different developmental stages (Waring and Mahowald, 1979; Petri et al., 1976). The chorion proteins have been designated sXX, where s denotes shell and XX indicates its approximate size based on its mobility in SDS gels. The six major chorion proteins include s38, s36, s19, s18, s15, and s16. s36 and s38 are synthesized during the early choriogenic stages (s11–13). Production of s19 and s16 peaks during stage 13, whereas s18 and s15 synthesis is maximal during stage 14. A similar approach was used to identify vitelline membrane proteins, except stage 10 egg chambers were used for the eggshell fragment preparations and 웁-mercaptoethanol was omitted from the sonication buffer. Vitelline membrane fragments were isolated from stage 10 egg chambers radiolabeled in vivo for 2–5 hr and solubilized in SDS buffers containing 5% 웁-mercaptoethanol. Six protein species were resolved by SDS-PAGE (Fargnoli and Waring, 1982). Analysis of pulse labeled extracts from egg chambers at different developmental stages verified the stage-specific synthesis pattern (stages 8–11) of five of the six proteins. Synthesis by the follicle cells was confirmed by one- and two-dimensional gel analysis of enriched follicle cell preparations obtained from pulse-labeled stage 10 egg chambers. A polyclonal antiserum directed against stage 14 eggshell fragments prepared in the absence of 웁-mercaptoethanol recognized three of the putative vitelline membrane proteins in stage 10 extracts. The vitelline membrane components identified in this study were named sV130, sV70, sV23, sV17, and sV14 in keeping with the chorion protein nomenclature established previously. The sV denotes shell, vitelline layer. Mindrinos et al. (1985) electrophoretically resolved three major protein species from handdissected vitelline membranes isolated from radiolabeled ovaries that had been cultured in vitro. Based on labeling intensity and their stage-specific labeling patterns, these three proteins, v24, v14.5, and v11, most likely correspond to the sV23, sV17, and sV14 proteins identified by Fargnoli and Waring (1982).

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B. Genetic Analysis An alternative approach used to identify eggshell components was to correlate aberrations in eggshell protein profiles with abnormalities in eggshell morphology. In general, eggshell mutants are found among the larger class of female sterile mutants which display a collapsed egg phenotype. Mutations that affect the outer endochorion layer can be readily identified by ultrastructural analysis of the eggshell. Potential vitelline membrane mutants are more problematic because of the homogeneous ultrastructural appearance of this layer. Some female sterile mutants lay flaccid eggs which readily take up neutral red and ooze their contents when chemically dechorionated. Eggs from a collection of second chromosome female sterile mutants (Schupbach and Wieschaus, 1991) were screened for the collapsed and oozy egg phenotype. One mutant with the requisite phenotypic properties, fs(2)QJ42, was also defective in vitelline membrane protein synthesis. Stage 10 egg chambers from fs(2)QJ42 females failed to accumulate the previously identified vitelline membrane protein, sV23. The mutation was mapped cytogenetically to region 25D7 to 26A7 of the second chromosome (Savant and Waring, 1989). Two female sterile mutants with altered endochorion structure have been correlated with defects in specific eggshell proteins, cor-36 and dec-1. s36 is not detected in cor-36 egg chambers, and the endochorion fails to organize (Digan et al., 1979). The mutation maps cytogenetically to region 7E108A4 of the X chromosome, a 16-band region to which the s36 and s38 structural genes had previously been mapped using electrophoretic variants of the s36 and s38 proteins (Spradling et al., 1979). In contrast to the analyses of the sV23 and s36 mutant alleles, analyses of several defective chorion (dec-1) alleles recovered from female sterile screens of the X chromosome (Gans et al., 1975; Mohler, 1977; Komitopoulou et al., 1983; Bauer and Waring, 1987) revealed new eggshell components. Bauer and Waring mapped several dec-1 alleles to region 7C1-7C3 of the X chromosome and established a correlation between the fs(1)410 mutant allele and the lack of two proteins normally found in stage 14 eggshells prepared in the absence of reducing agents: s85 and s67 (Bauer and Waring, 1987). In vivo labeling experiments suggested that s67 was not a primary translation product. Pulse chase studies showed that s67 was derived from s85. Pulse labeling of stage 10 egg chambers and the analysis of proteins translated in vitro from stage 10 poly-A containing RNA both indicated that s85 itself was not a primary translation product but rather a derivative of a yet larger dec-1-related protein, fc130, which was synthesized in stage 10 follicle cells. The fc prefix for the 130-kDa protein denoted its follicle cell origin. Earlier studies by Lineruth et al. (1985) had shown that two geographically distinct wild-type strains produced three high molecular

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weight follicle cell protein variants simultaneously and that this variation mapped to region 7C (Lineruth et al., 1985). Electrophoretic analysis of an independent dec-1 mutant allele, fs(1)384, showed all three variant follicle cell proteins were affected simultaneously (Komitopoulou et al., 1988), a result consistent with multiple dec-1 protein products. More recently, the dec-1-related proteins, fc130, s85, and s67, have been renamed fc106, s80, and s60, respectively, to more accurately reflect the sizes of the conceptual products determined by translating DNA sequence data (Waring et al., 1990). Figure 3 shows a summary of the major eggshell proteins identified biochemically and genetically along with the periods in which they are synthesized or accumulate.

IV. Eggshell Genes A. Cloning 1. Chorion Genes In general, stage-specific synthesis patterns have been exploited to clone the major eggshell genes. By in vivo labeling RNAs, Spradling and Mahowald (1979) identified a small set of stage-specific, poly-A containing RNAs that were synthesized during the stages of eggshell formation. The follicle cell origin of two prominent choriogenic RNA bands (E3 and E4) was verified by separating the contents of the follicle cells from the remainder

FIG. 3 Eggshell protein synthesis and accumulation. Maximal synthesis of the vitelline membrane proteins (sV XX) occurs during stages 9–10B with peak accumulation during stage 10B. The dec-1 protein, fc106, is synthesized during stages 9 and 10; a smaller derivative, s80, first appears in stage 10B egg chambers. The early chorion proteins, s36 and s38, are synthesized during stage 11–13; synthesis of the s16 and s19 chorion proteins and accumulation of the smallest dec-1 derivative, s60, begin in late stage 13 and continue into early stage 14. Synthesis and accumulation of the late chorion proteins, s15 and s18, are stage 14-specific. The relative lengths of each stage or period are indicated at the top of the figure.

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of the egg chamber by mild detergent lysis. The E3 and E4 RNAs were linked to the 7E-F region of the X chromosome by in situ hybridization to polytene chromosomes. To isolate the genes encoding the follicle cell mRNAs, a cDNA library constructed from poly-A-containing RNAs isolated from choriogenic egg chambers (11–14) was differentially screened with poly-A-containing RNAs from choriogenic follicle cells and preblastoderm embryos. Putative chorion clones were identified based on their selective hybridization to the follicle cell RNA probe. Relationships between the putative chorion cDNA clones and specific chorion proteins were established by hybrid selection translation experiments. Three chorion clones were unambiguously identified in this manner: s36, s38, and s18. The s36 and s38 clones hybridized to region 7E11 of the X chromosome, whereas s18 hybridized to region 66D of the third chromosome (Spradling et al., 1980). The gene encoding s15, an abundant chorion protein synthesized during late choriogenesis (s14), was cloned in a like manner. A cDNA library enriched in sequences present in stage 13 and 14 egg chambers was screened for clones that selectively hybridized with RNAs synthesized in stage 13 and 14 egg chambers. Because the synthetic activities of choriogenic stage egg chambers are devoted almost exclusively to the elaboration of the eggshell by the follicle cells, the RNA probe was expected to be selectively enriched in RNAs encoding chorion structural proteins. Hybrid selected translation and immunoprecipitation were used to verify the identity of the s15 cDNA clone. Like s18, the s15 cDNA hybridized to region 66D of the third chromosome (Griffen-Shea et al., 1980). 2. Vitelline Membrane Genes The 26A region of the second chromosome was initially implicated in vitelline membrane formation by the isolation of a recombinant DNA phage clone that contained two coding regions (CR1 and CR2) which hybridized in a stage-specific manner (8–10) to ovarian cDNA probes. The cross hybridization of the CR2 region with the s18 gene from the autosomal chorion cluster suggested its involvement in eggshell formation (Higgins et al., 1984). A firmer relationship between the 26A region and vitelline membrane formation was subsequently established by the isolation of cDNA clones complementary to small RNAs (650–750 nt) that accumulated specifically in stages 8–10 egg chambers (Mindrinos et al., 1985; Burke et al., 1987). In situ hybridization of the corresponding genomic clones to polytene chromosomes revealed two cytological localizations: 26A and 34C of the second chromosome. Hybrid select translation using the 26A region sequence gave a product whose size was compatible with sV17 (Mindrionos et al., 1985; Burke et al., 1987). The RNA complementary to the 34C region

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DNA gave a translation product more compatible with a precursor for one of the smaller vitelline membrane components (e.g., sV14) (Mindrinos et al., 1985). More refined analysis of the 26A region clone revealed four tightly linked genes (TU-1 to TU-4) within a 12-kb span which encoded follicle cell RNAs that accumulated specifically during the period of vitelline membrane deposition (Popodi et al., 1988). For the more abundant transcripts (TU2 and TU-4), the follicle cell specificity was shown by hybridization of radiolabeled single-stranded RNA probes to sectioned ovarian tissue. For the less abundant transcripts (TU-1 and TU-3), follicle cell-specific expression was shown by comparative S1 analysis. Stage 10 egg chambers were cut at the midline to generate follicle cell and nurse cell fractions. The intensities of the S1-resistant fragments from the follicle and nurse cell fractions were compared with the S1-resistant bands from whole stage 10 egg chambers. The similar intensities of the follicle cell and whole egg chamber bands and the lack of a detectable signal in the nurse cell fractions indicated that the TU-1 and TU-3 transcripts were also expressed in a follicle cell-specific manner. The TU-2 gene corresponds to the previously identified sV17 gene. Hybrid select translation suggested that TU-4 encoded sV23 and that TU-3 encoded a follicle cell product of approximately 20 kDa (fc 20) (Popodi et al., 1988). A small vitelline membrane-like gene was identified in a study designed to isolate ovarian cDNAs complementary to DNA from region 32 of the second chromosome, a region which contains the maternal effect genes hup, wdl, dal, and abo (Gigliotti et al., 1989). DNA sequence analysis predicted a 13-kDa secreted protein rich in alanine and proline. The stagespecific accumulation of the 32E RNA in stage 10 egg chambers was compatible with its assignment as a vitelline membrane gene. 3. dec-1 Even though most eggshell genes have been cloned by screening ovarian cDNA libraries for genes that are selectively expressed during the stages of eggshell formation, the dec-1 gene was isolated by positional cloning (Hawley and Waring, 1988). dec-1 was mapped cytogenetically to region 7C1-7C3 of the X chromosome. With one exception, the recessive female sterility of all the dec-1 mutant alleles tested was uncovered by Df(1)ct4b1, a deficiency chromosome with breakpoints in the 7B1,2 and 7C3 regions. The exceptional dec-1 allele, fs(1)1501, was fully complemented by Df(1)ct4b1. This unusual complementation behavior suggested that the deficiency chromosome and the fs(1)1501 chromosome retained complementary dec-1 genetic functions and that the 7C3 breakpoint of the deficiency chromosome was located within or near the dec-1 locus. A breakpoint-

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containing clone was isolated from a genomic library from flies carrying the Df(1)ct4b1 chromosome. The 7C sequences from the breakpoint clone were used to initiate a 20-kb chromosome walk which extended in both directions. The dec-1 gene within this 20-kb region was identified by several criteria including: (1) complementarity to a RNA species of sufficient size to encode a 106-kDa protein product, (2) stage-specific expression during the stages when fc 106 is produced (9–10), (3) the correlation of genomic alterations in the region with a dec-1 mutant strain encoding an aberrantly sized transcript, ( fs(1)764), and a wild-type strain encoding variant dec-1 proteins (Shahrinah) (Hawley and Waring, 1988), and (4) restoration of fertility in dec-1 null mutants by the introduction of a transgene containing a 9.6-kb Nco-Bg1I fragment from the region (Waring, unpublished).

B. Conceptual Translation Products 1. Vitelline Membrane Genes Conceptual translations of the putative eggshell genes showed that all the open reading frames were headed by a stretch of hydrophobic amino acids characteristic of a signal peptide followed by potential cleavage sites. This findings was consistent with EM autoradiography which suggested that eggshell proteins are synthesized by the follicle cells and secreted into the extracellular space in Golgi-derived vesicles (Giorgi, 1977). Most of the eggshell genes expressed during the period of vitelline membrane formation (34C, 32E, sV17, and sV23) are small, intronless, and encode proteins rich in alanine, proline, serine, and glycine. The mature proteins each contain a highly conserved hydrophobic domain 38 amino acids in length which has been termed the ‘‘vm domain’’ (Scherer et al., 1988). Edwards et al. (1998) have recently reported that vitelline envelop genes from the mosquito, Aedes aegypti, contain a conserved region which overlaps in sequence with the D. melanogaster ‘‘vm domain.’’ In the region of overlap (22 amino acids) eight amino acids, including three cysteine residues, have been conserved. Using this conserved consensus motif as a query sequence (Fig. 4, Ac/Dm), two additional D. melanogaster genes containing the vm domain have been identified in the Drosophila genome project database: fc20 and a previously unidentified X-linked sequence, ‘‘VM 78’’ (Fokta, F. J. and Waring, G. L., unpublished). Based on deduced amino acid sequence, fc20 (with its hydrophobic signal sequence, and glycine and proline-rich content) fits the profile of a typical vitelline membrane protein. The fc20 proteins also features a central region, consisting of ten tandem repeats of the pentamer PGFGG. The deduced amino acid sequence of the X-linked gene is striking it its similarity to sV23. The central region of sV23 contains an

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8 amino acid repeating motif, YSAPAAPA, which makes up approximately one-half of the entire protein (Popodi et al., 1988). The X-linked sequence has a 662 amino acid open reading frame, almost all of which consists of either perfect or slightly variant forms of the repeat sequence YSAPAAPS. Whereas the composition and inclusion of the vm domain in the X-linked open reading frame is consistent with an eggshell protein, other proteins may possess these features. In particular, using the conserved vm domain as a probe, Gigliotti et al. (1989) isolated cDNA clones with expression patterns that would exclude their participation in eggshell formation. The apparent inclusion of this highly conserved domain in other proteins suggests a common function that may be shared by other proteins participating in the formation of extracellular structures. Conceptual translation of TU-1 from the 26A region follicle cell gene cluster revealed remnants of a vm domain. Within a stretch of 15 amino acids, TU-1 contains 8 residues which are identical to those in the ‘‘vm’’ consensus sequence shown in Fig. 4.

FIG. 4 vm domain containing genes. Comparative sequence analysis of the sV23, sV17, VM34c, and VM32e genes revealed the highly conserved 38 amino acid region shown in the first four lines termed the vm domain. Conserved regions found in mosquito vitelline envelop genes, which overlaps with vm domain sequences are shown in the three lines at the bottom of the figure. Ae shows the positions of the eight amino acids which are identical in all 7 sequences (sV23, sV17, VM34c, VM32e, Ae-1, Ae-2, and Ae-3). vm domain-like sequences found in two additional D. melanogaster genes, fc20 and ‘‘VM 78’’ are shown. ‘‘VM 78’’ is an X-linked sequence with a coding potential of 78,000 Da. Based on its striking similarity to the vitelline membrane protein, sV23, it has been temporarily coined ‘‘VM 78’’ despite the lack of either RNA or protein expression data (Fokta and Waring, unpublished). A comparison of the six D. melanogaster sequences lead to a refinement of the vm domain consensus sequence (Dm) (Fokta and Waring, unpublished). Amino acids within the Dm consensus sequence are found in at least 4 of the 6 D. melanogaster sequences. The underlined residues are identical in all six sequences.

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2. Chorion Genes The open reading frames of the major eggshell genes expressed during the choriogenic stages have been sequenced in D. melanogaster as well as several evolutionarily related species. With the exception of s16 in the autosomal gene cluster, all the open reading frames are interrupted by a small intron located near the 5⬘ end of the gene in the region encoding the signal peptide. The structure of the s16 gene differs significantly in that it contains a small intron at the end of the gene, creating a small (27–29 nt) 3⬘ exon. The organization of both the X-linked and autosomal gene clusters has been conserved through at least 120 million years of evolution (Konsolaki et al., 1990; Aggeli et al., 1991; Vlachou et al., 1997). The s36 and s38 proteins both exhibit a tripartite structure with each containing a central domain rich in lysine, proline, and valine flanked by two arms rich in glycine, alanine, and serine (Levine and Spradling, 1985). The central regions of both s36 and s38 show high evolutionary conservation and are predicted to adopt a structure consisting of short 웁-sheets alternating with 웁-turns (Hamodrakas et al., 1989). The structure is formed by tandem repeats of X3Z(4-6), where X represents a large hydrophobic residue and Z represents a 웁-turn forming residue. Sequence comparisons between the Ceratitis capitata and Drosophila melanogaster s38 proteins (separated by approximately 120 million years) revealed only four variant residues within this central region and no insertions or deletions (Konsolaki et al., 1990; Tolias et al., 1990). The left arm of s36 is similar to the right arm of s38 in that both contain stretches of the repeated dipeptide-glycine/histidine. Tyrosine residues, at least some of which are involved in crosslinking, are abundant in the arms of both proteins but occur rarely in the central domains. The asymmetric distribution of amino acid residues within both proteins suggests that the arms and central regions have distinct functional and/or structural roles (Hamodrakas et al., 1989). The proteins encoded by the autosomal chorion genes bear little resemblance to one another other than having a rich alanine, glycine, and proline content. The genes do not cross hybridize and, in general, show extensive sequence divergence through evolution (Martinez-Cruzado et al., 1988; Fenerjian et al., 1989; Swimmer et al., 1990; Martinez-Cruzado, 1990; Vlachou et al., 1997). Sequence comparisons between subgenera that span 50 million to 80 million years show that s16 is strikingly conserved over most of its length, whereas s15 has evolved the most rapidly, exhibiting extensive sequence divergence. Secondary structural predictions indicate that, like the central domain of s36 and s38, the autosomal chorion proteins are all highly structured with 웁-sheets alternating with 웁-turns. The structure of s16 is unusual in that the putative 웁-sheets and 웁-turns are interrupted by a central highly conserved alpha helix structure (Vlachou et al., 1997).

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Unlike their X-linked counterparts, homologs of the autosomal chorion genes could not be detected in the medfly, Ceratitis capitata, by cross hybridization. Entry into the locus via a highly conserved adjacent paramyosin gene revealed highly conserved peptide segments within each chorion gene that allowed unambiguous comparisons with the Drosophila homologs (Table I) (Vlachou et al., 1997). In general, the medfly proteins are longer than their Drosophila counterparts. Even though s16 is slightly larger, the s19 and s18 medfly proteins are nearly double in length. In both proteins, most of the increase in length can be attributed to a segment consisting of tandem repeats of the heptapeptide YSAPAPS. Although absent in the Drosophila chorion genes, this heptapeptide bears a striking resemblance to the octapeptide YSAPAAPA found in the sV23 vitelline membrane gene. 3. dec-1 Gene The dec-1 eggshell gene is distinctive in that it is a large, intron-containing gene, which produces multiple RNAs through the use of alternative RNA processing pathways (Waring et al., 1990). Developmental Northern blots showed that the dec-1 sequences hybridized to two different size classes of poly-A-containing RNAs: a 4.0-kb class (3.7 kb based on DNA sequence analysis), whose accumulation peaked during stages 9 and 10, and a 5.8-kb class detected later in stages 11 and 12 egg chambers (5.7-kb based

TABLE I Distinctive amino acid sequences in eggshell proteins Proteins

Amino acid sequences

sV23 fc20 TU-1 s36 & s38 dec-1 s18 s15 s19 s16

YSAPAAPA PGFGG APAPPAPAYE XXXZ(4-6) QN-MMM–RQW-E(E/D)QAK-QQ VGGYAYQVVQPALTV-AI PSAAAAAAAA-A–NPG-Y-Q-AVP-YEL PRWTVQPAGATLLYPGQNSYR-Y-SPPEYSKV-LPVRAA-PVAKLY-PEN EAQAAALTNAAGAAASAAK

Repeating motifs found in the vitelline membrane proteins sV23, fc20, and TU-1 are shown in the first three rows (Fokta and G. L. Waring, unpublished). The evolutionarily conserved structural motif (X3Z(4-6)) of the s36 and s38 chorion proteins is shown in row four (Xhydrophobic residue; Z-beta-turn former)(Konsolaki et al., 1990; Tolias et al., 1990). Features of the central dec-1 repeat that have been conserved in D. melanogaster and D. virilis are shown (Badciong and Waring, unpublished) as are sequences that have been conserved in the fruit fly and medfly in the proteins encoded by the autosomal chorion gene cluster (Vlachou et al., 1997).

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on DNA sequence data). RNAse protection assays, using RNA from stage 9 or 12 egg chambers and a series of antisense probes that spanned the dec-1 gene, indicated that the only discernible difference between the 5.7-kb and the 3.7-kb transcripts was the inclusion of a 2-kb intron (IVS-4) in the former. Comparative DNA sequence analysis of the genomic dec-1 clone with 18 independent cDNA clones isolated from an ovarian cDNA library (Hawley and Waring, 1988) revealed more subtle differences in the processing of the transcripts. Ninety percent of the cDNA clones representing the 3.7-kb class had 14 more nucleotides at the 5⬘ end of exon 4 (Fig. 5, 3.7B) than the 5.7-kb RNA (5.7A), whereas 10% were identical (3.7A). Using antisense oligonucleotide probes complementary to the E3E4 splice junctions representing the A and B pathways, respectively, developmental Northern blots showed that, like 3.7B, accumulation of the 3.7A transcript peaked during stages 9–10. The 10- to 20-fold greater abundance of the 3.7B transcript suggests that in stage 9 and 10 egg chambers the B pathway predominates. The machinery needed for pathway A is not limiting in the cell as elevated levels of the 3.7A transcripts accumulate in splicing mutants. In fs(1)1501, the upstream AG splice acceptor site used to generate the 3.7B RNA is mutated to TG. In its absence, the alternative downstream acceptor site is utilized efficiently, and the 3.7A transcripts accumulate at approximately 10-fold greater levels than in wild-type stage 10 egg chambers (Waring et al., 1990). The higher levels of the 3.7A transcripts in the splicing mutant is consistent with 3.7B:3.7A ratios being regulated at the level of RNA processing rather than RNA stability. During stages 11 and 12 there is a precipitous drop in dec-1 RNA accumulation. Concomitant with this decrease is the emergence of the 5.7-kb transcript. The predominance of the 5.7-kb RNA in these stages suggests both preferential use of the A pathway and masking or negative regulation of the 5⬘ donor and/or 3⬘ acceptor sites in IVS-4 used to generate the 3.7-kb RNAs. Production of the 5.7-kb message does not appear to be restricted

FIG. 5 Alternative processing of dec-1 transcripts. The bold lines represent the exons (E1E5) which compose the three RNAs produced from the dec-1 gene. The 5.7A and 3.7A transcripts differ by the inclusion of intron 4 sequences in the former. The 3.7B transcript differs from the 3.7A transcript by the inclusion of 14 additional nucleotides at the 5⬘ end of exon 4. The extra nucleotides in exon 4 of the 3.7B transcript are shown. Use of an alternative 3⬘ splice acceptor site (AG) yields a E3/E4 junction characteristic of 3.7A and 5.7A.

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to a particular subset of follicle cells. A dec-1 antiserum raised against a peptide specified by sequences within the IVS-4 open reading frame reacted with a product that was distributed in all regions of the eggshell in stage 14 egg chambers (Mauzy-Melitz and Waring, unpublished). Open reading frame analysis of the dec-1 RNAs (Waring et al., 1990) predicted proteins of 106, 125, and 177 kDa (Fig. 6). The inclusion of the 14 additional nucleotides in the 3.7B transcript causes a frameshift that results in the termination of the open reading frame 19 nt downstream from the E3-E4 splice junction and the production of the 106-kDa product, fc106. The 3.7A open reading frame extends well into exon 5 leading to the production of fc125. The 5.7-kb transcript encodes fc177, the largest dec-1 product. All three proteins share an identical N-terminus that includes a central region (332 aa) consisting of tandem repeats of a 26 aa motif (QNPMMMQQRQWSEEQAKIQQNQQQIQ). The distinctive nature of this central region (39% glutamine, 16% methionine) is underscored by the fact that dec-1 is the only eggshell protein identified to date that contains internal methionines. Comparisons of the repeat sequences in D. melanogaster with those in its evolutionary distant D. virilis homolog (separated 50 million to 80 million years) (Badciong and Waring, in preparation) revealed several conserved features (see Table 1). Each dec-1 protein has a distinct C-terminus. Because of the extensive overlap between fc106 and the two larger derivatives, the C-terminus of fc106 contains only 6 amino acids that are not included in fc125 or fc177. The C-terminus of fc125 includes 138 fc125-specific amino acids, whereas the fc177-specific Cterminus contains 604 amino acids. Two conspicuous regions in the fc177specific C-terminus were revealed by comparisons of the D. melanogaster and D. virilis dec-1 homologs (Badciong and Waring, unpublished). Eight cysteine residues contained within a small stretch of 36 amino acids near the beginning of the fc177-specific sequence were strictly conserved. The

FIG. 6 Deduced protein products of the dec-1 locus. fc106, the translation product of the 3.7B RNA, contains a large central region B consisting of perfect and imperfect copies of a 26 amino acid repeat flanked by arms A and C. fc177 and fc125 essentially contain all the fc106 sequences. Region D shows the 42 amino acid segment encoded by exon 4, which is found in both the fc 125 and fc177 proteins. Regions E, F, and G are fc177-specific; H represents the fc125-specific C-terminus. E marks a small cysteine-rich region that heads the fc177specific sequence, whereas F denotes a highly conserved 65 amino acid block enriched in charged amino acids.

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cysteine-rich region was followed by a highly conserved block of 65 amino acids (93% identity) rich in charged amino acids (37%). Even though there were small patches of conserved residues scattered throughout the remainder of the sequence, extensive deletions in the D. virilis homolog reduced the fc177-specific C-terminus by 200 amino acids in this species. The evolutionary comparison of the fc125/C-terminal specific sequences revealed a conserved central domain (51 amino acids, 75% identity) flanked by more variable N-terminal (28 amino acids, 25% identity) and C-terminal (54 amino acids, 40% identity) sequences. Small insertions and deletions punctuated the fc125-specific C-terminus. In D. virilis the C-terminal end of fc125 was extended by 74 amino acids, 48% of which were charged residues. Translation in vivo of the conceptual products was confirmed by Western blot analysis (Noguero´n and Waring, 1995). Using antisera directed against peptide sequences encoded by the N-terminal region, both fc106 and fc125 were detected in wild-type stage 10 egg chambers at levels that reflected the relative abundances of their respective RNAs. Initial identification of the fc125 product was facilitated by its overexpression in fs(1)1501 mutant females. The use of an antiserum directed against peptide sequences within the fc177-specific C-terminus allowed visualization of fc177 in extracts from stages 11–12 egg chambers. A summary of the sequences or motifs which define some of the eggshell proteins is presented in Table 1.

V. Posttranslational Processing of Eggshell Proteins A. dec-1 Proteins Although in vivo labeling experiments suggested that fc106 was processed in two steps to 80- and 60-kDa derivatives, the complexity of dec-1 processing was not appreciated until serological reagents which provided a more definitive means to track dec-1 related products were developed. Open reading frames from selected regions of the dec-1 gene as well as other eggshell genes (s18, s36, sV17, and sV23) were subcloned in frame into pATH expression vectors under the control of the inducible tryptophan promoter (Noguero´n and Waring, 1995; Pascucci et al., 1996). Electrophoretically purified bacterial fusion proteins were used to produce rabbit polyclonal antisera. Western blot analyses of egg chamber extracts from different development stages revealed stage-specific processing of not only the dec-1 proteins (Noguero´n and Waring, 1995) but also the sV23 and sV17 vitelline membrane proteins (Pascucci et al., 1996).

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During late stage 10, fc106 is cleaved to a smaller s80 derivative. To determine whether sequence were cleaved from the N- or C-terminus, Western blots of stage 10 egg chamber extracts were reacted with antisera directed against either the N- or C-terminus of fc106 (Noguero´n and Waring, 1995). fc106 and a 25-kDa product (s25) reacted with the N-terminal serum, whereas fc106 and s80 reacted with the C-terminus serum. Nterminal sequencing of s80 allowed placement of the cleavage site between alanine-280 and serine-281 of fc106 (Waring et al., 1990), a site that is not a recognizable target for known proteases. The alanine, proline-rich nature of the N-terminal product, s25, is typical of vitelline membrane proteins. Recognition and utilization of the fc106 cleavage site does not appear to be dependent on the presence of most, if not all, of the N-terminal sequences (Mauzy-Melitz and Waring, unpublished). An N-terminal deletion mutant producing an 81-kDa protein lacking most of the s25 region was cleaved to s80 in late stage 10 egg chambers. Antisera directed against either the N- and C-termini of s80 were used to show that s80 is cleaved at its Nterminus during stages 13–14, yielding a 20-kDa N-terminal derivative, s20, and a 60-kDa C-terminal derivative, s60 (Fig. 7A). The less abundant dec-1 proteins, fc125 and fc177, are also processed in a stage-specific manner. The fc125 maturation pathway was elucidated in the overproducing mutant, fs(1)1501, and confirmed in wild-type egg chambers. Western blot analysis using an N-terminal serum and a more C-terminal serum showed that fc125 is cleaved to a 95-kDa C-terminal derivative, s95, through a series of processing intermediates. Unlike the fc106 pathway, a stable N-terminal product is not produced nor is s95

FIG. 7 Proteolytic processing of dec-1 proteins. fc106 is cleaved at its N-terminus (downward arrow) in late stage 10B egg chambers to s25 and s80. During late stage 13 and early stage 14, s80 is cleaved at its N-terminus (upward arrow) producing s20 and s60. fc125, shown below, is cleaved at its N-terminus in late stage 10 (downward arrow) to a 95 kDa C-terminal derivative, s95, which persists in soluble form through stage 13. The central gradient depicts the repeat region in fc106 and fc125; the filled and shaded regions in B represent regions shared with fc177 and unique to fc125, respectively.

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subjected to additional cleavage in late stage egg chambers (Fig. 7B). Because fc106 is essentially contained in its entirety within fc125, the distinct maturation pathways and differential use of potential cleavage sites was not anticipated. Clearly, residues that lie in the distant C-terminal regions play a role in how the proteins are processed. The importance of the fc125 C-terminus was established by analyses of a genetically engineered dec-1 truncation mutation. A dec-1 transgene harboring a premature termination codon in the fc125-specific C-terminus which eliminated 118 amino acids from its C-terminus was crossed into dec-1 null mutants. Although fc106, s80, and s60 were produced as anticipated, truncated forms of fc125 and s95 in the size ranges expected were not detected (Mauzy-Melitz and Waring, unpublished). These results are compatible with aberrant processing and/ or the generation of protein products that are rapidly degraded in the absence of a normal C-terminus. The sterility of the fc125-deficient flies confirmed its essential function in eggshell assembly (Mauzy-Melitz and Waring, unpublished). fc177, produced during stages 11–12, is processed to a stable 85-kDa Cterminal derivative (s85) during stages 12–13 via a 120-kDa processing intermediate (Noguero´n and Waring, 1995). Although fc177 is processed at a later time than fc106 or fc125 (stages 12–13 vs late 10), the fc177 maturation pathway does not appear to be stage dependent. When fc177 was expressed prematurely by creating a fc177 transgene under the control of normal dec-1 regulatory sequences, fc177 and s85 accumulated prematurely in otherwise wild-type stage 10 egg chambers (Mauzy-Melitz and Waring, unpublished). B. Vitelline Membrane Proteins Posttrranslational processing of eggshell proteins is not limited to dec-1. At least two vitelline membrane proteins, sV17 and sV23, are also procesed in a temporally regulated manner (Pascucci et al., 1996). During stages 13–14, both sV23 and sV17 display a marked increase in electrophoretic mobility on SDS gels (Fig. 8). The increase is compatible with the loss of about 25–30% of their original masses. For sV23, the origin of the smaller reactive species was definitely established as sV23 by its absence in latestage egg chambers from the sV23 null mutant, fs(2)QJ42. Processing of sV23 is associated with a loss of epitopes. An acetone powder from fs(2)QJ42 egg chambers was used to preabsorb cross-reacting antibodies from a sV23 antiserum. The purified serum recognized stage 10 vitelline membrane epitopes it situ and on Western blots. In marked contrast, it failed to react with stage 14 epitopes either in situ or on Western blots. These results indicate that epitopes found in the sV23 stage 10/11 product are removed as sV23 is processed to its smaller derivative during stages

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FIG. 8 Late-stage processing of vitelline membrane proteins. Proteins in extracts from staged egg chambers were separated by SDS-PAGE, transferred to nitrocellulose, and reacted with antisera directed against bacterial fusion proteins containing open reading frames from either the sV23 or sV17 vitelline membrane proteins. The reactive species found in choriogenic stage egg chambers (11–14) are shown. (Adapted from portions of Fig. 1, Pascucci et al., 1996)

13–14 (Pascucci et al., 1996). Although less definitive, epitope loss has also been implicated in the processing of sV17. Immunogold analysis using sV17 antisera showed a dramatic decrease in the density of gold particles over the vitelline membrane as the egg chamber developed from stages 10–12 to 14. The late stage decrease in the reactivity of the sV17 antiserum noted in situ was also observed on developmental Western blots (Fig. 8). Relative to the intensity of the sV17 signal in stages 10–12 egg chambers, the signal from the smaller stage 13–14 derivative was markedly reduced (Pascucci et al., 1996). Inspection of the conceptual open reading frames of sV23 and sV17 revealed one potential proteolytic target site, a paired basic motif (RXXR or RLRK) in the N-terminal regions of sV23 and sV17, respectively, whose use would be consistent with the sizes and the immune reactivities of the smaller stage 14 derivatives. Paired basic motifs are at the cleavage sites of many pro-proteins that are processed by eukaryotic convertases. A furin-like convertase has been implicated in the processing of provitellogenins in the mosquito, Aedes aegypti (Chen and Raikhel, 1996; Sappington and Raikhel, 1998). Homologs of mammalian furins, Dfurins, have been identified in Drosophila (Hayflick et al., 1992; Roebroek et al., 1995). Ovarian expression (Hayflick et al., 1992), as well as the secretion of soluble Dfurin isoforms, has been reported (De Bie et al., 1995), both features that would be required for the extracellular processing of vitelline membrane substrates. C. Chorion Proteins Unlike dec-1 and some of the vitelline membrane proteins, the chorion proteins do not appear to be processed or are processed in a manner

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that is not perceptible by electrophoretic analyses. Fluorographs of twodimensional gels of chorion proteins synthesized in stages 11–14 egg chambers and labeled chorion fragment preparations from stage 14 egg chambers run in parallel were compared. The superimposition of the newly synthesized (stages 11–14) and mature chorion proteins (stage 14 chorion fragments) showed that the sizes and the charges of the chorion proteins are not altered as the egg chambers mature (Waring and Mahowald, 1979). Developmental Western blots of the early chorion protein, s36, and the late chorion protein, s18, confirmed that these proteins are not subject to posttranslational modifications that alter their mobilities on SDS polyacrylamide gels (Pascucci et al., 1996).

VI. Assembly A. Production of Specialized Regions I. Follicle Cell Migrations Regional complexity is exhibited at the anterior and posterior ends of the egg. Specialized structures formed at the anterior end include the micropyle, operculum, and dorsal appendages. While there is little knowledge of whether, or how, the protein composition of the specialized regions differs from that in the main body, it is known that the specialized regions are elaborated by distinct subsets of migrating follicle cells. The first follicle cells to become differentiated from the remainder of the follicular epithelium are a pair of cells at each end of the newly formed egg chamber called polar follicle cells. The polar follicle cells express characteristic marker proteins (Margolis and Spradling, 1995) and do not proliferate. During stage 9 nearly all follicle cells migrate posteriorly over the oocyte. The anteriormost 6–10 follicle cells (border cells) which include the anterior polar follicle cells delaminate and migrate as a group posteriorly between the nurse cells toward the anterior pole. The border cells arrive at the anterior end of the oocyte by stage 10, where they participate in formation of the micropyle. The posteriorward migration of all the follicle cells is essentially complete by early stage 10 (10a). Approximately 95% of the follicle cells (columnar follicle cells) cover the oocyte; 5% remain stretched over the nurse cells. Most of the columnar follicle cells are involved in production of the mainshell. During late stage 10 (10b), a subset of about 150 follicle cells migrates centripetally between the nurse cells and oocyte. The centripetally migrating follicle cells (CMFC) produce the operculum and a small subset of this population collaborates with the border cells to form the micropyle.

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2. Formation of the Micropyle Formation of the micropyle begins in stage 10 egg chambers with the secretion of a paracrystalline region of the vitelline membrane by the border cells. During stage 11, follicle cells immediately adjacent to the border cells begin to secrete a spongy vitelline membrane component which is morphologically distinguishable from the paracrystalline region and the vitelline membrane that surrounds the rest of the oocyte. The polar follicle cells within the border cell cluster extend a long process which penetrates the paracrystalline region forming a pocket (Margaritis, 1984). The border cell process is rich in microtubules (Zarani and Margaritis, 1986) and actin cytoskeletal elements (Edwards et al., 1997). The vitelline membrane protrusion, consisting of the spongy vitelline membrane, the eccentric paracrystalline layer, and pocket, is completed by stage 12. The vitelline membrane protrusion fits into the hollow part of an outer endochorion cone which is secreted between stages 12 and 14 by a subset of approximately 36 CMFCs. The border cell process around which the vitelline membrane and endochorion components of the micropyle are secreted degenerates during stage 14B giving rise to the micropylar canal through which the sperm enter (Zarani and Margaritis, 1986).

3. Formation of the Operculum and Dorsal Appendages Proper patterning of the anterior follicle cells which produce the dorsal appendages and the operculum involves the convergence of two major signaling pathways. The anterior follicle cells are initially subdivided into two populations (midline and dorsal lateral) by a gurken signaling pathway. The pattern is then refined by decapentaplegic (dpp) signaling which commits the anterior-most CMFC to the production of the operculum. Torpedo (tor), a homolog of the mammalian epidermal growth factor receptor (EGFR), is activated in anterior-dorsal follicle cells by the localized secretion of the ligand gurken from the oocyte in stages 8 and 9 egg chambers. Even though activation of the epidermal growth factor receptor by gurken is necessary for correct patterning in the dorsal anterior region of the eggshell, it is not sufficient. Proper positioning of the dorsal respiratory appendages requires the expression of at least four additional genes in the follicle cell epithelium: spitz, vein, rhomboid, and argos (Wasserman and Freeman, 1998). Spitz, vein, and rhomboid participate in an autocrine regulatory circuit which increases the amplitude and extent of the initial EGFR signal in stage 10 egg chambers. In response to the amplified signal, argos is expressed in stage 11 follicle cells at the dorsal midline. The localized expression of argos inhibits EGFR activation, causing cells expressing activated EGFR to be split into two populations. The peaks of EGFR expres-

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sion on either side of the dorsal midline specify the positions of the dorsal appendage producing cells. Ectopic activation of tor/EGFR throughout the follicular epithelium produced one of two phenotypes—ectopic dorsal appendages or an expansion of operculum-like material (Queenan et al., 1997). The different phenotypes reflected different levels of EGFR expression. The highest levels of ectopic expression were associated with the operculum or dorsal-like fate. Lower levels of EGFR expression were associated with dorsolateral or appendage-producing cell fates. These observations are consistent with the notion that inhibitors of EGFR activation, such as argos, are induced above threshold levels of EGFR expression. Threshold levels of EGFR expression are normally encountered only in the follicle cells at the dorsal midline. The production of the operculum by the anterior-most follicle cells is influenced by TGF-웁 signaling. Using conditional lethal mutants Twombly et al. (1996) showed that diminished levels of dpp, a TGF-웁 family member, are associated with loss of the operculum and anterior most follicle cell fates. Conversely, ectopic expression of dpp through the expression of a hsp70-driven transgene led to the formation of an enlarged operculum. The dorsal appendages (Figs. 1 and 2) are specified by 55–65 cells in the dorsal lateral regions on either side of the dorsal midline (Deng and Bownes, 1997). In response to appropriate signals, the dorsal/anterior follicle cells migrate in two circles and secrete chorion proteins centripetally to form the base of the appendages. When the base is completed, a layer of follicle cells migrate outwardly past the previous cells, forms another circle, and secretes additional chorion material. Migration and chorion deposition continues in a similar manner, leading to the formation of the tubular structure. The follicle cells then spread out to form the flat paddle structure (Rittenhouse and Berg, 1995). Abnormal morphogenesis of the dorsal appendages has been associated with mutations that disrupt follicle cell migrations as well as those which alter follicle cell fates. P(Gal) enhancer trap lines revealed that a specific isoform encoded by the BR-C complex specifically marks the progenitor cells for dorsal appendages in stages 10b and 11. BRC function is required for dorsal appendage morphogenesis as partial loss of function mutants produce shorter and thinner appendages. The production of shorter and thinner appendages is consistent with a decrease in the number of follicle cells which are committed to secreting dorsal appendages (Deng and Bownes, 1997). The abnormal appendage morphology exhibited in bullwinkle (bwk) mutants reflects impaired migration rather than a change in the number of dorsal appendage-producing cells. By introducing a mutant bwk mutant allele into an enhancer trap line which marked appendage cell precursors, Rittenhouse and Berg (1995) showed that appendage-producing cells appeared over the dorsal/lateral regions of the oocyte in late stages 10 and 11 egg chambers as in wild type. However,

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their limited outward migration in the later stages of choriogenesis led to the production of short, broad, dorsal appendages. The analysis of germline clones produced by mitotic recombination and follicle cell clones produced by pole cell transplantation showed bwk function is required in the germ line. Germ-line expression of the bullwinkle product suggests that it is needed for follicle cell guidance or traction.

B. Production of Mutilayered Mainshell Whereas small subsets of follicle cells are involved in elaborating the specialized eggshell structures found at the anterior of the egg, the majority of follicle cells are involved in elaborating the multilayered mainbody or mainshell. Electron microscopic autoradiography suggests that the spatial distribution of proteins within the layers of the eggshell is dictated in large part by when the proteins are synthesized. The vitelline membrane was heavily labeled by pulse-labeled proteins secreted from stages 9–10 follicle cells. Grains were localized predominantly over the ICL and endochorion floor in pulse-labeled stage 12 egg chambers, whereas newly synthesized follicle cell proteins were found in the forming pillars in stage 13 egg chambers (Giorgi, 1977). Based on more recent pulse-chase experiments, Papassideri and Margaritis (1996) also suggested that proteins labeled during early choriogenesis (stages 10b–11) reside primarily in the ICL and inner endochorion layers. The acquisition of antisera directed against specific eggshell proteins provided a means to add molecular definition to this generalized picture of how the multiple layers form. Immunoelectron microscopy of ovarian tissue sections reacted with sV17 and sV23 antibodies showed that sV17 and sV23 are found only in the vitelline membrane layer after its completion at the end of stage 10 (Pascucci et al., 1996). These data are consistent with the build-up of the eggshell layers by zones, starting with the oocyte proximal vitelline membrane layer and ending with the outer endochorion roof. Immunolocalization of s18, s36, and the dec-1 proteins suggest added complexity highlighted by the transient sequestration and regulated release of several eggshell proteins from the vitelline membrane layer. 1. Changes in the Distribution of Major Chorion Proteins in the Assembling and Mature Eggshell s18 is one of two major eggshell proteins whose synthesis peaks in stage 14 egg chambers. As a member of the late chorion protein class, its localization within the outer endochorion roof was anticipated. Using antibodies directed against the C-terminal half of the protein, s18 was initially localized

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to the endochorion roof and apices of the pillars in late stage 13 and early stage 14 egg chambers. As s18 accumulation peaked in mid and late stage 14 egg chambers, it became distributed throughout the floor, pillars, and roof of the endochorion. In the anterior region of the eggshell, the pillar, roof, and floor substructure are apparent in late stage 12 and early stage 13 egg chambers, prior to synthesis and secretion of s18, suggesting that s18 is not required to establish either the roof or pillar framework and intercalates into a pre-existing scaffold or framework. Trougakos and Margaritis (1998a) recently reported that the Drosophila virilis homolog of the late chorion protein, s15, also immunolocalizes throughout the layers of the tripartite endochorion in late stage 14 egg chambers. The endochorion was almost devoid of immunogold particles during early stage 14 when the endochorion is essentially morphologically complete. Dvs 19, the D. virilis homolog of s19, was similarly immunolocalized over the entire endochorion, including the outer roof, pillars, and inner floor, at late stage 14. The temporal and immunolocalization patterns of both Dvs 15 and Dvs 19 support a model whereby ‘‘late’’ eggshell proteins intercalate into a preformed structure (Trougakos and Margaritis, 1998a). s36 is one of two major chorion proteins produced during early choriogenesis (stages 11–13) when the ICL, endochorion floor and pillar structures are forming. The lack of pillar formation in s36 null mutant egg chambers suggested that s36 may be a pillar component (Digan et al., 1979). Immunogold labeling of stage 14 eggshells with antibodies directed against the Cterminal half of the s36 protein showed that s36 distributes throughout the tripartite endochorion in a manner indistinguishable from that of s18 (Pascucci et al., 1996). If the chorion is built in zones, as envisioned by the autoradiographic data, as a member of the early temporal class, the spatial distribution of s36 was expected to be distinct from s18, a late chorion protein. More surprising, however, was the association of a large proportion of s36 with the vitelline membrane layer during the early choriogenic stages. Sectioned egg chambers often display regions where there is a pronounced separation between the vitelline membrane and forming chorion, an artifact of processing (Margaritis, 1985). Immunogold labeling of stage 12 egg chambers with the s36 antisera in such regions showed that the majority of gold particles associated with the vitelline membrane layer; the remainder were near the follicle cell border in the region where the chorion was forming (Fig. 9). Gold particles over the vitelline membrane were not detected in stage 14 egg chambers, suggesting that the s36 epitopes associated with the vitelline membrane during the early choriogenic stages are either masked during late oogenesis, turned over, or released to the assembling chorion during late choriogenesis. Developmental Western blots show that the accumulation of s36 peaks during stage 13 and peak levels of soluble s36 are maintained well into stage 14, a result incompatible with

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FIG. 9 Relocalization of s36 during late choriogenesis. Thin sections of stage 12 (A) or stage 14 egg chambers (B, C) were reacted with a rabbit polyclonal s36-specific antiserum. Goat anti-rabbit IgG, coupled to 20-nm immunogold particles, were used to visualize the bound primary antibodies. vm, vitelline membrane; ch, chorion; fc, follicle cell. (Figure adapted from portions of Fig. 6, Pascucci et al., 1996.)

extensive protein turnover. Whereas immune reactivity has proven to be problematic in stage 14 chorions (Trougakos and Margaritas, 1998b; Waring, 1999), several vitelline membrane epitopes have been readily detected in situ in late stage 14 egg chambers (Waring, 1999; Mauzy-Melitz and Waring, unpublished). Taken together, these data indicate that most of the s36 secreted by the follicle cells is not retained in the follicle cell proximal space for immediate assembly into the forming chorion but rather makes its way to the vitelline membrane layer where it becomes transiently sequestered. During late choriogenesis, the sequestered molecules are released and available for incorporation into the growing structure. Because small amounts of s36 (10–20%) appear to be sufficient for fertility (Digan et al., 1979), the s36 utilized for assembly during the early choriogenic stages may be functionally distinct from that incorporated late. The small proportion utilized early may be essential for establishing the framework, whereas that utilized late may intercalate along with the late chorion proteins (e.g., s18) into the preexisting framework. The incorporation of D. virilis early chorion

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proteins into the vitelline membrane has recently been reported by Trougakos and Margaritis (1998b). Using a rabbit antiserum directed against the D. virilis homologs of s36 and s38, Trougakos and Margaritis showed an immunolocalization pattern similar to that described for s36 in D. melanogaster, namely immunogold labeling over both the vitelline membrane and chorion in early choriogenic egg chambers followed by its restriction to the endochorion in stage 14 egg chambers. 2. Sequestration and Trafficking of dec-1 Proteins The importance of the dec-1 proteins in eggshell assembly is underscored by the lack of a structured endochorion framework and the eventual collapse of chorion material into the underlying vitelline membrane layer in dec-1 null mutants. Although peak dec-1 gene expression occurs during the stages of vitelline membrane formation, the genesis of mature dec-1 proteins during the choriogenic stages is compatible with the altered chorion morphology seen in dec-1 mutants. The multiplicity of dec-1 proteins raised the question of whether all have separate and essential roles in eggshell assembly or whether there is functional redundancy. Genetic data indicate several dec1 derivatives have distinct functions. The breakpoint of the deficiency chromosome Df(1)ct4bl falls within the dec-1 gene in the fc177 specific C-terminal region (Hawley and Waring, 1988). Because the breakpoint lies 3⬘ of the open reading frame for fc106, the deficiency chromosome produces wildtype quantities of fc106 and its derivatives (Waring et al., 1990). Females heterozygous for the deficiency chromosome and a dec-1 null allele such as fs(1)410 are sterile. The sterility of the Df(1)ct4bl/fs(1)410 transheterozygotes suggests normal expression of fc106 and its derivatives is not sufficient to produce a functional eggshell. At least one fc016 derivative is necessary because fs(1)1501 mutants produce nonfunctional eggshells despite production of the larger fc125 and fc177 proteins and their derivatives. s25 has been selectively eliminated by introducing a dec-1 transgene bearing an N-terminal deletion into homozygous fs(1)1501 females. The N-terminal deletion produces a 81-kDa pro-protein instead of fc106. Despite the normal appearance of s80 and s60 in fs(1)1501 females carrying the altered transgene, fertility is not restored (Mauzy-Melitz and Waring, unpublished). This suggests that s25, the N-terminal protein produced by the cleavage of fc106, has an essential function. Premature stop codons have been engineered separately into the fc177- and fc125-specific open reading frames to create dec-1 transgenes deficient for the s85 and s95 derivatives, respectively. Both mutant transgenes failed to rescue the sterility of fs(1)410 dec1 null mutant females, suggesting that both C-terminal derivatives, s95 and s85, are required for proper eggshell function (Mauzy-Melitz and Waring, unpublished).

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The distinct roles of the multiple dec-1 proteins in eggshell assembly implied by the genetic data are consistent with immunolocalization data. In wild-type egg chambers, fc106 and its derivatives are the major proteins produced by the dec-1 locus. The distribution of fc106 and its derivatives was followed with antisera directed against trypE fusion proteins containing peptides from different regions of fc106: the N-terminus of fc106 (Nfc106), the N-terminus of s80 (N-s80), and the C-terminus of fc106 (Cfc106) (Noguero´n and Waring, 1995). fc125, fc177, and their derivatives are recognized by some of these sera. However, because fc106 derivatives accumulate in wild-type egg chambers in amounts 10- to 20-fold greater than either fc125 or fc177 derivatives, fc106 and its derivatives presumably represent the predominant reactive species. Immunogold staining showed that fc106 is synthesized in the follicle cells, packaged in secretory vesicles, delivered to the follicle cell surface, and immediately secreted into the extracellular space where it becomes associated with the forming vitelline membrane (Noguero´n, 1996). During late stage 10, fc106 is cleaved within the vitelline membrane layer to its s25 N-terminal derivative and its s80 C-terminal derivative. s25 reacts with the Nfc106 serum, and s80 reacts with either the Nfc80 or Cfc106 serum. In stage 12 egg chambers, s25 localized both within the vitelline membrane layer and in the forming ICL-endochorion complex. In stage 14 egg chambers, s25 was found in the endochorion and ICL (Fig. 10A–C). Although eggshell peroxidase has been cytochemically localized in the ICL (Mindrinos et al., 1980; Trougakos and Margaritis, 1998b), s25 is the only eggshell protein that has been immunolocalized in this layer (Pascucci et al., 1996; Trougukas and Margaritis, 1998a; Noguero´n, 1996). The presence of s25 in the ICL is compatible with defects in the crystalline substructure of the ICL in fs(1)384 dec-1 mutants reported by Margaritis et al. (1991). The regional distribution of s25 has been assessed at the light microscope level by indirect immunofluoresence using biotin-labeled secondary antibodies complexed with streptavidin-alkaline phosphatase in conjunction with a fast red substrate. Aside from the mainshell, intense staining was evident in the micropyle and operculum regions (Fig. 10D). The behavior of the C-terminal derivative, s80, was distinct from s25 in that it remained homogeneously distributed within the vitelline membrane layer during stages 12 and 13. During late stage 13 and stage 14, s80 is cleaved at its N-terminus-yielding s20 and s60. The Cfc106 serum was used to track s60 in stage 14 egg chambers, and the Ns80 serum was used to track s20. Immunolocalization data suggested that even though some s60 was released to the endochorion layer in stage 14 egg chambers, a substantial portion remained in the vitelline membrane (Noguero´n, 1996). The partitioning of s60 between the two layers was verified by biochemical fractionation experiments. The chorion

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FIG. 10 Multilayered distribution of the N-terminal dec-1 derivative, s25, in late stage 14 egg chambers. (A) Indirect immunofluorescent staining of sectioned stage 14 egg chambers reacted with an antiserum directed against sequences in the N-terminus of fc106. In stage 14 egg chambers, this serum recognizes s25. Intense staining in the chorion (ch) and vitelline membrane(vm) layers is shown in A. Immunoelectron microscopy of the chorion layer shows immunogold particles over the endochorion (en) (B) and inner chorionic layer (icl) (C). Fluorescence microscopy also revealed intense staining in the micropyle (mp) and operculum (op) (D). The staining seen in the oocyte (oo) in A presumably reflects reactivity to a less abundant fc177 derivative as the oocyte does not stain in stage 14 egg chambers from Df(1)ct4bl/ fs(1)410 transheterozygotes but stains with comparable intensity in those derived from fs(1)1501 mutants (Mauzy-Melitz and Waring, unpublished).

was manually removed from stage 14 egg chambers following osmotic shock. Western blot analysis of the resulting chorion and vitelline membranebound oocytes revealed significant amounts of s60 in both fractions. In marked contrast to s60, the N-terminal derivative, s20, disappeared rapidly from the vitelline membrane layer after its genesis. Stage 14 egg chambers

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incubated with the Ns80 serum showed background labeling over the vitelline membrane layer and intense labeling of vesicles located within the peripheral regions of the oocyte (Noguero´n, 1996). Trafficking of s20 to the oocyte was confirmed by biochemical fractionation experiments. Western blot analysis showed the bulk of the s20 reactivity fractionated with the vitelline membrane-bound oocyte. Almost all the reactivity was lost when the vitelline membrane-bound oocyte was punctured and the oocyte contents were released (Noguero´n and Waring, unpublished). The multilayered distribution of s25 and s60 contrasts with the restricted distribution of sV23, sV17, s36, s18, s19, and s15 in the mature structure. Although s36 is found in both the vitelline membrane and the forming chorion during the early stages of eggshell formation, its complete removal from the vitelline membrane during late choriogenesis contrasts with s25 and s60. The functional significance of the multilayered distribution of s25 and s60, as well as the molecular basis for their gradual and incomplete release from the vitelline membrane, remain unknown. The immediate and seemingly complete release of s20 suggests that sequences in this region either do not play a role or are inadequate to tether s80 to the vitelline membrane. Although sufficient for release of s20, cleavage may be necessary but is certainly not sufficient for the redistribution of s60. Secondary modifications of s60 may be required for its movement from the vitelline membrane to the endochorion. The differential uptake of s20 by the oocyte may reflect late-stage pinocytotic activity by the oocyte or may be a receptormediated endocytotic process. In either case, the uptake suggests a mechanism by which the oocyte can continue to receive information from the follicle cells after the morphological completion of the vitelline membrane, a structure that has previously been perceived as an impenetrable physical barrier. The uptake of s20 may be functionally significant or it may represent a mechanism by which proteins in the extracellular milieu that do not become incorporated into the egg shell are removed. Oocyte uptake as a removal mechanism has been implicated by the distribution of the s85 dec1 derivative in an overproducing mutant. To facilitate immunolocalization of fc177 and its derivatives, a transgene consisting of fc177 cDNA driven by dec-1 regulatory sequences was created (Mauzy-Melitz, unpublished). In transgenic females, large quantities of fc177 are prematurely produced and processed in stage 10 egg chambers. Immunofluorescent staining showed fc177 and its derivatives associate with the vitelline membrane at this stage. In addition, intense staining was observed in the oocyte of stage 10 egg chambers. In wild-type egg chambers, fc177 and/or s85 initially accumulate in the vitelline membranes of stage 12 egg chambers. Unlike the overproducer, oocyte staining has not been detected with the fc177specific serum. This suggests that the amount of fc177/s85 that can be accommodated by the vitelline membrane at stage 10, and perhaps stage

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12, is limiting. In the overproducer, molecules which do not become initially incorporated into the vitelline membrane are taken up by the oocyte and thus removed from the assembly process.

C. Quantitative Considerations The s38 and s36 proteins accumulate in choriogenic egg chambers in proportion to the gene dosage. Introduction of an extra copy of the 7EF region via an insertional translocation resulted in a 1.5-fold increase in s36 and s38 relative to wild type; egg chambers from females heterozygous for a deficiency covering the 7EF region showed a 0.5⫻ decrease. In either case, there were no deleterious effects on eggshell assembly or function. Two copies of transgenes encoding either sV23 or dec-1 proteins have been successfully introduced into wild-type genetic backgrounds via P-elementmediated transformation without consequence. In some of the transformant lines, the transgenes were expressed at wild-type levels, suggesting that a twofold increase in these proteins is not deleterious. Substantial increases in the production of the minor dec-1 proteins, fc125 and fc177 (5–10⫻), were also tolerated in a wild-type genetic background. Although limited, data to date indicate that overproduction of the eggshell proteins does not interfere with the assembly or the function of the eggshell. The recovery of only null mutations in eggshell structural genes from numerous female sterile screen suggests that hypomorphic mutations may not be associated with an identifiable phenotype. Limited data are available on the minimum amounts of eggshell proteins that are needed to assemble a functional eggshell. Analysis of cor36/oc transheterozygotes suggests that less than 20% of wild-type levels of the s36 chorion protein are needed for proper eggshell function. The ocelliless (oc) mutation is associated with a small chromosome inversion whose distal breakpoint lies within a 90-kb central region which normally undergoes amplification in ovarian follicle cells (Spradling, 1981). In oc females, the EcoR1 fragment carrying the s36 gene is underreplicated two- to three-fold in late-stage egg chambers (Spradling and Mahowald, 1981). At the protein level, there is an approximate fivefold reduction in the accumulation of both major chorion proteins encoded by the region, s36 and s38 (Digan et al., 1979). Morphologically, egg chambers from homozygous oc females appear to retain a roof network but show a marked reduction in the number of endochorion pillars and lack the endochorion floor. Fertility and near wild-type eggshell morphology are restored in oc/cor 36 egg chambers. Although failing to produce s36, the cor36 chromosome complements other genetic defects associated with the oc mutation. This result suggests that, although necessary, small amounts of s36 (e.g., 10–20% of wild-type levels) are sufficient for proper

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eggshell assembly and function. While small amounts of s36 are sufficient for function in a background of near wild-type levels of the other eggshell proteins, the oc phenotype and the plethora of eggshell gene amplification mutants that have been recovered from female sterile screens suggests that threshold levels of other eggshell proteins are necessary for proper eggshell function (Orr et al., 1984). Based on the morphological and biochemical analyses of the K451 and K1214 amplification mutants (Komitopoulou et al., 1988), Trougakos and Margaritis (1998a) suggest that thin but organized endochorions can be formed unless chorion protein production drops below 10% of wild-type levels, in which case complete endochorion disorganization occurs.

D. Stabilization of the Eggshell In the mature egg, eggshell proteins are insoluble in denaturing solutions (2% SDS, 8M urea or 6M guanidine HCl). The chorion becomes insoluble during late stage 14, and its insolubility has been correlated with the appearance of di- and trityrsoine residues in eggshell hydrolysates (Petri et al., 1976). Insolubilization does not occur simultaneously throughout the eggshell; it is initiated at the poles and proceeds into the central mainshell region. Using an in vitro culturing system, Mindrinos et al. (1980) showed that endochorion proteins remain soluble if phloroglucinol, a potent inhibitor of peroxidase, is included in the culture medium. At the morphological level, the mature ICL also remains soluble provided that egg chambers are developed in the presence of phloroglucinol and 웁-mercaptoethanol is included in the solubilization buffer. Peroxidase activity has been detected histochemically in isolated endochorions from stage 14 egg chambers. Histochemical staining of ovarian whole mounts revealed peroxidase activity in the regions of the forming dorsal appendages as early as stage 11. By stages 13 and 14 the entire eggshell was heavily stained. More refined ultrastructural analysis revealed peroxidase activity in follicle cell secretory granules associated with chorion deposition in stage 11 egg chambers. Staining was associated only with the ICL and endochorion layer in choriogenic stage eggchambers. Peroxidase activity was never detected in the vitelline membrane layer. Trougakas and Margaritis (1998b) reported identical patterns of peroxidase activity in Drosophila virilis egg chambers. Immunolocalization of peroxidase in D. melanogaster using rabbit anti-horseradish peroxidase verified its absence in the vitelline membrane and its presence throughout the endochorion complex and ICL (Keramaris et al., 1991). Although peroxidase activity can clearly be demonstrated during the early choriogenic stages by the introduction of exogenous substrates, natural tyrosine cross-linking commences in stage 14 egg chambers. The natural

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trigger appears to be the production and secretion of hydrogen peroxide from the follicle cell plasma membrane during early stage 14 (Margaritis, 1985). Following secretion, hydrogen peroxide diffused throughout the chorion layers but was excluded from the vitelline membrane. Taken together, these data suggest that chorion proteins in both the ICL and endochorion are crosslinked via the formation of peroxidase-catalyzed interpolypeptide di- and trityrosine residues. Stabilization of the ICL appears to also entail disulfide bond formation. Western blot analysis shows that the major vitelline membrane proteins and their derivatives remain soluble well into stage 14 (Pascucci et al., 1996). Although the means by which vitelline membrane proteins become insoluble has not been established, the strict evolutionary conservation of cysteine residues within the vm domain suggests that disulfide linkages may be involved. It has been suggested that dec-1 derivatives which include the central glutamine-rich repeats may be cross-linked to other eggshell proteins via glutamyl-lysine cross-links (Waring et al., 1990). Glutamyl-lysine cross-links catalyzed by transglutaminases stabilize a variety of biological structures including the keratinocyte envelope (Eckert and Green, 1986), fibrin clots (Lorand and Conrad, 1984), and ionophore-induced hardening of the erythrocyte membrane (Lorand et al., 1976). Even though transglutaminase activity in egg chamber extracts has yet to be documented, it has been speculated that s36 and s38 with their evolutionarily conserved lysinerich central domains could provide the requisite amino acid residues. The similar distributions of s60 and s36 throughout the endochorion complex is compatible with this speculation. Apart from regional differences in the timing of eggshell insolubilization, there is variability in the timing in which individual eggshell proteins become insoluble. Western blot analysis indicates that three dec-1 derivatives–s25, s85, and s95—become insoluble prior to stage 14. Soluble s25 and s95 are produced in late stage 10 egg chambers and soluble s85 is produced during stages 12–13. Although all the products remain reactive with the appropriate dec-1 antiserum in situ through stage 14 (Mauzy-Melitz, unpublished), s25 becomes insoluble in SDS and 웁-mercaptoethanol-containing buffers at the end of stage 12, whereas s85 and s95 are no longer soluble after stage 13 (Noguero´n and Waring, 1995). Proteins within the vitelline membrane and chorion layers become crosslinked into insoluble matrices through the formation of covalent bonds. The collapsed eggshell phenotypes of the dec-1 and sV23 null mutants suggests that insolubilization may be necessary but is not sufficient to form a stable structure. In dec-1 mutants, an organized endochorion never forms, and chorionic material collapses into the vitelline membrane in late stage 14 egg chambers (Bauer and Waring, 1987). A disorganized endochorion in and of itself is not sufficient to cause a collapse as cor36 mutants fail

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to organize a tripartite endochorion yet accumulations of electron-dense chorionic material in the vitelline membrane are not observed (Digan et al., 1979). dec-1 proteins contribute to all three proteinaceous layers of the eggshell, the vitelline membrane, the ICL, and the endochorion. This multilayered distribution is compatible with the multilayer defects inferred by the dec-1 null morphological phenotype. Like dec-1 null mutants, sV23 null mutants show a collapse of endochorionic material into the vitelline membrane in late stage 14 egg chambers. The endochorionic nature of the electron-dense globules found in late stage vitelline membranes was confirmed by their reactivity with s18 and s36 antisera (Pascucci et al., 1996). Because sV23 has been immunolocalized only within the vitelline membrane layer, the multilayered defects suggested by the morphological analyses suggest that at least one vitelline membrane protein, sV23, plays a prominent role in the stabilization of the outer endochorion layers (Pascucci et al., 1996). With the ICL intervening between the vitelline membrane and endochorion layers, direct molecular interactions would appear to be precluded. Perhaps proteins within the ICL provide a molecular means to bridge vitelline membrane and endochorion components. Alternatively, late-stage processing of the sV23 protein may release a transient peptide which plays a critical role in stabilizing the outer eggshell. A third more intriguing possibility is that chorion subparticles form within the vitelline membrane layer during the early and mid choriogenic stages. During late choriogenesis, the subparticles are released and assemble into the structured endochorion. The immunolocalization of s60, s36, and s38 chorion proteins within the vitelline membrane layer during these stages is compatible with this possibility. Perhaps the repeating octapeptide in sV23 serves as an interactive surface upon which the subparticles can assemble. In the absence of sV23, aberrant subparticles assemble which, when released, form an unstable structure that collapses into the vitelline membrane. This model suggests that early protein–protein interactions within the vitelline membrane layer directed by a vitelline membrane protein(s) are critical for stabilization of the outer chorion layers during stage 14.

VII. Concluding Remarks Drosophila eggshell formation will provide insights into molecular strategies that are used in higher eukaryotes to build complex extracellular architectures in vivo, where components are assembled at physiological concentrations and where proper protein–protein interactions are established in a multicomponent environment. Significant strides have been made in identifying the major structural components, in uncovering complex postdeposi-

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tional protein-processing pathways, and in revealing unanticipated molecular dynamics in the vitelline membrane layer after its formation at the end of stage 10. Table II summarizes the molecular changes in the vitelline membrane layer that have been documented to date. At the very least, the temporally regulated dec-1 cleavages control the timing of the appearances of the mature dec-1 products. The functional significance of the processing of the vitelline membrane proteins, sV17 and sV23, has yet to be addressed. The enzymatic machinery utilized for these processing events as well as the means by which the cleavages are temporally regulated remain intriguing questions. How chorion components are anchored within the vitelline membrane as well as the trigger(s) for their release remain challenges for the future. If chorion subparticles assemble in the vitelline membrane, what proteins or motifs are used to nucleate the process? dec-1 derivatives differentially sort after their release from the vitelline membrane layer. What molecular tags instruct s25 to incorporate into the ICL, while others such as s20 are directed to the oocyte? The next layer in the analysis of eggshell formation will be defining protein–protein interactions and their interacting motifs. This can be accomplished by a combination of biochemical and genetic approaches. Using evolutionarily conserved motifs as a guide, by genetically engineering mutations, the sequences required for anchoring and trafficking specific proteins to their proper destinations can be established. The yeast two-hybrid system offers a promising means to isolate genes encoding interacting partners. Using domains deemed functionally significant by genetic analyses as bait, it should be possible to fish out genes encoding interacting proteins. The authenticity of the interaction can be verified by colocalizing tagged gene products in the morphological structure. The functional significance of

TABLE II Changes in the Molecular Composition of the Vitelline Membrane Layer during the Stages of Eggshell Formation Stages

Proteins

10

12

14

sV23 sV17 fc106

sV23 sV17 s80 s25 s36 s38

sV23 (17 kDa) sV17 (10 kDa) s60 s25 ND ND

The approximate sizes of the smaller stage 14 vitelline membrane derivatives are indicated (17 kDa for sV23 and 10 kDa for sV17). ND indicates that no reactivity was detected in the vitelline membrane layer at this stage.

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the interaction in the assembly process will ultimately require obtaining mutations in the genes which encode interacting proteins. By exploiting the genetic advantages of Drosophila and by making use of the spatial resolution provided by this large structure, the eggshell promises to continue to yield molecular insights into in vivo assembly processes in a developing system.

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Rittenhouse, K. R., and Berg, C. A. (1995). Mutations in the Drosophila gene bullwinkle cause the formation of abnormal eggshell structures and bicaudal embryos. Development 121, 3023–3033. Roebroek, A. J., Ayoubi, T. A., Creemers, J. W., Pauli, I. G., and Van de Ven, W. J. (1995). The Dfur2 gene of Drosophila melanogaster: Genetic organization, expression during embryogenesis, and pro-protein processing activity of its translational product Dfurin2. DNA Cell Biol. 14, 223–234. Sappington, T. W., and Raikhel, A. S. (1998). Molecular characteristics of insect vitellogenins and vitellogenin receptors. Insect Biochem. Mol. Biol. 28, 277–300. Savant, S. S., and Waring, G. L. (1989). Molecular analysis and rescue of a vitelline membrane mutant in Drosophila. Dev. Biol. 135, 43–52. Scherer, L. J., Harris, D. H., and Petri, W. H. (1988). Drosophila vitelline membrane genes contain a 114 base pair region of highly conserved coding sequence. Dev. Biol. 130, 786–788. Schupbach, T., and Wieschaus, E. (1991). Female sterile mutations on the second chromosome of Drosophila melanogaster. II. Mutations blocking oogenesis or altering egg morphology. Genetics 129, 1119–1136. Spradling, A. C. (1981). The organization and amplification of two chromosomal domains containing Drosophila chorion genes. Cell 27, 193–201. Spradling, A. C., and Mahowald, A. P. (1981). A chromosome inversion alters the pattern of specific DNA replication in Drosophila follicle cells. Cell 27, 203–209. Spradling, A. C. (1993). Developmental genetics of oogenesis. In ‘‘The Develoment of Drosophila melanogaster’’ (M. Bates and A. Martinez Arias, Eds.), Vol. 1, pp. 1–70. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Spradling, A. C., and Mahowald, A. P. (1979). Identification and genetic localization of mRNAs from ovarian follicle cells of Drosophila melanogaster. Cell 16, 589–598. Spradling, A. C., Digan, M. E., Mahowald, A. P., Scott, M., and Craig, E. (1980). Two clusters of genes for major chorion proteins of Drosophila melanogaster. Cell 19, 905–914. Spradling, A. C., Waring, G. L., and Mahowald, A. C. (1979). Drosophila bearing the ocelliless mutatation under produce two major chorion proteins both of which map near this gene. Cell 16, 609–616. Swimmer, C., Fenerjian, M. G., Martinez-Cruzado, J. C., and Kafatos, F. C. (1990). Evolution of the autosomal chorion cluster in Drosophila. III. Comparison of the s18 gene in evolutionarily distant species and heterospecific control of chorion gene amplification. J. Mol. Biol. 215, 225–235. Tolias, P. P., Konsolaki, M., Komitopoulou, K., and Kafatos, F. C. (1990). The chorion genes of the medly, Ceratitis capitata. II. Characterization of three novel cDNA clones obtained by differential screening of an ovarian library. Dev. Biol. 140, 105–112. Trougakos, I. P., and Margaritis, L. H. (1998a). The formation of the functional chorion structure of Drosophila virilis involves intercalation of the ‘‘middle’’ and ‘‘late’’ major chorion proteins: A general model for chorion assembly in Drosophilidae. J. Struct. Biol. 123, 97–110. Trougakos, I. P., and Margaritis, L. H. (1998b). Immunolocalization of the temporally ‘‘early’’ secreted major structural chorion proteins, Dvs38 and Dvs36, in the eggshell layers and regions of Drosophila virilis. J. Struct. Biol. 123, 111–123. Twombly, V., Blackman, R. K., Jin, H., Graff, J. M., Padgett, R. W., and Gelbart, W. M. (1996). The TGF-beta signaling pathway is essential for Drosophila oogenesis. Development 122, 1555–1565. Vlachou, D., Konsolaki, M., Tolias, P. P., Kafatos, F. C., and Komitopoulou, K. (1997). The autosomal chorion locus of the medfly, Ceratitis capitata. I. Conserved synteny, amplification and tissue specificity but sequence divergence and altered temporal regulation. Genetics 147, 1829–1842.

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Waring, G. L. (1999). Eggshell assembly in Drosophila. In ‘‘A Comparative Methods Approach to the Study of Oocytes and Embryos’’ ( J. D. Richter, Ed.), pp. 385–396. Oxford University Press, New York. Waring, G. L., and Mahowald, A. P. (1979). Identification and time of synthesis of chorion proteins in Drosophila melanogaster. Cell 16, 599–607. Waring, G. L., Hawley, R. J., and Schoenfeld, T. (1990). Multiple proteins are produced from the dec-1 eggshell gene in Drosophila by alternative RNA splicing and proteolytic cleavage events. Dev. Biol. 142, 1–12. Wasserman, J. D., and Freeman, M. (1998). An autoregulatory cascade of EGF receptor signaling patterns the Drosophila egg. Cell 95, 355–364. Zarini, F. E., and Margaritis, L. H. (1986). The eggshell of Drosophila melanogaster. Structure and morphogenesis of the micropylar apparatus. Can. J. Zool. 64, 2509–2519.

Cell Biology of Cytochrome P-450 in the Liver Shinsuke Kanamura and Jun Watanabe Department of Anatomy, Kansai Medical University, 10-15 Fumizono-cho, Moriguchi, Osaka 570-8506, Japan

Cytochromes P-450 (P-450) are members of a multigene superfamily of hemoproteins consisting the microsomal monooxygenase system with NADPH P-450 reductase (reductase) and/or reducing equivalents. Expression of many P-450 isoforms in hepatocytes is shown to be regulated at the level of transcription through interaction between cis-acting elements in the genes and DNA-binding (transacting) factors. Some isoforms of the CYP1A, 2B, 2E, and 3A subfamilies are regulated at the posttranscriptional level. For the topology of P-450 and reductase molecules in ER membrane of hepatocytes, models from stopped flow analysis and electron spin resonance are proposed. The densities of total P-450 and reductase molecules are revealed to be high enough to support the cluster model, suggesting that about ten P-450 molecules form an aggregate and surround one reductase molecule, and therefore the two enzymes form large micelles. ER proliferation after PB administration, which had been correlated with increase in P-450 level, is shown to be probably independent of the increase in P-450 level. There are considerable discrepancies among results reported on sublobular expression of various P-450 isoforms. Causes of the discrepancies are likely to be differences in experimental conditions of histochemical detection carried out and/or in species, strain, and/or sex. KEY WORDS: Cytochrome P-450, Hepatocyte, Gene regulation, Topology, Endoplasmic reticulum. 䊚 2000 Academic Press.

I. Introduction Cytochromes P-450 (P-450) are members of a multigene superfamily of hemoproteins and, together with NADPH cytochrome P-450 reductase International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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Copyright 䉷 2000 by Academic Press All rights of reproduction in any form reserved.

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(reductase) and/or reducing equivalents, are composed of the microsomal monooxygenase system. The microsomal monooxygenase system plays a role in the metabolism of a variety of xenobiotics and compounds such as drugs, carcinogens, environmental pollutants, natural plant products and alcohols and of endogenous compounds such as steroids, bile acids, fatty acids, prostaglandins, leukotrienes, and biogenic amines (Nebert et al., 1991). At least, 12 gene families with 19 subfamilies have been identified in mammals on the basis of amino acid sequence homology (Nebert et al., 1991; Nelson et al., 1993). A characteristic of P-450 is their inducibility by numerous structurally different chemicals. Phenobarbital (PB) and 3⬘methylchoranthrene (MC) are the representative inducing agents. The liver contains principally high levels of P-450 and is consequently the major site of the metabolism. However, P-450 at lower concentrations is also found in nearly every extrahepatic organs such as lung, kidney, and brain (Guengerich and Mason, 1979; De Waziers et al., 1990; Zerilli et al., 1995). Generally, P-450 isoforms are not uniformly expressed in all hepatocytes in the liver lobule and are present predominantly in perivenular hepatocytes. The majority of mammalian P-450 isoforms are bound to ER membranes (Marti et al., 1990; Nebert et al., 1991; Persohn et al., 1993). However, evolutionally closely related steroid metabolizing enzymes—CYP10, CYP11A, CYP11B, and CYP27—are located in the mitochondrial membrane (Nebert et al., 1991). In addition, CYP4A1 is localized in peroxisomes and mitochondria (Persohn et al., 1993). Much information, including numerous reviews, has been published on the nature and role of P-450. However, there is only one article on cellbiological aspects on P-450 (Tashiro et al., 1993), and it discusses ‘‘membrane topology, biosynthesis, intracellular distribution, induction and turnover and degradation’’ of P-450. In the present review, we describe new information on ‘‘regulatory mechanisms for P-450 expression in hepatocytes,’’ ‘‘membrane topology of P-450 and reductase in ER membrane of hepatocytes,’’ and ‘‘sublobular expression of P-450 in the liver’’ in mammals.

II. Regulatory Mechanisms for P-450 Expression in Mammalian Hepatocytes A. Transcriptional Regulation Expression of many P-450 isoforms is shown to be regulated at the level of transcription through interaction between cis-acting (cis) elements in P450 genes and DNA-binding (transacting) factors (Table I). Several cis elements have been found in the 5⬘ flanking region or intron of genes

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CELL BIOLOGY OF CYTOCHROME P-450 IN THE LIVER TABLE I Major cis Elements in the 5⬘ Flanking Region of Cytochrome P-450 Genes

Isoform 1A1 1B1 2B1/2/10 2C2 2C6 2C11/12 2C13 2D5 2D9

2E1 3A1 3A4/7

3A5 3A10 3A23 4A1/6 7A

cis element XRE BTE XRE Proximal promoter PBRE HPF-1 motif DBP-binding site GHNF-binding site HNF-3-binding site HNF-1-binding site HNF-3-binding site ⫺112/-83 element ⫺110/⫺92 element Demethylated ⫺100/⫺93 element NF-Y-binding site HNF-1-binding site Dexamethasoneresponsive element NFSE HFLaSE BTE Upstream GRE half-site STAT-binding element NF-Y-binding site Steroid receptor motif PPRE DBP-binding site

Transacting factor or (possible transacting factor)

Drug receptor

Ah receptor/Arnt complex (Sp1), BTEB Ah receptor/Arnt complex (28kD protein, C/EBP) (NF1) (HPF-1 ⫽ HNF-4) (DBP) (GHNF or STAT 1, 3 and 5) (HNF-3 or HNF-6) (HNF-1) (HNF-3) C/EBP beta-Sp1 complex NF2d9 GABP

Ah receptor ? Ah receptor ? ? ? ? ? ? ? ? ? ? ?

(NF-Y) (HNF-1) ? ? ? BTEB (GR) STAT 5a/5b (NF-Y) ? PPAR/RXR alpha complex (DBP or C/EBP)

? ? ? ? ? ? (GR) ? ? ? PPAR ?

XRE, xenobiotic-responsive element; BTE, basic transcription element; Ah receptor, arylhydrocarbon receptor; Arnt, arylhydrocarbon receptor nuclear translocator; Sp1, specificity protein 1; BTEB, basic transcription element binding factor; C/EBP, CCAATT enhancer binding protein; PBRE, phenobarbital-responsive element; NF-1, nuclear factor 1; HPF-1; HepG2 nuclear protein; HNF-4, hepatic nuclear factor-4; DBP, D-site binding protein; GHNF, growth hormone-regulated nuclear factor; STAT, signal transducers and activators of transcription; HNF-1, hepatic nuclear factor-1; HNF-3, hepatic nuclear factor-3; HNF-6, hepatic nuclear factor-6; NF2d9, nuclear factor for CYP2D9; GABP, gamma-aminobutyric acid-binding protein; NFSE, P450NF-specific element; HFLaSE, P450HFLa-specific element; GRE, glucocorticoid-responsive element; NF-Y, nuclear factor Y; GR, glucocorticoid receptor; PPRE, peroxisome proliferator-responsive element; PPAR, peroxisome proliferator activator receptor; RXR alpha, retinoid X receptor-alpha.

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encoding P-450 isoforms. Induction of the CYP1A1 by interaction between cis elements and dioxin-activated DNA-binding factor in rat hepatocytes has become a model for understanding mechanisms for the xenobioticsinduced gene regulation. 1. CYP1 Family a. CYP1A1 a. Cis Elements P-450 isoforms encoded by the CYP1A gene subfamily are induced markedly by polycyclic aromatic hydrocarbons such as 3methylchoranthrene (MC), benzopyrenes, heterocyclic amines and dioxins. There are at least four types of cis elements in CYP1A genes; xenobioticresponsive element (XRE), glucocorticoid-responsive element (GRE), negative regulatory element (NRE) and basic transcription element (BTE). XRE acts as a potent enhancer (Fujisawa-Sehara et al., 1986). There are six XRE sequences containing CACGC core motif in the 5⬘ flanking region in CYP1A1 gene (Denison et al., 1989). GRE acts as an enhancer with incorporation of XRE (Mathis et al., 1989). A GRE sequence is found in the first intron of the gene (Mathis et al., 1989). NRE acts as a silencer and is located in the 5⬘ flanking region (Bhat et al., 1996). BTE contains motif identical to GC box, and is essential for full activation of CYP1A1 gene transcription through the XRE (Yanagida et al., 1990). BTE is localized at upstream side of TATA box in the 5⬘ flanking region (Yanagida et al., 1990). b. Transacting Factor for XRE Transcription of CYP1A gene is activated by binding of DNA-binding factor to XRE (Fujisawa-Sehara, 1986). The XRE-binding factor is a transiently formed heterodimer of arylhydrocarbon (Ah) receptor and Ah receptor nuclear translocator protein (Arnt) (Probst et al., 1993). The Ah receptor–Arnt heterodimer is designated as an XRE-binding protein (Fujisawa-Sehara, 1986). Alone an Ah receptor or Arnt cannot interact with XRE (Pollenz et al., 1993; Reisz-Porszasz et al., 1994; Bacsi et al., 1995). The Ah receptor and Arnt contain nuclear targeting signals and period–Arnt–single (PAS) domains (Burbach et al., 1992). The two proteins also contain basic-helix-loop-helix (bHLH) motifs near their amino termini (Burbach et al., 1992). Therefore, Ah receptor and Arnt have been classified into members of bHLH/PAS transcription factor family (Poellinger, 1995). The basic domain in the bHLH of Ah receptor or Arnt is responsible for XRE binding (Whitlock et al., 1996). The helix-loop-helix domain in the proteins may form a dimerization interface (Bacsi et al., 1995; Swanson et al., 1995). Furthermore, a transcription-activating domain has been shown to localize to the carboxyl terminus of both proteins (Whitelaw et al., 1994; Sogawa et al., 1995). However, Reisz-Porszasz et al. (1994) and Ko et al. (1996) reported that the transcription-activating domain of Arnt plays mini-

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mal role in up-regulation of the CYP1A1, although Whitelaw et al. (1994) indicated that the domain is crucial for the up-regulation. c. Other Transacting Factors There are two possible binding factors for BTE. One is specificity protein 1 (Sp1), a common transcription factor for GC box (Imataka et al., 1992). Sp1 contains zinc-finger domains to interact with Ah receptor and Arnt (Kobayashi et al., 1996). The other is BTEbinding protein (Imataka et al., 1992). The protein consists of 244 amino acids and also contains zinc-finger domains in its carboxyl terminus (Imataka et al., 1992). In addition, a binding factor for NRE is shown to be octamer-binding transcriptional factor 1 (Oct-1) (Bhat et al., 1996). The factor for GRE is however presently not clearly defined. d. Regulatory Mechanism Through XRE and XRE-Binding Protein The Ah receptor forms a complex with a molecular chaperon, heat-shock protein 90 (HSP90), in the hepatocyte cytoplasm before inducing the CYP1A1 (Whitelaw et al., 1993). The PAS domain of the receptor plays an important role in its binding to HSP90 or inducer molecules such as MC or dioxin (Whitelaw et al., 1993). HSP90 masks the nuclear targeting signal within Ah receptor (Whitelaw et al., 1993). On the other hand, Arnt does not interact with HSP90 (Dogra et al., 1998). After stimulation of dioxin or MC, the Ah receptor-HSP90 complex binds to the inducer molecule (Dogra et al., 1998). Binding the inducer molecule to the complex may induce conformational change in the complex which allows exposure of the nuclear targeting signal and subsequent nuclear translocation of the receptor. Dogra et al. (1998) postulated that HSP90 is released from the Ah receptor after the nuclear translocation of the complex. However, Ma and Whitlock (1997) and Carver and Bradfield (1997) found in the cytoplasm nobel molecular chaperons, AIP and ARA9, to influence the nuclear targeting of Ah receptor. Thus, the Ah receptor may release HSP90 in the cytoplasm before the nuclear translocation and acquires ability to translocate from the cytoplasm to the nucleus. Previously, both Ah receptor and Arnt were considered to localize in the cytoplasm of hepatocytes. Thus, the following mechanism had been considered (Fujisawa-Sehara et al., 1986; Hankinson, 1995). After stimulation of MC or dioxin, the Ah receptor associates with Arnt in the cytoplasm, and the resulting Ah receptor-Arnt heterodimer (XRE-binding protein) translocates to the nucleus to bind to XRE. However, immunohistochemical studies of Pollenz et al. (1993) and Hord and Perdew (1994) showed exclusive localization of Arnt in the nucleus of Hepa-1 cells before and after dioxin stimulation. Furthermore, Asaka et al. (1998) found by in situ Southwestern hybridization that XRE-binding protein is exclusively localized in the nucleus, and therefore Ah receptor and Arnt does not form the binding protein in the cytoplasm but in the nucleus of hepatocytes. Thus, Arnt is not the nuclear translocator but a nuclear protein.

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The major role of XRE-binding protein is to disrupt the chromatin structure of the CYP1A1 promoter and to allow access to other transacting factors that play more direct roles in the production of mRNA by action of RNA polymerase II (Okino and Whitlock, 1995; Dogra et al., 1998). One of the transacting factors, Sp1, can bind to GC box in BTE sequence immediately upstream of the TATA box of the CYP1A1 gene (Dogra et al., 1998). The zinc-finger domain of Sp1 can interact with bHLH/PAS domain of Ah receptor or Arnt (Dogra et al., 1998). Thus, XRE-binding protein bound to XRE probably interacts with Sp1. This may be an important trigger in disrupting chromatin structure to allow anchoring of transcription factors to the CYP1A1 promoter. Structure and size of potent CYP1A1 inducer molecules, such as benzo (a)pyrenes, heterocyclic amines, and dioxins, are almost uniform. These inducers may similarly regulate the transcription of the gene. b. CYP1A2 The CYP1A2 gene contains at least one XRE sequence in the 5⬘ flanking region of the gene (Quattrochi and Tukey, 1989). Chung and Bresnik (1995) found two functionally important cis elements—distal and proximal elements—in the 5⬘ flanking region. The distal element (bp ⫺2352/⫺2094) contains consensus-binding sequences for activator protein1, nuclear factor E1.7, and hepatic nuclear factor-1 (HNF-1). The proximal element (bp ⫺72/⫺31) contains CCAAT and GC boxes. In livers of Ah receptor gene knockout (Ah receptor ⫺/⫺) mice, induction of the CYP1A2 is totally abolished as is CYP1A1 induction (FernandezSalguero et al., 1995; Schmidt et al., 1996). However, sequence of the 5⬘ flanking region of the CYP1A2 gene differs markedly from that of the CYP1A1 gene (Sogawa et al., 1985). The detailed regulatory mechanism of expression of CYP1A2 gene is not yet elucidated. c. CYP1B1 Savas et al. (1994) and Bhattacharyya et al. (1995) found a new member of the CYP1 gene family—CYP1B1. The CYP1B1 gene contains nine core XRE motifs in the 5⬘ flanking region (Tang et al., 1996). At least three of the nine motifs seems to be functional in mediating dioxininduced transcription of the gene (Tang et al., 1996). Analysis of the 5⬘ flanking region of the CYP1B1 gene shows a number of similarities to the region of the CYP1A1 gene (Tang et al., 1996). The regulatory mechanism for expression of the CYP1B1 gene probably resembles that of the CYP1A1 gene. 2. CYP2 Family a. CYP2B1 and 2B2 (2B1/2) P-450 isoforms encoded by the CYP2B gene subfamily are markedly induced by PB. The CYP2B1 and CYP2B2

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are major isoforms. The sequence in coding and upstream regions of the both genes are about 97% homologous (Gonzalez, 1989). Thus, the expression of the two genes seems to be regulated by a similar manner at the transcriptional level. The CYP2B1/2 genes contain two functionally important regions—proximal and distal promoters—in the 5⬘ flanking region (Dogra et al., 1998). a. Cis Elements in Proximal Promoter The proximal promoter resides within about 200 bp of upstream side of the start point of transcription in CYP2B1/2 genes (Dogra et al., 1998). Although detailed information on the promoter is various among investigators, there is a positive cis element (⫺98/⫺69) in this promoter, which contains Barbie box, a 15-bp sequence with AAAG core (Prabhu et al., 1995). The Barbie box is important for PB induction in the bacterial PB-inducible P-450 isoform—CYP102 and CYP106 (Liang et al., 1995; Liang and Fulco, 1995). In addition, Prabhu et al. (1995) found a negative control element between ⫺160 and ⫺126, and Hoffman et al. (1992) showed another negative control element (⫺103/ ⫺66), which overlaps the positive cis element. b. Transacting Factors for Proximal Promoter Several binding proteins for cis elements in the proximal promoter have been reported. Prabhu et al. (1995) found a 28-kDa protein that interacts with the proximal promoter after phosphorylation and suggested the following model. In the absence of inducers, dephosphorylated form of the 28-kDa protein binds to the negative control elements. When the protein is phosphorylated in the presence of an inducer, the phosphorylated form binds to the positive cis element. In this model, phosphorylation acts as molecular switch for druginducible CYP2B induction. Luc et al. (1996) found a binding site for members of CCAATT enhancer-binding protein (C/EBP) family between ⫺66 and ⫺42 bp in the proximal promoter and suggest that the members of C/EBP family act as DNA-binding factor. In addition, a repressor protein that interacts with Barbie box, characterized by Liang and Fulco (1995), is a candidate for DNA-binding factor. c. PB-Inducible CYP2B1/2 Expression Through Distal Promoter In the distal promoter, a cis element (⫺2318/⫺2155) that responds to PB (PBresponsive element; PBRE) has been found in the CYP2B1/2 genes (Trottier et al., 1995; Park et al., 1996). In the PBRE, the 88-bp stretch from ⫺2588 to ⫺2710 is the minimal sequence (PBRE core) that mediates PBinduction (Liu et al., 1998). The minimal sequence contains binding site for nuclear factor (NF)-1, and the binding site is important for PB-inducible expression of the CYP2B1/2 genes (Liu et al., 1998). In addition, Liu et al. (1998) found other unidentified regulatory elements in PBRE core, and suggest that the binding sites for NF-1 and the unidentified elements comprise PB-responsive unit. NF-1 and binding proteins for the unidentified elements may act as a transacting factor. A GRE is also found in the 5⬘

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flanking region about 1.4 Kb (⫺1359/⫺1339) upstream of the transcription start site of the CYP2B2 gene ( Jaiswal et al., 1990). Although the expression of the CYP2B1/2 genes has been presumed to be regulated at the transcriptional level through a intracellular receptor (Dogra et al., 1998), no PB receptor has been identified. If a PB receptor exists, it is questionable whether inducers for CYP2B1/2 such as PB, DDT, chlordan, isosafrol, and dieldrin bind to the receptor because the molecular size and stracture of the inducers are various. b. Other PB-Inducible CYP2B-Related Isoforms Honkakoski et al. (1996) found a PBRE in the distal promoter of the CYP2B10 gene, which encodes a major PB inducible P-450 isoform in mouse livers. Transcriptional regulation of a PB-inducible P-450 isoform in avian, CYP2H1, has been well examined (Dogra and May, 1997; Dogra et al., 1998). c. CYP2C2 The CYP2C2 gene encodes the arachidonic acid-inducible P450 isoform acting as an arachidonic acid epoxygenase in rabbit livers. Venepally et al. (1992) found that transcription of the CYP2C2 gene is activated through a cis element containing the HepG2 nuclear protein (HPF-1)-binding motif. The motif is located around ⫺100 bp in the CYP2C2 promoter, and is highly homologous to HNF-4-binding motif that is present in promoters of more than 20 other CYP2 genes (Chen et al., 1994a). Subsequently, Chen et al. (1994b) revealed that the HPF-1 motif is the HNF-4 binding site, and HPF-1 and HNF-4 are possibly identical. These suggest that HPF-1 (and probably HNF-4) acts as a common transactivator for the CYP2 gene family. Recently, Li et al. (1996) examined transcriptional regulation of the CYP2C2 gene with transgenic mice and indicated that the 5⬘ flanking region contains basal regulatory elements. d. CYP2C6 PB induces a P-450 isoform encoded by the CYP2C6 gene in rat livers in addition to CYP2B1/2. Dexamethasone also induces this isoform. Yano et al. (1992) examined the role of upstream DNA sequence of the gene and found that D-site binding protein (DBP) activates the CYP2C6 promoter. The DBP-binding site is localized between ⫺40 and ⫺65 bp upstream of the transcription start site (Yano et al., 1992). The transcription of the CYP2C6 gene and induction of its mRNA by PB treatment is blocked by antiprogestin–antiglucocorticoid, RU486, suggesting the involvement of steroid receptor in the induction of the enzyme by PB (Shaw et al., 1993). e. CYP2C9 P-450 isoform encoded by the CYP2C9 gene is one of the most important drug-metabolizing enzymes in human livers. The CYP2C9

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metabolizes various drugs such as fluoxetine, losartan, phenytoin, tolbutamide, torsemide, s-warfarin, and numerous nonsteroid antiinflammatory drugs. Xiang et al. (1998) found a 27-bp cis element immediately after the translation start site. A 100-kDa nuclear protein binds to two 6-bp direct repeats (5⬘-CTTGTG-3⬘) in the element. f. CYP2C11 The CYP2C11 is expressed in livers of male rats and acts as a steroid hydroxylase. Expression of the gene is regulated by growth hormone (GH) at the level of transcriptional initiation (Legraverend et al., 1992). Pulsatile GH release stimulates expression of the gene (Sundseth et al., 1992), while continuous GH secretion suppresses the expression and stimulates expression of a female-specific isoform, CYP2C12 (Section II.A.2.g). Recently, Waxman et al. (1996) found a specific binding site for GH-regulated nuclear factor (GHNF) in the 5⬘ flanking region of the CYP2C11 gene, and suggest that GHNF may contribute to down-regulation of the gene by continuous GH stimulation. Strom et al. (1995) found two silencer elements, an HNF-1-binding site-like element and HPF-1-binding site-like elements in the 5⬘ flanking region. Although the HPF-1-binding site-like elements bind to nuclear orphan receptors, the elements have limited functional importance (Strom et al., 1995). Transacting factors for the silencers and the HNF-1-binding site-like element are unidentified. g. CYP2C12 The CYP2C12 is a female-specific steroid hydroxylase. Expression of the CYP2C12 gene in rat livers is transcriptionally activated by continuous exposure to plasma GH. Waxman et al. (1996) found five binding sites for GHNF, two binding sites for C/EBP, two binding sites for DBP, and an HNF-1-binding site in the 5⬘ flanking region of the gene. As GHNF is enriched in nuclear extracts from livers of female rats and those from livers of male rats treated continuously with GH, the transcription of the gene relates to accumulation of the factor in hepatocyte nuclei (Waxman et al., 1996). The factor is distinguished from other GH-dependent transcriptional factors such as STAT (signal transducers and activators of transcription) proteins (Waxman et al., 1996). Thus, the transcription of the CYP2C12 gene is activated via a signaling pathway distinct from STAT pathway that is proposed to other CYP genes (Section II.A.3.e). Recently, Lahuna et al. (1997) found binding sites for HNF-3 (⫺138/ ⫺126) in the 5⬘ flanking region. However, GH treatment induces binding of HNF-6 to the HNF-3-binding sites (Lahuna et al., 1997). Furthermore, HNF-6 competes in the binding of HNF-3 to the sites, although HNF-6 and HNF-3 possess different DNA binding specificities and are immunologically distinct (Lahuna et al., 1997). These findings suggest that the expression of the CYP2C12 gene is regulated through interaction between the ⫺138/ ⫺126 site and the two nuclear orphan receptors.

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h. CYP2C13 The CYP2C13 gene encodes a constitutive male-specific P450 isoform that acts as steroid hydroxylase. The expression of the enzyme is restricted to the liver and suppressed by the female pattern of GH secretion at the level of transcriptional initiation. Cis elements have been found between ⫺2000 and ⫺117 nucleotides in the 5⬘ flanking region of the gene (Legraverend et al., 1994). Transactivation of the CYP2C13 gene through the elements requires both HNF-1 and HNF-3 (Legraverend et al., 1994), suggesting that the expression of the gene is influenced by members of nuclear orphan receptor subfamily. i. CYP2D5 Expression of the CYP2D5 gene commences a few days after birth in rat livers. The maximal mRNA level is achieved when animals reach puberty. Lee et al. (1994b) found a cis element between nucleotides ⫺112 and ⫺83 in the 5⬘ flanking region of the gene. Expression of the gene is markedly increased by cotransfection with a vector expressing C/EBP beta. However, the expression is unaffected by vectors producing C/EBP alpha, HNF-1 alpha, and DBP (Lee et al., 1994b). The ⫺112/⫺83 element displays some sequence similarity to Sp1 consensus sequence and is able to bind to Sp1 (Lee et al., 1994b). C/EBP beta alone is unable to bind to the ⫺112/⫺83 element, but C/EBP beta-Sp1 complex binds to the element (Lee et al., 1994b). Thus, the two transacting factors—C/EBP beta and Sp1—can work in conjunction to activate the CYP2D5 gene via protein– protein interaction. j. CYP2D9 The CYP2D9 gene encodes a male-specific P-450 isoform that catalyzes steroid 16 alpha-hydroxylation in mouse livers. There is a regulatory element in the 5⬘ flanking region (⫺100 TTCCGGGC ⫺93, Yokomori et al., 1995; ⫺110 CTCCTCCCTATTCCGGGCC ⫺92, Sueyoshi et al., 1995) of the gene. Nucleotide at position ⫺99 (T) is crucial for binding the element to a nuclear factor, NF2d9, and transcriptional activation of the ⫺110/⫺92 element (Sueyoshi et al., 1995). NF2d9 belongs to the CYP2 gene family and is the mouse homolog of human LBP-1a (Sueyoshi et al., 1995). Thus, the CYP2D9 gene is regulated through association of NF2d9 with the ⫺110/⫺92 element. Furthermore, Yokomori et al. (1995) found that demethylation occurs in the ⫺100/⫺93 element, and that binding factor for the demetylated element is gamma-aminobutylic acid-binding protein (GABP). GABP does not bind to the methylated ⫺100/⫺93 element, and is a methylation-sensitive transacting factor for the CYP2D9 gene (Yokomori et al., 1995). k. CYP2E1 The CYP2E1 is responsible for microsomal P-450-mediated ethanol oxidation. The enzyme also metabolizes halothane, acetaminophen, and chlorzoxazone. Furthermore, the enzyme is involved in metabolic acti-

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vation of compounds such as carbon tetrachloride. In addition, the enzyme is the primary component of the methylglyoxal and propandiol pathways of gluconeogenesis. The CYP2E1 gene is transcriptionally activated in rat livers within one day after birth (Ueno and Gonzalez, 1990). Three cis elements have been detected in the 5⬘ flanking region of the gene (Ueno and Gonzalez, 1990). One element is the binding site for the nuclear factor-Y (NF-Y), and another element is a palindrome sequence unique to the gene (Ueno and Gonzalez, 1990). However, the two elements do not contribute to transcriptional activation of the gene in vitro (Ueno and Gonzalez, 1990). The remaining element is binding site for HNF-1 and is considered to be ethanol response element (Liu and Gonzalez, 1995). Recently, however, McGehee et al. (1997) showed that the increase in the CYP2E1 gene transcription by ethanol is not mediated through the HNF-1-binding site, and ethanol response element is absent in the 5⬘ flanking region. On the other hand, the CYP2E1 has been shown to be posttranscriptionally regulated through mRNA and protein stabilization (Section II.B.3.a).

3. CYP3 Family P-450 isoforms encoded by the CYP3A genes are induced by glucocorticoids. The CYP3A5 (human) and 3A1 and 3A23 (rat) are major dexamethasone-inducible isoforms. The induction mechanism of classical glucocorticoid receptor-dependent proteins such as tyrosine aminotransferase follows: glucocorticoids bind to glucocorticoid receptors, and the ligand-receptor complex binds to GRE in of glucocorticoid receptor-dependent proteins. However, the CYP3A genes are induced not only by glucocorticoids but also by antiglucocorticoids such as pregnenolone 16 alpha-carbonitrile (PCN) and nonsteroids such as rifampicin, methylapone, and metapyrone (Pereira and Lechner, 1995). Furthermore, genes of major CYP3A isoforms do not contain intact GRE (Hashimoto et al., 1993; Quattrochi et al., 1995). Therefore, the induction mechanism of the CYP3A isoforms differs from that of the classical glucocorticoid receptor-dependent proteins. a. CYP3A1 The CYP3A1 is a major dexamethasone-inducible isoform in rat livers. The enzyme is induced markedly by dexamethasone in immature animals, whereas the induction is drastically impaired in adult animals. Pereira and Lechner (1995) found that expression of the CYP3A1 gene is activated after treatment with PCN in immature rats. Furthermore, the expression is synergistically induced by PCN plus dexamethasone (Pereira and Lechner, 1995). However, the induction by dexamethasone occurs through activation of gene transcription, whereas the induction by PCN

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takes place by additional synthesis of the CYP3A1 (Pereira and Lechner, 1995). The 5⬘ flanking region of the gene contains a dexamethasone-responsive element (Burger et al., 1992). However, the element does not contain GRE (Quattrochi et al., 1995), and does not bind to glucocorticoid receptor (Burger et al., 1992; Quattrochi et al., 1995). Transacting factors for the element have been unidentified. b. CYP3A4 The CYP3A4 is an adult-specific form of P-450 in human livers. There are two characteristic sequences termed as P450NF-specific element (NFSE) and P450HFLa specific element (HFLaSE) in the 5⬘ flanking region of the gene (Hashimoto et al., 1993). In addition, a basic transcription element locates in the region (Hashimoto et al., 1993). Transacting factors for these elements are unknown. c. CYP3A5 Two GRE half-sites are identified in the 5⬘ flanking region of the CYP3A5 gene (Schuetz et al., 1996). The two half sites are located 60 bp apart and are required for the dexamethasone response (Schuetz et al., 1996). One of the sites binds to the glucocorticoid receptor (Schuetz et al., 1996). These suggest that the glucocorticoid receptor is necessary for dexamethasone-induced CYP3A5 expression. The two sites presumably interact to facilitate the glucocorticoid receptor binding (Schuetz et al., 1996). d. CYP3A7 The CYP3A7 is an isoform of P-450 that was isolated from human fetal livers and termed as P-450HFLa. This isoform has been clarified to be expressed during fetal life specifically. The identify of the 5⬘ flanking sequences between CYP3A4 and CYP3A7 genes is 91% (Hashimoto et al., 1993). Within the 5⬘ flanking region of CYP3A7 gene, putative binding sites for several transcriptional regulatory factors exist, and basic transcription element-binding factor (BTEB) is shown to interact with one of the binding sites (Itoh et al., 1992). e. CYP3A10 The CYP3A10 is a male-specific P-450 isoform that catalyzes 6 beta-hydroxylation of lithocholic acid. There are two cis elements in the 5⬘ flanking region of the gene (Subramanian et al., 1998). One element binds to STAT 5a or 5b, and the other element, located adjacent to the STAT-binding element, binds to NF-Y. NF-Y bound to the element modulates the binding of STAT5a or 5b to the STAT-biding element (Subramanian et al., 1998). f. CYP3A23 A dexamethasone responsive region comprising two functional elements is identified in proximal promoter of the CYP3A23 gene

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(Huss et al., 1996). The elements do not contain GRE, but one element includes an imperfect direct repeat of the steroid receptor motif AGGTCA, separated by 4 bp. The other element contains a direct repeat of ATGAACT, separated by 2 bp. However, glucocorticoid receptor does not bind to the two elements (Huss et al., 1996).

4. CYP4 Family Major P-450 isoforms belonging to the CYP4A subfamily are known to be induced by peroxisome proliferators such as clofibrate, Wy14643, and di(2)-ethylhexyl phthalate (DEHP). The CYP4A1 in rats and the CYP4A6 in rabbits are major CYP4A isoforms. The peroxisome proliferators show no obvious structural similarity. Administration of the proliferators to animals leads to an increase in oxidation of fatty acid through the microsomal omega oxidation followed by the peroxisomal beta oxidation. CYP4A1 and 4A6 catalyze the omega oxidation. a. CYP4A1 and 4A6 Issemann and Green (1990) discovered peroxisome proliferator activator receptor (PPAR) alpha. The PPAR alpha is a member of nuclear orphan receptor family in rabbits (Mangelsdorf and Evans; 1995). The PPAR beta and PPAR gamma have been identified in mice (Green and Wahli, 1994). Leukotrien B4 and Wy14643 are found to be ligands for PPAR alpha (Devchand et al., 1996). In livers of PPAR knockout mice, peroxisomes do not proliferate, and transcription of CYP4A genes is not activated by treatment with peroxisome proliferators (Lee et al., 1995). In CYP4A6 gene, Muerhoff et al. (1992) found a cis element (⫺677/ ⫺644) containing imperfect direct repeat of consensus sequence for nuclear receptor family-binding site (AGGTGA) in the 5⬘ flanking region. Deletion of the ⫺677/⫺644 element abolishes the response of the gene to peroxisome inducers, and therefore the element is defined as peroxisome proliferatorresponsive element (PPRE) (Palmer et al., 1994). Recent studies, however, suggest that PPRE containing an additional 5⬘ sequence is necessary for full activation of the CYP4A6 gene transcription (Palmer et al., 1995; Johnson et al., 1996). PPAR requires retinoid X receptor-alpha (RXR alpha) as accessory partner protein to bind to PPRE (Aldridge et al., 1995; Devchand et al., 1996). The PPRE strongly binds to PPAR alpha/RXR alpha complex, but not to PPAR alpha alone (Devchand et al., 1996). In the CYP4A1 gene, a PPRE has been identified approximately 4300 nucleotide upstream of the transcription start site (Palmer et al., 1995). PPAR alpha and RXR alpha form a heterodimer, and the dimer binds specifically to the PPRE (Aldridge et al., 1995). In addition, PPRE motifs are present in genes responsive to peroxisome proliferators notably genes

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of acyl CoA oxidase and other enzymes for peroxisomal beta oxidation ( Johnson et al., 1996; Varanasi et al., 1996). See Johnson et al. (1996) and Dogra et al. (1998) for further explanation of the induction process of the CYP4A1/6 gene expression through PPAR. b. CYP4A4 The CYP4A4 gene encodes a P-450 isoform that increases during pregnancy and catalyzes the prostaglandin omega-hydroxylation. Although the intron/exon structure of the CYP4A4 gene is highly conserved in relation to genes of CYP4A1 and 4A6, the 5⬘ flanking sequences of the gene differs from those of the CYP4A1 and 4A6 genes (Palmer et al., 1993). Like other CYP4A genes, the CYP4A4 promoter does not contain a TATA box (Palmer et al., 1993). No consensus sequences for glucocorticoid receptor or progesterone receptors are found within 1086 bp of sequence upstream of the transcription start site (Palmer et al., 1993). The regulatory mechanism of the expression of CYP4A4 gene has been unclear. 5. CYP7A The CYP7A gene encodes a P-450 isoform that acts as cholesterol 7 alphahydroxylase and catalyses the first and rate-limiting step in the conversion of cholesterol to bile acids. Hoekman et al. (1993) found the major transcription-activating region in the proximal 145 nucleotide and thyroxine-responsive site further upstream. Lee et al. (1994a) showed five distinct DBP-binding sites distributing between nucleotides ⫺41 and ⫺295 from the transcription start site. The stimulating effect of DBP can in part be ascribed to its functional interaction with DBP-binding sites B (⫺125/ ⫺115), C(⫺195/⫺172), and D(⫺230/⫺214) (Lee et al., 1994a). C/EBP beta also binds to these sites but causes a more modest increase in CYP7A gene transcription (Lee et al., 1994a). 6. CYP51 The CYP51 is a P-450 isoform that acts as sterol 14 alpha-demethylase in plants and animals. Human CYP51 shows ubiquitous expression, and initiation of transcription at more than one site in this gene indicates that the CYP51 gene is a housekeeping gene (Stromstedt et al., 1996). The 5⬘ portion of intron 1 is GC-rich and contains potential binding sites for several transcription factors (Rozman et al., 1996). In the 5⬘ flanking region, GCrich sequence is present, but TATA and CAAT sequences are absent (Rozman et al., 1996). The 5⬘ flanking region, the 5⬘ portion of intron 1, and exon 1 show the characteristics of CpG island and sterol responsive element-like motifs are present in this island (Rozman et al., 1996).

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B. Posttranscriptional, Translational, and Posttranslational Regulations Including Degradation There have been divergences between amounts of P-450 mRNA and their transcripts and amounts of the transcripts and the mono-oxygenase activities mediated by P-450 isoforms belonging to the CYP1A, 2B, 2E, and 3A subfamilies. Therefore, the expression of these isoforms is also regulated in the posttranscriptional, translational or posttranslational process. 1. CYP1A Subfamily Regulation of the expression of the CYP1A genes occurs mainly at the transcriptional level (Section II.A.1.a). However, Adesnik and Atchison (1986) have proposed involvement of posttranscriptional mechanisms in the regulation of CYP1A expression. Kim et al. (1991) found that pyridine modulates the expression of the CYP1A gene subfamily in rat livers by elevating CYP1A1/2 mRNA levels through mRNA stabilization. Recently, Calleja et al. (1997) showed that interferon (IFN)-gamma inhibits induction of CYP1A1/2 proteins and their enzymatic activities but exhibits only a weak influence on CYP1A1/2 mRNA levels in cultured rabbit hepatocytes. Thus, IFN stimulates posttranscriptional suppressive pathway of the CYP1A1/2. The degradation mechanism of the CYP1A1/2 is unclear at present. 2. CYP2B Subfamily a. Posttranscriptional Regulation In general, a close relationship has been observed between the CYP2B mRNA expression and the amount of its transcripts in livers of animals. However, Nanji et al. (1994) found that CYP2B protein levels increase markedly but that the CYP2B1 mRNA level is unaffected in corn oil- and ethanol-fed rats. Agrawal and Shapiro (1996) found that mRNA level of CYP2B1 is unaffected but that the protein level increases by PB administration in rats. These suggest that posttranscriptional mechanisms are responsible for the increase in CYP2B protein levels. Recently, Mino et al. (1998) found that increase in the CYP2B1/2 content, seen in PB-treated rats, is suppressed in spite of retention of a large amount of the CYP2B2 mRNA in perivenular hepatocytes of animals injected first with PB and then with MC (PB⫹MC-treated animals). The divergence between the amount of mRNA and of the transcript can be explained by suppression of translation in the CYP2B2 mRNA. It is also possible that degradation of mRNA by ribonuclease after translation is repressed, and untranslated mRNA is retained in the cytoplasm. As the cause of suppression of translation, competition between the CYP2B1/2 mRNA and the

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CYP1A1/2 mRNA for ribosomes (Graves et al., 1987), phospholyration of transcriptional regulatory factors (Rhoads, 1993) or binding of repressor proteins to ribosomes (Nomura et al., 1984) may be considered. Mechanisms underlying the turnover and degradation of CYP2B isoforms have been an area of interest (Tashiro et al., 1993). Turnover of the heme moiety of the CYP2B is faster than that of the protein moiety (Shiraki and Guengerich, 1984). Phosphorylation of serine-128 in the CYP2B isoforms causes conformational changes of the reduced form of P-450 molecules to their inactive form—P-420 (Koch and Waxman, 1989; Pyerin and Taniguchi, 1989). P–420 changes into apo P-420 that is rapidly degraded (Koch and Waxman, 1989; Pyerin and Taniguchi, 1989). b. Degradation Three intracellular pathways to degrade P-450 molecules have been suggested (Tashiro et al., 1993). First, P-450 is transported to lysosomes via Golgi apparatus. Although PB-inducible P-450 isoforms are not detectable in Golgi apparatus after PB treatment (Yamamoto et al., 1985; Marti et al., 1990), Neve et al. (1996) found significant amounts of P450 such as the CYP4A1, CYP2E1 and CYP1A2 in Golgi fractions. Second, ER membrane containing P-450 is degraded in the autophagosome-autolysosome system. In livers of PB-treated rats, Masaki et al. (1987) showed that ER membrane loaded with CYP2B isoforms is degraded in autophagic vacuoles. Yamamoto et al. (1990) revealed that leupeptin treatment blocks proteolytic process in autolysosomes and degradation of the isoforms. Difference in the half lives of microsomal proteins is very small in PB- and/or leupeptin-treated rats. Thus, this system is a pathway to degrade CYP2B molecules in hepatocytes of PB-treated animals. However, wide differences in the half lives of microsomal proteins do not support the possibility that microsomal proteins are exclusively degraded by the autophagic process (Tashiro et al., 1993). Third, microsomal P-450 is degraded in ER by proteolytic enzymes. This has been found in the CYP2E (Eliasson et al., 1992) and the CYP3A (Eliasson et al., 1994) but not observed in the CYP2B. 3. CYP2E Subfamily a. Posttranscriptional Regulation The CYP2E1 has been known to be regulated by many of its substrates through substrate-induced stabilization of the enzyme molecules. Kim and Novak (1990) found that the CYP2E1 increases without elevation of its mRNA level after administration of pyridine in rat livers. Cycloheximide administration completely prevents the induction of the enzyme by pyridine, whereas actinomycin D administration has no effect (Kim and Novak, 1990). Wu et al. (1997) showed that pyridine prevents the decline

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of the CYP2E1 protein level and activity during culture of rat hepatocytes but does not prevent the fall in mRNA level. These indicate that pyridine modulates the CYP2E1 level by posttranscriptional mechanisms. The mechanisms may involve increased translational efficiency of mRNA or stabilization of the enzyme protein against degradation. Similar findings have been reported in livers of ethanol-fed rats (Nanji et al., 1994), hepatoma FGC-4 cells treated with ethanol (McGehee et al., 1994) and in livers of isoniazid-treated rats (Park et al., 1993). However, Damme et al. (1996) found a significant increase in the CYP2E1 mRNA and its transcript levels in rats treated with acetylsalicylic acid or sodium salicylate. b. Degradation The CYP2E1 is phosphorylated and degraded in ER (Eliasson et al., 1990). First, serine (Ser)-129 in the enzyme is phosphorylated by cAMP-dependent kinase. Then, the phosphorylated isoform is degraded rapidly in ER by Mg2⫹-ATP-activated proteolytic system. Glucagon activates the phosphorylation on Ser-129 and the subsequent degradation of the isoform (Eliasson et al., 1992). The effect of glucagon on the degradation is abolished by cycloheximide (Eliasson et al., 1992). However, the degradation is not influenced by inhibitors of the autophagosome– autolysosome system (Eliasson et al., 1992). Thus, the CYP2E1 is degraded by specific proteolytic system in ER.

4. CYP3A Subfamily a. Posttranscriptional Regulation Expression of the CYP3A1 gene is regulated not only at the transcriptional level by glucocorticoids but also, in some cases, at the posttranscriptional level by substrate-dependent stabilization. Ferrari et al. (1993) found that interleukin (IL)-1 induces both the CYP3A1 protein and the activity in female rats, but in males, IL-1 represses CYP3A2 activity without decreasing the protein level. The discrepancy between the protein level and activity of CYP3A2 in males indicates a posttranslational regulation. Yuan et al. (1994) found that ethylbenzene elevates both CYP3A-dependent 2 beta-hydroxylase and CYP3A protein levels. Despite the increase in the CYP3A protein levels, the mRNA levels of the CYP3A1 and CYP3A2 are not altered by the drug treatment (Yuan et al., 1994). Thus, the CYP3A1/2 can be induced by posttranscriptional mechanisms. Zangar et al. (1997) found that dimethyl sulfoxide (DMSO) elevates the protein levels of the CYP3A and CYP2E1 without increasing the respective mRNA level in cultured rat hepatocytes. This shows that DMSO enhances CYP3A and CYP2E1 gene expression by posttranscriptional mechanisms. Furthermore, Zangar and Novak (1998) showed that

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DMSO elevates CYP3A protein levels by blocking their degradation in cultured rat hepatocytes. b. Degradation The CYP3A1 is phosphorylated and degraded in ER by proteolytic enzymes by similar mechanism found in the CYP2E1. Phosphorylation of Ser-393 in the enzyme is catalyzed by cAMP-dependent kinase, and the phosphorylated isoform is degraded in ER (Eliasson et al., 1994). Glucagon activates the phosphorylation and the degradation of the isoform in ER, whereas substrates of the CYP3A1 prevent the phosphorylation and the degradation (Eliasson et al., 1994).

III. Membrane Topology of P-450 Molecules P-450 molecules are principal constituents of ER membrane of hepatocytes. Studies on the topology of P-450 in ER membrane, how P-450 molecules exist in the membrane, are important for a better understanding of the structure of biomembrane or of the microsomal monooxygenase system.

A. Biosynthesis and Insertion of P-450 Molecules into ER Membrane In hepatocytes, the mRNA of PB-inducible P-450 isoforms translocates from the nucleus to the cytoplasm through nuclear pores, and interacts with ribosomes bound to ER membrane. PB-inducible P-450 isoforms are synthesized exclusively in the membrane-bound ribosomes (Tashiro et al., 1993). Nascent P-450 peptides are inserted directly from the ribosomes onto the cytoplasmic surface of the rough ER membrane (Tashiro et al., 1993). The insertion of P-450 molecules onto ER membrane requires an insertion signal. Sakaguchi et al. (1987) and Monier et al. (1988) found that short N-terminal segment of the CYP2B molecules functions as the insertion signal. Furthermore, the N-terminal segment also acts as signal for ER retention (Sakaguchi et al., 1987). In contrast to typical secretory signal sequence, the N-terminal segment lacks positively charged amino acid residues but has negatively charged residues (Tashiro et al., 1993). Furthermore, hydrophobic segment adjacent to the N-terminal segment is longer than the secretory signal sequence (Sato et al., 1990). When basic amino acids are substituted into N-terminal amino acids, or hydrophobic core sequences are shortened, the insertion signal is converted into the secretory signal (Szczesna-Skorupa et al., 1988; Sato et al., 1990). The hydrophobic segment

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appears to pull the negatively charged N-terminal segment into ER membrane (Sakaguchi et al., 1992). The N-terminal segment functions as ER retention signal also in the CYP2C1 (Ahn et al., 1993; Szczesna-Skorupa et al., 1995), the 2C2 (Szczesna-Skorupa et al., 1995) and the 3A1 (Van den Broek et al., 1996) molecules. P-450 molecules thus synthesized and inserted are rapidly translocated from rough ER membrane to smooth ER membrane and distributed evenly in ER membrane by lateral diffusion within an hour (Tashiro et al., 1993).

B. Localization and Density of P-450 and P-450 Reductase in ER Membrane 1. Localization in ER Membrane a. Models from Molecular Biological and Biophysical Analyses Threedimensional structure of P-450 molecules in ER membrane is tried to presume by hydropathy analysis. Thirty and more isoforms of P-450 have been analyzed. Pattern of intramolecular sequence of hydrophobic and hydrophilic domains is shown to be similar in all isoforms. Therefore, all isoforms are presumed to have similar three-dimensional structure (Nelson and Strobel, 1988). There are two models of the topology of P-450 molecules in ER membrane. First, a model, suggesting that a P-450 molecule penetrates ER membrane 4앑8 times, was considered (Tarr et al., 1983; Hudecek and Anzenbacher, 1988). However, Nelson and Strobel (1988) proposed the other model wherein the N-terminal hydrophobic domain is anchored to the ER membrane and most of the molecule is exposed on the cytoplasmic surface of the membrane. This model is more suitable than the model of Tarr et al. (1983) and Hudecek and Anzenbacher (1988) for explaining interaction between P-450 and their reductase, and between P-450 and substrates. In addition, data supporting the anchor model have been shown in various P-450 isoforms (Szczesna-Skorupa and Kemper, 1993; Black et al., 1994; He et al., 1996; Menzel et al., 1996). However, Miller et al. (1996) analyzed the membrane topology of the CYP2B4 by X-ray diffraction analysis and suggested that the isoform is deeply embedded in ER membrane. Furthermore, Shank-Retzlaff et al. (1998) examined the CYP2B4 with the Langmuir-Blodgett monolayer system and showed that the isoform contains large membrane insertion areas that allow two to four transmembrane helices to be formed, although the isoform may not be deeply embedded. Thus, the topology of the CYP2B4 may differ from that of other microsomal P-450 isoforms.

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Nelson and Strobel (1988) suggested that the N-terminal hydrophobic domain of microsomal P-450 molecules forms a hairpin-loop and that the domain penetrates the ER membrane twice. However, Vergeres et al. (1989) assumed only one penetration through the membrane. Recent studies revealed that the domain forms a single transmembrane anchor projecting into lumen of ER (Szczesna-Skorupa and Kemper, 1993; Black et al., 1994; Menzel et al., 1996). Therefore, the domain spans ER membrane only once. In conclusion, except for CYP2B4, three findings currently describe the topology of P-450 molecules in ER membrane: (1) the N-terminal hydrophobic domain is anchored to the ER membrane, (2) the domain spans ER membrane only once, and (3) the active site is present in the cytoplasmic surface of the ER membrane. NADPH cytochrome P-450 reductase (reductase) has an N-terminal hydrophobic domain that anchors into the ER membrane (Porter and Kasper, 1986; Kida et al., 1998). The remaining part of the enzyme including the C-termunus is exposed to the cytoplasmic surface of the membrane (Kida et al., 1998). The reductase molecule is assumed to contain five key elements—ER membrane-binding domain; FMN-, FAD-, and NADPHbinding sites exposed to the cytoplasmic surface of ER membrane; and P450-binding site, as revealed by cDNA sequencing and X-ray diffraction analysis (Porter and Kasper, 1986). The P-450-binding site had been thought to be present in the ER membrane-binding domain (Masters et al., 1973). However, the interaction between CYP2E1, which lacks the ER membranebinding domain, and reductase is the same as that between complete CYP2E1 and reductase (Larson et al., 1991). Therefore, the P-450 binding site is unlikely to exist in the ER membrane-binding domain. b. Models from Immunoelectron Microscopy First, liver microsomes incubated with ferritin particles conjugated with anti-PB-inducible P-450 antibody (Matsuura et al., 1979) and with antireductase antibody (Morimoto et al., 1976) were examined. Matsuura et al. (1979) found ferritin particles on the outer surface (cytoplasmic surface) of microsomal membrane, and showed quantitatively that the particles form aggregates. Morimoto et al. (1976) observed clusters formed by 3–5 ferritin particles on the outer surface of microsomal membrane. From these results, Matsuura et al. (1979) supposed that 3–5 reductase molecules and 30–100 P-450 molecules form an aggregate, taking the cluster model of Peterson et al. (1976) into consideration. Later, P-450 were examined by the postembedding immunogold method. Marti et al. (1990) observed localization of PB-inducible P-450 in rat hepatocytes and calculated labeling density (number of gold particles per square micrometer of ER membrane) by combining the number of gold particles with the area of ER obtained by morphometry. The authors showed that

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the labeling density is not different between rough and smooth ER and that, after PB administration, it increases both in rough and smooth ER. However, they did not find aggregation of gold particles as reported on ferritin particles by Matsuurra et al. (1979). Nevertheless, the density of P450 molecules in the ER membrane is unknown by their method. Fukui et al. (1992) examined localization of the CYP2B in hepatocytes by postembedding immunogold method, and reported that particle density (number of gold particles per 1 애m length of rough ER membrane on an electron micrograph) is nearly zero in normal rats and increases markedly after PB administration. It is worth noticing that the authors also did not observe the aggregation of gold particles. Thus, the aggregate formation, observed on ferritin particles, is not generally found when the immunogold method is used in both sections and microsomes. Therefore, the aggregate formation can be considered as an artifact caused by use of ferritin. However, the evidence showing aggregate formation of P-450 molecules have been biochemically indicated (Section III.B.2.c). Even if P-450 molecules form aggregates in the ER membrane, the aggregates may not be detected electron-microscopically because of steric hindrance. In these immunoelectron-microscopic results, however, sublobular positions of hepatocytes examined are unclear. Hepatocytes show functional and morphological heterogeneity depending on positions within the liver lobule (Section IV). In addition, the method of sampling for quantitative analysis of immunoelectron micrographs is not described in these reports. 2. Densities in ER Membrane a. Microphotometry of Total P-450 The reduced form of P-450 binds to carbon monoxide and represents a characteristic absorption band at 450 nm (Omura and Sato, 1964). Gooding et al. (1978) developed a microspectrophotometric method for assay of P-450 in tissue sections, based on the biochemical method of Omura and Sato (1964). However, the sensitivity of method of Gooding et al. (1978) is low, and therefore the exact amount of P-450 in sections can hardly be determined by their method for two probable reasons. First, their method has a fundamental fault; they used an inadequate extinction (⌬OD450), although the difference of absorbance at 450 nm minus that at 490 nm (⌬OD450-490, true extinction of P-450) is necessary for calculation of the amount of P-450 (Omura and Sato, 1964). Second, Johannesen and DePierre (1978) claimed that hemoglobin and methemoglobin interfere with the extinction of P-450 as determined by the biochemical method of Omura and Sato (1964). Tissue sections usually contain large amounts of these hemoproteins. Watanabe et al. (1989) and Watanabe and Kanamura (1991) developed a microphotometric method in

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which the true extinction of P-450 was used and the effect of contaminating hemoproteins was minimized. b. Microphotometry and Image Analysis of P-450 Isoforms and Reductase The immunostaining intensity of an enzyme in sections can be semiquantitatively estimated by microphotometry or microfluorometry. However, the results obtained from microphotometry or microfluorometry are simply relative values such as absorbance or fluorescence intensity. If information on the relation between the amount of the enzyme and of final product of the immunohistochemical reaction is obtained, the relative values can be converted into molar contents in the cytoplasm. The nitrocellulose (NC) model system for obtaining the information has been introduced by Nibbering et al. (1986) and Nibbering and van Furth (1987). Watanabe et al. (1991) established a method for measuring the content of reductase in the cytoplasm by combination of immunohistochemistry with microphotometry and NC binding assay. In addition, Tanaka et al. (1997) and Mino et al. (1998) developed methods for measurement of the CYP1A1/2 and 2B1/2 respectively, on the same principle. c. Estimation of Densities Total 450, CYP1A1/2, CYP2B1/2, and reductase amounts per unit cytoplasmic volume can be measured by the methods of Watanabe et al. (1989, 1991), Watanabe and Kanamura (1991), Tanaka et al. (1997) and Mino et al. (1998). On the other hand, ER area per unit cytoplasmic volume is estimated by the point counting method (Weibel, 1979). When the amounts of the enzymes per unit cytoplasmic volume are divided by ER area per unit cytoplasmic volume, we can calculate amounts of the enzymes per unit area of ER membrane. Densities of the enzyme molecules in ER membrane (numbers of enzyme molecules per unit area of ER membrane) are obtained by multiplying the amounts of the enzymes per unit area of ER membrane by Avogadro’s number. This method is useful because it allows us to measure values in hepatocytes of periportal, midzonal, or perivenular zones. In mice, the density of total P-450 molecules in ER membrane, measured by the method of Watanabe et al. (1992), is 3700/애m2 ER membrane in hepatocytes adjoining the portal area, 4000 in hepatocytes in second and third layers from the portal area, 2900 in hepatocytes adjoining the central venule, and 5600 in hepatocytes in the second and third layers from the central venule. Thus, the value in hepatocytes in the second and third layers from the central venule is greatest. However, the values in rats are somewhat different from those in mice; 5600–6800/애m2 ER membrane in periportal hepatocytes, 6200–6600 in midzonal hepatocytes and 3900–4700 in perivenular hepatocytes. Thus, the value (3900–4700/애m2 ER) is unexpectedly smallest in perivenular hepatocytes in which drug-metabolizing capacity is

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thought to be highest (ER area per unit cytoplasmic volume is largest in rat perivenular hepatocytes) (Watanabe et al., 1993a). However, it is the same with mice that the value is smallest in hepatocytes adjoining the central venule. The peculiarity of hepatocytes adjoining the central venule is discussed in Section IV. The density of the CYP1A1/2 molecules in ER membrane is 100/애m2 ER membrane in periportal hepatocytes, 130 in midzonal hepatocytes, and 250 in perivenular heptocytes in rats (Tanaka et al., 1997). After MC treatment, the density becomes 2400 in peroportal, 3100 in midzonal, and 5000 in perivenular hepatocytes. Thus, the value is greatest in perivenular hepatocytes. The density of reductase molecules in ER membrane is measured in rats; 140–160/애m2 ER membrane in periportal hepatocytes, 170–180 in midzonal hepatocytes, and 110–130 in perivenular hepatocytes (Watanabe et al., 1993a). The sublobular gradient of the values is similar to that of total P450, and the value in hepatocytes adjoining the central venule is smaller than that in hepatocytes in the third layer from the central venule.

C. Interaction between P-450 and Reductase in ER Membrane On the dynamics of P-450 molecules in ER membrane, Peterson et al. (1976) proposed the cluster model suggesting that about ten P-450 molecules form an aggregate and surround one reductase molecule. The P-450 molecules are considered to move by the lateral diffusion in ER membrane to meet reductase molecules (French et al., 1980). The lateral diffusion of P-450 or reductase molecules in ER membrane is shown to play an important role for efficient drug metabolism by the two enzymes (Backes and Eyer, 1989). However, when P-450 or reductase molecules form aggregates, the speed of lateral diffusion is very low (Yang, 1975). Therefore, the density of P450 or reductase molecules in ER membrane should be very high for metabolizing drugs effectively, if the molecules form aggregates. However, the densities of P-450 and reductase molecules in ER membrane is not yet known. Therefore, the cluster model of Franklin and Estabrook (1971) and Peterson et al. (1976) was replaced by the noncluster model (Yang, 1975) suggesting that P-450 molecules are expected to exist solitarily for effective assembly of P-450 and reductase. Recently, the densities of P-450 and reductase molecules in ER membrane have been estimated (Section III.B.2.c). The densities of reductase molecules in ER membrane in hepatocytes of the three sublobular zones are 110–180 molecules/애m2 ER (Watanabe et al., 1993a). On the basis of the values, Watanabe et al. (1993a) calculated the time of reduction of P-

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450 molecules by reductase in the cluster model of Franklin and Estabrook (1971) and Peterson et al. (1976). The diffusion constant of the P-450 aggregate is calculated to be 0.05–0.07 애m2 ⭈ sec⫺1. The value and densities of reductase molecules show that all P-450 molecules in ER membrane can be reduced by reductase within 0.3 sec. Furthermore, if the density of reductase is 20/애m2 ER, the reduction of P-450 molecules finishes within 1 sec. Therefore, the densities of P-450 or reductase molecules in ER membrane are high enough to support the cluster model of Peterson et al. (1976). In fact, P-450 or reductase are again proposed to form aggregates. French et al. (1980), Stier et al. (1989), Rietjens et al. (1989), and Schwarz et al. (1990) have showed that P-450 molecules do not exist singly but as hexamers. Furthermore, Schwarz et al. (1990) observed P-450 molecules incorporated into liposomes by freeze-fracture electron microscopy and analyzed dynamics of the molecules by saturation transfer electron paramagnetic resonance. They considered that P-450 form aggregate consisting of six or more molecules in ER membrane and proposed a model suggesting that the aggregate is rotating around an axis through the center of the aggregate. Furthermore, French et al. (1980) considered that reductase also exists as hexamers in microsomal membrane from rabbit livers. Rietjens et al. (1989) incorporated reductase isolated from rat livers into liposomes resembling ER membrane in nature. They found that reductase molecules form 31-mers, and phospholipid is necessary for formation and maintenance of the polymers. Because the molecular weight of reductase is 78,000, reductase molecules must exist in ER membrane as large aggregates of molecular weight, about 500,000 (hexamer per aggregate) in rabbits and 2,400,000 (31-mer per aggregate) in rats. Speed of lateral diffusion of the aggregates is naturally very slow. In addition, possibility that the aggregates of P-450 and those of reductase form large micelles in ER membrane is shown (Rietjens et al., 1989). Assuming that P-450 molecules form hexamers and reductase molecules form 31-mers, the reduction of P-450 is calculated to be completed within 1 sec because of high densities of P-450 and reductase molecule in ER membrane (Section III.B.2.c). Although the ratio of P-450 to reductase required for the optimum monooxygenase activity is 1 : 1 (French et al., 1980; Backes and Eyer, 1989), the two enzyme molecules are not present in an equimolar ratio in microsomal or ER membrane. The ratio is shown to be 20 : 1 in liver microsomes from rats treated with PB (Franklin and Estabrook, 1971; Peterson et al., 1976), 15 : 1 in the microsomes from control and PB-treated rats, and 21 : 1 in the microsomes from 웁-naphthofravon-treated rats (Shephard et al., 1983). Watanabe et al. (1993a) measured the ratio to be 32앑45 : 1 by use of the previous method.

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D. Relationship between ER Proliferation and P-450 Induction PB administration to animals induces a marked increase in P-450 level in the liver and a prominent proliferation of ER in hepatocytes (Bresnick, 1980; Kanai et al., 1984, 1990; Watanabe et al., 1992). The proliferation of ER after PB administration was correlated with the increase in P-450 level. Watanabe et al. (1992) analyzed the increase in total P-450 level and ER proliferation in hepatocytes of PB-treated mice using a combination of microphotometry and morphometry. They found that the relationship between the increase in total P-450 level and ER proliferation exhibits differences depending on the location of hepatocytes within the lobule. In perivenular hepatocytes, ER proliferates generally without increase in the density of total P-450 molecules in ER membrane. In hepatocytes in the second and third layers from the portal area, the density increases markedly without ER proliferation or with slight ER proliferation. Therefore, ER proliferation appears not to relate closely to the increase in the density of total P-450 molecules in the hepatocytes. Hepatocytes adjoining the portal area are unique; the cells do not respond to PB, and increases in the density and ER proliferation are not observed in the cells. Administration of cobalt chloride is shown to inhibit increases in hepatic microsomal P-450 induced by PB treatment (Tephly and Hibbeln, 1971). Amatsu et al. (1995) examined whether ER proliferates in hepatocytes of mice injected with PB and cobalt chloride. They found that the ER area increases but that total P-450 level decreases or remains unchanged in periportal, midzonal, and perivenular hepatocytes of animals treated with the two compounds. They concluded that cobalt inhibits the increase in total P-450 level but has no effect on the ER proliferation in hepatocytes of mice treated with PB, indicating a dissociation of ER proliferation and P-450 increase after PB administration. Inducers for the enzymes of the microsomal monooxygenase system are classified into two broad classes: (a) compounds exemplified by PB that cause ultrastructural changes including prominent proliferation of smooth ER in hepatocytes and (b) MC and other compounds that have a less intensive effect on hepatocyte ultrastructure (Shull et al., 1983). It was, however, unclear whether smooth ER proliferates in hepatocytes after MC administration. Tanaka et al. (1997) found that in spite of a significant increase in the intensity of immunohistochemical staining of the CYP1A1/2 and total P-450 level, proliferation of smooth ER (and rough ER) does not occur in periportal, midzonal, and perivenular hepatocytes from rats injected with MC. The results indicate the increase in P-450 level without ER proliferation in rat hepatocytes after MC administration and additional

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evidence showing a dissociation of ER proliferation and P-450 increase after drug administration.

IV. Sublobular Expression of P-450 Hepatocytes of adult mammals show functional and structural heterogeneity depending on their locations within the lobule. The physiological significance of the hepatocyte heterogeneity appears still obscure, but Gumucio (1989) discusses possible hypotheses. The heterogenous expression of total P-450 and most isoforms in the lobule as well as the sublobular locationspecific induction of isoforms by inducers are manifestations of the hepatocyte heterogeneity. In addition, a layer of hepatocytes adjacent to the terminal hepatic venule or the portal area has been shown to have ultrastructure and/or functions differing from those of other perivenular or periportal hepatocytes (Hildebrand and Fuchs, 1984).

A. Total P-450 Sublobular distribution of total P-450 in rat livers has been studied by microphotometry (Gooding et al., 1978), by biochemical analysis of hepatocytes separated by gradient centrifigation (Gumucio et al., 1986), and by semiquantitative immunohistochemistry (Baron et al., 1982). These studies, except that of Gumucio et al. (1986), suggested that P-450 content in the cytoplasm is greater in perivenular hepatocytes than in periportal hepatocytes. Gumucio et al. (1986) reported that P-450 content (nanomoles per milligram of microsomal protein) in periportal hepatocytes is similar to that in perivenular hepatocytes. On the other hand, Kanai et al. (1990) found that P-450 content (amount per unit cytoplasmic volume) is greater in perivenular hepatocytes than in periportal hepatocytes, but P-450 density (molecule number per unit ER area) is not different in hepatocytes of all three zones in mice. Thus, the result of Gumucio et al. (1986) is similar to the sublobular distribution of the P-450 density, and the result of other investigators is similar to that of the P-450 content. Thus, a cause of the difference in the results between Gumucio et al. (1986) and other investigators may be the difference in the parameter used. In addition, the finding of Kanai et al. (1990) suggests that the lobular gradient seen in the P450 content simply reflects the lobular gradient of ER area. Furthermore, because the area of rough ER does not differ between periportal and perivenular hepatocytes in rats and mice (Kanai et al., 1986), the lobular

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gradient of P-450 content may correspond to the sublobular distribution of the area of smooth ER. The P-450 content in the cytoplasm and the density of the P-450 molecules in ER membrane are less in hepatocytes adjoining the terminal hepatic venule (1 perivenular hepatocytes) than in other perivenular hepatocytes in mice (Watanabe et al., 1992). In rats, however, the content and the density in 1 perivenular hepatocyte are not different from those in other perivenular hepatocytes (Watanabe and Kanamura, 1991). The rate of increase in the content and the density after PB (mice) (Watanabe et al., 1992) or MC (rats) (Tanaka et al., 1997) administration is lower in 1 perivenular hepatocyte than in other perivenular hepatocytes. Similarly, hepatocytes adjoining the portal area (one periportal hepatocyte) are also unusual; 1 periportal hepatocyte does not respond to PB stimulation in mice (Section III.D). The rate of increase in the content and the density (and the CYP1A1/2 content) after MC administration is lower in 1 periportal hepatocyte than in other periportal hepatocytes in rats (Tanaka et al., 1997).

B. Various Isoforms In normal mammals, P-450 isoforms are reported to be expressed restrictedly or predominantly in perivenular hepatocytes or evenly throughout the lobule (Table II). Administration of an inducer causes an increase in the inducer-specific isoform in perivenular hepatocytes or in the hepatocytes of the three zones. There are considerable discrepancies among results reported in normal mammals. For example, the CYP1A2 (human) is expressed restrictedly in perivenular hepatocytes (Ratanasavanh et al., 1991) and evenly throughout the lobule (Kirby et al., 1996); the CYP2B1/2 (rat) is observed restrictedly (Bu¨hler et al., 1992) and predominantly (Mino et al., 1998) in perivenular hepatocytes; the CYP2E1 (rat) is present restrictedly in perivenular hepatocytes (Bu¨hler et al., 1991, 1992), evenly throughout the lobule (Zerilli et al., 1995) and predominantly in perivenular hepatocytes (Weltman et al., 1996). A cause for these discrepancies is likely to be differences in experimental conditions of histochemical detection carried out by these investigators. A decrease in the intensity of immunostaining is known to occur by loss of antigenisity in sections during fixation, washing, and incubation with antibodies (Riederer, 1989). In addition, various factors such as steric hindrance, antibody trapping, and high antibody concentration impede binding of antibody to antigen in sections if antigen content in portions of sections is high (Chu et al., 1989; Riederer, 1989; Watanabe et al., 1991, 1994; Raivich

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TABLE II Sublobular Expression of Various P-450 Isoforms in the Liver Isoforms

Strain and sex

Expression

References

2B1/2

Alco mixed, male

2E1

Sprague-Dawley, male Alco mixed, male

Predominant in perivenular hepatocytesb Predominant in perivenular hepatocytesa Restricted in perivenular hepatocytesa Predominant in perivenular hepatocytesb Restricted in perivenular hepatocytesa Predominant in perivenular hepatocytesa Even throughout the lobulea Restricted in perivenular hepatocytesa Predominant in perivenular hepatocytesa

Tanaka et al., 1997

2A1

Sprague-Dawley, male Alco mixed, male

Rat 1A1/2

Wistar, male

3A1 3A1/2 Human 1A2

2A6 2B1 2E1

Wistar, male Alco mixed, male Wistar, male and female

Restricted in perivenular hepatocytesa Even throughout the lobulec Even throughout the lobulea Even throughout the lobulea Even throughout the lobulea Restricted in perivenular hepatocytesa Predominant in perivenular hepatocytesa

Bu¨hler et al., 1992 Bu¨hler et al., 1992 Mino et al., 1998 Bu¨hler et al., 1991, 1992 Weltman et al., 1996 Zerilli et al., 1995 Bu¨hler et al., 1992 Derbi et al., 1995

Ratanasavanh et al., 1991 Palmer et al., 1992 Kirby et al., 1996 Kirby et al., 1996 Kirby et al., 1996 Bu¨hler et al., 1991 Weltman et al., 1996

a

Immunohistchemistry. Quantitative immunohistchemistry. c In situ hybridization. b

et al., 1993). Thus, determining whether histochemical detections were carried out under suitable conditions in experiments of these investigators appears to be important (Table II). It is probable that the distribution of the CYP2B1/2 in Sprague-Dawley rats is predominantly perivenular because the experiments of Mino et al. (1998) were carried out upon careful consideration of the suitable conditions. The sublobular distribution of many P-450 isoforms is predominantly perivenular. Reductase, a valuable marker of the P-450 system because it is common to all isoforms other

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than certain mitochondrial forms, is expressed predominantly in perivenular hepatocytes (McManus et al., 1987; Watanabe et al., 1994). In addition, almost all the results (Table II) concerning sublobular expression of isoforms are simply visual observations of immunohistochemical reaction. This may be another cause for the discrepancies. Furthermore, species, strain, and/or sex differences may cause the discrepancies. The mRNA of the CYP2B1 and 2B2 is distributed homogeneously throughout the lobule in Sprague-Dawley rats, except for a narrow band of cells within the immediate vicinity of the periportal tract (Omiecinski et al., 1990). Rich and Boobis (1997) confirmed the same distribution for these enzymes in Sprague-Dawley rats, but found that the expression of these enzymes is perivenular in Wistar rats. Sex differences have been found in sublobular expression of isoforms. Although the CYP3A is expressed almost exclusively in perivenular hepatocytes, the absence of the enzyme in periportal hepatocytes is particularly clear in female rats (Oinonen and Lindros, 1995). The CYP4A, which is not expressed in normal rats, shows a marked lobular gradient with strong induction in perivenular hepatocytes in female rats after dehydroepiandrosterone treatment, but, in male rats, the enzyme is induced more uniformly thoughout the lobule (Beier et al., 1997). Sublobular location-specific induction of isoforms by inducers such as MC, PB, and ethanol has been reported. MC administration to rats causes an increase in the content of the CYP1A1/2 in the cytoplasm of hepatocytes of the three zones, although the increased value is greatest in perivenular hepatocytes (Tanaka et al., 1997). Dioxin administration to rats induces the CYP1B1 only in perivenular hepatocytes in spite of negligible expression in normal liver (Walker et al., 1998). In rats injected with a single dose of 80 mg/kg PB, the content of the CYP2B1/2 in the cytoplasm increases in perivenular and midzonal hepatocytes (Mino et al., 1998). Administration of 100 mg/kg PB once a day for 3 days to rats results in a relatively higher induction of the CYP2B1/2 in periportal hepatocytes, which becomes evenly distributed throughout the lobule, except for cells adjoining the portal tract (Bu¨hler et al., 1992). In rats treated with 16 alpha-carbonitrile, although CYP3A2 is induced in hepatocytes throughout the lobule, the induced enzyme shows greatest expression in perivenular hepatocytes (Debri et al., 1995). However, ethanol administration to rats causes an increase in the immunostaining of the CYP2E1 in perivenular hepatocytes (Bu¨hler et al., 1991; Weltman et al., 1996) and in perivenular and midzonal hepatocytes (Zerilli et al., 1995) and produces a perivenous ring pattern exhibiting the strongest staining in hepatocytes in six to eight cell layers from the terminal hepatic venule (Bu¨hler et al., 1992). Thus, compounds such as PB and MC appear to induce synthesis of inducer-specific isoform in hepatocytes of the three zones, although the induced isoforms generally reveal greatest

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expression in perivenular hepatocytes. Particularly, PB causes induction of the CYP2B1/2 first in perivenular and midzonal hepatocytes, and increases in the dosage of the inducer results in the enlargement of the area, in which induction of isoform occurs, to periportal hepatocytes. Similar enlargement of the area is found in total P-450 (Kanai et al., 1990; Watanabe et al., 1992). However, ethanol appears not to induce CYP2E1 in periportal hepatocytes. As mentioned already, hepatocytes of adult mammals show functional and structural heterogeneity within the lobule. At least two types of regulation of the hepatocyte heterogeneity were proposed (Gumucio, 1989): heterogeneity that may be controlled by factors transported by blood circulation and an imprinted pattern of zonal expression not susceptible to the sinusoidal microenvironment. The perivenular expression of total P-450 and various isoforms and the induction of P-450 isoforms in perivenular hepatocytes do not agree with the first hypothesis because of the blood flow from the portal tract to the terminal hepatic venule. Traber et al. (1989) found that rat CYP2B1/2 are induced 20- to 30-fold by PB treatment in only a fraction of hepatocytes transplanted 6 months earlier in the spleen, suggesting the location-specific induction of the CYP2B1/2 is not regulated by circulating factors. Similarly, retention of the zone-specific inducibility in isolated and cultured hepatocytes shows that the zonated response is independent of the lobular organization (Bars et al., 1989; Suolinna et al., 1989). Thus, most P-450 isoforms are expressed characteristically in a zonated pattern in the lobule. However, the factors responsible for the heterogenous expression are largely unknown. Oinonen et al. (1993) found that the expression of the CYP2B1/2 protein and mRNA in the normally silent periportal area becomes high after hypophysectomy in rats and that treatment with GH reestablishes perivenularly restricted expression that is original. In addition, Oinonen and Lindros (1995) observed a similar change in sublobular expression of the CYP3A proteins and the CYP3A2 mRNA after hypophysectomy and treatment with GH or tri-iodothyronine; they indicated that GH regulates the expression of the CYP2B1/2 and 3A genes and the thyroid hormone controls CYP3A genes sublobular location-specifically by suppressing their transcription in periportal hepatocytes. Because animals are usually subject to various inducers, it is probable that inductions of various types of isoform occur one after another in the liver. However, effects of an inducer on expression of the isoform induced antecedently by another inducer are unknown. Mino et al. (1998) found that MC administration drastically influences the pattern of expression of isoforms induced by PB in rats; the PB-induced increase in the content of the CYP2B1/2 in the cytoplasm is suppressed in perivenular hepatocytes but promoted in midzonal hepatocytes 48 hr after PB (80 mg/kg) and 24 hr after MC (25 mg/kg) injection. They also observed strong hybridiza-

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tion signal for the CYP2B2 mRNA in midzonal and perivenular hepatocytes and considered that the promotion of the increase in the isoform content in midzonal hepatocytes probably corresponds to the strong hybridization signal, whereas there appears to be an divergence between the intensity of the signal and the enzyme content in perivenular hepatocytes.

C. Postnatal Development Here we describe the development of sublobular expression of P-450 isoforms during formation of adult lobular organization. Rich and Boobis (1997) reviewed various data on expression and inducibility of isoforms during liver ontogeny. The higher level of total P-450 content in perivenular hepatocytes than in periportal hepatocytes is not present in fetal and newborn animals and forms during postnatal development. Watanabe et al. (1993b) showed in rats that the perinatal period is the time at which a marked increase in total P-450 content occurs in hepatocytes throughout the lobule, the subsequent period before weaning is the time at which sublobular heterogenous distribution appears, and the period after weaning is the time at which a slight increase in the content in periportal hepatocytes and a marked increase in perivenular hepatocytes takes place. A similar pattern of development was observed in reductase in rats, although the developmental increase in reductase content does not parallel that of total P-450 content in periportal or perivenular hepatocytes (Watanabe et al., 1996). The predominant expression in perivenular hepatocytes, which is seen in various isoforms of P-450, is also not present in fetal and newborn animals and is constituted during postnatal development. The CYP1A1 is detected in most rat hepatocytes in the lobule 72 hr before birth and is low to negligible 24 hr before and after birth. Five days after birth, the expression of the enzyme is greatest in hepatocytes (1–3 cell rows) lining the central venule (Rich and Boobis, 1997). The CYP1A2 was detected in most rat hepatocytes 72 hr before birth, through parturition and for the first 5 days after birth, and by 15 days, the expression is greatest in perivenular hepatocytes (Rich and Boobis, 1997). The CYP1A2 (Ratanasavanh et al., 1991) was not detected, and the CYP1A and 3A (Murray et al., 1992) were evenly distributed throughout the lobule in human fetus, and these enzymes were expressed only or mainly in perivenular hepatocytes in the adult. The CYP2C was generally absent and the CYP3A was present evenly throughout the lobule in fetal and newborn human livers; preferential localization in (midzonal and) perivenular hepatocytes appears during the first few weeks after birth and in 3-year-old and older humans (Ratanasavanh et al., 1991).

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V. Concluding Remarks Information on mechanisms of transcriptional regulation of P-450 genes has been accumulated to some extent. In particular, the regulatory mechanisms for the CYP1A and 4A genes have been well characterized. However, there are still many problems to be resolved. For example, although many cis-acting elements are found in genes of various P-450 isoforms, only a few transacting factors that actually bind to the elements are identified. Furthermore, intracellular receptors for P-450 inducer molecules are unknown for many isoforms, although only two receptors—Ah receptor for the CYP1A1 gene and PPAR for the CYP4A1/6 gene—are characterized. Thus, detailed mechanisms for transcriptional regulation of many P-450 genes through interaction between the elements and the factors are still unclear. Moreover, there is little information on posttranscriptional regulation of various P-450 isoforms. Studies on the topology of enzymes constituting the microsomal monooxygenase system have been carried out biochemically, biophysically, and immunoelectron-microscopically. However, almost all of the information obtained from these studies is based on assumptions. Molecular morphological methods by electron or atomic force microscopy are considered to be necessary for obtaining more accurate information. For example, the topology of enzymes of microsomal monooxygenase system should be analyzed in situ by using a region-specific monoclonal antibody and a 1-nm colloidal gold-labeling method. The interaction between P-450 and reductase molecules in ER membrane may be elucidated further by use of immunofreeze fracture method or atomic force microscope. Because the densities of P-450 and reductase molecules in ER membrane have been known, we can incorporate the same densities of the two enzymes into artificial lipid membrane and use this as the material of analysis. Using molecular morphological methods probably will bring about more progress in future studies on drug metabolism.

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Alternative Protein Sorting Pathways John Kim, Sidney V. Scott, and Daniel J. Klionsky Section of Microbiology, University of California, Davis, California 95616

The term ‘‘nonclassical protein targeting’’ has been used to describe those pathways that have been recently discovered and differ mechanistically from the more studied ‘‘classical pathways.’’ Because this nomenclature is rather arbitrary in terms of cellular relevance, we have chosen to group these protein sorting mechanisms under the heading ‘‘alternative protein sorting pathways’’ for the purpose of this review. Many of the alternative targeting pathways described are of primary importance. For example, without retrograde transport, both membrane material and targeting machinery accumulate at distal sites in the endomembrane system, preventing anterograde transport. Further, lysosome/vacuole delivery of degradative substrates by autophagic pathways is central to the role of this organelle as a primary site for intracellular degradation. Finally, targeting through the classical CPY pathway requires the ALP pathway for delivery of the vacuolar t-SNARE Vam3p. Analysis of these alternative targeting pathways provides a more complete understanding of eukaryotic cellular physiology. KEY WORDS: Aminopeptidase I, Autophagy, Cvt pathway, Lysosome, Protein targeting, Protein degradation, Retrograde transport. 䊚 2000 Academic Press.

I. Introduction Over the last 20 years, biologists have gained tremendous insight into the mechanisms used to achieve the correct localization of proteins throughout the cell. Initial observations led to the generation of specific models for protein transport. These models have undergone considerable revision but have resulted in the establishment of certain paradigms. For example, after initially translocating into the endoplasmic reticulum (ER), proteins destined for exit from this organelle were thought to leave by a bulk flow process that was signal-independent. These proteins would enter vesicles budding from the ER that would subsequently fuse with the Golgi complex. International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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Continued transport throughout the Golgi complex would occur via a series of forward-directed budding and fusion events. Even though the value of this basic model in directing our scientific thinking is evidenced by the molecular details that have been gained about this process, many of the basic details have been called into question. Proteins exit the ER due to the presence of specific transport signals, and are packaged into transport vesicles via interaction with cargo receptors. Resident organellar proteins are actively retained and retrieved. Transport through the Golgi complex is likely to require vesicle budding and fusion, but the direction of vesicle transport may be primarily in the retrograde direction. Similarly useful paradigms have been established for other transport processes. For example, delivery of resident hydrolases to the lysosome/ vacuole was shown to involve a portion of the secretory pathway. After transit to the trans-Golgi Network, lysosomal/vacuolar proteins are segregated from resident Golgi complex and secretory proteins and are localized to the vacuole via an endosomal intermediate. Each transport step was presumed to involve standard vesicle budding and fusion reactions. Continued research into this area has now revealed multiple pathways that are used for the delivery of proteins to the vacuole. Aminopeptidase I is a resident hydrolase that is targeted to this organelle independent of the secretory pathway. Even for those proteins that transit to the trans-Golgi Network, there are at least three different mechanisms used for the ultimate delivery step. The more recently discovered targeting pathways have been termed ‘‘nonclassical.’’ This term may be misleading because the number of exceptions to the ‘‘classical’’ rules continues to increase. In fact, the number of and variation in the mechanisms used in protein targeting are astounding. For each organelle, there are a range of processes used to deliver specific proteins to their correct destination. In this article, we have examined some of the recent research into alternative targeting pathways. This topic cannot be fully covered in a single review. Instead, it is our intention to point out how our current understanding of cellular transport processes has evolved and to highlight some of the directions for future research into this exciting and dynamic topic.

II. Retrograde Transport A. General The vectorial forward movement of transport vesicles carrying proteins destined for secretion remains a conceptual hallmark of cell biology (Palade,

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1975). In the classical model, proteins are cotranslationally translocated into the ER. They undergo proteolytic removal of the signal sequence and addition of core carbohydrate residues. The presence of transport signals allows these proteins to be sorted from resident ER proteins and packaged into vesicles that bud from the surface of the organelle. Fusion of the vesicles with the cis-Golgi complex releases the protein into the lumen. Movement of the proteins through the stacks of the Golgi complex is accompanied by additional glycosyl modifications. At the trans-Golgi Network (TGN), the proteins are packaged into secretory vesicles that fuse with the plasma membrane, releasing the proteins at the cell surface. According to this model, transport through the secretory pathway is defined by a series of vesicle budding and fusion events. Molecular components including cargo receptors, v- and t-SNAREs, regulatory factors, and vesicle coat proteins have been demonstrated to be necessary for this complex transport process (Rothman and Wieland, 1996; Schekman and Orci, 1996). 1. Coat Proteins Play an Integral Role in Vesicle Formation A major breakthrough in our understanding of the mechanism of vesiclemediated protein transport came with the discovery of coat protein complexes. The clathrin-coated vesicles were the first to be discovered and serve as a paradigm for the role of vesicle coat complexes in cargo selection and concentration, and vesicle budding (Schmid, 1997; Hirst and Robinson, 1998). Vesicle coat components bind to the appropriate membrane and assemble into hetero-oligomeric complexes that mark the site of vesicle budding. Coat protein subunits interact with sorting signals on cargo molecules and receptors, sorting and concentrating appropriate cargo to be packaged into the emerging vesicle. The coat complexes involved in the early secretory pathway are the COPI (coatomer) and COPII complexes (Lowe and Kreis, 1998; Barlowe, 1998; Springer et al., 1999). The COPI coat proteins have been characterized in detail at the molecular level and consist of seven subunits (움, 웁, 웁⬘ 웂, ␦, ␧, ␨) that self assemble in the cytoplasm and then nucleate coat formation on Golgi membranes in an ARF (ADP-ribosylation factor) GTPase-dependent manner. The COPI complex appears to be involved in retrograde transport within the Golgi as well as between the Golgi and ER. The COPII complex was isolated more recently (Barlowe et al., 1994) and consists of Sec13p, Sec31p, Sec23p, Sec24p, and the GTPase Sar1p. A recent study reconstituted the formation of COPII-coated vesicles from purified coat proteins and liposomes (Matsuoka et al., 1998). The sequential recruitment of Sec23p/ Sec24p to the GTP-bound form of Sar1p on liposomes or ER was followed by the binding of Sec13p/31p to this complex. Whereas COPI appears to play a role in various transport pathways (Gaynor et al., 1998; Geli and

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Riezman, 1998), COPII vesicles appear to be restricted to anterograde transport from the ER to the Golgi. Thus, the type of coat complex may, in part, determine the direction of cargo movement for the step in which it is employed. 2. Retrograde Transport Is Needed to Balance Forward Movement The continuous forward flow of membranes and transport machinery results in an accumulation of these components in addition to the expansion of membrane material at the acceptor compartment. Further, distinct intracellular compartments must maintain their unique membrane composition and resident proteins that define each organelle. Therefore, it stands to reason that, for every anterograde trafficking step of the endomembrane system, there must be a reciprocal retrograde step. Retrograde transport will allow recycling of v-SNAREs, cargo receptors, and resident proteins that have left the donor compartment. This coupling of anterograde and retrograde transport explains why many of the recycling components such as sorting receptors are rarely found in one distinct compartment. Here we discuss four targeting steps that employ retrograde movement of transport vesicles: (1) COPI-mediated Golgi-to-ER transport; (2) intra-Golgi traffic by cisternal maturation and COPI-mediated vesicles; (3) endosome to Golgi recycling mediated by a novel coat complex, the retromer; and (4) evidence for retrograde traffic out of the vacuole, which has been generally thought to be a terminal organelle of the endomembrane system.

B. Golgi to ER Retrograde Transport 1. Retrograde Transport Is Required for Retrieval of ER Resident Proteins and Recycling of Components for Anterograde Transit The Golgi-to-ER retrograde pathway serves not only to retrieve resident ER proteins that have escaped retention but also to recycle proteins required for subsequent anterograde movement. Two well-characterized retrieval sequences have been identified for the retrieval/retention of ER resident proteins (Pelham, 1994, 1996; Gaynor et al., 1998). For soluble, resident ER proteins, the C-terminal tetrapeptide sequence KDEL for mammals (Munro and Pelham, 1987) and HDEL for yeast (Pelham et al., 1988) serves as the Golgi-to-ER retrieval signal. K/HDEL-containing ER proteins that have escaped the ER bind Erd2p, the K/HDEL receptor. Erd2p is localized to the cis-Golgi complex and mediates the recycling of these proteins back to the ER (Lewis and Pelham, 1990; Semenza et al.,

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1990; Lewis and Pelham, 1992). Several lines of evidence demonstrate that the K/HDEL retrieval is mediated by COPI vesicles. First, SEC21, which encodes the 웂-COP subunit of the coatomer, is required for the Golgi-toER retrieval of Erd2p (Lewis and Pelham, 1996). In addition, the Erd2p/ KDEL-protein complex has been localized by immunogold electron microscopy to mammalian COPI vesicles, suggesting that the K/HDEL-containing proteins use COPI vesicles in retrograde transport (Orci et al., 1997). In vivo evidence of COPI involvement in Golgi-to-ER retrograde transport was demonstrated recently using cholera toxin (Majoul et al., 1998). After endocytosis, the A subunit of cholera toxin (CTX-A) binds to Erd2p via its C-terminal KDEL motif followed by retrograde transport to the ER. Injecting Vero cells with function-blocking antibodies to Erd2p, the 웁 subunit of COPI, or p23 (a member of the p24 protein family of cargo receptors) inhibited the retrograde transport of CTX-A from the Golgi to the ER (Majoul et al., 1998). Finally, a recent study reconstituted Golgi-to-ER retrograde transport in a cell-free system. Using the yeast pheromone 움-factor fused to the HDEL retrieval sequence as a marker (Dean and Pelham, 1990), Spang and Schekman demonstrated that a simple set of purified cytosolic components was required for the retrograde reaction including the coatomer complex, as well as the Golgi v-SNARE Sec22p, the ER t-SNARE Ufe1p, and Emp47p, a cargo protein containing the dilysine Golgi-to-ER retrieval signal (Lewis and Pelham, 1996; Spang and Schekman, 1998). For type I membrane proteins of the ER, the dilysine motif KKXX at the C-terminal cytosolic domain acts like a Golgi-to-ER retrieval signal ( Jackson et al., 1990, 1993; Gaynor et al., 1994; Townsley and Pelham, 1994). In vitro evidence demonstrated that coatomer subunits (움, 웁, 웁⬘, and 웂) bind to a yeast protein (Wbp1p) containing the dilysine retrieval motif, suggesting a physical interaction between the retrieval signal and some of the subunits of the COPI coat (Cosson and Letourneur, 1994). Yeast mutants in coatomer subunits 움-COP (retl-1), 웁⬘-COP (sec27-1), and 웂-COP (sec21-1 and sec21-2) were shown to be defective in the retrieval of a fusion protein consisting of the 움-factor receptor (Ste2p) and the cytosolic domain of Wbp1 containing the dilysine-retrieval motif (Letourneur et al., 1994). Additional analysis with two temperature-sensitive (ts) mutants for ␦-COP (ret2-1) and ␨-COP (ret3-1) also indicated a Golgi-toER retrograde transport defect of a marker protein containing the dilysine retrieval motif (Cosson et al., 1996). In vivo evidence for COPI-mediated Golgi-to-ER retrograde transport was also demonstrated by examining the fate of Emp47p, a cargo protein containing the dilysine motif in its cytosolic tail, and Erd2p, the HDEL retrieval receptor (Lewis and Pelham, 1996). Mutations in Sec21p (웂-COP) as well as Ufe1p, the ER-localized t-SNARE required for retrograde vesicle transport, caused a defect in the Golgi-to-

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ER recycling of Emp47p and Erd2p (Lewis and Pelham, 1996). A recent in vitro study defined the cytosolic requirements of COPI vesicle budding from artificial lipid bilayers whose composition mimicked mammalian cell membranes including the Golgi. In this study, COPI vesicle budding did not occur unless a bivalent interaction of the coatomer with membranebound ARF-GTP and with the cytosolic tails of transmembrane, dilysinetagged cargo occurred. Therefore, retrograde cargo uptake and coatomer assembly appear to be coupled events in transport vesicle formation (Bremser et al., 1999). 2. Coatomer May Function Primarily in Retrograde Transport Although the function of COPI vesicles in retrograde Golgi-to-ER transport is well established, its direct role in ER-to-Golgi anterograde transport has been a point of debate (Pelham, 1994; Cosson and Letourneur, 1997; Schekman and Mellman, 1997; Gaynor et al., 1998). Two general models emerge from COPI-mediated transport studies between the ER and Golgi. In one model, COPI vesicles transport cargo in both anterograde and retrograde directions. The alternative model postulates that the effects of COPI mutations on anterograde transport are indirect. Factors necessary for anterograde transport (e.g., cargo receptors) must be recycled back to the donor compartment for successive rounds of anterograde vesicle budding, docking, and fusion. If the retrograde pathway recycling these transport components is blocked, then anterograde transport would also cease. Therefore in this model, COPI vesicles are used only for retrograde traffic, whereas COPII vesicles mediate all anterograde transport between the ER and Golgi (see Fig. 1). Evidence for COPI-mediated anterograde transport between the ER and Golgi stems from temperature-sensitive yeast coatomer mutants sec21-1 (웂-COP), ret2-1 (␦-COP), sec27-1 (웁⬘-COP), as well as from the deletion mutant sec26⌬ (웁-COP), which all show defects in the anterograde transport of carboxypeptidase Y (CPY) from the ER to the Golgi (Hosobuchi et al., 1992; Duden et al., 1994; Cosson et al., 1996). COPI-rich regions of ER have been observed in mammalian cells (Orci et al., 1994), and budding of COPI-coated vesicles from purified nuclei (ER membranes) has been demonstrated in vitro (Bedmarek et al., 1995). However, unlike COPII vesicles, the ER-derived COPI vesicles were devoid of any known cargo (Bedmarek et al., 1995). Therefore, in vitro results may not faithfully represent physiological events. Coatomer interactions in vitro with various membrane sources indicate that these coat components can assemble onto every secretory compartment tested thus far (Cosson and Letourneur, 1997). Further, phenotypes of mutations can vary greatly depending on the type of screening procedures used to isolate the mutants. All the original COPI

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mutants were isolated based on screens that either selected for secretory defects (sec mutants) or retrieval defects (ret mutants) and subsequent analyses of these mutants merely confirmed the strategy of the original screens. For example, sec21-1 was isolated as a mutant defective in anterograde ER-to-Golgi transport (Hosobuchi et al., 1992) whereas sec21-2 was characterized as a retrograde-defective mutant (Letourneur et al., 1994). In order to eliminate the bias of screening procedures, a recent study employed a random PCR mutagenesis approach to obtain additional alleles of SEC21 based on temperature-conditional growth rather than screening for directional transport defects (Gaynor and Emr, 1997). In all the new sec21 ts mutants, only a subset of cargo molecules (CPY and 움-factor) was blocked from exiting the ER, whereas others (HSP150 and invertase) were secreted normally in these as well as in other COPI mutants. In contrast, COPII vesicle components, Sec18p (NSF) and Sec22p (v-SNARE for ER to Golgi transport), were required for the delivery of all cargo molecules examined (Gaynor and Emr, 1997). This study suggests that the forward transport defects in the sec21ts were indirect and were caused by the failure of mutant COPI vesicles to recycle proteins required for packaging of specific cargo into anterograde COPII vesicles. Taken together, these recent studies appear to indicate that the role of COPI vesicles in traffic between the ER and Golgi is exclusively in the retrograde direction, whereas COPII transport vesicles handle the anterograde delivery of cargo (Fig. 1).

C. Two Models of Intra-Golgi Transport The debate over the role of COPI vesicles in anterograde versus retrograde transport between the ER and Golgi also extends to the models of intraGolgi traffic (Schekman and Mellman, 1997; Glick and Malhotra, 1998; Gaynor et al., 1998). The classic paradigm of vesicular transport states that all traffic within the Golgi stacks is mediated by COPI transport vesicles carrying cargo both in the anterograde and retrograde directions (Rothman and Wieland, 1996). In this model, the Golgi stacks are viewed as stable compartments (cis, medial, trans) which are defined, in part, by the differential concentration of various resident Golgi enzymes. In the alternative, cisternal progression/maturation model, the Golgi stacks are proposed to be dynamic structures that begin as cis-Golgi and mature into more distal Golgi structures, until they are finally consumed as secretory vesicles at the trans-Golgi Network. Evidence for both models is discussed below. As with traffic between the ER and Golgi, both intra-Golgi transport models support COPI-mediated intra-Golgi retrograde transport. However, in the bidirectional vesicular transport model, retrograde transport functions primarily as a recycling mechanism, even though it is the primary mechanism

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by which the different Golgi stacks are formed in the cisternal maturation model. 1. Vesicle Transport Model The role of the coatomer complex was first identified in an in vitro analysis of intra-Golgi transport (Balch et al., 1984; Orci et al., 1986). In the transport assay, donor Golgi membranes were prepared from a cell line expressing the vesicular stomatitis virus (VSV) G protein and were deficient in N-acetylglucosamine (GlcNAc) transferase I, while wild-type Golgi membranes with functional GlcNAc transferase I were used as the acceptor. The addition of GlcNAc to VSV G protein was interpreted as evidence of anterograde intra-Golgi transport (Ostermann et al., 1993). The addition of GTP웂S caused the accumulation of nonclathrincoated vesicles which were subsequently identified as the subunits of the COPI complex (Malhotra et al., 1989; Waters et al., 1991). Even though the findings from this in vitro assay have long been used as key evidence for COPI-mediated anterograde intra-Golgi transport, it appears equally feasible that GlcNAc addition to VSV G protein could have occurred via retrograde transport of GlcNAc transferase I to the donor compartment (Featherstone, 1998). In addition, COPI yeast mutants that completely block ER-to-Golgi transport of specific cargo proteins (via a direct block in Golgi-to-ER retrograde transport) do not show any defects in intra-Golgi anterograde transport (Gaynor and Emr, 1997). Other evidence of COPI-mediated anterograde intra-Golgi transport include the use of brefeldin A (BFA). ARF, in its activated GTP-bound form, binds to Golgi and subsequently recruits coatomer assembly on the membranes for vesicle formation. BFA behaves as an uncompetitive inhibitor in that it binds the transient complex formed by ARF-GDP and the Sec7 domain of the nucleotide exchange factors, leading to an abortive ARFGDP:BFA:Sec7 domain complex; thus, BFA causes ARF-GDP to act as a dominant negative mutant (Peyroche et al., 1999). This block causes the Golgi complex to disassemble and redistribute to the ER, making BFA a potent inhibitor of anterograde secretion (Lippincott-Schwartz et al., 1989; Peyroche et al., 1999; Roth, 1999; Chardin and McCormick, 1999). Also, function-blocking antibodies to the 웁-COP subunit have been shown to inhibit intra-Golgi anterograde transport (Pepperkok et al., 1993). However, these effects may be indirect consequences of direct defects in intra-Golgi retrograde transport as discussed earlier. Nevertheless, a recent study provides intriguing evidence that COPI vesicles mediate both anterograde and retrograde intra-Golgi transport (Orci et al., 1997). Using immunogold electron microscopy, COPI vesicles containing both anterograde-directed cargo (proinsulin, VSV G protein) and retrograde-directed cargo (Erd2p, the KDEL receptor) were shown budding from every level of the Golgi stacks in whole cells. Interestingly, forward cargo and retrograde cargo

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were segregated into two distinct populations of COPI vesicles. The in vivo findings were confirmed in a cell-free Golgi transport system in which VSV G protein and the KDEL receptor were found in separable vesicles, suggesting that two populations of COPI vesicles exist, one mediating anterograde transport and the other for retrograde transport (reviewed in Schekman and Mellman, 1997). 2. The Cisternal Maturation Model The cisternal maturation/progression model of intra-Golgi transport postulates that COPI vesicles operate exclusively in the retrograde direction (see Fig. 1). No vesicle-mediated forward transport is required as the maturation of each Golgi stack occurs en bloc. In this model, COPII vesicles derived from the ER form the ERGolgi-intermediate compartment (ERGIC) which, in turn, forms the cisGolgi by a series of homotypic vesicle fusion events (Mironov et al., 1997; Glick et al., 1997; Glick and Malhotra, 1998). ER-resident proteins and recycling machinery (e.g., cargo receptors and SNAREs) are returned to the ER via COPI-mediated retrograde transport. Concurrently, enzymes from the medial-Golgi are delivered to the cis compartment via retrograde transport such that the cis-Golgi effectively ‘‘matures’’ into the medialGolgi. The medial-Golgi, in turn, progresses into the trans-Golgi as it receives, via retrograde transport, the set of proteins that define the trans compartment. Finally, cargo destined for secretion is released from the TGN through the formation of secretory granules or vesicles. Resident TGN proteins are delivered via retrograde transport to the trans-Golgi, thereby maturing this compartment into the next TGN. Therefore, in each ‘‘cycle’’ of cisternal maturation, a new cis-Golgi is formed, the TGN fragments and is converted into transport vesicles, and each of the other cisternae matures by one step (Glick et al., 1997). Early evidence for the cisternal progression model comes from observations of scale-covered green algae (Becker and Melkonian, 1996; Mironov et al., 1997). Scales are electron-dense macromolecular glycoconjugates that are assembled inside the Golgi cisternae and appear to be transported to the cell surface by the progression of Golgi cisternae. Direct fusion of the cisternae with the plasma membrane releases each scale. Because of their size (up to 20 times the size of a typical transport vesicle), scales would not fit inside a typical transport vesicle and are not seen in the vesicles that bud from the Golgi stacks. Other examples of transport of macromolecular structures in both plant and animal cells suggest that the mechanisms driving scale secretion are not unique to some algae. A recent study examining the transport of procollagen provided the best evidence for the cisternal maturation model (Bonfanti et al., 1998). Procollagen assembles into 300-nm rigid, rodlike triple helices in the ER and moves to the Golgi complex where it forms electron-dense aggregates. Examining serial section

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reconstructions of the Golgi stacks by immunolabeled electron microscopy, Bonfanti et al., observed that the procollagen aggregates remained in the lumen of the Golgi cisternae at all times during its movement across the Golgi stacks (Bonfanti et al., 1998). In addition, Bonfanti et al., were able to synchronize the transport of procollagen to the Golgi by a drug that reversibly inhibited the proper folding of procollagen in the ER, thereby preventing export to the Golgi. Washing the drug effectively restored proper folding and subsequent transport to the Golgi. In this way, a synchronous population of procollagen migration was observed. Procollagen moved throughout the Golgi stacks in the cis–trans direction without ever entering conventional transport vesicles or large dissociative elements, thus providing strong evidence for the cisternal maturation model. Although it would be most satisfying to have a single, unified model of Golgi transport, both anterograde vesicular transport and cisternal maturation may occur. Vesicular transport may deliver small proteins for secretion, whereas cisternal maturation may account for the transport of macromolecular structures. In addition, other models involving tubular connections among the Golgi cisternae may also play a role in intra-Golgi transport (Mironov et al., 1997). Regardless of the method of forward transport, COPI-mediated retrograde transport of Golgi cargo appears to occur (Sonnichsen et al., 1996; Orci et al., 1997; Love et al., 1998).

D. Endosome-to-Golgi Retrograde Transport: A Novel, Membrane Coat Complex, the Retromer Genetic screens in Saccharomyces cerevisiae have identified at least 40 genes (VPS for vacuolar protein sorting) whose products are required for the delivery of resident hydrolases to the vacuole (Bankaitis et al., 1986; Rothman and Stevens, 1986; Robinson et al., 1988). Among the transport machinery identified is the vacuolar hydrolase sorting receptor Vps10p which binds to its ligands CPY and other hydrolases at the TGN and delivers its cargo to the endosome or prevacuolar compartment (PVC; Marcusson et al., 1994; Westphal et al., 1996). After transport to the PVC, Vps10p, as well as resident Golgi transmembrane proteins that have escaped the late Golgi (e.g., Kex2p and DPAP A), are recycled back to the Golgi via retrograde transport (Cereghino et al., 1995; Cooper and Stevens, 1996). This recycling pathway requires a specific retrieval signal on the cytosolic domains that generally includes two important aromatic amino acids, F/YXF/Y. In addition, the cytosolic domains of the cargo are thought to be important regions for interactions with cytosolic factors that associate peripherally with the donor membranes and sort and package the cargo into transport vesicles (Burd et al., 1998).

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Initial studies demonstrated that a number of VPS genes were required for the proper localization of Vps10p, including VPS5, 17, 29, 30, and 35 (Horazdovsky et al., 1997; Nothwehr and Hindes, 1997; Seaman et al., 1997). Vps5p and Vps17p could be also biochemically crosslinked to each other, suggesting a physical interaction (Horazdovsky et al., 1997). Mutations in these genes resulted in the mislocalization of Vps10p to the vacuole and caused CPY to be secreted from the cell (Seaman et al., 1997). A recent study has demonstrated that five peripheral membrane proteins—Vps5p, 17p, 35p, 29p, and 26p—interact physically with each other to form a large protein complex, termed the retromer, on a prevacuolar membrane (Seaman et al., 1998). The proper function of the retromer components is essential for efficient endosome to Golgi retrograde movement of the CPY sorting receptor Vps10p. Further, cross-linking experiments in vps5 and/or vps17 mutants indicated that Vps35p, 29p, and 26p form a subcomplex as do Vps5p and 17p. Gel filtration experiments also indicated that Vps17p and 5p heterodimers may oligomerize into higher-order structures. Sucrose density gradients demonstrated that the complex colocalizes with the endosomal marker Pep12p and a vesicle fraction of intermediate density between the endosome and dense COPI vesicles. Immunolabeled electron microscopy also indicates that Vps5p of this complex localizes to endosomal membranes in vivo and to intermediate-sized vesicles. Taken together, these results indicate that the Vps35p, 29p, 26p, 17p, and 5p proteins may serve as a novel membrane coat complex required for retrograde transport from the endosome to the Golgi (Seaman et al., 1998). Analogous to cargo selection and concentration in anterograde transport, the subcomplex Vps35p, 29p, 26p may associate with the cytoplasmic tails of Vps10p as well as other cargo and package them into transport vesicles. The finding that these complexes can assemble into higher-order structures also supports the idea that the retromer may serve as a multiplyiterated vesicle coat complex (see Fig. 1). Purification of the retromercoated transport vesicles and the identification of additional components remain to be investigated.

E. A Novel Vacuole-to-Endosome Retrograde Transport Pathway The vacuole contains an array of digestive enzymes which function to degrade delivered cellular material (Klionsky et al., 1990). Therefore, the vacuole was considered the terminal compartment of the endosomal pathway. However, this view is problematic because a substantial influx of membranes and vesicle transport machinery would occur without a compen-

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satory mechanism to maintain vacuole size and recycle vesicle transport machinery for successive rounds of transport. Using a hybrid reporter protein, Bryant et al. recently demonstrated that retrograde movement out of the vacuole was possible (Bryant et al., 1998). Previous studies had demonstrated that the cytosolic tail domain of the vacuolar hydrolase alkaline phosphatase (ALP) encoded sequence determinants required for its transit to the vacuole directly from the TGN by the ALP Pathway (Cowles et al., 1997b; Piper et al., 1997; see Section IV.B). A chimeric marker protein, RS-ALP (retention sequence-ALP) was created by replacing the vacuolar localization motif of ALP with a 10 amino acid sequence containing the FXFXD retrieval motif from the cytosolic tail of the resident TGN protein dipeptidyl aminopeptidase (DPAP) A (Nothwehr et al., 1993). In wild-type cells, this aromatic amino-acid containing sorting motif is required for the efficient retrieval of DPAP A from the PVC to the TGN (Nothwehr et al., 1993; Bryant and Stevens, 1997). The transplantation of the FXFXD retrieval signal to RS-ALP resulted in the steady-state localization of RS-ALP to the TGN as observed by indirect immunofluorescence microscopy and colocalization with the Golgi marker Kex2p (Bryant et al., 1998). However, a series of pulse-chase immunoprecipitation experiments indicated that, like wild-type ALP, the lumenal domain of RS-ALP was processed in a PEP4-dependent manner. These experiments indicated that newly synthesized RS-ALP first traveled to the vacuole via the ALP pathway where it was proteolytically processed. Then, by virtue of the FXFXD retrieval signal, it returned to the PVC by a novel retrograde pathway before being retrieved back to the TGN (see Fig. 1). This provided the first evidence of retrograde transport out of the vacuole (Bryant et al., 1998). In order to prepare for successive rounds of vesicle-mediated transport between two compartments, sorting machinery must be recycled back from the acceptor to the donor compartment. For example, the CPY receptor binds CPY at the TGN and delivers it to the endosome for subsequent transport to the vacuole. The CPY receptor itself is retrieved back to the TGN for successive rounds of CPY delivery (Conibear and Stevens, 1998; Bryant and Stevens, 1998). Consistent with this model, Bryant et al. demonstrate that Vti1p, the v-SNARE required for the delivery of both CPY and ALP to the vacuole, also recycles between the PVC and the vacuole (von Mollard et al., 1997; Fischer von Mollard and Stevens, 1999; Bryant et al. 1998). This work further revealed that Vac7p, a vacuolar protein required for vacuole inheritance (Bonangelino et al., 1997), was also required for the retrograde movement of RS-ALP from the vacuole to the PVC. Other components of this pathway remain to be identified. Some candidates include AP2/clathrin coats, which have been demonstrated to specifically

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assemble on the surface of lysosomes and may participate in the formation of the retrograde transport vesicles (Traub et al., 1996).

III. Degradative Pathways A. Macroautophagy Delivers Cytosolic Components to the Lysosome/Vacuole for Degradation When cells are stressed for nutrients, they respond by targeting a portion of their cytosolic contents to the lysosome/vacuole compartment for turnover and recycling of macromolecular components. The macroautophagy pathway accomplishes this process. The morphology of macroautophagy has been studied in cultured hepatocytes subjected to serum deprivation (Blommaart et al., 1997; Kadowaki et al., 1996; Mortimore et al., 1996), in tobacco or sycamore suspension cultures stressed for sucrose (Aubert et al., 1996; Journet et al., 1986; Moriyasu and Ohsumi, 1996), and in S. cerevisiae in media lacking carbon or nitrogen (Takeshige et al., 1992). The basic morphology of the autophagic process appears to be conserved across all of these organisms. From these studies, it is clear that double membrane vesicles (termed autophagosomes in yeast and initial autophagic vacuoles (AVi) in mammalian cells) enclose portions of bulk cytosol (Baba et al., 1994; Dunn, 1990a, 1990b). The outer membrane of these vesicles fuses with the vacuole in yeast to release a still-intact vesicle into the vacuole lumen (called an autophagic body). In proteasedeficient yeast, autophagic bodies accumulate inside the vacuole, indicating that these vesicles are ultimately degraded in a protease-dependent manner (Takeshige et al., 1992). In mammalian cells, the autophagosomes appear to fuse with an intermediate endosomal compartment forming intermediate autophagic vacuoles (AVi/d). The AVi/d vesicles then acquire proteases and acidic pH in stepwise manner developing into degradative autophagic vacuoles (AVd) that degrade the remaining internal membranes and their cytosolic contents (Dunn, 1994; Lawrence and Brown, 1992; Liou et al., 1997). The subcellular origin of the membrane that forms autophagosomes has not been definitively identified. In addition, whether the vesicles form de novo or by an existing organelle extending to engulf a portion of cytosol is not known. In mammalian cells, studies using immunogold labeling have identified the presence of rough ER proteins on some AVi membranes (Dunn, 1990a). Because rough ER protein staining is not present on all AVi membranes and, further, these membranes do not appear to exactly resemble any known subcellular compartment, a specialized membrane

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source termed the phagophore has been proposed (Seglen et al., 1996). In S. cerevisiae, histochemical staining of autophagosomal membranes to assess their carbohydrate content, and therefore their origin, found only light staining suggesting that they were derived from an early secretory pathway membrane (Baba et al., 1994). Wherever the autophagosomal membrane originates, proteins such as v-SNARES that would be necessary for vacuole recognition and fusion must end up on the sequestering membranes. In this respect, the endosome would be a logical donor membrane. In fact, recent studies of the molecular components required for autophagy have implicated several endosomal proteins in the autophagic process (see following discussion). The significance of these findings in terms of the identification of the donor membrane for autophagosomes is not yet clear, but they suggest that endosomal membranes may be involved. 1. Molecular Components of Macroautophagy Two groups of autophagy mutants have been isolated in S. cerevisiae. These mutants are named apg (Tsukada and Ohsumi, 1993), and aut (Thumm et al., 1994). In addition, a number of mutants defective in the cytoplasm to vacuole targeting (cvt) pathway are also autophagy defective (Harding et al., 1996, 1995; Scott et al., 1996). The Cvt pathway is a constitutive biosynthetic pathway that targets aminopeptidase I (API) to the vacuole (see Section IV.A). This process utilizes double membrane vesicles (called Cvt vesicles) that are morphologically similar to autophagosomes to selectively target API to the vacuole when cells are grown under vegetative conditions (Baba et al., 1997; Scott et al., 1997). In addition, API is specifically targeted to the vacuole by autophagosomes during starvation (Baba et al., 1997; Scott et al., 1997). Because API is selectively transported by autophagosomes or by the similar Cvt vesicles, it is a useful molecular marker for studies of the autophagic process. Cloning of the genes responsible for the apg, aut, and cvt mutations, as well as analyzing API to characterize the Cvt/autophagy defects of other known yeast mutants has led to the identification of a number of the molecular components required for these pathways (Table I). The largest class of proteins involved in autophagy and Cvt transport identified so far are involved in early steps in the process before vesicle formation takes place. These proteins include Apg1p, Apg5p, Apg7p, Apg10p, Apg12p, Apg13p, and Apg16p. Apg5p is the best characterized protein in this class. Analysis of the localization of precursor API (prAPI) in a temperature sensitive apg5 allele revealed that prAPI was bound to membranes, but not enclosed within vesicles (George et al., 1999). This result indicates that Apg5p acts in a step before vesicle formation. Apg5p is a 32-kDa protein that is covalently

TABLE I Proteins Required for Precursor API Import and Autophagy Gene

Topology/localization

Function/interactions

APG1/AUT3/CVT10 APG5 APG6/VPS30 APG7/CVT2

Hydrophilic, cytosolic Hydrophilic, membrane associated Peripheral membrane Soluble, some punctate staining in SD-N

APG9/AUT9/CVT7 APG10 APG12 APG13 APG14/CVT12 APG16 AUT1 AUT2/APG4 AUT7/APG8/CVT5 CSC1/END13/VPS4 TOR2 VAC8 VAM3 VPS18

Integral membrane with 6–8 TMD — Hydrophilic, membrane associated Hydrophilic Peripheral membrane Hydrophilic Hydrophilic Cytosolic Cytosolic Prevacuolar compartment Phosphatidylinositol kinase homolog Vacuole peripheral membrane Vacuole membrane Hydrophilic, membrane associated in complex Golgi and prevacuole membranes Cyclical membrane association

SER/THR protein kinase, overexpression suppresses apg13 Conjugated by Apg12p, binds Apg16p In complex with Apg14p Homology to ubiquitin-activating enzymes (E1), required for Apg12p-Apg5p conjugation — Required for Apg12p-Apg5p conjugation Conjugated to Apg5p, phosphorylated Phosphorylated, suppressed by Apg1p In complex with Apg6p Binds Apg5p — Interacts with Tub1p, Tub2p, and Aut7p in vitro Homolog of rat microtubule-associated protein LC3, interacts with Aut2p in vitro Constitutive autophagy defect, ATPase activity needed for induction of autophagy Negative regulator of autophagy — t-SNARE required for fusion of cytosolic vesicles with vacuole RING finger zinc-binding domain protein required for fusion of cytosolic vesicles with vacuole v-SNARE Rab GTP-binding protein required for fusion of cytosolic vesicles with vacuole

VTI1 YPT7

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linked via an internal lysine residue to the C-terminal glycine residue of another autophagy protein, Apg12p (Kametaka et al., 1996; Mizushima et al., 1998a). Apg12p is a 21-kDa hydrophilic protein. Both the Apg5p monomer and the Apg5p/Apg12p conjugate are peripherally associated with cellular membranes (George et al., 1999; Mizushima et al., 1998a). Apg16p is another hydrophilic protein that interacts with Apg5p preferentially in the conjugated form (Mizushima et al., 1998a). The conjugated Apg5p/ Apg12p/Apg16p product is required for both autophagy and Cvt transport. In both the apg7/cvt2 and apg10 mutant strains the Apg5p/Apg12p conjugate does not form, suggesting that the proteins encoded by these genes are required for the conjugation reaction. Cloning of APG7 revealed that it contains a domain homologous to the E1 class of ubiquitin-activating enzymes. Interestingly, Apg12p shares no homology to ubiquitin, and, unlike ubiquitin, it appears to be conjugated to just one protein, Apg5p. Human homologs of Apg5p and Apg12p have been identified (Mizushima et al., 1998b). The human proteins are also linked by an isopeptide bond, suggesting that the novel protein conjugation reaction identified in yeast also acts in mammalian cells. Examination of the mRNA levels of APG12 in different tissues including heart, brain, placenta, lung, liver, skeletal muscle, kidney, and pancreas reveal that it is ubiquitously expressed (Mizushima et al., 1998b). This finding indicates that autophagy is carried out in many different cell types in mammals. Analysis of Apg7p indicates that it is required for Apg5p-Apg12p conjugation, and, like Apg5p, acts at the vesicle formation step (Kim et al., 1999). Apg7p is homologous to the ubiquitin activating enzyme Uba1p over an 85 amino acid region that contains the active cysteine required for ubiquitin activation and a conserved ATP binding domain. The rest of the protein shows no homology to ubiquitin activating enzymes. Mutation of the residues of Apg7p that correspond to the active site cysteine and the ATPase domain of Uba1p prevent Apg5p-Apg12p conjugation (Tanida et al., 1999). Physical interaction of Apg7p and Apg12p has been observed. Localization experiments using GFP-tagged Apg7p show Apg12p-dependent membrane localization when cells are incubated in nitrogen-poor media (conditions that induce autophagy; Kim et al., 1999). Further, these two proteins have been shown to be associated by both two-hybrid analysis and coimmunoprecipitation experiments (Tanida et al., 1999). Together, these data suggest that Apg7p is an Apg12p activating enzyme that is necessary for the formation of the Apg5p-Apg12p conjugate. Apg1p/Aut3p/Cvt10p is a 102-kDa phosphoprotein that contains a conserved Ser/Thr protein kinase domain (Matsuura et al., 1997; Schlumpberger et al., 1997). Characterization of this protein indicates that it has autophosphorylation activity in vitro, and that phosphorylation in vivo decreases in nitrogen starvation conditions (Matsuura et al., 1997). Additional substrates

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of this kinase are not known, but apg1 mutants are tightly blocked for both autophagy and Cvt transport. Overexpression of Apg1p suppresses the defects in the apg13 mutant strain, suggesting that Apg1p and Apg13p interact (Funakoshi et al., 1997). The gene encoding Apg13p has been cloned and is predicted to be 83 kDa. It has no homologs in the database (Funakoshi et al., 1997). Once autophagosomes are formed in the cytosol, they are delivered to the vacuole/lysosome. Based on experiments in mammalian cells using the microtubule depolymerizing drug nocodazole (Aplin et al., 1992) and the microtubule inhibitor vinblastine (Seglen et al., 1996), microtubules are predicted to be involved in the rapid delivery of vesicles to the vacuole/ lysosome. The yeast proteins Aut2p and Aut7p may be involved in this process (Lang et al., 1998). The gene encoding the aut2 mutant has been cloned and is predicted to encode a 56-kDa protein. Several homologs were identified in the database, but none of them has a known function. Aut7p was identified in a screen for suppressors of aut2. The aut7 mutant is allelic to previously identified mutants cvt5 and apg8. The protein encoded by AUT7/CVT5/APG8 is required for both Cvt transport and autophagy. Aut7p is a 14-kDa protein with homology to a rat-microtubule-associated protein. In vitro binding experiments indicate that Aut2p can bind to the yeast tubulins Tub1p and Tub2p. By two-hybrid analysis, Aut2p was found to interact with both Aut7p and the yeast tubulins. Electron microscopy studies identified vesicles in both aut2⌬ cells and aut7⌬ cells that may be autophagosomes (Lang et al., 1998), although their size is smaller than that reported previously for these compartments (Baba et al., 1994). Based on these findings, Aut2p and Aut7p are proposed to aid in the association of autophagosomes with microtubules that can then direct them to the vacuole surface. However, because fractionation experiments performed in both the cvt5 and apg8 mutants failed to identify vesicles containing API precursor in rich media conditions (Harding et al., 1996; Scott et al., 1996), it is not clear whether these proteins act in this manner in the Cvt pathway. Two additional proteins that appear to be involved in the regulation of autophagy have been identified. The first is the protein kinase Tor. In S. cerevisiae, Tor is encoded by the TOR1 and TOR2 genes. Tor is negatively regulated by the drug rapamycin, thus preventing phosphorylation of its substrates. In growing cultures, the addition of rapamycin induces autophagy (Noda and Ohsumi, 1998), suggesting that phosphorylation by Tor is involved in switching between the autophagy and Cvt pathways. A second component in this pathway, Csc1p, was identified in a screen for constitutive autophagy mutants (Shirahama et al., 1997). Cloning of csc1 revealed that it is allelic to END13/VPS4/GRD13. This protein is a member of the AAA family of ATPases and appears to play an important role both in maintenance of the structure of the endosome and in protein sorting through this

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organelle; cells expressing mutant forms of this protein contain an enlarged version of the endosome called the class E compartment (Babst et al., 1998; Munn and Riezman, 1994; Nothwehr et al., 1996). Morphological examination of mutants accumulating class E structures indicates that cuplike multilamellar structures are formed (Rieder and Emr, 1997). These studies make the endosome a compelling candidate for the donor membrane of autophagosomes in yeast. The macromolecular components involved in vesicle docking and fusion have been examined in other systems such as homotypic vacuole fusion. Vesicle fusion appears to be governed by a conserved set of proteins in these previously examined cases. N-ethylmaleimide-sensitive factor (NSF) and its soluble receptor SNAP are two such factors. These proteins modulate the activity of SNAREs. Vesicles targeted for a particular organelle contain a v-SNARE on their surface. This molecule recognizes a t-SNARE on the target organelle. Binding of these two molecules brings the two apposing membranes in close proximity to each other, and fusion occurs. SNAREs are not solely responsible for directing vesicles to fuse at the correct target organelle. One v-SNARE, Vti1p, has been found to be involved in at least three different subcellular targeting reactions (Fischer von Mollard and Stevens, 1999). In addition to these factors, a Sec1 protein family homolog, a member of the rab family of small GTPases, as well as proteins that regulate the GTP-binding state and membrane attachment of rabs are required. Molecular components required for the fusion step of the autophagy and Cvt targeting pathways accumulate cytosolic autophagosomes that can be identified by electron microscopy (Baba et al., 1994). They also contain precursor API inside cytosolic vesicles where it can be identified biochemically (Scott et al., 1997). A mutant in vacuolar protein sorting, vps18, has been found to be required for vacuole fusion of both Cvt vesicles (Scott et al., 1997) and autophagosomes (Rieder and Emr, 1997). Vps18p is part of a protein complex that contains Vps11p, Vps16p, and Vps33p. These proteins are important for formation or maintenance of the structure of the vacuole because cells bearing mutant forms of these four proteins lack a definable vacuole (class C mutants). A temperature-sensitive allele of vps18 causes the accumulation of vesicles containing prAPI immediately after the switch to a nonpermissive temperature, suggesting a direct role for Vps18p in the fusion of Cvt vesicles with the vacuole (Scott et al., 1997). Similarly, autophagosomes have been identified in a different temperature sensitive allele of vps18 (Rieder and Emr, 1997). In addition, other members of the class C family have been found to be involved in autophagy. Electron microscopy studies indicate that vps33 accumulates autophagosomes (Baba et al., 1994), and a mutant in API targeting, cvt15, was found to be allelic to vps16 (A. M. Fischer, S. V. Scott and D. J. Klionsky, unpublished data).

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Vps18p and Vps11p have been shown to bind to each other via RING finger motifs as part of a larger complex that also contains Vps16p and Vps11p (Rieder and Emr, 1997). This complex is thought to be localized on the vacuole membrane. Together, these studies suggest the involvement of the class C protein complex in autophagosome/Cvt vesicle fusion at the vacuole. The role of these proteins in either autophagy, Cvt transport, or vacuole protein sorting via the Vps pathway has not been determined. The t-SNARE Vam3p is localized to the vacuole and is required for homotypic vacuole fusion (Nichols et al., 1997; Ungermann et al., 1998). Studies utilizing a temperature-sensitive allele of this gene indicate that it accumulates both autophagosomes and Cvt vesicles, suggesting that it is the t-SNARE required for these pathways (Darsow et al., 1997). Overproduction of Vam3p suppresses sorting defects of a vps18 temperaturesensitive mutant, suggesting an interaction between these proteins. The v-SNARE Vti1p is also required for prAPI delivery (Fischer von Mollard and Stevens, 1999) and has been shown to interact with Vam3p, suggesting that it is present on Cvt vesicles and most likely on autophagosomes and may mediate their targeting/recognition at the vacuole surface through interaction with Vam3p. SNARE activity is regulated by members of a family of small GTPases called rabs. The rab protein Ypt7p is required for homotypic vacuole fusion (Haas et al., 1995). Biochemical analysis of precursor API localization in a ypt7 deletion mutant indicates that this protein is also required for fusion of Cvt vesicles and autophagosomes at the vacuole (Kim et al., 1999; George et al., 1999; S. V. Scott and D. J. Klionsky, unpublished data). Once released into the vacuole lumen, autophagic bodies and Cvt bodies are broken down allowing for the maturation of precursor API and the degradation of the cytosolic contents of the vesicles. In cells defective in proteinase A or proteinase B, degradation is blocked, and autophagic and Cvt bodies accumulate (Takeshige et al., 1992). Because proteinase A is required for activation of proteinase B and mutations in proteinase B alone block degradation, the activity of proteinase B is probably required for vesicle breakdown. In addition, vesicles also accumulate when cells are incubated in the presence of the proteinase B inhibitor PMSF, suggesting that proteinase B activity is directly requried for this process. Additional factors such as lipases that would likely be required for complete elimination of subvacuolar vesicles have not been identified. Several additional proteins required for autophagy have been identified, but their functional role in this process is not yet clear. Apg14p is a novel hydrophilic protein predicted to be 40 kDa. Overexpression of this protein partially suppresses the autophagy defect in the apg6-1 allele (Kametaka et al., 1996). Cloning apg6 revealed that it is allelic to the vacuole protein sorting mutant vps30. Overexpression of APG14 did not, however, over-

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come the vacuole protein sorting defect in vps30 or in the apg6-1 allele, and apg14 itself is not defective in vacuole protein sorting. Both Apg14p and Apg6p/Vps30p are membrane associated, and the two proteins can be coimmunoprecipitated, suggesting that these two proteins physically interact. Apparently, there are two distinct roles for Apg6p/Vps30p, one in vacuole protein sorting and the other in a multimeric protein complex with Apg14p in autophagy. Aut1p is predicted to be a 36-kDa hydrophilic protein. The role of this protein is not known, but it contains clusters of charged residues including KEKE motifs that may indicate that it assembles into a larger protein complex (Schlumpberger et al., 1997). 2. Physiological Role of Autophagy Examination of the general morphological properties as well as the more recent identification of required molecular components indicates that the principles of macroautophagy are conserved from yeast to mammals. This conservation across kingdoms suggests that this process is of critical importance. In yeast, autophagy is essential for survival during long periods of starvation (Tsukada and Ohsumi, 1993) and is induced by depletion of nutrients including nitrogen, carbon, and sulfur (Takeshige et al., 1992). In mammals, hepatocytes respond to diminishing nutrients by activating autophagy, presumably to enable the maintenance of constant pools of metabolites. Autophagy has also been implicated in many cellular remodeling processes including uterine changes during the pregnancy and postpartum periods, differentiation, and aging (Luzikov, 1999). Recent studies indicate that autophagy plays a role in apoptosis. The apoptosis specific protein (ASP1) was recently cloned and is a homolog of the yeast autophagy protein Apg5p (Hammond et al., 1998). Plant cells respond to sucrose starvation by inducing autophagy (Aubert et al., 1996; Moriyasu and Ohsumi, 1996), and autophagy is probably involved in cellular remodeling processes that take place during germination and senescence.

B. The KFERQ-Mediated Pathway Is Also a Starvation-Response Pathway In addition to macroautophagy, mammalian cells possess a second mechanism for lysosomal delivery of cytosolic proteins in response to starvation. This delivery mechanism is selective for proteins containing a KFERQlike motif. Thirty percent of cytosolic proteins contain this motif and are thus putative substrates for this targeting pathway (Dice, 1990). Experimentally, a number of cytosolic proteins including RNase A, GAPDH, aldolase, some subunits of the 20S proteasome, and the intracellular inhibitor of the

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nuclear factor ␬B have been shown to be specifically targeted to lysosomes (Cuervo and Dice, 1998; Dice; 1990). These proteins are directly transported into lysosomes after prolonged starvation in mammals or in response to serum deprivation in cultured cells (Cuervo et al., 1995). The transport process is facilitated by ATP and the constitutive form of cytosolic HSC73. KFERQ-containing proteins are first recognized by cytosolic HSC73 (Hayes and Dice, 1996). They then bind the lysosomal membrane protein LGP96, which acts as a receptor for the KFERQ motif (Cuervo and Dice, 1996). The action of a lysosomal form of HSC73 is then thought to power protein translocation into the lumen where the proteins are degraded by resident proteases (Agarraberes et al., 1997). Studies performed in fasting rat liver lysosomes provide some explanation of the relative cellular roles of macroautophagy and the KFERQ-mediated pathways in starvation (Cuervo et al., 1995). On initiation of starvation, macroautophagy proceeds first and typically results in the nonselective degradation of 30–40% of cytoplasmic proteins in the first 24 hr. If starvation persists, the KFERQ-mediated pathway is induced to degrade the polypeptides specifically containing KFERQ-like sequences nearly completely. By this combination of degradation pathways, maximal protein turnover is ensured while avoiding total destruction of essential enzymes. C. A Specific Targeting Pathway for Delivery of Degradative Substrates to the Yeast Vacuole It is not known if yeast contains a protein translocation pathway analogous to the KFERQ pathway in mammalian cells, but S. cerevisiae does specifically target some cytosolic enzymes to the vacuole for degradation. The gluconeogenic enzyme FBPase is induced when cells are grown on poor carbon sources, and is rapidly inactivated and then degraded when glucose is added to the media. This degradation can be either by the proteasome or by vacuolar hydrolases depending on conditions (Chiang and Schekman, 1994; Schork et al., 1994; reviewed in Scott and Klionsky, 1998; Klionsky, 1997). Mutants in the vacuolar import and degradation pathway (vid ) have been isolated (Hoffman and Chiang, 1996). Biochemical and morphological examination of vid mutants indicates that FBPase is imported into 30-nm vesicles, which then fuse with vacuoles (Chiang and Chiang, 1998; Huang and Chiang, 1997). A second set of mutants in glucose-induced degradation ( gid; Hammerle et al., 1998) may provide further insight into the relationship between the two pathways of FBPase degradation. D. Ubiquitin-Mediated Endocytosis In addition to being a major site for turnover of cytosolic proteins, proteins from the cell surface are also delivered to the lysosome/vacuole for degrada-

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tion. This process is mediated by endocytosis. Extracellular and plasma membrane proteins are packaged into vesicles which then fuse with endosomes and their contents are carried on to the vacuole where they are degraded (Bryant and Stevens, 1998; Geli and Riezman, 1998; Schmid et al., 1998).

1. Degradation of Plasma Membrane Proteins Endocytosis is also utilized for regulated degradation of specific cell surface receptors. In S. cerevisiae, this process has been studied using the G-protein coupled receptor Ste2p, which is expressed on MATa type cells and binds 움-factor. Binding of 움-factor to Ste2p triggers internalization of the receptor ligand complex. Rather than recycling, Ste2p is delivered to the vacuole where it is degraded (Hicke, 1997). The signal SINNDAKSS within the cytosolic domain of Ste2p is sufficient for this internalization process. In studies using a truncated form of Ste2p, binding of 움-factor triggered phosphorylation of the SINNDAKSS serines (Hicke et al., 1998). The phosphorylated form of Ste2p is then modified by conjugation of the 76-amino acid protein ubiquitin at the SINNDAKSS lysine and internalized. Studies using mutants in the ubiquitin conjugation pathway demonstrated that ubiquitination is required for efficient internalization (Hicke and Riezman, 1996).

2. Coordination between Degradation Pathways Degradation of ubiquitinated proteins by the cytosolic proteasome has been well documented (reviewed in Hochstrasser, 1996). The additional function of ubiquitin as a signal for down-regulation of surface receptor proteins requires accurate differentiation between proteins intended for degradation by the proteasome and the vacuole/lysosome. Misdirection of surface receptors to the proteasome could result in degradation of just their cytosolic domains, eliminating the ability for regulation. Proteasome recognition is mediated by polyubiquitin chains formed by conjugation of ubiquitin through internal lysine residues. In contrast, vacuole delivery of Ste2p requires the addition of just a single ubiquitin residue. Mutations that eliminate the internal lysine residues in ubiquitin or the enzyme required for formation of higher-order ubiquitin conjugates do not prevent vacuole delivery (Terrell et al., 1998). Studies of the uracil permease in yeast indicate that monoubiquitination in not the entire story. This protein requires lysine residue 63 of ubiquitin for efficient internalization and may utilize a diubiquitin chain as a vacuolar targeting signal (Galan and HaguenauerTsapis, 1997). These studies suggest that in contrast to recognition by the

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proteasome, which requires polyubiquitination, mono or small ubiquitin chain modifications direct plasma membrane proteins to the vacuole. Modification by ubiquitin is a major determinant of the life span of plasma membrane proteins. In S. cerevisiae, the majority of plasma membrane proteins known to be internalized appear to be ubiquitin modified (Hicke, 1997). Ubiquitination of Ste2p increases its degradation rate approximately tenfold (Hicke and Riezman, 1996). In addition, a large number of plasma membrane receptors in mammalian cells are ubiquitinated. There is evidence that the growth hormone receptor is targeted for degradation by ubiquitination (Strous et al., 1996), suggesting that ubiquitin-mediated degradation of plasma membrane proteins is conserved in mammalian cells.

E. Peroxisomes Are Delivered to the Vacuole by Autophagic Processes In order to sustain optimal growth while conserving nutrients, organelles must be turned over when they are no longer needed. This can allow for their macromolecular components to be recycled and reused. Of the organelle degradation processes, peroxisomal degradation has been the best characterized. When cells are grown in conditions where peroxisomal enzymes are not in high demand such as glucose, only a few peroxisomes per cell are present. In methylotrophic yeasts, peroxisomes proliferate when cells are grown using methanol as a carbon source. Under these conditions, peroxisomes can occupy as much as 50% of the total cell volume. Similarly, peroxisome number is increased in the nonmethylotrophic yeast, S. cerevisiae, when cells are grown on poor carbon sources such as oleic acid. Upon introduction of glucose, peroxisomes are no longer required in high numbers, and they are rapidly and specifically degraded. The destruction of unnecessary organelles is a vacuolar process and is dependent on the vacuolar hydrolases proteinase A and proteinase B (Tuttle and Dunn, 1995). 1. Two Pathways for Peroxisomal Degradation in Pichia pastoris Examination of peroxisome degradation in the methylotrophic yeast Pichia pastoris revealed that the mechanism utilized for degradation depends on the carbon source. When methanol-grown cells are transferred to ethanol, cytosolic membranes surround peroxisomes sequestering them by a processes analogous to macroautophagy called macropexophagy (Klionsky, 1997, 1998; Sakai et al., 1998; Tuttle and Dunn, 1995). In contrast, when methanol-adapted cells are instead transferred to glucose, vacuole protru-

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sions surround clumps of peroxisomes in a process called micropexophagy (Sakai et al., 1998; Tuttle and Dunn, 1995). These two pathways of degradation can be distinguished by their sensitivity to the protein synthesis inhibitor cyclohexamide. Whereas glucose-stimulated micropexophagy is blocked by this inhibitor, ethanol stimulated macropexophagy proceeds normally (Sakai et al., 1998; Tuttle and Dunn, 1995), suggesting that these pathways are biochemically different. Two sets of mutants in pexophagy called gsa (Tuttle and Dunn, 1995) and pag (Sakai et al., 1998) in P. pastoris have been isolated. Some of these mutants only affect glucose-stimulated peroxisome degradation, confirming that the two modes of pexophagy differ at the molecular level. Cloning of gsa1 revealed that this gene encodes the metabolic enzyme phosphofructokinase 1 (PFK; Yuan et al., 1997). This enzyme is required for initiation of glucose-induced micropexophagy. Interestingly, the catalytic domain of PFK does not appear to be involved because a mutation eliminating catalytic activity does not affect peroxisome degradation. A second gsa mutant, gsa7, was also cloned recently. Gsa7p contains regions of homology to ubiquitin E1 activating enzymes and is homologous to Apg7p in S. cerevisiae (Yuan et al., 1999). A human homolog of GSA7/APG7 has also been cloned (Yuan et al., 1999). Introduction of S. cerevisiae APG7 complements gsa7 cells in P. pastoris (Yuan et al., 1999) and in the corresponding experiment, introduction of P. pastoris GSA7 into apg7 cells partially complements both the autophagy and Cvt defects caused by this mutation (Kim et al., 1999). This finding suggests that GSA7 and APG7 are functional homologs, and that some of the same genes are involved in the selective degradation of peroxisomes and the nonspecific uptake of cytosolic components. Construction of point mutants in P. pastoris GSA7 revealed that the putative ATP binding domain and catalytic site of E1 activating enzymes are required for pexophagy (Yuan et al., 1999). When gsa7 and gsa7⌬ were examined by electron microscopy under glucose-stimulated pexophagy conditions, these cells contained clusters of peroxisomes partially surrounded by protrusions of the vacuole (Yuan et al., 1999). It appears from these images that Gsa7p is involved in a late step in peroxisome sequestration, possibly in the homotypic fusion of vacuole membrane that would be required to engulf peroxisomes fully. It is not known whether Apg7p plays a similar role in the formation of autophagosomes. In addition to the gsa mutants, six pag mutants defective in pexophagy in P. pastoris have been identified (Sakai et al., 1998). Of these, pag1, pag2, pag3, and pag6 were defective only in glucose-induced pexophagy (the micropexophagy pathway), whereas pag4 and pag5 were defective in both ethanol (macropexophagy) and glucose-stimulated pexophagy. These mutants were further characterized by determination of the step of the micropexophagy pathway that was disrupted. The mutants pag1 and pag2 are

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blocked at a very early step, before any peroxisome sequestration has taken place. The products of these mutations are likely involved in the signaling pathway that is necessary for initiating vacuole membrane protrusions around peroxisomes. In pag3 cells, the vacuole membrane distends and begins to surround peroxisomes, but intermediate structures accumulate that are not fully enclosed, suggesting that homotypic fusion of the vacuole membrane is blocked. Cells bearing the pag4, pag5, and pag6 mutations are defective in the breakdown step as they accumulate membrane-enclosed peroxisomes inside the vacuole. It is not surprising that the two mutants that block both macropexophagy and micropexophagy appear to act at the degradation step of the process because both delivery mechanisms result in subvacuolar vesicles containing peroxisomes. Further, degradation of vesicles resulting from both macropexophagy and micropexophagy is known to require proteinase A and proteinase B (Tuttle and Dunn, 1995). 2. Pexophagy in Hansenula polymorpha Involves a Macroautophagic Mechanism In contrast to P. pastoris, in Hansenula polymorpha peroxisomes are degraded by macropexophagy in either glucose- or ethanol-containing media. Five complementation groups of mutants in pexophagy have been identified in H. polymorpha ( pdd; Scott and Klionsky, 1998; Titorenko et al., 1995). These mutants define two different steps in the macropexophagy pathway. The pdd1 mutant is blocked in the sequestration step, and the pdd2 mutant appears to be defective in fusion of vesicles containing peroxisomes at the vacuole surface (Titorenko et al., 1995). Recent cloning of pdd1 revealed that it is homologous to VPS34 in S. cerevisiae (Kiel et al., 1999). Vps34p is a phosphatidylinositol-3 kinase that is required for formation of vesicles at the trans-Golgi that carry biosynthetic cargo proteins destined for the vacuole to the prevacuolar compartment (PVC). The role of this protein in pexophagy is not known. 3. Peroxisome Degradation in Saccharomyces cerevisiae Degradation of peroxisomes by pexophagy also occurs in S. cerevisiae. Although S. cerevisiae cannot grow with methanol as the sole carbon source, peroxisomes are induced by growth on oleic acid. Upon shifting to glucosecontaining media, peroxisomes are degraded with a half time of 1.5–5 hours (Chiang et al., 1996; Hutchins et al., 1999). Under these conditions, degradation of peroxisomes is specific and results in the turnover of the majority of peroxisomal material (Hutchins et al., 1999). In contrast, the nonspecific degradation of cytosolic enzymes by macroautophagy plateaus at approximately 30% (Scott et al., 1996). Peroxisomes are degraded in the

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vacuole. In cells containing mutations in vacuolar proteases, degradation of peroxisomal enzymes does not occur (Hutchins et al., 1999), and membraneenclosed peroxisomes accumulate in the vacuole lumen (Chiang et al., 1996; Hutchins et al., 1999). In S. cerevisiae, it is not yet clear whether pexophagy proceeds by micro- or macromechanisms, or by both as occurs in P. pastoris. Examination of pexophagy in cvt and apg mutants revealed that essentially all cvt and apg strains are defective in this process (Hutchins et al., 1999). This finding suggests that although degradation of peroxisomes is a specific process, it requires the bulk of the machinery required for the nonspecific degradation of cytosolic components by macroautophagy.

F. Mitochondria Are Also Delivered to the Vacuole by Autophagosomes Regulation of the number of mitochondria per cell is an important physiological process. Mitochondrial number increases when the concentration of respiratory substrates is high. Due to the energetic costs of maintaining a functional mitochondrion, it would be advantageous to degrade these organelles when they are no longer required. In plant cells, an increase in autophagic degradation and a corresponding decrease in the number of mitochondria per cell has been observed when low levels of substrates for mitochondrial respiration are present (Aubert et al., 1996; Journet et al., 1986). In both mammalian and S. cerevisiae cells subjected to starvation, mitochondria have been observed together with bulk cytosol inside autophagic vesicles (Baba et al., 1994; Dunn, 1990b). Together, these studies indicate that, along with degradation of cytosolic enzymes, the turnover of mitochondria contributes to the ability of cells to survive periods of starvation. In addition, because damaged mitochondria can cause oxidative damage to other cellular components, selective destruction of defective mitochondria would provide an evolutionary advantage. The mitochondria permeability transition (MPT) is a condition that causes molecules up to 1500 Da to permeate the mitochondrial inner membrane through the opening of an inner membrane channel. Fluorescent studies in hepatocytes indicate that mitochondria undergoing MPT are delivered to the lysosome for degradation by macroautophagy (Lemasters et al., 1998b). In fact, membrane depolymerization may provide the signal for sequestration of mitochondria by autophagosomes both in starvation and in cellular conditions that cause mitochondrial damage (Lemasters et al., 1998a). In this manner, macroautophagy can effectively protect cells from oxidative damage due to loss of mitochondrial membrane integrity.

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IV. Nondegradative Pathways A. Cvt Pathway 1. Aminopeptidase I Is Localized to the Vacuole Independent of the Early Secretory Pathway The biosynthesis of CPY established the paradigm for the transit of vacuolar proteins through a portion of the secretory pathway. PreproCPY, translocates into the endoplasmic reticulum where the signal sequence is removed and the protein is core glycosylated. As this p1 form transits through the Golgi complex, the carbohydrate residues undergo additional modification, generating the higher molecular weight p2 form. Both precursor species contain a propeptide that keeps the enzyme inactive. Following transport to the endosome and subsequent delivery to the vacuole, the propeptide is removed to produce the mature, active form of the hydrolase. This route to the vacuole, termed the CPY pathway (Fig. 2A), is utilized by various hydrolases including at least proteinase A and proteinase B, and depends on various Sec components (Klionsky et al., 1990). Initial studies of aminopeptidase I (API) indicated that it was a vacuolar glycoprotein (Metz and Rohm, 1976). This result suggested that API transited to the vacuole through a portion of the secretory pathway, similar to all the previously characterized vacuolar hydrolases. However, once the structural gene for API, LAP4 now called APE1, was cloned, it became apparent that the protein lacked a standard signal sequence or consensus signal sequence cleavage site (Chang and Smith, 1989; Cueva et al., 1989). Furthermore, the protein did not contain any predicted membrane domains that could function as internal uncleaved signal sequences, making it unclear as to its mechanism of translocation into the ER. Subsequent analyses of the biosynthesis of API revealed that the protein was initially synthesized as a 61-kDa precursor (prAPI) that was processed to a 50-kDa mature form with a half-time of 30–40 min (Klionsky et al., 1992). Treatment with the N-linked glycosylation inhibitor tunicamycin did not alter the molecular mass of prAPI. Similarly, the precursor protein failed to bind the mannose lectin concanavalin A even though the protein has potential N-linked glycosylation sites. Finally, the in vitro synthesized product migrates at the same position as the in vivo precursor confirming that the protein is not glycosylated. These data suggested that prAPI was delivered to the vacuole without first translocating into the ER. One other vacuolar hydrolase, 움-mannosidase or Ams1p, had been shown to enter the vacuole bypassing the secretory pathway (Yoshihisa and Anraku, 1990). Ams1p was found to localize to the vacuole and be processed

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to a lower molecular mass form in strains with sec mutations that block translocation into the ER or the formation/fusion of COPII vesicles that transit to the Golgi complex (Yoshihisa and Anraku, 1990). Finally, vacuolar proteins that transit through the secretory pathway are partially secreted when they are overproduced, due to saturation of a limiting component in the TGN. In contrast, Ams1p was not secreted upon overproduction. Similar results were seen with prAPI (Klionsky et al., 1992). Processing of the precursor protein was shown to occur in sec mutants that are blocked in early stages of the secretory pathway, and prAPI was not secreted upon overproduction. In addition, prAPI was not secreted to the cell surface in vps mutants that cause the missorting and secretion of vacuolar proteins that transit through a portion of the secretory pathway. Taken together, these data indicate that prAPI does not directly rely upon the early organelles of the secretory pathway for localization to the vacuole. 2. Biochemical Studies Suggest a Vesicle-Mediated Mechanism for prAPI Delivery The initial studies of API suggested that the mature protein was present in the vacuole as a dodecamer with a corresponding molecular mass of approximately 600 kDa (Lo¨ffler and Ro¨hm, 1979; Metz et al., 1977). To gain further insight into the delivery mechanism, the kinetics of prAPI oligomerization were examined through pulse/chase analyses and glycerol gradient centrifugation (Kim et al., 1997). Precursor API rapidly oligomerizes in the cytosol with a half-time of 2–3 minutes, suggesting that oligomerization is not rate-limiting in the import process. An altered form of prAPI containing a mutation of arginine to lysine at position 12 of the propeptide, K12R, accumulates the precursor form at a

FIG. 2 Nondegradative protein delivery pathways. (A) Several hydrolases utilize the CPY pathway where proteins translocate across the ER membrane and transit through the Golgi complex to the trans-Golgi Network. At the TGN, these proteins are diverted away from the secretory pathway and are transported to a prevacuolar compartment or endosome, followed by delivery to the vacuole. It is not known whether movement from the endosome to the vacuole occurs by direct fusion or via vesicular intermediates. (B) The Cvt pathway is used for the localization of aminopeptidase I. This route is relatively independent of the SEC gene products. Cytosolic precursor API is sequestered within a double-membrane vesicle, which fuses with the vacuole. (C) The vacuolar hydrolase alkaline phosphatase, as well as certain other vacuolar proteins, transits through the secretory pathway to the trans-Golgi Network. These proteins are then delivered directly to the vacuole through the ALP pathway, bypassing the endosome. (D) Carboxypeptidase S transits to the endosome where it enters invaginations of the endosomal membrane. The resulting multivesicular body fuses with the vacuole to release the protein into the lumen via the MVB sorting pathway.

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nonpermissive temperature (Oda et al., 1996). The K12R prAPI was shown to be in an oligomeric form, and the mutation was thermally reversible (Kim et al., 1997). The K12R mutation was used to accumulate a synchronized pool of precursor protein at the nonpermissive temperature that was competent for subsequent vacuolar delivery upon shift to permissive conditions. A pulse/chase analysis of K12R prAPI indicated that the precursor protein transited to and entered the vacuole in the dodecameric form. Precursor API is approximately 732 kDa in mass, making translocation through a protein channel highly unlikely. Large pore structures are not detectable in the vacuole membrane by electron microscopy, ruling out localization through this type of mechanism. Simultaneous to the analysis of oligomerization, prAPI import was analyzed in vitro (Scott and Klionsky, 1995). Based on treatment with protease and biotinylated cross-linker, precursor API was converted from a proteaseaccessible to an inaccessible form in a reaction that was dependent on energy and temperature. Inhibition of import at 14⬚C provided further evidence against translocation through a protein channel; translocation into the ER, mitochondria, or chloroplast can occur at temperatures as low as 0⬚C as long as unfolding is not rate-limiting. Direct evidence for a vesicle-mediated mechanism was obtained by examining prAPI import in mutant strains of yeast (Scott et al., 1997). The vps18 mutant is defective in fusion of vesicles with the vacuole. Precursor API was shown to be located within a membrane-enclosed compartment that was separate from the vacuole in a vps18 temperature-sensitive mutant. The cvt17 mutant (see following discussion) is blocked in the breakdown of intravacuolar vesicles. Vacuoles prepared from a cvt17 mutant contain prAPI as well as lumenal vacuolar marker proteins. The prAPI in the cvt17 mutant can be separated from vacuole lumenal proteins, however, by differential osmotic lysis using conditions that allow breakage of the vacuole membrane while largely retaining the integrity of smaller vesicles. These data suggest that prAPI is contained within cytosolic vesicles in strains blocked in fusion with the vacuole and within intravacuolar vesicles in mutants that prevent membrane breakdown in the vacuole lumen. 3. Genetic Analysis Reveals an Overlap with Mutants Defective in Autophagy A genetic screen was utilized to identify components of the subcellular machinery that delivers prAPI to the vacuole (Harding et al., 1995; Scott et al., 1996). This screen relied on the observation that wild-type yeast accumulate the mature form of API in the vacuole; API is stable in the vacuole lumen as a resident hydrolase. Mutants defective in localization and/or processing of prAPI would accumulate the precursor form. A series

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of mutants that displayed the expected phenotype was isolated. These mutants were termed cvt, for cytoplasm-to-vacuole targeting defective. With two exceptions, the cvt mutants accumulated prAPI in the cytosol, indicating that these mutants are defective in targeting and not processing of the precursor protein. The cvt1 mutant is in the same complementation group as prb1, the gene-encoding proteinase B, which is the processing enzyme that converts prAPI to the mature form. The only other mutant that accumulated vacuolar prAPI was cvt17 which is defective in the breakdown of intravacuolar vesicles (Scott et al., 1997). With a few exceptions, the cvt mutants do not accumulate the precursor forms of vacuolar hydrolases that transit through the secretory pathway, suggesting that they are specific for this alternate targeting mechanism. Some of the cvt mutants are in the same complementation groups as various vps mutants that were isolated based on defects in the vacuolar localization of carboxypeptidase Y (Bryant and Stevens, 1998). The block in prAPI localization in vps mutants may be an indirect effect. Some vps mutants lack a normal vacuole and may not present a proper target for vesicle fusion. It is also possible that some components needed for prAPI import are delivered to the vacuole through the secretory pathway. For example, delivery of the vacuolar t-SNARE Vam3p through the ALP pathway (see following discussion) is dependent on Vps proteins, and Vam3p is required for prAPI import. The most significant overlap between cvt mutants and other strains known to affect delivery of proteins to the vacuole was seen with two sets of mutants that had been isolated based on defects in autophagy (see Section III.A; Harding et al., 1996; Scott et al., 1996). Detailed information on the autophagic process was available based on extensive electron microscopy analyses (Baba et al., 1995, 1994; Takeshige et al., 1992). These studies allowed for a model of prAPI import that fit the biochemical data (Fig. 2B). Direct testing of this model through immunoelectron microscopy supplied further information about the mechanism of prAPI import through the Cvt pathway.

4. Morphological Analyses Confirm a Mechanistic Overlap between the Cvt Pathway and Macroautophagy Immunoelectron microscopy using antiserum to API revealed that the protein was not randomly dispersed throughout the cytosol. Instead, prAPI was seen to assemble into higher-order Cvt complexes consisting of multiple dodecamers (Baba et al., 1997; Scott et al., 1997). These complexes were seen to be devoid of membrane or partially enwrapped in the cytosol and also within intravacuolar vesicles in strains defective in vesicle breakdown

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such as cvt17 or pep4. In wild-type strains, API was shown to be uniformly dispersed throughout the vacuole lumen. By extending the analysis to include both vegetative and starvation conditions, it was demonstrated that prAPI import utilizes two types of vesicles. Under vegetative conditions, prAPI is detected as a Cvt complex within double-membrane Cvt vesicles, which are 140 to 160 nm in diameter. These vesicles appear to exclude cytosol based on particle distribution. When cells undergo starvation, prAPI is again seen in the form of Cvt complexes but now within autophagosomes. These vesicles are 300–900 nm in diameter and include cytosol along with the Cvt complex, consistent with their role in cytosolic protein degradation. These results suggest that prAPI enters the vacuole through two overlapping pathways that are topologically similar. 5. Functional Roles for the Cvt/Apg Proteins Insight into the molecular basis of the Cvt and autophagy pathways has been gained by analyzing the products of the APG, AUT, and CVT genes along with mutant strains that are defective in prAPI import and/or autophagy (Table I). The model for prAPI import presented in Fig. 2 suggests several specific stages where protein components may be involved in the delivery process. At present, no mutants that are defective in the earliest stages of import, oligomerization of prAPI or assembly of the Cvt complex, have been identified. Perhaps the most complex aspect of prAPI localization concerns the mechanism by which the Cvt complex is sequestered within cytosolic double-membrane vesicles. As is the case in macroautophagy, the origin of the sequestering membrane remains to be determined. The majority of the proteins that are required for prAPI import can tentatively be assigned to the step of enclosure within the cytosolic vesicle. This assignment is based on the presence of protease-sensitive prAPI following differential lysis of spheroplasts, using conditions that retain the integrity of internal compartments (Kim et al., 1999; George et al., 2000). Among the proteins involved in the sequestration event, the Apg5p, 7p, 10p, 12p, and 16p proteins have been shown to participate in the production of a novel protein complex (see Section III.A). The functions of the remaining Aut, Apg, and Cvt proteins are less well defined, although certain proteins have been shown to interact through genetic and/or biochemical experiments (Table I). As discussed earlier, prAPI is sequestered within either Cvt vesicles or autophagosomes, depending on the environmental conditions. The signal transduction mechanism that controls the type of vesicle used for this process is not understood. Four proteins—Apg1p, Apg13p, Csc1p/Vps4p and Tor2p—have been identified; they may be involved in the regulatory process (see Section III.A).

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The Cvt pathway converges with other biosynthetic pathways used for delivery of proteins to the vacuole at the step of vesicle fusion. Targeting of the Cvt vesicle to the vacuole and fusion of the respective membranes appears to involve v- and t-SNAREs, the rab protein Ypt7p (Kim et al., 1999; Scott et al., 1997), the NSF homolog Sec18p and the GDP dissociation inhibitor Sec19p (S.V. Scott and D.J. Klionsky, unpublished results). The t-SNARE Vam3p and the v-SNARE Vti1p may be involved in the final targeting event (see Section III.A), although it has not been demonstrated that prAPI in the vti1 mutant is present in membrane enclosed vesicles.

B. ALP Pathway Initial studies on the biosynthesis of alkaline phosphatase (ALP) revealed that it is synthesized as a type II integral membrane precursor protein (Klionsky and Emr, 1989). It is processed to the mature form by the removal of a carboxyl-terminal propeptide upon delivery to the vacuole. Processing occurs with a half-time of 6 minutes, similar to that seen for CPY. ALP transits to the vacuole through a portion of the secretory pathway. Delivery to the vacuole is dependent on sorting information contained within the cytosolic tail and/or transmembrane domain (Klionsky and Emr, 1990). In contrast to CPY, vacuolar delivery of ALP is not dependent on acidification by the vacuolar ATPase (Klionsky and Emr, 1989; Morano and Klionsky, 1994). Similarly, ALP targeting is relatively unaffected by vps mutants which missort CPY (Klionsky and Emr, 1989). These latter observations suggested that the membrane protein ALP utilized an alternative mechanism for vacuolar localization. 1. Vacuolar Delivery of ALP Occurs in vps Mutants Further analysis of ALP including immunofluorescence studies confirmed that its vacuolar delivery is not blocked in most vps mutants (Cowles et al., 1997b; Piper et al., 1997). In particular, Vps45p is a Sec1p homolog that is needed to move CPY from the Golgi complex to the endosome. Vps27p is involved in recycling from the endosome and in forward transit between the endosome and the vacuole. Vacuolar delivery of ALP is independent of both proteins, suggesting that it bypasses the endosome-dependent CPY pathway. In support of this observation, vacuolar localization of ALP is independent of the endosomal t-SNARE Pep12p (Cowles et al., 1997b). In contrast, sorting of ALP, but not CPY, was defective in a vps41 mutant. This observation suggests that Vps41p acts in an alternative pathway that is needed for delivery of ALP. Delivery of both ALP and CPY is dependent on the vacuolar t-SNARE Vam3p (Piper et al., 1997). Because delivery of

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ALP is independent of the CPY pathway, Vam3p must also be delivered to the vacuole by an alternative pathway. In agreement with this deduction, Vam3p localization was shown to be independent of Vps27p (Piper et al., 1997). The alternative route used for delivery of Vam3p and ALP to the vacuole is called the ALP pathway (Fig. 2C). 2. Genetic Analysis of the ALP Pathway A mutant screen devised to identify components involved in ALP sorting provided evidence that subunits of the AP-3 adaptor complex were involved in the alternative ALP pathway (Cowles et al., 1997a). Similarly, a synthetic lethal screen with the apm3 mutant suggested the involvement of AP-3 in the ALP pathway (Stepp et al., 1997). In contrast to defects in transport of ALP, vacuolar delivery of CPY is not dependent on AP-3. Low levels of ALP maturation in AP-3 mutant cells is dependent on Pep12p (Cowles et al., 1997a) and Vps45p (Stepp et al., 1997), indicating that when the AP3 route is blocked, delivery of ALP occurs through the CPY pathway. In agreement with the observation that Vam3p delivery is independent of Vps27p (Piper et al., 1997), it is blocked in AP-3 mutants (Cowles et al., 1997a). 3. Analysis of the Targeting Signal in ALP Domain exchanges and deletions revealed that ALP contains a vacuolar targeting signal within the cytoplasmic domain (Cowles et al., 1997b; Piper et al., 1997). The observation that either amino acids 1–16 or 22–33 of the ALP cytosolic tail are sufficient to target passenger proteins into the alternative pathway suggests that there may be two redundant targeting signals. To more clearly define the targeting signal in ALP, hybrid proteins were made containing the ALP cytoplasmic domain and the transmembrane and lumenal domains of the Golgi protein guanosine diphosphatase (GDPase; Vowels and Payne, 1998). The ALP targeting signal is able to override the Golgi localization signal present in GDPase, whereas mutations in the vacuolar targeting signal lead to Golgi retention rather than vacuolar localization. The advantage of this system is that it bypasses problems associated with default delivery of membrane proteins to the vacuole (Nothwehr and Stevens, 1994). The data suggest that the first 16 residues of the ALP cytosolic tail, including leucine and valine at positions 13 and 14, are critical for vacuolar targeting of ALP. Delivery by the LV-based sorting signal is clathrin-independent (Vowels and Payne, 1998), in agreement with studies from animal cells showing that AP-3 functions independently of clathrin. Mutation of these two residues in wild-type ALP diverts the protein into the Pep12p-dependent CPY pathway (Vowels and Payne, 1998). Simi-

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larly, a di-leucine motif is part of the signal that is required for targeting of Vam3p to the vacuole through the ALP pathway (Darsow et al., 1998). The presence of an alternate targeting route allows proteins such as Vam3p to bypass the endosome. This may be critical in allowing efficient targeting of proteins to the vacuole as a discrete organelle. That is, the specificity of Vam3p as a vacuolar t-SNARE is in part achieved by its localization to that organelle through a pathway that does not involve the endosome and may prevent premature interaction with its cognate v-SNARE.

C. MVB Sorting Pathway It appears that at least two distinct subsets of the Vps pathway are used for the delivery of resident membrane proteins to the vacuole. DPAP B and membrane components of the vacuolar ATPase transit through the CPY pathway, whereas ALP bypasses the endosome and utilizes the AP-3-dependent ALP pathway. Recently, it has been shown that a third pathway exists for delivery of proteins to the vacuole through a portion of the secretory pathway. Carboxypeptidase S (CPS) is synthesized as a type II integral membrane precursor protein of two different masses that vary due to variable levels of glycosylation (Spormann et al., 1992). Initial studies on the biosynthesis of CPS revealed that it is processed in a proteinase B-dependent manner upon delivery to the vacuole, to yield a soluble mature form of the enzyme. The processing occurs with a half-time of 20 minutes. This rate is substantially slower than that seen with CPY or ALP, suggesting some differences in the transport mechanism. Processing of precursor CPS is not required for activity, however, so it may not be a good indicator of the transport kinetics. CPS can be recovered from purified vacuoles in agreement with its role as a vacuolar hydrolase. However, subcellular fractionation of spheroplasts prepared from a pep4⌬prb1⌬ mutant strain under conditions that cause vacuolar lysis, results in the recovery of CPS in a 100,000 ⫻ g pellet (Odorizzi et al., 1998). In contrast, ALP is located exclusively in a 13,000 ⫻ g pellet, suggesting that the two proteins are not colocalized in the vacuolar membrane. Analysis of a GFP-CPS hybrid protein revealed staining throughout the vacuolar lumen, indicating that the cytosolic domain that is fused to GFP must be located inside the vacuole. Because CPS is made as a type II membrane protein, a lumenal localization of the cytosolic domain argues for an unusual mechanism of transport. An altered form of GFP-CPS containing a targeting domain for the ALP pathway is delivered to the vacuole membrane. This observation suggests that the lumenal distribution of CPS is dependent on transit through the

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endosome. Similarly, class E vps mutants that are defective in endosomal function showed substantial levels of GFP-CPS on the vacuole membrane. Immunoelectron microscopy of a vma4⌬ strain defective in the vacuolar ATPase, revealed GFP-CPS within 40 to 50 nm intravacuolar vesicles. Finally, fab1⌬ mutants which are unable to synthesize phosphatidylinositol3,5-diphosphate (Gary et al., 1998) accumulate GFP-CPS primarily on the vacuole membrane. These results led to a model for CPS transport to the vacuole by a novel mechanism. Precursor CPS transits through the secretory pathway to the Golgi complex and is then diverted to the endosome. The protein is subsequently localized within membrane vesicles that result from invagination of the endosomal membrane. The resulting endosome, or multivesicular body (MVB), fuses with the vacuole, delivering the vesicles into the lumen where they are ultimately degraded (Fig. 2D). A similar distribution within the vacuole lumen to that observed for GFP-CPS was seen with a Ste2pGFP hybrid (see Section III.D for Ste2p degradation), suggesting that turnover of this protein also occurs through the MVB sorting pathway. One advantage of this mechanism is that it allows a physical separation of membrane proteins, such as Ste2p, that are destined for degradation from most of the resident proteins of the vacuole membrane.

D. Folded Protein Pathways In addition to the Cvt pathway described earlier, both eukaryotic and prokaryotic cells utilize other systems for biosynthetic localization of folded proteins (Teter and Klionsky, 1999). For example, peroxisomes have the capacity to import oligomeric complexes (Glover et al., 1994; McNew and Goodman, 1994, 1996). The mechanism of import has not been determined but is proposed to involve either translocation through a transient pore or invagination of the peroxisomal membrane. Folded proteins can also cross the chloroplast thylakoid membrane (Clark and Theg, 1997; Hynds et al., 1998). A similar pathway exists in bacteria with the twin arginine translocation system (Dalbey and Robinson, 1999). As with these other alternative pathways, we do not yet have a biochemical understanding of the components involved in the translocation process.

V. Channel-Mediated Transport The alternative transport pathways we have discussed up to this point have primarily involved vesicle-mediated mechanisms. The degradation of

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KFERQ-containing proteins and FBPase described in Section III may be exceptions in that these proteins appear to translocate across a membrane bilayer. In both cases, however, the molecular details of protein localization into the vacuole or the Vid vesicles, respectively, have not been determined. The various anterograde and retrograde pathways rely on an initial translocation event to allow the targeted protein to cross the bilayer. These translocation components are exemplified by the Sec machinery in the bacterial plasma membrane and ER of eukaryotic cells and have been extensively characterized. In brief, integral membrane proteins provide hydrophilic channels through the hydrophobic bilayer. The channel-mediated translocation of proteins across biological membranes encompasses a diverse range of transport processes. Here we briefly describe some of these alternative pathways and refer to recent reviews addressing each topic.

A. ER Dislocation Signal-mediated translocation of proteins into the ER lumen through the Sec61p translocation complex is a well-characterized process in the biosynthesis of secretory proteins and resident organellar proteins of the endomembrane system (Corsi and Schekman, 1996; Rapoport et al., 1996). To ensure the export of correctly folded proteins, an ER quality control mechanism exists so that proteins that are misfolded, aberrantly modified, or fail to oligomerize properly are retained and degraded before they can escape the ER to more distal compartments (Hammond and Helenius, 1995; Kopito, 1997; Brodsky and McCracken, 1997; Bonifacino and Weissman, 1998). Both misfolded membrane and soluble proteins undergo a rapid, reverse translocation step from the ER lumen, through the translocation channel, and are released into the cytosol as soluble proteins or remain associated with the ER membrane. Chaperone molecules both in the ER lumen (e.g., Cne1p/calnexin or Kar2p/BiP) as well as in the cytosol (e.g., Hsp70 family) are thought to play a role in maintaining the substrate in an unfolded state as well as in providing the push in this ER ‘‘dislocation’’ or retrotranslocation event (Bonifacino and Weissman, 1998). After the substrates are exposed to cytosol, they are deglycosylated by a cytosolic N-glycanase and polyubiquitinated and degraded by the proteasome machinery. A growing number of proteins are being identified as substrates for ER dislocation. Interestingly, mutant forms of several ABC transporters serve as ER dislocation substrates including the cystic fibrosis transmembrane conductance regulator (CFTR). Mutations cause this polytopic membrane protein to misfold, undergo ER dislocation, multi-ubiquitination, and proteasomemediated degradation, thus leading to the human genetic disease cystic fibrosis (Kopito, 1999).

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B. ABC Transporters ABC transporters comprise a superfamily of proteins that couple the energy of ATP hydrolysis to the translocation of substrates ranging from ions, amino acids, sugars, vitamins, complex hydrophobic drugs, phospholipids, peptides, and large proteins across biological membranes (Higgins, 1992). The specificity of ABC transporters can be restricted to a single compound, as in the case of bacterial amino acid permeases, or to a group of related substrates, as in the case of the mammalian multidrug resistance protein which mediates the transport of a wide range of lipophilic compounds. The clinical relevance of ABC transporters in disease states as well as normal human physiology is significant. Defects in human ABC transporter genes have been implicated in a number of genetic diseases including the peroxisomal disorder adrenoleukodystrophy (ALDP1; Dubois-Dalcq et al., 1999), abnormal insulin release in hyperinsulinemic hypoglycemia of infancy (SUR; Aguilar-Bryan and Bryan, 1999) and cystic fibrosis (CFTR; Kopito, 1999). ABC proteins are found in all organisms including prokaryotes, eukaryotes, and archaea (Croop, 1998). All ABC transporters are defined by a common set of structural domains (Schneider and Hunke, 1998). A typical ABC transporter is composed of four parts—two integral membrane domains, each of which spans the membrane six times, and two ATP hydrolyzing domains located on the cytosolic face of the membrane. Whereas the hydrophobic transmembrane domains share little sequence homology, ABC transporters share 30–40% sequence identity in the ATP binding domains including two short sequences (Walker A and B motifs) and a typical ‘‘signature’’ sequence that distinguishes these ATP-binding cassettes from other nucleotide binding proteins (Higgins, 1995; Schneider and Hunke, 1998). In eukaryotes, the four domains of an ABC transporter are generally expressed by a single polypeptide, whereas bacterial ABC transporters are typically assembled from four separate subunits. ABC transporters are integral to a number of cellular processes. The development of multidrug resistance in humans is due to the action of several ABC transporters including P-glycoprotein, which acts as an ATPdependent efflux pump to extrude a broad range of anticancer drugs (Ambudkar et al., 1999). Two well-characterized examples of ABC transporters involved in protein export are the TAP transporters in antigen presentation and the Ste6p transporter for the cell-surface presentation of the yeast mating pheromone, a-factor. The TAP transporter is a heterodimer encoded by the TAP1 and TAP2 genes. Localized to the ER membrane, the TAP complex transports short peptide antigens (8–16 amino acids long) from the cytosol into the lumen of the ER where they can assemble with newly synthesized major histocompatibility complex class I (MHC I) molecules.

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Therefore, TAP function is essential for the subsequent cell surface presentation of MHC I-antigen complex to cytotoxic T lymphocytes (Lehner and Trowsdale, 1998; van Endert, 1999). Ste6p is transiently localized on the plasma membrane of yeast cells and exports the yeast-mating pheromone a-factor by translocation across the plasma membrane. This transport process is termed nonclassical because a-factor export occurs normally in mutants, sec, which are blocked in the secretory pathway (Michaelis, 1993). In addition to its transport function in various cellular processes, there is increasing evidence that some ABC transporters may have a bifunctional role in regulating the activity of other channels and membrane proteins. For example, CFTR has been shown to regulate other ion channel proteins in addition to its intrinsic chloride ion channel activity (Higgins, 1995; Kunzelmann and Schreiber, 1999).

VI. Concluding Remarks Although the alternative protein sorting pathways discussed here have only been recently described, they play key roles in the structure and function of eukaryotic cells. Retrograde transport is now a central tenet of anterograde movement through the endomembrane system. Pathways mediating protein turnover are recognized as an important regulatory process. The alternative vacuolar delivery pathway used by ALP is essential for vacuole biogenesis. Finally, the ABC superfamily of transporters provides a specialized transport system for many important molecules including the peptides that invoke T-cell immune response. Continuing investigation into the molecular mechanisms of these pathways will provide new insight into cellular organization and function.

Acknowledgment The authors would like to thank Greg Rogers for providing Fig. 1.

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Structural Correlates of the Transepithelial Water Transport Ekaterina S. Snigirevskaya and Yan Yu. Komissarchik Institute of Cytology, Russian Academy of Sciences, St. Petersburg, Russia

Transepithelial permeability is one of the fundamental problems in cell biology. Epithelial cell layers protect the organism from its environment and form a selective barrier to the exchange of molecules between the lumen of an organ and an underlying tissue. This chapter discusses some problems and analyzes the participation of intercellular junctions in the paracellular transport of water, migration of intramembrane particles in the apical membrane during its permeability changes for isotonic fluid in cells of leaky epithelia, insertion of water channels into the apical membrane and their cytoplasmic sources in cells of tight epithelia under ADH (antidiuretic hormone)-induced water flows, the osmoregulating function of giant vacuoles in the transcellular fluxes of hypotonic fluid across tight epithelia, and the role of actin filaments and microtubules in the transcellular transport of water across epithelia. KEY WORDS: Epithelia, Transepithelial water transport, Tight junction, Microfilaments, Microtubules, Golgi complex. 䊚 2000 Academic Press.

I. Introduction Epithelial tissues line a cavity or cover a surface of the animal body and have various functions in animals—protective, sensory, and transport. Transporting epithelia participate in such processes as absorption, secretion, and excretion. The general features of epithelial cells are their morphological and functional polarity and the complete absence of the ability to move. The polarity provides the necessary direction for the transport of substance across epithelial cells toward or away from the lumen. Understanding how polarized epithelial cells transport different substances is one of the fundamental problems in cell biology. Physiologists have been studying the transport of different molecules across epithelia since the second half of the International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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nineteenth century. However, many questions are still unanswered, new discoveries continue to be added to this field of investigation. Particularly significant progress in understanding mechanisms and pathways of transepithelial transport has been made in the last 30 years due to a combination of current physiological, morphological, and molecular biological methods. From the great amount of available data about these processes, we have chosen to review only the subject of absorption processes, specifically water absorption. Every cell is bordered by a plasma membrane providing differences in solute concentration between the interior and exterior of the cell. This in turn generates the osmotic water movements across the plasma membrane (Oschman, 1978; Finkelstein, 1984, 1987; Hill, 1995). The experimental data obtained in recent years have confirmed the ideas of some physiologists that osmotic flow across plasma membranes is mediated by specific pore proteins (Finkelstein, 1987; Hill, 1980). A tremendous development in the physiology of the water permeability has been the isolation, cloning, and functional expression of water channel proteins, aquaporins (Preston et al., 1992; Sabolic´ et al., 1992, 1995; Nielsen et al., 1993a, 1993b; Zhang et al., 1993; Abrami et al., 1994, 1995, 1996, 1997; Yamamoto et al., 1995; Granquist et al., 1996; Ma et al., 1996; Siner et al., 1996). Organisms with continuous processes of intake of water and solutes have universal mechanisms of maintaining an osmotic balance and stabilizing cell volume with the aid of Na⫹-K⫹-ATPase. It controls the solute concentration inside the cell by pumping out Na⫹ ions, thereby maintaining a relatively stable cell volume (Cantly, 1981). Simpler fresh-water organisms (Protozoa, Spongia, and some Coelenterata) are protected from hypotonic stress by a specialized osmoregulatory organelle, the contractile vacuole (McKanna, 1976; Patterson, 1980; Zeuhten, 1992). Epithelia of functionally different organs are different in their water and ion permeability (Fro¨mter and Diamond, 1972; Diamond, 1974). Leaky epithelia perform isotonic absorption when solute and water transport are coupled. Tight epithelia have a low basal water and ion permeability regulated by the antidiuretic hormone (ADH), vasopression. In this type of epithelia, there is no coupling between solute and water fluxes. There are some discrepancies in the literature about pathways of fluid transport across different epithelia. Some authors support the idea that the route for the fluid transport includes paracellular spaces, whereas others claim that the predominant part of fluid movement is transcellular. With respect to leaky epithelia, morphological data on the trans- or paracellular water pathways are rather contradictory. Some work demonstrated changes in the ultrastructural tight junction (TJ) organization of small intestine epithelium during hydroplilic substance load (Madara and Pappenheimer, 1987; Atisook and Madara, 1991; Pappenheimer, 1993); others do not ob-

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serve any obvious alterations (Ugolev et al., 1995; Snigirevskaya et al., 1997). In any case, it is clear that the TJ barrier is not absolute and that its permeability is regulated by many cellular signaling mechanisms. Morphological investigations have contributed substantially to our understanding of mechanisms and pathways of water movement across tight epithelia. Thus, much indirect and direct evidence of transcellular routes of fluid movement across this type of epithelia has been obtained. The most significant achievement was visualization of high water permeability domains in the apical membranes of epithelial cells responding to ADH hydroosmotic action (Chevalier et al., 1974). Until now, this fact was the main proof for the transcellular water movement across the tight epithelium. Granular cells are responsible for water transport in the amphibian urinary bladder and are the principal cells in renal collecting ducts (Chevalier et al., 1974; Harmanci et al., 1980; Orci et al., 1981; Wade et al., 1981; Komissarchik et al., 1982). Several different electron microscopic (EM) techniques can be used for the ultrastructural analysis of changes in cell organization after stimulation of transport processes: standard methods of EM (double fixation, embedding in epoxy resins, ultrathin sections); the method of electron-dense tracers such as lanthanum chloride, horseradish peroxidase, ferritin for analysis of the barrier function of tight junction; cryomethods of EM (freezefracture and freeze-substitution); methods of immuno-EM for identification of the chemical nature of cell components and their localization; scanning EM; and electron probe X-ray microanalysis for identification of the elemental composition of the cell. This chapter discusses results of morpho-functional investigations of both the tight and the leaky epithelia on transport stimulation by different substances. Mainly, attention will be paid to changes in the plasma membrane, intercellular junctions, vacuolar system, and cytoskeletal elements during induced water transport.

II. Features of Epithelia As mentioned previously, epithelial tissues are characterized by their structural–functional asymmetry (Koefoed-Johnson and Ussing, 1953, 1958; Simons and Fuller, 1985; Cereijido et al., 1993; Mellman et al., 1993; Rodriguez-Boulan and Zurzolo, 1993). The apical and basolateral asymmetry can determine a vectorial transport of substances across an epithelial layer: from apical surface to basolateral as absorption, and in the opposite direction as secretion (Cerijido et al., 1993). The functional asymmetry can be explained by the polarized distribution of ion, water, and nonelectrolyte

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translocating mechanisms in membranes of epithelial cells (KoefoedJohnson and Ussing, 1958; Cereijido et al., 1989a, 1989b). The apical and basolateral distribution of membrane components has been shown to be provided by the specialized intercellular junctions which restrict the lateral diffusion of proteins and lipids along the membrane (Farquhar and Palade, 1963; Rodriguez-Boulan and Nelson, 1989; Woods and Bryant, 1993; Mitic and Anderson, 1998).

A. Functional Asymmetry of Transporting Epithelia Physiological analysis suggests extensive variations in the ion and water permeability of different epithelia. Features of transepithelial ion transport determine a wide variety of epithelia in functionally different organs (Fro¨mter and Diamond, 1972; Diamond, 1974, 1978, 1979; Ussing et al., 1974; DiBona and Mills, 1979; Lewis et al., 1984; Fro¨mter, 1987). All transporting epithelia can be classified according to their transepithelial electrical resistance in tight and leaky varieties (Fro¨mter and Diamond, 1972; Diamond, 1974, 1978, 1979; Ussing et al., 1974; Erlij and Martinez-Palomo, 1978; DiBona and Mills, 1979; Lewis et al., 1984; Fro¨mter, 1987). Leaky epithelia are characterized by a low resistance, from 5 to 300 ⍀/cm2. Tight epithelia have a high electrical resistance, from 300 to 80,000 ⍀/cm2 (Martinez-Palomo, 1975).

1. Leaky Epithelia Leaky epithelia are epithelia of the gallbladder, small intestine, choroid plexus, and proximal tubules of the kidney. Leaky epithelia have been known to be involved in the isotonic fluid transport when the reabsorbate is isotonic to the bathing solution. It has been established that ion and water fluxes across the leaky epithelia are tightly coupled (Whitlock and Wheeler, 1964; Sackin and Boulpaep, 1975; Hill, 1980; Loo et al., 1996; Spring, 1998). The exact mechanisms for the solute–solvent coupling in epithelial layers are yet to be determined. Various transport processes in the leaky epithelia are supposed to be driven by transmembrane ion gradients that are formed mainly by Na⫹ ions. The Na⫹ that enters the cell with the fluid is pumped out by the Na⫹-K⫹-ATPase which, by maintaining the Na⫹ gradient, indirectly drives the transport. There are some discrepancies in the literature about pathways of fluid transport across different epithelia. Investigators have proposed alternative explanations: the pathways provide either exclusively paracellular (Hill,

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1980; Steward, 1982), or exclusively transcellular transport (Fischbarg and Montoreano, 1982; Persson and Spring, 1982). For the explanation of mechanisms of water transport across the epithelia, several alternative models have been proposed: the compartmental model (Curran and MacIntosch, 1962; Whitlock and Wheeler, 1964), and arising from it the standing-osmotic gradient model (Diamond and Bossert, 1967), and the solvent drag model (Andersen and Ussing, 1957; Pappenheimer and Reiss, 1987). However, none of these models is able to reliably explain driving forces for water movement in the absence of externally detectable gradients in osmolality and leakness of TJs. Zeuhten and co-workers (Zeuhten and Stein, 1994; Zeuhten, 1995; Loo et al., 1996) proposed a new idea about the mechanism of the route of transcellular fluid across several leaky epithelia, including the small intestine. This mechanism assumes that water transport is directly linked to solute transport by cotransport proteins such as the brush border Na⫹/glucose transporter and/or the basolateral K⫹/Cl⫺ transporter. They concluded that a secondary active mode of water transport was independent of the osmotic gradient (Zeuhten, 1992; Loo et al., 1996). As indicated in their work, one of the crucial factors for isotonic transport is the cytosol osmolarity (Fischbarg et al., 1985; Zeuhten, 1995). At present there is evidence that the cells of the water-transporting epithelia can be hypertonic to the external solution (Zeuhten, 1995). Because the cell membranes in the leaky epithelia have much higher resistance values than those of the transepithelial resistance, a paracellular shunt pathway for ion flows takes place (Fro¨mter, 1987; Ussing et al., 1974; Cereijido et al., 1989a). It seems likely that water also crosses the junction zone. Apart from TJ, an important role in fluid transport across the epithelia is played by lateral intercellular spaces (LIS). Permeability of TJ and LIS is regulated by many signaling factors and differs in different epithelia. 2. Tight Epithelia Tight epithelia perform hypertonic absorption. These are epithelia of the collecting ducts (CD) of mammalian kidney, amphibian skin, and amphibian urinary bladder. The latter is useful for studies on the salt and water transport as model of CDs that are samples of relative structural simplicity. Note that permeability of tight epithelia to ions and water is regulated by antidiuretic hormones (ADH—vasopressin and oxytocin) secreted by the posterior pituitary gland in response to a deviation in plasma osmolarity. It has been established that Na⫹ and water transport in the tight epithelia are uncoupled processes (Ussing et al., 1974; Martinez-Palomo, 1975; Diamond, 1978). The experimental ADH-induced stimulation of ion and water transport across the amphibian epithelia has allowed researchers to follow the

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sequence of events initiated by the interaction of the hormone with its receptors. 3. Mechanisms of ADH-Stimulation of Water Transport The interaction of the hormone with its receptor at the basolateral membrane triggers the final effector process in the apical membrane of the epithelial cell (Hays, 1968, 1972; Ussing et al., 1974; Finkelstein, 1976; Dousa, 1980; Handler and Orloff, 1981; Sousa and Grosso, 1981; Ausiello and Orloff, 1982; Grosso et al., 1982; Levine, 1985; Sousa, 1985, 1986; Taylor et al., 1987; Franki et al., 1989; Yorio and Satumiira, 1989; Natochin, 1994). Our understanding of the ADH action has been significantly improved during the last decade because of the application of biochemical and molecular–biological methods. It is generally acknowledged that there are two types of receptors binding with ADH—the so-called V1 and V2 of the basolateral membranes of the target cells. V2-receptors binding with ADH activate adenylate cyclase (AC) via a stimulatory G-protein. AC, in turn, initiates an intensive synthesis of cAMP that, alongside with Ca2⫹, is the most important intracellular transmitter in the cell response to ADH. The calcium release is triggered by the interaction of ADH with V1 receptors. The ADH-receptor complex also activates phospholipase C via G-protein. Phospholipase C cleaves phosphoinositol to generate two products—inositol triphosphate (InsP3) and diacylglycerol (DAG). This process leads to the release of Ca2⫹ from calcium-sequestring compartments in the cytosol. DAG has been shown to take part in the synthesis of prostaglandins and related lipid signaling molecules. In addition, DAG activates protein kinase C which, in turn, phosphorylates a number of important cell proteins (Breyer, 1991; Natochin, 1994). Apart from the hormonal effects, a great number of chemical and physical factors are known to affect the ADH-induced water transport. These include sodium concentration in extracellular fluid, local hyperosmolarity, local and circulating hormones, and prostaglandins (Orloff and Zusman, 1978; Schlondorff, 1985; Natochin, 1994). A particularly important role has been shown for autacoids (PGE2) in modulating ADH-action in amphibian osmoregulatory epithelium (Natochin, 1994; Natochin et al., 1996; Parnova et al., 1997). Exciting data about the stimulation of water transport across the tight epithelium of the frog urinary bladder in the absence of ADH have been obtained in recent years (Natochin and Schakhmatova, 1995; Natochin et al., 1996, 1998; Parnova et al., 1997). It seems that several changes of the serosal-bathing Ringer solution leads to an increase in water permeability of the apical membrane of epithelial cells. This phenomenon is accompanied by a decrease in serosal prostaglandin E2 (PGE2) concentration from

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14.8 ⫾ 1.0 in the first solution to 0.6 ⫾ 0.1 nM in the fifth solution. A concentration of 10⫺6 M PGE2 in vitro inhibited activity of membrane AC from highly permeable bladders by 33.4%. A significant activation of AC was observed by PGE2 at a concentration lower than 10⫺10 M. In this case, the trigger role in activation of water transport is played by a decreased level of PGE2 which could stimulate AC. Hence, the stimulation of the water transport is provided by the same molecular mechanisms as in the case of treatment with ADH: activation of AC, accumulation of cAMP, and insertion of water channels into the apical membrane of epithelial cells (Natochin et al., 1996; Parnova et al., 1997; Komissarchik et al., 1998). Similar changes in water permeability occur during the incubation of Ringer solution in the trout urinary bladder, which is composed of ADHinsensitive tight epithelium (Komissarchik et al., 1998; Natochin et al., 1998). These results suggest that not only neuropituitary control but also osmoregulatory epithelium itself can strongly modify its permeability properties via mechanisms of regulation of PGE2, its synthesis, and release (Natochin et al., 1996; Parnova et al., 1997; Komissarchik et al., 1998). B. Ultrastructural Features of Transporting Epithelia Epithelial cell layers are composed of inidividual cells, closely packed in hexagonal arrays and connected by specialized intercellular junctions. The epithelial junctional complex was described as long ago as the beginning of this century as ‘‘occluding bar’’ (Bonnet, 1895; Zimmerman, 1911). Then this fact was confirmed by the classical EM work of Farquhar and Palade (1963). At present, the study of structural and chemical organization of intercellular junctions is one of the most interesting directions of cell biology. The following junctions are known in epithelia: tight junction (zonulae occludens), intermediate junction (zonulae adhaerens), desmosomes (maculae adhaerens), and gap junction (nexus) (Staehelin, 1974; Snigirevskaya and Komissarchik, 1980). One of the essential features of the transporting epithelia is cell polarity. The epithelial cell polarity is seen in the position of cell organelles, the composition of the extracellular matrix, and the distribution of specialized plasma membrane domains. The epithelial cell layer is underlaid by the basal membrane or basal lamina forming a kind of barrier between the epithelium and supporting tissue beneath it. It has been shown that there are certain differences in the ultrastructural organization of absorbing leaky and tight epithelia. 1. Comparison of the Ultrastructure of Leaky and Tight Epithelia Available morphological work indicates that different epithelium types differ in the predominant cell shape: leaky epithelium cells are usually

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extended perpendicularly to the epithelial layer plane, whereas the tight epithelium cells are more flat (Fig. 1) (DiBona et al., 1969a, 1969b; DiBona and Mills, 1979; Maunsbach, 1980). Leaky epithelia are characterized by more developed intermediate junctions and terminal web (TW) than tight epithelia (Farquhar and Palade, 1963; Mooseker and Tilney, 1975; Rodewald et al., 1976). The cells of most tight epithelia have short and irregular microvilli. On the contrary, leaky epithelium cells contain many microvilli packed in a special structure—the brush border (Fig. 1) (DiBona and Mills, 1979; Mooseker, 1985; Bretscher, 1991). Morphological investigations have shown the transepithelial permeability to fluid is higher in epithelia with larger areas of TJs on the luminal absorption surface. A different contribution of the paracellular pathway in the permeability of tight and leaky epithelia has been demonstrated in work studying the relations of the area occupied by apical membranes and tight junction zones (Welling and Welling, 1976, 1979; Mills and Malick, 1977; DiBona and Mills, 1979; Sousa and Grosso, 1981). The morphometric analysis of the mucosa surface of different organs of the same animal (bullfrog) obtained with Nomarski microscopy has shown that in the leaky gallbladder epithelium the total length of the TJ zone inside a square area of 1 cm2 is 26 m, whereas in the tight urinary bladder epithelium it is 6.5 m (DiBona and Mills, 1979). In the proximal tubules of mammalian kidney, with highest

FIG. 1 Diagrams to compare structure of leaky (A) and tight (B) epithelia. In contrast to tight epithelia, leaky epithelia have well-developed brush border, that consists of numerous, closely packed microvilli (Mv), highly expressed intermediate junction (IJ), and terminal web (TW). Golgi complex (Gc) in the tight epithelia is characterized by more narrow and flat cisternae than in the leaky epithelia.

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permeability among all epithelia, the total length of the tight junction zone on the mucosa surface is 50–100 m (Welling and Welling, 1976). Many leaky epithelia (mammalian small intestine, choroid plexus of early sheep fetuses, rat proximal tubules, rabbit gallbladder) have a welldeveloped intracellular system of tubules and cisternae of ER (Thiery and Rambourg, 1976; Møllgaa˚rd and Rostgaard, 1978; Maunsbach, 1980; Vinnichenko, 1980; Mosevitch et al., 1987; Parisi et al., 1995). The system seems to be most abundant in leaky epithelia with low transepithelial resistance involved in transcellular transport (Møllgaa˚rd and Rostgaard, 1978). The special feature of leaky epithelia is their extensively folded lateral membrane. Quantitative measurements indicate that folding significantly increases the membrane area (Hill and Hill, 1978; Bungaard and Zeuhten, 1982). The features of tight junction and apical membrane structure in different epithelia will be discussed in Sections III and IV, respectively.

2. Leaky Epithelium of the Rat Small Intestine The morpho-functional investigations of structural correlates of the small intestine is rather difficult due to the complex tissue geometry and heterogeneity of its cell types. The proximal small intestine of the rat consists of one layer of cylindrical cells closely packed in an hexagonal array so that small spaces (18–20 nm) are formed between the plasma membranes of adjacent lateral cell surfaces. There are several types of epithelial cells, the predominant one being the columnar absorptive cell, enterocyte (Fig. 2). The luminal (apical) surface of enterocytes has a unique specialized structure revealed for the first time in the light microscope and called the striated or brush border (Ham and Cormack, 1979; Mooseker, 1985). EM has revealed the striated border to be a series of finger-shaped cytoplasmic projections or microvilli (Mv), each limited with the plasma membrane of the enterocyte (Figs. 2 and 3a). They expand the membrane surface available for absorption and transport of nutrients (Bretscher, 1991; Madara, 1998). The microvillar membrane is rich in membrane transporters, membrane-anchored hydrolases, and other multienzyme complexes (Ugolev and Iezuitova, 1991). In the apical part of intercellular boundary, the epithelial cells are connected with each other by the following specialized intercellular junctions: tight junction, intermediate junction, and desmosomes (Farquhar and Palade, 1963). In the basolateral part of the epithelium, the plasma membranes form interdigitating folds attaching the adjacent cells to each other (Fig. 2). The epithelial cells are supported by the basement membrane.

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Enterocytes are characterized by the presence of a well-developed cytoskeleton consisting of actin filaments (microfilaments), intermediate filaments, and microtubules (Fig. 2). Microfilaments forming the dense bundles within Mv expand into the cytoplasm where they form the terminal web (TW) associated with the junctional complex (Figs. 2 and 3a). In addition, the TW contains myosin I that forms a complex with calmodulin molecules, fimbrin, tropomyozin, villin, and minor component ezrin (Bretscher, 1991; Balda and Matter, 1998). Intermediate filaments of a keratin nature are attached to the desmosomes and form thick dense bundles expanding up to the nucleus (Figs. 2 and 3c) (Franke et al., 1981; Snigirevskaya et al., 1997). Intracellular organelles are mainly observed beneath the TW. These are nucleus and usual cytoplasmic constituents, such as mitochondria, ER cisternae, Golgi complex (Gc), and ribosomes. 3. Tight Epithelium of the Frog Urinary Bladder Morphology of the amphibian urinary bladder is described in a series of works (Wade and DiScala, 1971; DiBona and Civan, 1973; Chevalier et al., 1974; Bourguet et al., 1976; Humbert et al., 1977; DiBona, 1978; Reaven et al., 1978; Wade, 1978; Kachadorian et al., 1979, 1987; DiBona and Walker, 1985; Komissarchik et al., 1982, 1985). The epithelium of the amphibian urinary bladder represents the squamous transitional epithelium. It is composed of a single layer of thin cells supported by the basal lamina. There are four cell types in the epithelium—goblet, basal, granular, and mitochondria-rich. The most numerous cells are the granular cells; for example, in Dominican toads the mucosal surface is ⬎95% occupied by granular cells (Wade, 1978). In the scanning EM each mitochondria-rich cell is seen to be surrounded by three to five granular cells (not shown). All the cells are connected by a junctional complex consisting of tight junction, a weakly developed intermediate junction, and several desmosomes (Figs. 4a and 4b). The apical surface forms finger- or ridge-shaped Mv (Davis et al., 1981; Komissarchik et al., 1985). The apical membrane is coated with the well-developed

FIG. 2 Electron micrograph of parts of two enterocytes of a small intestine. The cells are joined by the junctional complex, consisting of tight junction (TJ), intermediate junction (IJ), desmosomes (D), and basolateral interdigitating folds (arrows). Microvilli (Mv) are revealed on the apical surface. The cytoplasm contains mitochondria, ER cisternae, ribosomes, and cytoskeletal elements: microtubules (MT), microfilaments (MF), and intermediate filaments (IF). Bar ⫽ 0.5 애m.

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glycocalyx, and basolateral cell borders have interdigitations forming a space 15–50 nm wide (Figs. 4a and 5b). Granular cells of the unstimulated amphibian urinary bladder contain numerous granules of various shape and density, mitochondria, ribosomes, and cytoskeletal elements (7-nm microfilaments, 10-nm intermediate filaments, and 20-nm microtubules) (Figs. 4a, 4b, 5a, and 5b). Single microtubules are mainly oriented along the long axis of the cell, whereas microfilaments are distributed along the cytoplasm, randomly condensing in the Mv, basolateral processes, and submembrane apical zone (Komissarchik et al., 1982; Snigirevskaya and Komissarchik, 1987, 1993; Gorshkov and Komissarchik, 1998). As in other tight epithelia, the TW of the granular cells is poorely developed. Antiactin antibodies conjugated with colloidal gold particles label the submembrane apical zone rather intensively (Figs. 5c and 5d). The vacuolar system of granular cells is represented by cisternae of the smooth and rough ER, stacks of the Golgi complex (Gc) and sometimes electron dense canaliculi lying commonly in the apical region of the cell.

III. The Role of Tight Junctions in Transepithelial Transport Tight junctions (TJ) (also called occluding junctions or zonulae occludens) form a continuous belt around the apical end of the cell. They are present in both vertebrate and invertebrate epithelial tissues (Farquhar and Palade, 1963; Friend and Gilula, 1972; Snigirevskaya and Komissarchik, 1980; Lane, 1981; Woods and Bryant, 1993). Tight junction is the most apical specialized zone of the intercellular junctional complex including the tight junction, intermediate junction, desmosomes, and gap junction. Tight junctions are known to determine the transport functions of epithelia. Together with the intermediate junction, they hold neighboring cells through a family of Ca⫹2-dependent cell–cell adhesion molecules (cadherins) that are linked to actin and myosin filaments (Denker and Nigam, 1998).

FIG. 3 Immunogold labeling of cytoskeletal elements in rat enterocytes. (A) Antiactin label in the control enterocyte is revealed on microfilaments of the microvilli (Mv) and roots; a weak label is seen near the tight junction (TJ). (B) After glucose load the label between the root filaments and in the TJ region is more intensive (arrows) (Reproduced from Snigirevskaya et al., 1997, with permission). (C) Antikeratin antibodies label bundles of intermediate filaments (IF). Bar ⫽ 0.5 애m.

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A. Structure of Tight Junctions We will not describe the TJ structure in detail because there are many excellent reviews on this subject (Lane, 1981; Pinto da Silva and Kachar, 1982; Schneeberger and Lynch, 1992; Citi, 1993; Woods and Bryant, 1993; Anderson and van Itallie, 1995; Balda and Matter, 1998; Cereijido et al., 1998; Denker and Nigam, 1998; Mitic and Anderson, 1998; Matter and Balda, 1999). In ultrathin sections, tight junction is revealed in the apical part of lateral boundaries between epithelial cells (Figs. 2, 3a, 3b, 4a–4c, and 5d). This zone appears as a series of spot contacts or kisses where the adjoining membranes are fused by their outer leaflets into an electron-dense line, 2–3 nm thick (Farquhar and Palade, 1963; Snigirevskaya and Komissarchik, 1980; Schneeberger and Lynch, 1992). These spotlike contacts correspond to continuous branching strands revealing as ridges on the P-face (PF) of fractured lateral membranes of epitheliocytes (Staehelin, 1973, 1974). The strands consist of intramembrane particles (IMPs) and completely circumscribe the cell (Figs. 6a and 6b). Most authors think that these IMPs have a protein nature. According to another model proposed by Pinto da Silva and Kachar (1982), the TJ strands represent intramembranous, cylindrical, inverted lipid micelles stimulated to form and perhaps to be maintained at the junctional site by integral membrane proteins. Immediately below the TJ, the intermediate junction forms also a continuous circumferencial belt around the cell (Figs. 2, 3a, 4a, and 5d). In the lateral cell membranes in the intermediate junction region, the circumferential ring of actomyosin filaments is inserted (Hirokawa and Tilney, 1982). For the past several years, a long list of proteins comprising the tight junction zone has been reported: occludin, ZO-1, ZO-2, cingulin, 7H6, semplekin, numerate signaling proteins (Stevenson et al., 1986; Gumbiner et al., 1991; Citi, 1993; Keon et al., 1996; Denker and Nigam, 1998; Mitic and Anderson, 1998; Yap et al., 1998). Occludin is considered to be an integral protein functioning as a cell–cell adhesion molecule, whereas proteins ZO-1 and ZO-2 bind occludin to the cytoplasmic plaques. Using immunogold labeling, freeze-fracture techniques, Fujimoto (1995) showed occludin

FIG. 4 Reaction of lateral intercellular spaces (LIS) to the stimulated fluid transport. (A) Boundary between two granular cells of the control epithelium of the frog urinary bladder. In the apical part of the epithelial layer, the cells are joined by intercellular junctions, in the basolateral part, by interdigitating folds. LIS are narrow. (B) After ADH stimulation of water transport the LIS are significantly dilated. The intercellular junctions are not changed. (C) After glucose load of rat small intestine, the large dilations of LIS are observed (Reproduced from Snigirevskaya et al., 1997, with permission). Bar ⫽ 0.25 애m.

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FIG. 6 Freeze-fracture replicas of tight junction (TJ) region in epithelia. (A) The TJ in the frog urinary bladder. PFA and EFA, P-face and E-face of the apical membrane, respectively; PFL and EFL, P-face and E-face of the lateral membranes. The strands of TJ are revealed as ridges on PFL, and grooves, on EFL. (B) The same in the rat small intestine. On the top of the picture fractured microvilli (Mv) are seen (Reproduced from Snigirevskaya et al., 1997, with permission). Bar ⫽ 0.25 애m.

FIG. 5 Parts of granular cells of the frog urinary bladder. (A) Numerous granules are crowded in the middle part of control granular cells. The apical membrane is coated by the glycoclyx (Gl). (B) After ADH stimulation of water transport, granules migrate to the apical surface. (C) Antiactin antibodies label the cortical actin preventing the approach of granules to the apical membrane. (D) After ADH action the cortical actin is partly depolymerized, and granules approach the apical membrane (arrows). Bar ⫽ 0.5 애m.

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to be located in the tight junction strands. The structure and function of occludin is the subject of many studies (Furuse et al., 1993; Balda et al., 1996; McCarty et al., 1996; van Itallie and Anderson, 1997; Haskins et al., 1998; Mitic and Anderson, 1998). Transmembrane TJ domains are linked to the cytoskeleton via a complex of peripheral membrane proteins. Studies of the effect of Cytochalasin B and D on epithelial structure and function indicate that the actin filament organization might play an important role in maintaining epithelial cell shape and barrier function (Meza et al., 1980; Stevenson and Begg, 1994).

B. Participation of the Tight Junction in the Maintenance of Cell Polarity The essential role of TJ in establishment and maintenance of epithelial polarity has been shown in many studies (Koefoed-Johnson and Ussing, 1953, 1958; Simons and Fuller, 1985; Rodriguez-Boulan and Nelson, 1989; Mellman et al., 1993; Rodriguez-Boulan and Zurzolo, 1993; Cereijido et al., 1989a, 1989b, 1993, 1998; Balda and Matter, 1998; Mitic and Anderson, 1998; Matter and Balda, 1999). By circling the cell at the boundary between the apical and lateral regions, the TJ provides a strong physical attachment between cells, regulates transepithelial transport, and forms a fence in the membrane bilayer between apical and basolateral domains of the plasma membrane. The TJ allows segregating cell surface membrane proteins and lipids into the apical and basolateral membrane domains and restricts their free diffusion in the plane of the plasma membrane.

C. Tight Junction as a Diffusion Barrier for Solutes In addition to apical cell membranes, the TJs are considered to be one of the main barriers to diffusion of solutes between epithelial cells (Schneeberger and Lynch, 1992; Mitic and Anderson, 1998). The tight and leaky epithelia can be distinguished depending on the tightness of TJ for ion diffusion (Fro¨mter and Diamond, 1972; Ussing et al., 1974; Diamond, 1974, 1978, 1979; DiBona and Mills, 1979; Lewis et al., 1984; Fro¨mter, 1987). Unlike the leaky epithelia, the tight epithelia can support high osmotic and ion gradients and a marked transepithelial potential difference. This is determined by the low water and ion permeability of their apical membranes and TJs (Ussing, 1980; Kirk et al., 1984a, 1984b; Lewis et al., 1984; Fox, 1986; Finkelstein, 1987; Strange and Spring, 1987).

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As shown by morphological studies, the larger area of TJ zones on the luminal absorption surface of the epithelium, the higher its transepithelial permeability to fluid (Welling and Welling, 1976; DiBona and Mills, 1979). Application of the freeze-fracture method has revealed a correlation between the ultrastructural organization of TJ and epithelial permeability (Claude and Goodenough, 1973; Claude, 1978; Oschman, 1978; Sardet et al., 1979). In some cases there is a strong correlation between the amount of junction strands and epithelial permeability; this is seen in the tight epihelium of distal channel and leaky epithelium of mouse kidney proximal tubule. The former contains five strands; the latter contains only one strand. However, this regularity cannot be found in all epithelia. Thus, the mammalian small intestine epithelium with the high ion and osmotic permeability has many strands (Fig. 6b). In recent years, researchers have obtained physiological evidence has showing that TJ domains of many leaky epithelia ˚ that can explain their contain cation selective pores in the range of 8–20 A high permeability (Reuss, 1991; Mitic and Anderson, 1998). Morphologically, the barrier function of TJ can be assessed by the ability of TJ to block the passage of electron dense molecules, such as ruthenium red, lanthanum, and cationic ferritin (Mullin et al., 1997). 1. Mechanisms of Regulation of the Tight Junction Permeability in Leaky Epithelia At present, it is clear that TJ is a highly dynamic and physiologically regulated structure. The TJ does not constitute an absolute diffusion barrier, but it allows the passage of water and certain solutes (Cereijido et al., 1993; Balda et al., 1996). Details of the molecular events that result in TJ regulation are not completely elucidated; however, several clues are available. First, structural alterations of perijunctional cytoskeleton participate in the TJ changes. An example of physiological regulation by physiological signals of the junctional solute permeability includes the TJ regulation during intestinal nutrient absorption (Pappenheimer, 1993; Madara, 1988, 1998; Turner and Madara, 1995). Electron microscopic study of the small intestine loaded with glucose at high concentrations has identified focal discontinuities in TJ at which water and macromolecular permeation can occur (Madara and Pappenheimer, 1987). In addition, Madara and Pappenheimer have revealed structural alterations in the perijunctional actin-based cytoskeleton, indicating an enhanced cytoskeletal tension. These data as well as results of physiological experiments have led them to a conclusion that, under physiological conditions at relatively high luminal concentrations of glucose, its absorption predominantly occurs by solvent drag through the paracellular channels, whereas the active membrane transport of glucose plays a

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secondary role as a trigger mechanism. Glucose transport across the apical membrane activates contraction of the enterocyte cytoskeleton, thereby opening tight junctions, and forms a concentration gradient of the substrate across the junctions for an increased fluid transport. However, our study of glucose absorption by the rat small intestine has shown the TJ permeability alterations to affect nutrient uptake very little (Ugolev et al., 1995, Snigirevskaya et al., 1997). In binding intestine loops in vivo in acute and chronic experiments, we showed a close correlation between the absorption rates of glucose and water; even at a maximal glucose concentration (75 mmol/ l) the rate of the solvent drag transfer of fluid does not exceed 10% of the total glucose absorption. In our EM study we have never observed structural changes in the TJ structure; however, some condensations of actin associated with TJ did occur. Immune EM has shown that the density of antiactin labeling increases in the area close to TJ (Figs. 3a and 3b) (Ugolev et al., 1995). So, we can think that certain alterations of TJ organization can take place at extremely high luminal concentrations of glucose, indicating tension of the enterocyte perijunctional ring. In addition to actin, the perijunctional ring contains myosin II. By contracting this complex, it is possible to promote paracellular solute permeation (Madara, 1988, 1998). The ability of enterocytes to contract in the perijunctional ring has been shown during isolation of brush border (Rodewald et al., 1976; Mosevitch et al., 1982). The experiments with cytochalasin B and D treatment confirm that microfilaments are involved in a dynamic regulation of TJ barrier function (Meza et al., 1980; Kovbasnjuk et al., 1998). Thus, an important role of the perijunctional actin cytoskeleton in the TJ regulation is demostrated in the series of studies. One of essential factors of physiological regulation of junctional permeability is Ca2⫹, which is known to be one of many agents involved in synthesis and maintenance of TJ and apical basolateral cell polarity (Cereijido et al., 1998). Recent studies have shown that Ca2⫹ promotes cell–cell contacts and activates a cascade of intracellular reactions including phospholipase C, protein kinase C, and calmodulin (Balda et al., 1991; Cereijido et al., 1998; Mitic and Anderson, 1998). In the early 1970s we showed role of Ca2⫹ in TJ function and structure regulation in the rat liver with treatment by EDTA in a Ca2⫹-free medium (Komissarchik et al., 1976). To follow the TJ alterations, a morphologically detected tracer, lanthanum chloride, was used. When applied basolaterally in the normal liver, La3⫹ does not permeate the junctional zone between hepatocytes. After withdrawal of extracellular Ca2⫹, lanthanum enters the lumen of bile canaliculi. Similar results were obtained after Ca2⫹ removal from the bathing solution in the presence of EGTA in the rat small intestine and MDCK cells (Contreras et al., 1992). After this treatment the permeability of the rat small intestine to phenol red increases tenfold.

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Considering recent data on this subject, we can suggest that removal of extracellular Ca2⫹ leads to the washing out of cadherin, mediating the calcium-dependent cell–cell adhesion. The essential role of E-cadherin in the establishment and maintenance of TJs in epithelia has been recently shown by several authors (Behrens, 1993; Madara, 1998). For the last few years it has been found that a number of signaling molecules are implicated in the regulation of TJ function, including thyrosine kinase, cAMP, protein kinases, phospholipase C, small GTPases rho, rac, and Cdc 42 (Cereijido et al., 1993; Anderson and von Itallie, 1995; Balda et al., 1996; Balda and Matter, 1998; Madara, 1998). As mentioned earlier, regulatory factors might mediate the selective diffusion across TJ by modifying aqueous channels embedded into tight junctional strands and composed of the integral protein occludin (Fujimoto, 1995; Balda et al., 1996; Matter and Balda, 1999). Data in the literature show that occludin expression and localization in the TJ correlates with the establishment of the paracellular permeability barrier in cultured epithelial monolayers (Wong and Gumbiner, 1997). It is possible that in the regulation of TJ during physiological and patophysiological events the dynamics of assembly/depolymerization of occludin plays an important role. Note that the lateral intercellular space also contributes to paracellular resistance. In Necturus gallbladder this contribution is minor (only 5%) compared with that of the TJ. However, treatments which collapsed the lateral intercellular space increased its contribution up to 30% of the total paracellular resistance (Kottra et al., 1993). 2. Behavior of Tight Junctions in Tight Epithelia During Rise in Water Absorption The permeability of tight epithelia for water, sodium ions, and urea significantly rises under effect of vasopressin (Bentley, 1958; Pak Poy and Bentley, 1960; Peachey and Rasmussen, 1961; Natochin, 1963, 1994; Natochin and Shakhmatova, 1966; Ganote et al., 1968; DiBona et al., 1969a, 1969b; Pietras and Wright, 1975; DiBona, 1978; Ussing, 1980; Kirk et al., 1984a, 1984b; Fox, 1986; Strange and Spring, 1987). Using the freeze-fracture technique to compare TJ structure of the frog urinary bladder epithelium at low and increased water permeability, we revealed no changes in the amount, configuration, and distribution of TJ fibrils. Meanwhile, TJ of tight epithelia also represents a dynamic cell structure changing its organization under the effect of certain factors. Thus, intercellular spaces in the TJ zones in the frog urinary bladder epithelium can be opened by serosal hypertonicity (Ripoche et al., 1973; Fischbarg et al., 1976; Parisi et al., 1995), by removal of extracellular Ca2⫹ (Komissarchik et al.,

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1976, 1978b; Meldolesi et al., 1978; Hardy and DiBona, 1982), or by serosal treatment with sulfhydryl reagent, ␳-chloromercuryphenylsulfonic acid (Bagrov et al., 1993). The preliminary treatment of the urinary bladder by low concentrations of glutaraldehyde stabilized peripheral proteins of basolateral membranes and prevented their washing out, saving TJ integrity (Komissarchik et al., 1976).

IV. Ultrastructure of Apical Membranes of Epitheliocytes A most important area of contact between the organism and its environment is the apical surface of epithelial cells. The apical membrane of epitheliocytes contains specific proteins involved in various transmembrane transport processes. In many epithelial cells the absorbing surface is greatly increased by the presence of numerous microvilli forming a specialized structure, the so-called brush border. As mentioned in Section II.B, there is a substantial difference between the structure of the apical surface in tight and leaky epithelia. Leaky epithelia are richer in pronounced and regularly packed Mv than tight epithelia. Apical membranes of both the leaky and tight epithelia have the same trilaminar structure revealed by conventional EM methods. The widespread use of freeze-fracture techniques, however, has allowed discovery of specific structural features of apical membranes in different types of epithelia.

A. Structure of Apical Membranes 1. Leaky Epithelia The enterocyte apical membrane has numerous, tightly and regularly packed Mv with a mean diameter of 0.2 애m. In thin sections, the apical membrane consists of two osmiophilic layers and one electron transparent layer. The outer membrane leaflet is thicker than the inner one and is coated with long threads of glycocalyx. Between the Mvs in the apical membrane, occasional coated pits can be revealed. In freeze-fracture replicas the middle hydrophobic part of the membrane is exposed (Branton, 1966). As in most cell membranes, intramembrane particles (IMPs) of the fracturing apical membrane are divided between P-face and E-face (EF). The PF contains more particles per square micro-

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meter of surface than the EF. IMP of both fracturing surfaces between MVs are scattered more or less uniformly (Fig. 7a). 2. Tight Epithelia Apical cell membranes of tight epithelia are characterized by a low water and ion permeability (Peachey and Rasmussen, 1961; DiBona et al., 1969a, 1969b; Wade et al., 1975; Natochin and Chapek, 1976; Masur et al., 1984). In ultrathin sections the apical membrane of the frog urinary bladder epithelium cells has a typical three-layer structure with wider outer layer coated with glycocalyx (Komissarchik et al., 1985). The glycocalyx is especially well developed on Mv radiating from their tops as long threads (Figs. 4b and 5b). Some specific granules of the granular cells have been suggested to be contributors of glycocalyx precursors (Masur et al., 1986; Komissarchik et al., 1978a). Freeze-fracture studies of the epitheliocyte apical membrane structure have established that there is a correlation between membrane ultrastructural organization and its water permeability (Chevalier et al., 1974; Wade et al., 1975; Komissarchik et al., 1985). It seems that the distribution of intramembrane particles in the apical membrane of tight epithelium cells, specifically in granular and mitochondria-rich cells of amphibian skin and

FIG. 7 Freeze-fracture replicas of the apical membranes of epithelial cells. (A) PF of the apical membrane of the rat enterocyte. The uniform distribution of intramembrane particles (IMPs) is revealed between microvilli. (B) After glucose load, the uniform distribution of IMPs is disturbed, and numerous IMPs are seen in the bases of Mv (arrows). (C) PF of the granular cell apical membrane of the frog urinary bladder contains a few small IMPs. (D) EF of the apical membrane of the granular cell contains numerous large IMPs. Bar ⫽ 0.25 애m. (Parts A and B reproduced from Snigirevskaya et al., 1997, with permission).

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urinary bladder, and of the principal cells of the mammalian collecting ducts is inverted in comparison with the rest of the cell membranes (Chevalier et al., 1974; Kachadorian et al., 1975, 1977, 1986, 1987; Humbert et al., 1977; Hays, 1983; Kachadorian and Mu¨ller, 1984; Komissarchik et al., 1985). Unlike most of the cell membranes containing numerous IMPs on the P-face and a few IMPs on the E-face, the apical membranes of tight epithelium cells have a few IMP on PF and many IMP on EF (Figs. 7c and 7d). In the frog urinary bladder, distribution of IMPs on the apical membrane fracture surfaces is rather uniform, whereas their amount in different cells of the same epithelium varies significantly, which suggests that these cells are in different functional states (Komissarchik et al., 1985). Note that the IMP diameter of PF is lower than the diameter of IMP on EF (14.5 and 17 nm, respectively) (Komissarchik et al., 1985). According to the data of Chevalier et al., (1985), large IMPs on EF consist of two to four subunits. Using the label-fracture method, these authors have shown that large particles of EF correspond to protein structures that reach the outer surface and/or are associated with the glycoprotein components of the cell coat. Analysis of the distribution of cholesterol labeled by filippin with the freeze-fracture method has demonstrated that the density of filippin– cholesterol defects in the basolateral membranes are higher (270⫾10) than in the apical ones (162⫾11). On ADH-stimulation on water flow, both density and polarity of filippin-induced deformations are altered differently in the apical and basolateral regions of the plasma membrane. The filippininduced deformation density in the apical membrane is decreased to 170⫾14. The results obtained may suggest that the appearance of ADHinduced IMP aggregates is accompanied by a relative cholesterol decrease in the apical membrane (Kever et al., 1988). Similar data have been obtained in the works of Orci et al. (1980) and Stetson and Wade (1983).

B. Dynamics of Structural Changes during Stimulated Fluid Transport 1. Leaky Epithelia The thin section technique fails to observe any changes in the enterocyte apical membrane during the nutrient absorption event at a concentration up to 70 mM. Meanwhile, the freeze-fracture method makes it possible to see redistribution of IMPs on the enterocyte apical membrane during fluid absorption. On PF the uniform scattering of IMPs is disturbed, and smooth areas appeared between microvilli (Snigirevskaya et al., 1997). The most abundant IMPs are present around the bases of microvilli (Fig. 7b).

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Snigirevskaya et al., (1997) suggested that the replacement of IMPs along the membrane has some connection to the conformational changes of the cotransport proteins of the enterocyte apical membrane. Recently, Zeuhten and his coauthors (Loo et al., 1996) proposed that the secondary active water transport is performed by the Na⫹-glucose cotransporters. Thus, these changes of IMP distribution can be affected not only by the transfer of glucose and Na⫹ but also by water. However, we cannot forget that the enterocyte apical membrane contains other integral proteins, aquaporins, participating in the transmembrane water transport. However, the data in the literature about the presence of aquaporins in small intestine enterocytes are insufficient. At present there are only a few molecular biological works indicating that AQP3, AQP4 (Granquist et al., 1996), and AQP9 (Kuriyama et al., 1997) are expressed by the small intestine cells. 2. Tight Epithelia During stimulated transepithelial water transport, significant changes of the apical membrane structure have been observed in granular cells of the amphibian urinary bladder and principal cells of mammalian CDs (Chevalier et al., 1974, 1979; Kachadorian et al., 1975, 1977; Wade et al., 1975; Bourguet et al., 1976; Harmanci et al., 1980; Lacy, 1981; Mu¨ller et al., 1980, 1981; Sousa and Grosso, 1981; Hays et al., 1982; Komissarchik et al., 1985; Wade, 1978). In these tissues, with increased water permeability of epithelia, aggregates of large IMPs appear on the PF of the fractured apical membrane (Fig. 8a). The complementary surface of EF contains a few big IMPs and thin parallel ridges (not shown). The aggregates of IMPs on PF vary on their shape (e.g., roundish, asterisk-like and rectangular). In the apical membranes of the amphibian urinary bladder granular cells, the aggregates are characterized by a dense array of IMPs (Bourguet et al., 1976; Humbert et al., 1977; Wade, 1978; Chevalier et al., 1979; Mu¨ller et al., 1980), often tetragonal (Komissarchik et al., 1985, 1998; Komissarchik and Snigirevskaya, 1990, 1991). The apical membrane of the CD principal cells of mammalian kidney, on the contrary, contains aggregates with loose packed IMPs (Lacy, 1981; Orci et al., 1981; Brown et al., 1988). In the frog urinary bladder granular cells, they are often located in the vicinity of the sites of the fusion of the granule membranes with the apical membrane (Fig. 8a) (Komissarchik et al., 1985, 1998; Komissarchik and Snigirevskaya, 1990, 1991). Calculations have shown that the membrane area occupied by IMP aggregates is proportional to the transepithelial water permeability (Kachadorian et al., 1977; Brown and Orci, 1983; Hays, 1983; Komissarchik et al., 1985; Mu¨ller and Kachadorian, 1985). It has been shown that the amount of IMP aggregates in the toad urinary bladder is related to a concentration

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of hormone bathing the bladder and to the duration of its action (Kachadorian et al., 1975, 1977, 1985; Chevalier et al., 1979; Wade, 1978; Kubat et al., 1989). A comparison of the principal cell ultrastructure of CDs of normal mice and of mice with nephrogenic diabetes insipidus unable to synthesize vasopressin has also revealed the presence of a correlation between the apical membrane area occupied by IMP aggregates and transepithelial water permeability. The apical membranes of the latter mice contained no IMP aggregates at all (Brown et al., 1980, 1985, 1988; Harmanci et al., 1980). A similar correlation between the membrane area with aggregates of IMPs and the water flow intensity was demonstrated under the inhibitory effect of Co2⫹ ions on the ADH-stimulated water transport in the frog urinary bladder (Natochin, 1985; Komissarchik et al., 1988). The relative membrane area occupied by IMP aggregates during ADH action is 1.5% of apical membrane area, meanwhile after the Co2⫹ inhibition of the induced water flows, this area is as low as 0.3% (Komissarchik et al., 1988). These results and the fact that the aggregate distribution density is not affected by urea and sodium inhibitors of the ADH-stimulated transport of these substances confirm with certainty the suggestion that these aggregates are the membrane domains with high water permeability (Chevalier et al., 1974, 1979, 1985; Wade et al., 1975; Bourguet et al., 1976; Sousa and Grosso, 1981; Wade, 1978; Komissarchik et al., 1985). The discovery of the IMP aggregates in the apical membranes of the ADH-treated urinary bladders in the absence of an osmotic gradient proves that they are not a consequence of the water flow across the membrane but rather that they function as transmembrane water permeation (Kachadorian et al., 1977; Rappoport et al., 1981; Kachadorian, 1985; Mu¨ller and Kachdorian, 1984, 1985). For the last few years, similar changes in the apical membrane structure of the frog urinary bladder epithelium, which occurred after an increase in water permeability by repeated changes of the serosal Ringer solution, in the absence of ADH, leading to washing out of autacoids (Parnova et al., 1997), have been of special interest.

FIG. 8 Fusion of specific granules with the apical membrane in the frog urinary bladder epithelial cells. (A) Freeze-fracture replica of PF. The aggregates of IMPs are seen near the fusion site of the granule with the apical membrane. (B) The fusion of specific granules with the apical membrane (freeze-substitution). (C) Freeze-fracture replica of the cytoplasm of two granular cells. Arrows indicate the fusion sites of elongated granules with the apical membrane. (D) After the fusion of granule membrane with the apical membrane, the granule content remains in the cytoplasm. Bar ⫽ 0.25 애m. (Parts A and C reproduced from Komissarchik et al., 1989, with permission).

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a. Cytoplasmic Sources of IMP Aggregates Being able to determine the origin of IMP aggregates in the apical membranes of ADH-sensitive epithelial cells during stimulated water transport is extremely important for understanding mechanisms of the transmembrane water permeability. The main contribution to this problem can be made using current EM methods such as cryomethods and immunogold labeling. As mentioned earlier, high water permeable IMPs are concentrated in limited domains but are not distributed uniformly along PF of the apical cell membrane. Such a localization of water channels can possibly be explained by inserting them into the plasma membrane from some cytoplasmic sources. This suggestion is confirmed by quantitative analysis of the distribution of IMPs on PFs and EFs of the fractured apical membrane. The amount of IMPs on EF and PF after the ADH action has been shown to rise by 30% (Komissarchik et al., 1985). According to measurements of electrical capacity, the total area of the apical membrane also increased by 8–30% (Ellis et al., 1980; Stetson et al., 1982; Palmer and Lorenzen, 1983; Lewis and Hanrahan, 1986; Erlij et al., 1989). Moreover, using the freeze-fracture method, a direct proof of the intracellular origin of high water permeable domains in the apical membrane was obtained. Many authors reported that the IMP aggregates on PF are located in the close vicinity of fusion sites of the apical membrane with intracellular membranes (Eggena, 1972a, 1972b; Parisi et al., 1981; Komissarchik et al., 1984, 1985, 1988; Masur et al., 1984, 1985; Mu¨ller and Kachadorian, 1984, 1985; Harris et al., 1986; Coleman et al., 1987; Komissarchik and Snigirevskaya, 1990). Several interpretations of similar pictures are presented in the literature. The most widely held opinion is that some specialized intracellular tubular membrane structures, aggrephores, fuse with, and are inserted into, the apical membrane IMP aggregates during ADH action. This kind of aggrephore was observed in granular cells of the toad urinary bladder (Humbert et al., 1977; Hays et al., 1979, 1982, 1985, 1987; Ellis et al., 1980; Mu¨ller et al., 1980; Kachadorian et al., 1979; Wade et al., 1981, 1984; Sasaki et al., 1984; Ding et al., 1985; Franki et al., 1986). Wade and co-authors (1981) have proposed a hypothesis of a shuttle mechanism of aggrephore movements during ADH action. Later on, this hypothesis was accepted by most investigators. With respect to aggrephores, they have not been found in CD epithelial cells and have seldom been observed in frog urinary epithelia (Komissarchik et al., 1982, 1985). In addition, on the freeze-fracture replica through the granular cell cytoplasm, they can be erroneously interpreted as specific granules that often have an elongated shape (Figs. 5b and 8c) (Komissarchik et al., 1985, 1998). The membrane structure of aggrephores described in the literature and some specific granules observed by us is very similar (Komissarchik et al., 1998).

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The data obtained in our studies have allowed us to assume that the water permeable domains of the apical membrane in frog urinary bladder epitheliocytes are originated from membranes of the specific granules (Komissarchik et al., 1984, 1985). The granular cells of the frog urinary bladder contain several types of specific granules representing derivatives of Gc. One type of these specific granules has been shown to be involved in renewal of the cell glycocalyx (Komissarchik et al., 1978b; Masur et al., 1986). Like some other authors, we have observed in ultrathin sections the increased migration of these specific granules to the epithelial cell apical surface during stimulation of water transport across the frog urinary bladder wall (Figs. 5b and 8b–8d) (Masur et al., 1971, 1972; Gronowitz et al., 1979; Rubin et al., 1982; Komissarchik et al., 1985; Masur and Massardo, 1987; Komissarchik and Snigirevskaya, 1990, 1991). Using EM cryomethods (freeze-fracture, freeze-substitution), which are especially valuable for revealing the fast membrane fusion processes, we observed the processes of fusion of the apical and granule membranes (Figs. 8b and 8c) (Komissarchik and Snigirevskaya, 1990, 1991). It has been suggested that the process of fusion between granules and the apical membrane are regulated and triggered by the ADH stimulatory action. Surprisingly, we have found that the contents of some granules remain in the cytoplasm (Fig. 8d). The granule contents seem to be used during cell response to the increased water transport across the epithelium. Monomere tubulin and calcium ions may be stored by some granules that seem to be released into the cytoplasm in response to the ADH action (Schakhmatova et al., 1982; Davis et al., 1987; Snigirevskaya and Komissarchik, 1987, 1993). This hypothesis regarding the origin of IMP aggregates from the specific granules cannot be considered universal. It is known that such vasopressinsensitive epithelia as mammalian collecting ducts and skin cells of some amphibia do not have specific granules (Brown et al., 1980, 1988; Ding et al., 1985; Hays et al., 1985; Harris et al., 1986; Coleman et al., 1987), whereas the Xenopus laevis urinary bladder epithelial cells containing the granules do not respond to vasopressin (Harris and Handler, 1988). The third conception considers the water channels to be delivered to the plasma membrane by the small vesicles located in the apical cytoplasm (Brown and Orci, 1983; Brown et al., 1985, 1988; Brown and Stow, 1996). These data were obtained predominantly on kidney CD cells. Most observations have shown that intracytoplasmic vesicles, whose limiting membranes contain water channels, move to, and fuse with, the apical plasma membrane after hormonal stimulation. The water channels are thereby inserted into the apical membrane. b. Internalization of IMP Aggregates under Decreasing Water Transport In either case, washing out ADH affects the fast reduction of water

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permeability of epithelia and the consequent disappearance of IMP aggregates on the apical membrane. Most authors believe that internalization of water channels with clathrin coated pits occurs after the ADH-action (Brown and Orci, 1983; Masur et al., 1984; Brown et al., 1985, 1988; Harris et al., 1986; Coleman et al., 1987; Erlij et al., 1989; Brown and Stow, 1996). An increase in the endocytic activity of epithelial cells under ADH effect has been described in the mammalian CDs and amphibian urinary bladder in both the apical and basolateral surfaces (Masur et al., 1971, 1981, 1984; Wade and DiScala, 1971; Gronowitz et al., 1979; Wade et al., 1981; Ding et al., 1985; Hays et al., 1985; Harris et al., 1986; Coleman et al., 1987). Moreover, Brown et al., (1988) have demonstrated that the ability to endocytose the horseradish peroxidase in the normal rat CD cell apical membrane is five to six times higher than in mutant rats unable to synthesize vasopressin. At the same time, the treatment of the mutant rats with exogenous vasopressin increases markedly the number of endocytic vesicles in the apical cytoplasm of principal cells of their CDs. The discovery of the apical membrane specialized domains with high water permeability in tight epithelia with stimulated water transport raises the question about their chemical nature and their origin. Recent studies have made good progress, revealing specialized proteins, aquaporins, that form the water channels.

C. Aquaporins—Proteins of Membrane Water Channels From extensive available material on aquaporin (AQP) permeability in transporting epithelia, this section will consider a few topics such as immunolocalization and distribution of AQPs in leaky and tight epithelia. In recent years, many excellent reviews on nomenclature, classification, immunolocalization, molecular structure, function, gene cloning, and sequencing of AQPs have appeared (Verkman, 1992; Agre et al., 1993, 1995; Brown and Sabolic´, 1993; Sabolic´ and Brown, 1994; Brown et al., 1995; Verbavatz, 1995; Agre and Nielsen, 1996; Brown and Stow, 1996; Knepper et al., 1996, 1997; Verkman et al., 1996; King and Agre, 1996; Wintour, 1997; Deen and van Os, 1998; Dibas et al., 1998; Echevaria and Hundain, 1998; Hamann et al., 1998; Ishibashi et al., 1998; Sasaki et al., 1998; Yamamoto and Sasaki, 1998). The exciting discovery of the first specific water channel protein, CHIP28, in a red cell membrane was made in 1992 at Agre’s laboratory (Preston et al., 1992). Since this discovery, the number of proteins known to have an essential role in membrane water transport, called aquaporins, has been rapidly increasing (Agre et al., 1995). At present, several AQPs are referred by multiple names. Thus, AQP1 was originally called CHIP28 (Preston et

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al., 1992), AQP2 ⫺ WCH-CD (Fushimi et al., 1993), and AQP3 ⫺ GLIP (Ma et al., 1996) (see Table I). Aquaporins are members of a large family of small hydrophobic channelforming proteins known as membrane integral proteins (MIPs). In the unglycosylated state, all AQPs are approximately 30 kDa (265–282 amino acids). Most AQPs are permeable only to water and impermeable to small organic and inorganic molecules, except for AQP3, 7, and 9 that are also permeable to water and glycerol. At present, ten different AQPs (AQP0–AQP9) are identified in a wide variety of secretory and resorptive epithelia (Deen and van Os, 1998). Some of them are colocated in the same tissue; others are distributed in the tissues with certain physiological functions. Different AQPs in the same tissue may work cooperatively. Phylogenetic analysis of AQPs has shown that two groups of AQPs originated from two divergent bacterial paralogues—a glycerol facilitator and aquaporin (Park and Saier, 1996). These authors have found that AQP3 and AQP7 belong to the glf group derived from the prototype of Escherichia coli—GlpF functioning as a water/glycerol facilitator. Note that a bacterial GlpF has no water channel function. So the glp group of eucaryotic cells has acquired the water channel function in their evolution (Sasaki et al., 1998). All other mammalian and plant AQPs are of another origin. Their prototype is the aquaporin from E. coli—Aqp-Z, a water channel protein. For the aqp group the exclusive selectivity for water is a characteristic feature (Sasaki et al., 1998). Studies of distribution of AQPs in cells were carried out successfully with the EM cryomethods: freeze-fracture (Rash et al., 1998; Eskandari et al., 1998), freeze-substitution (Nielsen et al., 1995, 1997). Good progress in studying AQPs was made when polyclonal antibodies were grown against them and a series of studies with immunolabeling appeared in the late 1990s (Agre et al., 1995; Abrami et al., 1997; Hamann et al., 1998; Kim et al., 1998). The AQP biophysical properties are examined in the oocyte and yeast expression systems and in the reconstituted proteoliposomes. In recent years, the best characterized expression system is the sec6-4yeast (Coury et al., 1998). Using methods of high-resolution cryoelectron crystallography, atomic force microscopy, and EM (freeze-fracture of reconstituted proteoliposomes and negative staining of purified protein), the three-dimensional structure of several AQPs (AQP1, AQP2, AQP3) was determined (Verbavatz et al., 1993; Jung et al., 1994; Walz et al., 1995, 1997; Bai et al., 1996; Cheng et al., 1997; Kuwahara et al., 1997; Eskandari et al., 1998). All the AQPs studied have a more or less similar structure. They are tetrameres and consist of six transmembrane domains connected by five loops. It has been shown that each tetramere contains four water pores. The pore radius

TABLE I AQP Distribution in Mammalian and Amphibian Tissues Aquaporins AQP0 (MIP26–major intrinsic protein 26; CHIP–channel intrinsic protein) AQP1 (CHIP28)

Animal Mouse Frog

Localization Lens fiber cells

Features and function

234

Important role in water and/or glycerol homeostasis; participates in maintenance of lens transparency Human rat LLC-PK Red cells; Kidney: proximal tubules, thin In water reabsorption in proximal tubules and (pig cells) MDCK descending limbs of Henle’s loops; choroid Henle’s loops; in the urine concentration up plexus; bile duct; capillary and venula to 3000 mOsM in mice and 1200 mOsM in endothelia; lacteals and lymphatics; retina human; a role in secretion of spinal fluid, reproductive fluids, aqueous humor (in retinal homeostasis), bile; in vascular permeability AQP2 (AQP-CD–aquaporin in Rat human Collecting duct (CD) principal cells (apical Vasopressin-regulated water channel; collecting duct; membrane and subapical vesicles) participates in reabsorption of water in kidney CD WCH-CD–water channel in collecting duct) AQP3 (GLIP–glucose intrinsic Human, rat Kidney CD principal cells (basolateral Permeable to water, urea and glycerol; protein) membrane); gastrointestinal tract; airway paricipates in water reabsorption in CD, in epithelium; eye conjunctiva; brain reabsorption of humor, of the secreted fluid ependymal cells; spleen in lung disease AQP4 (MIWC–mostly inactive Human, rat High expressed in glial and ependymal cells in With highest single channel water conductance water channel or mercury the brain, weakly—in the retina, airway among AQP 0-5; participates: in insensitive water channel) epithelium, gastrointestinal tract, kidney CD cerebrospinal fluid transport in brain, water principal cells (basolateral membrane) transport in kidney CD, aquous humor transport in the eye, airway hydration in the lung, in osmoreception in CNS and in volume regulation

AQP5

Rat

AQP6 AQP7

Human Rat

AQP8

Rat, mouse

AQP9

235

FA-CHIP (frog aquaporin–channel intrinsic protein) AQP-TB (aquaporin in toad bladder)

Frog

Toad

Secretory epithelium in upper airway and salivary gland alveolar epithelium (apical membrane), type I pneumocytes Kidney (uncertain localization) Highly expressed in late spermatids, maturing sperm; weakly—in kidney, heart Highly expressed in hepatocytes, pancreatic glandular lobes, testis; weakly—in colon, salivary gland; placenta, liver, heart Highly expressed in adipocytes, weakly—in heart, kidney, small intestine Urinary bladder, skin, brain, gall bladder, lung

Participates in the secretion of saliva, tears, and sputum (pulmonary fluid) A potential role in water transport Permeable to water, urea, glycerol; important role in spermatogenesis and fertilization Necessary for the secretion of pancreatic juice

Permeable to water, urea, glycerol; most gomologous to AQP3 and AQP7; 79% identity AQP1, 42%—AQP2 not vasopressin regulated

Urinary bladder, skin, brain, gall bladder, lung Not vasopressin regulated, functional homologue of AQP1 and FA-CHIP

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˚ ⬍ 2.1 A ˚ (Coury et al., 1998). With respect to is suggested to be ⬎ 1.6 A AQP1 and AQP2, they have a narrow pore that is highly selective for water. The results of studying the AQP1 and AQP2 functions suggest that the water pore of these channels has a barrier to the flow of charged species, such as protons (Coury et al., 1998). All currently known AQPs of mammals, plants, and bacteria have a great resemblance in their amino acid sequences, up to 20–40% identity (Reizer et al., 1993). All of them, except AQP4, are mercury-sensitive, i.e., mercurial sulfhydryl reagents inhibit the high water permeability of plasma membranes containing AQPs (Preston et al., 1992; Agre et al., 1995; Yang et al., 1996; Wintour, 1997). AQP4 has been shown to have no cysteine residue considered as a mercury-sensitive site near its putative aqueous pore (Preston et al., 1993; Verkman et al., 1996).

1. Leaky Epithelia Agre et al. (1993) discovered AQP1 in high water permeable renal proximal tubules and the descending thin limb. They suggested that AQP1 contributes to the countercurrent multiplier mechanism responsible for water conservation by the proximal nephron (Agre et al., 1993, 1995). Immune EM shows that AQP1 is abundant in the microvillar membrane. Of AQP1 95% is located in the apical membrane and only 5% of this protein is in vesicles and vacuoles (Preston et al., 1992; Nielsen et al., 1993b; Agre et al., 1995; Elkjær et al., 1995). Mutations in the AQP1 gene have no clinical abnormality. AQP1 is presumed to be a simple constitutive activating water pore inserted in the apical membrane by vesicular translocation (Marinelli et al., 1997). The dependence of the constitutive pathway of AQP1 proteins to the plasma membrane on microtubule integrity was demonstrated by treating proximal tubules with colchicine (Elkjær et al., 1995). The microtubule disruption prevents formation of endocytic invaginations and vacuoles, that is AQP1 recycling. Meanwhile, the membrane recycling is a fundamental biological feature necessary for the normal functioning of membrane proteins. Several AQPs (AQP1–AQP5) have been detected by immunocytochemical methods in various tissues of another water-transporting organ, the eye. The physiological importance of AQPs in maintaining lens transparency was demonstrated (Hamann et al., 1998; Kim et al., 1998). In addition, AQP1 expression was found to be abundant in capillary endothelium, in which it may contribute to vascular permeability (Nielsen et al., 1993a, 1993b).

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The most conflicting data on AQP distribution have been obtained in the small intestine. Studying expression of the mRNA of AQP1–AQP5 in the small intestine of infant and adult rats, Granquist with coauthors (1996) revealed that the proximal small intestine contains AQP3 and AQP4 but no AQP1, AQP2, and AQP5. AQP3 has been shown to undergo postnatal development. The expression of AQP3 increases dramatically in adult tissue as compared with the infant intestine. It is possible that the low expression of AQP3 in infant rats may account for the immaturity of their intestinal water transport. By molecular cloning of a novel human aquaporin, Kuriyama et al. (1997) have shown a weak expression of AQP9 in the small intestine. However, so far there is no immunocytochemical confirmation of distribution of AQPs in the small intestine epithelial cells.

2. Tight Epithelia The first data on protein composition of water permeable membrane domains in the apical membrane of the tight epithelium were obtained by Wade and co-authors (1984). They isolated new proteins appearing only during the vasopressin stimulation of water transport from the apical membranes of amphibian urinary bladder epitheliocytes and raised the antibodies against these proteins (Wade et al., 1984; Harris et al., 1986, 1987, 1988; Harris and Handler, 1988). Three proteins with molecular weights of 7, 15–17, and 55 kDa, which were absent in control bladders, were revealed. Dassouli et al. (1989) described two polypeptides with molecular weights of 76 and 14 kDa in the apical surface of ADH-treated amphibian urinary bladder epithelial cells. With immunofluorescent microscopy, these authors showed predominant localization of these proteins in apical membranes of epitheliocytes. Subsequently, in amphibian urinary bladder epithelia, by molecular biological techniques, freeze-substitution, and antibodies against AQP conjugated with colloidal gold, their own specific channel proteins were discovered. These are FA-CHIP in the frog (Abrami et al., 1994, 1995, 1996, 1997) and AQP-TB in the toad (Ma et al., 1996; Siner et al., 1996). Abrami and coauthors (1994, 1997) isolated and cloned the frog AQP and showed that it has 79% identity with rat AQP1 and only 42% identity with the kidney CD AQP2. It appeared that pAB against human red blood cell CHIP recognized and labeled FA-CHIP (Calamita et al., 1995). No difference in the labeling patterns was observed between resting and vasopressintreated bladders. Unlike AQP2 and AQP3 in kidney CD cells, FA-CHIP has not been found in the apical membranes of the frog urinary bladder epitheliocytes.

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FA-CHIP is highly expressed in endothelial and mesothelial cells of blood capillaries and is poorly expressed in epithelial and red blood cells. In the toad urinary bladder, an aquaporin homologue, the so-called AQPTB (AQP toad bladder) was revealed (Siner et al., 1996). Apart from the urinary bladder, this protein is expressed abundantly in the skin, lung, skeletal muscle, kidney, and brain. AQP-TB has been reported to increase during dehydration or chronic vasopressin stimulation. Cloning cDNA of a water channel protein from the toad urinary bladder, Ma et al. (1996) have shown that it has 76% identity with mammalian AQP1 and 88% identity with frog water channel FA-CHIP. These authors obtained data indicating that this protein is a functional homologue of AQP1 and FACHIP and is probably not vasopressin-regulated. FA-CHIP and AQP-TB are suggested to be constitutively delivered to the plasma membrane by vesicular transport. Boom et al. (1995) showed that the cycling of water channels in the toad urinary bladder could be controlled by one or more G-proteins. The ADH-regulated channel protein, AQP2, has not yet been revealed so far in the apical membranes of amphibian urinary bladder cells. With respect to another vasopressin-dependent osmoregulatory epithelium, it has been shown that mammalian kidney collecting duct epithelium cells contain three types of AQPs: AQP2 (Nielsen et al., 1993a, 1995, 1997; Agre et al., 1995; Yamamoto et al., 1995), AQP3 (Echevarria et al., 1994; Ishibashi et al., 1994, 1998; Sasaki et al., 1998), AQP4 (Frigeri et al., 1994). AQP2 has been revealed in apical vesicles and in the apical membranes of the CD principal cells. After the treatment of CD by vasopressin, AQP2 is especially abundant in the apical membrane (Nielsen et al., 1995; Sabolic´ et al., 1995; Yamamoto et al., 1995). The redistribution of AQP2 is accompanied by a vasopressin-induced increase in epithelial water permeability measured in the same tubules. The question whether these AQP2 correspond to the aggregates of IMPs described in the apical membrane of these cells remains to be answered. It is possible that this information can be obtained with the freeze-fracture labeling techniques, as in the case of AQP4 in the rat brain astrocytes and ependymocytes where 84% of labeling protein was present beneath square arrays (Rash et al., 1998). However, after the transfection of cDNA encoding AQP2 in the Chinese Hamster ovary cells, IMP aggregates have never been observed (van Hoek et al., 1998). Nielsen and coauthors (1995) showed AQP2 to be translocated from apical vesicles into the apical membrane by exocytosis. After withdrawal of vasopressin, AQP2 protein is internalized into the cytoplasm by endocytosis (Brown and Sabolic´, 1993; Marples et al., 1995; Nielsen et al., 1995; Katsura et al., 1997; Saito et al., 1997; Maric et al., 1998).

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Thus, the immunocytochemical approach has allowed confirming the shuttle hypothesis put forward by Wade et al. (1981) with respect to water permeable membrane domains. However, what organelles are involved in this process is not finally clear. In addition to AQP2, CD principal cells contain two more aquaporins— AQP3 and AQP4. These AQPs are located in basolateral membranes and rarely in the cytoplasmic vesicles (Frigeri et al., 1994; Ecelbarger et al., 1995; Yamamoto et al., 1995; Knepper et al., 1996, 1997). The colocalization of AQP3 and AQP4 could account for the high water permeability of the basolateral membranes of these cells. Although all these AQPs are colocalized in the same cells, it has been shown that characteristics, physiological roles, and response to vasopressin of these aquaporins are different (Sasaki et al., 1998). The expression level (long-term regulation) of all these AQPs is increased by vasopressin. However, contrary to AQP3 and AQP4, which are constitutively inserted into the apical membrane and whose subcellular localization is not changed during vasopressin action (Ecelbarger et al., 1995; Knepper et al., 1997), the translocation of intracellular vesicles bearing AQP2 (shortterm regulation) to the apical membrane is stimulated by vasopressin. After binding of vasopressin to the V2 receptor, the trafficking AQP2 is triggered by cAMP. Sorting is dependent on the PKA-mediated phosphorylation of AQP2. Subsequently, transport vesicles containing the AQP2 water channels translocate to, and fuse with, the apical membrane (Fushimi et al., 1997; Yasui et al., 1997; Mulders et al., 1998). It has been demonstrated that the process of AQP2 targeting is dependent on the microtubule integrity. The data obtained regarding the inhibition of the hydroosmotic response of the CD principal cells by colchicine treatment could be explained by the dependence of AQP2 targeting to the apical membrane on microtubule integrity (Taylor et al., 1974; Sabolic et al., 1995). Marples et al. (1998) demonstrated by immunocytochemical methods the colocalization of dynein and dynactin with AQP2 in intracellular vesicles. This is consistent with the viewpoint that the microtubule-associated motor protein dynein and the associated dynactin complex participate in the vesicle transport (Marples et al., 1998). The molecular machinery for exocytotic insertion and endocytotic retrieval of AQP2-bearing vesicles is largely unknown, but recent studies suggest that this process is similar to the regulated exocytosis of synaptic vesicles (Brown and Stow, 1996; Deen and van Os, 1998). AQP2 is a single water channel whose physiological functions have been clearly demonstrated in pathological states. Thus, mutations in the human AQP2 cause nephrogenic diabetes insipidus (Deen et al., 1994; Nielsen et al., 1997; Mulders et al., 1998). Down-expression of AQP2 is also observed in a vasopressin-deficient strain of rats, Brattleboro rats, which exhibits

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central diabetes insipidus (Digiovanni et al., 1994; Sabolic´ et al., 1995). Administration of vasopressin to the rats increases the amount of AQP2 proteins in apical membranes of CD principal cells. This is consistent with the data that show an increase in the aggregates with IMPs in them (Brown et al., 1980, 1985, 1988; Harmanci et al., 1980). On the contrary, the increased expression of AQP2 protein is revealed in rats with congestive heart failure (Nielsen et al., 1997). The disregulation is associated with an increase in CD water permeability and in water absorption from CDs. Thus, studies on the mammalian kidney function also confirmed the fact that vasopressin regulation of AQP2 is important to the regulation of the body water balance (Sasaki et al., 1998). The question whether the vasopressin-regulated water channel protein AQP2 is present in the frog urinary bladder epithelium is an important fundamental question of the membrane physiology.

V. Participation of the Vacuolar System in the Cell Response on Induced Water Transport The vacuolar system of eucaryotic cells consists of such organelles as the ER, Golgi complex (Gc), endosomes, and lysosomes occupying stable positions within the cytoplasm.

A. Changes of Epithelial Cell Structure during Stimulated Water Transport Vasopressin stimulation of osmotic flows across tight epithelia produces remarkable changes of cell structure. It has been shown that an increase in water absorption in the amphibian urinary bladder and mammalian CD is accompanied by the swelling of granular and principal cells, respectively (Fig. 4b) (Bentley, 1958; Peachy and Rasmussen, 1961; Natochin, 1963; Mashansky et al., 1966; Masur et al., 1971; Tisher et al., 1971; Wade and DiScala, 1971; Eggena, 1977; Davis et al., 1982; Snigirevskaya et al., 1982). The cell volume is increased 1.5–2 times (Peachy and Rasmussen, 1961). X-ray microanalysis of toad urinary bladder epithelial cells has revealed that ADH action decreases the general contents of Na⫹, K⫹, and Cl⫺ ions (Rick et al., 1978; Rick and DiBona, 1987) providing the cytoplasm dilution. Intercellular spaces in the basolateral region of epithelia are diluted (Bentley, 1958; Pak Poy and Bentley, 1960; Grantham et al., 1969; Grantham, 1970; Komissarchik et al., 1982; Snigirevskaya et al., 1982). Cell apical parts are evaginated into the lumen, and preferentially ridgelike microvilli are

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replaced by the finger-like ones (Davis et al., 1981). The use of an extracellular marker, ferritin, reveals small endocytic vesicles in the apical cell cytoplasm of both resting and stimulating bladders (Komissarchik and Snigirevskaya, 1990). According to the data in the literature, the endocytosis is essentially increased during the ADH-action (Wade and DiScala, 1971; Wade et al., 1981; Brown and Orci, 1983; Masur et al., 1984). Intracellular structures of epithelial cells also undergo significant alterations. The most essential changes have been observed in the cell vacuolar system. Studying changes of the vacuolar system of the frog urinary bladder epithelium under ADH action in the presence of an osmotic gradient, we have revealed the appearance of giant vacuoles originated from Golgi cisternae in the granular cells (Komissarchik et al., 1982, 1985; Snigirevskaya et al., 1982; Snigirevskaya, 1983). Similar large vacuoles were demonstrated only in a few studies during maximal transepithelial water flows in tight epithelia of the amphibian urinary bladder and mammalian CDs (Ganote et al., 1968; Kirk et al., 1984a, 1984b; DiBona et al., 1985; Shinovara et al., 1989). The structure and behavior of these vacuoles will be considered in more detail in Section V.D.3.

B. The Structure and Function of Golgi Complex 1. Epithelial Cells During the past few years, great strides have been made toward understanding the Gc structure–function relationships by applying new approaches. Apart from EM cryomethods and immunolabeling, green fluorescent protein technology has made it possible to visualize Gc in the living cell (Chalfie et al., 1994). The detailed pathway of cargo and Golgi resident proteins within Gc and the dynamics of microtubules associated with Golgi membranes can be followed with the aid of video movies (Presley et al., 1997, 1998). Its unique structure and function have drawn the attention of many authors. Rather, many reports dealing with the Gc role in the intracellular transport of proteins, its membrane dynamics, and its three-dimensional reconstructions have appeared for recent years (Farquhar and Palade, 1981, 1998; Rambourg et al., 1981; Rodriguez-Boulan and Zurzolo, 1993; Rothman, 1994; Rothman and Wieland, 1996; Keller and Simons, 1997; Mironov et al., 1997, 1998; Presley et al., 1997, 1998; Bloom and Goldstein, 1998; Kaiser and Ferro-Novik, 1998). The important role of Gc in cell functioning is well known. The Gc is involved in secretory membrane traffic and sorting. Newly synthesized proteins and lipids are covalently modified in the Gc, packed, and delivered to secretory granules, lysosomes, or plasma membrane.

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The Gc consists of stacked membrane cisternae and associated tubules and vesicles and is a typical polarized organelle. The cis-surface is associated with ER, and the trans-surface is where the trans-Golgi network (TGN) is located (Farquhar and Palade, 1981; Kreis, 1990; Rambourg et al., 1981). The medial compartments are located between them. At the trans-surface, condensing vacuoles, secretory granules, and lysosomes are observed. To the morphological polarity, functional polarity also corresponds. Each compartment has presumptive markers used as guidposts (Farquhar and Palade, 1998). However, many aspects of intra-Golgi transport remain unclear so far. Proposed models of mechanisms of moving cargo through Golgi stacks include the maturation or cisternal progression model (Hicks, 1966; Severs and Hicks, 1979) and the vesicular transport/stationary cisterna model (Farquhar and Palade, 1981; Rothman, 1994; Rothman and Wieland, 1996), and each is controversial. Therefore, existing models have been reexamined in the last few years, and new modified models have appeared as hybrids between the maturation and vesicular transport models (Farquhar and Palade, 1998; Pelham, 1998; Warren and Malhotra, 1998). Mironov and his coworkers (1997, 1998) in their synthetic model underline a special importance in the cargo transfer through the tubular elements of Gc.

2. Rat Enterocytes and Granular Cells in the Frog Urinary Bladder The rat enterocytes and frog urinary bladder granular cells have welldeveloped Gc consisting of four or five cisterna stacks and its derivatives— lysosomes, small smooth and coated vesicles, and tubules (Snigirevskaya et al., 1982; Snigirevskaya, 1983; Snigirevskaya and Komissarchik, 1987, 1995). In general, the Gc structure of both cell types is consistent with the available data on its structure in secreting cells (Farquhar and Palade, 1981; Rambourg et al., 1981; Farquhar, 1982, 1985; Thyberg and Moskalewsky, 1985). But cisternae in enterocytes are always dilated, whereas cisternae in granular cells are very thin, not more 20 nm wide (Snigirevskaya and Komissarchik, 1987, 1995; Snigirevskaya et al., 1997). With respect to other cell types, in the granular cells there is an association of Golgi membranes with microtubules radiating from the centrioles (Maurice et al., 1983; Pavelka and Ellinger, 1983; Tassin et al., 1985; Veselov and Snigirevskaya, 1985; Ho et al., 1989; Kreis, 1990). Agents that change distribution of microtubules disturb the integrity and location of the Gc. Thus, treatment of cells with nocodazole leads to fragmentation of the Gc into distinct elements dispersed throughout the cytoplasm (Ho et al., 1989; Kreis, 1990; Cole et al., 1996).

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In addition, actin microfilaments are essential for the integrity and positioning of Gc inside the cell (Valderrama et al., 1998). It has been suggested that, in the granular cells, Gc plays a significant role in the processing of secretory product of specific granules crowded inside the cytoplasm (Fig. 5a).

C. Behavior of Golgi Complex during Induced Water Transport One of the most reactive organelles in the response to the stimulated transepthelial water and fluid transport is the Gc. 1. The Golgi Complex in Rat Enterocytes during Increased Transepithelial Fluid Transport During glucose load of the proximal small intestine loop, we observed significant dilations of all Golgi cisternae (Figs. 9a–9c) (Snigirevskaya et al., 1997), which indicates some dilution of the enterocyte cytoplasm during isoosmotic fluid transport across the epithelium. It has been suggested that the Gc has the ability to absorb an excess of water from the cytoplasm, thereby saving the cell from osmotic shock and performing an osmoregulatory function in the cell. 2. Changes during Stimulated Water Transport across Frog Urinary Bladder Epithelium Analysis of Gc ultrastructure of granular cells in the frog urinary bladders in different functional states has revealed a high reactivity of this organelle in the response of ADH action (Snigirevskaya and Komissarchik, 1988, 1995). On the basis of the observations of Gc ultrastructure, we can present the following dynamics of its structural changes during ADH stimulation of water flows across epithelium (Figs. 10a, 10b, and 11). Under low basal transepithelial water transport, Gc is characterized by an extremely narrow intracisternal space—15–20 nm (Fig. 10a). The insignificant water flow induced by the brief ADH treatment of the bladders (5 min) is accompanied by both the fragmentation of the complex on individual stacks and the small dilations of cis-cisternae (Fig. 10b). The fragmentation of the Golgi complex is supposed to be the result of a disruption of microtubules determining its integrity under the conditions of diluted cytoplasm (Snigirevskaya and Komissarchik, 1987, 1995). Scattering Golgi elements during microtubule disruption has been described in the literature (Cole and LippincottSchwartz, 1995; Yang and Storrie, 1998). Under the maximal ADH effect

FIG. 9 Golgi complex of the rat enterocytes. (A) The control enterocyte. Some cisternae of Gc are dilated, whereas the rest of the cisternae are narrow. (B) Freeze-fracture replica of the control enterocyte cytoplasm containing parallel Gc cisternae (arrows). At the left the nucleus (N) is seen. (C) After glucose load, all cisternae of Gc are dilated. Bar ⫽ 0.25 애m. (Reproduced from Snigirevskaya et al., 1997, with permission).

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FIG. 10 Golgi complex of the frog urinary bladder. (A) In the control urinary bladder, the Golgi cisternae are flat and narrow (Reproduced from Snigirevskaya and Komissarchik, 1995, with permission). (B) After short action of ADH (5 min), a small dilation of cis-cisterna is seen. Bar ⫽ 0.5 애m.

on the frog urinary bladder epithelium, the giant vacuoles appear in the granular cell cytoplasm (Figs. 11, and 12a, 12b). As mentioned in Section IV.E, Golgi derivatives (specific granules) migrate to the apical surface and fuse with the apical membrane during ADH action. We proposed a regulated process of inserting water channel proteins into the apical membrane in response to the ADH stimulatory action. Whether granules or other cytoplasmic structures contain aquaporins should be clarified in the future. 3. Specific Vacuolation during Maximal Water Flows across Frog Urinary Bladder Epithelium In the current literature only a few studies describe the presence of large vacuoles during ADH stimulation of water flow in the principal cells of another tight epithelium, the mammalian epithelium of CDs, both at the EM level (Ganote et al., 1968) and in phase-contrast observations (Kirk et al., 1984a, 1984b; DiBona et al., 1985). The authors previously cited think

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FIG. 11 Schematic drawing of dynamics of the Gc structure changes in tight (A,B,C,D) and leaky (B,C) epithelia during stimulation of fluid transport across epithelia. (A) In the tight epithelium with very low water permeability, the Gc stack consists of flat and narrow parallel cisternae. (B) Golgi cisternae are slightly dilated in the control rat enterocytes and in the granular cells during a weak increase of water transport. (C) During water transport in tight epithelia and isotonic fluid transport in leaky epithelia, the cis-cisternae of Gc are dilated. (D) During the large water flow stimulated by ADH, the cis-cisternae of Gc form giant vacuoles. Microfilaments and microtubules are associated with the vacuole membrane.

that an insignificant part of water is transported with the aid of the large vacuoles. In granular cells of the toad urinary bladder epithelium, large vacuoles were revealed during inhibition of ADH-induced water transport by Cytochalasin B (Davis et al., 1974a, 1974b; Davis and Goodman, 1986; Davis and Finn, 1987). Davis et al. consider these vacuoles to be a result of the disruption of microfilaments associated with basolateral membrane and subsequent cell alteration. However, we think that the disruption of microfilaments by Cytochalasin B stops the migration of the vacuoles along the cytoplasm; therefore, the vacuoles can be revealed in this case in ultrathin sections more often. Note that giant vacuoles are preserved only by

FIG. 12 During the maximal increase of water transport across the frog urinary bladder, the giant vacuoles are formed in granular cells. (A) Association of the giant vacuole with other elements of Gc. Bar ⫽ 0.5 애m. (B) The vacuole revealed in the granular cell in the frozen ‘‘hemithin’’ section. (C) Distribution of K⫹ in granular cells after ADH stimulation of water transport across the bladder wall. K⫹ concentration sharply falls inside the vacuole (X-ray microanalysis). Bar ⫽ 5 애m. (Part A reproduced from Snigirevskaya and Komissarchik, 1995, with permission; Parts B and C reproduced from Snigirevskaya, 1990, with permission).

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gentle fixation methods so the use of standard EM techniques does not always allow the giant vacuoles to be discovered. By the freeze-substitution method and osmium fixation, the giant vacuoles are revealed in granular cells only in the presence of an osmotic gradient. Besides, they are absent when epithelial cells are swollen as a result of bathing by hypotonic media from the serosal side of the epithelium (Snigirevskaya et al., 1982; Komissarchik et al., 1985). Some components of Gc (e.g., cisternae, small smooth and coated vesicles, electron-dense canaliculi, as well as cytoskeletal elements—20- and 40-nm microtubules and microfilaments) are located in the close vicinity of these giant vacuoles (Komissarchik et al., 1985). The vacuole membrane IMP distribution revealed with the freeze-fracture method is similar to that of the Gc cis-cisterna membrane (Snigirevskaya and Komissarchik, 1987). The striking feature of the cell ultrastructure of stimulated urinary bladders is the appearance of a great number of ‘‘thick’’ microtubules in the apical cytoplasm of granular cells and around the giant vacuoles (Fig. 13) (Snigirevskaya and Komissarchik, 1993). The localization of these vacuoles in the vicinity of nuclei, the association of the vacuoles with Gc cisternae and small vesicles, microtubules, and microfilaments, and the distribution of IMPs in their membranes indicate that the vacuoles are derivatives of Golgi cis-cisterna. The application of fillipin has shown that the vacuole membranes are enriched in cholesterol (Kever et al., 1988), and this may account for the peculiar water permeability features of vacuole membranes. The associations of electron-dense canaliculi with vacuole reservoir can sometimes be observed.

FIG. 13 Immunocytochemical confirmation of the tubulin nature of the ‘‘thick’’ microtubules. Arrows indicate the colloidal gold particles on the MT (freeze-substitution). Bar ⫽ 0.25 애m.

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4. The Homology between the Granular Cell Giant Vacuoles and Protozoan Contractile Vacuoles Comparison of electron micrographs of the giant vacuoles of granular cells and contractile vacuoles of Paramecium has demonstrated a prominent morphological similarity of both vacuolar systems (Snigirevskaya, 1983). The connection of electron-dense canaliculi with the vacuole reservoir in granular cells is similar to the connections of spongium canaliculi with vacuoles in the protozoan contractile vacuole complex (McKanna, 1976). As in the contractile vacuole complex, numerous microfilaments and microtubules are associated with the giant vacuoles. An electron microprobe analysis has revealed a marked decrease in contents of potassium ions inside the giant vacuoles, whereas the potassium ion concentration in the cytoplasm does not differ from the normal one, 110–120 mM (Figs. 12b and 12c) (Snigirevskaya and Komissarchik, 1984; Rick et al., 1978). These results also agree with similar data on the protozoan contractile vacuole (McKanna, 1976). The preceding results have allowed us to put forward a hypothesis about the morpho-functional homology of giant vacuoles in granular cells and the contractile vacuoles of Protozoa (Snigirevskaya and Komissrachik, 1984; Snigirevskaya, 1983). According to this hypothesis, the giant vacuoles (‘‘contractile’’ vacuoles) are involved in cell osmoregulation and water transport across the granular cells from their apical surface to their basolateral surface. The ‘‘thick’’ microtubules and microfilaments are proposed to participate in the giant vacuole migration along the cytoplasm (Snigirevskaya and Komissarchik, 1993). The origin of the giant vacuoles from Golgi cis-cisterna confirms the point of view of Nassonov (1924) about the functional homology between protozoan contractile vacuole and Golgi complex.

5. Evolutionary Considerations of Giant Vacuoles The discovery of giant vacuoles with features of contractile vacuoles in the frog urinary bladder epithelial cells is of great interest with respect to the evolution of mechanisms of intracellular osmoregulation (Snigirevskaya, 1983, 1990). The contractile vacuoles were previously thought to be present only in cells of primitive organisms lacking osmoregulatory organs (e.g., fresh water Protozoa, Spongia, and Algae at some life stages—without cell walls). These organisms which live in hypotonic media avoid lysis caused by the contractile vacuole that regulates the cell volume. During evolution, cells of organs and tissues of multicellular organisms have acquired the ability to live in isoosmotic surroundings, except for tight epithelia lining cavities with hypotonic contents. Tight epithelia are characterized by water impermeable tight junctions and apical cell membranes. Under the effect

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of ADH, tight epithelia become highly water permeable. In circumstances of high and fast transepithelial water flows, the volume seems to be regulated not only by the sodium pump but also by giant vacuoles playing the role of contractile vacuoles. This indicates a more widespread distribution of contractile vacuoles than was thought previously.

VI. The Role of Cytoskeleton in Transcellular Water Transport One of the important problems in cell biology is the regulation of the behavior of the cytoskeleton during cell functioning. Cytoskeletal elements play a crucial role in cell functioning, enabling cells to respond effectively to changing membrane and secretory needs (Cohen and Smith, 1985; Ausiello et al., 1987; Hugon et al., 1989; Cole and Lippincott-Schwartz, 1995). Understanding the mechanisms underlying cytoskeletal changes can be achieved using combined approaches of biochemistry, biophysics, immune EM, and molecular biology. Cytoskeleton is composed of three main components—actin filaments (microfilaments), intermediate filaments, and microtubules, which closely interrelate structurally and functionally in cells. Recent investigations have shown that microfilaments and microtubules are involved in the organelle transport and membrane trafficking in cells, both containing microtubule and actin-dependent motors (Fath et al., 1994; Cole and Lippincott-Schwartz, 1995; Langford, 1995). They are highly dynamic structures able to assemble (polymerize) and disassemble (depolymerize) very rapidly.

A. Participation of Cortical Actin in Docking and Inserting of Water Channel Proteins into Apical Membrane An ADH-produced increase in osmotic water permeability is believed to be accompanied by cytoskeleton reorganization (Taylor et al., 1978; Pearl and Taylor, 1983; Sasaki et al., 1984; Komissarchik et al., 1985, 1996; Snigirevskaya and Komissarchik, 1987, 1993). Using confocal and electron microscopes, it has been established that vasopressin depolymerizes F-actin in the apical region of amphibian urinary bladder granular cells (Ding et al., 1985; Komissarchik et al., 1992, 1996, 1998). Immune EM of actin filament distribution revealed a 2.5- to 4-fold decrease in antiactin label density in the apical area after stimulation of water transport by both vasopressin and repeated changes of serosal Ringer solution (Komissarchik et al., 1996,

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1998). At the same time, there were no significant changes in immunogold labeling over the microvillar actin and in the deep layers of the cytoplasm. This indicates functional heterogeneity of the actin filament pools in the cell. In ADH-treated epithelium, density of the actin labeling of cortical actin is reduced, and specific granules might come close to, and fuse with, the apical membrane (Figs. 5c and 5d). The cortical actin layer described in different epithelial cells is largely considered an apparatus that limits and controls exocytosis (Orci et al., 1972; Sontag et al., 1988). In regulating cytoskeletal function the alterations in the cytosolic calcium have been shown to play the essential role (Margulis, 1983; Mia et al., 1983, 1987, 1988; Stevenson and Begg, 1994). As mentioned earlier, during ADHaction, calcium is released from intracellular storage sites and serves as one of the important messengers in the cell hydroosmotic response. Both in frog and toad urinary bladders, such storage sites might be specific granules of granular cells, which accumulate calcium at high concentrations (Schakhmatova et al., 1982; Davis et al., 1987). During the fusion of granule membranes and the apical membrane, the granule Ca ion contents remain in the cytoplasm and can promote actin depolymerization. Ausiello et al., (1984) identified a calcium-activated protein, villin, that binds actin at micromolar calcium concentrations and causes actin filament dissociation. It cannot be excluded that gelsolin also might be involved in this process (Ausiello et al., 1984; Ausiello and Hartwig, 1985). Therefore, depolymerization of apical submembraneous F-actin is an obligatory condition for increase in water permeability, whereas the depolymerization of all cytoplasmic F-actin inhibits this cell response (Komissarchik et al., 1998).

B. The Role of Microtubules in the Hydroosmotic Cell Response on ADH Microtubules are found in all eucaryotic differentiated cell types and perform a wide variety of cell functions. They organize the cytoplasm, position the nucleus and organelles, and serve as the principal structural elements of flagella and cilia (Cooper et al., 1988; Klausner et al., 1992; Cole and Lippincott-Schwartz, 1995). One of the important functions of microtubules is their participation in mitosis. Concerning osmoregulatory epithelia, earlier studies limited the role of cytoskeleton exlusively to the maintenance of the cell shape (Dousa and Valtin, 1976). The first proof that microtubules and microfilaments are involved in vasopressin-induced transcellular water transport was obtained in the 1970s by physiological studies of Taylor (1977), Taylor et al., (1974, 1978), and Reaven et al., (1978). They showed that such antimitotic agents as colchicine, podophyllotoxin, and vinca alca-

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loids inhibit the effect of vasopressin on the transcellular water movement but not on sodium or urea transport in the toad urinary bladder. However, morphological studies utilizing conventional preparations for EM have failed to demonstrate a precise role of cytoskeletal elements in transcellular water transport (Reaven et al., 1978). 1. Microtubules in Granular Cells of Untreated Frog Urinary Bladder (at Low Water Permeability) Granular cells of the unstimulated urinary bladder with a great number of granules of various inner structure contain usual 20-nm microtubules. Microtubules are sparsely and apparently randomly distributed in the cytoplasm, concentrating around the centrioles and the Golgi complex (Reaven et al., 1978; Snigirevskaya and Komissarchik, 1987, 1993; Gorshkov and Komissarchik, 1998). 2. Microtubules in Granular Cells of the Vasopressin-Stimulated Frog Urinary Bladder (at High Water Permeability) Studying the peculiarities of the distribution of cellular cytoskeletal elements in the frog urinary bladder epithelium, we discovered that their preservation was significantly improved by using fixation with glutaraldehyde without osmium or by preparing the material with the freezesubstitution method (Snigirevskaya and Komissarchik, 1987). Embedding the material in acrylic resins has allowed the immunocytochemical identification of tubulin (Snigirevskaya and Komissarchik, 1991, 1993; Komissarchik et al., 1992; Komissarchik et al., 1998). As mentioned in Section C.2, after vasopressin stimulation of water transport, numerous changes occur in the cytoplasm of urinary bladder granular cells. The cytoplasm of granular cells is swollen, and the intercellular spaces are enlarged. Most granules are partially fused with the apical membrane. Giant vacuoles appear in the cells, and they are usually located in the central area in the cells and associated with Gc and cytoskeletal elements. With conventional chemical fixation, only 20-nm microtubules are seen to be associated with centrioles and the limiting membrane of giant vacuoles. According to the data obtained by Taylor et al., using conventional preparations, only an insignificant increase of the number of microtubules, about 30%, was revealed (Reaven et al., 1978). 3. The Appearance of ‘‘Thick’’ Microtubules in the Granular Cells of ADH-Stimulated Frog Urinary Bladder Epithelium A dramatic difference in the cytoskeleton of the granular cell is seen after preparation of the bladder by the freeze-substitution method or after fixa-

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tion using glutaraldehyde alone. In addition to the usual 20-nm microtubules, actin filaments, and intermediate filaments, ‘‘thick’’ microtubules (35–40 nm in diameter) can be observed in the cytoplasm of these cells (Fig. 13). Under high magnification, the subunit organization of ‘‘thick’’ microtubules was revealed; they consist of 17 subunits, 7.5 nm each (Snigirevskaya and Komissarchik, 1993). Microtubules with a similar structure, called accessory tubules, were described in the tails of spermatozoa of the spermatids of some stick insects (Afzelius, 1988). In the granular cells of the stimulated bladder, the ‘‘thick’’ microtubules are much more abundant than the usual 20- to 25-nm ones. Most ‘‘thick’’ microtubules are located in the apical portions of the cells, and they are often concentrated around the giant vacuoles. ‘‘Thick’’ microtubules also are seen in the membrane-limited granules located in the apical cytoplasm, which are revealed with conventional methods as specific granules with structured contents. In addition to single ‘‘thick’’ microtubules, some form clusters of four to six microtubules. The ‘‘thick’’ microtubules are packed very tightly in bundles forming a structure reminiscent of paracrystals observed in some cells after microtubule poison treatment (Dustin, 1984). Some of the ‘‘thick’’ microtubules are decorated by electron-dense globular particles located regularly along the microtubule length, 30 nm from each other (Snigirevskaya and Komissarchik, 1993). The mean diameter of these globules is 21–23 nm. These globular particles seem to represent microtubule-associated proteins (Murphy and Tilney, 1974; Dustin, 1984); however, many questions remain unanswered concerning the newly found ‘‘thick’’ microtubules and their relationship to the 20- to 25-nm microtubules. It is unclear, for example, whether both 20- to 25-nm and 35- to 40-nm microtubules consist of the same or different isotypes of tubulin. Thus, the application of several EM techniques for the analysis of the cytoskeleton of the frog urinary bladder has allowed detection of significant differences in the number, distribution, and structure of microtubules in the granular cells under conditions of low and high water permeability. 4. Tubulin Nature of ‘‘Thick’’ Microtubules Single ‘‘thick’’ microtubules, as well as those organized in clusters, are positively labeled for tubulin by antitubulin antibodies conjugated with colloidal gold (Fig. 13). The paracrystalline formations were also labeled by the gold particles (not shown). On the basis of ultrastructural study and tubulin immunolabeling, we suggest that, in the bladder granular cells, tubulin exists in different molecular states—monomeric, polymeric, and crystalline. The assembly of microtubules is likely to be triggered by the action of vasopressin. To produce such a large number of microtubules, the cells should contain a pool of

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monomeric tubulin. As Mandelkow and Mandelkow (1995) indicate, microtubules ‘‘must be nucleated at the right time and in the right place. This normally happens at the centrosome.’’ Whether the centrosomes are the centers of organization of the ‘‘thick’’ microtubules (COMT) in the granular cells during ADH induction of water transport is not clear. 5. Specific Granules as the Depot of Monomeric Tubulin and Ca2⫹ One of the mechanisms that could prevent spontaneous polymerization of tubulin in the absence of the hormonal signal is the storage of tubulin inside some intracellular compartments. The immuno-positive granules possibly serve as a storage for monomere tubulin whose polymerization is triggered by vasopressin activation. It has been shown that specific granules of granular cells of the bladder epithelium accumulate calcium ions (Schakhmatova et al., 1982; Davis et al., 1987). Calcium ions in large concentration are known to inhibit polymerization of tubulin (Solomon, 1982). Vasopressin could induce a release of calcium from the granules by the inositoltriphosphate pathway and thus trigger the assembly of microtubules and depolymerization of cortical actin filaments (Snigirevskaya and Komissarchik, 1993; Komissarchik et al., 1998). It can be ruled out that some specific granules in these cells are the numerous COMTs. 6. Participation of ‘‘Thick’’ Microtubules in the Migration of Giant Vacuoles through Cells A marked increase in the number of microtubules in granular cells of the frog urinary bladder after treatment with vasopressin suggests their involvement in transcellular water transport. This suggestion is confirmed by physiological data on the inhibition of vasopressin-induced fluxes by antimitotic agents (Taylor et al., 1978). We believe that ‘‘thick’’ microtubules participate in the migration of large vacuoles containing hypotonic fluid through the cytoplasm of the granular cells (Snigirevskaya and Komissarchik, 1991, 1993). As mentioned earlier, it is possible that these vacuoles with hypotonic contents are involved in the transcellular transport of water from the lumen into the blood. The role of microtubules in the movements of intracellular organelles has been shown for derivatives of the vacuolar system, particularly for extensions of ER membranes, early endosomes, lysosomes, and the Gc (Cooper et al., 1988; Cole and LippincottSchwartz, 1995). Two hypotheses are in the literature to explain the role of microtubules in transcellular water transport across the tight epithelia. According to one, the microtubules as well as microfilaments are involved in the process of

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translocation of tubular elements (aggrephores) toward the apical membrane (Kachadorian et al., 1979; Mu¨ller et al., 1981; Parisi et al., 1985). The second hypothesis suggests that microtubules take part in the fusion of aggrephores with the apical plasma membrane and in the migration of aggregates of IMPs (high water permeability membrane domains) in the plasma membrane (de Sousa, 1985, 1986; Mu¨ller and Kachadorian, 1985; Pearl and Taylor, 1985). Recent studies have shown the role of microtubules in membrane recycling and polarized distribution of Aquaporin-1 in epithelial cells (Elkjær et al., 1995). This question was discussed in Section IV.E. According to the data obtained in our earlier work, ‘‘thick’’ microtubules participate in the vacuole migration through the cytoplasm (i.e., in the water transport across the cell).

VII. Conclusions Water transport across different epithelia is known to be coupled with such important functions of an organism as osmosis, the nervous system, temperature regulation, digestion, and breathing. Studies carried out for the past three decades have greatly increased our knowledge of the mechanisms responsible for transmembrane, transcellular, and paracellular water transport. The combination of physiological and morphological methods, particularly different ultrastructural approaches, have provided strong evidence for the idea that the main pathway for water, both across the leaky and tight epithelia, is the transcellular one. However, in different leaky epithelia water transfer through the TJ region can occur. Substantial progress has been achieved in identifying apical membrane domains with high water permeability in tight epithelium cells after water transport stimulation by ADH or washing out of autacoids. The latter is suggested to wash out prostaglandin E2 that stabilizes the basal water permeability of the apical membrane. A further achievment of the early 1990s was the discovery of specialized water channel proteins, aquaporins. They have been identified in most bacterial, plant, and animal plasma membranes. However, a series of questions are not yet completely answered. For instance, the aquaporin presence in leaky epithelium plasma membranes cannot fully explain how small driving forces can perform such a prominent water flow across the epithelial layer. What is the participation of such diversity of different aquaporins? What is the participation of Gc in processing, sorting, and docking aquaporins into the plasma membrane? Does Gc participate in intracellular osmoregulation in leaky and tight epithelia, and, if so, what is its functional role? Is there a possibility that numerous centers of microtubule organization exist in the frog urinary bladder granular cells? Do the

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usual microtubules and ‘‘thick’’ microtubules belong to the same or to different tubulin classes? These and other problems can be solved by applying up-to-date approaches from cell and molecular biology.

Acknowledgments The authors thank Prof. L. Z. Pevzner and Prof. J. Holt for their help in preparing the manuscript and Mr. G. V. Sabinin for providing the photographs. This work was supported by projects #97-04-48901 and #99-04-49554 of the Russian Foundation of Fundamental Research.

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Membrane Trafficking and Processing in Paramecium Richard D. Allen* and Agnes K. Fok† Pacific Biomedical Research Center*, Department of Microbiology* and Biology Program†, University of Hawaii, Manoa

Cellular membranes are made in a cell’s biosynthetic pathway and are composed of similar biochemical constituents. Nevertheless, they become differentiated as membrane components are sorted into different membrane-limited compartments. We summarize the morphological and immunological similarities and differences seen in the membranes of the various interacting compartments in the single-celled organism, Paramecium. Besides the biosynthetic pathway, membranes of the regulated secretory pathway, endocytic pathway, and phagocytic pathway are highlighted. Paramecium is a multipolarized cell in the sense that several different pools of membrane-limited compartments are targeted for exocytosis at very specific sites at the cell surface. Thus, the method used by this cell to sort and package its membrane subunits into different compartments, the processes used to transport these compartments to specific locations at the plasma membrane and to other intracellular fusion sites, the processes of membrane retrieval, and the processes of membrane docking and fusion are reviewed. Paramecium has provided an excellent model for studying the complexities of membrane trafficking in one cell using both morphological and immunocytochemical techniques. This cell also promises to be a useful model for studying aspects of the molecular biology of membrane sorting, retrieval, transport, and fusion. KEY WORDS: Acidosomes, Endocytic membranes, Membrane biosynthetic system, Membrane replacement, Membrane trafficking, Phagosomal–lysosomal membranes, Quick-Freeze deep-etch, Secretory membranes. 䊚 2000 Academic Press.

I. Introduction Intracellular membrane trafficking and, more generally, protein trafficking have received widespread attention during the past decade. Several reviews International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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and symposia have focused on various aspects of this theme (e.g., Hernandez et al., 1996; Mellman, 1996; Pfeffer, 1996; Teasdale and Jackson, 1996; Mironov et al., 1997; Aroeti et al., 1998). Several reviews deal with the mechanisms behind the formation, targeting, docking and fusion of membrane vesicles. Others deal with vesicle membrane trafficking within a particular cell, tissue or organ [e.g., the nephron (Brown and Stow, 1996) or the trypanosome cell (Clayton et al., 1995)]. In the present review, we will focus on the total membrane trafficking, as far as it is known, within one free-living single-celled organism, (i.e., the protozoan Paramecium) and on the processing of the membrane as the membrane is modified, retrieved, or recycled. This cell embodies all the usual membrane-bounded cellular organelles found in an aerobic, eukaryotic, nonphotosynthetic organism. In addition, it contains a few specialized organelle systems uniquely designed to allow a single cell to function as a separate and complete organism independently of other cells (Allen, 1988). The only known exception to this cellular independence is the need to eventually exchange genetic information with another organism of the same species for survival. Biochemical and molecular studies of Paramecium have only been performed in a few laboratories. Consequently much of the information relating to its membrane trafficking has been obtained from electron microscopic and immunocytochemical studies. However, the ease with which these cells can be grown, manipulated, and observed microscopically has compensated, in part, for the difficulties encountered at the biochemical and molecular level. Paramecium, which is not a polarized cell in the same sense as an epithelial cell, does have numerous specialized areas at its plasma membrane which are designed for exocytosis and endocytosis. Thus, Paramecium can be thought of as a multipolarized cell where (1) the discoidal vesicles are transported to the cytopharynx, a specialized region at the cell surface designed for digestive vacuole (DV) formation, (2) the spent DVs are targeted to the cytoproct, a specialized region designed for vacuole defecation, (3) the trichocysts (secretory organelles) are transported to multiple and identical specialized docking sites on the plasma membrane designed for secretion, (4) contractile vacuoles periodically discharge their fluid at two permanent expulsion sites on the plasma membrane, and (5) clathrincoated pits are distributed at regular intervals over the cell surface through which endocytosis and, possibly constitutive exocytosis, can occur. Such specific domains within the plasma membrane show that paramecia, like polarized epithelial cells, are able to direct specific vesicular and vacuolar packages to very specific docking and fusion sites. In some cases, the sites are very limited in number, and only one vesicle or vacuole can be accepted for docking and fusion at a time. In other cases, the sites are numerous or

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more extensive in area so that many vesicles can be accepted simultaneously for docking and fusion. Along with the variety of docking and fusion sites at the cell surface, this cell also contains in its cytosol a number of vesicles that must recognize, dock, and fuse with specific membrane-bound motile compartments. These motile compartments are scattered among a very large and varied population of membrane-bound compartments within the cell. Failure of a vesicle to find its appropriate docking and fusion partner will presumably result in an inhibited or aborted cellular function. In this chapter, the discussion of membrane trafficking will be divided into four sections: (1) the biosynthetic pathway, (2) the regulated and constitutive secretion, (3) the endocytic system, and (4) the phagosomal system. Trafficking of membranes of the nuclei, the mitochondria, the peroxisomes, the contractile vacuole systems, and the calcium-storing alveolar sac system are not covered in this review, as little or no direct information is available on how they fit into the overall scheme of membrane trafficking in this cell. Figure 1 presents, in cartoon form, an overall view of how membrane trafficking in Paramecium is thought to occur and how its trafficking systems may be interrelated. After describing and discussing each membrane system, we will review the overall packaging and modulation of these membranes. Finally, we will discuss the schemes used by this cell for distribution and routing its packages to their proper destinations.

II. Membrane Trafficking A. Biosynthetic Pathway 1. ER and Its Transition Zone Most intracellular membranes, like the plasma membrane, are initially synthesized by the expansion of the endoplasmic reticulum (ER) (Fig. 2). Here exist the enzymes and molecular complexes required for the synthesis of integral membrane proteins and the phospholipids, the two major components of all biological membranes. As the proteins are synthesized on ribosomes, those destined to become membrane proteins possess both a signal sequence and one or more domains of hydrophobic amino acids that became embedded in the membrane. These hydrophobic domains will anchor the proteins in the membrane in a permanently asymmetrical orientation with respect to the membrane’s cytosolic and exoplasmic surfaces. Phospholipids, as synthesized, also remain as an integral part of the

FIG. 1 Membrane trafficking in Paramecium. A cartoon illustrating the various routes of trafficking that are discussed here. DV, digestive vacuole; I to IV, stages of DVs. For details refer to the text.

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ER membrane until they are transported to other membranes either by phospholipid transport proteins or by vesiculation. Thus one of the first steps in the trafficking of intracellular membranes in Paramecium, as in other cells, occurs at the transition zone of the ER. This is a ribosome-free membrane domain that contains protein complexes bound to its cytosolic surface. These protein complexes may enable the membrane to bud toward the cytosol and to eventually pinch off (Fig. 3). These transition vesicles (Fig. 4), derived from the ER membrane, pass to the forming face of the Golgi stack. 2. Membranes of the Golgi Stack A separate cis-Golgi reticulum or intermediate compartment, similar to that described in some mammalian cells (Rambourg and Clermont, 1990), has not been observed in Paramecium. Also, evidence of continuous vesicular–tubular clusters in the ER-to-cis-Golgi transition zone has not been demonstrated, even in physically fixed quick-frozen cells. The endoplasmic reticulum and Golgi stack lie so close together in Paramecium that this zone is wide enough for only a single layer of vesicles. The Golgi stack in this ciliate, as in other ciliates as well, is often reduced to only two cisternae that are abundantly fenestrated and/or reticulated (Esteve, 1972; Allen and Fok, 1993a; Garreau de Loubresse, 1993). In quick-frozen deepetched (QF-DE) preparations of chemically unfixed cells, the transition vesicles in the process of formation from the ER are observed to be coated with 11-nm diameter globules (Allen and Fok, 1993a). These globules are presumably Paramecium’s version of coatomer protein II (COP II) (Barlowe et al., 1994; Barlowe, 1995; Scales et al., 1997). These coated vesicles lose their coats and fuse with the cis-Golgi cisterna of the Golgi stack. 3. Filaments in the ER Transition Zone A unique feature of the transition zone between the ER and the Golgi stack in Paramecium is a meshwork of nonetchable filaments which spans the narrow space between the ER and the Golgi-stack (Fig. 4) (Allen and Fok, 1993a). Non-clathrin-coated buds and vesicles of 60-nm diameter (Table I) are encased in or attached to this meshwork and are presumably, therefore, not free to escape into the general cytoplasm (Allen and Fok, 1993a). These 60-nm vesicles, after they have separated from the ER, appear to lose their 11-nm-diameter coats and become the 40-nm vesicles that can be seen lying near the cis-face of the Golgi-stack (Fig. 4). The chemical nature of these transition-zone and Golgi filaments is not known. In kidney and muscle cells (Devarajan et al., 1996, 1997; De Matteis and Morrow, 1998), however, a homolog of ankyrin has been found to link a

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Golgi-specific spectrin to the Golgi apparatus. In Chinese hampster ovary (CHO) cells, Golgi-specific proteins such as p115, GM130, and giantin (So¨nnichsen et al., 1998) may be involved in linking COP I vesicles to the Golgi stack. Freeze-fracture and thin-section electron microscopy show that vesicles in CHO cells are linked to each other by filaments and with the Golgi cisternae, suggesting that vesicle procession through the Golgi stack is controlled by these attachments (Orci et al., 1998). This has been termed the ‘‘string theory’’ of vesicle procession. The cargo proteins of these transition vesicles, putatively, would consist of both membrane-bound and soluble proteins which are destined for transport (1) to the lysosomes, (2) to the secretory granules (trichocysts), and (3) to other cellular membranes or membrane-bound compartments such as the acidosomes, the plasma membrane, the alveolar sacs, the contractile vacuole system, and the early endosomes. Within the ER and the Golgi stack, the luminally exposed proteins are modified by glycosylation. One difference between Paramecium and higher organisms is that sialic acid does not appear to be attached to the ends of its putative complex-oligosaccharide chains (Pape et al., 1988). Thus the Golgi stacks of these ciliates may not need to have as many cisternae that compartmentalize the glycosylating enzymes to ensure that they function in the appropriate sequence. The presence of N-acetylglucosamine has been demonstrated at the trans-face of the Golgi stack of Paramecium (Allen et al., 1989) which suggests that the enzyme N-acetylglucosamine transferase is located at the trans-most cisterna of the Golgi stack. 4. The trans-Golgi Network A trans-Golgi network (TGN) is present in Paramecium and is composed of a mass of membrane-bound vesicles and cisternae with clathrin-like

FIGS. 2–5 Membranes of the biosynthetic pathway of Paramecium as seen in QF-DE images. FIG. 2. Ribosomes (arrowheads) attached to the endoplasmic reticulum (er). Arrows indicate what may be intramembrane protein complexes to which ribosomes dock (e.g., translocons). FIG. 3. Transition vesicles budding from the endoplasmic reticulum (er). Remnants of a COPlike coat composed of 11-nm globules surround the buds (arrowheads). Other globules line the neck (arrow). FIG. 4. The transition zone between the ER (er) and the Golgi stack (G). Transition vesicles formed from the ER are at first covered with a coat (arrowhead) which then disassembles before the vesicles (arrows) fuse with the Golgi stack. Nonetchable filaments link the ER to the vesicles and the Golgi stack. FIG. 5. The trans-Golgi network, where protein sorting occurs. Clathrin-like coats (arrowheads) are found around some of the vesicles or blebs in this area. The trans-Golgi network consists of a clump of vesicles and cisternae held together by a bundle of filaments (F). The E-fracture face (E) of these vesicles contain a relatively small number of particles as well as a few pits. (All figures in this chapter are printed at 100,000⫻ magnification.) Bar ⫽ 0.1 애m.

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TABLE I Membrane Trafficking Compartments in Paramecium Compartment Biosynthetic pathway Uncoated transition vesicles Coated transition vesicles Golgi cisternae Golgi vesicles TGN compartments Coated vesicles in TGN Secretory vesicle membrane Pretrichocyst vesicles Retrieved trichocyst vesicles Trichocysts Endocytic system Coated vesicles at cell surface Preendosomal vesicles Early endosomes Carrier vesicles Acidosomes at DV-I Phagosomal System Discoidal vesicles Phagosomes (DV-I) Retrieved DV-I membrane Phagoacidosomes (DV-II) Primary lysosomes Secondary lysosomes Phagolysosomes (DV-III) Retrieved DV-III membrane Spent vacuoles Retrieved membrane at cytoproct ‘‘Knobby’’ vesicles at DV Autophagosomes

Relative sizea

Shape

40 nm 60 nm ⬍700 nmb 100 nm 135 nm 110 nm

Spherical Spherical Flattened to discoidal Spherical Spherical to flattened Spherical

100 nm to 2.5 애mc 75 nm 3–4 애m ⫻ 1 애m

Spherical Spherical ‘‘Carrot’’ shaped

180 nm 180–200 nm 200–700 nm 80–100 nm 650–700 nm

Spherical Spherical Flattened to discoidal Spherical Spherical with indentations

300 nm 10–12 애m 75 nm diameter ⬍10애m 75 nmc 400 nm ⬎10 애m 40 nm diameter 앑10 애m 60–75 nm diameter 50–150 nm 1 애m

Discoidal Spherical Tubular to varied Spherical Spherical Spherical to prolate Spherical Tubular with bulbous ends Spherical Tubular to varied Spherical Spherical

a Measurements are from P. multimicronucleatum except where sizes from P. tetraurelia were used as in b(Esteve, 1972) or combined together as in c(Garreau de Loubresse, 1993).

coated evaginations extending from their limiting membranes (Fig. 5) (Allen and Fok, 1993a). These vesicles seem to be bound together by a tight meshwork of filaments. The TGN is the site of protein sorting and packaging for the secretory, lysosomal, and acidosomal proteins. It may also receive recycling vesicles returning receptors to the TGN. The membrane components destined for the alveolar sac system and the contractile vacuole complexes may also be sorted into separate vesicles at the TGN; however, no studies have yet determined how or where these two membrane systems arise in Paramecium.

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5. Mechanisms of Membrane Recognition, Docking, and Fusion As stated earlier, membrane trafficking in Paramecium has been studied most thoroughly using electron microscopy and immunocytochemical and cryo-immunogold techniques. Studies on the aspects of membrane recognition, docking, and membrane fusion are scarce for Paramecium. In one study, seven small GTP-binding proteins were isolated from Paramecium tetraurelia (Peterson, 1991). Three of the proteins were shown to be associated with the trichocyst membranes and with the cilia. The locations of the remaining four proteins within the cell were not identified. Using polymerase-chain-reaction (PCR) techniques, genes encoding 15 small GTP-binding proteins similar to the rab and rho subfamilies of proteins of mammalian cells (Balch, 1990; Novick and Brennwald, 1993; Schimmo¨ller et al., 1998) and their homologous proteins (Ypt) in yeast (Riezman, 1993; Lazar et al., 1997) were shown to be present in Paramecium (Fraga and Hinrichsen, 1994). But again, the locations and functions of the gene products have not been investigated. Thus, the mechanisms responsible for membrane trafficking between the ER and the Golgi stack, within the Golgi stack, and between the Golgi stack and the TGN wait to be systematically studied in Paramecium. From what has been determined by electron microscopy and the relatively few published biochemical and molecular biological studies, it is reasonable to expect that mechanisms for membrane synthesis and membrane trafficking similar to those used by yeast and mammalian cells will be found also in Paramecium. B. Regulated and Constitutive Secretion 1. The Trichocyst Paramecium exhibits a bona fide regulated or stimulated secretion when its trichocysts are expelled (reviewed by Plattner et al., 1993). Trichocysts are membrane-enclosed crystalline bodies which, when exposed to calcium ions, expand eight times in length into a long needle-like shaft (Hausmann, 1978). In nature the trichocysts apparently serve as a defense mechanism as they are expelled when a predator cell such as Entodinium or Dileptus nasutum approaches (Harumoto and Miyake, 1991; Knoll et al., 1991b; Miyaki and Harumoto, 1996). However, this is not an invariable response to all predators as the predator Didinium does not elicit the same response (Haacke-Bell et al., 1990; Miyake and Harumoto, 1996). 2. Trichocyst Development Trichocyst precursor proteins are synthesized in the biosynthetic pathway. These proteins have been found in the ER and Golgi stack within 20 to 30

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minutes following massive trichocyst exocytosis when immunocytochemical techniques had been used (Garreau de Loubresse, 1993; Garreau de Loubresse et al., 1994). In the P. tetraurelia mutant, tam 38, which forms aberrant football-shaped trichocysts (Ruiz et al., 1976), non-clathrin-coated vesicles of 앑55–60 nm in diameter accumulate in the region of the Golgi stack (Gautier et al., 1994). Such vesicle accumulation is not observed in wildtype cells. Small 100- to 200-nm vesicles lying near Golgi stacks were shown to contain trichocyst core proteins (Fok et al., 1988; Hausmann et al., 1988). Vesicles of this size near Golgi stacks can also be distinguished by regular thin-section electron microscopy (Gautier et al., 1994; Hausmann et al., 1988). Frequently, they are the smallest in a field of trichocyst precursor vesicles whose sizes extend from 100 nm up to 1–2 애m in diameter. The appearances of the contents of these vesicles vary and are indicative of the stage of their maturity. The smallest and youngest vesicles contain an amorphous flocculent material. The next larger group of vesicles contains an increasingly large core of crystalline material in the center of their flocculent content. By 60–90 minutes, those vesicles with small crystalline cores will have developed the carrot-shaped body of the mature trichocyst which is surmounted by a more electron-opaque tip (Garreau de Loubresse, 1993). The flocculent material, that had occupied much of the younger vesicles, will have disappeared in mature trichocysts, and the membrane will come to lie close to the trichocyst’s crystalline body.

3. The Trichocyst Membrane The trichocyst membrane buds from the TGN as vesicles that may or may not be coated. In trichless mutants (Pollack, 1974) large numbers of coated vesicles are seen at the TGN suggesting that coated vesicles may contain the precursor trichocyst proteins (Garreau de Loubresse, 1993). In any event, pretrichocyst vesicles (100–200 nm) coalesce and fuse together to form the ‘‘condensing’’ trichocyst vacuoles (see Fig. 6). A large excess of membrane must surround the developing trichocyst vesicles as the trichocyst contents condense. It is reasonable to assume that such excess membrane is removed and recycled back to the TGN. However, currently no experimental markers that can be used to follow such a recycling event in Paramecium have been found. Trichocyst membrane can be distinguished from other cellular membranes in QF-DE images, by the presence of large angular pits on the membrane’s E-fracture face. Trichocyst membrane has this appearance in both the condensing vesicle stage (Fig. 6) and the mature trichocyst stage (Fig. 7). This suggests the presence of large protein complexes, or clumps of complexes, that have been pulled out of this face during fracturing.

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Membrane trafficking is therefore an indispensable aspect of regulated secretion for moving the secretory proteins through the Golgi stack, for sorting and concentrating the numerous proteins making up the trichocyst within the TGN, and for removing excess membrane to produce a final discrete secretory package. 4. Trichocyst Transport within the Cell Other critical aspects of secretory membrane trafficking include: (1) the movement of the trichocysts to their highly specialized docking sites at the cell surface, (2) the process of docking preparatory to becoming competent to react to an external secretory signal, (3) the processes of membrane fusion, (4) the fate of the trichocyst membrane following secretion, and, finally, (5) the restoration and reconfiguration of the docking site to its original state ready to accept another trichocyst. Trichocysts, assembled in the cytosol, are said to be moved by saltation (Aufderheide, 1977) along microtubules to their docking sites (Plattner et al., 1982; Glas-Albrecht et al., 1991). The microtubules along which trichocysts move grow from microtubule initiation sites near the proximal ends of the basal bodies and extend deep into the cytosol. Because trichocyst movement is a minus-end-directed movement, it could conceivably be mediated by cytoplasmic dynein (Glas-Albrecht et al., 1991). Cytoplasmic dynein, a minus-end-directed motor, has been isolated from Paramecium and has been shown to be capable of actively moving microtubules in an in vitro assay system (Schroeder et al., 1990). However, the movements produced by a cytoplasmic dynein motor usually do not exhibit saltation but are smooth and more rapid than those reported for trichocysts. Thus the motor(s) that mediates trichocyst movement waits to be confirmed. 5. Trichocyst Docking and Fusion Docking of trichocysts occurs between the trichocyst tip (Fig. 8) and the cell surface. The multiple docking sites are distinguished in the freezefractured plasma membrane by a double circle of intramembrane particles (IMPs) located on the P-fracture face (Fig. 9). This double circle of particles, the outer circle being more prominent, lies at the junction between the plasma membrane and the margin of the underlying alveolar sac system. When no secretory vesicle is present under the docking site, the double circle of IMPs collapses to form a set of ‘‘parentheses.’’ If a trichocyst is present, the parenthesis of IMPs rounds out into a double ring. A rosette of about nine larger IMPs is found in the center of the ring principally on the E-fracture face on the plasma membrane in QF-DE images (Fig. 10) (Plattner et al., 1973). This rosette of IMPs forms the fusogenic site.

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Several minutes are required at the docking site before the newly arrived trichocyst can attain a state of fusion-competent readiness (Pape and Plattner, 1985). A stimulus presented after fusion-competence is achieved will result in membrane fusion during which an exocytosis-sensitive 63-kDa phosphoprotein (Gilligan and Satir, 1983) will be dephosphorylated (Ho¨hne-Zell et al., 1992). This 63-kDa protein, called parafusin, has recently been localized by confocal microscopy (Zhao and Satir, 1998) and other methods (Kissmehl et al., 1998) at the exocytic sites of the plasma membrane, to the alveolar sacs and to the trichocyst membrane itself. Also, stores of calcium in the alveolar sacs are involved in trichocyst exocytosis (Stelly et al., 1991, 1995; Knoll et al., 1993; Erxleben and Plattner, 1994; La¨nge et al., 1995). A calsequestrin-like protein, 53 kDa, has been identified in the alveolar sacs (Plattner et al., 1997). The release of the stored calcium from the alveoli seems to be activated by a local Ca2⫹ influx from outside the cell (Erxleben et al., 1997). Membrane fusion starts within the 10-nmwide zone defined by the rosette of IMPs and results in the apparent disassociation of the nine large IMPs into about six times as many smaller IMPs (Knoll et al., 1991a). The fusion site then expands beyond the diameter of the initial double ring of IMPs during the time (30 to 80 msec) required for the trichocyst to be discharged (Knoll et al., 1991a). 6. Trichocyst Membrane Retrieval and Fate The trichocyst membrane ghost remains behind after the trichocyst is discharged (Hausmann and Allen, 1976). The plasma membrane reseals within 앑350 msec (Knoll et al., 1991a). The ghost, that can be labeled with exogenous horseradish peroxidase (HRP), at first remains attached to the alveolar sacs (Haacke and Plattner, 1984; Plattner et al., 1985a) and then gradually vesiculates over a period of a few minutes (t1/2 ⫽ 30 min) (Pape and Plattner,

FIGS. 6–10 Membranes of the regulated secretory pathway. FIG. 6. Trichocysts, the secretory organelles of Paramecium, develop from condensing vesicles in which the trichocyst proteins (T) crystallize. During development the vesicles fuse (arrow) together. The E-fracture face (E) of these vesicles has large angular pits. FIG. 7. The E-face of the membrane of the mature trichocyst resembles that of the small trichocyst condensing vesicles. T, trichocyst crystalline core; arrowhead, rim of fractured cytosolic leaflet. FIG. 8. The elaborately decorated Pfracture face of the membrane of the trichocyst tip, shown here as an indentation into the replica (no rim indicating a fractured membrane leaflet is seen), lies under the docking site at the plasma membrane. FIG. 9. The mature trichocyst docks at the plasma membrane at a site marked on its P-fracture face by a double ring of intramembrane particles (IMPs) and a central rosette of pits located at the ultimate site of membrane fusion. FIG. 10. The rosette of IMPs at the tip of the trichocyst remains with the E-fracture face of the plasma membrane. Bar ⫽ 0.1 애m.

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1985). These 75-nm remnant vesicles (Hausmann and Allen, 1976) may remain for some time in the vicinity of the Golgi stacks, but eventually they will fuse with secondary lysosomes (Allen and Fok, 1984a) which fuse, in turn, with the digestive vacuoles (Lu¨the et al., 1986). A recent study in which trichocyst ghosts and digestive vacuoles were independently labeled and then viewed by confocal microscopy confirms the transfer of the contents of trichocyst ghosts to the phagosome/lysosome system (Ramoino et al., 1997). 7. Constitutive Secretion Studies on regulated secretion have also provided evidence of a constitutive secretory pathway in Paramecium. Among the mutants defective in the processing of the trichocyst precursor protein, the trichless mutant (Pollack, 1974) secretes unprocessed trichocyst proteins into the surrounding medium (Madeddu et al., 1994). This means that Paramecium must have other methods of protein secretion separate from secreting intact trichocysts because intact trichocysts are never secreted from these mutant cells. These observations argue for a constitutive secretory pathway. However, such a pathway has yet to be observed unequivocally, although a recent paper by Flo¨tenmeyer et al., (1999) proposes that constitutive secretion occurs via the secretion of uncoated vesicles arising from the TGN at the parasomal sacs. The problem with the data presented is that it is impossible to distinguish between vesicles coming into the cell by endocytosis and those leaving the cell by exocytosis because the same marker may be present in both types of vesicles. In summary, Paramecium offers distinct advantages for the study of membrane trafficking during regulated secretion as well as the process of secretion itself. The secretion steps can be studied during the triggering of massive exocytosis (Plattner et al., 1985b), and a number of secretory mutants which are defective in specific steps in this pathway as well as in trichocyst biosynthesis are available. Also, the molecular biology of trichocyst formation is now at a stage where the proteins and the genes involved can be identified and studied (Madeddu et al., 1995).

C. Endosomal Trafficking 1. Endocytosis at the Parasomal Sac/Coated Pit Membrane trafficking also occurs during endocytosis. The plasma membrane of Paramecium possesses multiple endocytic sites regularly arranged over the somatic surface of the cell and over the buccal cavity within the

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oral region. These sites, classically called parasomal sacs by protozoologists, are in fact morphologically analogous to the clathrin-coated pits of other organisms. Thus, for clarity, the term ‘‘parasomal sac’’ will be used for the permanent porelike indentation where the plasma membrane passes inward through a circular gap in the alveolar sac system, and the term ‘‘coated pit’’ will be used to refer to the coated cytosolic extension of the plasma membrane that lies internal to this pore (Figs. 11 and 12) (Allen and Fok, 1993b). Usually there is one clathrin-like coated pit lying to the right side of a basal body or a pair of basal bodies within each polygonally shaped cortical depression. There are about 8000 such cortical depressions on the surface of Paramecium multimicronucleatum (앑1000 in P. tetraurelia). In addition, there are 앑500 coated pits that are arranged in rows that separate the three ciliary membranelles in the buccal cavity of this cell. The coated pits average nearly 180 nm in diameter in P. multimicronucleatum and are encased in a clathrin-like cage, as can be visualized in favorable thin sections and even more clearly in QF-DE images (Allen et al., 1992). It is reasonable to assume that these coated pits are sites for receptormediated endocytosis. However, little information is known about what receptor–ligand complexes might be endocytosed in Paramecium by this method. Receptors have been postulated for biotin (Bell et al., 1998), and a receptor for GTP and a 58-kDa receptor for lysozyme are known to be present on Paramecium’s surface (Kim, et al., 1997), but whether they are taken into the cell via the coated pits has not been investigated. 2. Preendosomal Vesicles and Early Endosomes Coated pits pinch off to form coated preendosomal vesicles of 앑200 nm in diameter (Fig. 13). These vesicles take up HRP (Allen et al., 1992) or ferritin (Westcot et al., 1985). The loss of the clathrin-like coat occurs soon after the vesicle is released from the plasma membrane. These uncoated preendosomal vesicles dock and fuse with the membrane of an early endosome, one of many flattened cisternae situated near the level of the proximal ends of basal bodies (Figs. 14 and 15). These early endosomes range from 0.2 to 0.7 애m in diameter and are circular or cup-shaped in three-dimensional morphology (Table I); they characteristically have small blebs bearing a clathrin-like coat budding from their membranes (Allen et al., 1992). These buds are only 80 nm in diameter so that they are only about half the diameter of the coated pits at the plasma membrane. Several antigens have been immunologically localized in these cisternae. Capdeville et al., (1993) have used polyclonal antibodies to localize both a 250-kDa GPI-anchored surface antigen and a 40-kDa transmembrane antigen on the cell surface and in the early endosomes in P. tetraurelia. Flo¨tenmeyer et al. (1999) have also raised several polyclonal antibodies to the

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surface proteins of this species. In P. multimicronucleatum a 190-kDa antigen was located with a monoclonal antibody (mAb) on the plasma membrane, the preendosomal vesicles and the flattened portions of the early endosomes but not on the 80-nm evaginations extending from the margins of early endosomes (Allen et al., 1992). Such coated and uncoated vesicles (80 to 100 nm) when labeled with exogenous HRP marker that had entered the early endosomes by the endocytic pathway do not carry the 190-kDa antigen. That the 190-kDa antigen does not pass deeper into the endoplasm suggests that this antigen must be recycled back to the plasma membrane.

3. Carrier Vesicles Arise from Early Endosomes A second antigen of 40 kDa was also located by mAb labeling at the plasma membrane of P. multimicronucleatum. This antigen enters the early endosomes, but unlike the 190-kDa antigen, it enters and accumulates in the 80 nm coated buds. These coated buds are pinched off the early endosomes as 100-nm vesicles that are called carrier vesicles (Fok et al., 1993). After separation from the early endosomes, they enter the endoplasm where they soon appear next to acidosomes (see Fig. 18). These carrier vesicles then fuse with and add their cargo and membrane to the acidosomes (discussed in Section II.D.2) (Allen et al., 1993; Fok et al., 1993). Thus Paramecium has the components required of a classical ‘‘receptormediated’’ endocytic pathway. Multiple coated pits at the plasma membrane give rise to preendosomal vesicles that are rapidly uncoated. These uncoated preendosomal vesicles are then incorporated into the early endosomes that are typically found in the cortex (exoplasm) of the cell. Similar to mammalian cells, the early endosomes also function as sorting compartments where surface proteins can be sorted from the soluble cargo and other membrane proteins that are destined to be delivered to the late endosomes and eventually to the phagocytic system. Recycling of the sur-

FIGS. 11–15 Membranes of coated vesicles and early endosomes. FIGS. 11 and 12. The parasomal sac, a pore in the plasma membrane (small arrows) is connected to a coated pit, seen here with its E-face exposed (E). A clathrin-like coat (large arrow) covers the cytosolic surface of the coated pit or coated vesicle. FIG. 13. Coated pits pinch off to form large preendosomal vesicles (arrows). Few IMPs are found on the E-fracture face of these vesicles (E). FIGS. 14 and 15. The preendosomal vesicles merge with an early endosome seen here with only a few IMPs on its exposed E-face (E). Smaller coated pits and vesicles (small arrows) arise from the margins of these early endosomes. Large arrow in Fig. 14 is a basal body cross-fractured through its proximal end. Bar ⫽ 0.1 애m. Figures 12–15 are from Allen et al. (1992).

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face proteins must occur, but for technical reasons this has not yet been documented. 4. Parasomal Sacs as Possible Sites of Exocytosis and Noncoated Forms of Endocytosis As is the case for constitutive secretion (Flo¨tenmeyer et al., 1999), the exact site on the plasma membrane for docking and fusion of small recycling vesicles is not known, though both Allen (1988) and Capdeville et al. (1993) have reached the same conclusion that the membrane must be returned to the parasomal sacs for exocytosis and reentry into the plasma membrane. Except for the parasomal sacs, the close and uniform apposition of the alveolar sac system with the plasma membrane would seem to preclude any other site for exocytic vesicles to be attached to the plasma membrane. Although usually coated with clathrin-like cages, an occasional much larger and mostly uncoated membrane tubule or invagination has been observed to be continuous with a parasomal sac in Paramecium [see Fig. 67 in Allen (1978)] as well as in Tetrahymena (Allen, 1967). Exocytosis at the parasomal sacs has been better documented in Vorticella (Suchard and Goode, 1982). These observations suggest that parasomal sacs may be used as sites for both endocytic and exocytic activities and not just clathrin-coated receptormediated endocytosis.

D. Phagosomal System The phagosomal system of Paramecium represents a membrane trafficking system equal to or of greater complexity than phagosome systems described in mammalian cells (Fok and Allen, 1988, 1990, 1993). It is the intent of this section (1) to briefly summarize previous findings on phagosome membrane trafficking, (2) to present recent findings, and (3) to show how the endosomal, secretory, and biosynthetic pathways are associated with the phagosomal system. In this chapter the term ‘‘phagosome’’ is interchangeable with the first stage of the digestive vacuole, the DV-I, DV-II with the phagoacidosome, DV-III with the phagolysosome, and DV-IV with the spent DVs. 1. Discoidal Vesicles, Phagosomal Membrane Precursors Phagosomes arise at the cytopharynx. Membrane is added to this permanent single membrane-lined region along the left side of the posterior half of the buccal cavity. The cytopharynx is supported by a complex cytoskeletal architecture (Allen, 1974). Phagosomes grow as discoidal vesicles fuse with

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the cytopharyngeal membrane (Allen, 1974) and merge into the plane of the nascent digestive vacuole (NDV) membrane. The discoidal vesicles are transported to the cytopharynx along 앑40 microtubular ribbons (Fig. 16) (Schroeder et al., 1990). These ribbons are anchored at the cytopharygeal membrane (Allen, 1974). From their initiation sites in the filamentous reticulum, the ribbons pass under and then curve back sharply over a cytostomal cord. The ribbons then extend laterally into the cytosol for several micrometers. This orientation gives the ribbons an anterior and posterior side relative to the cell’s axis. Schroeder et al. (1990) have shown that the discoidal vesicles are moved toward the cytopharynx mostly, if not entirely, along the anterior sides of these ribbons, and in a minus-end direction toward the origin of the microtubules. Not only has the minusend-directed motor, cytoplasmic dynein, been isolated from Paramecium (Schroeder et al., 1990), it has also been localized between the discoidal vesicles and these ribbons (Fok et al., 1994). This strongly suggests that cytoplasmic dynein is the motor responsible for the transport of the discoidal vesicles to the cytopharynx. That the discoidal vesicles move only forward on one side of the ribbons suggests that binding sites for cytoplasmic dynein are available only on the anterior sides of the ribbons. This is shown to be the case as bridging filaments seen in QF-DE images, that are presumably cytoplasmic dynein heavy chains, link the discoidal vesicles to the anterior side of the microtubular ribbons (Fig. 16). Thus discoidal vesicles are directed to their docking and fusion sites by first binding to the cytoplasmic dynein which is then temporarily but repetitively attached to the microtubules. Vesicles are thereby moved along these microtubules to their fusion sites at the membrane of the cytopharynx. The cytostomal cord that lies along the cytopharyngeal ends of the microtubular ribbons is a centrin-containing myonemal-like filamentous bundle (Klotz et al., 1997; Allen et al., 1998) thus it must contract when exposed to calcium. Such contraction may lead to a close association of the discoidal vesicles with the cytopharyngeal membrane and thereby the cord may promote fusion. The source of calcium for this contraction undoubtedly comes from the alveolar sacs, compartments that are known to store calcium (Stelly et al., 1991, 1995; Knoll et al., 1993; La¨nge et al., 1995) and that send finger-like tubular projections to this cord (Allen, 1974; Allen et al., 1998). These projections are bound tightly to the cord by short fibrous links (Allen, 1974). The origin of the discoidal vesicles has not been completely resolved. Many discoidal vesicles are formed from retrieved DV-I membrane and others are formed from the spent vacuole membrane at the cytoproct. This problem is more fully discussed in Section III.A.3.

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2. Phagosome Formation As the NDV grows, it becomes filled with food particles that are swept into and along the buccal cavity by the beating of the oral ciliary membranelles. Under an as yet unknown trigger, the NDV begins to move along the postoral microtubular bundles in a microtubular plus-end direction. This movement appears to be responsible for pulling the NDV away from the cytopharynx resulting in the fission of the vacuole membrane. The final pinching off, that results in the formation of the DV-I, resembles the formation of a soap bubble when one blows on a soap film covering a wire or plastic loop. The motor for this movement is actin and myosin based because anti-heavy meromyosin antibodies labeled the postoral microtubular bundles during and shortly after this phagosome movement (Cohen et al., 1984a, 1984b). Cytochalasin B, a potent inhibitor of actin polymerization, inhibits the parturition of the NDV from the cytopharynx (Allen and Fok, 1985; Fok et al., 1985), suggesting that actin polymerization may be required just before the commencement of the movement of the NDV along the microtubular bundle. This movement itself is very rapid, 앑20 애m/sec, which is faster than movements produced by known microtubule-based motors (Schroeder et al., 1990). 3. Acidosomes: ‘‘Late Endosomes’’ That Fuse with Carrier Vesicles and Then with Phagosomes At the time the NDV is growing in size, a pool of relatively large vesicles is transported to the cytopharynx along the same set of cytopharyngeal microtubular ribbons that transport the discoidal vesicles. These are called the acidosomes because they are responsible for the acidification of the DV-I (Allen and Fok, 1983c). They seem to be too large to move under the cytostomal cord at the minus ends of the cytopharyngeal ribbons. Upon encounteng the NDV membrane which lies along the minus ends of the cytopharyngeal ribbons, these acidosomes become firmly docked at the NDV membrane and leave the microtubular ribbons as the NDV membrane

FIGS. 16–18 Vesicles that move along the cytopharyngeal microtubular ribbons. FIG. 16. Discoidal vesicles (DC) are linked to microtubular ribbons (small arrow) by bridges (arrowheads). The E-fracture face (E) of these vesicles is packed with IMPs. FIG. 17. Small 100-nm vesicles are also transported along these ribbons (arrows). Bridges (arrowheads) link these vesicles to the ribbons. A few IMPs can be seen on the E-faces (E) of these vesicles. FIG. 18. Acidosomes (AC) also move along these ribbons. The acidosome seen here has its P-face (P) exposed and is lying close to a 100-nm vesicle with which it is known to be able to fuse. Bar ⫽ 0.1 애m. Figure 18 is from Allen et al. (1993).

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grows. Ultimately, these acidosomes, that remain bound to the NDV as well as the released phagosome, will fuse with the DV-I (see Section II.D.4). Acidosomes can first be seen at the TGN, where they have a flattened or rounded shape (Allen et al., 1993). They then become more spherical and attain a size of roughly 1 애m in diameter. Their limiting membranes often exhibit characteristic indentations that protrude into their lumens. The reason for these indentations is unclear. In QF-DE images acidosome membranes sometimes show signs of regular packing of lipid subunits (see Fig. 19, right side of the acidosome on the figure’s right). They contain vacuolar-type proton pumps (V-ATPase) on their membranes (Ishida et al., 1997). These ATPase complexes may correspond to some of the prominent IMPs in their P-fracture face (Figs. 18 and 19) as well as the prominent particles that protrude from the true cytosolic face of the acidosomes (Fok et al., 1996). A third set of vesicles, the 100 nm vesicles, are found aligned along the same cytopharyngeal microtubular ribbons (Fig. 17) (Schroeder et al., 1990). These vesicles have the same size as the carrier vesicles arising from the early endosomes (see Section II.C.3) (Allen et al., 1992). Such carrier vesicles containing exogenous HRP can be seen docked on the acidosome membrane. The carrier vesicles then fuse with acidosomes and transfer their HRP cargo and membrane to the acidosomes (Fig. 18) (Allen et al., 1993). Thus the acidosomes can be considered to be late endosomes as defined in mammalian cells. By transferring the combined membranes and cargo of these carrier vesicles and the acidosomes to the phagosomes, the endosomal system merges into the phagosomal system, as also happens in the macrophage (De Chastellier and Thilo, 1998). 4. Passage of DVs through the Cell From the time a DV-I is released from the postoral microtubular bundles in the posterior quarter of the cell, it progresses slowly through the cell by cyclosis whose motive force has yet to be determined. In general, a DVI (phagosome) is located in the posterior quarter of the cell, a DV-II (phagoacidosome) can be found in the posterior half of the cell, a DV-III (phagolysosomes) is usually found in the anterior half, and a DV-IV (spent vacuole) at times can be found in the anterior half but is most commonly located in the posterior half of the cell near the cytoproct (Allen and Fok, 1984b). 5. Conversion of a Phagosome into a Phagoacidosome via Membrane Replacement When the DV-I pinches off from the cytopharynx, along with its load of docked acidosomes (Fig. 19), it moves into the posterior end of the cell

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(Allen and Fok, 1983a). Within the next 15 sec to 1 min, the acidosomes fuse with the phagosome. At about the same time, the original discoidal vesicle-derived phagosomal membrane becomes tubulated (Allen and Fok, 1983a, 1983b, 1983c). The membrane tubules are then pinched off from the phagosomes. This leads to the formation of (1) a DV whose content, for the first time, can become acidified, and (2) a pool of membrane tubules capable of being remodeled into discoidal vesicles for recycling back to the cytopharynx. The end result is an essentially complete membrane replacement (Allen and Fok, 1983b) where the ‘‘discoidal vesicle’’ type of membrane (Fig. 19) is replaced with an ‘‘acidosome’’ type of membrane, thus converting the DV-I into a DV-II or a phagoacidosome (Fig. 20) (Allen et al., 1993; Fok and Allen, 1993). This dramatic replacement has been supported by images of vacuole membranes of known ages as seen in thin sections, freeze-fracture replicas (Allen and Staehelin, 1981), and QF-DE replicas (Fok and Allen, 1993). Furthermore, such replacement is supported by immunofluorescence (Fok et al., 1986) and immunogold studies. Cryo-immunogold labeling using mAbs specific for antigens of the discoidal vesicle membrane (B2, Q2) and acidosome membrane (L1, E9) (Allen et al., 1995) verified the membrane replacement process. The discoidal vesicle and the DV-I (prior to fusing with acidosomes) contain only the B2 and Q2 antigens. The acidosomes that are docked at the DV-I or are in the process of fusing with the DV-I and the resulting DV-II contain the two acidosomal membrane antigens. Thus, it appears that the replacement is essentially complete and that the membrane surrounding the DV-II has a different set of antigens than is found in the membrane around the DV-I (Allen et al., 1995). The membrane tubules that come off the DV-I also contain only the discoidal vesicle membrane antigens, B2 and Q2. It can thus be concluded that these tubules are destined to be reformed into discoidal vesicles and returned to the cytopharynx. Together with the discoidal vesicles derived from the spent DVs, these retrieved tubules form the pool of discoidal vesicles at the cytopharynx. No other pool of vesicles bearing the discoidal vesicle antigens has been found in the cell. The phagoacidosome or DV-II soon becomes strongly acidic (Fok et al., 1982). A recent study showed that these DV-II have a V-ATPase in their membranes (Ishida et al., 1997). Using three antibodies raised against the subunits of V-ATPases, one or two large DVs in the posterior half of the cell were found to be strongly labeled and several other DVs were found to be more weakly labeled. Cryo-immunogold techniques showed that the membranes of the docked acidosomes carried the V-ATPases on their membranes that resulted in the one or two brightly fluorescent DVs. These V-ATPases are then transferred to the DV-II membrane when the acidosomes fuse with the DV-I. Concanamycin B (at 10 nM), a V-ATPase specific

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inhibitor, can significantly inhibit the rate of vacuole acidification (Ishida et al., 1997). This shows that this proton pump is responsible for the acidification of the phagoacidosomes. 6. Conversion of Phagoacidosome into a Phagolysosome The stage of the DV at which lysosomes can bind appears to be the phagoacidosome or DV-II stage (Fig. 20) (Fok et al., 1984, 1987). Lysosomes have not been found docked at the phagosomes or DV-I. Lysosomes are present in at lease two populations, the 75-nm primary lysosomes arising from the TGN and the larger 0.5-애m secondary lysosomes that bind to the DV-II (Fig. 20). The primary lysosomes appear to contain a higher relative concentration of hydrolases because they are usually filled with acid phosphatase reaction product in cytochemically stained cells, whereas secondary lysosomes contain only scattered foci of the same reaction product in their lumens (Fok et al., 1984). The primary lysosomes may either fuse with the secondary lysosomes or with the DV-II to deliver their load of hydrolases. Docking of secondary lysosomes at the DV-II can be so extensive that a seemingly solid layer of lysosomes can be seen surrounding the DV-II (Allen and Fok, 1984b). Fusion of lysosomes with the DV-II membrane appears to be synchronous—it is all or none. This suggests that a specific trigger has led to a simultaneous fusion, although what this trigger is has not been determined. However, as a result of this fusion, a phagolysosome (DV-III) is formed and digestion commences. The freeze-fracture appearance of the phagolysosome membrane now resembles that of the lysosome rather than the acidosome (Fig. 21) (Allen and Staehelin, 1981). The period of active digestion lasts for 10 minutes or longer depending on the type of food contained in the phagolysosome (Fok and Allen, 1990). Another potential source of secondary lysosomal membrane is that contributed by trichocyst membrane retrieval. The vesicular remnants of trichocyst membrane apparently fuse with the secondary lysosomes. Particularly

FIGS. 19–20 Membranes of digestive vacuoles, acidosomes, and lysosomes. FIG. 19. The highly particulate E-fracture face of the phagosome (DV-I), resembling discoidal vesicle E-faces, lies next to docked acidosomes. Both the P-fracture face (P) and E-fracture face (E) of acidosomes can be seen. Arrowheads indicate pits in the E-face where protein particles, presumably remaining with the P-face, have been pulled out of this leaflet. FIG. 20. The E-fracture face of the phagoacidosome (DV-II) has very few IMPs but does contain pits (arrowhead). Lysosomes, seen here, one in cross-fractured view with its lumen exposed (L) and two with their E-fractured faces exposed (E), contain some prominent IMPs on their E-faces.

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following a massive discharge of trichocysts, huge amounts of secretory membrane can be returned to the cytosol as 75-nm vesicles. Presumably this membrane will become part of the lysosomal membrane pool because these remnants eventually fuse with secondary lysosomes (Allen and Fok, 1984a). How such large quantities of secretory membrane may alter the function of the lysosomal membrane, which has a distinctive and extensive glycocalyx on its luminal side (Allen and Fok, 1984b), is unknown. 7. Formation of the Spent Vacuole and Vacuole Membrane Retrieval at the Cytoproct At the end of digestion, a new round of vacuole membrane tubulation commences, and membrane tubules, 앑45 nm wide bearing bulbous distal ends and containing acid phosphatase, form from the surface of the DVIII (Allen and Fok, 1984c). These tubules appear to arise in patches, with a few tubules per patch. These patches are randomly scattered over the vacuole surface (Allen and Staehelin, 1981; Allen and Fok, 1984c). These tubules with bulbous ends separate from the phagolysosome and presumably recycle as secondary lysosomes. At times these tubules contain some acidosome antigens (Allen et al., 1993). The acid phophatase-negative spent vacuole is now a defecation-competent vacuole or DV-IV. This occurs about 20 minutes from the beginning of the digestive cycle in cells growing on a partially chemically-defined culture medium (Fok and Allen, 1979). Ultimately the spent vacuole comes in contact with microtubule bundles extending into the cytoplasm from the proximal ends of those basal bodies bordering the cytoproct, its defecation site (Allen and Wolf, 1974). The spent DVs are thought to move along these microtubular bundles in a minus-end direction toward the cytoproct. Single microtubules extend into the cytoplasm from the ridge of the cytoproct where they also appear to make contact with the spent vacuole membrane as it approaches the cytoproct. The position of these single microtubules may move over the surface of the DV-IV causing the cytoproct ridge to be pulled down toward the DV membrane, thus allowing the DV membrane to approach the plasma membrane at the cytoproct and then to fuse with it. An opening formed between these two membranes would result in a small focal fusion that would presumably quickly expand, due to membrane fluidity, to the diame-

FIG. 21. The phagolysosome (DV-III) membrane has a moderate number of prominent IMPs on its E-face (E) which resembles the E-face of secondary lysosomes. Similar to the lysosome (Ly) the phagolysosome membrane has a few small pits on its E-face (arrows). Bar ⫽ 0.1 애m. Part of Fig. 21 is from Fok and Allen (1990).

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ter of the vacuole or to the extent that the cytoproct can physically expand. This would lead to the rapid defecation of vacuole contents (Allen and Wolf, 1974). The closing of the cytoproct requires the complete retrieval of the spent DV membrane. This process of membrane retrieval can be inhibited by drugs that prevent actin polymerization such as cytochalasin B (Allen and Fok, 1985) or by the calmodulin antagonist, trifluoperazine (Fok et al., 1985). Membrane retrieval of spent DV membrane is accomplished by tubulation of the membrane. As would be expected based on the drug studies, microfilamentous material has been found around the 60- to 75nm-diameter membrane tubules that form at the cytoproct (Allen and Wolf, 1974; Allen and Fok, 1985). During vacuole defecation, exogenous markers such as HRP can enter the lumens of tubules and become trapped during fission (Allen and Fok, 1980). Under such conditions a mass of HRP-labeled tubules and vesicles of varied morphologies accumulates adjacent to the cytoproct soon after defection. These tubules and vesicles eventually bind to the cytopharyngeal microtubular ribbons. Such binding seems to result in the transformation of the tubular vesicles into a uniform discoidal shape. The discoidal shape may be induced as the number of cytoplasmic dynein motors binding the vesicles to the cytopharyngeal microtubular ribbons (Fok et al., 1994) becomes maximized. Evenly distributed motors attached to the tubular vesicles may pull the tubules flat against the ribbon. Because each ribbon consists of 앑12 microtubules oriented parallel to each other and lying in the same plane, the average ribbon is wider than the diameter of a typical discoidal vesicle. Thus, tubules formed from the spent vacuole membrane become transformed into discoidal vesicles and are transported to the cytopharynx for a new round of phagosome formation.

III. Membrane Modulation and Membrane Transport Systems A. Budding, Tubulation, and Sorting Events A comparison of membrane-bound packages that must be formed and then distributed to specific locations within Paramecium (Table I) shows that such containers range in size from the uncoated transition vesicles (as small as 40 nm in diameter) to the DVs which are 14 애m or larger. These containers also vary in shape; most are more or less spherical, but some are flattened or have a discoidal shape.

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1. QF-DE Images Can Be Used to Identify Different Membranes Quick-frozen deep-etched images of the various membrane packages show how membranes of these trafficking compartments differ in their relative protein content based on their freeze-fracture morphology. The membranes of physically fixed cells (chemically unfixed) can often be distinguished by the number and size of pits left in their E-fracture faces following freezefracturing and deep-etching. The fracture faces of the ER have a heterogeneous array of IMPs and pits of various sizes that are present in specific patterns (Fig. 2). The transition vesicles and small vesicles in the Golgi stack have unpitted E-fracture faces (Figs. 3 and 4), whereas the vesicles in the TGN are perforated by a relatively low number of pits of different sizes on their E-fracture faces (Fig. 5). The membranes that give rise to the phagosomes (i.e., the discoidal vesicles) have few pits but are studded with IMPs on their E-fracture faces (Fig. 16). The 100-nm carrier vesicles have very few pits (Figs. 17 and 18). The acidosomes have a moderate number of relatively small pits with larger ones forming a dispersed ‘‘necklace’’ around the sites where carrier vesicles (regions without pits) seem to have fused with the acidosomes (Fig. 19) (Allen et al., 1993). The lysosomes have a few small pits on their E-fracture faces (Figs. 20 and 21). The E-fracture faces of the DV-I, DV-II, and DV-III are similar in appearance to their membrane precursors, the discoidal vesicles (Fig. 19), acidosomes (Fig. 20), and lysosomes (Fig. 21). Membrane of early endosomes is relatively smooth with a few scattered pits of small diameter (Figs. 14 and 15) and in this respect resembles preendosomal (Fig. 13) and coated pit membranes (Figs. 11 and 12). In contrast, precursor and mature trichocyst membranes are characterized by many large and angular pits (Figs. 6 and 7). The E-fracture face of the membranes of the contractile vacuole system is also highly pitted. The smooth spongiome has numerous pits (Fig. 22), whereas the decorated spongiome in its tubular form fails to fracture at all. Only when the decorated spongiome rounds up, possibly during cell division or when the cell is under stress, does this membrane fracture. At such times, the E-fracture face of the decorated spongiome is so porous that it barely resembles a membrane at all (Fig. 23). These QF-DE images indicate differences in the composition of the membranes, particularly in their transmembrane protein complexes or in the way these proteins are associated with the underlying cytoskeleton. These complexes leave their distinctive imprints on the exoplasmic leaflet of the membrane when they are fractured. In some cases their imprint is so distinctive that QF-DE images can be used to identify the vesicles without additional markers.

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FIGS. 22–23 Membranes of the contractile vacuole complex. FIG. 22. A collecting canal (CC) has many IMPs on its P-face (P), whereas its E-face (E) has a spongy appearance because of the many pits left behind by the removal of IMPs. FIG. 23. Decorated tubules normally do not fracture as a result of their very high protein content. However, when they do round up (DS), their fractured appearance resembles no other membrane because of their extremely pitted appearance (E). L, lumen of DS; arrowheads, outline the fractured edge of a DS vesicle. Bar ⫽ 0.1 애m.

2. Coated Membranes Lead to Forward Processing (Anterograde) Movements There are four regions where coated pits give rise to coated vesicles in Paramecium. COP-like coats occur at the transition zone of the ER (Fig. 3), clathrin-like coats occur at the endocytic sites at the plasma membrane (Figs. 11 and 12) and clathrin-like coated vesicles bud from the early endosomes (Figs. 14 and 15) and the TGN (Fig. 5). In each case the coated vesicles can be said to be moving in an anterograde direction with reference to the cargo (i.e., forward from the site of cargo synthesis and packaging).

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In all cases the membrane curvature has become modulated by receiving a cytosolic coat so that a more or less planar topography takes on a highly curved topography. In these cases the bud that forms from the planar membrane ultimately pinches off to form a vesicle of a characteristic size. The COP-like coats lead to 40-nm vesicles after the coats are removed and the clathrin-like coats give rise to vesicles of two sizes—the 75- to 100-nm or 앑180-nm vesicles depending on their origins. It has been determined that in mammalian cells clathrin binds to adaptor proteins (AP) and that at least three different adaptor complexes exist (Schmid, 1997; Odorizzi et al., 1998); AP-1 attaches clathrin to the TGN, AP-2 attaches clathrin to the plasma membrane, and AP-3 attaches clathrin to the early endosomes. A similar situation can be anticipated to occur in Paramecium. 3. Tubulation Leads to Membrane Recycling (Retrograde) Movements and to Changes in Fusogenic Properties of the Membrane Membrane tubules form in association with a microfilamentous material on both the membranes of the DV-I and of the cytoproct-attached DV-IV. These tubules have diameters of 60 to 75 nm (Allen and Fok, 1980, 1983a), and after fission both give rise to tubular vesicles that are modified in shape and recycled as discoidal vesicles. In contrast, the tubules arising from the DV-III are smaller in diameter (45-nm) and return both membrane and acid phosphatase to the pool of secondary lysosomal membrane (Allen and Fok, 1984c). Because of the smaller diameter of these membrane tubules, the mechanism for retrieval of lysosomal membrane may differ from that utilized on the DV-I and DV-IV membranes. Recycled membrane plays a key role in the phagocytic cycle in Paramecium. The accumulated evidence suggests that four possible sources of new membrane enter the phagosome system. New membrane from the plasma membrane and endosomal system enters via the carrier vesicles that arise from the early endosomes and that fuse with the acidosomes. A second source of membrane comes from the primary lysosomes that originate from the TGN and fuse with the secondary lysosomes or that fuse directly with the DV-II. The third source of membrane comes from the acidosomes that also originate from the TGN. A fourth possible source of new membrane is from remnant vesicles of trichocysts that fuse with the secondary lysosomes. Interestingly, vesicles containing the acidosome antigen L1 and primary lysosomes containing acid phosphatase have been observed in the TGN but vesicles containing the discoidal vesicle membrane markers (B2 and Q2) have not been found in association with the Golgi stacks or the TGN. Part of the discoidal vesicle pool is derived from the recycled membrane

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of the DV-I, and this pool contains the B2 and Q2 markers. Another part of this pool is derived from retrieved DV-IV membranes at the cytoproct, and, like the DV-IV, they lack the B2 and Q2 antigens. How the discoidal vesicles recycled from the cytoproct obtain these antigens or if they actually do acquire these antigens that are absent in the spent vacuole membrane is a question that requires further study. The essential need for a phagosome to be able to form tubules has recently been demonstrated in mammalian macrophage. Under some conditions, when a macrophage takes a single particle into its phagosome, such as a 1-애m hydrophobic polystyrene latex bead or a single virulent Mycobacterium avium, the particle binds so tightly to the phagosomal membrane that the phagosome membrane will not be able to be modified into tubules (De Chastellier and Thilo, 1997). Phagosomes that contain such particles will still be able to fuse with early endosomes but they are prevented from fusing with lysosomes. Thus, normal maturation of the phagosome will be inhibited. If the particle is a parasitic bacterium such as M. avium, it can remain alive and divide inside the phagosome (De Chastellier and Thilo, 1998). Tubulation is apparently required for the phagosome membrane to be able to change its fusogenic properties from a membrane that is capable of fusing only with early endosomes to one that is capable of fusing only with lysosomes. As in the macrophage, tubulation in Paramecium is probably required for the modification of the fusogenic properties of the DV membrane. Changes in the DV’s fusogenic properties are probably needed at the end of the DV-I stage and at the end of the DV-III stage where tubulation has been observed. However, changes may also occur at the end of the DV-II stage, although extensive tubulation has not been seen at this stage. Controlling the fusogenic properties of the membrane will likely ensure that fusion between compartments will occur in an appropriate physiological sequence. 4. Cognate Membrane Recognition, Docking, Priming, and Fusion Studies of yeast and mammalian cells, particularly the synaptic vesicle studies of neurons, indicate that, for an acceptor membrane to recognize and fuse with the appropriate donor membrane, both membranes require membrane-specific protein complexes (Su¨dhof, 1995). For example, for the synaptic vesicles to recognize and dock with the presynaptic nerve membrane at the neuromuscular junction, a t-SNARE in the nerve plasma membrane must bind to a v-SNARE in the synaptic vesicle. Various soluble protein complexes then associate with the docked vesicles to form a fusion complex. Applying these studies to membrane trafficking in Paramecium

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promises to be an exciting area of research. For example, different v-SNAREs should be found in discoidal vesicles and acidosomes that bind, respectively, to two different t-SNAREs located in the DV-I. Secondary lysosomes should possess SNAREs recognizing cognate SNAREs in the DV-II, the remnant secretory membrane vesicles, as well as primary lysosomes. Besides the SNARE in the acidosome membrane that recognizes the DV-I, the acidosomal membrane should also contain a SNARE for carrier vesicles coming from early endosomes. In addition to the SNAREs, each specific membrane fusion reaction requires a small GTP-binding protein of the rab family. Over 30 rab proteins have been identified in mammalian cells and 11 are known in yeast (reviewed in Aroeti et al., 1998 and Lazar et al., 1997). In Paramecium, PCR studies have identified genes for 15 rab- and rho-like proteins. In Tetrahymena, 11 small GTPases have been localized specifically with isolated phagosomal membranes alone (Meyer et al., 1998). In a very recent study we have cloned a soluble protein that is associated with discoidal vesicles, acidosomes, and lysosomes in Paramecium. Microinjecting the monoclonal antibody specific for this 37-kDa protein into the cell reduced DV formation by 80%, showing that this 37-kDa protein is involved at some step in the recognition, docking, priming, or fusion of discoidal vesicles with the cytopharynx membrane (Yamauchi et al., 1999). Because this protein is located on several types of vesicles (i.e., discoidal vesicles, acidosomes and lysosomes), it probably functions in a more general membrane–membrane binding or fusion step rather than in a membranespecific role such as that played by SNAREs and rabs. Because the 37-kDa protein had little homology with other known proteins, its precise function waits to be determined.

B. Distribution and Routing 1. Long Distance Movements along Microtubules and Microfilaments Appropriately addressed vesicular compartments must be delivered to cognate docking sites at the surface of Paramecium. In most cases microtubules, arising near the cell surface are used to guide these compartments to their docking and fusion sites. Thus bundles of microtubules direct the spent DV to its single exocytic site, the cytoproct, while individual microtubules direct trichocysts to their multiple docking sites. Finally, cytopharyngeal microtubular ribbons direct three different types of membrane-limited compartments toward the cytopharynx—the discoidal vesicles, acidosomes, and 100-nm carrier vesicles.

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The movement of vesicles in Paramecium occurs at different rates. Vesicles moving along cytopharyngeal ribbons move smoothly and rapidly at 5.8 ⫾ 0.9 애m/sec (Schroeder et al., 1990). Moving at this speed are both the discoidal vesicles and acidosomes (100-nm vesicles are too small to be seen in whole-cell video microscopy). Trichocysts, on the other hand, are said to move by saltation with a mean velocity of around 1 애m/sec (Aufderheide, 1977; Glas-Albrecht et al., 1991) which is 20% of the rate of the vesicles moving along the cytopharyngeal ribbons. At present there is no information available on the speed of the movement of spent vacuoles toward the cytoproct. The fastest movement recorded is of the DV-I moving from the cytopharynx to the posterior end of the cell. This occurs at 20 애m/sec (Schroeder et al., 1990). The DV-I follows the path of bundles of microtubules when it is first released from the cytopharynx. However, this movement differs from the other types of movement listed earlier in that the DV-I moves in a plusend direction relative to the origin of the microtubule bundles. Furthermore, it has been shown that an actin-like protein may be involved in this movement (Cohen et al., 1984b). This movement is not restricted just to phagosomes because a large number of vesicles of different sizes continue to stream toward the cell’s posterior end along this same track for several seconds following the passage of the phagosome. Therefore, this type of movement is more nonspecific as to the type of membrane-bound particles that can be moved along the track. Thus, Paramecium has an elaborate distribution and routing system for moving membrane vesicles both along microtubule arrays as well as along an actin-myosin-like track. In most cases, selection of the appropriate vesicles, that are to be transported to a particular docking site at the cell surface, seems to be made at the point of contact between the vesicle and the track of microtubules or microfilaments, and this selection can occur a long way from the final docking site. Thus, the kind of motor that can bind to a vesicle as well to a particular microtubular system or microfilamentous system must determine what vesicles will move along a particular track. In Paramecium we have shown that cytoplasmic dynein is the motor used to move discoidal vesicles and acidosomes, and presumably 100-nm carrier vesicles, along the cytopharyngeal ribbons in a minus-end direction to the cytopharynx (Schroeder et al., 1990; Fok et al., 1994). 3. Controlling Short-Distance Movements In the ER–Golgi transition zone, entrapment of vesicles within a mesh of filaments or tethering of vesicles may ensure proper and efficient routing. Coated transition vesicles initially of 60-nm diameter pinch from the ER and are surrounded by nonetchable filaments. Movement of these vesicles

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toward the cis-face of the Golgi stack within this meshwork seems to be the case because uncoated vesicles (40 nm) are seen lying nearer the Golgi stack. In the endosomal system, preendosomal vesicles are always released in close proximity to the early endosomes with which they fuse. Possibly, the nearby proximity of the early endosome will ensure a high probability that these two compartments will make contact and fuse. Although no special cytoskeletal elements that might ensure fruitful encounters between preendosomal vesicles and the early endosomes have been observed, there is some morphological indication that the early endosomes themselves or the preendosomal vesicles might be kept in the cell’s cortex by microtubules (Allen et al., 1992).

IV. Concluding Remarks By studying the membrane trafficking of a number of membrane pools in a single cell, it can be demonstrated that such a cell is well equipped to (1) retrieve membrane from compartments that have fulfilled their function, (2) recycle this membrane so that it can package newly incoming exogenous material, and (3) modify the membrane as it changes its function during a complex physiological process. Thus membranes that are used are not discarded either by discharge into the environment or by digestion in autolysosomes. The economic advantage to the cell in energy saved that would be needed for new membrane biosynthesis is evident. It is also evident that cells have a unique capacity to select specific membrane compartments from a vast array of vesicles and vacuoles of different sizes and shapes and to be able to move these selected vesicles and vacuoles to only those target membranes that they can recognize and with which they can dock and fuse. Again energy is potentially conserved as the selection of the right vesicles for a particular fusion site is made early so that energy is not wasted in transporting compartments to inappropriate docking sites. The interactions of membranes between the various membrane systems in the cell suggest that many if not all membranes could merge under the right conditions or when supplied with the appropriate recognition proteins and fusion complexes. However, in the cell, fusion of different compartments does not happen indiscriminately. For example, secretory membrane can fuse with lysosomes and carrier vesicles can merge with acidosomes when they encounter each other. However, acidosomes do not fuse with the NDV nor with the DV-I until the DV-I has moved to the cell’s posterior pole, and lysosomes do not normally fuse with acidosomes even though lysosomes will fuse with acidosome-type membrane once the DV-II has

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become lysosome-fusion competent. Another example is that trichocyst membrane will not fuse with the lysosomes until the trichocyst membrane has been retrieved from the plasma membrane. Thus docking and fusion are highly regulated and will probably not occur until the appropriate cognate SNARE proteins in both the acceptor and donor membranes are uncovered and are allowed to interact, as has been postulated for mammalian cells (Schimmo¨ller et al., 1998). The next broad area of research required to further our understanding of membrane trafficking in Paramecium will be an intensive molecular biological study of these fusion complexes. When combined with cell-free studies on isolated membrane compartments, many new insights on membrane trafficking in this cell can be anticipated.

Acknowledgments The authors thank our former students, postdoctoral fellows, and associates for their contributions to our investigations on membrane trafficking over the years. We especially acknowledge the contributions of Dr. C. C. Burgess (Schroeder) for his skill in providing the QF-DE replicas, Dr. Tomomi Tani for providing the summary drawing, and Marilynn S. Aihara, our long-time research associate, for keeping our Lab running and for her biochemical and immunocytochemical expertise. Our research has been funded by a series of NSF grants of which MCB 98 09929 is the most recent. The Biological EM Facility has been partially supported by NIH RCMI grant RR-03061 and has also benefited over the years from several NSF instrumentation grants.

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Role of Natural Benzoxazinones in the Survival Strategy of Plants Dieter Sicker*, Monika Frey†, Margot Schulz‡, and Alfons Gierl† *Institute of Organic Chemistry, University of Leipzig, D-04103 Leipzig, Germany; †Institut fu¨r Genetik, Technische Universita¨t Mu¨nchen, D-85747 Garching, Germany; and ‡Institute of Agricultural Botany, University of Bonn, D-53115 Bonn, Germany

Benzoxazinoid acetal glucosides are a unique class of natural products abundant in Gramineae, including the major agricultural crops maize, wheat, and rye. These secondary metabolites are also found in several dicotyledonous species. Benzoxazinoids serve as important factors of host plant resistance against microbial diseases and insects and as allelochemicals and endogenous ligands. Interdisciplinary investigations by biologists, biochemists, and chemists are stimulated by the intention to make agricultural use of the benzoxazinones as natural pesticides. These natural products are not only constituents of a plant defense system but also part of an active allelochemical system used in the competition with other plants. This review covers biological and chemical aspects of benzoxazinone research over the last decade with special emphasis on recent advances in the elucidation of the biosynthetic pathway. KEY WORDS: Benzoxazinones, Cyclic hydroxamic acids, Acetal Glucosides, Biosynthesis, Defence, Allelopathy. 䊚 2000 Academic Press.

I. Introduction Natural benzoxazinones were discovered 40 years ago in rye when resistance against pathogenic fungi was investigated. The first benzoxazinone aglucone and the first glucoside were reported in two successive papers (Virtanen and Hietala, 1960; Hietala and Virtanen, 1960). The discovery that natural benzoxazinones function as pesticides resulted in interdisciplinary research for understanding all aspects of this unique class of natural products. The natural occurrence of benzoxazinoids and their role in host plant resistance International Review of Cytology, Vol. 198 0074-7696/00 $35.00

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to pests and diseases have been reviewed by several authors (Meyer, 1988; Niemeyer, 1988; Gross, 1989). Structural requirements for the biological activity of benzoxazinoids and possible molecular mechanisms of action have been discussed (Hashimoto and Shudo, 1996). Recently, synthetic approaches to acetal glucosides and aglucones as well as to analogues have been documented (Sicker et al., 1997). A substantial part of benzoxazinoid biosynthesis has been elucidated (Frey et al., 1997). It is the aim of this review to summarize the genetic, biochemical, and cellular information on biosynthesis and to relate this information to plant host defense and allelopathic aspects of benzoxazinone research of the last decade.

II. General Characteristics of Benzoxazinones A. Structure and Distribution Benzoxazinones are stored in the vacuole of plant cells as d-glucosides. Because of active endogenous glucosidases, aglucones are produced when aqueous extracts are made from plant material. The 2-hydroxy-2H-1,4benzoxazin-3(4H )-one skeleton and the d-glucose unit show (2R)-2-웁linkage without exception, although four stereochemical possibilities exist (Fig. 1). Acronyms are used in the literature that are derived from the chemical designation, e.g. DIMBOA, one of the best-studied benzoxazinones, is derived from 2,4-dihydroxy-7-methoxy-2H-1,4-benzoxazin3(4H)-one. The term cyclic hydroxamic acids or hydroxamic acids is often used for these natural products, although some of them do not contain a cyclic hydroxamic acid but the related lactam. However, it is only with the combination of hydroxamic acid with the cyclic hemiacetal unit that these secondary metabolites receive their unique bioactive properties. Furthermore, the bioactivity can be clearly enhanced by a 7-methoxy or 7-hydroxy group as a donor substituent. To avoid misinterpretation of the chemical structures responsible for biological effects, a neutral term like benzoxazinones or benzoxazinoids is used here. A unique feature of benzoxazinones is the presence of a nitrogen atom in the cyclic hemiacetal ring of the aglucone. From the chemist’s point of view, this is the structural source of a certain instability essential to obtain the chemical reactivity required for the defence reaction. An overview on benzoxazinones is given in Fig. 1. Note that TRIBOA has only been detected as an aglucone, not as a corresponding 2-웁-dglucoside, whereas HM2BOA was only described as glucoside. Further-

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FIG. 1 Bezoxazinods from plants. # All aglucones are released from corresponding (2R)-2웁-d-glucosides by enzymatic cleavage. References for isolation of the glucosides are omitted, except for HM2BOA. References:a Virtanen and Hietala (1959),b Niemeyer (1988),c Woodward et al. (1979),d Barria et al. (1992).e Schulz et al. (1994),f Wolf et al. (1985),g Todorova ¨ zden et al. (1992),i Pratt et al. (1995),k Wahlroos and Virtanen (1959),l Harteet al. (1994),h O nstein et al. (1992),m Friebe et al. (1995),n Hedin et al. (1993),o Chatterjee et al. (1990),p Bailey and Larson (1991),q Kamperdick et al. (1997),r Nagao et al. (1985),s Grambow et al. (1986),t Hofman and Masojidkova (1973).

more, the hydroxamic acid ester HDIBOA could only be proved by some spectroscopic methods (Hedin et al., 1993). In the aglucone a racemic mixture of the (2R)- and (2S )-enantiomers is always found with respect to the C(2)UO bond. Benzoxazinones have been predominantly found in genera of the Gramineae. Outside the Gramineae, they have been isolated from various species

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of the Acanthaceae, Ranunculaceae, and Scrophulariaceae (Fig. 1). A systematic search for benzoxazinones has yet to be performed and might identify more species containing these natural products. There is a single report that bezoxazinones are synthezised in bacteria. The antibiotic C-1027, an antitumor chromoprotein has been isolated from Streptomyces globisporus (Otani et al., 1988). The absolute configuration of the C-1027 chromophore has been determined (Fig. 2; Iida et al., 1996). C-1027 belongs to the enediyne antibiotics and contains a 1,4-benzoxazin3(4H )-one structural moiety which resembles the benzoxazinoids from plant origin except for the 2-methylene group in place of the hemiacetalic 2-OH group. Evidence has been presented that the benzoxazinone moiety confers intercalative DNA binding of the C-1027 chromophore (Yu et al., 1995).

B. Degradation of Acetal Glucosides to Benzoxazolin-2-(3H)-ones Benzoxazinoid acetal glucosides are compounds of low toxicity that can undergo enzymatic and chemical degradation. In the intact plant cell, the acetal glucosides and the 웁-glucosidases are stored in two different cell compartments, the vacuole, and the plastid, respectively (Esen, 1992; Babcock and Esen, 1994). The benzoxazinone glucoside content in seedlings can reach concentrations between 1 and 10 mmoles per kilogram of fresh weight (Long et al., 1974; Tang et al., 1975). After cell damage, the glycoside

FIG. 2 C-1027 Chromphor from Streptomyces globisporus.

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and the enzyme are released and the hemiacetalic aglucone is produced (Fig. 3). In maize the enzymatic release of DIMBOA after wounding is complete within half an hour. The aglucone has been found to be the toxic principle against microbial and insect pests (see Section IV). Interestingly, all aglucones containing the 2,4-dihydroxy-2H-1, 4-benzoxazin-3(4H )-one skeleton (i.e., the direct combination of cyclic hydroxamic acid and cyclic hemiacetal unit) have been found to be chemically instable. As a result of the chemical degradation, benzoxazolin-2(3H )-ones are formed in a ring contraction reaction accompanied by the loss of formic acid. The half-life of DIMBOA in the exudate of injured maize cells is about 24 hr (Woodward et al., 1978). In fact, benzoxazinone acetal glucosides were detected from this end of the cascade, starting with the discovery of benzoxazolin-2(3H )-one (BOA) from rye (Virtanen and Hietala, 1955). Meanwhile, several substituted benzoxazolin-2(3H )-ones have been isolated from plants (Fig. 4). The glucosidic origin has only been established in the case of BOA and its 6-methoxy derivative MBOA. Very likely, DMBOA, 4-ABOA, 4-ClDMBOA and 5-Cl-MBOA are also derived from glucosides, although this has yet to be proven.

FIG. 3 Enzymatic and chemical degradation of a benzoxazinoid acetal glucoside.

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FIG. 4 Structure and occurrence of natural benzoxazolin-2(3H )-ones. BOA was isolated from rye (Virtanen and Hietala, 1955; Barnes et al., 1987), from maize (Kosemura et al., 1995), from Blepharis edulis (Chatterjee et al., 1990), and from Aphelandra tetragona (Werner et al., 1993). MBOA was isolated from maize and wheat (Virtanen et al., 1957), from Aphelandra tetragona (Werner et al., 1993), from Coix-lachrymajobi (Nagao et al., 1985), and from Scoparia dulcis (Chen and Chen, 1976; Hayashi et al., 1994). DMBOA, (Klun et al., 1970; Kosemura et al., 1995), 4-ABOA, (Fielder et al., 1994), and 4-Cl-DMBOA, (Kosemura et al., 1995), and 5-Cl-MBOA (Kato-Noguchi et al., 1998) were each isolated from maize.

III. Biosynthesis of Benzoxazinoids The benzoxazinoid pathway has been elucidated in maize. First, the genes encoding enzymes integrated in biosynthesis were isolated. The expression of these genes in microbial systems has enabled the determination of the enzymatic reactions in vitro. Gene isolation was facilitated by the genetic tools that are available for maize and by the substantial biochemical data that had accumulated. The pathway is genetically defined by the Bx1 gene. The maize mutation bx1 (benzoxazineless) abolishes DIMBOA synthesis (Hamilton, 1964). The DIMBOA and tryptophan biosynthetic pathways share certain intermediates because labeled tryptophan precursors such as anthranilic acid (Tipton et al., 1973) and indole (Desai et al., 1996) are incorporated into DIMBOA, although incorporation of labeled tryptophan was not detected. The conversion of HBOA to DIBOA was identified as a cytochrom P450 monooxygenase catalysed reaction (Bailey and Larson, 1991). Later it was determined that all five oxygen atoms of DIMBOA are derived from molecular oxygen (Glawischnig et al., 1997). Therefore, key features of the pathway were expected to be a mechanism that produces indole and the incorporation of oxygen atoms by P450 enzymes in order to convert indole to a benzoxazinone.

A. Isolation and Function of Bx1 The Bx1 gene was molecularly identified by directed transposon tagging using the Mutator (Mu) transposon system of maize (Chomet, 1994). A

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Mu-induced recessive bx1 allele was identified. Subsequently, a genomic DNA fragment flanking the Mu insert was isolated by the so-called AIMS method (a PCR-based method, Frey et al., 1998). This fragment was used to isolate the wild-type Bx1 and the recessive bx1 alleles from genomic lambda-libraries as well as the full length cDNA clones (Frey et al., 1997). Another allele of Bx1 was fortuitously isolated (Kramer and Koziel, 1995) by searching for genes that were not expressed in the maize seed. It was found that Bx1 is similar to tryptophan synthase alpha subunits (TSA) and when expressed in E. coli, Bx1 complemented a bacterial TSA mutation. Therefore, originally this gene was falsely classified as TSA from maize. However, BX1 is not involved in trypthophan biosynthesis but represents the mechanism that produces indole for secondary metabolism (see discussion that follows) and manifests the branch point from primary metabolism for benzoxazinoid biosynthesis (Fig. 5). The bx1 mutation can be complemented with indole (Frey et al., 1997). The immersion of shoots of bx1 seedings into a 1 mM solution of indole restored the formation of DIMBOA. This confirmed that indole is a specific intermediate for DIMBOA biosynthesis. BX1 catalyzes the conversion of indole-3-glycerol phosphate to indole (Frey et al., 1997). BX1 protein was expressed in E. coli and purified to homogeneity. The steady-state kinetic constants (Km 0.013 mM, Kcat 2.8 s⫺1) indicate that BX1 acts efficiently as an indole-3-glycerol phophate lyase (Frey et al., 1997). In a separate study, BX1 enzyme was given the name indole synthase (Melanson et al., 1997). The sequence similarity to TSA suggests that BX1 function originated from the penultimate reaction in tryptophan biosynthesis and was modified during evolution to open the secondary metabolic pathway. These modifications essentially permitted BX1 to act as a free enzyme, whereas tryptophan synthase is typically a tetrameric heterosubunit complex that is formed by two TSA-TSB (tryptophan synthase beta) complexes that are linked via TSB (Creighton and Yanofsky, 1966). These complexes were originally described in bacteria; however, an analogous heterosubunit tryptophan synthase exists in plants (Radwanski et al., 1995). Indole is usually not released from the tryptophan synthase complex but is directly converted to tryptophan. TSA activity is almost completely dependent on complex formation. Tryptophan levels in the bx1 mutants are normal; the natural maize TSA subunit must, therefore, be encoded by gene(s) different from Bx1. The product of these genes would presumably form heterosubunit complexes with TSB for which two genes have been characterized in maize (Wright et al., 1992). The BX1 protein, like tryptophan synthase, is localized in the stroma of the chloroplast. In vitro import experiments with spinach plastids revealed the cleavage of a signal peptid. When the mature BX1 protein is isolated

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FIG. 5 DIMBOA biosynthesis. The branchpoint of DIMBOA and tryptophan biosynthesis is indicated on top. Abbreviations for enzymes are indicated for each reaction (see text). The enzymes for conversion of DIBOA to DIMBOA have not yet been identified.

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from maize, the amino terminus is characteristically shorter than that predicted from the Bx1 cDNA sequence (C. Stettner, unpublished). The synthesis of several other secondary metabolites in plants, such as the indole glucosinates, anthranilate-derived alkaloids, and tryptamine derivatives (Radwanski and Last 1995, Kutchan 1995), could depend on indole as an intermediate. Indole-3-glycerol phosphate was proposed as a branchpoint from the tryptophan pathway for the synthesis of the indolic phytoalexin camalexin (3-thiazol-2’yl-indole) in Arabidopsis thaliana (Tsuji et al., 1993, Zook, 1998). In maize and cotton, indole is produced as part of the volatile mixture or ‘‘cocktail’’ that is released following attack by army beet worm catapillars and other insect larvae (Turlings et al., 1990). The indole-3-glycerol phosphate lyase function of BX1 exemplifies how indole can be generated in plants to serve either as intermediate for secondary metabolism or as an end product.

B. Isolation and Function of Bx2–Bx5 The following reactions in DIBOA biosynthesis are catalyzed by four cytochrome P450-dependent monooxygenases. These enzymes are membranebound heme-containing mixed function oxidases. They use NADPH or NADH to reductively cleave molecular oxygen to produce functionalized organic products and a molecule of water. In this generalized reaction, reducing equivalents from NADPH are transferred to the P450 enzyme via a flavin-containing NADPH-P450 reductase. In plants, P450 enzymes are involved mainly in hydroxylation or oxidative demethylation reactions of a large variety of primary and secondary metabolites including hormones, phytoalexins, xenobiotics, and pharmaceutically relevant compounds. Four maize P450 genes, one of which was isolated by substractive cDNA cloning from high versus low DIMBOA accumulating lines (Frey et al., 1995), are in the CYP71C subfamily of plant cytochrome P450 genes. These genes are strongly expressed in young maize seedlings and share an overall amino acid identity of 45–65%. The observation that all oxygen atoms of DIMBOA are incorporated from molecular oxygen (Glawischnig et al., 1997) led to the speculation that these cytochrome P450 enzymes might be involved in this pathway. The genes encoding these enzymes are designated Bx2, Bx3, Bx4 and Bx5. Direct evidence for the involvement of Bx3 in DIMBOA biosynthesis is provided by a mutant allele (Bx3::Mu) isolated by a reverse genetic approach to screen for Mu insertions in the P450 genes (Frey et al., 1997). In maize seedlings homozygous for the recessive mutant allele, no DIMBOA could be detected by HPLC analysis. In contrast, DIMBOA

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was detected in seedlings that were either heterozygous or homozygous wild type. Thus, a functional Bx3 gene is required for DIMBOA biosynthesis. The cDNAs of the four P450 enzymes were expressed in a yeast system (Truan et al., 1993; Urban et al., 1994). Microsomes were isolated from the transgenic yeast strains in order to demonstrate a function in benzoxazinoid biosynthesis. BX2–BX5 convert indole to DIBOA by catalyzing a series of four specific hydroxylations (Frey et al., 1997, Fig. 5). First, BX2 catalyzes the formation of indolin-2(1H )-one, which is converted to 3-hydroxyindolin-2(1H )-one by BX3. Then, BX4 catalyzes the conversion of 3hydroxy-indolin-2(1H )-one to 2-hydroxy-2H-1,4-benzoxazin-3(4H )-one (HBOA). The mechanism for the unusual ring expansion associated with this reaction is presently being analyzed. Finally, HBOA is hydroxylated by BX5 to DIBOA. Although the four cytochrome P450 enzymes are homologous proteins, they are substrate specific. Only one intermediate in the pathway was converted by each respective P450 enzyme to a specific product. The question of enzyme specificity was addressed with the yeast expression system. The intermediate metabolites of the DIBOA pathway indole, indolin2(1H )-one, 3-hydroxy-indolin-2(1H )-one, and HBOA (Frey et al., 1997) were incubated with microsomal preparations each containing one of the P450 enzymes. No detectable conversions occurred in other enzyme/substrate combinations. Each enzyme is therefore specific for the introduction of only one of the oxygen atoms in the DIBOA molecule. Enzymatic reactions, identical to the ones with the different yeast microsomal preparations, could be performed with maize microsomes, indicating that identical reactions occur naturally in maize. In addition, feeding bx1 mutant seedlings with all the intermediates from indole to DIBOA results in the formation of DIMBOA in the plantlets. These findings suggest that the reaction sequence in maize from indole to DIMBOA is as depicted in Fig. 5. In addition to the reactions already described, 2H-1,4-benzoxazin-3(4H )one and 2-hydroxy-7-methoxy-2H-1,4-benzoxazin-3(4H )-one (HMBOA) were tested as possible substrates for BX2–BX5. The hydroxylation of 2H1,4-benzoxazin-3(4H )-one to HBOA is catalyzed by BX3 (Glawischnig et al., 1999). This reaction is approximately half as efficient as the hydroxylation of indolin-2-one. In both cases, a C atom at an equivalent position is hydroxylated (Fig. 5). In contrast, for HMBOA, a derivative present in maize seedlings, no enzymatic conversions are detectable with BX2–BX5 expressed in yeast or with maize microsomes. In particular, HMBOA is not N-hydroxylated by BX5. This indicates that the 7-methoxy group of HMBOA would interfere with BX5 action. The relatively high specificity of the enzymes seems to support the idea that plant P450s generally have a much greater substrate specificity than their animal homologues. However, there is emerging evidence that

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plant P450s in addition to their normal physiological function, can also convert certain xenobiotics with varying efficiencies. The artificial substrate p-chloro-N-methylaniline (pCMA) is for example efficiently demethylated by BX2 and by several other plant P450 enzymes (Glawischnig et al., 1999). The cellular compartments that comprise the benzoxazinoid biosynthetic enzymes and the glucosidases required for producing the active aglucones have also been identified (Fig. 6). Formation of indole by BX1 takes place in the plastid (C. Stettner, unpublished). The conversion of indole to DIBOA by consecutive oxidation is catalized by BX2–BX5. These P450 enzymes are localized in the endoplasmatic reticulum (Frey et al., 1997). Very likely, further hydroxylation and methylation reactions take also place in the cytoplasm. Biosynthesis commences by glycoylzation followed by transport and storage of the glucosides in the vacuole. The glycosidases required for activation of the glucosides are stored in the plastid (Cicek and Esen, 1998). In the case of cell wounding, the two cellular organelles are damaged, and the toxic aglucones are produced.

C. Genomic Organization of the Bx Gene Cluster Bx2–Bx5 have been grouped into the CYP71C subfamily (Frey et al., 1995) of cytochrome P450-dependent monooxygenases. These genes have probably evolved by duplication events as indicated by the clustering of the genes on the short arm of chromosome four, their similar exon/intron organization, and their sequence homology (Fig. 7). Such a case illustrates that a substantial part of a pathway can evolve by gene duplication followed by sequence modification. The cluster of P450 genes is tightly linked to the Bx1 gene encoding the indoleglycerol-3-phosphate lyase that produces indole which is hydroxylated by the P450 enzyme BX2. The genes of Bx1 and Bx2 are separated by only 2.5 kb. At present, there is no other example of two nonhomologous genes that are comparably closely linked in maize. All genes of the Bx gene cluster are located within 6 cM. These genes will therefore frequently be transferred to the next generation as one functional unit, encoding all enzymes required for the biosynthesis of DIBOA.

D. Evolution of the DIBOA Pathway The benzoxazinoids are widely distributed in grasses and are also found in several dicotyledoneous species (Fig. 1), suggesting that the acquisition of this pathway occurred relatively early in the evolution of the Gramineae

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FIG. 6 Cellular compartimentation of DIMBOA biosynthesis. Indol, synthesized in the chloroplast, is the substrate for P450 enzymes localized in the endoplasmic reticulum. The benzoxazinones are readily glucosylated by cytosolic UDP-glucosyl-transferases and the glucoside is stored in the vacuole. The specific glucosidase is found in the chloroplast. When the structural integrity of the cell is destroyed, the glucosidase and the glucosides encounter each other, and the toxic aglucon is produced.

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FIG. 7 Structure and chromosomal location of the Bx genes in maize. A schematic presentation of the Bx gene cluster at the short arm of chromosome 4 is given; genetic distances are indicated in centimorgans. The exon–intron structure of Bx1 through Bx5 is outlined. Exons are represented by boxes, and the position of translation start and stop codons are indicated. Introns of Bx2–Bx5 are present at equivalent positions but differ in sequence and size.

and probably even before the monocots and dicots diverged. The activity of the DIBOA-specific P450 enzymes has been assayed in two other cereals: rye (containing benzoxazinoids) and barley (without benzoxazinoids). These two species are much more closely related to each other than either is to maize (Devos and Gale, 1997). The predominant benzoxazinoid of rye is DIBOA. As in maize, there are relatively high concentrations (up to 1 mg/g fresh weight) present in the rye seedling. The cytochrome P450-dependent reactions are very similar to those detected in maize. Indole, indolin-2(1H )-one, 3-hydroxy-indolin2(1H )-one, and HBOA were converted to the same products that were obtained with maize microsomes (Glawisching et al., 1999). No additional products were detected. The reactions were strictly dependent on NADPH, indicating true cytochrome P450 enzyme reactions. The similarity of the reactions in maize and in rye suggests identical DIBOA biosynthetic pathways for both species. In microsomes prepared from barley seedlings, no activities of the P450 enzymes of the DIBOA pathway were detectable, although the total P450 content in microsomes and the NADPH-P450 reductase activity were similar to maize (Glawischnig et al., 1999).

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Although maize and rye are distantly related, the DIBOA biosynthetic pathway seems to be identical in both species. Therefore, a set of proteins homologous to BX1–BX5 could be expected to exist in other grasses. If this were the case, the gene duplications responsible for the evolution of the Bx2–Bx5 gene cluster must have occurred early in the development of the Gramineae, possibly even before the devergence of the monocots and the dicots when the presence of benzoxazinones in the Acanthaceae, Ranunculaceae, and Scrophulariaceae is taken into account. The isolation of genes homologs to Bx1–Bx5 from other species will give an insight into the evolution of the Bx gene cluster. Barley microsomes show no significant DIBOA-specific P450 activities. It remains to be shown whether the loss of enzyme activity is due to gene inactivation, or whether the whole Bx2–Bx5 cluster has been lost. This might have occurred during agricultural breeding from wild barley varieties in which DIBOA was initially present (Niemeyer, 1988; Barria et al., 1992). A similar loss of enzyme activity has also been observed for the UDPglucose : DIBOA glycosyltransferase. Glycosyltransferase activity is present in wild varieties and was susequently lost during barley cultivation (Leighton et al., 1994).

IV. Mode of Action Benzoxazinoid Aglucones A. Interaction of Benzoxazinoid Aglucones with Pests Benzoxazinones are part of the so-called nonhost or general defense of plants (Health, 1985). In contrast to specific or cultivar resistance, a wider range of pathogens and pests are controlled by the general toxicity of such ecochemicals. A decrease of pathogen or pest injury has been positively correlated with increased benzoxazinone content by several authors for maize and wheat (Givovich and Niemeyer 1996; references in Niemeyer 1988) and Triticum (Thackray et al., 1990; Niemeyer et al., 1992). High benzoxazinone levels reduce the growth rate and enhance the mortality of bacteria, fungi, and insects; the spread of plant viruses is indirectly affected by controlling their insect hosts (Bravo and Lazo, 1993; Freymark et al., 1993; Givovich and Niemeyer, 1996). The antialgal, antifungal, and antimicrobial potential of benzoxazinones can be demonstrated by including benzoxazinones in the growth media for these organisms (Bravo and Lazo, 1996; Bravo et al., 1997). Interestingly, DIMBOA blocks both virulence and growth of Agrobacteria (Sahi et al., 1990). This protection against infection by tumor-

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inducing bacteria might be one reason why cereals have been recalcitrant to Agrobacteriaum tumefaciens-mediated transformation. The correlation between benzoxazinone content and insect defense in relation to loss of yield is perhaps the most important aspect of study. Two main approaches are used: (i) larval development and insect performance are analyzed on benzoxazinone diets (Argandona et al., 1983, Campos et al., 1989), and (ii) insect damage is monitored in the greenhouse or by field trials on high and low benzoxazinone cultivars (references in Niemeyer, 1988). These investigations have shown that benzoxazinones can act as feed deterrents and reduce the viability of larvae. In maize, the control of the main insect pest, the European corn borer (ECB, Ostrinia nubilalis), can be correlated with high DIMBOA content. The genetic basis of benzoxazinone-mediated resistance against ECB was demonstrated very early on, and the DIMBOA content was successfully elevated in breeding programs (Klun et al., 1970; Grombacher et al., 1989). The proposed major gene on short arm of chromosome 4 and the major quantitative trait locus (QTL) at this position turned out to coincide with the Bx gene cluster. Benzoxazinone-mediated resistance is restricted to the first brood of the European corn borer (Barry et al., 1994). The second brood of ECB can circumvent benzoxazinone control since DIMBOA synthesis and concentration is highest in young plants and is much reduced when the second generation larvae develop. Also in wheat, rye, and perennial triticeae, benzoxazinone levels are highest in the seedling (Copaja et al., 1991a, 1991b). A detailed accumulation pattern has been elucidated in wheat (Iwamura et al., 1996), and the observed benzoxazinone distribution fits exactly with the expression pattern of the genes Bx2–Bx5 (Frey et al., 1995). As would be expected for a defense-related compound, the organs facing the environment like the coleoptile or the root tip have the highest Bx gene activity and benzoxazinone content. In hexaploid and tetraploid Triticum, an elevated benzoxazinone content is not restricted to the seedling, but the concentration is also high in young leaves (Thackray et al., 1990). This finding is paralleled by in situ hybridization work that shows Bx2–Bx5 gene expression not only in the seedling root but also in young side roots and young emerging adventitious roots (M. Frey unpublished). Clearly, an intrinsic ‘‘seedling’’ and ‘‘young tissue’’ expression program exists for benzoxazinone synthesis in the grasses, and protection of these tissues might be especially important in the defense strategy of the plant. In the seedling, the benzoxazinone content and Bx2–Bx5 gene expression is not detectably influenced by wounding. However, a different expression program might exist in older plants. A localized, nonsystemic increase of benzoxazinone was detected in reaction to aphid damage in wheat and wild wheat (Niemeyer et al., 1989; Gianoli and Niemeyer, 1998) and mechanical

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damage in maize (Morse et al., 1991). The induction is variable with respect to the cultivar and requires high levels of aphid infestation.

B. Molecular Mode of Action of Aglucones Several proposals have been made to explain the defense mechanism of benzoxazinoids based on model experiments with DIMBOA and its structural analogs. DIMBOA is known as an enzyme inhibitor of the chloroplast ATPase coupling factor (Queirolo et al., 1983), papain (Perez and Niemeyer, 1989a), 움-chymotrypsin (Cuevas et al., 1990), aphid cholinesterases (Cuevas and Niemeyer, 1993), and plasma membrane H⫹-ATPase (Friebe et al., 1997). DIMBOA, but not its glucoside, has a range of different activities: ● Reduction of the electron transport in both mitochondria and chloro-

plasts of maize (Massardo et al., 1994). ● Stimulation of the rate of NADH oxidation catalyzed by horseradish

peroxidase isoenzyme C that catalyzes H2O2 formation, required for oxidative cross-linking of polysaccharides and proteins of the cell wall (Rojas G. et al., 1997). ● Reduction of the activity of glutathione S-transferase of the grain aphid, Sitobion avenae (Leszczynski and Dixon, 1992). It was suggested that these effects may result from reaction with NH2 and SH nucleophilic groups of biomolecules. The oxo-cyclo tautomerism of DIMBOA offers two possibilities for a structural understanding. (i) The aldehyde group of the oxo form should be a strong electrophile. It has been shown that it reacts with the ␧-NH2 group of N-움-acetyl lysine used as a model for lysine residues of proteins (Perez and Niemeyer, 1989b). (ii) Toward mercapto groups, the hemioxidized cyclic hydroxamic acid may behave as an oxidant, with the corresponding lactam as a reduction product. Hence, reactions of DIMBOA with mercaptanes have been studied in detail (Perez and Niemeyer, 1985); 2,4-dihydroxy-2H-1,4-benzoxazin-3(4H )-ones substituted at the aromatic ring with 2-mercaptoethanol gave evidence that an electron-donating substituted at position 7 ( para to the hydroxamic acid N atom) is required to obtain reduction (Atkinson et al., 1991). Siderophores of microorganisms of the hydroxamate type are known to be efficient carriers of Fe3⫹ ions in the form of iron complexes from the soil into the microorganisms. Benzoxazinoids secreted by maize, wheat, and rye seedlings have a phytosiderophore-like function (Petho¨ et al., 1997). The importance of the substitution pattern of DIMBOA was studied by removing either the 2-OH group or the 7-MeO group with concomittant activation of the analogs by acetylation at N&B1;OH. Both model

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compounds, 4-acetoxy-7-methoxy-2H-1,4-benzoxazin-3(4H )-one and 4acetoxy-2-hydroxy-2H-1,4-benzoxazin-3(4H )-one, were shown to react with various C-, N-, or S-nucleophiles, regioselectively (Hashimoto et al., 1991). Also DNA reacted covalently with the hydroxamic acid N atom (Ishizaki et al., 1982). The results suggest that DIMBOA, after metabolic O-acylation, may act as an alkylating agent toward biomolecules, like proteins or nucleic acids, due to the hemiacetal and the 7-donor-activated cyclic hydroxamic moiety (Hashimoto and Shudo, 1996). Recently, it was reported that a DIMBOA dehydration product is a reactive formyl donor (Hofmann and Sicker, 1999). Therefore formylation could also contribute to the biological effects of benzoxazinoids. In summary, the high reactivity of DIMBOA is the basis for the wide antimicrobial and mutagenic activities of benzoxazinoids.

V. Molecular Allelopathy of Benzoxazinones and Benzoxazolinones Allelopathic interactions between individuals of different plant species or those of the same species are caused by plant-produced allelochemicals. Once released into the environment, passively or actively, they can influence germination, growth, and development of neighboring plants either negatively or positively (Harborne, 1977; Rice 1984). Most allelochemicals are characterized by multifunctional phytotoxicity and are often also important for general defense. The allelopathic potential of cereals has been extensively studied in rye. A dramatic reduction (more than 90%) of the biomass of dicotyledonous weeds, such as Chenopodium album by rye mulch (Putnam and DeFrank, 1983) and living rye plants was reported by Perez and Ormeno-Nunez (1993). In 1987, Barnes et al. identified DIBOA and BOA as first-order reaction allelochemicals in rye. According to the calculations of Barnes et al. (1986), 35-day-old rye can release 14.3 kg/ha of DIBOA. Fresh rye mulch contains 20–50 mmol benzoxazinoids. After 12 days, this concentration is reduced to half, and after about four months benzoxazinoids are no longer detectable (Yenish et al., 1995). Rye actively releases the DIBOA into the environment via root exudation. The exudate should influence the species composition of the neighboring vegetation and the composition of rhizosphere-colonizing microorganisms. DIBOA exudation was also observed with Agropyron repens (common couch grass), another member of the grass family. Here, phytotoxic effects were highest in exudates of young rhizome-borne roots, that

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appear during spring time. The effects decreased with increasing age of the roots (Schulz et al., 1994). The position and the nature of substitutions of benzoxazolinone molecules may influence effects on monocots and dicots. Barnes et al. (1987) observed that DIBOA had a significant effect on monocots. Perez and Ormeno-Nunez (1991) demonstrated a 100% suppession of Avena fatua radicle growth in the presense of 1 mM MBOA. In a recent study, KatoNoguchi et al. (1998) reported on 5-chloro-6-methoxy-benzoxazolin-2(3H )one as a potent growth inhibitor of several Gramineae. The relatively high capacity of certain Gramineae to detoxify benzoxazinoids could reflect the abundance of these secondary products in this family. The biosynthesis of endogenous benzoxazinoids commences with glycosylation and storage of the compounds in the vacuole. It seems that the same mechanism is used for detoxification of exogeneous benzoxazinoids (Fig. 8). Glycosylation of BOA was observed when BOA uptake experiments with oats, wheat, rye, and corn were analyzed (Wieland et al., 1998, 1999). BOA-6-O-glucoside was the major metabolite in oat, wheat, and rye. In maize, however, BOA-N-glucoside was the dominant

FIG. 8 Metabolization products of BOA. References for plants (Wieland et al., 1998, 1999) and for soil bacteria (Chase et al., 1991, 1991b; Friebe et al., 1996; Gerber and Le Chevalier, 1964; Kumar et al., 1993; Nair et al., 1990).

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metabolite. Whereas BOA-N-glucoside shows no phytotoxicity, BOA-Oglucoside is still inhibitory but drastically lower than BOA. Thus, BOAN-glucoside presents a true detoxification product, particularly as it is not hydrolysable by common 웁-glucosidases. In contrast, BOA-6-O-glucoside can be easily hydrolyzed, resulting in highly toxic BOA-6-OH. However, in the plant cell, the O-glucoside is certainly transported into and stored within the vacuole. Generally, the phytotoxic effects of benzoxazinoids depend on the dose and the target species, with monocots showing a lower sensitivity compared to dicots. Avena sativa, Avena fatua, and Triticum aestivum were compared with the dicot Vicia faba. Roots or whole plants were incubated in the presence of BOA (Wieland et al., 1999). The four species were able to absorb the substance when 100 애M were applied, however with different kinetics (Table I). With 500-애M BOA, the root tips of Vicia faba were damaged during the course of incubation, showing extensive blackening. The cereals were still able to absorb BOA, Triticum aestivum without a lag phase, and Avena sativa after 10–15 hr. Microorganisms that live within the rhizosphere or colonize roots seem to be also an important factor in overcoming the phytotoxic effects of BOA. The benzoxazolinone was converted to 2-amino-H-phenoxazin-3one and 2-acetylamino-3H-phenoxazin-3-one (Friebe et al., 1996) by bacteria present on the root surfaces of oats and rye (Table I). Both substances are known as the natural antibiotics questiomycin and Nacetylquestiomycin (Gerber and LeChevalier, 1964). BOA-mediated growth inhibition was reduced in the absence of antibiotics, when phenoxazinone production was allowed. Phenoxazinones can also be produced by bacteria associated with the roots of several dicotyledonous species (Schulz and Wieland, 1999). The different detoxification capacities seem to explain the variation in sensitivity to BOA observed in the plant species. Constitutive and inducible enzymes involved in detoxification reactions must be regarded as causative factors. The ability to detoxify may not always be essential, but it is an essential function during germination or seedling development, which are very sensitive stages in the life cycle of the plant. In a balanced plant community, individuals must endure allelochemicals released by neighboring plants of the same or of different species. There is a certain selection pressure for taxa of the community to detoxify characteristic allelochemicals highly abundant in that plant association. This can be regarded as a co-evolutionary process that comprises a whole plant community. In order to test this hypothesis, 15 species were chosen and evaluated (Schulz and Wieland, 1999): seven species of field weed communities (vegetation class Secalieta); three species of hoed vegetable communities (vegetation class Chenopodietea); one species belonging to hoed vege-

TABLE I Detoxification Capacity of Avena sativa, Triticum aestivum, Avena fatua, and Vicia faba

BOA uptake

Exudation Products in root extracts

Avena sativa

Triticum aestivum

Avena fatua

Vicia faba

Fast 0.1 mM in ⬍ 3 days, 0.5 mM in ⬍ 5 days (with antibiotica) BOA-6-OH BOA-6-OH, BOA6-O-glucoside, BOA-N-glucoside

Fast 0.1 mM in ⬍ 4 days, 0.5 mM in ⬍5 days (without antibiotica) BOA-N-glucoside BOA-6-OH, BOA-6O-glucoside, BOAN-glucoside

Fast 0.5 mM ⬍ 4 days phenoxazinone production in absence of antibiotica

Slow 0.1 mM ⬍ 5 days root tip blackening in presence of 0.5 mM Not detectable BOA-6-OH traces of BOA-6-Oglucoside after 10 days of incubation with 0.1 mM BOA

(Wieland et al. 1999).

BOA-6-OH, BOA-N-glucoside BOA-6-OH, BOA-6-O-glucoside, BOA-N-glucoside

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table communities but occurring in grain field weed communities as well; two species of the vegetation subclass Artemisietea vulgaris; Plantago major, a member of Agrostietea stolonifera communities; and Galinsoga ciliata, native to the Andes and a neophyte in Germany since 1850. Data obtained resulted in four groups of species with high to low detoxification capacity (Table II): Group I consists of Plantago major and Coriandrum sativum. Both species exhibited a high detoxification capacity but are in some respect exceptional: Plantago major is a perennial species in contrast to the others, and Coriandrum sativum is a cultivated plant that occasionally behaves as a weed in wheat fields. Group II represents species of grain field weeds and Artemisietea vulgaris communities with a rather good detoxification capacity. Carduus nutans and Daucus carota often grow closely associated with Agropyron repens, which exudes DIBOA, at least temporarily. The other species are weeds occurring in wheat and rye fields. Group III, with Consolida regalis, Agrostemma githago, Capsella bursapastoris, and Legousia speculum veneris, is a group with moderate to low detoxification capacity. Capsella bursa-pastoris and Chenopodium album (group IV), can be found only occasionally in wheat fields, although they belong to Chenopodietea communities. Both species are able to perform some N-glucosylation. Chenopodium album was integrated into group IV, because N-glucosylation broke down at higher BOA concentrations. Polygonum aviculare, Urtica urens, and Galinsoga ciliata in group IV are characterized by a low detoxification potential and cannot perform N-glucosylation, but O-glucosylation is detectable. Polygonum aviculare represents an exception in group IV because it can appear in both Secalietea and Chenopodieta vegetation classes but lacks the ability to perform N-glucosylation. Polygonum aviculare and Chenopodium album have, however, a high allelopathic potential and may inhibit hydroxamic acid containing species in their close neighborhood. In summary, there seems to be indeed a correlation between the occurrence of a species in certain plant communities and the existence of effective metabolization pathways that result in detoxification products. Evidently, the ecobiochemical potential of a species to detoxify benzoxazolinone can be regarded as an essential secondary function in rye and wheat fields, which is manifested in those European plant associations. Exact explanations at the molecular level for the differences in species reactions to allelochemicals are not common. Benzoxazinoids are, however, an example where biochemical and physiological mechanisms that improve our knowledge about allelochemical interactions have been identified. The understanding of molecular aspects of allelopathic interactions is a prerequisite if the application of allelochemicals in modern agrotechnology is to be considered.

TABLE II Metabolization Capacity of Species Belonging to Different Vegetation Classes (European Plant Communities)

I

II

III

IV

Vegetation class

Species

Family

Agrostietea stolonifera Secalietea Secalietea Artemisietea vulgaris Secalietea Secalietea Artemisietea vulgaris Secalietea Secalietea Chenopodietea Secalietea Chenopodietea Chenopodietea Chenopodietea Neophyte

Plantago major Coriandrum sativum Centaurea cyanus Carduus nutans Papaver rhoeas Matricaria chamomilla Daucus carota Consolida regalis Agrostemma githago Capsella bursa-pastoris Legousia speculum-veneris Chenopodium album Polygonum aviculare Urtica urens Galinsoga ciliata

Plantaginaceae Apiaceae Asteraceae Asteraceae Papaveraceae Asteraceae Apiaceae Ranunculaceae Caryophyllaceae Brassicaceae Campanulaceae Chenopodiaceae Polygonaceae Urticaceae Asteraceae

Metabolization capacity High High High to moderate High to moderate Moderate Moderate Moderate Moderate to low Moderate to low Moderate to low Moderate to low Low No BOA-N-glucoside No BOA-N-glucoside No BOA-N-glucoside

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VI. Concluding Remarks Benzoxazinones are an important component of general defense in certain plants. Substantial data concerning the biosynthesis, its realization in cellular compartments, the temporal and spatial expression pattern, the molecular mode of action, and the allelopathic interactions have accumulated for these secondary metabolites. There are also indications as to how the pathway could have evolved. Some enzymes still remain to be isolated and characterized. Completely absent from this relatively complex picture are data about the factors that govern the concerted expression of the biosynthetic genes during plant development. This information could be relevant for elevating benzoxazinone levels and extending the expression of the biosynthetic genes into older developmental stages. The biosynthetic pathway to DIBOA is relatively short and begins with indole-3-glycerol phosphate, a metabolite ubiquitous to plants. Therefore, DIBOA biosynthesis could be transgenically introduced into other plant species. If successful, such species would gain a powerful component of general defense that should improve their disease resistance against a wide range of pathogens.

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SUBJECT INDEX

A

yeast vacuole targeting pathway, 173 nondegradative pathways, 179–191 ALP pathway, 185–187 genetic analysis, 186 targeting signal analysis, 186–187 vps mutants, 185–186 Cvt pathway, 179–185 aminopeptidase I localization, 179–181 Apg protein role, 184–185 autophagic defective mutants, 182–183 Aut protein role, 184–185 Cvt protein role, 184–185 genetic analysis, 182–183 macroautophagic overlap, 183–184 morphological analysis, 183–184 prAPI delivery, 181–182 vesicle-mediated mechanisms, 181–182 folded protein pathways, 188 MVB sorting pathway, 187–188 overview, 153–154, 191 retrograde transport, 154–165 coat proteins role, 155–156 endosome-to-Golgi transport, 162–163 function, 156 Golgi-to-endoplasmic reticulum transport, 156–159 anterograde transit components recycling, 156–158 coatomer function, 158–159 endoplasmic reticulum resident protein retrieval, 156–158

ABC transporters, channel-mediated transport mechanisms, 190–191 Acidosomes, membrane trafficking in Paramecium, 297–298, 311 Actin, transepithelial water transport mechanisms, 250–251 Aglucone, plant survival role, 319, 332–335 ALP pathway, 185–187 genetic analysis, 186 targeting signal analysis, 186–187 vps mutants, 185–186 Alternative protein-sorting pathways, 153–191 channel-mediated transport, 188–189 ABC transporters, 190–191 endoplasmic reticulum dislocation, 189 degradative pathways, 165–178 autophagosomes role, 178 KFERQ-mediated pathway, 172–173 macroautophagy, 165–172 molecular component delivery, 166–172 physiological role, 172 mitochondria delivery to vacuoles, 178 peroxisome delivery to vacuoles, 175–178 in Hansenula polymorpha, 177 in Pichia pastoris, 175–177 in Saccharomyces cerevisiae, 177–178 starvation-response pathway, 172–173 ubiquitin-mediated endocytosis, 173–175 coordination, 174–175 plasma membrane protein degradation, 174

347

348 Alternative protein-sorting pathways (continued ) intra-Golgi transport, 159–162 cisternal maturation model, 161–162 vesicle transport model, 160–161 vacuole-to-endosome transport, 163–165 vesicle formation, 155–156 Aminopeptidase I, in Cvt pathway, 179–181 Anterograde protein transport membrane trafficking in Paramecium, 306–307 retrograde transport compared, 154–165 coat proteins role, 155–156 endosome-to-Golgi transport, 162–163 function, 156 Golgi-to-endoplasmic reticulum transport, 156–159 anterograde transit components recycling, 156–158 coatomer function, 158–159 endoplasmic reticulum resident protein retrieval, 156–158 intra-Golgi transport, 159–162 cisternal maturation model, 161–162 vesicle transport model, 160–161 vacuole-to-endosome transport, 163–165 vesicle formation, 155–156 Antero-posterior axis, somite formation, 14–18 intrinsic determination, 16–17 polarity relationship, 17–18 resegmentation theory, 15–16 sclerotome compartmentalization, 14–16 Antidiuretic hormone, transepithelial water transport role, 204, 207–209, 252–253 Apg protein, in Cvt pathway, 184–185 Apical membrane, transepithelial water transport mechanisms, 224–240 leaky epithelia, 224–227, 236–237 structural change dynamics during transport, 226–232 aquaporins role, 232–240 intramembrane particle aggregate sources, 230–231 tight epithelia, 225–232, 237–240 Aquaporins, transepithelial water transport role, 232–240 Autophagy, protein sorting mechanisms degradative pathways autophagosomes role, 178 macroautophagy, 165–172

SUBJECT INDEX nondegradative pathways, Cvt pathway, 182–184 Aut protein, Cvt pathway, 184–185

B Benzoxazinones, plant survival role, 319–341 acetal glucoside degradation to benzoxazolin-2-(3H)-ones, 322–324 aglucone action mechanisms, 332–335 benzoxazinoid aglucone action mechanisms, 332–335 biosynthesis, 324–332 Bx2-Bx5 isolation function, 327–329 Bx1 isolation and function, 324–327 Bx gene cluster organization, 329 DIBOA pathway, 329–332 distribution, 320–322 molecular allelopathy, 335–340 overview, 319–320, 341 structure, 320–322 bHLH transcription factor, somite formation role, 13 BMP4, medio-lateral polarity in somitogenesis, 42–47 concentration-dependent mechanisms, 42–43 indirect medial somitic identity promotion, 46–47 Noggin antagonism, 43–46

C Calcium, accumulation in transepithelial water transport, 254 Cellular transport, see Transepithelial water transport Channel-mediated protein transport, 188–189 ABC transporters, 190–191 endoplasmic reticulum dislocation, 189 Chorion genes, Drosophila eggshell morphogenesis role cloning, 76–77 conceptual translation products, 81–82 genetic analysis, 75–76

SUBJECT INDEX Chorion proteins, Drosophila eggshell morphogenesis role biochemical analysis, 73–74 distribution changes in assembly process, 92–95 posttranslational processing, 88–89 Clock theories, in somitogenesis, 10–11 Coat proteins, in vesicle-mediated protein transport, 155–159 Conceptual translation products, in Drosophila eggshell morphogenesis, 79–85 chorion genes, 81–82 dec-1 gene, 82–85 vitelline membrane genes, 79–80 Cvt pathway, 179–185 aminopeptidase I localization, 179–181 Apg protein role, 184–185 autophagic defective mutants, 182–183 Aut protein role, 184–185 Cvt protein role, 184–185 genetic analysis, 182–183 macroautophagic overlap, 183–184 morphological analysis, 183–184 prAPI delivery, 181–182 vesicle-mediated mechanisms, 181–182 Cytochrome P-450, cellular biology in liver cells, 109–140 degradation regulation, 123–126 endoplasmic reticulum membrane topology analytical models, 127–128 densities, 129–131 endoplasmic reticulum proliferationP450 induction relationship, 133–134 image analysis, 130 immunoelectron microscopy models, 128–129 insertion into endoplasmic reticulum membrane, 126–127 localization, 127–129 microphotometry models, 129–130 P-450 reductase interactions, 131–132 overview, 109–110, 140 posttranscriptional regulation, 123–126 posttranslational regulation, 123–126 sublobular expression, 134–140 isoforms, 135–139 postnatal development, 139–140 total P-450, 134–135

349 transcriptional regulation, 110–126 translational regulation, 123–126 specific isoform characteristics CYP1A1/2, 123–124, 130, 135–137 CYP1A1, 111–114, 123, 133, 136, 139 CYP1A2, 114, 123–124, 133, 135–136, 139 CYP1B1, 111, 114, 137 CYP2A6, 136 CYP2B1/2, 111, 130, 135–138 CYP2B1, 114–116, 123–124, 136–137 CYP2B2, 114–116, 123–124, 137, 139 CYP2B4, 127–128 CYP2C1, 127 CYP2C2, 111, 116, 127 CYP2C6, 111, 116 CYP2C9, 116–117 CYP2C11, 111, 117 CYP2C12, 111, 117 CYP2C13, 111, 118 CYP2D1, 118 CYP2D5, 111 CYP2D9, 111, 118 CYP2E1, 111, 118–119, 124–125, 128, 135–138 CYP3A1/2, 125, 136 CYP3A1, 111, 119–120, 125–126, 136 CYP3A2, 125, 137–138 CYP3A4, 111, 120 CYP3A5, 111, 120 CYP3A7, 111, 120 CYP3A10, 111, 120 CYP3A23, 111, 119–121 CYP4A1, 111, 121–122, 124 CYP4A4, 122 CYP4A6, 111, 121–122 CYP7A, 111, 122 CYP11A, 110 CYP11B, 110 CYP27, 110 CYP51, 122 Cytochrome P-450 reductase, endoplasmic reticulum membrane topology in liver cells, cytochrome P-450 interactions, 131–132 Cytoskeleton, transepithelial water transport mechanisms, 250–255 cortical actin participation, 250–251 microtubules participation, 251–255 antidiuretic hormone role, 252–253 calcium ion accumulation, 254

350

SUBJECT INDEX

Cytoskeleton (continued ) giant vacuole migration, 254–255 granular cells in frog urinary bladder, 252–253 high water permeability, 252 low water permeability, 252 thick microtubules, 253–255 tubulin polymerization, 254 water channel protein docking and insertion, 250–251

D dec-1 gene, Drosophila eggshell morphogenesis role cloning, 78–79 conceptual translation products, 82–85 genetic analysis, 75–76 dec-1 protein, Drosophila eggshell morphogenesis role posttranslational processing, 85–87 sequestration and trafficking assembly process, 95–99 Degradative protein-sorting pathways, see Protein degradation Diffusion, see also Transepithelial water transport barriers, tight junction role, 220–224 permeability regulation in leaky epithelia, 221–223 water absorption, 223–224 2,4-Dihydroxy-7-methoxy-2H-1,4benzoxazin-3(4H)-one, see Benzoxazinones DIMBO, see Benzoxazinones Dorso-ventral patterning, in somitogenesis, 18–37 control mechanisms, 23–29 dorsalizing controls, 25–27 ectoderm role, 25–29 floor plate role, 24–25, 27–29 neural tube role, 25–29 notochord role, 24–25, 27–29 ventralizing controls, 24–25 lineage segregation, 18–20, 29–37 dermis formation, 18–19 external cues, 20 skeleton formation, 18–19 myogenic differentiation, 27–29, 33–37 self differentiation, 20–23 cell-autonomous mechanisms, 23

commitment evolution, 21–22 environmental signals, 22–23 historical perspectives, 20–21 Shh protein role, 29–36 spatial regulation, 21–22, 34–37 temporal regulation, 21–22, 34–37 Wnt protein role, 29–30, 33–35 Drosophila, eggshell morphogenesis, 67–104 assembly mechanisms, 89–102 chorion protein distribution changes, 92–95 dec-1 protein sequestration and trafficking, 95–99 dorsal appendages formation, 90–92 eggshell stabilization, 100–102 follicle cell migration, 89 micropyle formation, 90 multilayered mainshell production, 92–99 operculum formation, 90–92 quantitative considerations, 99–100 chamber development, 69 chorion genes cloning, 76–77 conceptual translation products, 81–82 chorion proteins distribution changes in assembly process, 92–95 posttranslational processing, 88–89 conceptual translation products, 79–85 dec-1 gene cloning, 78–79 conceptual translation products, 82–85 dec-1 protein posttranslational processing, 85–87 sequestration and trafficking assembly process, 95–99 gene cloning, 76–79 overview, 67–68, 102–104 proteins biochemical analysis, 72–74 genetic analysis, 75–76 posttranslational processing, 85–89 radial complexity, 69–72 regional complexity, 72–73 vitelline membrane genes cloning, 77–78 conceptual translation products, 79–80 vitelline membrane proteins, posttranslational processing, 87–88

351

SUBJECT INDEX

E Ectoderm myogenic differentiation role, 27–29 in somitogenesis dorso-ventral patterning, 25–29 medio-lateral polarity, 41 Eggshell morphogenesis, see Drosophila, eggshell morphogenesis Endoplasmic reticulum alternative protein-sorting pathways channel-mediated transport, dislocation, 189 retrograde transport from Golgi, 156–159 anterograde transit components recycling, 156–158 coatomer function, 158–159 endoplasmic reticulum resident protein retrieval, 156–158 membrane topology in hepatocytes, cytochrome P-450 biology analytical models, 127–128 densities, 129–131 endoplasmic reticulum proliferationP450 induction relationship, 133–134 image analysis, 130 immunoelectron microscopy models, 128–129 insertion into endoplasmic reticulum membrane, 126–127 localization, 127–129 microphotometry models, 129–130 P-450 reductase interactions, 131–132 membrane trafficking in Paramecium, transition zones, 279–283 Endosomes, trafficking in Paramecium, 290–294 carrier vesicles, 293–294 coated pit, 290–291 early endosomes, 291–294 endocytosis, 290–291 exocytosis sites, 294 noncoated endocytosis, 294 parasomal sac, 290–291, 294 preendosomal vesicles, 291–293 Epithelia, water transport, see Transepithelial water transport Extracellular trafficking, dec-1 protein in Drosophila eggshell morphogenesis, 95–99

F Floor plate myogenic differentiation role, 27–29 somite dorso-ventral patterning role, 24–25, 27–29 Follicle cell migration, in Drosophila eggshell morphogenesis, 89

G Gene cloning, in Drosophila eggshell morphogenesis, 76–79 chorion genes, 76–77 dec-1 gene, 78–79 vitelline membrane genes, 77–78 Golgi complex membrane trafficking pathway stack membranes, 281 trans-Golgi network, 283–284 retrograde protein transport mechanisms endosome-to-Golgi transport, 162–163 Golgi-to-endoplasmic reticulum transport, 156–159 anterograde transit components recycling, 156–158 coatomer function, 158–159 endoplasmic reticulum resident protein retrieval, 156–158 intra-Golgi transport, 159–162 cisternal maturation model, 161–162 vesicle transport model, 160–161 transepithelial water transport mechanisms, 241–250 enterocytes in rat, 242–243 epithelial cells, 241–242 giant vacuole evolution, 249–250 granular cell giant vacuole and protozoan contractile vacuole homology, 249 urinary bladder in frog, 242–248 Granular cells, transepithelial water transport mechanisms giant vacuole and protozoan contractile vacuole homology, 249 microtubules participation, 252–253

H Hansenula polymorpha, peroxisome degradation pathways, 177

352

SUBJECT INDEX

Hemoproteins, see Cytochrome P-450 Hepatocytes, cytochrome P-450 cellular biology, 109–140 degradation regulation, 123–126 endoplasmic reticulum membrane topology analytical models, 127–128 densities, 129–131 endoplasmic reticulum proliferationP450 induction relationship, 133–134 image analysis, 130 immunoelectron microscopy models, 128–129 insertion into endoplasmic reticulum membrane, 126–127 localization, 127–129 microphotometry models, 129–130 P-450 reductase interactions, 131–132 overview, 109–110, 140 posttranscriptional regulation, 123–126 posttranslational regulation, 123–126 sublobular expression, 134–140 isoforms, 135–139 postnatal development, 139–140 total P-450, 134–135 transcriptional regulation, 110–126 translational regulation, 123–126

I Intracellular protein degradation, see Protein degradation Intramembrane particles, transepithelial water transport role, 230–231 aggregate internalization, 231–232 cytoplasmic sources, 230–231

K KFERQ-mediated pathway alternative protein-sorting pathways, degradative pathways, 172–173 protein degradation, 172–173

L Lateral plate, medio-lateral polarity in somitogenesis, 40–43 antagonism, 40–41 dorso-ventral-medio-lateral linkage, 41 trunk versus limb hypaxial lineage specification, 41–42 Liver cells, see Hepatocytes

M Macroautophagy, see Autophagy Medio-lateral polarity, in somitogenesis, 37–47 BMP4 role, 42–47 dorso-ventral-medio-lateral linkage, 41 ectoderm role, 41 external controls, 39–40 gastrulation, 39 lateral plate role, 40–43 lateral sclerotome, 47 lineages, 37–40 muscular differentiation, 38–39 Noggin role, 43–46 rib origin, 47 Shh pathway, 45–46 trunk versus limb hypaxial lineages, 41–42 Wnt pathway, 45–46 Membrane trafficking, 277–312 anterograde movements, 306–307 biosynthetic pathway, 279–285 docking mechanisms, 285 endoplasmic reticulum transition zones, 279–283 filaments, 281–283 fusion mechanisms, 285 Golgi stack membranes, 281 recognition mechanisms, 285 trans-Golgi network, 283–284 budding, 304–309 coated membranes role, 306–307 cognate membrane recognition, 308–309 distribution and routing, 309–311 long distance movements, 309–310 microfilaments role, 309–310 microtubules role, 309–310

353

SUBJECT INDEX short distance movement regulation, 310–311 docking mechanisms, 285, 308–309 endosomal trafficking, 290–294 carrier vesicles, 293–294 coated pit, 290–291 early endosomes, 291–294 endocytosis, 290–291 exocytosis sites, 294 noncoated endocytosis, 294 parasomal sac, 290–291, 294 preendosomal vesicles, 291–293 fusion mechanisms, 285, 308–309 fusogenic property changes, 307–308 membrane recycling, 307–308 modulation, 304–309 overview, 277–279, 311–312 phagosomal system, 294–304 acidosomes, 297–298 cytoproct role, 303–304 digestive vacuole passage, 298 discoidal vesicles, 294–296 membrane replacement, 298–301 phagoacidosome into phagolysosome conversion, 301–303 phagosomal membrane precursors, 294–296 phagosome formation, 297 phagosome into phagoacidosome conversion, 298–301 spent vacuole formation, 303–304 vacuole membrane retrieval, 303–304 priming mechanisms, 308–309 quick-frozen deep-etched imaging, 305 retrograde movements, 307–308 sorting events, 304–309 transport systems, 304–309 trichocyst secretion, 285–290 characteristics, 285–287 constitutive secretion, 290 docking, 287–289 fusion, 287–289 membrane retrieval, 289–290 transport, 287 tubulation, 304–309 Mesp1 and 2 transcription factors, somite formation role, 13 Micropyle, formation in Drosophila eggshell, 90

Microtubules membrane trafficking role in Paramecium, 309–310 transepithelial water transport mechanisms, cytoskeleton role, 251–255 antidiuretic hormone role, 252–253 calcium ion accumulation, 254 giant vacuole migration, 254–255 granular cells in frog urinary bladder, 252–253 high water permeability, 252 low water permeability, 252 thick microtubules, 253–255 tubulin polymerization, 254 Mitochondria, autophogosome-mediated degradation, 178 Molecular clock theories, in somitogenesis, 10–11 Morphogenesis, see Drosophila, eggshell morphogenesis Muscle development, see Myogenesis MVB sorting pathway, nondegradative protein sorting, 187–188 Myogenesis dorso-ventral patterning, lineage segregation, 18–19 initiation, 27–29, 33–34 lineage differentiation, 37–40 myogenic factor activity regulation, 36–37

N Neural tube myogenic differentiation role, 27–29 somite dorso-ventral patterning role, 25–29 Noggin, medio-lateral polarity in somitogenesis, BMP4 antagonism, 43–46 Nonclassical protein targeting, see Alternative protein-sorting pathways Notch signaling pathway, in somitogenesis dorso-ventral patterning, 34–35 somite formation, 11–13 Notochord myogenic differentiation role, 27–29 somite dorso-ventral patterning role, 24–25, 27–29

354

SUBJECT INDEX

O Occluding junctions, see Tight junctions Oogenesis, eggshell morphogenesis, see Drosophila, eggshell morphogenesis Operculum, formation in Drosophila eggshell, 90–92 Osmotic balance, see Transepithelial water transport

P P-450, see Cytochrome P-450 Paramecium, membrane trafficking, 277–312 anterograde movements, 306–307 biosynthetic pathway, 279–285 docking mechanisms, 285 endoplasmic reticulum transition zones, 279–283 filaments, 281–283 fusion mechanisms, 285 Golgi stack membranes, 281 recognition mechanisms, 285 trans-Golgi network, 283–284 budding, 304–309 coated membranes role, 306–307 cognate membrane recognition, 308–309 distribution and routing, 309–311 long distance movements, 309–310 microfilaments role, 309–310 microtubules role, 309–310 short distance movement regulation, 310–311 docking mechanisms, 285, 308–309 endosomal trafficking, 290–294 carrier vesicles, 293–294 coated pit, 290–291 early endosomes, 291–294 endocytosis, 290–291 exocytosis sites, 294 noncoated endocytosis, 294 parasomal sac, 290–291, 294 preendosomal vesicles, 291–293 fusion mechanisms, 285, 308–309 fusogenic property changes, 307–308 membrane recycling, 307–308 modulation, 304–309 overview, 277–279, 311–312

phagosomal system, 294–304 acidosomes, 297–298 cytoproct role, 303–304 digestive vacuole passage, 298 discoidal vesicles, 294–296 membrane replacement, 298–301 phagoacidosome into phagolysosome conversion, 301–303 phagosomal membrane precursors, 294–296 phagosome formation, 297 phagosome into phagoacidosome conversion, 298–301 spent vacuole formation, 303–304 vacuole membrane retrieval, 303–304 priming mechanisms, 308–309 quick-frozen deep-etched imaging, 305 retrograde movements, 307–308 sorting events, 304–309 transport systems, 304–309 trichocyst secretion, 285–290 characteristics, 285–287 constitutive secretion, 290 docking, 287–289 fusion, 287–289 membrane retrieval, 289–290 transport, 287 tubulation, 304–309 Paraxial mesoderm, in somitogenesis commitment, 6–8 differentiation, 1–3 precursor cells, 6–8 segmentation prepattern formation, 8–9 Paraxis transcription factor, somite formation role, 13 Patterning, somitogenesis, see Somites Peroxisomes, degradation, autophagic delivery to vacuoles, 175–178 in Hansenula polymorpha, 177 in Pichia pastoris, 175–177 in Saccharomyces cerevisiae, 177–178 Phagoacidosomes, membrane trafficking in Paramecium phagoacidosome into phagolysosome conversion, 301–303 phagosome into phagoacidosome conversion, 298–301 Phagosomes, trafficking in Paramecium, 294–304 acidosomes role, 297–298 cytoproct role, 303–304

355

SUBJECT INDEX digestive vacuole passage, 298 discoidal vesicles, 294–296 membrane replacement, 298–301 phagoacidosome into phagolysosome conversion, 301–303 phagosomal membrane precursors, 294–296 phagosome formation, 297 phagosome into phagoacidosome conversion, 298–301 spent vacuole formation, 303–304 vacuole membrane retrieval, 303–304 Pichia pastoris, peroxisome degradation pathways, 175–177 Plants, survival, benzoxazinones role, 319–341 acetal glucoside degradation to benzoxazolin-2-(3H)-ones, 322–324 benzoxazinoid aglucone action mechanisms, 332–335 biosynthesis, 324–332 Bx2-Bx5 isolation function, 327–329 Bx1 isolation and function, 324–327 Bx gene cluster organization, 329 DIBOA pathway, 329–332 distribution, 320–322 molecular allelopathy, 335–340 overview, 319–320, 341 structure, 320–322 Plasma membrane, ubiquitin-mediated protein degradation, 174 Presomitic mesoderm, somitogenesis role, 3, 7–13 Protein degradation, 165–178 autophagosomes role, 178 KFERQ-mediated pathway, 172–173 macroautophagy, 165–172 molecular component delivery, 166–172 physiological role, 172 mitochondria delivery to vacuoles, 178 peroxisome delivery to vacuoles, 175–178 in Hansenula polymorpha, 177 in Pichia pastoris, 175–177 in Saccharomyces cerevisiae, 177–178 starvation-response pathway, 172–173 ubiquitin-mediated endocytosis, 173–175 coordination, 174–175 plasma membrane protein degradation, 174 yeast vacuole targeting pathway, 173

Protein targeting, see Alternative proteinsorting pathways

Q Quick-frozen deep-etched imaging, membrane trafficking in Paramecium, 305

R Resegmentation theory, in somitogenesis, antero-posterior compartmentalization, 15–16 Retrograde transport membrane trafficking in Paramecium, 307–308 protein-sorting pathways, 154–165 coat proteins role, 155–156 endosome-to-Golgi transport, 162–163 function, 156 Golgi-to-endoplasmic reticulum transport, 156–159 anterograde transit components recycling, 156–158 coatomer function, 158–159 endoplasmic reticulum resident protein retrieval, 156–158 intra-Golgi transport, 159–162 cisternal maturation model, 161–162 vesicle transport model, 160–161 vacuole-to-endosome transport, 163–165 vesicle formation, 155–156

S Saccharomyces cerevisiae, peroxisome degradation pathways, 177–178 Sclerotome compartmentalization, in somitogenesis antero-posterior compartmentalization, 14–16 mechanisms, 14–15 resegmentation theory, 15–16 sclerotomal fate, 15–16 medio-lateral polarity, 47

356 Shh protein, in somitogenesis dorso-ventral patterning, 29–36 expression control, 35–36 myogenesis promotion, 33–34 ventral cell fate promotion, 30–33 Wnt protein combinatorial signaling, 33–34 medio-lateral polarity, 45–46 Somites, somitogenesis, 1–49 antero-posterior compartmentalization, 14–18 intrinsic determination, 16–17 polarity relationship, 17–18 resegmentation theory, 15–16 sclerotome compartmentalization, 14–16 commitment, 5–8 paraxial mesoderm-forming precursor cell specification, 6–8 somite origin, 5–6 dorso-ventral patterning, 18–37 cell-autonomous mechanisms, 23 commitment evolution, 21–22 control mechanisms, 23–29 dermis formation, 18–19 dorsalizing controls, 25–27 ectoderm role, 25–29 environmental signals, 22–23 external cues, 20 floor plate role, 24–25, 27–29 lineage segregation, 18–20, 29–37 muscle formation, 18–19 myogenic differentiation, 27–29, 33–37 neural tube role, 25–29 notochord role, 24–25, 27–29 self differentiation, 20–23 Shh protein role, 29–36 skeleton formation, 18–19 spatial regulation, 34–37 temporal regulation, 34–37 ventralizing controls, 24–25 Wnt protein role, 29–30, 33–35 formation, 8–13 bHLH transcription factors role, 13 cell-cell interactions, 9–10 cell-matrix interactions, 9–10 molecular clock linkage, 10–11

SUBJECT INDEX paraxial mesoderm segmentation prepattern, 8–9 tissue interactions, 9 medio-lateral polarity, 37–47 BMP4 role, 42–47 dorso-ventral-medio-lateral linkage, 41 ectoderm role, 41 external controls, 39–40 gastrulation, 39 lateral plate role, 40–43 lateral sclerotome, 47 lineages, 37–40 muscular differentiation, 38–39 Noggin role, 43–46 rib origin, 47 Shh pathway, 45–46 trunk versus limb hypaxial lineages, 41–42 Wnt pathway, 45–46 overview, 1–5, 47–49 Sorting pathways, see Alternative proteinsorting pathways Starvation-response pathway, protein degradation, 172–173

T Targeting pathways, see Alternative protein-sorting pathways Tight junctions, transepithelial water transport role, 204–205, 215–224 cell polarity maintenance, 220 solute diffusion barrier, 220–224 permeability regulation in leaky epithelia, 221–223 water absorption, 223–224 structure, 217–220 Trafficking dec-1 protein in Drosophila eggshell morphogenesis, 95–99 membranes in Paramecium, 277–312 anterograde movements, 306–307 biosynthetic pathway, 279–285 docking mechanisms, 285 endoplasmic reticulum transition zones, 279–283 filaments, 281–283 fusion mechanisms, 285 Golgi stack membranes, 281

SUBJECT INDEX recognition mechanisms, 285 trans-Golgi network, 283–284 budding, 304–309 coated membranes role, 306–307 cognate membrane recognition, 308–309 distribution and routing, 309–311 long distance movements, 309–310 microfilaments role, 309–310 microtubules role, 309–310 short distance movement regulation, 310–311 docking mechanisms, 285, 308–309 endosomal trafficking, 290–294 carrier vesicles, 293–294 coated pit, 290–291 early endosomes, 291–294 endocytosis, 290–291 exocytosis sites, 294 noncoated endocytosis, 294 parasomal sac, 290–291, 294 preendosomal vesicles, 291–293 fusion mechanisms, 285, 308–309 fusogenic property changes, 307–308 membrane recycling, 307–308 modulation, 304–309 overview, 277–279, 311–312 phagosomal system, 294–304 acidosomes, 297–298 cytoproct role, 303–304 digestive vacuole passage, 298 discoidal vesicles, 294–296 membrane replacement, 298–301 phagoacidosome into phagolysosome conversion, 301–303 phagosomal membrane precursors, 294–296 phagosome formation, 297 phagosome into phagoacidosome conversion, 298–301 spent vacuole formation, 303–304 vacuole membrane retrieval, 303–304 priming mechanisms, 308–309 quick-frozen deep-etched imaging, 305 retrograde movements, 307–308 sorting events, 304–309 transport systems, 304–309

357 trichocyst secretion, 285–290 characteristics, 285–287 constitutive secretion, 290 docking, 287–289 fusion, 287–289 membrane retrieval, 289–290 transport, 287 tubulation, 304–309 Transcription, see also specific transcription factors cytochrome P-450 expression regulation in liver cells posttranscriptional regulation, 123–126 transcriptional regulation, 110–126 somite formation, 13 Transepithelial water transport, 203–256, see also Diffusion apical membrane ultastructure, 224–240 leaky epithelia, 224–227, 236–237 structural change dynamics during transport, 226–232 aquaporins role, 232–240 intramembrane particle aggregate sources, 230–231 tight epithelia, 225–232, 237–240 cytoskeleton role, 250–255 cortical actin participation, 250–251 microtubules participation, 251–255 antidiuretic hormone role, 252–253 calcium ion accumulation, 254 giant vacuole migration, 254–255 granular cells in frog urinary bladder, 252–253 high water permeability, 252 low water permeability, 252 thick microtubules, 253–255 tubulin polymerization, 254 water channel protein docking and insertion, 250–251 epithelia features, 205–215 functional asymmetry, 206–209 antidiuretic hormone effects, 208–209 leaky epithelia, 206–207 tight epithelia, 207–208 ultrastructure, 209–215 leaky and tight epithelia compared, 209–211

358 Transepithelial water transport (continued ) leaky epithelia in rat intestine, 211–213 tight epithelia in frog bladder, 213–215 overview, 203–205, 255–256 tight junction role, 215–224 cell polarity maintenance, 220 solute diffusion barrier, 220–224 permeability regulation in leaky epithelia, 221–223 water absorption, 223–224 structure, 217–220 vacuolar system role, 240–250 epithelial cell structure changes, 240–241 Golgi complex function, 241–250 enterocytes in rat, 242–243 epithelial cells, 241–242 giant vacuole evolution, 249–250 granular cell giant vacuole and protozoan contractile vacuole homology, 249 urinary bladder in frog, 242–248 Trans-Golgi network, membrane trafficking role in Paramecium, 283–284 Translation, see also Alternative proteinsorting pathways cytochrome P-450 expression regulation in liver cells posttranslational regulation, 123–126 translational regulation, 123–126 eggshell morphogenesis in Drosophila, 79–85 chorion genes, 81–82 dec-1 gene, 82–85 vitelline membrane genes, 79–80 Transport, see Transepithelial water transport Trichocysts, secretion mechanisms in Paramecium, 285–290, 312 characteristics, 285–287 constitutive secretion, 290 docking, 287–289 fusion, 287–289 membrane retrieval, 289–290 transport, 287 Tubulin, polymerization in transepithelial water transport, 254

SUBJECT INDEX

U Ubiquitin, endocytosis mediation, 173–175 coordination, 174–175 plasma membrane protein degradation, 174

V Vacuoles membrane trafficking in Paramecium, phagosomal system digestive vacuole passage, 298 spent vacuole formation, 303–304 vacuole membrane retrieval, 303–304 protein degradation mitochondria delivery, 178 peroxisome delivery, 175–178 in Hansenula polymorpha, 177 in Pichia pastoris, 175–177 in Saccharomyces cerevisiae, 177–178 yeast vacuole targeting pathway, 173 protein-sorting pathways, vacuole-toendosome transport, 163–165 transepithelial water transport, 240–250 epithelial cell structure changes, 240–241 evolution, 249–250 Golgi complex function, 241–250 enterocytes in rat, 242–243 epithelial cells, 241–242 giant vacuole evolution, 249–250 granular cell giant vacuole and protozoan contractile vacuole homology, 249 urinary bladder in frog, 242–248 migration, microtubules participation, 254–255 protozoan contractile vacuole homology, 249 Vasopressin, transepithelial water transport role, 204, 207–209, 252–253 Vitelline membrane, Drosophila eggshell morphogenesis role genes cloning, 77–78 conceptual translation products, 79–80 proteins biochemical analysis, 73–74, 103 posttranslational processing, 87–88

359

SUBJECT INDEX VPS proteins, vacuolar protein recycling role, 162–163

W Water transport, see Transepithelial water transport Wnt protein, in somitogenesis dorso-ventral patterning, 29–35 dorsal lineage specification, 30–33

inhibitors, 35 myogenesis promotion, 33–34 Shh protein combinatorial signaling, 33–34 medio-lateral polarity, 45–46

Z Zonulae occludens, see Tight junctions

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