The Golgi Apparatus: State of the art 110 years after Camillo Golgi's discovery

SpringerWienNewYork

Alexander A. Mironov and Margit Pavelka (eds.)

The Golgi Apparatus State of the art 110 years after Camillo Golgi’s discovery

SpringerWienNewYork

Prof. Dr. Alexander A. Mironov Consorzio Mario Negri Sud Laboratory of Intracellular Traffic Department of Cell Biology and Oncology, S. Maria Imbaro (Chieti), Italy

Prof. Dr. Margit Pavelka Department of Cell Biology and Ultrastructure Research Institute of Histology and Embryology Center for Anatomy and Cell Biology Medical University of Vienna Vienna, Austria € tzung des Bundesministeriums fu € r Wissenschaft und Forschung Gedruckt mit Unterstu in Wien

This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically those of translation, reprinting, re-use of illustrations, broadcasting, reproduction by photocopying machines or similar means, and storage in data banks. Product Liability: The publisher can give no guarantee for all the information contained in this book. This does also refer to information about drug dosage and application thereof. In every individual case the respective user must check its accuracy by consulting other pharmaceutical literature. The use of registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use.  2008 Springer-Verlag/Wien Printed in Austria SpringerWienNewYork is a part of Springer Science þ Business Media springer.at Typesetting: Thomson Press (India) Ltd., Chennai Printing: Holzhausen Druck & Medien, 1140 Wien Printed on acid-free and chlorine-free bleached paper SPIN: 12054541 With 106 partly coloured figures Library of Congress Control Number: 2008932528 ISBN 978-3-211-76309-4 SpringerWienNewYork

Preface

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Preface In 2008, we celebrate the 110th anniversary of the first description of the complex apparato reticolare interno by Camillo Golgi in the Bollettino della  Medico-Chirurgica di Pavia. The biography of Camillo Golgi, and the Societa Golgi apparatus history have impressively been described by Paolo Mazzarello in his book “The Hidden Structure. A Scientific Biography of Camillo Golgi” (Oxford University Press, 1999). During the 20th century, Camillo Golgi’s discovery had a changing up and down and up history, timely being assessed as an artefact, and then again coming into the centre stage of cell biologic research. Today, it is well established that the Golgi apparatus constitutes a main crossroads in the intracellular transport routes of the biosynthetic, endocytic, and recycling systems. During the past decades, multiple new discoveries contributed to the understanding of the organization and the functions of the complex organelle (see Chapter 1.1). In 1997, the excellent book about the Golgi apparatus edited by J. Roth and E. Berger provided a comprehensive summarizing presentation of the state of research 100 years after the first description of the organelle. Now, after further 10 years, it is necessary to summarize again what it is known about the complex organization of Camillo Golgi’s apparatus. Our book is an attempt to bring together multiple new results obtained by different techniques, and addressing different aspects of the Golgi apparatus and intracellular transport. We are hopeful that the presentation of the state of the art 110 years after Camillo Golgi’s discovery of the complex apparato reticolare interno will lead to an improved understanding, novel insights, and new perspectives for future research.

Acknowledgements The editors cordially thank all authors of the Chapters, all our colleagues involved in the works presented in the Chapters of this book written by us and by others. We thank the Springer Company for the possibility to publish this book and in particular we thank Mag. Franziska Brugger, Mag. Angelika Heller, and Mrs. Ursula Szorger for a huge help in our work. A.M. is especially thankful to Dr. A. Fusella and D. Gaindomenico for the technical help and to Chris Berrie and Raman Parashuraman for critical reading of the manuscripts, in which A.M. is a co-author. A. A. Mironov and M. Pavelka S. Maria Imbaro/Vienna, May 2008

Movies can be viewed online at: www.springer.com/springerwiennewyork/lifeþsciences/book/978-3-211-76309-4

Contents

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Contents Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

1. General considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 1.1. The Golgi apparatus and main discoveries in the field of intracellular transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 1.2. The Golgi apparatus as a crossroads in intracellular traffic . . . 16 2. Main machineries operating at the Golgi apparatus . . . . . . . . . . . . . 2.1. SNAREs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Rabs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. COPII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. COPI: mechanisms and transport roles . . . . . . . . . . . . . . . . . . . 2.5. Arfs and Arls: models for Arf family members in membrane traffic at the Golgi . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. COG complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7. The TRAPP complex. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8. The role of Ca2þ in the regulation of intracellular transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9. Golgi glycosylation enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10. Nucleotide sugar transporters of the Golgi apparatus. . . . . . . 2.11. Luminal lectins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12. The Golgi ribbon and the function of the golgins . . . . . . . . . . 2.13. Functional cross talk between membrane trafficking and cell signalling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.14. The role of the cytoskeleton in the structure and function of the Golgi apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15. The dynamin–cortactin complex as a mediator of vesicle formation at the trans-Golgi network . . . . . . . . . . . . . . . . . . . 2.16. The geometry of organelles of the secretory pathway . . . . . . 3. Main transport steps . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. ER-to-Golgi transport . . . . . . . . . . . . . . . . . . . . . 3.2. Intra-Golgi transport . . . . . . . . . . . . . . . . . . . . . . 3.3. Structure and domain organization of the trans-Golgi network . . . . . . . . . . . . . . . . . . . . . . 3.4. Golgi-to-PM transport . . . . . . . . . . . . . . . . . . . . . 3.5. Protein transport from the trans-Golgi network to endosomes . . . . . . . . . . . . . . . . . . . . . . . . . . .

41 43 66 78 87 106 120 130 143 161 190 207 223 247 270 301 314

. . . . . . . . . . . 331 . . . . . . . . . . . 333 . . . . . . . . . . . 342 . . . . . . . . . . . 358 . . . . . . . . . . . 375 . . . . . . . . . . . 388

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3.6. 3.7. 3.8. 3.9. 3.10. 3.11. 3.12. 3.13. 3.14. 3.15.

The transport of soluble lysosomal hydrolases from the Golgi complex to lysosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . Transport of lysosomal membrane proteins from the Golgi complex to lysosomes . . . . . . . . . . . . . . . . . . . . . . . . Retrograde endosome-to-TGN transport . . . . . . . . . . . . . . . . . Retrograde plasma membrane-to-Golgi apparatus transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Interactions between endocytosis and secretory transport . . Origins of the regulated secretory pathway . . . . . . . . . . . . . . Secretion and endocytosis in endothelial cells . . . . . . . . . . . . . Formation of mucin granules . . . . . . . . . . . . . . . . . . . . . . . . . . Golgi apparatus and epithelial cell polarity . . . . . . . . . . . . . . . Golgi apparatus inheritance . . . . . . . . . . . . . . . . . . . . . . . . . . .

4. Peculiarities of intracellular transport in different organisms . . 4.1. Features of the plant Golgi apparatus . . . . . . . . . . . . . . . 4.2. Yeast Golgi apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Morphodynamics of the yeast Golgi apparatus . . . . . . . . 4.4. Structure and function of the Golgi organelle in parasitic protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. Evolution of the Golgi complex . . . . . . . . . . . . . . . . . . . .

.... .... .... ....

402 414 425 459 475 485 520 535 563 580 609 611 623 630

. . . . 647 . . . . 675

5. General conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 693 Contributors in alphabetical order . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 697 Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 706

Introduction

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Introduction Alexander A. Mironov and Margit Pavelka

All cells secrete a diversity of macromolecules to modify their environment or to protect themselves. On the other hand, there is the necessity to replace membrane proteins and lipids that are being constantly degraded in compartments of the secretory and endocytic pathways. Therefore, eukaryotic cells synthesize proteins for either export (secretion) or delivery to the secretory and endocytic compartments and to the plasma membrane (PM) for replacement of degraded proteins and lipids. The synthesis is carried out by ribosomes attached to the cytosolic surface of the endoplasmic reticulum (ER). The synthesis of most of cellular lipids and all fatty acids also occurs in the smooth ER. During or after the synthesis, the polypeptide chains containing transmembrane domains composed of hydrophobic amino acids are inserted into the ER membrane, whereas soluble proteins are transferred into the ER lumen. After the cleavage of their leading hydrophobic signal peptides and after protein folding both the groups of proteins are transported along the secretory pathway. The Golgi apparatus (GA) is the central station along this pathway. While passing through the GA, proteins and lipids undergo posttranslational modifications (mainly glycosylation by Golgi glycosidases and transferases) and sorting. The transport of proteins and lipids from the ER to their destinations may be carried out in several ways: via the dissociation mechanism, the progression mechanism, and/or the lateral diffusion mechanism (see Chapter 1.2). Over the past 30 years, the field of intracellular traffic has seen tremendous advances towards the identification of the relevant molecular machineries. Substantial progress in the isolation, cloning, and characterization of proteins involved in intracellular transport and its regulation, as well as in deciphering corresponding molecular events was achieved. Proteins involved in budding, fission, fusion, and sorting have been discovered, and, in some cases, a picture of how such proteins are assembled and work has been glimpsed. In contrast, perhaps surprisingly, a satisfactory understanding of how transport occurs in vivo at the organelle level has not been achieved, and the general picture of this process remains obscure. As a consequence, our present view of the overall mode of intracellular traffic (the “physiology” of traffic) in living cells is rather poorly developed. Although in vitro reconstitution experiments have been crucial in establishing the minimum number of components required for carrier budding using biochemical techniques, a more critical evaluation of intracellular transport could be complemented by in vivo experiments. The reasons for this lag are both technological and conceptual. Technological, because intracellular traffic is an essentially dynamic event in time and

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space. Yet, methods to study its dynamic features have been lacking until very recently. The recent advent of green fluorescent protein (GFP) technology has now partially filled this gap. Conceptual, because the field has been dominated until very recently by the model of intracellular traffic by anterograde vesicles. The vesicular scheme has been very successful in providing a framework to integrate an enormous amount of biochemical and genetic evidence collected over the past two decades into a coherent picture. In the past years, the progress in development of new scientific methods accelerated. New microscopic and biochemical methods, new scientific instruments together with the development of molecular tools has given the possibility to study biochemical reactions and molecules in single cells with a resolution impossible to achieve before. Significant progress was made in the development of new synchronization protocols suitable for study of transport of several cargoes. On the other hand, the complete deciphering of the full genetic code of yeast, humans, plants and several other species together with the development of proteomics provided a significant number of new proteins and details on protein–protein and protein–lipid interactions involved into the traffic. In the literature, there are thousands of details regarding protein interactions, their sorting signals, and the effects of their inhibition and deletion. As such, it seems that the field of intracellular transport appears to be facing a serious crisis. There are now so many proteins and inter-protein interactions involved that it has become almost impossible to follow and understand the meaning of all of these details. The main idea of this book is thus to collect the full range of expertise and to examine the problems from different points of view and with different approaches. This book is devoted to the molecular mechanisms of morpho-functional organization of the GA and summarizes most of new data related to the GA. The book is a collection of chapters written by different groups and therefore expresses different views especially on the mechanisms of traffic. The possibility to follow the evaluation of the intracellular transport from different points of view and on the basis of different expertises could help to resolve this contradiction in the field. There are several levels in the book. The main is the description of cell physiology with emphasis on the physiology of intracellular transport. The second level is the presentation of the morphology in wide term including light microscopy, analysis of live cells and so on. Finally, there are several chapters devoted to the molecular mechanisms involved in different physiological processes related to intracellular transport. In this book, we could not exclude completely some degree of overlapping. First, because the authors have different opinions about models of transport, and the second, it was necessary to illustrate some ideas from different points of view. We apologize to colleagues, whose relevant work has not been mentioned because of space limitations and focusing on work published most recently.

Introduction

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In the first part of the book there are two chapters. One is an overview of the history of this field of cell biology. The second one is a morpho-functional overview of the main steps of intracellular transport including functional organization and architecture of the GA. In the second part of the book, main machineries operating along the secretory pathway are presented. These include SNAREs, Rabs, COPII, COPI, ARFs, ARFGEFs and ARLs, COG, TRAPP, dynamin and cortactin, Golgi enzymes and sugar transporters, ERGIC-53, and other luminal lectins, Golgins and Golgi matrix proteins. Lipids and lipid signalling, as well as common signalling mechanisms will be assessed. Additionally, the role of calcium and other ions in the regulation of intracellular transport, and the role of the cytoskeleton in Golgi function will be described. In the third part of the book, different transport steps of intracellular transport will be evaluated, namely, ER-to-Golgi transport, intra-Golgi transport, Golgi-to-PM transport, Golgi-to-endosome transport, transport of lysosomal enzymes, retrograde endosome-to-TGN transport, and retrograde PM-to-Golgi transport. Mechanisms of regulated secretion will be explained in a separate chapter and in particular, mechanisms involved in formation of mucin granules are described. Interactions between endocytosis and secretory transport, the relationship between the GA and cell polarity, as well as structure and domain organization of the trans-Golgi network, and the questions of GA formation and inheritance will be discussed in separate chapters. Finally, in the last part of the book the reader will find some aspects about the peculiarities of intracellular transport in different organisms: plants, yeast, and protists. A separate chapter is devoted to the endomembrane ultrastructure and dynamics in yeasts. Finally, the models of Golgi evolution will be discussed in the final chapter of this part. This book is intended for cell biologists and histologists, who work with students, and also for scientists working in other fields of biology as well as for students per se. The most important item for teaching is the understanding of not a single or several mechanisms but the comprehensive view upon the full drama of development of scientific models. The readers should understand the logic of model replacements. Therefore, the aim of the book is to make the field of intracellular transport more understandable by keeping it as simple as possible, but also as full as possible, and at minimal cost. Finally, we would like to mention the question of terminology. Since the original term was “Golgi apparatus”, this term is used in most of the chapters. However, both terms, Golgi apparatus and Golgi complex, are customary today.

General considerations

The Golgi apparatus and main discoveries

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The Golgi apparatus and main discoveries in the field of intracellular transport Alexander A. Mironov and Margit Pavelka

In this chapter, we summarize important findings in the field of intracellular transport, which have considerably contributed to the understanding of the function and organization of the Golgi apparatus (GA). It is not possible to mention all authors in this huge field. We apologize for gaps and incompleteness, and are thankful for suggestions and corrections. The GA is named after its discoverer Camillo Golgi, who first described the complex apparato reticolare interno in 1898 (Golgi 1898a,b; reviewed € scher 1998). Although Camillo Golgi had presented his by Berger 1997; Dro discovery convincingly, for a long time his data have been considered as an artifact of cell staining (Farquhar and Palade 1981). Only after the electron microscopic confirmation of the existence of the GA in cells by Dalton in 1951, scientists started to believe in its reality. Therefore, we will not list the discoveries within the area of intracellular transport made in the time, before the existence of the GA was confirmed electron microscopically. However, the names of A. Negri, H. Fuch, A. Perroncito, S. Ramon y Cajal, D.N. Nassonov, R.H. Bowen, G.S. Carter, H.W. Beams and R.L. King, V.M. Emmel, H.W. Deane and E.M. Dempsey, W.C. Schneider et al. should be mentioned, because they have considerably contributed to the understanding of the Golgi function (reviewed in Berger 1997). Here, we want to address most important discoveries within the area of intracellular transport after 1951 (Table 1). Additionally, we would like to mention further important contributions to this field. The hypothesis of lipid rafts was proposed and developed by van Meer and Simons. The Lodish group made the invention of the synchronization of the transport of cargoes. The role of lectins in ER-to-Golgi transport was discovered by H.-P. Hauri. The most important contribution to the characterization of Rab machinery (although in the endocytic pathway) was made by M. Zerial. W. Hong, R. Sheller and R. Jahn made important contributions to the understanding of the function of the SNARE machinery. R. Schekman and W. Balch deciphered the functions of the COPII coat. A. Rambourg, Y. Clermont, G. Griffiths, A. Staehelin and K. Howell made significant contributions to the 3D-analysis of the GA in different cell types. J. Slot and H. Geuze provided new insight into the morphology of the endocytic system and its interaction with exocytosis. The important contribution into the analysis of the kinetics of the plant GA was made by C. Hawes. The characterization of the 3D-structure of several proteins important for intracellular transport, and protein coat complexes in their crystal state is linked with W. Balch and J. Goldberg’s names. We apologise again for possible

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Table 1. The Golgi apparatus and main discoveries in the field of physiology of intracellular transport 1898 1951 1961 1964 1964 1964 1964 1966

1966 1967 1969 1976 1977 1980 1981 1982

Discovery of the GA Confirmation of the presence of the GA (Dalton 1951) The regional distribution of the thiamine-pyrophosphatase activity within the GA (Novikoff and Goldfischer 1961) The trans ER (Novikoff 1964; Novikoff et al. 1964) GERL concept (Novikoff 1964)  and Mollenhauer 1964) Isolation of Golgi membranes from cells (Morre The process of sulphation in the GA (Godman and Lane 1964) The sugar–nucleotide transport from the cytosol to the Golgi lumen across the Golgi membranes, the role of the GA in glycosylation (Neutra and Leblond 1966) The origin of lysosomes and the function of clathrin-coated vesicles during protein absorption (Bainton and Farquhar 1966; Friend and Farquhar 1967) The intracellular transport (Jamieson and Palade 1967a,b)  et al. 1969) Galactosyltransferase as a Golgi marker (Whur et al. 1969; Morre Isolation of clathrin-coated vesicles (Pearse 1976) The PM-to-Golgi transport of the endogenously added marker (Herzog and Farquhar 1977) M6P-mediated sorting of Golgi enzymes at the GA (Tabas and Kornfeld 1980) Clathrin-coated buds in the trans side of the GA (Griffiths et al. 1981) Immunocytochemical localization of galactosyltransferase (Roth and Berger 1982) Topology of N-glycosylation (Dunphy and Rothman 1983) Reconstitution of intra-Golgi transport in vitro (Balch et al. 1984) The 15 C temperature block (Saraste and Kuismanen 1984) Clathrin-independent endocytosis (Moya et al. 1985; Sandvig et al. 1985)

1983 1984 1984 1985 1985– 1987 The mitotic form of the GA and mechanisms of mitotic Golgi transformation in animal cells (Featherstone et al. 1985; Lucocq et al. 1987) 1986 The COPI-coated vesicles and characterization of molecular mechanisms involved into the function of COPI coat (Orci et al. 1986; Serafini et al. 1991) 1986 The structure and function of the TGN and the 20 C temperature block (Griffiths and Simons 1986) 1987 KDEL-signal for the retention of luminally located proteins (Munro and Pelham 1987) 1989 BFA was applied for the study of intra-Golgi transport (Doms et al. 1989; Lippincott-Schwartz et al. 1989) 1990 SNAREs (Newman et al. 1990) 1990 The main genes involved in intracellular transport, the genetic evidence in favour of the vesicular model of the transport in yeast (Kaiser and Schekman 1990) 1991 A Golgi retention signal in the membrane-spanning domain (Swift and Machamer 1991) 1993 The role of oligomerization for the retention of Golgi enzymes (Weisz et al. 1993) 1993 The role of PM-derived signalling for intra-Golgi transport (De Matteis et al. 1993) 1994 Golgi matrix (Slusarewicz et al. 1994) and cis-Golgin, GM130 (Nakamura et al. 1995) 1994 COPI-dependent retrieval sorting signals (Cosson and Letourneur 1994)

The Golgi apparatus and main discoveries

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Table 1. (Continued)

1994 1996 1996 1997 1997 1997 1998 1998 1998 1999 2001 2003 2003 2004 2006 2006 2007

COPII coat. Isolation of COPII-dependent small vesicles in cell-free system (Barlowe et al. 1994) Application of GFP-technology for the study of the GA in living cells (Cole et al. 1996) Characterization of the ER exit sites (Bannykh et al. 1996) The AP3 and AP4 coats (Dell’Angelica et al. 1997, 1999) Characterization of ER-to-Golgi transport carriers in living cells (Presley et al. 1997; Scales et al. 1997; Mironov et al. 2003) Characterization of post-Golgi transport carriers in living cells (Wacker et al. 1997; Hirschberg et al. 1998; Polishchuk et al. 2000) Intra-Golgi transport of large cargo aggregates (Bonfanti et al. 1998) The role of endocytic TGN in the formation of the most-trans Golgi cisterna (Pavelka et al. 1998) Discovery of R- and Q-SNAREs (Fasshauer et al. 1998) Tomographic reconstruction of the GA (Ladinsky et al. 1999) The concentration of regulatory secretory proteins within the Golgi cisternae (Oprins et al. 2001) The understanding of the evolution of small GTPases had changed the model kely 2003) of the Golgi evolution (Je Characterization of Golgi-to-apical PM transport carriers in living cells (Kreitzer et al. 2003) Intercisternal connections in transporting GA (Marsh et al. 2004; Trucco et al. 2004) Characterization of the Golgi-to-endosome carriers in living cells (Polishchuk et al. 2006) The role of GM130 in the maintenance of the Golgi ribbon (Puthenveedu et al. 2006) The role of ER-to-Golgi transport in the maintenance of the Golgi ribbon (Marra et al. 2007)

gaps (all authors quoted in the consecutive chapters deserve to be listed here). The list is open for suggestions. The development of the research in the field of intracellular transport has been comprehensively discussed in 1998 at the conference in Pavia devoted to the 100th anniversary of the Golgi discovery.

History of models of intracellular transport Historically, the first mechanism that had been proposed for intracellular transport was the progression. The origin of the progression model (or the concept of cis-to-trans flow) links to Grasse’s name (1957) who proposed that the continuous formation of cis Golgi cisternae balances the conversion of trans one into secretory granules. However, the first experimental data in favour of the progression concept were obtained in 1971 (Franke et al. 1971). In 1967, it has been demonstrated that proteins newly synthesized in the ER appeared, after a few minutes, not only over Golgi stacks but also over

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round profiles surrounding the GA and the conclusion that secretory proteins bypass the GA was made (Jamieson and Palade 1967a,b, 1968a,b). Then, in 1981, the vesicular model replaced the progression model because the main support for the progression model, the cis–trans movement of scales in algae has been considered to be a rare formula connected with unusual geometry and size of the product (Farquhar and Palade 1981). Ironically, the major supporting data for the vesicular model at that time was based on the isolation of Golgi-derived clathrin-coated vesicles (Rothman et al. 1980). However, after the discovery of coat protein I (COPI) (Orci et al. 1986), the vesicular model was changed, and instead of clathrin-dependent vesicles, COPI-dependent vesicles were proposed to serve as anterograde carriers. The strongest support for the vesicular model appeared from the experiments in yeast with the temperature sensitive Sec genes (Kaiser and Schekman 1990). The in vitro isolation of functional (containing VSVG and able to fuse with acceptor Golgi membranes) COPI-coated vesicles (Osterman et al. 1993) was interpreted as the second proof for the role of COPI-coated vesicles in the anterograde intra-Golgi transport. Importantly, however, that the first author of this paper later stressed, that actually, these data support the cisterna maturation model (Ostermann 2001). On the other hand, it has also been demonstrated that 20 min after fusion of two (or more) cells (one cell is VSV-infected, another is a non-infected cell) and formation of a heterokaryon, VSVG seems to move from the GA derived from the infected cell to the GA derived from non-infected cells (Rothman et al. 1984). These results were interpreted as confirmation of the ability of vesicular carriers to diffuse through the cytosol of the heterokaryon from one GA to another. However, later, the Rothman group (Orci et al. 1998) laid less emphasis on the heterokaryon experiments, suggesting that those observations appeared as a result of the treatment of cells with an acidic medium. Instead, the “string theory” was proposed, according to which a proteinaceous-like string links vesicles to cisternal elements and prevents budded vesicles from diffusing away, while still allowing them to diffuse laterally. With time, due to accumulation of contradictions, the current vesicular paradigm became less and less effective in the explanation of growing body of observations (Mironov et al. 1997). As a result, the original version of the vesicular paradigm began to be modified not only by the opponents of the vesicular model but also by its authors and proponents (Orci et al. 1998). In order to resolve accumulated contradictions within the field, almost simultaneously several groups (Bannykh and Balch 1997; Mironov et al. 1997; Glick et al. 1997; Schekman and Mellman 1997) have published the cisterna maturation-progression model based on the COPI vesicles-mediated Golgi enzyme recycling. The first experimental confirmation that large aggregated cargo, such as procollagen I, can be transported through the GA by maturation mechanism came in 1998 (Bonfanti et al. 1998). Previous stereological observations in

The Golgi apparatus and main discoveries

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P. scheffelii suggesting that their scales being much too large to be packaged into vesicles are transported by the progression of Golgi cisternae towards the plasmalemma were published not in an original paper but in a review (Becker et al. 1995) and were not confirmed later because glycoprotein and polysaccharide synthesis are uncoupled during flagella regeneration (Perasso et al. 2000). Next, it has been demonstrated (Mironov et al. 2001) that both diffusible and non-diffusible cargoes are transported in the same carriers through the Golgi stacks. It has been proved that vesicles are not transport carriers for cargo in the intra-Golgi transport not only in situ, but also in vitro, in cell-free assay (Happe and Weidman 1998). After these publications, there was a short period when the cisterna maturation model became dominant. With time new contradictions not compatible with the cisterna maturation-progression model have accumulated (Mironov et al. 2005). The attempts to use transport models based on combination of basic principles were not successful (see Chapter 3.2). Therefore now, there is no consensus on the models of intra-Golgi transport. The existence of the maturation mechanism is almost finally established for the secretion of large polymeric structures incompatible in size with COPI-dependent vesicles in many types of cells and under the infection of some viruses.

References Bainton DF, Farquhar MG (1966) Origin of granules in polymorphonuclear leukocytes. Two types derived from opposite faces of the Golgi complex in developing granulocytes. J Cell Biol 28(2): 277–301 Balch WE, Dunphy WG, Braell WA, Rothman JE (1984) Reconstitution of the transport of protein between successive compartments of the Golgi measured by the coupled incorporation of N-acetylglucosamine. Cell 39: 405–416 Bannykh SI, Rowe T, Balch WE (1996) The organization of endoplasmic reticulum export complexes. J Cell Biol 135: 19–35 Bannykh SI, Balch WE (1997) Membrane dynamics at the endoplasmic reticulum–Golgi interface. J Cell Biol 138: 1–4 Barlowe C, Orci L, Yeung T, Hosobuchi M, Hamamoto S, Salama N, Rexach MF, Ravazzola M, Amherdt M, Schekman R (1994) COPII: a membrane coat formed by Sec proteins that drive vesicle budding from the endoplasmic reticulum. Cell 77: 895–907 Becker B, Bolinger B, Melkonian M (1995) Anterograde transport of algal scales through the Golgi complex is not mediated by vesicles. Trends Cell Biol 5: 305–307 Berger EG (1997) The Golgi apparatus: from discovery to contemporary studies. In: Berger EG, Roth J (eds) The Golgi apparatus. Basel et al., Birkhauser Verlag, pp 1–35  rguez JA, Martella O, Fusella A, Baldassarre M, Bonfanti L, Mironov AA Jr, Martínez-Mena Buccione R, Geuze HJ, Mironov AA, Luini A (1998) Procollagen traverses the Golgi stack without leaving the lumen of cisternae: evidence for cisternal maturation. Cell 95(7): 993–1003 Cole NB, Smith CL, Sciaky N, Terasaki M, Edidin M, Lippincott-Schwartz J (1996) Diffusional mobility of Golgi proteins in membranes of living cells. Science 273 (5276): 797–801

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Cosson P, Letourneur F (1994) Coatomer interaction with di-lysine endoplasmic reticulum retention motif. Science 263: 1629–1631 Dalton AJ (1951) Observations of the Golgi substance with the electron microscope. Nature 168(4267): 244–245 De Matteis MA, Santini G, Kahn RA, Di Tullio G, Luini A (1993) Receptor and protein kinase C-mediated regulation of ARF binding to the Golgi complex. Nature 364: 818–821 Dell’Angelica EC, Ohno H, Ooi CE, Rabinovich E, Roche KW, Bonifacino JS (1997) AP-3: an adaptor-like protein complex with ubiquitous expression. EMBO J 16(5): 917–928 Dell’Angelica EC, Mullins C, Bonifacino JS (1999) AP-4, a novel protein complex related to clathrin adaptors. J Biol Chem 274: 7278–7285 Doms RW, Russ G, Yewdell JW (1989) Brefeldin A redistributes resident and itinerant Golgi proteins to the endoplasmic reticulum. J Cell Biol 109: 61–72 € scher A (1998) Camillo Golgi and the discovery of the Golgi apparatus. Histochem Dro Cell Biol 109: 425–430 Dunphy WG, Rothman JE (1983) Compartmentation of asparagine-linked oligosaccharide processing in the Golgi apparatus. J Cell Biol 97(1): 270–275 Farquhar MG, Palade GE (1981) The Golgi apparatus (complex)-(1954–1981)-from artifact to center stage. J Cell Biol 91(3 Pt 2): 77s–103s Fasshauer D, Sutton RB, Brunger AT, John R (1998) Conserved structural features of the synaptic fusion complex: SNARE proteins reclassified as Q- and R-SNAREs. Proc Natl Acad Sci USA 95(26): 15781–15786 Featherstone C, Griffiths G, Warren G (1985) Newly synthesized G protein of vesicular stomatitis virus is not transported to the Golgi complex in mitotic cells. J Cell Biol 101(6): 2036–2046 Franke WW, Morre DJ, Deumling B, Cheetham RD, Kartenbeck J, Jarasch E-D, Zengtraf HW (1971) Synthesis and turnover of membrane proteins in rat liver: an examination of the membrane flow hypothesis. Z Naturforsch 26b: 1031–1039 Friend DS, Farquhar MG (1967) Functions of coated vesicles during protein absorption in the rat vas deferens. J Cell Biol 35(2): 357–376 Glick BS, Elston T, Oster G (1997) A cisternal maturation mechanism can explain the asymmetry of the Golgi stack. FEBS Lett 414: 177–181 Godman GC, Lane N (1964) On the site of sulfation in the chondrocyte. J Cell Biol 21: 353–366 Golgi C (1898a) Intorno alla struttura della cellula nervosa. Boll Soc Med Chir Pavia 13: 1–14 Golgi C (1898b) Sur la structure des cellules nerveuses des ganglions spinaux. Arch Ital Biol 30: 60–71 Grasse PP (1957) Ultrastructure, polarity and reproduction of Golgi apparatus. C R Hebd Seances Acad Sci 245(16): 1278–1281 Griffiths G, Warren G, Stuhlfauth I, Jockusch BM (1981) The role of clathrin-coated vesicles in acrosome formation. Eur J Cell Biol 26(1): 52–60 Griffiths G, Simons K (1986) The trans Golgi network: sorting at the exit site of the Golgi complex. Science 34: 438–443 Happe S, Weidman P (1998) Cell-free transport to distinct Golgi cisternae is compartment specific and ARF independent. J Cell Biol 140(3): 511–523 Herzog V, Farquhar MG (1977) Luminal membrane retrieved after exocytosis reaches most Golgi cisternae in secretory cells. Proc Natl Acad Sci USA 74(11): 5073–5077 Hirschberg K, Miller CM, Ellenberg J, Presley JF, Siggia ED, Phair RB, LippincottSchwartz J (1998) Kinetic analysis of secretory protein traffic and characterization of Golgi to plasma membrane transport in living cells. J Cell Biol 143: 1485–1503

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Jamieson JD, Palade GE (1967a) Intracellular transport of secretory proteins in the pancreatic exocrine cell. I. Role of the peripheral elements of the Golgi complex. J Cell Biol 34(2): 577–596 Jamieson JD, Palade GE (1967b) Intracellular transport of secretory proteins in the pancreatic exocrine cell. II. Transport to condensing vacuoles and zymogen granules. J Cell Biol 34(2): 597–615 Jamieson JD, Palade GE (1968a) Intracellular transport of secretory proteins in the pancreatic exocrine cell. 3. Dissociation of intracellular transport from protein synthesis. J Cell Biol 39(3): 580–588 Jamieson JD, Palade GE (1968b) Intracellular transport of secretory proteins in the pancreatic exocrine cell. IV. Metabolic requirements. J Cell Biol 39(3): 589–603 kely G (2003) Small GTPases and the evolution of the eukaryotic cell. Bioessays 25(11): Je 1129–1138 Kaiser CA, Schekman R (1990) Distinct sets of SEC genes govern transport vesicle formation and fusion early in the secretory pathway. Cell 61(4): 723–733 Kreitzer G, Schmoranzer J, Low SH, Li X, Gan Y, Weimbs T, Simon SM, Rodriguez-Boulan E (2003) Three-dimensional analysis of post-Golgi carrier exocytosis in epithelial cells. Nat Cell Biol 5(2): 126–136 Ladinsky MS, Mastronarde DN, McIntosh JR, Howell KE, Staehelin LA (1999) Golgi structure in three dimensions: functional insights from the normal rat kidney cell. J Cell Biol 144: 1135–1149 Lippincott-Schwartz J, Yuan LC, Bonifacino JS, Klausner RD (1989) Rapid redistribution of Golgi proteins into the ER in cells treated with Brefeldin A: evidence for membrane cycling from the Golgi to ER. Cell 56: 801–813 Lucocq JM, Pryde JG, Berger EG, Warren G (1987) A mitotic form of the Golgi apparatus in Hela cells. J Cell Biol 104: 865–874 Marra P, Salvatore L, Mironov A Jr, Di Campli A, Di Tullio G, Trucco A, Beznoussenko G, Mironov A, De Matteis MA (2007) The biogenesis of the Golgi ribbon: the roles of membrane input from the ER and of GM130. Mol Biol Cell 18(5): 1595–1608 Marsh BJ, Volkmann N, McIntosh JR, Howell KE (2004) Direct continuities between cisternae at different levels of the Golgi complex in glucose-stimulated mouse islet beta cells. Proc Natl Acad Sci USA 101(15): 5565–5570 Mironov AA, Weidman P, Luini A (1997) Variations on the intracellular transport theme: maturing cisternae and trafficking tubules. J Cell Biol 138: 481–484 Mironov AA, Beznoussenko GV, Nicoziani P, Martella O, Trucco A, Kweon HS, Di Giandomenico D, Polishchuk RS, Fusella A, Lupetti P, Berger EG, Geerts WJ, Koster AJ, Burger KN, Luini A (2001) Small cargo proteins and large aggregates can traverse the Golgi by a common mechanism without leaving the lumen of cisternae. J Cell Biol 155: 1225–1238 Mironov AA, Mironov AA Jr, Beznoussenko GV, Trucco A, Lupetti P, Smith JD, Geerts WJ, Koster AJ, Burger KN, Martone ME, Deerinck TJ, Ellisman MH, Luini A (2003) ER-toGolgi carriers arise through direct en bloc protrusion and multistage maturation of specialized ER exit domains. Dev Cell 5: 583–594 Mironov AA, Beznoussenko GV, Polishchuk RS, Trucco A (2005) Intra-Golgi transport. A way to a new paradigm? BBA Mol Cell Res 1744: 340–350  DJ, Mollenhauer HH (1964) Isolation of Golgi apparatus from plant cells. J Cell Morre Biol 23: 295–305  DJ, Merlin L, Keenan T (1969) Localization of glycosyl transferase activities in a Morre Golgi apparatus-rich fraction isolated from rat liver. Biochem Biophys Res Commun 37(5): 813–819 Moya M, Dautry-Varsat A, Goud B, Louvard D, Boquet P (1985) Inhibition of coated pit formation in Hep2 cells blocks the cytotoxicity of diphtheria toxin but not that of ricin. J Cell Biol 101: 548–559

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Munro S, Pelham HRB (1987) A C-terminal signal prevents secretion of luminal ER proteins. Cell 48: 899–907 Nakamura N, Rabouille C, Watson R, Nilsson T, Hui N, Slusarewicz P, Kreis TS, Warren G (1995) Characterization of a cis-Golgi matrix protein, GM130. J Cell Biol 131: 1715–1726 Neutra M, Leblond CP (1966) Radioautographic comparison of the uptake of galactoseH and glucose-H3 in the Golgi region of various cells secreting glycoproteins or mucopolysaccharides. J Cell Biol 30: 137–150 Newman AP, Shim J, Ferro-Novick S (1990) BET1, BOS1, and SEC22 are members of a group of interacting yeast genes required for transport from the endoplasmic reticulum to the Golgi complex. Mol Cell Biol 10(7): 3405–3414 Novikoff A, Goldfischer S (1961) Nucleosidediphosphatase activity in the Golgi apparatus and its usefulness for cytological studies. Proc Natl Acad Sci USA 47: 802–810 Novikoff AB (1964) GERL, its form and function in neurons of rat spinal ganglia. Biol Bull 127: 358 Novikoff AV, Essner E, Quintana N (1964) Golgi apparatus and lysosomes. Fed Proc 23: 1010–1022 Oprins A, Rabouille C, Posthuma G, Klumperman J, Geuze HJ, Slot JW (2001) The ER to Golgi interface is the major concentration site of secretory proteins in the exocrine pancreatic cell. Traffic 2: 831–838 Orci L, Glick BS, Rothman JE (1986) A new type of coated vesicular carrier that appears not to contain clathrin: its possible role in protein transport within the Golgi stack. Cell 46: 171–184 Orci L, Perrelet A, Rothman JE (1998) Vesicles on strings: morphological evidence for processive transport within the Golgi stack. Proc Natl Acad Sci USA 95(5): 2279–2283 Ostermann J, Orci L, Tani K, Amherdt M, Ravazzola M, Elazar Z, Rothman JE (1993) Stepwise assembly of functionally active transport vesicles. Cell 75(5): 1015–1025 Ostermann J (2001) Stoichiometry and kinetics of transport vesicle fusion with Golgi membranes. EMBO Rep 2(4): 324–329 Pavelka M, Ellinger A, Debbage P, Loewe C, Vetterlein M, Roth J (1998) Endocytic routes to the Golgi apparatus. Histochem Cell Biol 109: 555–570 Pearse BM (1976) Clathrin: a unique protein associated with intracellular transfer of membrane by coated vesicles. Proc Natl Acad Sci USA 73(4): 1255–1259 € linger B, Melkonian M, Becker B (2000) The Golgi € ntrup IM, Bo Perasso L, Grunow A, Bru apparatus of the scaly green flagellate Scherffelia dubia: uncoupling of glycoprotein and polysaccharide synthesis during flagellar regeneration. Planta 210(4): 551–562 Polishchuk RS, Polishchuk EV, Marra P, Alberti S, Buccione R, Luini A, Mironov AA (2000) Correlative light-electron microscopy reveals the tubular–saccular ultrastructure of carriers operating between Golgi apparatus and plasma membrane. J Cell Biol 148 (1): 45–58 Polishchuk RS, San Pietro E, Di Pentima A, Tete S, Bonifacino JS (2006) Ultrastructure of long-range transport carriers moving from the trans Golgi network to peripheral endosomes. Traffic 7: 1092–1103 Presley JF, Cole NB, Schroer TA, Hirschberg K, Zaal KJ, Lippincott-Schwartz J (1997) ER-toGolgi transport visualized in living cells. Nature 389: 81–85 Puthenveedu MA, Bachert C, Puri S, Lanni F, Linstedt AD (2006) GM130 and GRASP65dependent lateral cisternal fusion allows uniform Golgi-enzyme distribution. Nat Cell Biol 8: 238–248 Roth J, Berger EG (1982) Immunocytochemical localization of galactosyltransferase in HeLa cells: codistribution with thiamine pyrophosphatase in trans-Golgi cisternae. J Cell Biol 93(1): 223–229 Roth J, Berger EG (eds) (1997) The Golgi apparatus. Basel. Birkhauser

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Rothman JE, Bursztyn-Pettegrew H, Fine RE (1980) Transport of the membrane glycoprotein of vesicular stomatitis virus to the cell surface in two stages by clathrincoated vesicles. J Cell Biol 86(1): 162–171 Rothman JE, Miller RL, Urbani LJ (1984) Intercompartmental transport in the Golgi complex is a dissociative process: facile transfer of membrane protein between two Golgi populations. J Cell Biol 99: 260–271 Sandvig K, Sundan A, Olsnes S (1985) Effect of potassium depletion of cells on their sensitivity to diphtheria toxin and pseudomonas toxin. J Cell Physiol 124: 54–56 Saraste J, Kuismanen E (1984) Pre- and post-Golgi vacuoles operate in the transport of Semliki Forest virus membrane glycoproteins to the cell surface. Cell 38(2): 535–549 Scales SJ, Pepperkok R, Kreis TE (1997) Visualization of ER-to-Golgi transport in living cells reveals a sequential mode of action for COPII and COPI. Cell 90: 1137–1148 Schekman R, Mellman I (1997) Does COPI go both ways? Cell 90: 197–200 Serafini T, Orci L, Amherdt M, Brunner M, Kahn RA, Rothman JE (1991) ADP-ribosylation factor is a subunit of the coat of Golgi-derived COP-coated vesicles: a novel role for a GTP-binding protein. Cell 67(2): 239–253 Slusarewicz P, Nilsson T, Hui N, Watson R, Warren G (1994) Isolation of a matrix that binds medial Golgi enzymes. J Cell Biol 124(4): 405–413 Swift AM, Machamer CE (1991) A Golgi retention signal in a membrane-spanning domain of coronavirus E1 protein. J Cell Biol 115(1): 19–30 Tabas I, Kornfeld S (1980) Biosynthetic intermediates of beta-glucuronidase contain high mannose oligosaccharides with blocked phosphate residues. J Biol Chem 255 (14): 6633–6639 Trucco A, Polishchuk RS, Martella O, Di Pentima A, Fusella A, Di Giandomenico D, San Pietro E, Beznoussenko GV, Polishchuk EV, Baldassarre M, Buccione R, Geerts WJ, Koster AJ, Burger KN, Mironov AA, Luini A (2004) Secretory traffic triggers the formation of tubular continuities across Golgi sub-compartments. Nat Cell Biol 6 (11): 1071–1081 Varki A, Cummings R, Esko J, Freeze H, Hart G, Marth J (1999) Essentials of glycobiology. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Wacker I, Kaether C, Kromer A, Migala A, Almers W, Gerdes HH (1997) Microtubuledependent transport of secretory vesicles visualized in real time with a GFP-tagged secretory protein. J Cell Sci 110: 1453–1463 Weisz OA, Swift AM, Machamer CE (1993) Oligomerization of a membrane protein correlates with its retention in the Golgi complex. J Cell Biol 122(6): 1185–1196 Whur P, Herscovics A, Leblond CP (1969) Radioautographic visualization of the incorporation of galactose-3H and mannose-3H by rat thyroids in vitro in relation to the stages of thyroglobulin synthesis. J Cell Biol 43: 289–311

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The Golgi apparatus as a crossroads in intracellular traffic Alexander A. Mironov and Margit Pavelka

In this chapter, we will briefly describe the structure and list main functions of different compartments along the secretory pathway.

ER–Golgi interface After its synthesis, folding and quality control a cargo exits from the endoplasmic reticulum (ER) and moves to the Golgi apparatus (GA) through the ER–Golgi interface. Here, the most peculiar structures are vesicular–tubular clusters (VTC) defined as clusters of a few small vesicles and tubular–saccular elements associated with the region of the rough ER and even with the nuclear envelope containing COPII-coated buds (Bannykh et al. 1996; Mironov et al. 2003). On routine EM sections, the size of VTCs varies from 200 to 1,000 nm. They are positive for glucose-6-phosphatase (Thorne-Tjomsland et al. 1991) and at immunofluorescence level, appear as COPII-positive distinct sites (0.5–1 mm in size) associated with the ER (Stephens et al. 1997). In interphase cells at steady state, the average number of VTC remains constant varying from several dozens to several hundreds of VTC per one cell (Bannykh et al. 1996; Aridor et al. 1999; Hammond and Glick 2000). Typically, COPII-coated buds are described as elevations on the surface of the ER with a width of 65–85 nm, extruded from the membrane by at least 50% of their diameter. Buds are covered with an 8–10 nm thick electron-dense COPII coat. On grazing sections, the buds possess a lattice-like appearance due to semi-regular array of 4–5 nm elongated particles arranged in a semi-regular pattern with mostly tetrahedron organization (Bannykh et al. 1996). Separated COPII-coated vesicles do exist (Zeuschner et al. 2006) although they are few and mostly are devoid of secretory proteins (Mironov et al. 2003). Some elongated profiles (which might represent cross-sections of saccules or tubules) within VTCs have a dense COPI-like coat at their tips and on their central parts (Martinez-Menarguez et al. 1999). VTCs are not carriers, which undergo centralization and deliver cargo to the GA (Stephens et al. 2000; Mironov et al. 2003; see details in Chapter 3.1). For this purpose, ER-to-Golgi carriers (EGCs) are used. These appear mostly as saccular containers filled with either the large supramolecular cargo (i.e. procollagen) or the small diffusible cargo proteins. They arise through cargo concentration and direct en bloc protrusion of specialized ER domains in the vicinity of VTC (Mironov et al. 2003).

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ER–Golgi connections Direct membrane continuity between the ER and the GA has been described in many papers over the years in variety of tissues and cell types under different functional conditions (Flickinger 1969, 1973; Maul and Brinkley 1970; Claude 1970; Bracker et al. 1971; Holzman 1971; Morre et al. 1974; Franke and Kartenbeck 1976; Novikoff and Yam 1978; Uchiyama 1982; Broadwell and Cataldo 1983; Sasaki et al. 1984; Williams and Lafontane 1985; Lindsey and Ellisman 1985a,b; Tanaka etal. 1986; Krijnse-Locker et al. 1994; Sesso et al. 1994; Stinchcombe et al. 1995; Trucco et al. 2004) and using different methods of analysis, including three-dimensional (3D) observation in high voltage electron microscope (Lindsey and Ellisman 1985a,b), scanning electron microscopy (Tanaka et al. 1986), reconstruction of serial sections (Sesso et al. 1994) and even functional analysis of transport (Krijnse-Locker et al. 1994). Connections were described between the ER and EGC (Stinchcombe et al. 1995; Mironov et al. 2003). After 3D tomographic reconstruction (Ladinsky et al. 1999), a connection between the ER and the membrane disk integrated between Golgi cisternae has been found. However, the nature of this disk is not established. The author interpreted this disk as the specialized domain of the ER. Recently, two reports about ER-to-Golgi connections have been published. In one (Koga and Ushiki 2006) the existence of connections between the ER and the GA was not confirmed. However, the method used is not completely free from artefacts. ER-to-Golgi connections cannot be a result of fixation, because fixative usually disrupt pre-existing tubules rather than induce their formation (McIntosh 2001). In the report by Vivero-Salmeron et al. (2008), the existence of ER-to-Golgi connections was confirmed. The relatively low frequency of these observations might simply be due to the fact that thin sections are technically unsuitable for revealing a convoluted and transient (Vivero-Salmeron et al. 2008) structure extending through a large three-dimensional space or due to temporality of the connections (Fig. 1, 2c–e).

The function of compartments within the ER-to-Golgi interface Within the ER-to-Golgi interface, several important posttranslational functions are performed: COPII-mediated concentration of defined membrane and soluble cargoes (to improve the efficiency of transport), delivery of cargo to the GA (see Chapter 3.1), O-glycosylation (Tooze et al. 1988), acylation (Rizzolo et al. 1985), generation of mannose-6-phosphate signal for lysosomal protein targeting (Pelham et al. 1988), protein palmitoylation (Bonatti et al. 1989), retrieval of misfolded proteins (Hammond and Helenius 1994), and segregation of secretory cargoes, namely, regulatory secretory

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Figure 1. Scheme of structures within the ER–Golgi interface. The ER is shown as the structure filled with a grey content and covered with ribosomes (dark dots). The ER has a specialized domain that is attached to the Golgi stack (trans-ER). At the cis-side of the stack, the ER contains the ER exit site (ERES, or early intermediate compartment [IC] or vesicular tubular cluster; arrow). The ERES can include COPII-dependent vesicles (arrow). Near the ERES, there is the forming precursor of an ER-to-Golgi carrier (EGC, arrow). The cis Golgi network is composed of several domains. One represents the highly perforated disk similar in shape with Golgi cisternae. It is attached to the Golgi stacks (G) and is named as attached CGN (AC). There is a small part of the total CGN that appears as the three-dimensional tubular network or a cage (indicated as CGN) near the Golgi stacks. This part produces tubules moving towards the ERES (in the center of the image). Another part of the CGN is connected with the AC by tubules and has similar shape with the CGN. It localizes out of the Golgi stack and appears as the late IC that could be rather stable compartment. At the trans-side of the Golgi stack (G), there is the trans-Golgi network (TGN), parts of which reside apart of the stack, and are connected with the other part, the attached TGN (AT). The TGN contains clathrin-coated buds (double arrow). Precursors of Golgi-to-PM carriers (GPC) could form from the last two COPI-positive (with COPI-buds) cisternae, or from the entire TGN. COPI-vesicles are present near the rims of Golgi cisternae.

proteins, constitutive secretory proteins, proteins destined for the apical plasma membrane (PM) and basolateral PM, endosomal and lysosomal proteins by elimination of the mechanisms responsible to their retention within the ER. The role of microtubules in centralization of EGCs is described in Chapter 2.14. However, this function is absent in plant and yeast cells (Nebenfuhr and Staehelin 2001) and even in several cell types in mammals (such as oocytes, Motta et al. 1995) and, thus, might be not directly related to ER–Golgi transport per se. Transient ER–Golgi connections could serve for the diffusion of cargo proteins and recycling of resident proteins.

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Morphology of the Golgi apparatus The structural organization of the GA varies among species (see Chapters 4.1–4.4). In yeast (Chapter 4.3), and some protists (Chapter 4.4), the GA is composed of tubular networks and isolated disks. In animal and plant cells, the GA appears as a series of closely associated flattened membrane ‘sacs’ aligned in parallel to form a stack (Polishchuk and Mironov 2004; Chapter 4.1). In plants, S. cerevisiae, protists and some insect cells, stacks remain separated from each other whereas in mammalian cells stacks form the single ribbon (Chapter 2.12). Here, we describe mainly the mammalian GA. The GA is embedded into a (so-called) “zone of exclusion”, a polymer-based derivative of the cytosol that is especially evident in plant. This zone does not contain ribosomes but cytoskeleton elements can pass through it (Mollenhauer and Morre 1978). In epithelial cells of epididymis, goblet cells in the jejunum, gonadotrophs in pituitary glands and dorsal root ganglion cells, an “empty” zone of exclusion also surrounds the GA with the thickness of about 200 nm (Koga and Ushiki 2006). The GA represents 2% of hepatocellular membrane (Blouin 1983) or 20% of that of the ER system (Griffiths et al. 1989). The GA is capable to undergo rapid and reversible reorganization in response to a variety of experimental manipulations (Polishchuk and Mironov 2004). According to Griffiths et al. (1995), the GA begins from the place where mannosidase I is localized. Man I exhibits a highly polarized staining at the cis-pole of Golgi stacks usually composed of the first, sometimes of the first two cisternae (Marra et al. 2001). The canonical GA consists of a series flattened cisternal membranes closely associated, aligned in parallel and forming a stacked structure, abundant tubular–reticular networks and vesicles. In the perinuclear area, dozens or hundreds of Golgi stacks are linked together to form an interconnected, ribbon-like structure as a single organelle with alternating compact (stacked cisternae) and non-compact (tubular–reticular) zones (Mogelsvang et al. 2004; see also Chapter 2.12). The GA can be viewed schematically as being composed of three main compartments: the cis-, medial- and trans-Golgi. Both the cis- and the transmost Golgi elements are largely tubular. The medial-Golgi stacked cisternal compartment resides between these two networks (Rambourg and Clermont 1997; Polishchuk and Mironov 2004). Several gradients exist within a Golgi stack: (i) a gradient in the cisterna fenestration; (ii) a gradient in the cisterna thickness; (iii) a gradient in the localization of the Golgi enzymes; (iv) a gradient in the lipid bilayer thickness; (v) a gradient in the pH; (vi) a gradient in concentration of cholesterol. The concentration of cholesterol is higher at the trans-side of a Golgi stack. Especially high concentration of cholesterol is found in endosomes (Orci et al. € bius et al. 2003). 1981; Cluett et al. 1997; Mo

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The cis-to-trans changes in cisterna fenestration and thickness mean that the size of the fenestrae and the wells initially become smaller towards the medial cisternae, while the thickness of the cisternae decreases from the cis to the trans-pole. Then, still in a trans-wise direction, the cisternae become more perforated again (Ladinsky et al. 1999). The enzymes involved in the early stages of glycosylation are located mostly at the cis-side, whereas the late and terminal glycosylation enzymes are situated at the trans-side of the stacks. The trans-compartments are enriched in terminal processing enzymes involved in sialylation. However, a significant difference in the distribution of the Golgi enzymes can be detected only between the cis-most and trans-most cisternae of the central

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Golgi domain, and not between adjacent cisternae of the same stack (Rabouille et al. 1995). Thus, a single cisterna does not necessarily represent a separated Golgi compartment from either the biochemical or the structural point of view, potentially representing instead a part of a larger compartment.

Cis-Golgi network (CGN) The cis-Golgi network is composed of several domains. One represents the highly perforated disk similar in shape with Golgi cisternae. It is attached to the Golgi stacks and is named as the attached CGN or the Cis-perforated cisternae of the intermediate compartment or CGN (CISCIC). CISCIC appears as a disk with 30 nm perforations. This is especially evident in epithelial cells of epididymis, goblet cells in the jejunum, gonadotrophs in pituitary glands and dorsal root ganglion cells (Koga and Ushiki 2006). It is highly labelled for COPI (Oprins et al. 1993; Kweon et al. 2004). The second part of the CGN is a small tubular part of the total CGN that appears as the three-dimensional tubular network or a cage near the Golgi stacks. Let’s name it as the free CGN. This part produces tubules moving towards the ER-export sites (ERES; Marra et al. 2001; Mironov et al. 2003). Another part of the CGN is connected with the attached CGN by tubules and has similar shape with the CGN. It localizes out of the GA and appears as the late intermediate compartment (Marra et al. 2001) that could be rather a stable compartment (Ben-Tekaya et al. 2005). In spermatides, the cis-most cisterna appears as a regular network of anastomotic membranous tubules, and the medial saccules usually have fewer but larger irregular fenestrations in them (Ho et al. 1999). There, the cis-elements of Golgi stacks were slightly reactive for G6Pase. Labelling can be seen in some cis-Golgi cisternae. The CGN contains also early processing enzymes such as alpha-mannosidases that trim high mannose N-linked

"

Figure 2. Tomographic 3D reconstruction of different compartments of the secretory pathway. A,B Comparison of the resting (A) and transporting ministacks (B) in fibroblast-like cells. Ministacks are formed after the complete depolymerization of MTs (3 h of treatment with nocodazole). The resting stack (the stack before the release of the block of intra-Golgi transport) contains more vesicles (white spheres) and less cisternae. In resting stack (A), the cis-most and the trans-most cisternae are almost absent whereas in the transporting stack (B) both the transcisterna (white arrow) and the cis-cisterna (red arrow) are visible. In the resting stack (A), the cismost cisterna is replaced by the tubular network (white arrow). C–E Structure of moving (C,D) and stationary (E) ER-to-Golgi carriers. The ER is pictured in green. The uncoated domains of the EGCs are pictured into brownish. The domains coated with COPII-like coat are pictured into yellow whereas the buds coated with COPI-like coat are indicated by light-blue colour. Samples were prepared according to the correlative light-electron microscopy (Mironov et al. 2003) 20 min after the restoration of ER-to-Golgi transport of tsVSVG. F,G Structure of the Golgi exit site at the moment of the formation of Golgi-to-PM carriers filled with tsVSVG (12 min after the release of the ER exit block according to the small pulse-chaise protocol (Mironov et al. 2001). Connections (arrow) between the COPI-positive Golgi cisterna and the TGN (yellow network) are shown from different points of view. Red arrows show the Golgi stack. The ER is pictured in green. Models A and B were made by A. Trucco, Models C–G were made by G. V. Beznoussenko. Bar: 120 nm.

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oligosaccharides added to the nascent chain in the ER (Thorne-Tjomsland et al. 1991). The CGN receives newly synthesized or recycled polypeptides from the endoplasmic reticulum, which are then posttranslationally modified by glycosylation, sulphation, phosphorylation, palmitoylation, myristoylation or methylation (De Graffenried and Bertozzi 2004).

Cisternal shape Cisternae are oriented along microtubules. Although the number of cisternae in the Golgi stacks varies from one cell type to another (three to eight cisternae, in the majority of cases), however within the same cell line, the number of cisternae could be constant representing a specific characteristic of cell type (Ladinsky et al. 1999) or varies depending on the functional state. At least, this number is reproduced in cells washed out after brefeldin A and nocodazole treatment. The number of not perforated COPI-positive cisternae is almost not changed after the arrival of cargo (Trucco et al. 2004; see Chapter 2.16). All Golgi cisternae have roughly the same surface area, although they can differ in volume by as much as 50% (Ladinsky et al. 1999). The length of all of the cisternae within the stack is equal. When the GA becomes fragmented, this feature becomes particularly evident even when cargo is being transported through the GA. Even after arrival of cargo to the cis-side of the GA, the length of all Golgi cisternae rapidly became equal (Trucco et al. 2004). Both the cis-most perforated cisterna and all medial cisternae contain COPI-coated buds, whereas the trans-most perforated cisterna(e) contains only clathrin-coated buds and usually have no COPI-coated buds (Ladinsky et al. 1999). All Golgi cisternae are fenestrated; the existence of Golgi cisternae without these fenestrae has yet to be demonstrated, at least in mammalian cells. The large openings in cisternae can often form “wells” (Ladinsky et al. 1999). These fenestrae are necessary for movement of secretory granules (SGs; Rambourg and Clermont 1997). The lumen of a Golgi cisterna is usually quite narrow (10–20 nm). There is a systematic decrease in luminal diameter in the trans-direction in quick-frozen NRK cells (Ladinsky et al. 1999). Mechanisms responsible for Golgi cisterna stacking and maintenance of the cisterna shape remain mostly unknown. Attempts to explain stacking by the presence of so-called Golgi matrix proteins or Golgins (see Chapter 2.12) were not successful (Seemann et al. 2000) because these proteins are not present between all Golgi cisternae. Low affinity antiparallel dimerization of cytosolic domains of sugar transporters might be responsible for attachment of Golgi cisternae to each other. At least overexpression of GDP-mannose transporter in the yeast Saccharomyces cerevisiae induces formation of the stacked GA (Hashimoto et al. 2002). Stacks are also formed in S. cerevisiae after deletion of function of Sec7

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(Rambourg et al. 1993). In any case, additional analysis is necessary to resolve this problem.

Golgi vesicles There are several known coats along the secretory pathway participating in the formation of coat-dependent vesicles. The most important of them are the following: COPII (see above), COPI, clathrin. Each of them contains several variants of the coat. COPII-based coat has two main forms (see Chapter 2.3), COPI-based coat also has two forms (see Chapter 2.4), and clathrin coat has two or four variants interacting with different adaptors, namely, AP1, AP2, AP3 and AP4 (reviewed by Traub 2005). Membrane budding with the help of these coats could induce generation of corresponding coat-dependent vesicles. So far, only four types of coat-dependent small vesicles have been found in cells, namely, clathrin-coated vesicles that could be formed from clathrin/AP1coated buds present on the TGN attached to COPI-positive stacked Golgi cisternae, secretory granules, endosomes and clathrin/AP2-coated buds found on the PM. COPII-dependent vesicles are formed near the ER exit sites. COPI-dependent vesicles are formed at the level of the ER-to-Golgi carriers, the intermediate compartment, the cis-Golgi and medial Golgi (reviewed by McMahon and Mills 2004). One of the most important features of the GA is the presence of small 52 nm COPI-dependent vesicles surrounding each Golgi cisterna (Ladinsky et al. 1999; Marsh et al. 2001). COPI vesicles do not appear to be really “free” because most of them are unmistakably tethered to neighbouring vesicles and/or to the Golgi membranes (Orci et al. 1998). This explains why vesicles do not diffuse towards the cell periphery. In contrast, most of the clathrindependent or irregularly shaped “vesicles” are clustered away from the GA (Ladinsky et al. 1999). A dense COPI coat observed on Golgi buds/vesicles on thin sections appears as a lace-like cytoplasmic structure closely attached to the lipid bilayer and composed of a series obtuse spikes separated by an average of 20 nm centerto-center. These spikes do not have the “bristle” or “spiny” appearance of clathrin subunits in basketworks (Orci et al. 1986). The thickness of COPI coat is about 10 nm whereas the thickness of clathrin coat is 18 nm (Oprins et al. 1993). The number of vesicles is the result of the equilibrium between two processes – the activity of COPI machinery and the activity of SNARE/Ca machinery. Inhibition of COPI activity causes reduction of the number of COPI-dependent vesicles. Similar effects are observed, when the COPI/ARF machinery is inhibited with brefeldin A In contrast, when the SNARE machinery is inhibited, the number of vesicles increases (Kweon et al. 2004). Most of data suggest that these vesicles are formed from COPI-coated buds abundant along the cisternal rims. COPI-dependent vesicles could derive

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from varicosities within tangential tubules along the rims of Golgi cisternae (Weidman et al. 1993). There could be following functions for COPI vesicles. 1. Formation of COPI vesicles could be specific mechanisms controlling the geometry of the Golgi elements (see Chapter 2.16). 2. COPI vesicles could control the fusion between adjacent Golgi cisternae extracting Qb SNAREs from there. COPI-dependent vesicles are two-fold enriched in Qb-SNAREs of the same type: membrin and GOS28 (Trucco et al. 2004; our unpublished observations) and depleted of syntaxin 5 (Orci et al. 2000a,b). Extracting GOS28 and membrin from the Golgi membrane, COPI vesicles prevent fusion between the cisternae (Trucco et al. 2004). 3. Formation of coat-dependent vesicles could be the way for fast uncoating. The Role of COPI vesicles as anterograde or retrograde carriers will be analyzed in Chapter 3.2.

Intercisternal connections One of the important questions of Golgi morphology is the issue of intercisternal heterogeneous connections (Tanaka et al. 1986; Rambourg and Clermont 1990, 1997; Sesso et al. 1994). In some cells, the Golgi forms even a single continuous membranous system (Tanaka et al. 1986; Inoue 1992; Rambourg and Clermont 1990, 1997). At steady state and in unstimulated cells, the connections are rare (Marsh et al. 2004). Intercisternal connections are augmented after arrival of cargo to the GA (Trucco et al. 2004) or when the islet beta cells have been stimulated for 1 h with 11 mM glucose (Marsh et al. 2004). Recently, the existence of such connections has been confirmed at steady state (Beznoussenko et al. 2006; Vivero-Salmeron et al. 2008). The connections between cisternae at different levels of the GA are of three types. The first type is observed at points, where the Golgi ribbon branches (Rambourg and Clermont 1997). Cisternae could be connected at both equivalent and non-equivalent levels. This “Y” configuration of cisternae at the branch in the Golgi ribbon also means that there is direct continuity between cisternae at different levels around the periphery of the upper stack of Golgi membranes. The second type of connection occurs when one cisterna projects through an opening (fenestration) in an adjacent cisterna to form a continuous lumen with its next-nearest neighbour. In the third type of connection, membrane tubules connecting non-equivalent cisternae bypass interceding cisternae at the periphery of the stack in Golgi regions where the ribbon is unbranched (Marsh et al. 2004; Trucco et al. 2004). Intercisternal connections were detected rarely because on conventional EM sections luminal continuity between consecutive cisternae is almost undetectable (Marsh 2005).

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The tubules connecting different stacks have been proposed to participate in intra-Golgi traffic (Mellman and Simons 1992; Weidman 1995). These connections could be used for the transport of soluble cargo, like albumin, or serve for the movement of proton from the trans- to the cis-compartments, or for Golgi enzyme diffusion (Trucco et al. 2004). The speed diffusion of ions, lipids and even transmembrane proteins along Golgi lumen and membranes is high. For instance, the speed of cholesterol diffusion in clear lipid bilayer is about 2 mm/s. The speed of ceramide diffusion is about 0.4–0.5 mm/s (Cooper et al. 1990). If connections are constant, the ionic, lipid and protein gradients that are known to exist between the Golgi poles (reviewed in Mironov et al. 1998, 2005) have to be expected to dissipate through these continuities. However, since this is not the case the connections have to be transient and highly regulated as within the framework of kiss-and-run models of transport (see Chapter 3.2).

Function of the Golgi apparatus In mammalian cells, occupying a central position, the GA plays a central role in the classical secretory pathway, as well as in endocytic pathways, and in multiple recycling routes. 1. The GA exchanges membrane components with several other subcellular organelles, including endosomes, caveosomes, autophagosomes, and lipid droplets participating in sorting (Mironov et al. 2005). 2. The GA is the main station of cellular glycosylation. During movement along the Golgi membranes, cargoes undergo glycosylation. The Golgi stack is composed of a series of compartments containing oligosaccharide processing and other enzymes that are generally arranged in a cis-totrans orientation (Rabouille et al. 1995). 3. Assembly of triglycerides with apoB and other apolipoproteins occurs in the GA. During this process, apoB undergoes conformational changes, and the expanding lipoproteins recruit more apoE (Gusarova et al. 2007). 4. The GA is involved in various other cellular processes such as transcription, apoptosis, and mitosis via signalling pathways mediated by Ras proteins, protein kinases, and G proteins (Helms et al. 1998; DeBose-Boyd et al. 1999; Lane et al. 2002; Sutterlin et al. 2002; Bivona et al. 2003; Nardini et al. 2003; Preisinger et al. 2004). 5. The GA provides a connection between exocytosis and endocytosis.

Structure of the Golgi exit site After its passage through the GA cargo exits from it, and moves to the sites of its destinations. In the literature, this Golgi exit site (GES) is usually called as the trans-Golgi network (TGN, Griffiths and Simons 1986; Ladinsky et al. 1994; Clermont et al. 1995) suggesting that the TGN is composed of distinct tubules

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with little indication of anastomosis. However, this name is misleading because the term TGN reflects mostly the structural organization of only one compartment, the whole TGN, within GES (Fig. 2f, g). The identity of the GES could be defined by membrane components delivered to the TGN from both anterograde and retrograde transport, together with the recruitment from the cytosol of coat proteins, regulatory GTPases, fusion and Golgi matrix and motor proteins. Among the most important proteins localized within the TGN, one could observe clathrin, M6PR, TGN38, AP1, furin, GGA, EEA1, clathrin, Golgin-97, and other transGolgins (reviewed in Robinson and Bonifacino 2001). Two to four Golgi cisternae are stained with ceramide (Pagano et al. 1989) and TPPase (Novikoff et al. 1971). The GESs are composed of the two last (trans) COPI-positive cisternae of the Golgi stack (Ladinsky et al. 2002), the trans-most perforated cisterna(e) with clathrin-coated buds, and the network of tubules surrounding the stack near its trans-pole and sometimes being continuous with either multiple trans-cisternae (Rambourg et al. 1979) or with only the trans-most cisterna in the stack (Griffiths and Simons 1985; Griffiths et al. 1989). “Peeling off” configurations of the last Golgi cisterna with clathrin-coated buds are frequently described in the past (Ladinsky et al. 1994; Clermont et al. 1995). This trans-most cisterna contains exclusively clathrin-coated buds, whereas the other cisternae have COPI-coated buds only (Ladinsky et al. 1999, 2002). Only 12% of the total TGN surface area is attributable to the flattened cisternal part of the TGN which is labelled by the presence of TPPase and which is morphologically indistinguishable from the other cisternae of the Golgi stack. In many cases most of the tubules located within the TGN area are devoid of the reaction for TPPase (Griffiths et al. 1989). The trans-most cisterna containing clathrin-coated buds actually represents a highly perforated disk containing clathrin-coated buds along its rims (Ladinsky et al. 1999). This perforated cisterna is accessible for WGAHRP added from outside (Pavelka et al. 1998). Thus, it represents a part of the endocytic TGN connected with endosomes. As such, this trans cisterna of the endocytic trans-Golgi network could be named as the TRANSCET or the attached TGN. In epithelial cells of the epididymis, goblet cells in the jejunum, gonadotrophs in pituitary glands and dorsal root ganglion cells, the TRANSCET appears as perforated sheet, varicose tubules or small plates connected with each other two-dimensionally by tubules (Koga and Ushiki 2006). Structures within GES are highly dynamic and continuously undergo renewal (Clermont et al. 1995). Tubules continuously emanate from the Golgi cisternae going towards the cell periphery (Cooper et al. 1990). The structure of the GES depends on the cell type. In cells, where secretory granules (SGs) are not seen being associated with the GA, the non-attached TGN appears as a tubular network connected with last two Golgi cisternae. However, the TGN does not form the continuous ribbon along the Golgi

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ribbon. In the second group of the cells, where the SGs are seen exclusively within the trans-most cisterna of the stack or within the TGN, the TGN here is small. The SGs are formed within the TGN. In the third group of cells, immature secretory granules (ISGs) are formed as distensions of all Golgi cisternae with the exception of the cis-most perforated cisternae. The TGN here is almost invisible (Clermont et al. 1995). In spermatides, the TGN is also composed of irregular saccules fenestrated near the edge, and long strings of connected vesicles. However, only the edge portions of the trans-Golgi cisternae but not the whole cisternae are fenestrated (Ho et al. 1999). In the cells not possessing secretory granules, where MTs are depolymerized, the non-attached TGN still exhibits tubular organization but its size is reduced (Trucco et al. 2004). The ER cisternae attached to the TRANSCET or to one of the two of the last medial cisternae (these last medial cisternae usually exhibit the presence of TPPase activity; Paavola 1978a,b,c) are one of the most fundamental features of the Golgi exit site. This trans-ER described in multiple cell types has ribosomes bound to their surface, while their other side associates over considerable distances with Golgi cisternae (Novikoff et al. 1964; Pavelka and Ellinger 1983, 1986; Hermo and Smith 1998; Ladinsky et al. 1999; Marsh et al. 2001). The contact between the trans-ER and the trans-GA could serve for direct lipid transfer by, i.e. the ceramide-transfer protein, the oxysterolbinding protein and others (De Matteis et al. 2007). At the level of the GES, two morphologically distinct coats have been identified, namely, clathrin-based and lace-like (Ladinsky et al. 1994). Here, clathrin coat could form three different variants using adaptors AP1, AP3, AP4 and the GGAs. It has been postulated (but never proved) that there exists an additional coat composed of p62 and p200. The p200 is BFA-sensitive protein localized on Golgi membranes (Narula et al. 1992; Narula and Stow 1995). TGN tubules have varicosities coated by either clathrin or lace-like coat. Each individual tubule is covered by only one type of coat (Ladinsky et al. 1994). Within GES, there are only few clathrin-coated vesicles (Ladinsky et al. 1994, 1999). Many vesicles seen in the routine sections through TGN are actually continuous strings of vesicles (Ho et al. 1999).

Function of the Golgi exit site The first main functions of the GES are protein and lipid posttranslational and postsynthetic modifications. The compartments of GES contain a number of resident enzymes involved in the processing of cargo molecules, such as glycosyltransferases involved in the addition of terminal sugars (Rabouille et al. 1995), several pro-protein convertases including furin (Thomas 2002), and tyrosine sulphation enzymes. For instance, glycosamino glycan chains are synthesized and sulphated in the trans-Golgi/TGN, when cells are incubated with a membrane permanent xyloside (Farquhar 1985; Velasco et al. 1988).

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On the other hand, GES is at the crossroads of many transport pathways (Pavelka et al. 1998; Medigeshi and Schu 2003; Shewan et al. 2003; Sannerud et al. 2003; Young et al. 2005; Bonifacino and Rojas 2006). Therefore, the precise sorting is one of the main functions of the GES. This occurs when proteins undergo packaging into their appropriate membrane carriers for delivery not only to the cell surface but also to a number of other compartments (Griffiths and Simons 1986; Traub and Kornfield 1997; Keller and Simons 1998). Signals used for the sorting of proteins at GES for the apical PM (O-glycosylation and N-glycosylation) and endosomal/lysosomal deliveries are discussed in Chapters 3.5–3.7.

Role of cargoes in the structure of the Golgi apparatus Cargo proteins modulate structure and function of the GA. In yeast, a 3 h treatment with cyclohexamide leads to the complete disappearance of the GA (Morin-Ganet et al. 2000). The size of the GA is changed under different conditions. In mammalian cells, the GA tends to become larger when transport intensifies, while it declines in the absence of cargo input (Rambourg et al. 1993; Mironov and Mironov 1998; Trucco et al. 2004). The volume of the TGN depends on the cargo transported (Griffiths et al. 1989). When transport through the GA was inhibited, tubular–reticular membranes in the trans-Golgi area were not detected in tomograms (Trucco et al. 2004). In contrast, when more lysosomal enzymes are produced, there is also proliferation of the TGN (Decker 1974; Novikoff and Novikoff 1977; Paavola 1978a,b,c; Blest et al. 1978; Morre et al. 1979). There are many descriptions of transporting and not transporting stacks, although these are not systematic. Here, we list the main features of transporting and non-transporting stacks (Fig. 2a, b). 1. In the absence of intra-Golgi transport, the Golgi ribbon is fragmented (Marra et al. 2007; see also Chapter 2.12). 2. In the resting stacks, the volume of the GA is smaller. For instance, after accumulation of VSVG at the no permissive temperature the surface area of Golgi membrane reduces 1.6-fold (Aridor et al. 1999). During the starvation of cells, the volume of the GA declines (Mironov and Mironov 1998). In random sections, cisternae of resting stacks are shorter (Marra et al. 2007). In mammalian prolactin cells, blockage of prolactin secretion leads to a reduction of size of the GA (Rambourg et al. 1993). 3. Blockade of protein synthesis with cyclohexamide for 4 h reduces the size of the GA and leads to the formation of very thin cisternae in onion-like Golgi stacks (Taylor et al. 1997). However, in mammals, even 6 h after the blockage of protein synthesis the GA is still present (Beznoussenko et al. submitted). During the stimulation and inhibition of prolactin transport in lactating rats, the GA tends to decline in the absence of cargo input and

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5.

6. 7.

8.

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becomes larger when transport intensifies and when the medial Golgi cisternae increase in size (Rambourg et al. 1993). In contrast, when cargo influx increases sharply (e.g. 5 min after ERtransport-block release), the GA immediately grows in size (Aridor et al. 1999). For instance, synchronous release of the pre-Golgi transport block could induce a 3-fold augmentation of the Golgi surface (Trucco et al. 2004). Amplification of the GA is observed in cells secreting human growth hormone (Rudick et al. 1993). In transporting stacks, there are more COPI-dependent 52 nm vesicles than in resting stacks (Rambourg et al. 1993; Trucco et al. 2004). In starved cells, the volume of COPI-dependent vesicle is 20% higher (Mironov and Mironov 1998). In resting stacks, there are no intercisternal connections (Trucco et al. 2004). In resting stacks, there is no cis-most highly perforated cisterna positive for GM130. For instance, at 40 C, the no permissive temperature for tsVSVG, when VSVG is accumulated in the ER after 3 h of the temperature block these perforated cisternae were not visible (Marra et al. 2007; our unpublished observations). Two hours after the blockage of hormonal stimulation of prolactin producing cells the cis Golgi network disappears from the stacks. Lindsey and Ellisman (1985b) demonstrated that in neurons in the very same cell the cis side of the GA could be in five different stages forming the circle of function. In resting stacks, there is a reduction of the TRANSCET (Rambourg et al. 1993; Beznoussenko et al. submitted). The TGN augments when cells are stimulated to secrete (Rambourg et al. 1993, Fig. 3) or when more lysosomal enzymes are produced (Novikoff and Novikoff 1977; Paavola 1997, 1978a,b,c; Blest et al. 1978). In resting stacks, the trans-ER is attached not to the TRANSCET but to the last COPI-positive cisterna, or not attached at all.

Usually, at steady state, the features of the GA are more similar with transporting stacks than to the resting ones.

General principles of intracellular transport Currently, there are three basic principles for the intracellular and in particular intra-Golgi transport, namely, progression, dissociation and diffusion (Fig. 4). Their modifications and combinations of two or more principles in one model give the whole list of existing models of intra-Golgi transport. We think the same situation is present not only for intra-Golgi transport but also for all main steps of intracellular transport, such as ER-to-Golgi and post-Golgi transport. The essence the of progression model could be expressed as the following. At the ER exit sites, there is a formation of large containers for

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Figure 3. Tomographic 3D reconstruction of the Golgi exit site during the formation of the precursors of GPCs containing procollagen I (PCI). Samples are prepared at 12 min after the release of the ER exit block for PCI, according to the small pulse-chase protocol (Mironov et al. 2001). A Virtual tomographic section. B–D The 3D model. Each PCI distension (empty cavities coloured in yellow) is covered by the pieces of the trans-most cisterna (green) at least from one side. The single exception is the distension indicated with white arrows in B and D, where the distension appears between the trans-most cisterna and the additional cisternae (pictured in orange). The late endosome (red arrows) contains internal vesicles (blue). The TGN (green) has the connection (blue arrow in B) or a very close association with the plasma membrane (indicated by red lines). During maturation of GPCs these are surrounded by the trans-most cisterna. Models were made by G. V. Beznoussenko. Bars: 150 nm.

transport of cargo and these containers move through the entire pathway of intracellular transport without change of their composition and size. The compartments along the secretory pathway are not stable and each of them contains all proteins necessary for posttranslational modification of secretory proteins. In contrast, the dissociation models implies that the specialized small (much smaller than the compartments) transport carriers that are formed on the proximal (closer to the ER along the secretory pathway) compartment, detach from that compartment, and are delivered (by simple diffusion through the cytosol or with the participation of cytoskeleton motors) to the consecutive but more distal (closer to the PM) compartment, and then dock and fuse with it. Importantly, the fission of TCs occurs before the fusion. Therefore consecutive compartments do not have a common lumen at any time. The single variant of the dissociation model is the vesicular model (VM). One of the less known version of the vesicular model implied that tubules serve as retro-

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Figure 4. Scheme showing the main principles of intracellular transport. A The dissociation (mainly vesicular) model of intracellular transport. Initially, membrane buds are formed on the first (proximal) compartment with the help of protein coat and then, after fission and subsequent uncoating, coat-dependent vesicles move to the second (distal) compartment and are captured by tethering system. Then using SNAREs, the vesicle fuses with the second compartment. B The progression model of intracellular transport. Initially the large membrane protrusion is formed from the first (proximal) compartment with the help of not-coat mechanisms. This protrusion concentrates cargoes or contains them at the same concentration as in the first compartment. The large carrier is captured by the tethering system and with the help of SNAREs undergoes fusion with the second (distal) compartment. C The lateral diffusion model of intracellular transport.

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grade carriers (Lippincott-Schwartz 1993). The VM has been proposed by Palade (1975) and further developed by Rothman (1994). Within the framework of VM, membrane budding is the function of the coat proteins (Antonny and Schekman 2001; Bonifacino and Glick 2004; see also Chapter 2.3). Now, several coat complexes functioning within the secretory pathway are known: COPII (ER-to-CISCIC), COPI (CISCIC-to-Golgi; Golgi-to-ER; intra-Golgi: anterograde and retrograde transport), clathrin-AP1 (trans-Golgi networkto-lysosomes). At the TGN level, the existence an additional coat composed of p62 and p200 has been postulated (Zehavi-Feferman et al. 1995; Narula and Stow 1995; Ikonen et al. 1997; as well as clathrin-AP-3 and AP-4 (see Chapters 3.5–3.9). According to the VM, the compartments are stable, isolated entities. Secretory proteins move through the secretory pathway from each compartment to the next in discrete membrane-bound small coat-dependent vesicles. The process of transport at each transport step includes formation of coated buds, concentration of cargoes inside of them, detachment of coated buds with the formation of small 50–100 nm spheres and then their uncoating and fusion with the consecutive compartment. The diffusion mechanism is based on the presence of membrane and luminal connections along the secretory pathway and transport occurs by simple diffusion from the proximal to the distal compartment. In the following chapters, mechanisms of transport will be specified for each step of exocytosis.

Conclusion Structure and functions of the compartments along the secretory pathway are extremely complicated and their morphology depends on their functional state. Analysis of their functional roles within the framework of the transport models together with the precise mapping of known protein machines will be presented in the following chapters.

Abbreviations CISCIC CGN EGC EM ER ERES GA GES PM TRANSCET VM VTC

cis-cisterna of the CGN cis-Golgi network ER-to-Golgi carrier electron microscopy endoplasmic reticulum ER exit site Golgi apparatus Golgi exit site plasma membrane trans-cisterna of the endosomal TGN vesicular model vesicular–tubular cluster

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Main machineries operating at the Golgi apparatus

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SNAREs David K. Banfield and Wanjin Hong

Introduction Eukaryotic cells contain multiple membrane-bound compartments between which proteins and lipid molecules are continually shuttled via membranebound vesicular carriers. Despite the constant flux of proteins and lipid through these compartments their functional and composition integrity is maintained. While the molecular machinery involved in vesicle recognition and fusion can often be transport-step/fusion-event specific, one group of proteins – the SNAREs (soluble N-ethylmaleimide-sensitive factor attachment protein receptors) play a common and central role in this process. Transport-step-specific combinations of SNARE proteins, localized to the vesicle and the target organelle, form complexes that facilitate the final step leading to the fusion of vesicles with their cognate target organelles. In general, the role of SNAREs appears to be conserved irrespective of their location of function in the cell, and much of what has been established for SNAREs in a particular trafficking pathway or organelle, is broadly applicable to SNAREs that function in the Golgi. Here we review Golgi SNAREs and the role they play in membrane and protein trafficking in the Golgi apparatus with, a particular emphasis on their functions in yeast and human cells.

General features of Golgi SNAREs The majority of SNARE proteins that function in the Golgi are type II integral membrane proteins anchored in the lipid bilayer by virtue of their single C-terminal transmembrane domain (TMD) see Fig. 1. The TMDs of SNAREs are presumably crucial for the stable association of SNARE proteins with membranes, but also play a role in establishing the steady-state distribution of SNAREs in the Golgi (Banfield et al. 1994; Rayner and Pelham 1997; Watson and Pessin 2001). In addition, in vitro fusion assays have established that the transmembrane domains of v-SNAREs (Xu et al. 2005) and of Qa-SNAREs (Han et al. 2004) are important for the formation of the hemi-fusion intermediates that precede membrane fusion and vesicle content mixing with the target compartment. Adjacent to the TMD is a short stretch of amimo acids (10 in length) referred to as the membrane proximal region (MPR). The amino acid sequence and length of this region is not evolutionarily conserved. The MPR

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Figure 1. General features of Golgi SNAREs. Based on their structural and functional features Golgi SNAREs are grouped into four categories. Category I is comprised of the Qa-SNAREs and some Qb- and Qc-SNAREs. Category II is mainly comprised of the Qc-SNAREs. Category III the R-SNARE Sec22p/Sec22b and category IV the R-SNARE Ykt6. The filled rectangles denote the location of the membrane proximal region (MPR) see Table 1 and the text for further details.

serves to separate the TMD from the SNARE-motif, which precedes it. The length of the MPR appears to be important for the function of some SNAREs, at least in vitro (McNew et al. 1999, 2000; Melia et al. 2002) however, whether these observations extend to Golgi-localized SNAREs is presently not known. The SNARE-motif is comprised of a number of heptad-repeats, typically 7–8, which are responsible for the formation of the amphipathic helical bundles characteristic of SNARE complexes. An evolutionarily conserved amino acid residue that occupies a central position in the SNARE-motif, and which contributes to the zero ionic layer of SNARE complexes, is the basis of a SNARE protein family classification scheme (see below). In addition to the so-called SNARE-motif or core domain, SNARE proteins also contain N-terminal extensions (N-terminal domain (NTD)) of varying length and folds (see Figs. 1, 2 and Table 1). Golgi Qa- and Qb-SNAREs contain a domain which adopts a three-helix fold, termed an Habc domain, whereas Golgi R-SNAREs, with the exception of VAMP4, contain a longin fold. Golgi Qc-SNAREs typically contain short N-terminal regions that are predicted to be unstructured, although the NTDs of the Qc-SNAREs Tlg1p and Syntaxin 6, likely adopt an Habc fold. The longin fold, found in Golgi R-SNAREs, is also present in several sub-units of the Golgi-localized vesicle tethering complexes TRAPPI and TRAPPII (Kim et al. 2006) and is predicted to be present in two sub-units of the Golgi vesicle coat complex – coatomer, although the significance of this is not presently understood (Schlenker et al. 2006). In some cases the N-terminal domains of Golgi SNAREs are capable of binding to their respective SNARE-motif, in which case the SNARE is said to adopt a closed or folded-back conformation. Folded-back conformations are known to occur for the R-SNAREs Ykt6p (Tochio et al. 2001) and Sec22p

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Figure 2. The N-terminal domain folds of Golgi SNAREs. (A) Cartoon representation of the crystral structure of the human Vti1b Habc domain (Miller et al. 2007; pdb accession number 2qyw) viewed from the side. The three helices of the domain are labelled from N – to C  a, b and c. (B) The same structure is in (A) but viewed from the N-terminus down the three helix bundle. (C) The NMR-derived solution structure of the longin domain of yeast Ykt6p (Tochio et al. 2001; pdb accession number 1h8m). The cartoons represent 180 rotations of one another.

(Mancias and Goldberg 2007). For, Ykt6p this conformation appears to be important for the protein’s stability and likely plays a key role in the targeting of this protein by regulating the association of the cytoplasmic prenylated form of Ykt6 with membranes (Tochio et al. 2001; Fukasawa et al. 2004;

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Table 1. Yeast and human Golgi-resident SNAREs Type

Human

Yeast homolog

TMD

N-terminal extention

N-terminal fold

Qa

Syntaxin 5

Sed5p

Yes

Yes

Habc

Syntaxin 16

Tlg2p

Yes

Yes

Habc (predicted)

Syntaxin 10

–

Yes

Yes

Habc (predicted)

Syntaxin 11

–

No

Yes

Habc (predicted)

GS27 (membrin, GOS-27)

Bos1p

Yes

Yes

Habc (predicted)

Vti1a (Vti1-rp2)

Vti1p

Yes

Yes

Habc

GS28 (GOS-28)

Gos1p

Yes

Yes

Habc (predicted)

Syntaxin 6

Tlg1p

Yes

Yes

Habc (predicted)

Bet1

Bet1p

Yes

No

Random coil

GS15

Sft1p

Yes

No

Random coil

Sec22b (ERS-24)

Sec22p

Yes

Yes

Longin

Ykt6

Ykt6p

No (prenyl)

Yes

Longin

VAMP4

–

Yes

Yes

Unstructured

SNAP-29 (GS32)

–

No

No

–

Qb

Qc

R

Qb þ Qc

Hasegawa et al. 2004). For Sec22p, a folded-back conformation appears to be a prerequisite for this SNARE’s efficient incorporation into COPII-coated vesicles (Liu et al. 2004; Mancias and Goldberg 2007). The N-terminal domain of the Golgi syntaxins Sed5p (yeast)/Syn5p (mammals) are known to bind to the Sec1–Munc18 (SM) family member protein Sly1 (Yamaguchi et al. 2002; Dulubova et al. 2003; Arac et al. 2005). The association of Sly1p with Sed5p is important for the specificity of Golgi SNARE complex assembly (Peng and Gallwitz 2002) whereas the association of Sly1 with Syntaxin 5 is important for ER–Golgi transport (Williams et al. 2004). A folded-back conformation of Sed5p may be involved in the efficient packaging of this SNARE into COPIIcoated vesicles (Mossessova et al. 2003). Apparently, COPII preferentially binds Sed5p when the protein is part of the Sed5p–Bos1p–Sec22p SNARE

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Figure 3. The topological arrangements of v- and t-SNAREs.

complex because t-SNARE assembly presumably removes the auto-inhibitory contacts of the closed conformation of the protein, exposing its COPII sorting signal (Mossessova et al. 2003).

SNARE protein classification and nomenclature Functionally, SNAREs can be classified as either v-SNAREs or t-SNAREs. v-SNAREs are localized to the transport vesicle, whereas t-SNAREs are predominantly localized to the vesicle’s target compartment. Currently, the generally accepted view is that a single membrane-anchored v-SNARE forms a SNARE-complex in trans with a heterotrimeric t-SNARE. See Fig. 3 and Table 1 for a description of yeast and human Golgi-resident SNARE proteins. SNARE proteins can be further sub-divided based on their amino acid sequence similarities and the position their homologs occupy in SNARE complex macromolecular structures (Fasshauer et al. 1998; Bock et al. 2001). The macromolecular structures of the exocytic and endocytic SNARE complexes revealed that they are parallel four-helical bundles (Sutton et al. 1998; Antonin et al. 2002). In the case of the endocytic SNARE complex, four different SNARE proteins contribute a single helix each to the complex (Fig. 4), this arrangement is also very likely to be the case for Golgi SNARE-complexes. Thus, the syntaxin sub-family has been termed the Qa-SNAREs whereas SNAREs that share the greatest degree of amino acid similarity with the Nterminal SNARE-motif of SNAP-25 (SNAP-25N) are referred to as Qb-SNAREs. Similarly, SNAREs that are most similar to the C-terminal SNARE-motif of SNAP-25 (SNAP-25C) are referred to as Qc-SNAREs. Members of the so-called VAMP family are collectively referred to as R-SNAREs. The Qa-, Qb-, Qc- and R-SNARE nomenclature refers to the presence of a highly evolutionarily conserved amino acid residue at the so-called zero ionic layer of the four-helical bundle – a glutamine for the Q-SNAREs and an arginine for the R-SNAREs (Fig. 4). The general expectation is that members of each family will occupy the equivalent position in their respective SNARE complexes as the corresponding SNARE in the exocytic and endocytic SNARE complexes – adopting a Qa:Qb:Qc:R stoichiometry often referred to as the 3Q:1R rule.

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Figure 4. The SNARE complex is a four-helical bundle. (A) An elongated side view cartoon representation of the macromolecular structure of the endocytic SNARE complex (Zwilling et al. 2007). The position of the zero ionic layer is indicated by the arrow. Syntaxin 6 is represented by the yellow helix, whereas Syntaxin 13, Vti1a and VAMP4 are represented by the blue, magenta and green helices, respectively. (B) An enlarged and skewed side view of the SNARE complex cartoon. The colour scheme used is as in (A). (C) A view down the helical bundle of a cartoon representation of the SNARE complex in which the amino acid residue side-chains defining the zero ionic layer are indicated. The colour scheme used is as in (A). (D) The zero ionic layer residues of the endocytic SNARE complex. Note that while Vti1a is classified as a Q-SNARE, it contributes an aspartic acid, rather than glutamine to the layer. Cartoons where generated using MacPyMOL and the pdb file 2nps.

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Although the 3Q:1R rule is crucial for the formation of properly functioning SNARE complexes, amino acid substitution experiments have shown that it is not necessary, for example, that arginine be contributed by an R-SNARE per se (Katz and Brennwald 2000; Graf et al. 2005). However, exceptions to these general rules exist. For example, Sft1p and Bet1p, two yeast Golgi resident Qc-SNAREs, contain an aspartic acid and a serine (respectively) at the zero layer position, however the biological significance of this variability is presently unknown.

The general mode of SNARE protein function It is now generally accepted that the predominate function of SNARE proteins is to act as facilitators of intra-cellular membrane fusion events within the endomembrane system, through the formation of complexes between SNAREs on vesicles and SNAREs on organellar membranes. This association of SNAREs in trans is thought to be important in bringing the vesicle and organellar membranes close enough together to facilitate membrane fusion. In general, the SNARE complex that forms conforms to the 3Q:1R rule (Katz and Brennwald 2000). How are the individual SNARE proteins contributed to the SNARE complex? In vitro fusion assays with yeast Golgi SNAREs revealed that the heterotrimeric t-SNARE is comprised of a heavy chain – a Qa-SNARE (a syntaxin such as Sed5p or Syn5) and two different SNAREs which comprise the two t-SNARE light chains. Thus the t-SNARE consists of one Qa-SNARE together with either a Qb þ Qc, Qb þ R or Qc þ R pair defining the t-SNARE light chains. Employing this scheme the v-SNARE would be contributed by the remaining SNARE, i.e., either a Qb-, Qc- or R-SNARE, depending on the composition of the t-SNARE complex (Fig. 2). The v-SNARE is often an RSNARE, but this may not be so for SNAREs in the Golgi, as liposome fusion assays have established that the Qc-SNAREs, Bet1p and Sft1p, function as vSNAREs in this context (McNew et al. 2000; Parlati et al. 2002). However, an in vitro transport employing yeast Golgi SNAREs revealed that, in addition to Bet1p, the Qb-SNARE Bos1p and the R-SNARE Sec22p may also function as vSNAREs in transport between the ER and Golgi (Spang and Schekman 1998). Although the composition of the t-SNARE complex and it’s respective vSNARE appears to be quite rigid in vitro (Parlati et al. 2000, 2002) it seems likely that greater compositional flexibility exists in cells (Tsui and Banfield 2000; Tsui et al. 2001; Banfield 2001). In yeast, for example, some Golgi SNAREs interact with Qa-SNAREs other than Sed5p and in so doing participate in multiple transport steps (e.g. Vti1p, Ykt6p and Tlg1p). In addition, several yeast Golgi SNAREs are not essential for yeast cell growth. Given the importance of SNAREs in membrane fusion these observations have been viewed as being consistent with a functionally redundant role of SNAREs and reflexing a lack of selectivity in the composition of SNARE complexes. Despite the apparent flexibility in SNARE pairing interactions in cells, adherence to

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Figure 5. The Golgi SNARE cycle.

the 3Q:1R rule remains important. This requirement has been successfully exploited as a means to identify functionally interacting SNARE complexes (Graf et al. 2005). While the importance of residues in the immediate vicinity of the zero ionic layer have been documented (Stone et al. 1997; Graf et al. 2005) a comprehensive examination of the relative importance of other regions of the SNARE-motif in Golgi SNARE function is lacking. What directs the specificity of SNARE complex formation? Prior to SNARE complex formation, the v-SNARE and t-SNAREs encounter each other in cis (Fig. 5) and SNARE complex formation appears to proceed from the N- to Cterminus (Sorensen et al. 2006; Pobbati et al. 2006). The close opposition of the v- and t-SNAREs is mediated by a variety of factors, so-called tethering factors, which presumably function to ensure that only the correct SNAREs form biologically meaningful trans-complexes. The formation of cognate, fusogenic SNARE complexes between opposing membranes drives fusion. Although cartoons, such as the one depicted in Fig. 5, often show the formation of trans-SNARE complexes comprised of 1–2 complexes (for the sake of simplicity) the average number of complexes participating in one fusion reaction, based on studies on the neuronal exocytic SNARE complex, is likely to be on the order of 3–8 (Han et al. 2004; Rickman et al. 2005; Montecucco et al. 2005). The rosette-like structures that are observed to form from the association of multiple SNARE complexes may be important for mediating membrane fusion, perhaps via the transmembrane domain of SNAREs (Han et al. 2004). Following fusion of the vesicle with the Golgi, SNAREs remain bound to one another in trans. Trans-SNARE complexes are dissociated through the combined actions of a-SNAP/Sec17p and NSF/Sec18p after which, SNAREs are free to be recycled and reused in another round of vesicular transport and membrane fusion. Several Golgi SNAREs have been shown to cycle between

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the Golgi and the ER (Ballensiefen et al. 1998; Wooding and Pelham 1998; Ossipov et al. 1999; Cosson et al. 2004), reflecting the requirement for SNAREs in anterograde as well as retrograde vesicle-mediated transport (Spang and Schekman 1998). In addition, apart from the requirement that cells reuse SNARE proteins in successive rounds of transport, it seems likely that this recycling process is intimately linked to the establishment and dynamic nature of the Golgi apparatus itself (Cosson et al. 2004).

The specificity of SNARE complex formation Table 2 lists SNARE complexes known to function in transport to the Golgi in yeast and mammalian cells. The complexes that mediate such traffic in mammalian cells have predominantly been identified by co-immune precipitation experiments. In contrast, in budding yeast this information has been obtained from a variety of approaches, including co-immune precipitation, genetic studies and in vitro mixing and fusion assays. An observation that has

Table 2. SNARE complexes known to function in transport to the Golgi Mammals

Yeast

Complex

Transport step (s)

Complex

Transport step (s)

Syntaxin 5 (Qa) GS28 (Qb) GS15 (Qc) Ykt6 (R)

Recyling endosome–TGN

Sed5p Gos1p Sft1p Ykt6p

Intra-Golgi

Syntaxin 5 GS28 Bet1 Ykt6

ERGIC – Golgi

Sed5p Bos1p Bet1p Sec22p

ER–Golgi

Syntaxin 5 GS27 Bet1 Sec22p

ER–ERGIC

Sed5p Bos1p Bet1p Ykt6pa

ER–Golgi

Syntaxin 16 Vti1a Syn6 VAMP4

Early endosome–TGN

Sed5p Gos1p Bet1pa Ykt6p

Intra-Golgi

Syntaxin 16 Vti1a Syntaxin 10 VAMP3

Late endosome–TGN

a

Assumed on the basis of over-expression experiments in sec22D (Liu and Barlowe 2004) and sft1D cells (Tsui et al. 2001). SNAREs in bold, italicized font are encoded by non-essential genes. ERGIC (ER-Golgi intermediate compartment).

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dogged the role of SNAREs in the specificity of membrane fusion events has been the apparent lack of specificity among many SNARE–SNARE associations. This lack of specificity is particularly apparent in in vitro mixing experiments using bacterially expressed mammalian as well as yeast SNARE proteins (Yang et al. 1999; Fasshauser et al. 1999; Tsui and Banfield 2000). Recent evidence suggests that non-cognate SNARE complexes form in cells, but that cells have some, as yet to be identified mechanism, which selects only the physiologically relevant complexes for use in membrane fusion reactions (Bethani et al. 2007). Thus, the identification of SNARE–SNARE interactions by co-immune precipitation may not be sufficient to assign particular SNAREs to a functional complex (Bethani et al. 2007). In in vitro fusion assays using the theoretical maximum tetrameirc combinations of SNAREs encoded by the yeast genome, only 9/275 were found to be fusogenic (McNew et al. 2000; Parlati et al. 2000, 2002; Paumet et al. 2004). Two of the nine fusogenic complexes contained the Golgi Qa-SNARE Sed5p: Sed5p/Bos1p/Sec22p (t-SNARE) þ Bet1p (v-SNARE) and Sed5p/Gos1p/Ykt6p (t-SNARE) þ Sft1p (v-SNARE), complexes which mediate fusion of vesicles with the cis- and trans-Golgi, respectively. These two complexes correspond to the mammalian Syntaxin 5-containing complexes: Syntaxin 5/membrin/ Sec22b (t-SNARE) þ Bet1 (v-SNARE) (Hay et al. 1998), although some studies suggest that the v-SNARE may be Sec22b (Xu et al. 2000; Joglekar et al. 2003) and to Syntaxin 5/GS28/Ykt6 (t-SNARE) þ GS15 (v-SNARE) (Xu et al. 2002). These in vitro fusion assay data suggest that SNARE proteins encode the necessary information to direct the formation of fusogenic SNARE complexes. In yeast, Sed5p is the only syntaxin required for transport through the Golgi, however, Sec22p and Gos1p are encoded by non-essential genes. Thus in cells lacking either the SEC22 or GOS1 genes (presumably) only a single Sed5p-containing SNARE complex would remain. Additional Sed5pcontaining fusogenic SNARE complexes have been proposed to form in cells on the basis of biochemical as well as genetic studies (Tsui and Banfield 2000; Liu and Barlowe 2002) see Table 2. In cells, Ykt6p appears to be able to substitute for Sec22p (Liu and Barlowe 2002). Similarly, under conditions when the Qc-SNARE Bet1p is ectopically over-expressed, cells can survive without the Qc-SNARE Sft1p (Tsui and Banfield 2000). Thus, with the exception of the QaSNARE, yeast Golgi Qb-, Qc- and R-SNAREs display varying degrees of presumptive functional redundancy. Whether these additional complexes constitute functionally overlapping SNARE complexes, redundant complexes or complexes that form as a result of the absence of the cognate SNARE, requires further investigation. The observation that some Sed5p interacting SNAREs also form complexes with other Qa-SNAREs functioning on other organelles suggests that a single SNARE is likely to be insufficient to direct complex specificity. For example, Vti1p (Lupashin et al. 1997; Von Mollard et al. 1997) and Ykt6p (Kweon et al. 2003) bind to multiple Qa-SNAREs and function in multiple transport pathways. Combinatorial binding interactions may therefore influence the specificity of

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SNARE complex formation (Banfield 2001) in vivo. Finally, the extent to which regions of the SNARE-motif, other than the zero ionic layer, contribute to the specificity of Golgi SNARE complex assembly is an important issue which remains to be addressed.

i-SNAREs In vitro mixing studies with the soluble forms of Sed5p and its Golgi SNARE binding partners revealed far more ternary complexes than were identified on the basis of SNARE-mediated liposome fusion assays (Tsui et al. 2001; McNew et al. 2000; Parlati et al. 2002). The presence of more than two fusion competent Sed5p-containing SNARE complexes would help to reconcile conceptual problems arising from the fact that one SNARE from each of these complexes is non-essential – Sec22p and Gos1p, respectively (see Table 2) (McNew et al. 2000; Parlati et al. 2002). However, another explanation has been proposed to account for the additional Sed5p-containing SNARE complexes observed in in vitro mixing studies with soluble SNAREs. Using their well established SNARE-mediated liposome fusion assay Varlamov et al. (2004) established that certain sub-units of the cis-Golgi SNARE complex could inhibit fusion mediated by the trans-Golgi SNARE complex and vice versa (Varlamov et al. 2004) – the authors termed these SNAREs as i-SNAREs. While the opposing distribution of cis- and trans-Golgi SNAREs (Volchuk et al. 2004) could in principle account for the distribution of the fusogenic SNARE complexes, the authors argue that i-SNAREs would enhance this phenomena – essentially fine-tuning the specificity of membrane fusion events in the Golgi. While this is a particularly attractive notion, the concept of i-SNAREs functioning in the Golgi still awaits in vivo validation.

Localization of SNAREs to the Golgi In general SNAREs are predominantly localized to the vesicles and compartments on which they function and are absent from those on which they do not. How are SNAREs localized to the Golgi? Accumulating evidence suggests that both the transmembrane domain of SNAREs as well as signals in their cytoplasmic domains accounts for their steady-state distributions. A requirement of the transmembrane domain in the localization of Golgi enzymes is well established in mammalian cells. Such studies have led to the proposals that (1) the length of the transmembrane is an important factor in Golgi localization and that sorting/or localization is the result of Golgi membrane bilayer thickness (Bretcher and Munro 1993) or (2) that the TMDs of Golgi residents oligomerize and are prevented from exiting the Golgi (Nilsson et al. 1993). The transmembrane domains of the yeast SNAREs Sed5p and Sft1p contribute to their Golgi localization (Banfield et al. 1994; Rayner and Pelham 1997) and TMD length has been shown to be important for the Golgi localization of Syn5 in mammalian cells (Watson and Pessin 2001). However

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in yeast, TMD length alone is not sufficient to ensure exclusive Golgi localization of SNARE protein chimeras (Rayner and Pelham 1997). The steady-state distribution of SNAREs proteins in the Golgi appears be dynamic – relying on the active retrieval of SNAREs from distal cisternae and their retrieval to earlier sub-compartments or to the ER, from where they return to Golgi. In yeast, Sed5p (Wooding and Pelham 1998), Sec22p (Ballensiefen et al. 1998) and Bos1p (Ossipov et al. 1999) have been shown to cycle between the Golgi and the ER. Although yeast Bet1p does not undergo such cycling (Ossipov et al. 1999), in mammalian cells Bet1 and Sec22b have been shown to continually cycle between the Golgi and the ER (Hay et al. 1998). The recycling of Bet1 does not appear to require interactions with its cognate SNAREs (Joglekar et al. 2003). In the case of yeast Sec22p and Bos1p, the retrieval of these SNAREs from the Golgi requires a functional COPI coat (Ballensiefen et al. 1998; Ossipov et al. 1999). An analysis of the lateral distribution and vesicle incorporation of SNAREs in the mammalian Golgi using electron microscopy is also consistent with a dynamic localization mechanism (Cosson et al. 2005). The Golgi SNARE Ykt6 does not contain a TMD, but rather is dually lipid modified at its C-terminus (Fukasawa et al. 2004). The farnesylated form of Ykt6 resides in the cytoplasm whereas farnesylated, palmitoylated Ykt6 is found predominantly on Golgi membranes in non-neuronal mammalian cells (Fukasawa et al. 2004; Hasegawa et al. 2004). How Ykt6 is targeted to Golgi membranes remains to be determined.

SNAREs and COPI interactions Several SNAREs have been shown cycle within or from the Golgi in a COPIdependent manner, data that implies an interaction between these SNAREs and the coat protein complex. The COPI coat is comprised of the heptameric complex termed coatomer, together with the GTPase Arf1. Arf1 cycles on and off Golgi membranes as a function of its nucleotide-bound state. GTP-bound Arf localizes to membranes whereas GDP-bound Arf is found in the cytoplasm. The nucleotide status on Arf1 is controlled through the action of its exchange factor (Arf GEF) and its activating protein (Arf GAP). In vitro studies using yeast Golgi SNAREs revealed that the Arf1p GAPs, Glo3p and Gsc1p, act catalytically on Golgi SNAREs promoting a conformational change that facilitates stoichiometric recruitment of Arf1p to SNAREs (Rein et al. 2002). In agreement with these findings, studies in mammalian cells have identified a motif on Arf that is required for the recruitment of Arf to Golgi membranes by the Qb-SNARE membrin (Honda et al. 2005). Schindler and Spang (2007) have shown that Gcs1p accelerates the formation of SNARE complexes in vitro and suggested that Arf GAPs may function as folding chaperones for SNAREs. Such mechanisms may function to couple SNARE recruitment to vesicle formation in cells, thus ensuring that each vesicle carries sufficient SNAREs capable of forming cognate SNARE complexes at its target compartment.

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The role of SNAREs in the morphological and functional organization of the Golgi Presumably recycling Golgi SNAREs is important for ensuring efficient vesiclemediated transport, as this would allow these proteins to be employed in successive rounds of trafficking. Mathematical modeling using a minimal system, in which the variables were restricted to cytoplasmic coat protein complexes and SNAREs, was sufficient to generate stable non-identical compartments (Heinrich and Rapoport 2005). A requirement of Heinrich and Rapoport’s (2005) model was that each vesicle generating coat complex preferentially bound and packaged a characteristic set of SNAREs. The lateral distribution of Golgi SNAREs observed by Cosson et al. (2005) may similarly reflect differential affinity of Golgi SNAREs for the COPI coat in vivo. Thus, the affinity of vesicle coats, or their cargo sorting affiliated partners, may function to promote and maintain the compositional integrity of Golgi cisternia through their intrinsic ability to bind different SNAREs with varying affinities.

Regulators of Golgi SNARE function The activity of SNAREs is regulated at various stages of their action including the assembly post-translational of the t-SNARE and the assembly of the trans-SNARE complex (Fig. 5). A variety of proteins have been identified that modulate the activity of SNAREs. In addition, post-translational modifications such prenylation, palmitoylation and phosphorylation also influence the activity and or localization of SNAREs.

NSF/Sec18p and a-SNAP/Sec17p NSF and a-SNAP represent two co-operating core regulators of SNARE protein activity. These proteins are responsible for the disassembly of cis-SNARE complexes (Fig. 5), an activity that frees-up SNAREs to be used in successive rounds of vesicle fusion. Three molecules of a-SNAP link the cis-SNARE complex with a hexamer of the ATPase NSF/Sec18p and together this complex is referred to as the 20 S complex (Hohl et al. 1998; Wimmer et al. 2001; Furst et al. 2003; Brunger and DeLaBarre 2003). NSF contains two ATPase domains termed D1 and D2. The D2 ATPase domain mediates the formation of the NSF hexamer whereas the D1 ATPase domain effects the dissociation of the cisSNARE complex. The association of NSF with a-SNAP into the 20 S complex stimulates the ATPase activity of NSF (Marz et al. 2003).

The Sec1/Munc-18 like (SM) proteins Sec1/Munc-18 (SM) proteins bind directly to SNAREs and act downstream of vesicle tethering events. Sly1p, the yeast SM protein which binds to Sed5p, was identified because a mutant of this protein (sly1-20p) could suppress the loss of the essential Rab/Ypt GTPase Ypt1p (Dascher et al. 1991). The association of Sly1p with Sed5p has been shown to enhance Sed5p-containing transSNARE complexes (Kosodo et al. 2002), and to be important for the specificity

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of Golgi SNARE complex assembly (Peng and Gallwitz 2002) whereas the assocation of Sly1 with Syntaxin 5 has been demonstrated to be important for ER–Golgi transport (Williams et al. 2004). However, the interaction between Sed5p and Sly1p is dispensible for transport (Peng and Gallwitz 2004). Unlike the exocytic and neuronal syntaxins and their requisite SM protein interactions, the interaction between Sly1p and Sed5p is mediated by a short stretch of amino acids at the N-terminus of the protein which does not involve either the Habc or SNARE-motif domains (Yamaguchi et al. 2002; Bracher and Weissenhorn 2002; Dulubova et al. 2003) and thus this association does not promote a folded-back conformation for Sed5p. A similar mode of binding is also evident between the SM protein Vps45/Vps45p and the Qa-SNARE Syntaxin 16/Tlg2p (Dulubova et al. 2002). It is now apparent that Sly1p is capable of binding to non-syntaxin SNAREs as well as to SNARE complexes and that this SNARE binding property of Sly1p is important in the specificity of cognate SNARE complex formation (Peng and Gallwitz 2002, 2004; Li et al. 2005).

The Golgins The Golgins are a class of large coiled-coil containing Golgi localized proteins with roles in tethering vesicles to the Golgi. Some golgins contain a single Cterminal transmembrane domain whereas other members of the family are peripherally associated with Golgi membranes. The peripheral membrane protein p115 has been shown to bind directly to SNAREs involved in ERGolgi intermediate compartment (ERGIC) as well as ERGIC-Golgi transport (Allan et al. 2000; Shorter et al. 2002). A SNARE-related coiled-coil region of p115 interacts with many Golgi SNAREs and such interactions likely promote the formation of trans-SNARE complexes (Shorter et al. 2002). The functional consequences of such interactions may be to ensure a direct connection between the tethering machinery and SNAREs as well as to facilitate the recruitment of p115 to membrane sites where unassembled SNAREs are located (Brandon et al. 2006; Bentley et al. 2006). Mutational analysis of p115 suggests that the SNARE-modulating activity of the protein is more important than its tethering activity in maintaining the structure and function of the Golgi (Puthenveedu and Lindstedt 2004). Uso1p is the yeast homologue of p115 (Sapperstein et al. 1995, 1996; Cao et al. 1998). Other Golgins have also been shown to bind to Golgi SNAREs, including GM130 which interacts directly with Syntaxin 5 (Diao et al. 2007) and GCC185, which binds directly to Syntaxin 16 (Ganley et al. 2008). The emerging picture of the role of Golgins is one in which these proteins sequester Rab GTPases and Qa-SNAREs/syntaxins, and in so doing keep these two key proteins in close proximity to the tether.

Conserved oligomeric Golgi (COG) The COG complex is a member of the oligomeric vesicle tethering factor family which comprise a structurally diverse group of peripheral membrane

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protein complexes involved in vesicle-organellar tethering events prior to SNARE complex assembly (Oka and Krieger 2005; Stzul and Lupashin 2006). The COG complex is involved in retrograde trafficking of Golgi-resident proteins and extensive genetic interactions have been documented between the COG complex and SNAREs. In addition the yeast and mammalian COG complexes co-immune precipitate with Golgi SNAREs (Suvorova et al. 2002; Zolov and Lupashin 2005) and the localization and stability of Golgi SNAREs is altered in cells with defective COG complex components (Oka et al. 2004; Fotso et al. 2005; Zolov and Luphasin 2005; Shestakova et al. 2007). More recently, the yeast COG complex has been shown to interact with the SNAREmotif of Sed5p and to preferentially bind to Sed5p-containing SNARE complexes, leading the authors to propose that one function of the COG complex is to stabilize intra-Golgi SNARE complexes (Shestakova et al. 2007).

GATE-16 GATE-16, a member of the ubiquitin-fold (UF) protein family, is localized to the Golgi and interacts with NSF as well as GS28 (Sagiv et al. 2000). NSF/a-SNAP facilitates the interaction of GATE-16 with GS28 in a manner that requires ATP-binding but not ATP hydrolysis. Interestingly, GATE-16 binding prevents GS28 from interacting with Syntaxin 5 and in so doing prevents the formation of a functional t-SNARE (Muller et al. 2002). In addition, the yeast GATE-16 homologue, Aut7p, interacts with Bet1p, a Qc-SNARE involved in ER-to-Golgi transport and shows genetic interactions with BET1 and the ER-Golgi RSNARE SEC22 (Legesse-Miller et al. 2000).

FIG FIG (also known as CAL, PIST and GOPC) localizes to the TGN where it interacts with the Qc-SNARE Syntaxin 6 (Charest et al. 2001). FIG contains two coiledcoil regions and a single PDZ domain and the protein’s interaction with Syntaxin 6 is mediated via the second coiled-coil region and its C-terminal flanking region. Although the biological significance of this interaction remains to be determined, knock-out of the FIG gene in mice results in selective ablation of acrosome formation during spermatogenesis (Yao et al. 2002). The acrosome is believed to form from the Golgi apparatus and the absence of FIG leads to fragmented acrosomal vesicles suggestive of a role for FIG in the fusion of vesicles into the acrosome. Curiously, FIG also interacts with Golgin-160 (Hicks and Machamer 2005).

Phosphorylation Many SNAREs and their regulatory proteins are known to be phosphorylated by a variety of kinases (Gerst 2003; Snyder et al. 2006). The yeast Golgi Qa-SNARE Sed5p is a phosphoprotein and Weinberger et al. (2005) have shown that amino acid substitutions to an evolutionarily conserved protein kinase A phosphorylation site adjacent to the transmembrane domain of the protein has dramatic effects on Golgi morphology. While expression of the

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pseudophosphorylated form of Sed5p (Ser317Asp) results in the accumulation of ER membranes and vesicles, expression of the non-phosphorylatable form of the protein (Ser317Ala) results in the accumulation of Golgi membranes reminiscent of the mammalian cell Golgi (Weinberger et al. 2005). The Ser317Ala mutant also shows an increased affinity for the COPI coat, suggesting that phoshorylation status of Sed5p in cells may play a role in regulating Golgi morphology.

Palmitoylation Several SNAREs are known to be palmitoylated. Some of these SNAREs lack transmembrane domains and are anchored to the membrane by their palmitate moieties, such is the case for SNAP-25, SNAP-23 and Syntaxin 11 (Vogel and Roche 1999; Veit 2000; Prekeris et al. 2000). In contrast, the Golgi SNARE Ykt6p is anchored by a combination of prenylation and palmitoylation (Fukasawa et al. 2004). In addition, it is now apparent that several SNAREs bearing TMDs are also palmitoylated (Valdez-Taubas and Pelham 2005) including the TGN/ endosomal Qc-SNARE, Tlg1p. The yeast DHCC-CDR family member Swf1p is required for palmitoylation of Tlg1p and prevention of Tlg1p palmitoylation results in its ubiquitination and transportation, via the multivesicular body, to the vacuole for degradation. While palymitoyation of TMD-anchored SNAREs does not appear to be essential for their function, this modification may play a role in the membrane partitioning of these SNAREs and or in dissociation of these modified proteins from other SNAREs, following fusion (Valdez-Taubas and Pelham 2005). Based on amino acid sequence similarities with Tlg1p, the mammalian Golgi resident SNAREs Syntaxin 6, Syntaxin 10 and VAMP4 may also be substrates for palymitoylation (Valdez-Taubas and Pelham 2005). Unlike the TMD-anchored SNAREs, which are modified via DHHC-CDR palmitoyltransferases, Ykt6 appears to be capable of mediating its own palymitoylation (Veit 2004) via its longin fold (Dietrich et al. 2004). Ykt6 is found in two pools in cells–a cytoplasmic pool and a membrane associated pool. Ykt6 lacks a proteinaceous membrane anchor but contains a prenylation consensus sequence (a so-called CAAX box) at its C-terminus. The cytoplasmic pool of Ykt6 has been shown to farnesylated and the farnesylation of Ykt6 is prerequiste for the subsequent palymitoylation and membrane association of the protein (Fukasawa et al. 2004). Fukasawa et al. (2004) propose a cycle of membrane association of Ykt6 in which the farnesylated fold-back conformation of Ykt6p (mediated by an interaction between the longin domain and SNARE-motif, Tochio et al. 2001) is targeted to membranes, whereupon the protein is palymitoylated. This dual lipid modification may be required for stable membrane association of Ykt6 (Fukasawa et al. 2004).

Golgi SNAREs and apoptosis During programmed cells death (apoptosis) the Golgi loses its cisternal organization and is fragmented into clusters of tubulovesicular elements

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(Lane et al. 2001) and the early secretory pathway is blocked (Lane et al. 2002). While this phenotype is associated with the proteolytic cleavage of members of the Golgin family (Mancini et al. 2000; Chiu et al. 2002; Lane et al. 2002) Lowe et al. (2004) have shown that Syntaxin 5 is cleaved by caspase during apoptosis. Caspase-3 cleaves Syntaxin 5 at Asp 263, separating the SNARE-motif from the N-terminal Habc domain. Syntaxin 5 participates in several SNARE complexes in the Golgi, including partnerships with Bet1 (Qc-), membrin/GS27 (Qb-) and Sec22b (R-); Bet1 (Qc-), GS28 (Qb-) and Ykt6 (R-) as well as with GS15 (Qc-), GS28 (Qb-) and Ykt6 (R-) (Hay et al. 1998; Zhang et al. 2001; Xu et al. 2002). Thus, cleavage of Syntaxin 5 by caspase-3 is likely to affect several trafficking steps to and within the Golgi.

Future perspectives While much has been learned about the role of SNAREs in the Golgi many important questions remain to be addressed. These include identification of the sorting signals/motifs on SNAREs as well as the macromolecular details governing interactions between SNAREs and the COPI vesicle generating machinery. Establishing, whether like SNAREs and the ER vesicle coat COPII (Morsomme et al. 2003), Golgi SNAREs influence the incorporation of particular cargo proteins into COPI-coated vesicles. Finally, a more detailed understanding of the mechanisms governing the steady-state localization of SNAREs to the Golgi will make important contributions to our understanding of how Golgi SNARE trafficking impacts the morphological and functional organization of this fascinating organelle.

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Bethani I, Lang T, Geumann U, Sieber JJ, Jahn R, Rizzoli SO (2007) The specificity of SNARE pairing in biological membranes is mediated by both proof-reading and spatial segregation. EMBO J 26: 3981–3992 Bock JB, Matern HT, Peden AA, Scheller RH (2001) A genomic perspective on membrane compartment organization. Nature 409: 839–841 Bracher A, Weissenhorn W (2002) Structural basis for the Golgi membrane recruitment of Sly1p by Sed5p. EMBO J 21: 6114–6124 Brandon E, Szul T, Alvarez C, Grabski R, Benjamin R, Kawai R, Sztul E (2006) On and off membrane dynamics of the endoplasmic reticulum-Golgi tethering factor p115 in vivo. Mol Biol Cell 17: 2996–3008 Bretscher MS, Munro S (1993) Cholesterol and the Golgi apparatus. Science 261: 1280–1281 Brunger AT, DeLaBarre B (2003) NSF and p97/VCP: similar at first, different at last. FEBS Lett 555: 126–133 Cao X, Ballew N, Barlowe C (1998) Initial docking of ER-derived vesicles requires Uso1p and Ypt1p but is independent of SNARE proteins. EMBO J 17: 2156–2165 Charest A, Lane K, McMahon K, Housman DE (2001) Association of a novel PDZ domaincontaining peripheral Golgi protein with the Q-SNARE (Q-soluble N-ethylmaleimide-sensitive fusion protein (NSF) attachment protein receptor) protein syntaxin 6. J Biol Chem 276: 29456–29465 Chiu R, Novikov L, Mukherjee S, Shields D (2002) A caspase cleavage fragment of p115 induces fragmentation of the Golgi apparatus and apoptosis. J Cell Biol 159: 637–648 Cosson P, Ravazzola M, Varlamov O, Sollner TH, Di Liberto M, Volchuk A, Rothman JE, Orci L (2005) Dynamic transport of SNARE proteins in the Golgi apparatus. Proc Natl Acad Sci USA 102: 14647–14652 Diao A, Frost L, Morohashi Y, Lowe M (2007) Coordination of golgin tethering and SNARE assembly: GM130 binds syntaxin 5 in a p115-regulated manner. J Biol Chem 283: 6957–6967 Dietrich LE, Gurezka R, Veit M, Ungermann C (2004) The SNARE Ykt6 mediates protein palmitoylation during an early stage of homotypic vacuole fusion. EMBO J 23: 45–53 Dulubova I, Yamaguchi T, Arac D, Li H, Huryeva I, Min SW, Rizo J, Sudhof TC (2003) Convergence and divergence in the mechanism of SNARE binding by Sec1/Munc18like proteins. Proc Natl Acad Sci USA 100: 32–37 Dulubova I, Yamaguchi T, Gao Y, Min SW, Huryeva I, Sudhof TC, Rizo J (2002) How Tlg2p/ syntaxin 16 ‘snares’ Vps45. EMBO J 21: 3620–3631 Fasshauer D, Antonin W, Margittai M, Pabst S, Jahn R (1999) Mixed and non-cognate SNARE complexes. Characterization of assembly and biophysical properties. J Biol Chem 274: 15440–15446 Fasshauer D, Sutton RB, Brunger AT, Jahn R (1998) Conserved structural features of the synaptic fusion complex: SNARE proteins reclassified as Q- and R-SNAREs. Proc Natl Acad Sci USA 95: 15781–15786 Fotso P, Koryakina Y, Pavliv O, Tsiomenko AB, Lupashin VV (2005) Cog1p plays a central role in the organization of the yeast conserved oligomeric Golgi complex. J Biol Chem 280: 27613–27623 Fukasawa M, Varlamov O, Eng WS, Sollner TH, Rothman JE (2004) Localization and activity of the SNARE Ykt6 determined by its regulatory domain and palmitoylation. Proc Natl Acad Sci USA 101: 4815–4820 Furst J, Sutton RB, Chen J, Brunger AT, Grigorieff N (2003) Electron cryomicroscopy structure of N-ethyl maleimide sensitive factor at 11 A resolution. EMBO J 22: 4365–4374 Ganley IG, Espinosa E, Pfeffer SR (2008) A syntaxin 10-SNARE complex distinguishes two distinct transport routes from endosomes to the trans-Golgi in human cells. J Cell Biol 180: 159–172

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Peng R, Gallwitz D (2002) Sly1 protein bound to Golgi syntaxin Sed5p allows assembly and contributes to specificity of SNARE fusion complexes. J Cell Biol 157: 645–655 Pobbati AV, Stein A, Fasshauer D (2006) N- to C-terminal SNARE complex assembly promotes rapid membrane fusion. Science 313: 673–676 Prekeris R, Klumperman J, Scheller RH (2000) Syntaxin 11 is an atypical SNARE abundant in the immune system. Eur J Cell Biol 79: 771–780 Puthenveedu MA, Linstedt AD (2004) Gene replacement reveals that p115/SNARE interactions are essential for Golgi biogenesis. Proc Natl Acad Sci USA 101: 1253–1256 Rayner JC, Pelham HR (1997) Transmembrane domain-dependent sorting of proteins to the ER and plasma membrane in yeast. EMBO J 16: 1832–1841 Rein U, Andag U, Duden R, Schmitt HD, Spang A (2002) ARF-GAP-mediated interaction between the ER-Golgi v-SNAREs and the COPI coat. J Cell Biol 157: 395–404 Rickman C, Hu K, Carroll J, Davletov B (2005) Self-assembly of SNARE fusion proteins into star-shaped oligomers. Biochem J 388: 75–79 Sagiv Y, Legesse-Miller A, Porat A, Elazar Z (2000) GATE-16, a membrane transport modulator, interacts with NSF and the Golgi v-SNARE GOS-28. EMBO J 19: 1494–1504 Sapperstein SK, Lupashin VV, Schmitt HD, Waters MG (1996) Assembly of the ER to Golgi SNARE complex requires Uso1p. J Cell Biol 132: 755–767 Sapperstein SK, Walter DM, Grosvenor AR, Heuser JE, Waters MG (1995) p115 is a general vesicular transport factor related to the yeast endoplasmic reticulum to Golgi transport factor Uso1p. Proc Natl Acad Sci USA 92: 522–526 Schindler C, Spang A (2007) Interaction of SNAREs with ArfGAPs precedes recruitment of Sec18p/NSF. Mol Biol Cell 18: 2852–2863 Schlenker O, Hendricks A, Sinning I, Wild K (2006) The structure of the mammalian signal recognition particle (SRP) receptor as prototype for the interaction of small GTPases with Longin domains. J Biol Chem 281: 8898–8906 Shestakova A, Suvorova E, Pavliv O, Khaidakova G, Lupashin V (2007) Interaction of the conserved oligomeric Golgi complex with t-SNARE Syntaxin5a/Sed5 enhances intraGolgi SNARE complex stability. J Cell Biol 179: 1179–1192 Shorter J, Beard MB, Seemann J, Dirac-Svejstrup AB, Warren G (2002) Sequential tethering of Golgins and catalysis of SNAREpin assembly by the vesicle-tethering protein p115. J Cell Biol 157: 45–62 Siddiqi SA, Siddiqi S, Mahan J, Peggs K, Gorelick FS, Mansbach CM II (2006) The identification of a novel endoplasmic reticulum to Golgi SNARE complex used by the prechylomicron transport vesicle. J Biol Chem 281: 20974–20982 Snyder DA, Kelly ML, Woodbury DJ (2006) SNARE complex regulation by phosphorylation. Cell Biochem Biophys 45: 111–123 Sorensen JB, Wiederhold K, Muller EM, Milosevic I, Nagy G, De Groot BL, Grubmuller H, Fasshauer D (2006) Sequential N- to C-terminal SNARE complex assembly drives priming and fusion of secretory vesicles. EMBO J 25: 955–966 Spang A, Schekman R (1998) Reconstitution of retrograde transport from the Golgi to the ER in vitro. J Cell Biol 143: 589–599 Stone S, Sacher M, Mao Y, Carr C, Lyons P, Quinn AM, Ferro-Novick S (1997) Bet1p activates the v-SNARE Bos1p. Mol Biol Cell 8: 1175–1181 Sutton RB, Fasshauer D, Jahn R, Brunger AT (1998) Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature 395: 347–353 Suvorova ES, Duden R, Lupashin VV (2002) The Sec34/Sec35p complex, a Ypt1p effector required for retrograde intra-Golgi trafficking, interacts with Golgi SNAREs and COPI vesicle coat proteins. J Cell Biol 157: 631–643 Sztul E, Lupashin V (2006) Role of tethering factors in secretory membrane traffic. Am J Physiol Cell Physiol 290, C11–C26

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Tochio H, Tsui MM, Banfield DK, Zhang M (2001) An autoinhibitory mechanism for nonsyntaxin SNARE proteins revealed by the structure of Ykt6p. Science 293: 698–702 Tsui MM, Banfield DK (2000) Yeast Golgi SNARE interactions are promiscuous. J Cell Sci 113(Pt 1): 145–152 Tsui MM, Tai WC, Banfield DK (2001) Selective formation of Sed5p-containing SNARE complexes is mediated by combinatorial binding interactions. Mol Biol Cell 12: 521–538 Valdez-Taubas J, Pelham H (2005) Swf1-dependent palmitoylation of the SNARE Tlg1 prevents its ubiquitination and degradation. EMBO J 24: 2524–2532 Varlamov O, Volchuk A, Rahimian V, Doege CA, Paumet F, Eng WS, Arango N, Parlati F, Ravazzola M, Orci L, Sollner TH, Rothman JE (2004) i-SNAREs: inhibitory SNAREs that fine-tune the specificity of membrane fusion. J Cell Biol 164: 79–88 Veit M (2004) The human SNARE protein Ykt6 mediates its own palmitoylation at C-terminal cysteine residues. Biochem J 384: 233–237 Veit M (2000) Palmitoylation of the 25-kDa synaptosomal protein (SNAP-25) in vitro occurs in the absence of an enzyme, but is stimulated by binding to syntaxin. Biochem J 345(Pt 1): 145–151 Vogel K, Roche PA (1999) SNAP-23 and SNAP-25 are palmitoylated in vivo. Biochem Biophys Res Commun 258: 407–410 Volchuk A, Ravazzola M, Perrelet A, Eng WS, Di Liberto M, Varlamov O, Fukasawa M, Engel T, Sollner TH, Rothman JE, Orci L (2004) Countercurrent distribution of two distinct SNARE complexes mediating transport within the Golgi stack. Mol Biol Cell 15: 1506–1518 Von Mollard GF, Nothwehr SF, Stevens TH (1997) The yeast v-SNARE Vti1p mediates two vesicle transport pathways through interactions with the t-SNAREs Sed5p and Pep12p. J Cell Biol 137: 1511–1524 Watson RT, Pessin JE (2001) Transmembrane domain length determines intracellular membrane compartment localization of syntaxins 3, 4, and 5. Am J Physiol Cell Physiol 281: C215–C223 Weinberger A, Kamena F, Kama R, Spang A, Gerst JE (2005) Control of Golgi morphology and function by Sed5 t-SNARE phosphorylation. Mol Biol Cell 16: 4918–4930 Williams AL, Ehm S, Jacobson NC, Xu D, Hay JC (2004) rsly1 binding to Syntaxin 5 is required for endoplasmic reticulum-to-Golgi transport but does not promote SNARE motif accessibility. Mol Biol Cell 15: 162–175 Wimmer C, Hohl TM, Hughes CA, Muller SA, Sollner TH, Engel A, Rothman JE (2001) Molecular mass, stoichiometry, and assembly of 20 S particles. J Biol Chem 276: 29091–29097 Wooding S, Pelham HR (1998) The dynamics of Golgi protein traffic visualized in living yeast cells. Mol Biol Cell 9: 2667–2680 Xu D, Joglekar AP, Williams AL, Hay JC (2000) Subunit structure of a mammalian ER/ Golgi SNARE complex. J Biol Chem 275: 39631–39639 Xu Y, Martin S, James DE, Hong W (2002) GS15 forms a SNARE complex with syntaxin 5, GS28, and Ykt6 and is implicated in traffic in the early cisternae of the Golgi apparatus. Mol Biol Cell 13: 3493–3507 Xu Y, Zhang F, Su Z, McNew JA, Shin YK (2005) Hemifusion in SNARE-mediated membrane fusion. Nat Struct Mol Biol 12: 417–422 Yamaguchi T, Dulubova I, Min SW, Chen X, Rizo J, Sudhof TC (2002) Sly1 binds to Golgi and ER syntaxins via a conserved N-terminal peptide motif. Dev Cell 2: 295–305 Yang B, Gonzalez LJr, Prekeris R, Steegmaier M, Advani RJ, Scheller RH (1999) SNARE interactions are not selective. Implications for membrane fusion specificity. J Biol Chem 274: 5649–5653

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Yao R, Ito C, Natsume Y, Sugitani Y, Yamanaka H, Kuretake S, Yanagida K, Sato A, Toshimori K, Noda T (2002) Lack of acrosome formation in mice lacking a Golgi protein, GOPC. Proc Natl Acad Sci USA 99: 11211–11216 Zhang T, Hong W (2001) Ykt6 forms a SNARE complex with syntaxin 5, GS28, and Bet1 and participates in a late stage in endoplasmic reticulum-Golgi transport. J Biol Chem 276: 27480–27487 Zolov SN, Lupashin VV (2005) Cog3p depletion blocks vesicle-mediated Golgi retrograde trafficking in HeLa cells. J Cell Biol 168: 747–759

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Rabs Christoph Claas, Alexander A. Mironov and Vytaute Starkuviene

In this chapter we will describe the Rab family of proteins, one of the most important molecular machines operating within the secretory pathway. Rab proteins (Ras-related proteins in brain) are evolutionary conserved regulators of membrane traffic. They are of key importance for proper functioning of the exocytic secretory pathway and endocytosis. In fact, Rabs seem to play a role in all events of intracellular transport, i.e. they take part in cargo selection and budding of vesicular (or tubular) structures in donor compartments, they regulate the transport of these membrane-bound structures along cytoskeletal tracks, they control their docking to the target membrane, and they are involved in the final fusion of donor and target membrane. Moreover, they guarantee that organelles keep their protein and lipid composition as well as their right position within the cell (Seabra et al. 2002; Grosshans et al. 2006). Whereas 11 Rabs have been described in Saccharomyces cerevisiae, more than 70 Rabs and Rab-like proteins exist in mammals (Pereira-Leal and Seabra 2001; Gurkan et al. 2005). Consequently, together with the SNAREs the Rabs are the largest of the protein families involved in membrane trafficking.

Membrane attachment of Rabs Rabs are synthesized as soluble molecules, but their function depends on association with the cytoplasmic leaflet of cellular membranes – only after post-translational addition of two isoprenoid molecules (prenylation) at their C-terminus they are able to become membrane-associated (Pereira-Leal et al. 2001; An et al. 2003). This post-translational modification is catalyzed by socalled Rab escort protein (REP). REP binds to the newly synthesized Rab in GDPbound form and simultaneously to Rab geranylgeranyl transferase (RGGT or protein prenyl transferase, Pylypenko et al. 2003). After modification is accomplished, RGGT dissociates from REP–Rab complex, and REP delivers the Rab to the membrane (Thoma et al. 2001). Then, REP recycles back to bind new molecules of freshly synthesized Rab in the cytoplasm. Since there are only two isoforms of REP in mammalian cells, REP is able to bind many distinct Rab proteins via conserved residues (Alory and Balch 2001; Pfeffer 2005). Membrane-associated Rabs can either be converted into the active GTPbound form (see below) or they are retrieved from the membrane as Rab–GDP by cytoplasmic proteins called GDP dissociation inhibitor (GDI, Wu et al. 2007). GDI is able to bind to prenylated Rabs in the GDP-bound form (Wilson et al. 1996). Similar to the Rab-binding subunit of REP, GDIs hide

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prenyl groups attached to Rabs within a hydrophobic groove (Pylypenko et al. 2006), thereby keeping Rabs hydrophilic and enabling them to reside in the cytoplasm. Upon the action of GDI displacement factor (GDF), it is proposed that the complex of Rab–GDP and GDI dissociates, prenyl anchors can be inserted into the membrane, and as a result the Rab molecule becomes associated with the membrane (Dirac-Svejstrup et al. 1997). Little is known about the detailed mechanism leading to removal of GDI in the course of Rab membrane attachment. In fact, to date only one protein with GDF activity has been identified in yeast and mammals. This protein, Yip3 (PRA1), belongs to the Yip (Ypt-interacting proteins) family of proteins and is able to dissociate the Rab9–GDI complex and to recruit Rab9 to endosomal membranes (Silvar et al. 2003). At least five GDI isoforms are expressed in mammalian cells (Alory and Balch 2003), and structural analysis of some of them highlighted the mechanisms of their association to membranes and Rabs (Schalk et al. 1996; Luan et al. 2000). Human aGDI is highly enriched in brain tissue with low abundance in other cells, whereas bGDI is the main, “housekeeping” isoform being expressed ubiquitously (Nishimura et al. 1994). Therefore, their function could be different: aGDI is involved in sorting of highly specialised vesicles in brain such as neurosecretory vesicles, in difference to bGDI, which plays a general role in vesicular trafficking in diverse types of cells. Consequently, GDI isoforms are not specific and recognize a broad range of Rab species (Ullrich et al. 1993). Curiously, REP and GDI proteins are sharing common Rab-binding properties and high sequence similarity, so that they can be grouped into one evolutionary conserved family of REP/GDI proteins (Alory and Balch 2001). Still, the functions of both proteins are not interchangeable: GDI cannot assist in the prenylation of newly synthesized Rab proteins and REP cannot retrieve Rab proteins from membranes.

Rabs as molecular switches Rabs function as GTPases that cycle between a GTP- and a GDP-bound state (Dumas et al. 1999; Pasqualato et al. 2004). Only the GTP-bound form is functionally active and can recruit diverse proteins, so-called effectors (see later), which, in turn, participate in different steps of vesicular traffic (Grosshans et al. 2006). The inactive GDP-bound form can be either cytoplasmic or membrane-associated (Wittmann and Rudolph 2004). Because of the low rates of nucleotide exchange and hydrolysis, the activity switch function of Rabs is under the control of GDP–GTP exchange factors (GEFs) and GTPase activating proteins (GAPs) (Bos et al. 2007). Hydrolysis of GTP leads to inactivation of the Rab and the GDP-bound form can then be extracted from the membrane by GDI (Ullrich et al. 1993). GDI then periodically delivers cytosolic Rab to the appropriate organelle (Pfeffer and Aivazian 2004). When Rab–GDP becomes membrane-bound, a Rab GEF can promote exchange of GDP with GTP, thus converting Rab into the active GTP-bound

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form (Bos et al. 2007). Standard models of Rab dynamics suggest a continuous cycle of organelle association and dissociation (Segev 2001; Pfeffer and Aivazian 2004; Bos et al. 2007). In contrast to general regulators of Rab function such as REP and GDI proteins, GEFs and GAPs are specific for one or a few Rab family members, tissues or even subcellular localization (Bernards 2003; Bos et al. 2007). For instance, over-expression of Rab GAPs can be used to specifically inactivate the endogenous pool of a Rab and, thus, to interfere with the process this Rab is involved in as it was shown by dissecting the internalisation pathways of Shiga toxin and epidermal growth factor EGF (Fuchs et al. 2007). The human genome potentially encodes 39 Rab GAPs; therefore one Rab GAP should be acting upon several Rabs (Bernards 2003). Still, the identification of specific Rab–Rab GAP pairs is not a trivial task (Fuchs et al. 2007). Rab-specific GEFs comprise a diverse group of proteins; therefore their prediction presents a daunting task. Most Rab GEFs are Vps9-, Sec2- or Mss4like proteins (Bos et al. 2007). Little data is available about specific pairs of Rabs and GEFs. By inducing conformational changes to the Switch regions of an associating Rab protein, GEFs are decreasing the affinity for binding GTP or GDP As GEFs and Rabs have no preference for a particular nucleotide, it is the higher concentration of GTP within the cell that results in increased amounts of Rab–GTP upon interaction with GEF (Bos et al. 2007). Finally, since Rabs are regulators of all steps in vesicular traffic, a timely coordinated exchange in the Rabs being “in charge” of a given step has to be ensured (Grosshans et al. 2006). This so-called Rab conversion can be achieved by organizing Rab recruitment and activation into so-called “Rab cascades”. This concept postulates that the GEF of a downstream Rab GTPase will serve at the same time as an effector of an upstream Rab protein (Rink et al. 2005). For instance, Sec2 serves as GEF for the Rab Sec4, which is a regulator of Golgi to plasma membrane (PM) transport. Sec2 is recruited to membranes by the upstream acting Rabs Ypt31/32, thus ensuring timely and spatially coordinated regulation of secretory granule transport to the PM (Ortiz et al. 2002).

Structure of Rabs The structure of no other protein family was so extensively analysed as that of Rabs, including active and non-active states, complexes with REP, GDI and numerous effectors. The interest was raised due to the universality of Rab function, but their high specificity with respect to their target compartments and functional effectors. The versatility of Rabs contrasts with their high sequence homology and similarity in the three-dimensional fold (Pfeffer 2005). As members of the Ras superfamily of proteins the overall structure of Rabs is very similar to other small GTPases (Wennerberg et al. 2005). They share a set of G-box GDP/GTP-binding motifs at their N-terminus, and two switch regions (Switch I and Switch II), which change their conformation depending on which nucleotide is bound (GDP or GTP). Within Rab molecules,

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Switch regions are the only structural elements changing their conformation upon binding of GDP or GTP (Stroupe and Brunger 2000). Despite functional similarities, Switch domains of Rab GTPases have a different orientation than those of Ras proteins, serving as family discriminants. Switch domains of various Rabs crystallized to date overlap significantly in terms of their overall lengths and boundaries (Pfeffer 2005). Therefore, considerable effort was made to find specific structural features that could provide the basis for functional heterogeneity of Rabs. Taking into consideration that Rab effectors are preferentially interacting with GTP-bound proteins, they have to be able to “sense” whether GTP or GDP is bound by the Rab. Consequently, they should interact with sequences close to or within the Switch regions. By this, Switch regions will act not only as providers of information about the activity state of Rabs, but also as recognition domains for effectors. By comparative sequence analysis Pereira-Leal and Seabra (2000) identified five so-called Rab family sequences, F1–F5, that are conserved among Rabs but not Ras or Rho GTPases. Indeed some of them, namely F1, F3 and F5, are located within Switch domains. Moreover, four Rab subfamily-conserved (RabSF) regions were identified by sequence comparison, and have been used to define 10 subfamilies of Rab GTPases (Pereira-Leal and Seabra 2000). Molecular determinants of such specificity were described for some Rab proteins (Constantinescu et al. 2002). Similar to RabF motifs, some of the RabSFs are located within Switch regions (Stroupe and Brunger 2000). Likely, specificity of interactions between Rabs and their effectors are achieved by co-operative binding to RabF and RabSF regions (Pereira-Leal and Seabra 2000). However, regardless of the possibility to cluster Rab molecules according to their primary sequence features, function–structure predictions of uncharacterized Rabs might be not accurate enough as several additional specificity determinants were described (Schwartz et al. 2007). The variety of Rab interactions was attributed not only to differences in the primary sequence, but also to conformational heterogeneity. For instance, structural studies on Rab5C and Rab3A demonstrated that an invariant hydrophobic triad at the Switch region interface might be positioned by diverse angles, thereby creating very distinct surfaces of even closely related Rabs (Merithew et al. 2001). Variability between Rabs is also extended towards the C-terminus. C-termini of Rabs have different lengths and composition and, being the most distinct structural elements of Rabs (Chavrier et al. 1991), contribute to generation of distinct Rab protein surfaces and diversification of their function. Usually, they are called “hyper variable domains”; it was shown that Rab protein localization depends on these domains (Chavrier et al. 1991). However, whereas effectors interacting with multiple Rabs hardly can use these regions for recognition of their target molecules, these regions contain highly conserved residues important for the interaction with GDI and REP proteins (Pylypenko et al. 2003, 2006).

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Compartmentalisation of Rabs Rabs are widely distributed along the secretory pathway, but each member of the Rab family has its particular function inside the cell (Schwartz et al. 2007). Each Rab protein is localized at the membrane of specific intracellular compartments and is highly specific for a particular transport step and its effectors. For instance, Rabs 1 and 2 are important for ER-to-Golgi traffic (Tisdale et al. 1992; Saraste et al. 1995), Rab6 for intra-Golgi transport (Jiang and Storrie 2005), and Rab8 and Rab10 act in the late secretory pathway (Peranen et al. 1996; Babbey et al. 2006). Different Rabs are involved in various types of secretory processes, e.g., Rab3 and Rab27 are playing a role in regulated secretion (Oberhauser et al. 1992; Stinchcombe et al. 2001), Rab5, Rab7 and Rab11 in endocytosis (Ullrich et al. 1996; Pelkmans et al. 2004; Rink et al. 2005). Although some Rabs are tissue-specific (Gurkan et al. 2005), many are ubiquitous in their expression. Often multiple Rabs play a role in the same chain of trafficking events, so their functions need to be coordinated. Rab5, for example, is found on the plasma membrane and early endosomes. Early endosomes also carry Rab4, which acts downstream of Rab5. Rab4 and Rab11 then are localized together on recycling endosomes (Sonnichsen et al. 2000; Zerial and McBride 2001). Similarly, late endosomes carry both Rab7 and Rab9 (Barbero et al. 2002). How is correct delivery of Rabs to destination compartments achieved? Because geranylgeranylation is a common feature of essentially all Rabs and serves for membrane anchoring, it cannot account for their organelle-specific targeting. Specific targeting of Rab proteins via the hyper variable region (Chavrier et al. 1991; Stenmark et al. 1994) is a widely accepted model. It was shown, that replacing the C-terminus of Rab6 with the equivalent C-terminal region of Rab5 resulted in re-localization of the hybrid Rab to Rab5-positive structures, like early endosomes (Stenmark et al. 1994). Similar observations were reported for Rab5 and Rab7 proteins (Ali et al. 2004). However, it has been shown recently that the hyper variable region does not contain a general Rab targeting signal. Reciprocal exchanges of the hyper variable domains of Rab1a, Rab2a, Rab5a, Rab7 and Rab27a failed to re-direct the hybrid proteins away from their original compartment to the new compartment designated by the hyper variable region (Chavrier et al. 1991; Ali et al. 2004). Other regions have been demonstrated to be required within the RabF and RabSF motifs for specific targeting of Rab27a to secretory granules or melanosomes, and Rab5a to endosomes (Ali et al. 2004). Mutations in these targeting-determining regions frequently induced localization to the ER Indeed, the ER and the Golgi apparatus (GA) are preferential sites of membrane translocation for Rabs that have lost targeting information. Interestingly, the localization of the Rab27a was shown to be dependent on the presence of luminally expressed proteins, thus suggesting that cargo molecules might influence their environment including recruitment of specific Rabs (Hannah et al. 2003). In addition, there is evidence that certain receptors

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for Rabs are present in membranes and that those receptors show some preference for the interaction with certain Rab isoforms. Potentially, Yip proteins aid in specific Rab targeting as they can bind to Rab proteins that are in complex with GDI (Pfeffer and Aivazian 2004). However, binding to Yip cannot completely explain highly specific localization of Rabs within a cell as 16 Yip family members in humans (Pfeffer and Aivazian 2004) can most probably not account for proper delivery of roughly 70 Rabs. Many researchers, therefore, assume that Rabs are initially delivered nonspecifically. Only if the Rab protein does not encounter activating proteins and effector proteins already present at this localization or does not recruit these molecules itself to establish a proper Rab-defined complex, GDI will eventually extract Rab–GDP from the membrane and initiate a new cycle of membrane delivery. If the former events take place, positive feedback loops between the function of Rabs and of Rab effectors might lead to stabilization of the Rab localization resulting in a creation of a transient scaffold, a so-called Rab microdomain (Zerial and McBride 2001; Pfeffer and Aivazian 2004; Cai et al. 2007).

Rab effectors Effectors of Rab proteins are defined as proteins that are able to interact with the active, GTP-bound form of a specific Rab, and they exert at least one specific function downstream of this Rab. In contrast to some GDI, GAP, and Rabs themselves, effector proteins display a high degree of structural diversity and belong to a multitude of different protein superfamilies. Numerous interactions of Rabs with effectors have been described and the list of effector molecules is ever growing (Grosshans et al. 2006). Examples for such associations, therefore, have to be limited to a few examples covering each step in membrane trafficking. The first step in trafficking encompasses the selection and concentration of cargo molecules in a subdomain of a donor membrane and the formation of a vesicle. It has been demonstrated that the association of Rab9–GTP with tail-interacting protein of 47 kDa (TIP47) is important for the recycling of mannose6-phosphate receptors from late endosomes to the GA Mannose6phosphate receptor binds to hydrolases bearing mannose6-phosphate in the trans-Golgi network and transports them to late endosomes. Recycling back to the GA enables it to initiate a new cycle of transport. Rab9 on late endosomes increases the affinity of associated TIP47 for binding to the cytoplasmic tail of mannose6-phosphate receptor, thus regulating cargo selection for recycling vesicles (Carroll et al. 2001). Moreover, interaction of Rab9 with its effector TIP47 is required for the correct localization of TIP47 on late endosomes (Carroll et al. 2001). Vice versa, the presence of TIP47 on late endosomes stabilizes the steady-state localization of Rab9 on these organelles (Aivazian et al. 2006). This interplay of a Rab and its effector demonstrates the importance of these interactions for the correct targeting of the trafficking machinery.

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After budding from a donor membrane, newly formed carriers have to become associated with motor proteins moving them along actin filaments or along the microtubule network. Association can be established by direct interaction with a motor protein or indirectly via an adaptor protein. A wellstudied example for the latter case is the association of Rab27a expressed by melanocytes with myosin Va. The effector melanophilin links Rab27a on melanosomes to the actin motor myosin-Va (Fukuda et al. 2002; Nagashima et al. 2002; Strom et al. 2002). Without these interactions melanosomes loose their localization at the periphery of melanocytes and cannot be transported to neighboring keratinocytes (Wu et al. 1998). In retinal cells the same Rab27a was demonstrated to interact with another myosin motor, myosin VII (Gibbs et al. 2004). This interaction depends on the Rab effector MyRIP (Desnos et al. 2003; El-Amraoui et al. 2002), showing that Rab-effector complexes can be cell-type specific. Carriers on the way to their destination compartment have to recognize and to bind to structures on the target membrane. This is achieved by tethering factors, whose function is also regulated by Rabs. In general, tethering factors are either extended proteins with a coiled-coil structure or they are multi-subunit complexes. An example of the former is p115, which is present on ER-derived COPII-coated vesicles. P115 binds to the GM130/ GRASP65 complex present at the cis-GA, and both proteins have been shown to be effectors of Rab1. An intricate net of interactions between Golgi matrix proteins, tethering factors and Rab1 was shown to be essential for material flow through the early secretory pathway (Allan et al. 2002; Moyer et al. 2001; Beard et al. 2005). An example for a Rab-regulated multisubunit tether is represented by the exocyst. The exocyst is an octameric complex that tethers secretory granules to the PM. The yeast Rab Sec4p in its active state is able to interact with the exocyst subunit Sec15p (Guo et al. 1999). Biochemical analyses revealed that another component, Sec8p, also associates with Sec4p, suggesting that Sec4p serves as central regulator of exocyst function (Toikkanen et al. 2003). Later studies indicate that this interaction of Rab and exocyst is conserved in mammals (Zhang et al. 2004). Finally, Rabs play a role in membrane fusion, the last step in trafficking. An influence of Rabs on SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor, see Chapter 2.1) function has been shown by many investigators. The effect seems to be indirect, however, and is exerted by effector molecules directly interacting with Rabs (Grosshans et al. 2006). SNARE proteins induce fusion by bringing the bilayers of opposing vesicular and target membranes into close proximity (Pfeffer 2007; see Chapter 2.1). Shorter et al. (2002) describe that in the process of intra-Golgi transport p115 catalyzes the assembly of a trans-SNARE complex by linking the v-SNARE GOS28 to the t-SNARE Syntaxin5. Another role of Rabs in vesicle fusion could be the prevention of premature complex formation from monomeric SNAREs situated in the same compartment.

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Figure 1. Scheme of the Rab cycle. After its synthesis the Rab molecule (1) is prenylated by RGGT (A). Alternatively, initially Rab binds GDP (2) and then undergoes prenylation. After prenylation Rab binds REP (4) that accompanies Rab to the membrane (tree-layered arc filled with grey color). After detachment of REP (4), Rab (1) bound to GDP (2) binds RabGEF (5) that exchanges GDP for GTP (3). RabGTP interacts with SNARE preventing formation of a SNARE complex with the membrane plane. After fusion, the SNARE detaches from the Rab and the Rab binds to Rab GAP (6). This leads to hydrolysis of GTP and liberation of phosphate (P). After detachment of RabGAP, RabGDP can detach from the membrane and bind to Rab GDI (7). Rab GDI delivers the Rab back to a membrane and a new cycle of Rab activation can start.

Conclusion A plethora of different proteins is involved in the trafficking of membranebound structures within a cell. Besides Rabs, coat proteins, SNAREs, and motor proteins are of key importance. The number of Rabs during evolution, however, has increased much more than those of other proteins in this context, matching the needs of increasing complexity of membrane systems from yeast to mammals. This reflects the central role that Rabs play in the coordination of diverse steps during trafficking – some authors conceptualize Rabs as the central organizers or central hubs of membrane trafficking (Gurkan et al. 2005). Certainly, their function as binary switches oscillating between an active and an inactive form alone cannot explain how these molecules can contribute to complex spatial and timely regulation which is needed for proper trafficking. Presumably, positive feedback loops between Rab, Rab function modulating proteins and effector molecules stabilize the activity and localization of Rabs, and contribute to the establishment of Rabdefined microdomains within the membrane (Zerial and McBride 2001). Moreover, protein machineries in such microdomains have to be coordinated

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in terms of assembly, disassembly and interplay with other machineries to ensure proper regulation of successive steps during vesicle transport and, consequently, to ensure cellular homeostasis. The concept of organizing different Rabs into Rab cascades as described above may explain at least in part how this task is achieved (Rink et al. 2005). Still, studies on so far uncharacterized Rabs, the identification of more Rab-interacting proteins and, most importantly, knowledge about detailed mechanisms of the interplay of all these components will be required to fully understand the surprising versatility of Rab–GTPases.

Abbreviations GA GAP GDF GDI GEF PM REP RGGT Yip

Golgi apparatus GTPase-activating protein GDI displacement factor GDP dissociation inhibitor GDP–GTP exchange factor plasma membrane Rab escort protein Rab geranylgeranyl transferase Ypt-interacting protein

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COPII Ken Sato and Akihiko Nakano

Introduction Vesicular traffic provides a dynamic and elaborate communication network between the subcellular compartments that define the structure and identity of membrane-bound organelles (Bonifacino and Glick 2004). The molecular and structural mechanisms that direct lipid and protein cargo flow between discontinuous subcellular organelles involve specialized multiprotein machineries that are defined by the molecular and structural properties of cytosolic coat protein complexes (Bonifacino and LippincottSchwartz 2003). The Golgi apparatus is certainly involved in this flow. The endoplasmic reticulum (ER) is responsible for the synthesis of the proteins of most of the cellular organelles. Newly synthesized secretory proteins are translated at the rough ER and translocated into the ER lumen or ER membrane through the translocation channel, where they undergo folding, assembly and post-translational modifications with the aid of a variety of ER chaperones. Correctly folded and assembled secretory proteins are then segregated from ER resident proteins and transported to the Golgi apparatus for further processing and secretion. The ER-to-Golgi transport step is thought to occur via membrane-bound vesicles or carrier intermediates, which are formed by the assembly of the coat protein complex II (COPII) on the ER membranes. COPII is the name given to a cytosolic protein complex required for direct capture of cargo molecules and for the physical deformation of the ER membrane that drives the formation of the so-called “COPII vesicles” or carrier intermediates in anterograde transport from the ER to the Golgi. Protein export by COPII vesicle from the ER is the default ER-toGolgi route that has been proposed in yeast and mammals. Cargo proteins that are destined for delivery to the Golgi apparatus need not only refer to newly synthesized biosynthetic cargo molecules, but also a variety of other machinery proteins that constantly cycle between the ER and the Golgi are included. The molecular mechanistic details of COPII vesicle formation and following cargo delivery to the Golgi have been defined with greater precision through yeast genetics and in vitro reconstitution. Orthologues of almost all of these yeast genes have now been shown to play an equally important role in higher eukaryotes COPII function. Although none of the COPII components are predominantly localized to the Golgi, the COPII-mediated ER-to-Golgi transport is essential for maintaining the function and identity of the Golgi. In this

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section, we focus on the individual contributions of the COPII components to the ER-to-Golgi trafficking.

COPII coat recruitment and assembly Pioneering studies by Schekman and colleagues first identified the components of the COPII coat, Sar1 (Nakano and Muramatsu 1989), Sec23/24 (Hicke et al. 1992), Sec13/31 (Salama et al. 1993), and Sar1 regulator Sec12 (Nakano et al. 1988) in the yeast Saccharomyces cerevisiae as proteins required for ER exit in a genetic screen. Their physical interaction and ability to generate COPII vesicles in vitro from isolated ER membranes was shown a few years after the original discovery of these proteins (Barlowe et al. 1994). These COPII components generate COPII vesicles through a sequence of events (Fig. 1). A wealth of genetic and biochemical experiments has led to consensus that in the first step of COPII coat assembly is initiated by activation of the small Ras-like GTPase Sar1. Similar to other small GTPases, conversion of Sar1-GDP to Sar1-GTP is mediated by guanine nucleotide exchange factor (GEF). Sec12 is an ERanchored transmembrane GEF for Sar1 (Barlowe and Schekman 1993). Since Sec12 is strictly regulated to localize to the ER by static retention (Sato et al. 1996), Sar1 activation is restricted to the ER. The GTP binding triggers the exposure of the N-terminal amphipathic a-helix element of Sar1 that inserts into the ER membrane (Huang et al. 2001; Bi et al. 2002). Membrane insertion of the N-terminal helix is a prerequisite for GTP exchange and thus the GTP loading to Sar1 proceeds only in the presence of a membrane surface. Sar1GTP recruits the Sec23–Sec24 heterodimer by binding to the Sec23 portion to form a so-called “prebudding complex” (Kuehn et al. 1998). The Sec23 subunit of Sec23/24 complex is the GTPase-activating protein (GAP) for Sar1 (Yoshihisa et al. 1993) and therefore stimulates Sar1 GTP hydrolysis upon binding to Sar1, which leads to the loss from the membrane of both Sar1 and Sec23/24 complex (Antonny et al. 2001). However, kinetically stable Sec23/24–Sar1 complexes are maintained on the membrane by the presence of the GEF Sec12, which counteracts the GTPase-stimulating activity of the Sec23 by continually recharging Sar1 with GTP (Futai et al. 2004). Subsequently, the prebudding complex recruits Sec13–Sec31 heterotetramer onto the prebudding complex, providing the outer layer of the coat (Lederkremer et al. 2001). Since the Sec31 subunit interacts with both the Sec23 and Sec24 (Shaywitz et al. 1997), but not with Sar1 or directly with the membrane, it is likely that Sec13/31 complex cross-links the preassembled prebudding complexes and drive membrane deformation to form COPII vesicles (60–70 nm in diameter). A recent cryoelectron-microscopy study has proposed that Sec13/31 complex forms a cuboctahedral lattice whose faces are squares and triangles (Stagg et al. 2006). COPII vesicles could be generated from synthetic liposomes incubated with only the above-defined proteins Sec23/24 complex, Sec13/31 complex and GTP-locked Sar1 with non-hydrolyzable GTP analog (Matsuoka et al. 1998). They are therefore “core” components together with the Sec12 GEF.

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Figure 1. COPII vesicle formation and the selective uptake of cargo proteins. The COPII vesicle formation is initiated by GDP–GTP exchange on Sar1 catalyzed by Sec12. Activated Sar1–GTP binds to the ER membrane and recruits the Sec23/24 subcomplex. The cytoplasmically exposed signal of transmembrane cargo is captured by direct contact with Sec24, forming the prebudding complex. These prebudding complexes are clustered by the Sec13/31 subcomplex, generating COPII-coated vesicles. The assembled cargo has a high affinity for the Sec23/24 because of the combined export signals. This cargo-Sec23/24 association persists during the GDP–GTP exchange of Sar1 catalyzed by Sec12 (upper panel). In contrast, the Sar1 GTP hydrolysis dissociates the weak association between the coat and unassembled cargo or lipid before polymerizing into COPII coats (lower panel). Thus, the prebudding complex stabilities are biased towards the complex including assembled cargo, ensuring that the fully assembled cargo is preferentially incorporated into COPII vesicles.

In addition to the core COPII components, given the likely specialization of trafficking components to accommodate an extraordinary variety of cargo proteins with different structures, sizes and functions with COPII coat assembly, many other proteins are seem to be required to export cargo from the ER. The evolutionarily conserved Sec16 is a large peripheral protein that associates with the ER membrane, and shown to be an essential gene in S. cerevisiae and Pichia pastoris (Espenshade et al. 1995; Connerly et al. 2005). In vitro COPII vesicle budding reaction performed with isolated ER membranes stripped of their Sec16 reveals that vesicle formation is significantly reduced. Sec16 contains domains that make direct contact with the

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COPII coat components and may act as a scaffold for assembly of the coat (Gimeno et al. 1996; Supek et al. 2002). Another related protein isolated as a dosage-dependent suppressor of temperature-sensitive SEC16 mutations is Sed4. Sed4 is an integral membrane protein located at the ER membrane, and deletion of the SED4 gene from wild-type cells retards transport from the ER to the Golgi (Gimeno et al. 1995). Although the cytoplasmic domain of Sed4 shares significant homology with that of Sec12, no GEF activity has been found (Saito-Nakano and Nakano 2000). This cytoplasmic domain interacts directly with Sec16 at the ER membrane (Gimeno et al. 1995) and these factors are likely to function together, though their roles are less understood. Yip1p is a member of a conserved family of integral membrane proteins that interact with Rab/Ypt GTPases (Yang et al. 1998). Yip1 forms a complex with at least two other proteins Yif1 and Yos1, and Yip1/Yif1/Yos1 complex cycles between the ER and the Golgi. This complex is required for COPII vesicle biogenesis as Yip1 antibodies inhibit cargo export from the ER and yeast yip1 mutants are defective in COPII vesicle generation (Heidtman et al. 2003, 2005). The mammalian Yip1 is found to interact with the Sec23/24 complex (Tang et al. 2001). A role for Yip1/Yif1/Yos1 complex in COPII assembly is not still elucidated but obviously required for COPII-mediated cargo exit. COPII does not assemble randomly throughout the ER membranes in vivo but instead is concentrated at specialized regions termed transitional ER (tER) or ER exit sites (ERES) (Orci et al. 1991; Bannykh et al. 1996). Immunoelectron and fluorescence microscopy of COPII in coated buds shows a restricted localization to these sites, which represent domains of the ER responsible for the generation of COPII vesicles. At least, Sec16 is shown to be required for normal tER organization (Connerly et al. 2005), but it is not certain how these distinct zones are maintained and the proteins that build these sites are not fully identified. Moreover, the functional consequence of these specialized budding zones is not known, although the extent of organization of these ER sites might influence the morphology of newly forming Golgi elements (Rossanese et al. 1999).

Cargo selection by COPII components It is now widely accepted that the majority of cargo proteins are actively sorted into COPII vesicles. The formation of the prebudding complex consisting of Sec23/24–Sar1 bound to cargo protein is the cargo recognition step prior to polymerization by Sec13/31. The Sec24 subunit in the prebudding complex is generally responsible for interactions with cargo molecules (Mossessova et al. 2003; Miller et al. 2003). The selective capture is basically driven by export signals within the amino acid sequence of each transmembrane cargo protein contained on their cytoplasmically exposed regions, but some transmembrane and most soluble cargo proteins require

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specific transmembrane cargo receptors/adaptors to mediate the interaction with the COPII coats. These signals are quite diverse, and several classes of ER export signals have so far been identified on the cytoplasmic regions of transmembrane cargo proteins and transmembrane receptors/ adaptors from various organisms (Barlowe 2003). For example, di-acidic ((D/E)x(D/E), with x representing any amino acid residue) and di-hydrophobic (FF, YY, LL, or FY) motifs are well characterized. Interestingly, multiple distinct Sec24 family members are able to pair with Sec23 and bind to export signals different from those recognized by Sec24 (Pagano et al. 1999; Miller et al. 2003), expanding the cargo multiplicity captured by COPII coat. The ER contains a certain amount of newly synthesized unfolded or unassembled cargo proteins, which should be segregated from secretory proteins to be exported. To ensure efficient incorporation of fully folded and assembled cargo proteins into COPII vesicles, many exported proteins are required multiple signals in a specific display that is achieved only through proper assembly. In vitro experiments with cargo-reconstituted proteoliposomes have demonstrated that the Sec23/24 can remain transiently associated with properly assembled cargo even after Sar1 GTP hydrolysis, while the interaction between Sec23/24 and unassembled cargo or lipid on membranes is disrupted immediately upon Sar1 GTP hydrolysis (Sato and Nakano 2005). This is probably due to combined export signals displayed on assembled complex might increase the affinity to Sec23/24. In the presence of Sec12, stable binding of Sec23/24 with assembled cargo proteins has been observed due to continuous reactivation of Sar1 to its GTP-bound form before Sec23/24 release. In contrast, continuous Sec23/24 binding and release occurs on unassembled cargo and lipids accompanied by Sar1 GTPase cycles (Fig. 1). Thus, the prebudding complex stability is biased toward the complex containing properly assembled cargo proteins during Sar1 GTPase cycles. Thus, Sar1 selectively promotes exclusion of unassembled cargo proteins from emerging COPII vesicles by virtue of its GTP hydrolysis, explaining how only proper cargo molecules can be efficiently concentrated into COPII vesicles.

COPII vesicle budding and pinch-off Sar1 provides not only as a spatial landmark that recruits Sec23/24 and Sec13/31 but also as a mediator of membrane deformation and vesicle scission. There is evidence that Sar1-GTP can deform liposomes into tubules about 26 nm in diameter (Lee et al. 2005). This was observed only with activated Sar1-GTP, suggesting an involvement of the N-terminal helix into the outer leaflet of the ER membrane would act to displace the lipid headgroups, and this asymmetric expansion may promote curvature toward the cytoplasmic region (Farsad and De Camilli 2003). Sar1-GTP then stimulates the binding of an inner shell of Sec23/24 to form prebudding complex. It has

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been shown in structural studies that the Sec23/24 complex has a concave surface that matches the size of a 60-nm vesicle and is thus thought to facilitate membrane bending (Bi et al. 2002). Such prebudding complexes only give a random distribution of local membrane deformations because they have no ability to interact with each other. Sec13/31 alone is shown to self-assemble in solution into COPII cage-like structures (Stagg et al. 2006), and hence the spherical shape of the COPII coat seems to be formed by prebudding complex clustering by Sec13/31. Importantly, although Sec23/24 and Sec13/31 complexes can form budded vesicles, the buds rarely closed off to generate free vesicles when Sar1 is anchored in the nickel-conjugated lipids containing liposomes by a polyhistidine tag in place of N-terminal helix (Lee et al. 2005). So, the neck of a COPII coated bud is not likely to break spontaneously and the N-terminal helix of Sar1 may have an active role in membrane fission. Other experiments suggest that fission is more efficient when Sar1 hydrolyses GTP (Bielli et al. 2005), suggesting that the Sar1 GTP hydrolysis may play some roles in vesicle fission. Further analyses are required to determine how Sar1 N-terminal helix initiates and completes the fission of a COPII vesicle.

Building a Golgi with COPII vesicles COPII vesicles shed their coats before fusion with Golgi membrane and this uncoating reaction is thought to be achieved by the Sar1 GTP hydrolysis (Oka and Nakano 1994). In mammalian cells COPII vesicles derived from the tER do not fuse directly to the Golgi membrane, instead, they appear to tether and fuse to each other (homotypic fusion) to form carrier intermediates that lie adjacent to the tER (Xu and Hay 2004). COPII vesicles continue to fuse with the intermediates, which becomes larger and eventually thought to fuse with the Golgi. Numerous names have been given to the discontinuous carriers that move from the ER compartment to Golgi such as ER–Golgi intermediate compartment (ERGIC), pre-Golgi intermediate and vesicular tubular complex (VTC). Although machineries of the ERto-Golgi trafficking are highly conserved from yeast to human, no equivalent to a pre-Golgi compartment has been identified in the yeast, and COPII vesicles are thought to fuse directly to the Golgi membranes (heterotypic fusion). However, the high degree of homology between yeast and mammalian components required for the late stages of ER-to-Golgi traffic raises the possibility that yeast COPII vesicles may undergo homotypic tethering and fusion before they fuse with the Golgi. The other way round, the recent observation that the pre-Golgi compartment is stable (Ben-Tekaya et al. 2005; Sannerud et al. 2006) has raised the possibility that homotypic fusion is not exclusive and mammalian COPII vesicles may also heterotypically fuse with stable ER–Golgi intermediate compartments. Much is yet to be determined on how the pre-Golgi structure is formed from COPII vesicles.

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References Antonny B, Madden D, Hamamoto S, Orci L, Schekman R (2001) Dynamics of the COPII coat with GTP and stable analogues. Nat Cell Biol 3: 531–537 Bannykh SI, Rowe T, Balch WE (1996) The organization of endoplasmic reticulum export complexes. J Cell Biol 135: 19–35 Barlowe C (2003) Signals for COPII-dependent export from the ER: what’s the ticket out? Trends Cell Biol 13: 295–300 Barlowe C, Orci L, Yeung T, Hosobuchi M, Hamamoto S, Salama N, Rexach MF, Ravazzola M, Amherdt M, Schekman R (1994) COPII: a membrane coat formed by Sec proteins that drive vesicle budding from the endoplasmic reticulum. Cell 77: 895–907 Barlowe C, Schekman R (1993) SEC12 encodes a guanine-nucleotide-exchange factor essential for transport vesicle budding from the ER. Nature 365: 347–349 Ben-Tekaya H, Miura K, Pepperkok R, Hauri HP (2005) Live imaging of bidirectional traffic from the ERGIC. J Cell Sci 118: 357–367 Bi X, Corpina RA, Goldberg J (2002) Structure of the Sec23/24-Sar1 pre-budding complex of the COPII vesicle coat. Nature 419: 271–277 Bielli A, Haney CJ, Gabreski G, Watkins SC, Bannykh SI, Aridor M (2005) Regulation of Sar1 NH2 terminus by GTP binding and hydrolysis promotes membrane deformation to control COPII vesicle fission. J Cell Biol 171: 919–924 Bonifacino JS, Glick BS (2004) The mechanisms of vesicle budding and fusion. Cell 116: 153–166 Bonifacino JS, Lippincott-Schwartz J (2003) Coat proteins: shaping membrane transport. Nat Rev Mol Cell Biol 4: 409–414 Connerly PL, Esaki M, Montegna EA, Strongin DE, Levi S, Soderholm J, Glick BS (2005) Sec16 is a determinant of transitional ER organization. Curr Biol 15: 1439–1447 Espenshade P, Gimeno RE, Holzmacher E, Teung P, Kaiser CA (1995) Yeast SEC16 gene encodes a multidomain vesicle coat protein that interacts with Sec23p. J Cell Biol 131: 311–324 Farsad K, De Camilli P (2003) Mechanisms of membrane deformation. Curr Opin Cell Biol 15: 372–381 Futai E, Hamamoto S, Orci L, Schekman R (2004) GTP/GDP exchange by Sec12p enables COPII vesicle bud formation on synthetic liposomes. Embo J 23: 4146–4155 Gimeno RE, Espenshade P, Kaiser CA (1995) SED4 encodes a yeast endoplasmic reticulum protein that binds Sec16p and participates in vesicle formation. J Cell Biol 131: 325–338 Gimeno RE, Espenshade P, Kaiser CA (1996) COPII coat subunit interactions: Sec24p and Sec23p bind to adjacent regions of Sec16p. Mol Biol Cell 7: 1815–1823 Heidtman M, Chen CZ, Collins RN, Barlowe C (2003) A role for Yip1p in COPII vesicle biogenesis. J Cell Biol 163: 57–69 Heidtman M, Chen CZ, Collins RN, Barlowe C (2005) Yos1p is a novel subunit of the Yip1p–Yif1p complex and is required for transport between the endoplasmic reticulum and the Golgi complex. Mol Biol Cell 16: 1673–1683 Hicke L, Yoshihisa T, Schekman R (1992) Sec23p and a novel 105-kDa protein function as a multimeric complex to promote vesicle budding and protein transport from the endoplasmic reticulum. Mol Biol Cell 3: 667–676 Huang M, Weissman JT, Beraud-Dufour S, Luan P, Wang C, Chen W, Aridor M, Wilson IA, Balch WE (2001) Crystal structure of Sar1-GDP at 1.7 A resolution and the role of the NH2 terminus in ER export. J Cell Biol 155: 937–948 Kuehn MJ, Herrmann JM, Schekman R (1998) COPII-cargo interactions direct protein sorting into ER-derived transport vesicles. Nature 391: 187–190

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Lederkremer GZ, Cheng Y, Petre BM, Vogan E, Springer S, Schekman R, Walz T, Kirchhausen T (2001) Structure of the Sec23p/24p and Sec13p/31p complexes of COPII. Proc Natl Acad Sci USA 98: 10704–10709 Lee MC, Orci L, Hamamoto S, Futai E, Ravazzola M, Schekman R (2005) Sar1p N-terminal helix initiates membrane curvature and completes the fission of a COPII vesicle. Cell 122: 605–617 Matsuoka K, Orci L, Amherdt M, Bednarek SY, Hamamoto S, Schekman R, Yeung T (1998) COPII-coated vesicle formation reconstituted with purified coat proteins and chemically defined liposomes. Cell 93: 263–275 Miller EA, Beilharz TH, Malkus PN, Lee MC, Hamamoto S, Orci L, Schekman R (2003) Multiple cargo binding sites on the COPII subunit Sec24p ensure capture of diverse membrane proteins into transport vesicles. Cell 114: 497–509 Mossessova E, Bickford LC, Goldberg J (2003) SNARE selectivity of the COPII coat. Cell 114: 483–495 Nakano A, Brada D, Schekman R (1988) A membrane glycoprotein, Sec12p, required for protein transport from the endoplasmic reticulum to the Golgi apparatus in yeast. J Cell Biol 107: 851–863 Nakano A, Muramatsu M (1989) A novel GTP-binding protein, Sar1p, is involved in transport from the endoplasmic reticulum to the Golgi apparatus. J Cell Biol 109: 2677–2691 Oka T, Nakano A (1994) Inhibition of GTP hydrolysis by Sar1p causes accumulation of vesicles that are a functional intermediate of the ER-to-Golgi transport in yeast. J Cell Biol 124: 425–434 Orci L, Ravazzola M, Meda P, Holcomb C, Moore HP, Hicke L, Schekman R (1991) Mammalian Sec23p homologue is restricted to the endoplasmic reticulum transitional cytoplasm. Proc Natl Acad Sci USA 88: 8611–8615 Pagano A, Letourneur F, Garcia-Estefania D, Carpentier JL, Orci L, Paccaud JP (1999) Sec24 proteins and sorting at the endoplasmic reticulum. J Biol Chem 274: 7833–7840 Rossanese OW, Soderholm J, Bevis BJ, Sears IB, O’Connor J, Williamson EK, Glick BS (1999) Golgi structure correlates with transitional endoplasmic reticulum organization in Pichia pastoris and Saccharomyces cerevisiae. J Cell Biol 145: 69–81 Saito-Nakano Y, Nakano A (2000) Sed4p functions as a positive regulator of Sar1p probably through inhibition of the GTPase activation by Sec23p. Genes Cells 5: 1039–1048 Salama NR, Yeung T, Schekman RW (1993) The Sec13p complex and reconstitution of vesicle budding from the ER with purified cytosolic proteins. EMBO J 12: 4073–4082 Sannerud R, Marie M, Nizak C, Dale HA, Pernet-Gallay K, Perez F, Goud B, Saraste J (2006) Rab1 defines a novel pathway connecting the pre-Golgi intermediate compartment with the cell periphery. Mol Biol Cell 17: 1514–1526 Sato K, Nakano A (2005) Dissection of COPII subunit-cargo assembly and disassembly kinetics during Sar1p–GTP hydrolysis. Nat Struct Mol Biol 12: 167–174 Sato M, Sato K, Nakano A (1996) Endoplasmic reticulum localization of Sec12p is achieved by two mechanisms: Rer1p-dependent retrieval that requires the transmembrane domain and Rer1p-independent retention that involves the cytoplasmic domain. J Cell Biol 134: 279–293 Shaywitz DA, Espenshade PJ, Gimeno RE, Kaiser CA (1997) COPII subunit interactions in the assembly of the vesicle coat. J Biol Chem 272: 25413–25416 Stagg SM, Gurkan C, Fowler DM, LaPointe P, Foss TR, Potter CS, Carragher B, Balch WE (2006) Structure of the Sec13/31 COPII coat cage. Nature 439: 234–238 Supek F, Madden DT, Hamamoto S, Orci L, Schekman R (2002) Sec16p potentiates the action of COPII proteins to bud transport vesicles. J Cell Biol 158: 1029–1038 Tang BL, Ong YS, Huang B, Wei S, Wong ET, Qi R, Horstmann H, Hong W (2001) A membrane protein enriched in endoplasmic reticulum exit sites interacts with COPII. J Biol Chem 276: 40008–40017

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COPI: mechanisms and transport roles Victor W. Hsu, Jia-Shu Yang, and Stella Y. Lee

Coat protein I (COPI) is considered one of the best characterized coat complexes, which represent the core machinery by which vesicle formation and cargo sorting are coupled to initiate vesicular transport (Bonifacino and Lippincott-Schwartz 2003; McMahon and Mills 2004). Our understanding of the molecular mechanisms by which COPI acts and the transport pathways in which it operates has evolved significantly over the years, and with considerable accompanying controversy. These aspects of COPI research will be reviewed. See also Fig. 1 for a timeline that summarizes its key discoveries.

Historic background The origin of COPI could be traced to two lines of investigations, which initially seemed distinct. One avenue of research had sought to reconstitute transport among the Golgi cisternae using a cell-free system (Balch et al. 1984). This reconstitution took advantage of a mutant cell line that was defective in a specific medial Golgi glycosylation reaction. These cells were infected with the vesicular stomatitis virus (VSV). A Golgi-enriched fraction was then collected and incubated with a similarly enriched fraction from uninfected wild-type cells. Transport between the two Golgi fractions was marked by the transfer of the major G protein expressed by the VSV infection (and thus known as VSVG), which was monitored by its glycosylation pattern upon arrival to the acceptor fraction (Balch et al. 1984). Subsequent characterization of this reconstitution system revealed that it could be blocked by the addition of a nonhydrolyzable analog of guanosine triphosphate (GTP), known as GTPgS. Electron microscopy (EM) revealed that this block induced the accumulation of coated vesicles (Orci et al. 1986). As this coating was distinct from the only other coat protein known at the time, clathrin with the AP2 adaptor, the Golgi-derived vesicles were proposed to be formed by a novel, non-clathrin coat complex (Orci et al. 1986). Later, key components of this coating were identified to be part of a multimeric complex, known as coatomer (Waters et al. 1991). Moreover, coated vesicles formed in the presence of GTPgS contained coatomer in stoichiometric level to the small GTPase ADP Ribosylation factor 1 (ARF1) (Serafini et al. 1991a). Thus, the novel coat was considered to consist of ARF1 and coatomer (Orci et al. 1993; Serafini et al. 1991a), and given the name coat protein (COP) (Serafini et al. 1991a; Waters et al. 1991). Subsequent-

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Figure 1. A timeline summary of key insights in COPI research. Key mechanistic insights or events are highlighted above the arrow line, while key components are highlighted below this line.

ly, this name was changed to COPI when COPII was identified (Barlowe et al. 1994). Another line of research had sought to elucidate how a pharmacologic agent, brefeldin-A (BFA), blocked cellular secretion. Initially, BFA was found to induce the redistribution of the Golgi complex to the endoplasmic reticulum (ER), resulting in a mixed organellar system that was incapable of supporting transport to the plasma membrane (Lippincott-Schwartz et al. 1989). To understand how BFA induced this redistribution of the Golgi complex, an early insight was that certain Golgi-localized proteins were observed to become released to the cytosol upon treatment by BFA. The redistribution of one such protein, dubbed the 110 kDa protein, was notable due to its additional regulation by GTPgS, such that pre-incubation with GTPgS prevented BFA from releasing this protein on Golgi membrane (Donaldson et al. 1991). Notably, this line of investigation merged with the ongoing studies on reconstituted intra-Golgi transport, when the 110 kDa protein was found to be identical to a subunit of coatomer, named b-COP (Duden et al. 1991; Serafini et al. 1991b). Subsequent studies revealed that activation of ARF1 by its binding of GTP recruited coatomer from the cytosol onto Golgi membrane to initiate COPI vesicle formation (Donaldson et al. 1992a; Orci et al. 1991). Moreover, BFA inhibited ARF1 activation by blocking the guanine nucleotide exchange activity that catalyzed this activation (Donaldson et al. 1992b; Helms and Rothman 1992). These findings in COPI research have provided seminal contributions to our understanding of how the ARF family of small GTPases regulates coat proteins to initiate vesicle formation. However, subsequent studies on this regulation have revealed surprising mechanistic complexities. Moreover, whereas the role of COPI vesicles was originally envisioned to mediate anterograde (forward) transport through the Golgi complex, this role has also undergone dramatic revisions subsequently. Below, we will first discuss the key components discovered for COPI vesicle formation, and then review the intracellular pathways in which COPI has been implicated to act.

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Key components in COPI vesicle formation Much of our current mechanistic understanding of how the different factors act in COPI vesicle formation has come largely from a vesicle reconstitution system that evolved from the original intra-Golgi transport assay. Experimentally, this involves the incubation of Golgi membrane with purified protein components, which initially suggested how the GTPase cycle of ARF1 regulates coatomer to initiate COPI vesicle formation (Orci et al. 1991; Serafini et al. 1991a; Tanigawa et al. 1993). Later, in its more refined form, this reconstitution system has been instrumental in identifying additional key factors (Lee et al. 2005; Ostermann et al. 1993; Yang et al. 2002, 2005, 2006). Summarizing how these factors are currently thought to act, Fig. 2 proposes a model.

Coatomer Coatomer is a multimeric complex (composed of a, b, b0 , g, d, e, and z subunits) (Harrison-Lavoie et al. 1993; Stenbeck et al. 1993; Waters et al. 1991). Its role as the principal coating on COPI vesicles was initially suggested through the COPI vesicle reconstitution system (Orci et al. 1993; Serafini et al. 1991a). Examining a minimal system using liposomal membrane rather than Golgi membrane, later studies further suggested that coatomer and ARF1 were sufficient to form vesicular-like structures (Bremser et al. 1999; Spang et al. 1998).

Figure 2. A model proposed for how key components act in COPI vesicle formation. Besides coatomer, ARFGAP1 is now realized to be another coat component, while BARS/endophilin B have been identified to have mechanistically interchangeable roles in the fission step.

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However, more recent vesicle reconstitution studies that have re-visited the use of Golgi membrane have revealed additional factors needed, not only for COPI vesicle formation but also for its proper cargo sorting (see subsections below that discuss such factors in more detail). Sub-complexes of coatomer have been delineated based on physical interactions among its subunits (Eugster et al. 2000; Lowe and Kreis 1995; Pavel et al. 1998). One sub-complex consists of a, b0 , and e subunits, while the other consists of the remaining subunits. With their sequencing, the b, g, and z subunits of coatomer have been revealed to have detectable levels of sequence similarity to b, m, and s subunits of the clathrin AP2 adaptors, respectively (Cosson et al. 1996; Serafini et al. 1991b). Supporting this parallel, structural analysis also indicates that g-COP shares folding similarity to the appendage domains of a and b subunits of the clathrin AP2 adaptor (Hoffman et al. 2003). Altogether, these findings have led to the suggestion that coatomer subunits are organized in structurally similar ways as the clathrin coat complex (McMahon and Mills 2004). For cargo sorting, coatomer recognizes two distinct types of di-basic sequences on cargo proteins, known as sorting signals. Di-lysine signals are located near the carboxyl terminus within the cytoplasmic domains of COPI cargo proteins. In contrast, di-arginine signals occur near the amino terminus within the cytoplasmic domain of cargo proteins. Moreover, whereas the dilysine motif shows more strict spacing requirement with respect to the end of the carboxy terminus, the di-arginine motif exhibits more flexibility within the amino terminal region of cargo proteins (Teasdale and Jackson 1996; Zerangue et al. 2001). These two sorting signals are also recognized by distinct components of coatomer, with the di-lysine motif interacting with the a, b0 , and g subunits (Cosson and Letourneur 1994; Harter et al. 1996), and the diarginine motif interacting with b and d subunits (Michelsen et al. 2007). Recently, further complexity in COPI cargo sorting has been revealed by studies on ion channels. Phosphorylation near some di-basic sequences in potassium channels was found to inhibit the binding of coatomer to these motifs. As a mechanistic basis for this inhibition, isoforms of 14-3-3 were identified to recognize such phosphorylated sites, and thereby providing steric hindrance to prevent binding by coatomer (O’Kelly et al. 2002; Yuan et al. 2003). As 14-3-3 is a large protein family that participate in diverse biological events (Aitken, 1996), an intriguing aspect of this finding is that regulation of cargo sorting by the COPI complex may be coordinated with other cellular events through the actions of 14-3-3.

ARF1 ARF1 is the founding member of the ARF family of small GTPases, which are now well known to act in vesicular transport, actin rearrangement and signaling (D’Souza-Schorey and Chavrier 2006). ARF1 was originally discovered as a co-factor needed for the ADP-ribosylation of cholera toxin (Kahn and Gilman 1984), hence the origin of its name as ADP-ribosylation factor 1.

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However, in contrast to this role for which the physiologic meaning still remains relatively obscure, its subsequently elucidated role in regulating COPI transport has considerably advanced our understanding of how vesicle formation is regulated to initiate vesicular transport (D’Souza-Schorey and Chavrier 2006). Like all small GTPases, guanine nucleotide exchange factors (GEFs) are needed to catalyze the activation of ARF1 (Casanova 2007), while GTPaseactivating proteins (GAPs) are required to catalyze its deactivation (Inoue and Randazzo 2007). As another mode of regulation, ARF1 is myristoylated at its amino terminus, which is critical for its stabilization on target membrane upon its binding to GTP (Franco et al. 1996; Randazzo et al. 1995). ARF1 has been shown to interact with multiple effectors, including coatomer, specific cargo proteins, and multiple lipid-modifying enzymes (D’Souza-Schorey and Chavrier 2006). Moreover, a recently appreciated effector has been its GAP, which has been shown to act as a coat component in COPI vesicle formation (Lee et al. 2005; Yang et al. 2002). Altogether, these findings suggest that, rather than one specific interaction, the coordination of multiple interactions of activated ARF1 results in the stable recruitment of the coat complex onto membrane for COPI vesicle formation.

GEF The first GEFs identified for ARF small GTPases were Gea1p and Gea2p, two homologous yeast proteins (Peyroche et al. 1996). While neither was essential for yeast viability, deletion of both led to lethality (Peyroche et al. 1996). Moreover, similar to perturbation of coatomer subunits, perturbing these GEFs also led to secretion abnormalities and perturbation of the Golgi complex (Peyroche et al. 2001). Thus, these two yeast GEFs are currently thought to act redundantly in activating ARF1 for COPI transport. The first mammalian GEF identified for ARF small GTPases was ARNO, based on its sequence similarity to the yeast Gea1p and Gea2p, particularly within the region responsible for catalysis, known as the Sec7 domain (Chardin et al. 1996). However, ARNO turned out to be insensitive to BFA (Chardin et al. 1996). Moreover, it had a significant distribution on the plasma membrane, where it acted on ARF6 for the regulation of actin rearrangement (Frank et al. 1998a, b; Santy and Casanova 2001). GBF1 was subsequently identified as the more likely GEF that acts on ARF1 in COPI transport. Overexpression of GBF1 conferred resistance to cells treated with BFA (Kawamoto et al. 2002), while expression of a catalytic dead mutant mimicked the effect of BFA in redistributing coatomer from the Golgi (Garcia-Mata et al. 2003). Moreover, a viral protein expressed by picornavirus has recently been shown to inhibit GBF1, resulting in trafficking defects likely attributable to perturbations in COPI transport (Wessels et al. 2006). However, in contrast to all other key components currently known for COPI vesicle formation, the mechanistic details of how GBF1 participates in COPI transport have not been studied using the COPI vesicle reconstitution system.

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GAP The first GAP identified to act on an ARF small GTPase is ARFGAP1 (Cukierman et al. 1995). However, the manner by which this GAP participates in COPI transport has turned out to be surprisingly complex. In early studies, when Golgi membrane was incubated with purified ARF1 and coatomer, introduction of either an ARF1 mutant with impaired GTPase activity (ARF1-Q71L) or the addition of GTPgS to inhibit this activity, led to the formation of coated vesicles that could not undergo uncoating (Tanigawa et al. 1993). Moreover, later COPI vesicle reconstitution studies that involved the incubation of liposomal membrane with purified protein components also concluded that the GAP activity acted in COPI vesicle uncoating (Bigay et al. 2003; Reinhard et al. 2003). Altogether, these findings suggested that the GAP would act simply in destabilizing coatomer on membrane, leading to its mechanistic assignment in COPI vesicle uncoating. A need to revise this view was initially suggested by the finding that the GAP activity promoted COPI cargo sorting (Lanoix et al. 1999; Nickel et al. 1998; Pepperkok et al. 2000). As cargo sorting is tightly coupled to vesicle formation (Springer et al. 1999), early attempts to reconcile the seemingly disparate roles of the GAP activity (in promoting cargo sorting and yet also in inhibiting vesicle formation) focused on how it may be regulated. One notable example was based on the principle of kinetic proofreading, for which distinct members of the p24 family of cargo proteins were shown to exhibit differential regulatory effects on the GAP activity (Goldberg 2000; Lanoix et al. 2001). Moreover, another mechanism was suggested by the observation that the GAP activity was regulated by membrane curvature (Bigay et al. 2003). Recently however, the prevailing view that the GAP activity antagonizes COPI vesicle formation has been challenged by examining the behavior of ARFGAP1 in a refined COPI vesicle reconstitution system that used Golgi membrane rather than the simpler liposomal membrane. Remarkably, reconstituted COPI vesicles were found to be coated with coatomer and ARFGAP1 in stoichiometric levels (Yang et al. 2002). Moreover, ARFGAP1 was found to play a direct role in recruiting coatomer to bind cargo proteins (Lee et al. 2005). Also, its catalytic activity was found to be required for vesicle formation (Lee et al. 2005). Altogether, these findings have led to a new conclusion that, rather than simply acting as a negative regulator of the ARF1 GTPase cycle, the GAP also acts as an effector by being a component of the COPI complex.

Fission factors Besides GAP, the COPI vesicle reconstitution system that uses Golgi membrane has been further refined recently, leading to the identification of brefeldin-A ADP-ribosylated substrate (BARS) and endophilin B to act mechanistically interchangeable for a late step of COPI vesicle formation, known as the fission step.

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BARS BARS has a complex history. Its role in membrane traffic was originally suggested by the discovery that BFA activated an endogenous ADP-ribosylation reaction with BARS identified as one target (Di Girolamo et al. 1995; Spano et al. 1999). With its sequencing, BARS was realized to be a splice variant of C-terminal binding protein 1 (CtBP1) (Spano et al. 1999), which belonged to a protein family previously characterized to act as transcription co-repressors (Chinnadurai 2002). Thus, BARS is also known as CtBP3. Initial mechanistic insight on BARS suggested that it possessed an acyltransferase activity, which was associated with its ability to induce the fission of Golgi tubules into vesicles (Weigert et al. 1999). Subsequently however, this acyltransferase activity was shown not to be intrinsic to BARS (Gallop et al. 2005). In the meantime, when Golgi membrane was washed more stringently in the refined COPI vesicle reconstitution system, the existing known factors in COPI vesicle formation no longer became sufficient, implying additional factor(s) needed. By examining cytosol for a fraction that could complement the existing factors, a recent study has identified BARS (Yang et al. 2005). Characterization of its role revealed that BARS played a critical role during the fission process, which is late stage of vesicle formation when the neck of coated buds underwent constriction for their eventual release as vesicles from compartmental membrane (Yang et al. 2005). Also, a minimal domain of BARS was shown to be sufficient for this role, and this domain did not possess acyltransferase activity (Yang et al. 2005). Thus, even though these recent findings in COPI transport has confirmed a role for BARS in membrane fission, how this action is achieved remains to be determined. How BARS is regulated to act in COPI vesicle formation has also been revealed to be surprisingly complex. For transcription, CtBP members required NAD as a co-factor (Chinnadurai 2002). However, binding to this cofactor inhibited the role of BARS in COPI vesicle formation by preventing its association with ARFGAP1 (Yang et al. 2005). Instead, binding to p-coA by BARS as an alternate co-factor allowed its association with ARFGAP1 for COPI vesicle fission (Yang et al. 2005). Intriguingly, these observations have raised the possibility that transcription and transport, two intracellular events that had not been previously appreciated to have much in common, could be coregulated through the action of BARS.

Endophilin B Endophilin B belongs to a protein family for which endophilin A had been the best characterized member. Endophilin A has been shown to act in the fission of clathrin coated vesicles (with the AP2 adaptor) from the plasma membrane (Ringstad et al. 1999; Schmidt et al. 1999; Simpson et al. 1999). In contrast, endophilin B was initially found to localize at the Golgi but its function had remained unknown (Farsad et al. 2001). A role for endophilin B in COPI transport was recently discovered in an unexpected manner. As background, three forms of CtBP have been found: CtBP1, CtBP2, and CtBP3 (also known as

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BARS). CtBP2 is encoded by one gene, while the other two members are expressed through another gene. Deletion of both genes that encode for all CtBP members in the mouse, while being lethal with respect to the generation of the whole animal, led to viable embryonic cells (Hildebrand and Soriano 2002). This result seemingly contradicted the finding that COPI transport is critical for cell viability (Guo et al. 1994; Hosobuchi et al. 1992). Providing an explanation, a recent study examined embryonic cells derived from mice that had all CtBP members deleted, and found that these cells used endophilin B instead of BARS for COPI vesicle formation (Yang et al. 2006). Remarkably, endophilin B was found also to act in COPI vesicle fission in different adult mouse cell types, in a manner mutually exclusive to that of BARS (Yang et al. 2006). This latter finding is surprising, because all previously identified critical factors in COPI vesicle formation had been shown to play inflexibly roles from yeast to mammals.

COPI transport pathways Since its original discovery, COPI vesicles are now implicated to act in both intra-Golgi transport and also transport from the Golgi to the ER. While the discovery of the latter role was initially considered surprising, it has become more widely accepted in recent years. In contrast, the precise role of COPI vesicles in intra-Golgi transport remains debated. Notably, COPI has also been suggested recently to have roles other than in forming transport vesicles, such as potentially in organizing membrane domains on organellar compartments (Bonifacino and Lippincott-Schwartz 2003). However, as this proposed role has been relatively uncharacterized, we will focus instead on its intensely investigated roles in intracellular transport pathways.

Intra-Golgi transport The intra-Golgi transport assay originally designed to monitor the transfer of VSVG between Golgi cisternae (Balch et al. 1984). Thus, as VSVG is transported anterograde through the secretory system to reach the plasma membrane, the simplest explanation at the time was that COPI vesicles functioned in anterograde transport. However, a series of subsequent findings have led to major changes in this view. Initially, coatomer was found to bind to the cytoplasmic domain of cargo proteins that contained a di-lysine-based motif (Cosson and Letourneur 1994), with functional evidence suggesting that this binding led to the retrograde transport of di-lysine-containing cargo proteins from the Golgi to the ER (Letourneur et al. 1994). Other yeast studies at this time also revealed that perturbation of coatomer subunits perturbed anterograde transport between the ER and the Golgi by affecting more directly the retrograde arm of bidirectional transport that connected these two organelles (Gaynor and Emr 1997; Lewis and Pelham 1996). A role for COPI in retrograde transport was further suggested subsequently by the realization that cisternal maturation played a predominant role in

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anterograde intra-Golgi transport (Bonfanti et al. 1998; Mironov et al. 2001). In this model, rather than being static membrane compartments, which was the prevailing view at the time, Golgi cisternae dynamically transformed themselves by earlier stacks becoming later stacks. This dynamic transformation provided the basis by which anterograde intra-Golgi transport was accomplished. A key implication of this model was that retrograde transport must also exist, as markers of Golgi cisternae (which were typically transmembrane Golgi glycosylation enzymes) were noted to remain in their distinct distribution in the face of anterograde transport by cisternal maturation. Thus, COPI vesicles were scrutinized for a potential role in retrograde intraGolgi transport. For these studies, two complementary approaches were taken: (i) a biochemical approach that examined the content of COPI vesicles reconstituted from Golgi membrane, and (ii) a morphologic approach that visualized COPI vesicles and their content by immunogold electron microscopy. The biochemical approach initially found that Golgi enzymes were concentrated in COPI vesicles, and thus, concluding that these vesicles played a role in retrograde intra-Golgi transport. Specifically, in an initial re-visit of the intra-Golgi transport assay, Golgi enzymes were found to be transferred among Golgi stacks in a COPI-dependent manner (Love et al. 1998). Moreover, COPI vesicles reconstituted in the presence of GTPgS that blocked ARF1 deactivation were found to have reduced cargo sorting (Lanoix et al. 1999; Nickel et al. 1998; Pepperkok et al. 2000). When vesicles were generated instead by incubating Golgi membrane with cytosol in the presence of GTP, Golgi enzymes, but not examples of anterograde cargo protein, were detected to be concentrated in the reconstituted vesicles (Lanoix et al. 1999). However, these biochemical studies could be criticized that their conclusion rested on indirect evidence. As cytosol contained factor(s) that prevented the stabilization of COPI on vesicular membrane, these studies examined vesicles that did not have COPI coating. Instead, these vesicles were suggested to have been coated by COPI and then having undergone uncoating, because their generation was inhibited when coatomer was depleted from cytosol (Lanoix et al. 1999). In the other main approach, immunogold EM studies were undertaken. However, these studies led to conflicting results, with some studies concluding that Golgi enzymes were not transported by COPI vesicles (Cosson et al. 2002; Orci et al. 2000a), while others concluding the opposite (Martinez-Menarguez et al. 2001). A limitation of the traditional EM approach that involves thinsectioning has been the ability to distinguish vesicles from buds and tubules with certainty. To overcome this hurdle, EM tomography has been used more recently. These studies concluded that Golgi enzymes were not significantly concentrated in COPI vesicles (Kweon et al. 2004). Instead, an intriguing possibility was raised that Golgi enzymes could be transported retrograde through tubular connections among the Golgi stacks (Trucco et al. 2004). A recent review has summarized the conflicting conclusions regarding the role

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of COPI in retrograde intra-Golgi transport, suggesting potential methodologic and interpretational differences (Rabouille and Klumperman 2005). As it currently stands however, whether COPI vesicles plays a significant role in this transport remains disputed. Even more uncertain has been whether COPI vesicles participate in anterograde intra-Golgi transport, its originally proposed role. As alluded to above, early evidence for this role came from the cell-free reconstitution of intra-Golgi transport that monitored the transfer VSVG, which is an anterograde cargo protein (Balch et al. 1984). However, later studies suggested experimental caveats for this observation. First, because the relative fraction of VSVG detected in reconstituted COPI vesicles was low as compared to host proteins, a suggestion was made that the detection of VSVG in reconstituted COPI vesicles represented their mis-sorting due to viral-mediated overexpression of this cargo protein (Love et al. 1998). Second, GTPgS that was used to reconstitute COPI vesicles in the original studies was subsequently revealed to prevent the proper sorting of cargo proteins (Lanoix et al. 1999; Nickel et al. 1998; Pepperkok et al. 2000). When vesicles that were reconstituted in the presence of GTP and shown to be dependent on COPI, anterograde cargo proteins were not enriched in these vesicles (Lanoix et al. 1999; Love et al. 1998). Finally, EM studies have led to conflicting conclusions regarding whether anterograde cargo proteins are concentrated in COPI vesicles (Martinez-Menarguez et al. 2001; Mironov et al. 2001; Orci et al. 2000b, 1997). Recent studies have focused on the possibility that subpopulations of COPI vesicles may exist, for which one potential role would be in anterograde intraGolgi transport. Variants of different subunits of coatomer have been identified (Wegmann et al. 2004), and shown to have similar but non-overlapping distributions at the Golgi (Moelleken et al. 2007). These findings raise the specter that they may form distinct subpopulations of COPI vesicles. Moreover, another recent study has found that COPI vesicles could be distinguished based on their association with different tethering complexes and also in cargo content (Malsam et al. 2005). However, COPI vesicles reconstituted in this study did not include ARFGAP1 as a purified component. Thus, as GAP activity plays an important role in proper cargo sorting (Lanoix et al. 1999; Nickel et al. 1998; Pepperkok et al. 2000), whether subpopulations of COPI vesicles exist remains an interesting possibility that will require additional supporting evidence in the future.

Retrograde Golgi-to-ER transport Early studies on some chaperones that functioned in protein folding and assembly in the ER suggested that a fraction of these proteins leaked from the ER, and was then retrieved by retrograde transport from the Golgi (Munro and Pelham 1987). Subsequent studies revealed the molecular basis of this retrieval, which identified a receptor that recognized a common KDEL motif on the leaked ER proteins. Thus, the receptor was named the KDEL receptor (KDELR) (Lewis and Pelham 1990; Lewis et al. 1990; Semenza et al. 1990).

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Subsequent studies on the KDELR revealed that it was retrieved from the Golgi to the ER through COPI transport (Girod et al. 1999; Lewis and Pelham 1996; White et al. 1999). Moreover, the KDELR had a more complex role than simply being a passive passenger in COPI vesicles, as it was also found to play a critical role in recruiting a GAP that acts on ARF1 to Golgi membrane in regulating COPI transport (Aoe et al. 1997, 1998). Altogether, these findings not only showed that the KDELR was a COPI cargo protein in retrograde transport from the Golgi to the ER, but also revealed that it was a special cargo protein that regulated its pathway of transport. Another cargo protein that has been well-established to be transported retrograde from the Golgi to the ER by COPI is ERGIC-53. It was originally identified as a marker of the intermediate compartment (Saraste et al. 1987; Schweizer et al. 1988), and hence the basis for its name as ER–Golgiintermediate compartment—53 kDa. Early studies on this cargo protein using temperature blocks revealed its cycling between the ER and the Golgi (Lippincott-Schwartz et al. 1990). Subsequently, the luminal domain of ERGIC-53 was revealed to act as a lectin that recognized a subset of glycosylated soluble luminal proteins, which was critical for these soluble proteins to be transported through the early secretory system to reach peripheral lysosome-like organelles (Appenzeller et al. 1999). Moreover, the physiologic relevance of this binding was revealed by mutations in ERGIC-53 causing human coagulopathies (Nichols et al. 1998). The cycling of ERGIC-53 between the ER and the Golgi has been shown to involve COPII recognizing a diphenylalanine motif in the cytoplasmic domain of ERGIC-53 for its anterograde transport (Kappeler et al. 1997), and COPI recognizing di-lysine motif for its retrograde transport (Kappeler et al. 1997; Tisdale et al. 1997). Like ERGIC-53, the p24 family of transmembrane proteins also cycle between the ER and the Golgi complex. Moreover, a phenylalanine-based motif in their cytoplasmic domain is recognized by COPII and a di-lysine-based motif is recognized by COPI (Dominguez et al. 1998). However, their precise cellular roles remain debated. The p24 family was originally identified based on their abundance in purified COPI vesicles (Sohn et al. 1996; Stamnes et al. 1995). As such, they were proposed to be critical for the binding of COPI to membrane for its role in forming transport vesicles. Studies on one member in particular, known as p23, supported this contention (Bremser et al. 1999; Reinhard et al. 1999). Remarkably however, when all p24 family members were deleted in the yeast, a surprisingly mild phenotype was observed (Springer et al. 2000). Thus, as COPI is essential for cell viability from yeast to mammals (Guo et al. 1994; Hosobuchi et al. 1992), an essential role for p24 family members in COPI vesicle formation has been questioned. Instead, other evidence suggests that the luminal domain of p24 members binds to different ligands to mediate their exit from the ER (Muniz et al. 2000). In this light, p24 is essentially acting as a receptor that facilitates the exit of select cargo proteins from the ER, which is fundamentally similar to the role currently attributed to ERGIC-53.

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Studies on SNAREs (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) have revealed yet another class of cargo proteins for which retrograde transport from the Golgi to the ER by COPI plays a critical role. SNAREs mediate the fusion of transport vesicles with their compartment of destination (Jahn and Scheller 2006). Studies on Ufe1p, a yeast syntaxin, revealed that its distribution at the ER depended on COPI (Lewis and Pelham, 1996). Moreover, yeasts with mutations in either Ufe1p or subunits of coatomer had defective anterograde transport between the ER and the Golgi, which was deduced to be the result of a more direct defect in retrograde COPI transport (Lewis and Pelham 1996).

Perspective Studies on COPI transport have made fundamental contributions to our understanding of how coat proteins are regulated by the ARF family of small GTPases during vesicle formation. However, methodologic issues have led to controversies regarding the precise function of COPI vesicles and how they are formed. Notably, recent mechanistic insights on COPI vesicle formation have revealed that the GAP which deactivates ARF1 has surprisingly complex roles. Rather than simply acting as an upstream regulator of the GTPase cycle, the GAP also acts as a key downstream effector, by being a component of the COPI complex. Thus, it is now appreciated to be intimately involved in vesicle formation, rather than in vesicle uncoating as originally postulated (Fig. 3)

Figure 3. A revised model for the role of the GAP that acts on ARF1 in COPI transport. Rather than simply acting an upstream regulator of the GTPase cycle, ARFGAP1 also acts as a key downstream effector, by being a component of the COPI complex. A key implication of this new model is that COPI vesicle uncoating will involve factor(s) yet to be identified.

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This shift in our mechanistic understanding of COPI vesicle formation has three key implications. First, it has allowed further mechanistic insights into COPI vesicle formation, including the identification of novel critical factors. Second, it predicts that, rather than the GAP that acts on ARF1, COPI vesicle uncoating will be more directly regulated by other factor(s). Third, it simplifies models to explain how COPI vesicle formation and cargo sorting are coupled. In this last case, an intriguing prospect is that further studies on mechanisms of cargo sorting may eventually result in a more precise understanding of transport roles by COPI, for which some are still currently debated. Acknowledgements. We thank Jian Li for helpful discussions. This work is funded by a grant from the NIH to VWH. We apologize to our colleagues for not having cited all their work related to COPI due to space constraint.

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mediated COP-I assembly by inhibiting the guanine nucleotide exchange factor GBF1. Dev Cell 11: 191–201 White J, Johannes L, Mallard F, Girod A, Grill S, Reinsch S, Keller P, Tzschaschel B, Echard A, Goud B, Stelzer EH (1999) Rab6 coordinates a novel Golgi to ER retrograde transport pathway in live cells. J Cell Biol 147: 743–760 Yang J, Lee S, Spanò S, Gad H, Zhang L, Nie Z, Bonazzi M, Corda D, Luini A, Hsu V (2005) A role for BARS at the fission step of COPI vesicle formation from Golgi membrane. EMBO J 24: 4133–4143 Yang JS, Lee SY, Gao M, Bourgoin S, Randazzo PA, Premont RT, Hsu VW (2002) ARFGAP1 promotes the formation of COPI vesicles, suggesting function as a component of the coat. J Cell Biol 159: 69–78 Yang JS, Zhang L, Lee SY, Gad H, Luini A, Hsu VW (2006) Key components of the fission machinery are interchangeable. Nat Cell Biol 8: 1376–1382 Yuan H, Michelsen K, Schwappach B (2003) 14-3-3 dimers probe the assembly status of multimeric membrane proteins. Curr Biol 13: 638–646 Zerangue N, Malan MJ, Fried SR, Dazin PF, Jan YN, Jan LY, Schwappach B (2001) Analysis of endoplasmic reticulum trafficking signals by combinatorial screening in mammalian cells. Proc Natl Acad Sci USA 98: 2431–2436

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Arfs and Arls: models for Arf family members in membrane traffic at the Golgi Richard A. Kahn

Introduction The ADP-ribosylation factor (Arf) family GTPases are highly conserved 20 kDa GTP-binding proteins that play a number of roles in the regulation of cellular physiology, most relevant here is that they act to recruit soluble proteins to a membrane surface and coordinate the assembly of multi-protein complexes that are required for the biogenesis of nascent carriers of membrane traffic. The Arfs can also recruit and directly activate lipid-modifying enzymes, providing important functional links between localized changes in lipid composition and protein assemblies. The Arf family GTPases and their interactions have been the subject of a recent book (Kahn 2004) and reviews (Gillingham and Munro 2007; Inoue and Randazzo 2007). Molecular aspects of Arf family members acting at the Golgi and models for their actions are summarized in this chapter.

The Arf family of regulatory GTPases and their actions at the Golgi The Arf family includes both the Arf sub-family, with six members in mammals (Arf1–6) that share >60% primary sequence identity, and the more divergent Arf-like (Arl; typically 40–60% identity to Arfs or each other), and Sar (20–30% identical to Arfs) sub-families (Kahn et al. 2006; Li et al. 2004; Logsdon and Kahn 2004). This is an ancient gene family with already six members in the earliest eukaryotes, prior to the emergence of Ras or heterotrimeric G proteins, and likely arising in prokaryotes (Dong et al. 2007). Members of each of these sub-families play important roles in membrane traffic, while several Arls have distinct roles in the regulation of microtubule dynamics (Arl2 and Arl8 (Antoshechkin and Han 2002; Bhamidipati et al. 2000; Hoyt et al. 1990; McElver et al. 2000; Okai et al. 2004; Zhou et al. 2006)), ciliogenesis (Arl3, Arl6, and Arl13b (Caspary et al. 2007; Chiang et al. 2004; Fan et al. 2004; Sahin et al. 2004; Schrick et al. 2006; Zhou et al. 2006)), and cytokinesis (Arl3, Arf1, and Arf6 (Altan-Bonnet et al. 2003; Schweitzer and D’Souza-Schorey 2005; Zhou et al. 2006)). Sar1 (two genes/proteins in mammals) is the most divergent member of the Arf family yet acts to regulate carrier biogenesis at ER exit sites in a fashion that is very analogous to the actions of other family members at the Golgi (Barlowe et al. 1994; Kuge et al. 1994; Lee et al. 2005; Nakano and

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Muramatsu 1989; Oka et al. 1991). In many ways the mechanism of action of Sar1 in regulating exit of cargo from the ER and recruitment of the COPII complex is a paradigm for the actions of Arfs at the Golgi, though the latter have additional complexities as a result of the increased (a) diversity in highly homologous proteins, (b) number of coat complexes involved, and (c) role of lipid/membrane interactions, as summarized above. Ten of the thirty members of the mammalian Arf family are implicated as regulators of membrane traffic at the Golgi; Arf1–5, Arl1, and Arl5A–C, and Arfrp1. Arf6 is active predominantly in endocytosis at the plasma membrane and also impacts the actin cytoskeleton so is not included in this list. Our current models for Arf and Arl actions at the Golgi come from studies of Arf1–5, (though overwhelmingly only Arf1) and Arl1 so will form the focus of the rest of this chapter. Arf2 has been lost in humans and no specific functional studies have been reported for this isoform so will not be discussed further. Arl5s (three genes/proteins in mammals) localize to the Golgi and either knockdowns by siRNA or expression of dominant mutants have only quite subtle effects on Golgi morphology (Yawei Li and R. A. Kahn; unpublished observations), but no further information is available on their actions. Arfrp1 is discussed with Arl1 as they are genetically and functionally linked. With 10 proteins all active at the Golgi, where is the specificity in Arfregulated membrane traffic found? With clear differences in genetic and physical interactions it was relatively easy to distinguish functions of Arfs from those of Arl1 or Arfrp1, but this has not been true for Arf1–5. The high degree of conservation of primary sequence of the Arfs throughout eukaryotic evolution (human and S. cerevisiae Arf1 are 74% identical) is accompanied by functional conservation. For example, each of the five human Arfs (Arf1, Arf3–6) or the single Arf gene from Giardia lamblia can complement the lethality resulting from deletion of the two yeast Arf genes, ARF1 and ARF2 (Kahn et al. 1991; Lee et al. 1992), but neither the S. cerevisiae or other ARL1 genes can complement the deletion of the ARFs. In addition, in several in vitro assays of Arf activities the specific activities of the different human Arf proteins are indistinguishable. We still lack reagents that can distinguish between the Arf proteins in cells because the isoform specific antisera generated to date do not work for immunofluorescence and epitope tagging is fraught with artifacts resulting from (often poorly characterized) alterations in protein localization and affinities for protein- and lipid-binding partners. Thus, truly specific roles for any Arf1–5 have not been described, while the actions of Arfs and Arl1 are clearly distinct and will be discussed separately. In stark contrast to our failure to generate useful antibodies for immunocytochemistry and the difficulties in interpretation from the use of dominant mutants of such a highly conserved protein family, the use of siRNA to specifically deplete cells of each Arf, alone or in combination, has provided an opportunity to address the question of specificities within the Arf proteins. This approach has led to the identification of a number of distinctive roles for

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Arf1–5 in ER–Golgi traffic, exit from the Golgi, and in endosome recycling (Volpicelli-Daley et al. 2005). Surprisingly, the most profound phenotypes were only observed with different combinations of depletion of two Arfs. While these data may be interpreted in different ways, one intriguing model to arise from them is that Arfs act in pairs to regulate carrier biogenesis at a donor membrane. Indeed, a growing number of the proteins involved in membrane traffic are being found to dimerize on the membrane surface and it is predicted that this will be a central theme in molecular models in the future (e.g., see Burguete et al. (2008)).

Arfs bind lipids and lipid-modifying enzymes Arfs act to control aspects of carrier biogenesis at the Golgi, trans-Golgi network (TGN), ER–Golgi intermediate compartment (ERGIC), and likely endosomes. As regulatory GTPases, Arfs cycle between GDP- and GTP-bound conformations in a cycle at rates that are determined by binding to guanine nucleotide exchange factors (GEFs), GTPase activating proteins (GAPs), and effectors. In addition to these functionally critical protein interactions, Arfs are unusual among regulatory GTPases in that their affinity for biological membranes is directly impacted by the nucleotide bound. Specifically, Arfs are N-myristoylated and this hydrophobic anchor functions as a “myristoyl switch” to dock and orient the protein on the surface of the membrane when the Arf is GTP-bound, but the protein binds the myristate and shields it from aqueous interactions to promote its solubility when “inactive”, i.e., when GDP is bound (Franco et al. 1996; Kahn et al. 1988, 1992; Ames et al. 1997; Goldberg 1998). Thus, activation of Arf (i.e., GTP-binding) promotes a more stable membrane attachment as well as increasing its affinity for effectors. This myristoylation-dependent membrane translocation is predominantly nonspecific as far as lipid interactions. In contrast, some specific phospholipids are capable of altering the guanine nucleotide-binding properties of Arfs and some Arf GAPs (Brown et al. 1998; Kam et al. 2000; Randazzo 1997; Randazzo and Kahn 1994; Terui et al. 1994; Zheng et al. 1996). In a related fashion, some Arf GAPs appear capable of sensing membrane curvature and become more active as the curvature increases, providing a link between carrier maturation and Arf inactivation (Bigay et al. 2005, 2003). Together these data highlight the intimate interplay between Arfs and the membranes at which they act to regulate membrane traffic. In addition to Arfs and Arf GAPs being capable of sensing localized changes in the lipid environment, Arfs also have roles in both recruitment to a membrane and direct activation of a number of lipid-modifying enzyme. Activated Arfs increase the recruitment to membranes of PI 4-kinase and PI(4P) 5-kinase (De Matteis et al. 2005; Godi et al. 1999; Honda et al. 1999; Jones et al. 2000) and increase their activity. Arfs are also potent and direct activators of phospholipase D (Brown et al. 1993; Cockcroft et al. 1994; Ktistakis et al. 1995, 1996). The intimate co-dependence of lipid

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modifications and protein recruitment to Arf action and membrane traffic at the Golgi is evident from studies of the FAPP and related proteins, that have both Arf-binding and PI4P-binding domains, each of which are required for their action in export from the TGN (Godi et al. 2004; Vieira et al. 2005). Molecular details of how the lipid-modifying activities of Arfs and adaptor recruitment are integrated in the cell and the respective contributions of each activity at different sites remain controversial. But there is no disagreement among researchers that our understanding of molecular membrane traffic will require detailed understanding of the localized changes in the lipid bilayer, membrane curvature, protein recruitment, and enzyme activities; thus assuring that Arfs are key players in this complex and essential aspect of cell biology.

Activated Arfs recruit protein adaptors to nascent carriers of membrane traffic – models for Arf action at the Golgi/TGN Three observations from the late 1980’s and early 1990’s were central to the current models of the actions of Arfs at the Golgi. First was that the Arfs are GTPases that act at the Golgi to regulate aspects of membrane traffic (Kahn and Gilman 1986; Kahn et al. 1992; Stearns et al. 1990). The GTPase field that the Arfs arose from was defined by the plasma membrane localized, heterotrimeric G proteins, and thus the models involved initiation of GTPase signaling in response to agonist binding to a specific receptor, that has GEF activity. No such agonist has been defined for Arfs and thus the initiator of Arf signaling remains uncertain. While Arfs clearly operate through the canonical GTP/GDP cycle for regulatory GTPases, it is not necessarily true that agoniststimulated G protein signaling is the best molecular model for Arfs as regulators of membrane traffic. Second was the observation that Arfs were required to recruit the COPI coat to Golgi membranes in an in vitro assay of intra-Golgi transport (Serafini et al. 1991). Although several controversies grew out of this early work, some of which remain unresolved, it has become clear that a central role for Arfs as regulators of membrane traffic is the recruitment to membranes of protein adaptors or adaptor complexes. This is true not only for the heptameric COPI complex, but also the tetrameric adaptin complexes (AP-1 and AP-4 at the TGN (Dell’Angelica et al. 1999; Hirst et al. 1999; Stamnes and Rothman 1993; Traub et al. 1993), AP-2 at the plasma membrane and endosomes (Krauss et al. 2003; Paleotti et al. 2005; West et al. 1997), and AP-3 at endosomes (Drake et al. 2000; Faundez et al. 1998; Faundez and Kelly 2000; Ooi et al. 1998)), and monomeric GGA (Boman et al. 2000; Dell’Angelica et al. 2000; Hirst et al. 2001; Puertollano et al. 2001a, b; Takatsu et al. 2001) and Mint (Hill et al. 2003; Shrivastava-Ranjan et al. 2008) families (which each contain three different genes/proteins).

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The third landmark observation was that the macrolide antibiotic brefeldin A (BFA) was found to be a rapidly acting, membrane permeant, specific inhibitor of a subset of Arf GEFs that act at the Golgi/TGN (Donaldson et al. 1991, 1992; Fujiwara et al. 1988; Lippincott-Schwartz et al. 1989; Misumi et al. 1986; Robinson and Kreis 1992). Arf GEFs and their sensitivity to BFA have been the subject of an excellent recent review so will not be discussed in detail here (Casanova 2007). Use of BFA has allowed live cell imaging of Arfdependent processes and the emergence of one imporant criterion for an Arf-dependent adaptor at the Golgi – rapid release (10 mM (Pezzati et al. 1997). However, targeting aequorin to the GC to measure the [Ca2þ]GC of the Golgi cisternae has demonstrated that in the cisternal lumen of unstimulated cells the free [Ca2þ] is around 0.3 mM; therefore, a gradient in [Ca2þ] exists between the lumen of the Golgi and the cytosol (Pinton et al. 1998). These storage compartments are also equipped with Ca2þ-release channels: the inositol 1,4,5-trisphosphate (IP3) receptor and/or the ryanodine receptor (RyR, Dolman and Tepikin 2006). Ca2þ release from GC and subsequent reuptake are faster than in the ER (Missiaen et al. 2004a,b). Thus, cells can actively maintain a [Ca2þ]i that is some 40,000-fold lower than that outside of the cell, and some 14,000-fold lower than that in the lumen of endomembranes. To maintain such low [Ca2þ]i will also expend a lot of energy.

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Figure 1. Scheme for the role of Ca2þ at different transport steps. The arrows indicate the direction of the delivery of membrane carriers. The Ca2þ indications show where Ca2þ is important in the processes involved in the fusion of these carriers with their target compartments. The bold Ca2þ indicate the need for an increased [Ca2þ] for membrane fusion at the different stages of intracellular membrane transport. These include the following fusion steps: between the cargo domain in the GC and the TGN (arrow 2 and double-directed arrow near the Golgi-to-PM carrier [GPC]); between the Golgi-to-basolateral PM and Golgi-to-apical PM carriers and basolateral PM (arrow 3) and apical PM (arrow 6), correspondingly; between the Golgi-to-endosome carrier (GEC) and endosomes (E) (arrow 4); between the GPC ferrying albumin and other soluble cargoes to the basolateral PM (arrow 5); between apically directed mature secretory granules (AMSG) and apical PM (arrow 7), between different endosomes (double directed arrow near E) and between endosomes and lysosomes (not shown). The indications of grey Ca2þ show where even basal [Ca2þ]i in quiescent cells (50–100 nM) are sufficient for membrane fusion: between the ER-to-Golgi carrier and the Golgi cisternae (arrow 1), and between COPI-dependent vesicles and the Golgi cisternae (double directed arrow near “cis”). The role of Ca2þ for the fusion of the putative clathrin/AP-2-dependent vesicles and endosomes is not known (arrow 8, indicated as no Ca2þ). Dark squares show the border between the apical PM and the basolateral PM. Further abbreviations: AISG, apically directed immature secretory granules; AGPC, precursor of apically directed Golgi-to-PM carrier; APM, apical PM; AT, attached TGN; BLPM, basolateral PM; CCB, clathrin-coated buds; ER, endoplasmic reticulum; ERES, ER exit site; Nu, nucleus; PM, plasma membrane; TGN, trans-Golgi network.

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Ca2þ pumps Ca2þ ATPase pumps use a lot of energy to maintain the gradients of [Ca2þ] between the cytosol and the lumen of the ER and the GC. Golgi membranes contain two types of Ca2þ ATPase pumps that contribute to Ca2þ uptake in the GC. One of these belongs to the thapsigargin-sensitive, sarcoendoplasmic reticulum calcium ATPase (SERCA) family, that sequester Ca2þ mainly in the ER, and to a lesser extent in Golgi compartments. The other ATPase pump belongs to the thapsigargin-insensitive, secretory pathway calcium ATPase (SPCA) family, which are present on the medial-trans region of the GC (Behne et al. 2003). All of the Ca2þ uptake by the ER is mediated by the SERCA Ca2þ pumps (Vanoevelen et al. 2004). Various relative contributions of the SPCA and SERCA Ca2þ pumps to the total Ca2þ uptake in the GC have been reported, which could be due to celltype dependency. For instance, according to Rojas et al. (2000), Ca2þ uptake in a Golgi-enriched fraction of rat liver depended totally on a SERCA Ca2þ pump, since it was almost completely inhibited by thapsigargin. In contrast, Taylor et al. (1997) showed only 50% inhibition by thapsigargin of Ca2þ uptake into a stacked Golgi fraction from rat liver. A thapsigargin-independent Ca2þ uptake has also been ascribed to PMCA Ca2þ pumps in transit through the GC to the PM. This thapsigargin-resistant Ca2þ uptake disappears when SPCA1 expression is disrupted using RNA interference (Van Baelen et al. 2003). Using aequorin to measure Ca2þ uptake, it has been demonstrated that SERCA Ca2þ pumps are responsible for 50–85% of Ca2þ uptake in the Golgi compartment of HeLa cells (Pinton et al. 1998; Van Baelen et al. 2003). In HeLa and CHO cells overexpressing the Ca2þ-binding protein CALNUC, about 70% of the Ca2þ uptake by the GC depends on the SERCA pumps (Lin et al. 1999). In contrast, it has been reported that the SPCA1 Ca2þ pump is mainly used (67%) to load the GC with Ca2þ in human keratinocytes (Callewaert et al. 2003). Whereas the SERCA pumps are expressed in both the ER and the GC, the SPCAs appear to be more specifically confined to the latter compartments of the secretory pathway, i.e. the Golgi stack, the TGN, and maybe secretory granules. The cis-Golgi region appears to express SERCA and IP3 receptors (Missiaen et al. 2004a,b; Vanoevelen et al. 2004), while the trans-Golgi region appears to contain SPCA1 and to lack IP3 receptors (Missiaen et al. 2004a,b). Thus, SPCA1 is responsible for the uptake of Ca2þ into the trans area of the GC (Michelangeli et al. 2005). Two isoforms of the SPCA Ca2þ pump, known as SPCA1 and SPCA2, have so far been identified, although only SPCA1 has been shown to be active. Unlike SERCA, the SPCA1 Ca2þ pump can transport Mn2þ in addition to Ca2þ (Van Baelen et al. 2001; Missiaen et al. 2004a,b). SPCA2 is present in mammalian cells, although its level of expression and distribution in tissues remains controversial (Vanoevelen et al. 2005; Xiang et al. 2005) and it is not yet well characterized. However, it is known to have similar functions to SPCA1, which

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include of the same capacity for transporting Mn2þ and Ca2þ in a thapsigargin-insensitive way, and the same GC localization. Over-expression of SPCA can cause significant alterations in several of the Ca2þ transport molecules in the cell and dramatically increased the celldivision rate (Reinhardt et al. 2004). In Darier Disease, where SERCA2 function is impaired, the overexpression of SPCA1 might compensate for this SERCA2 dysfunction, suggesting a role in changes of resting [Ca2þ]i (Foggia et al. 2005). Thus, the SPCAs have different crucial roles: (a) they contribute in a significant manner to luminal Ca2þ uptake; (b) they are responsible for the cytosolic regulation and storage in the GC of Mn2þ, promoting the correct functioning of the few Golgi enzymes that require Mn2þ as a cofactor. The ATP2C1 gene, encoding for the SPCA1 in mammals cells, have an homolog in yeast and Caenorhabditis elegans. In fact, the PMR1 gene product was shown to be a Ca2þ-ATPase that is located in the GC (Sorin et al. 1997; Van Baelen et al. 2001). For the latter, these are seen as type IIA (ER-type) Ca2þ-ATPases, and type IIB (PM-type) Ca2þ-ATPases (Sze et al. 2000). The mechanisms by which Ca2þ is transported into the GC and maintained at high levels has not been completely defined yet. Ca2þ accumulation in isolated Golgi membranes varies according to both Ca2þ and Mg-ATP concentrations, and it can be inhibited by thapsigargin, but not stimulated by calmodulin (Rojas et al. 2000). Thapsigargin-independent Ca2þ accumulation was not affected by pre-treatment with agents such as NH4Cl or chloroquine, which collapse the H+ gradient across cell membranes (Pinton et al. 1998). Thus, there are two main Ca2þ pumps (SERCA and SPCA) that are responsible for the accumulation of Ca2þ in the ER and the GC.

Protein buffers for Ca2þ in the lumen of the GC To save on the energy that is needed to maintain such high gradients of Ca2þ between these intracellular Ca2þ stores and the cytosol, cells express specific Ca2þ-binding proteins, which can sequester most of the free Ca2þ in these stores. In the ER, there are several Ca2þ-binding proteins, including calreticulin (Michalak et al. 1999), calnexin (Ohsako et al. 1994), reticulocalbin, calumenin and ERC-55 (Vorum et al. 1999). Some of these, although not calumenin, have also been found in the lumen of the Golgi cisternae (Vorum et al. 1999). However, the Ca2þ sequestered in the Golgi lumen is mainly buffered by CALNUC (nucleobindin; Lin et al. 1998, 1999; Kawano et al. 2000), Cab45, the first resident protein that was described for the Golgi lumen (Scherer et al.  and Vorum 2000), and P54/NEFA (Morel-Haux et al. 2002). 1996; Honore CALNUC has also been localized to both the ER and the Golgi lumen (Lin et al. 1998, 1999).

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When the influx of Ca2þ into the cytosol is increased chronically, cells can express more of the Golgi calcium-binding protein CALNUC in order to prevent calcium cytotoxicity. Additionally, the overexpression of SPCA1 increases CALNUC levels (Rejnhardt et al. 2000). On the other hand, over expression of CALNUC leads to an enhancement of agonist-evoked calcium release (Lin et al. 1999). Thus, cells have developed specific molecular mechanisms to maintain these Ca2þ gradients between the Ca2þ stores and the cell cytosol.

Ca2þ release What is the role of these Ca2þ gradients and what are the mechanisms regulating the local [Ca2þ]i, during membrane transport? These mechanisms primarily involve Ca2þ channels. It appears that during the synchronous passage of cargo through the GC, the local [Ca2þ]i increases due to the activation of Ca2þ channels. Ca2þ is released from either the Golgi cisternae per se or from the trans ER (see Chapter 1), which is closely attached to the GC (Micaroni et al. submitted). Both the ER and the SERCA-expressing part of the GC are involved in the setting up of [Ca2þ]i signals. The release of Ca2þ from the ER can be triggered not only by the activation of IP3 and ryanodine receptors (Pinton et al. 1998) but also increase in [Ca2þ]i can themselves promote further Ca2þ release from these intracellular Ca2þ stores via the direct binding of Ca2þ to IP3 and RyR channels (Roderick et al. 2003). The GC has been shown to act in concert with the ER, albeit with different kinetics, in the elevation of [Ca2þ]i in response to agonist stimulation (Missiaen et al. 2004a,b). IP3 receptors have been immunolocalized in the GC (Lin et al. 1999), and agonist stimulation of the production of IP3 can activate these channels on the Golgi membranes, resulting in Ca2þ release from the GC. All of the 0.3 mM of free Ca2þ inside the Golgi lumen can be released upon stimulation with agonists coupled to IP3 production; for example, addition of histamine to cells, which is an agonist coupled to IP3 production, results in a rapid and extensive drop in the free Ca2þ in the GC. However, this drop in [Ca2þ]GC caused by histamine has been shown to be slightly smaller and slower than that observed in the ER (Pinton et al. 1998; Vanoevelen et al. 2004). Ca2þ release from the GC is also inactivated faster than that from the ER (Michelangeli et al. 2005), and it has been shown that the SPCA1-based part of the GC does not contribute to these changes in [Ca2þ]i (Vanoevelen et al. 2004; Missiaen et al. 2004a,b). This release of Ca2þ from the GC does not depend on the functions of the ARF/COPI machinery. Indeed, in digitonin-permeabilized cells, the release and uptake of Ca2þ from the GC was not affected by GTPgS, with neither the loading of Ca2þ into the GC nor the rapid emptying of the GC in thapsigargintreated cells (see above) is affected (Pinton et al. 1998).

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Thus, both the ER and the GC participate in local increases in [Ca2þ]i, which also facilitates vectorial membrane transport. Upon stimulation of a receptor coupled to IP3 production, not only does the GC contribute in part to the increased [Ca2þ]i, but there are also major changes in [Ca2þ]GC (Pinton et al. 1998). During intra-Golgi transport, the [Ca2þ]i local to the GC increases, with this increase not occurring immediately after the arrival of cargo at the GC, but a little time (5–7 min) later (Micaroni et al. submitted), which corresponds to the need for fusion between the cargo domain and the TGN (Mironov et al. 2005). Increases in the local [Ca2þ]i at the trans side of the GC can thus facilitate the transfer of a cargo domain from the cis to the trans side of the GC. A possible scheme of this process is shown in Fig. 2.

The positioning of the Ca2þ source The position of these Ca2þ stores is indeed very important for the regulation of intracellular transport (Dolman et al. 2005; Dolman and Tepikin 2006). For instance, in polarized cells, the positioning of the GC is such that both mitochondria and GC are segregated from the lateral regions of the PM, the nucleus and the basal part of the cytoplasm. Here, the ER and nucleus are located in the basolateral part of the cell, whereas the secretory granules are located at the apical pole (Gerasimenko et al. 2002). The GC is therefore positioned between the main Ca2þ release sites in the apical region of the cell and the important Ca2þ sink formed by the perigranular mitochondria. During acetylcholine-induced [Ca2þ]i signalling in the apical region, large Ca2þ gradients can form over the GC because the GC is sandwiched between the Ca2þ source (release sites in the apical region) and the Ca2þ sink (mitochondrial uniporters), Ca2þ gradients are formed with higher [Ca2þ]i over the trans-Golgi than over the cis-Golgi (Dolman et al. 2005). When low doses of acetylcholine were given apically, the Ca2þ gradient had almost completely dissipated at a distance of 2 mm from the GC, which was the region of the cell occupied by the perigranular mitochondrial belt (Dolman and Tepikin 2006). These [Ca2þ]i gradients can also reach hundreds of nanomoles per micrometer when measured along a line drawn from the apical to the basal part of an acinar cell (Gerasimenko et al. 1996). The well-established cell-free assay for intra-Golgi transport that measures glycosylation of the VSV-G protein is inhibited by BAPTA (IC50 0.8 mM) but not by EGTA. This indicates that luminal Ca2þ is required for transport (Porat and Elazar 2000) and suggesting a role for Ca2þ release from the luminal stores in the membrane fusion that accompanies intracellular transport. As the local release of Ca2þ occurs from the luminal stores of the organelles involved in fusion, can this explain the differential effects of BAPTA and EGTA? The on-rate of Ca2þ binding to BAPTA (Naraghi 1997) is similar to that for calmodulin (Falke et al. 1994), so that this Ca2þ chelator

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Figure 2. Behaviour of the GC and Ca2þ during the passage of cargo through the GC. A Exit of cargo (black lines) from the ER in ER-to-Golgi carriers. B,C Arrival of cargo at the GC triggers Ca2þ exit from the GC (grey shadow) that shifts the equilibrium between formation and consumption of COPI-dependent vesicles (blue circles) towards consumption, and that induces the SNAREdependent disappearance of vesicles and the formation of intracisternal connections. Ca2þ channels open, leading to release of Ca2þ (black dots in C), or there is a temporary reduction in SPCA activity. Increased [Ca2þ]i leads to recruitment of cPLA2 to GC membranes. D Shift of cargo domain to the trans side of the GC. E Reuptake of Ca2þ on the GC region due to closure of Ca2þ channels and maturation of the cargo domain into a post-Golgi carrier. F Enhanced activity of SPCA pumps restore basal [Ca2þ]i. Exit of post-Golgi carrier and restoration of vesicle equilibrium at the Golgi level.

can compete with calmodulin in its initial binding of Ca2þ. In contrast, EGTA has a considerably slower on-rate (Naraghi 1997). These differences in onrate would only be relevant for Ca2þ gradients lasting for less than 1 ms. It has been calculated that buffering by 10 mM BAPTA would reach equilibri-

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um in around 3 s, whereas that due to EGTA would take around 1.2 s (Adler et al. 1991). From the known diffusion rate for Ca2þ in the cell cytosol, it is clear that over 1 ms such a [Ca2þ]i gradient would dissipate within a distance of around 20 nm. This is a possible situation for a fast calmodulin-dependent fusion process that requires [Ca2þ]i increases in the micromolar range (or higher, as in yeast vacuole–vacuole fusion; Peters and Mayer 1998). If the basal [Ca2þ]i was sufficient, a differential effect of these two chelators on a Ca2þ/calmodulindependent fusion process would not be expected. Thus these differential effects indicate that the requirement must be for a local increase in [Ca2þ]i due to short-lived pulses of Ca2þ release from very close to the site of membrane fusion. This Ca2þ release would, therefore, have to be tightly coupled to the fusion, both spatially and temporally. One way in which this can be achieved for synaptic exocytosis is through the direct interactions of neuronal SNARE proteins with voltage-gated Ca2þ channels (Sheng et al. 1996). Perhaps similar interactions can occur with intracellular Ca2þ release channels. Thus, the special organization and localization of the Ca2þ stores can regulate the directionality of vectorial membrane transport.

Other functions of Ca2þ in trafficking Transient gradients in [Ca2þ]i also regulate the assembly and disassembly of the coat proteins that are responsible for vesicular trafficking between Golgi stacks and beyond the TGN. For instance, experiments with BAPTA and EGTA have demonstrated that some of their effects can be explained by an important role for Ca2þ in the stabilizing of the coat of the forming coated buds (Ahluwalia et al. 2001). As with Ca2þ-sensitive membrane fusion, COPI coat assembly is more sensitive to BAPTA than EGTA (Pryor et al. 2000). Also, after 90 min treatment of NRK cells with BAPTA-AM, while the GC was of similar size and retained a stacked structure, the number of cisternae within each stack was smaller. At the same time, accumulation of COPI-dependent vesicles was not seen because low Ca2þ blocks the formation of the COPI coat; indeed, the coatomer dissociates from the GC after treatment with BAPTAAM (Chen et al. 2002). Similar effect of Ca2þ on COPII coat have also been described. Sec31A (a subunit of COPII)-positive spots have been shown to increase in number and to be concentrated in a juxtanuclear region in response to Ca2þ mobilization. In contrast, Ca2þ chelation by BAPTA-AM decreases the number of punctate dots associated with Sec31A. The distribution pattern of the Sec23 (another subunit of COPII)-interacting protein p125 is not affected by treatment with either a Ca2þ ionophore or a Ca2þ chelator (Shibata et al. 2007). Additionally, at least at the level of the fusion with the PM, Ca2þ channels have been seen to associate physically and functionally with the Q-SNAREs, syntaxin and SNAP-25 (Yoshida et al. 1992; Sheng et al. 1994; Mochida et al.

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1996; Wiser et al. 1996; Rettig et al. 1997). Trans interactions between SNAREs on opposite membranes have been proposed to facilitate or trigger [Ca2þ]i signals in response to docking (Bezprozvanny et al. 1995; Scheckman 1998). Local increases in [Ca2þ]i near the GC lead to the recruitment of the cytosolic, Ca2þ-sensitive phospholipase A2 isoform cPLA2a to Golgi membranes (Ghosh et al. 2006; Polishchuk R, personal communication). The role of this event is not clear. Thus, normal and transiently increased [Ca2þ]i appears to be important not only for membrane fusion, but also for the formation of specialized membrane coats.

Roles of luminal Ca2þ in the Golgi apparatus Why it is necessary to have high concentrations of Ca2þ in the lumen of the ER and the GC? Fairly constant and rather high [Ca2þ]GC suggest that Ca2þ may be needed for the correct execution of luminal functions. However, little is known about the role of Ca2þ of this compartment, and its dynamic changes under physiological conditions. [Ca2þ]GC controls a variety of important functions, including protein and lipid synthesis, chaperone-dependent processing, glycosylation, sorting and eventual breakdown of newly formed proteins (Carnell and Moore 1994; Ivessa et al. 1995; Austin and Shields 1996; Duncan and Burgoyne 1996; Meldolesi and Pozzan 1998), as well as transport of proteins, cargo condensation, and precursor processing (Chanat and Huttner 1991; Carnell and Moore 1994; Austin and Shields 1996; Duncan and Burgoyne 1996; Corbett and Michalak 2000; Wuytack et al. 2003). Of interest, a sufficient supply of Mn2þ is also an absolute requirement for correct glycosylation of secretory proteins in the GC (Durr et al. 1998). Indeed, the activities of most enzymes are Ca2þ and Mn2þ dependent (Sharma et al. 1974; Parodi 1979). In mammalian cells, the endoproteolytic proprotein convertases (Davidson et al. 1988; Schmidt and Moore 1995; Steiner 1998) and the secretases (LaFerla 2002) in the GC and secretory vesicles are Ca2þ dependent. Ca2þ dyshomeostasis in these compartments could contribute to various amyloidoses, including Alzheimer’s disease. The role of luminal Ca2þ for human pathology can be illustrated by the intracellular pathogenic bacteria, such as Mycobacteria and Salmonellae, that have developed means to control fusion reactions in their host cells. They persist in phagosomes, the fusion of which with lysosomes is actively suppressed to ensure the bacteria survival inside host cells (Michelangeli et al. 2005). However, the maintenance of the high [Ca2þ]ER and [Ca2þ]GC appears not to be vital. Indeed, depletion of calcium from the ER and possibly from the GC, either by treatment with A23187 or thapsigargin, has no effects on the folding or secretion of newly synthesized albumin (Lodish et al. 1992). Moreover, cells can adapt their growth when the pumping of Ca2þ from the

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cytosol is blocked by thapsigargin, by expressing large amounts of thapsigargin-resistant Ca2þ-ATPase pumps (Waldron et al. 1995). Since under these conditions the Ca2þ pumps in the PM are functional, Ca2þ levels in the ER and the cytosol are expected to equilibrate at a low level. Thus, the maintenance of a high [Ca2þ] in the lumen of the ER and of the GC facilitates some important functions. These are necessary for enzymatic activity in these organelles and for the prevention of the early exit of unfolded proteins from the ER. The trans ER represents the source of the rapid increases in local [Ca2þ]i near the trans side of the GC. However, these high [Ca2þ] in the ER and GC lumen are not vital per se.

The role of Ca2þ in the TGN and endosomes The [Ca2þ] in the TGN and post-Golgi carriers is lower than in the GC (Pezzati et al. 1997). However, it has been shown that the selective aggregation of regulated secretory proteins in the TGN depends on luminal [Ca2þ] (Chanat and Huttner 1991; see also Chapter 3.11). On the other hand, it is known that in cells with regulatory secretion, most of the constitutively secreted proteins pass through endosomes (see Chapter 3.10). It is important to stress that these endosomal clathrin-coated buds and clathrin-independent tubules contain millimolar levels of Ca2þ when they pinch off from the PM, because clathrin-coated buds are in continuity with the external fluid that contains 1–2 mM Ca2þ. The fusion of recently uncoated clathrin-dependent buds or vesicles with endosome could provide a flux of Ca2þ from the extracellular space towards an endosome along the very thin tube that connects a bud with the PM, destabilizing the internal lipid leaflet. Ca2þ-sensitive probes that undergo endocytosis have demonstrated that the luminal [Ca2þ] is rapidly reduced from 1–2 mM to around 3 mM, within 20 min of endocytosis (Gerasimenko et al. 1998). In phagosomes, the [Ca2þ] can be as high as 400–600 mM (Christensen et al. 2002). Thus, the [Ca2þ] in the lumen of post-Golgi compartments and within the endosomal pathway is lower than in the [Ca2þ]ER and the [Ca2þ]GC. However, even this lower concentration is important for the execution of some functions that occur there. One of these functions could be the uptake of Ca2þ from outside.

Conclusion The cellular [Ca2þ] has an important role not only in signalling, but also in intracellular transport, and in particular in the directionality of consecutive fusion events along the secretory pathway. The GC contains a significant amount of Ca2þ that is involved not only in the regulation of intraluminal functions, but that is also released into the cytosol as part of the regulation of local [Ca2þ]i that is needed to maintain vectorial membrane transport during exocytosis.

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Abbreviations [Ca2þ] ER GC PM RyR STB TGN

Ca2þ concentration endoplasmic reticulum Golgi complex plasma membrane ryanodine receptor Shiga toxin-B trans-Golgi network

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Golgi glycosylation enzymes Eric G. Berger and Jack Rohrer

Historical perspective: the Golgi apparatus as the main site of glycosylation From time to time the question is posed by colleagues and research pupils about an object blackened by the classical Golgi techniques [black reaction or reazione nera developed by C. Golgi to identify the Golgi Apparatus], “Is this a Golgi body?” I suggest the proper answer would be: “I do not think that your question has a meaning. It is framed in terms of an improbable hypothesis” (Baker 1953). This telling citation coincides with the end of the long-lasting Golgi controversy about its mere existence; it was finally resolved by the clear definition of the GA1 as an ultrastructural entity (Dalton and Felix 1954). In fact, at these times the believers already recognized that the GA is likely to contain mono- and polysaccharides by virtue of specific histochemical staining (discussed by Bensley (1951)). The breakthrough to recognize the GA as the main cellular site of glycosylation can be traced back to the metabolic incorporation of glucose into cellular components shown by autoradiography to occur in the GA (Neutra and Leblond 1966). The procedure applied by these authors was inspired by Palade’s pioneering work on the secretory pathway (Caro and Palade 1964). The next milestone in associating glycosylation mechanisms with the GA was the advent of fractionation techniques combined with identification of subcellular fractions by marker enzymes such as galactosyltransferase [EC 2.4.1.22]. In the late 1960s a number of groups introduced a corresponding enzyme to identify Golgi fractions which were morphologically assigned to the GA (for review see Farquhar and Palade (1981)). The circle was then closed by the first immunocytochemical staining of the GA using antibodies to this enzyme (Berger et al. 1981), as shown in Fig. 1. Hence biochemical characterization of Golgi fractions was rendered feasible leading to a wealth of data allowing a coherent view on the stepwise assembly of glycans, mainly those of the N-glycosylation pathway of glycoproteins 1 The abbreviations are: AA, amino acids; BFA, brefeldin A; CT, cytoplasmic tail; CFP, cyan fluorescent protein; CDG, congenital disorder of glycosylation; cer, ceramide; fuc, fucose; fuc-T, fucosyltransferase, GA, Golgi Apparatus; GAG, glycosaminoglycan; gal, galactose; GalNAc-T, N-acetylgalactosaminyltranasferase; GalNA, N-acetylgalactosamine; GFP, green fluorescent protein; Glc, glucose; GlcA, glucuronic acid; GlcNAc, N-acetylglucosamine; Gn-T, N-acetylglucosaminyltransferase; GT, glycosyltransferase; hyl, hydroxylysine; man, mannose; PM, plasma membrane; PI, phosphatidyl-inositol; sia-T, sialyltransferase; TGN, trans Golgi network; TMD, transmembrane domain; wt, wild type; xyl, xylose

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Figure 1. Immunocytochemical staining of b4gal-T, the classical marker enzyme for the Golgi apparatus using polyclonal antibodies to a soluble form of this enzyme secreted in human milk (Berger et al. 1981).

(Kornfeld and Kornfeld 1985). This review is the milestone of the biochemical era. Remarkably, the pathway outlined therein is still valid, but has since been refined and complemented in cell-type-, species-, and development-dependent aspects. Introduction of antibodies to specific Golgi components initiated the cell biological era (Berger et al. 1981; Louvard et al. 1982) soon followed by cloning of the first genuine Golgi enzymes (Narimatsu et al. 1986; Shaper et al. 1986; Weinstein et al. 1987) introducing the molecular era. Both these new approaches have been fully accounted for in the monograph edited for the centenary of its discovery by C. Golgi (Berger and Roth 1997). Where do we stand now? In the past 10 years, new concepts of trafficking, biogenesis and organellar proteomics (Au et al. 2007) have emerged which all will push our current level of ignorance a bit farther away.

The basics Golgi glycosylation enzymes comprise two classes: processing glycosidases and glycosyltransferases. Glycosidases cleave and glycosyltransferases (GTS) create glycosidic linkages. Both are class II membrane proteins with their catalytic portion oriented to the lumen of the GA. Bioinformatics concerning these enzymes are found at http://www.cazy.org (Coutinho and Henrissat 1999). Among the glycosidases, only class I and class II mannosidases are associated with the GA (Herscovics 1999; Mast and Moremen 2006), the remaining glycosylation enzymes being glycosyltransferases. Their general function is depicted on Fig. 2. Catalytic properties are described in the legend of Fig. 2. A hallmark of these enzymes is their high substrate specificity. This property gave rise to the

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Figure 2. Glycosyltransferases catalyze the transfer of a sugar residue from a nucleotide-sugar (those acting in the GA are listed on the left side; the abbreviations are given in footnote 1). Catalysis of sugar transfer is classified into retaining or inverting mode depending on the anomeric configuration of the sugar before and after transfer. Usually, glycosyltransferases are activated by bivalent cations (Mn2+ or Ca2+) and follow an ordered compulsory mechanism by first binding the donor, then the acceptor substrate. Michaelis constants for the donor substrates are within the micromolar range whereas those for the acceptors are difficult to estimate; in vitro they are between micro- and millimolar. Optimum pH is around 7. The products are glycosylated acceptor and the nucleotide (NDP) which is immediately cleaved by a luminally acting nucleoside phosphatase to prevent kinetic inhibition. The resulting nucleoside monophosphate (NMP) is exchanged by a specific antiporter with the NDP-sugar synthesized in the cytoplasmic compartment (Caffaro and Hirschberg 2006). The fate of luminally produced phosphate is not known.

one enzyme-one linkage paradigm (Hagopian et al. 1968) predicting a specific glycosyltransferase, thus a gene encoding it, for the formation of each glycosidic bond. While this prediction was a fruitful basis for the search of distinct activities, molecular cloning has now revealed the existence of gene families for many glycosyltransferases with overlapping specificities. These are compiled in the Cazy database. An overview of glycosidic linkages found in mammals has been published (Ohtsubo and Marth 2006). The sequences of human Golgi glycosyltransferases forming the cytoplasmic, transmembrane and stem regions have recently been compared (Patel and Balaji 2007). Their respective topogenetic functions are discussed below. Like all genuine membrane proteins of the secretory pathway with type II topology, glycosyltransferases are synthesized in the ER with an internal signal sequence. They usually contain N- and/or O-linked glycans and follow the membrane flow of the secretory pathway. As discussed in ‘Trafficking of Golgi glycosyltransferases’ section, at some specific stage

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Figure 3. Golgi GTs are type II membrane proteins composed of four domains: a short cytoplasmic tail (CT), a transmembrane (TMD), a stem and a catalytic domain. They may be proteolytically cleaved in a post Golgi compartment and released as a soluble enzyme. PDB ID: 1O0R (Qasba et al. 2005).

within the Golgi cisternal stack GTs accumulate by mechanisms which are still poorly understood and which may involve complex formation (retention) and/or recycling by vesicular transport (retrieval) (see ‘Trafficking of Golgi glycosyltransferases’ section). Finally, they can move beyond the GA to undergo processing by furin-type proteases and to be released into the extracellular space as catalytically active, soluble enzymes devoid of their membrane anchor and part of the stem region (Fig. 3). Clearly, the mechanisms governing assembly and sequential arrangement of Golgi GTs are at the center stage of current efforts to understand the molecular mechanisms of Golgi-associated glycosylation.

Structural aspects Domain structure, topology Elucidation of the domain structure of Golgi GTs (see Fig. 3) was an important milestone (Paulson and Colley 1989). In a bioinformatics approach, the sequences of the domains of human GTs have recently been characterized (Patel and Balaji 2007): the cytoplasmic

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domain usually comprises 6–10 AAs but varies between a few and 80. They may contain motifs important for topogenesis (‘Summary and outlook’ section). The transmembrane domain contains 17–33 residues which would correspond to a length of up to 4.2 nm in an a-helical conformation. It is flanked by more positively charged residues (Arg, Lys) on the N-terminal than on the C-terminal side as predicted for type II membrane proteins according to the “positive-inside” rule (Higy et al. 2004). Moreover, in several cases intramembrane Cys residues possibly involved in dimer formation have been found (Qian et al. 2001; Sousa et al. 2003). The stem domain tethers the catalytic portion and enables molecular encounters of enzyme and acceptor, both anchored to the membrane; thus, the stem containing 70% of disorderpromoting AAs is believed to be flexible. The length varies between 10 and 165 AA and probably reflects the distance needed to bind the corresponding acceptor substrate; in accordance with this idea, glycolipid-specific GT appear to have shorter stem domains than those elongating N- or O-glycans.

3D structures of glycosyltransferases 3D structures are available for the catalytic portions of a number of Golgi GTs all belonging to the GT-A class as reviewed by Qasba et al. (2005) and Breton et al. (2006). A synopsis of the hitherto available structures is available at the following website: http://www.cermav.cnrs.fr/glyco3d/. These structures reveal common binding motifs (DXD or EXD) for the metal ion, the nucleotideactivated sugar followed by the acceptor substrate in an ordered sequential reaction. They also infer on the molecular mechanisms of the two catalytic processes, i.e. inversion and the less well understood retention of anomericity of the donor sugar. Moreover, substrate-induced conformational changes of a loop which acts as lid covering the donor substrate could be visualized; this in turn opens the space to bind the acceptor substrate (Qasba et al. 2005). Moreover, they form a rationale for designing GTs with defined specificities (Hancock et al. 2006).

Glycosylation pathways The GA is an impressive biosynthetic machine of glycans which accounts for the bulk of biomass in the living world. Notwithstanding the fact that an important glycosaminoglycan (hyaluronic acid) is synthesized at the plasma membrane in animal cells (Rilla et al. 2005) most glycans are assembled in the GA. These include peripheral sugars of protein-bound N-glycans and Oglycans, glycolipids and proteoglycans. A widely accepted view on this process predicts an assembly-line in which sequentially ordered GTs would elongate a glycan chain on a protein or lipid substrate along its passage through the GA. Several predictions can be made from this assumption: (1) glycosylation enzymes are expected to build-up in the GA in a steady state (i.e. inflow from endoplasmic reticulum (ER)-associated biosynthesis into the GA matches their outflow); (2) glycosylation enzymes would not be uniformly distributed

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along the Golgi cisternal stack; instead they would be sequentially arranged along the cis–trans axis of the GA; (3) co-localization of enzymes specific for the same acceptor substrate would result in competition. The first postulate has amply been confirmed and can be used as a litmus test for the specificity of antibodies to Golgi glycosylation enzymes: they have to provide the characteristic picture of the usually compact juxtanuclear localization of the GA at the level of the light microscope (cf. Fig. 1). There are cell-type specific variations but a close topographic relationship to the MTOC is invariably seen (Tassin et al. 1985). The best evidence for the second postulate relates to Golgi glycosylation enzymes involved in chain elongation and termination of N-glycans shown to be sequentially arranged (Rabouille et al. 1995). The third postulate has been documented in many instances; a recent striking example addresses competition of overexpressed fuc-T IV (EC 2.4.1.-) with a-gal-T (EC 2.4.1.87) for the terminal LacNAc structure leading to downregulation of the a-gal epitope (Hansen et al. 2005). The study of the topography of glycosylation enzymes with respect to the Golgi cisternal organization relies on different methods, each having intrinsic limitations as reviewed by Varki (1998): these included in the early days biochemical purification of GTs (Beyer et al. 1981) or their heterologous expression, immunocytochemical localization of the enzymes (Roth and

Figure 4. Glycan classes. The scheme depicts the sugars attached to the aglycone. Relevant to Golgi associated reactions are the elongation and termination of N-glycans, initiation of glycans, polymerization of glycosaminoglycans (GAG) and biosynthesis of glycolipids. The symbols designate: &GlcNAc; *Gal; Man; 4 Fuc; & GalNAc; Glc; rXyl.



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Taatjes 1998; Roth 1998), fractionation of the GA (Bretz et al. 1980), use of Golgi disturbing agents (Dinter and Berger 1998), intercompartmental transport assays (Pfeffer and Rothman 1987). Newer approaches include studies by life microscopy of GFP-tagged GTs (Lorenz et al. 2006). The concerted actions of Golgi GTs result in specific glycosylation pathways which can be classified according to (i) type of aglycone (lipid or protein), (ii) the first sugar linked to it and (iii) the species. An overview of all known start points of glycosylation pathways is given on Fig. 4: out of them, only those which are initiated or elongated in a mammalian GA are described in more detail. These include chain extension and termination of N-glycans and Oglycans as paradigms for further work aimed at understanding the molecular organization of these enzymes.

N-glycosylation N-glycosylation of proteins is initiated in the endoplasmic reticulum by en bloc transfer of the preformed oligosaccharide Glc3Man9GlcNAc2 from its dolichol carrier to the nascent polypeptide chain by the multimeric enzyme oligosaccharyltransferase. ER-associated glycans then fulfill a variety of intracellular functions such as quality control, sorting, degradation and secretion (for recent review see Helenius and Aebi (2004)). Their detailed description is beyond the scope of this chapter. Briefly, following transfer of the oligosaccharide to the peptide chain, one man residue is cleaved in the ER by mannosidase I (for review see Spiro (2004)). Glycoproteins then move to the GA (as outlined in ‘Domain structure, topology’ section) to arrive to the cis cisterna. The basic scheme of Golgi-associated trimming, chain elongation and termination reactions as proposed 20 years ago by (Kornfeld and Kornfeld 1985) is still basically valid and is depicted in Fig. 5. This pathway can be viewed as a theme with many species and cell-type specific variations. The final products are glycans exerting their functions (i) outside of the cell as part of secreted or shed glycoproteins, (ii) on the plasma membrane exposing the glycans on the cell surface, (iii) in post Golgi recycling compartments and (iv) in lysosomes. The enzymes modifying the N-glycans act sequentially from cis to trans as part of the multiglycosyltransferase system (Roseman 1970) characteristic for a given cell-type. Thus, the product of one becomes the substrate for the next enzyme. Annotations of these enzymes are available at the database CAZy (http://www.cazy.org/). Another useful website addressing GTs is found at http://www.functionalglycomics.org/glycomics/molecule/jsp/ glycoEnzyme/geMolecule.jsp. In the following, the main steps restricted to a human GA will be reviewed and recent findings mentioned. 1. 1a. The first Golgi-associated step is the conversion of the Man8GlcNAc2 N-glycan by mannosidases IA and IB to Man5GlcNAc2 (EC 3.2.1.113) as reviewed by Herscovics (Herscovics 1999). Recently, an additional type designated mannosidase IC has been cloned from a human fetal brain

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Figure 5. General N-glycosylation pathway in the Golgi apparatus. N-glycosylated proteins carrying a Man8GlcNAc2 glycan move by vesicular transport to the cis-Golgi where further processing (arrow 1a) takes place. Alternatively, lysosomal enzymes are substituted by one to two GlcNAc residues linked by a phosphodiester bond (arrow 1b). They are not further modified until removal of the GlcNAc by an uncovering enzyme in the TGN to expose the mannose-6 phosphate recognition marker. All other N-glycans may be elongated and terminated in the medial and trans-Golgi cisternae as depicted. Many alternative or additional substitutions are possible depending on the cell-type. These include core fucosylation (step 5), branching up to pentaantennary N-glycans and a number of different terminal structures. All symbols are explained in the legend to Fig. 4. ¤ designates sialic acid. The numbered arrows refer to the steps explained in the text. Detailed glycosidic linkages are shown on Fig. 6.

cDNA library and shown to have a tissular expression pattern different from IA and IB and a slightly different fine specificity in the order of cleavage leading to different intermediate compounds (for review see Herscovics (2001)). The final product of the class I mannosidases is the Man5GlcNAc2 species. 1b. An alternative glycosylation reaction substitutes lysosomal enzymes with a GlcNAc residues linked by a phosphodiester bond. The corresponding enzyme, the UDPGlcNAc: lysosomal enzyme-1-phosphotransferase (EC 2.7.8.15), a hexameric enzyme, has been purified (Bao et al. 1996), cloned (Kudo et al. 2005) and analyzed for pathogenic mutations (Tiede et al. 2005). The a/b subunits may carry loss of catalytic function mutations leading to mucolipidosis II (OMIM 252500) whereas defects of the g subunit which specifically recognizes lysosomal enzymes cause mucolipidosis III (OMIM 252600).

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2. Man5GlcNAc2-R then serves as a substrate for Gn-T I (EC 2.4.1.101) thereby initiating synthesis of hybrid or complex type N-glycans. This enzyme is of critical importance in the development of higher eukaryotes since mice in which the gene was ablated died around embryonic day 10 (Campbell et al. 1995). In fact, this enzyme has been extensively investigated in C. elegans leading to the surprising observation that worms have very little complex or hybrid N-glycans (Schachter 2004). A homologous enzyme encoded by a different gene has been described and designated Gn-T I.2 which may also be involved in extension of O-linked mannose (Zhang et al. 2002). 3. The ensuing hybrid-type glycan GlcNac1Man5GlcNAc2 becomes a substrate for processing mannosidase II (EC 3.2.1.114), a class II mannosidase (Moremen 2002). The alternative homologous enzyme mannosidase X (EC no entry; for recent discussion see: Akama et al. (2006)) may substitute for the former in case of its ablation. Persistence of the two remaining mannose residues by mannosidase II leads to the formation of hybrid glycans with different biological properties. Hence the development of inhibitors to this enzyme other than the classical compound swainsonine has been a subject of considerable interest in recent years (Kawatkar et al. 2006). 4. Following removal of mannose residues, Gn-T II (EC 2.4.1.143) adds another GlcNAc residue to the 1!6 branch to form GlcNAc2Man3GlcNAc2-R. Transcriptional regulation of this enzyme has been investigated and shown to depend on the Ets transcription factors but not src or neu (Zhang et al. 2000). The biosynthetic importance of this enzyme is also underlined by its defect in CDG IIa characterized by frequent postnatal lethality with multiple defects (OMIM 212066). 5. The formation of GlcNAc2Man3GlcNAc2-R permits a6fuc-T8 (EC 2.4.1.68) to transfer a fucose residue to the innermost GlcNAc, a process called core fucosylation. Also triantennary N-glycans with terminal GlcNAc residues can be core fucosylated whereas bisected glycans (see below) are not substrates. Expression of this enzyme is upregulated in a variety of cancer cells (Miyoshi et al. 1999). In addition, the crystal structure of the human enzyme has recently been reported (Ihara et al. 2007).

Figure 6. Branching of N-glycans. Specific N-acetylglucosaminyltransferases act in a cellspecific manner to form up to five branches (pentaantennary N-glycan). The depicted hexaantennary structure has not been observed in vivo.

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6. At this critical stage, either further branching or chain elongation to biantennary complex N-glycans may take place. A detailed insight into the numerous possibilities of branching by Gn-Ts is provided in the classical GO-NOGO lecture by Schachter (1986). One of the six branching reactions (Fig. 6) is catalyzed by “bisecting” Gn-T III (not shown on Fig. 5; EC 2.4.1.143), which received a great deal of attention in recent years (for review see Ikeda and Taniguchi (2001)). Remarkably, this branch is never extended and prevents further processing by mannosidase II, branching by core fuc-T, or importantly, Gn-T II, Gn-T IV (EC 2.4.1.145) and Gn-T V (EC 2.4.1.155) (Zhao et al. 2006). However, the enzyme can act on any bi-, triand tetraantennary form of N-glycans provided that they contain no galactose. Expression of Gn-T III appears to exert a pivotal role as it impairs branching by Gn-T V, a branch conferring metastatic potential to melanoma or NIH3T3 cells (for review see Gu and Taniguchi (2004)). 7. All the intermediate species are substrates for Gn-T IV which adds a b4GlcNAc to the 3-branch of core mannose leading to a triantennary species, or tetraantennary species if Gn-T V has already acted on the 6branch of core mannose. A second isoform exists with an ubiquitous tissue distribution (Yoshida et al. 1998). An intriguing increase of its expression has been observed in choriocarcinoma where overexpression of this enzyme forms so-called abnormal biantennary glycans, i.e. glycans lacking the product of Gn-T II (Takamatsu et al. 1999). 8. The a6 branch of core mannose can be substituted by Gn-T V (not shown on Fig. 5) which is also represented by a genetically distinct isoform designated Gn-T VB (Kaneko et al. 2003) or Gn-T IX with predominant expression in the brain (Inamori et al. 2006). Most interestingly, Gn-T VA has been implicated in a number cancer-associated changes of glycosylation as this enzyme was found to be responsible for increased branching of N-glycans in highly metastatic cancer cells (Dennis 1988); moreover this branch may carry polylacNAc structures which also contribute to the increased size of the N-glycans, the “Warren phenomenon” known since the seventies (Warren et al. 1978). Gn-T VA has a complex promoter set-up with several binding sites for transcription factors (Saito et al. 1995) and, remarkably, PEA3/Ets binding sites which can by activated by RAS-RAFMAPK signalling (Buckhaults et al. 1997). An entirely new connection of the b1,6 branch introduced by Gn-T V has been revealed in Mgat5(  /  ) mice: they appeared to be less sensible to anabolic cytokines since their receptors have reduced surface expression in absence of this branch (Cheung et al. 2007). While pentaantennary structures are commonly found, hexaantennary structures, although theoretically possible, have not been reported. 9. N-glycans substituted with GlcNAc residues then move by mechanisms addressed in ‘Trafficking of Golgi glycosyltransferases’ section to the transGolgi compartment, the site of chain elongation and termination. TransGolgi cisternae harbour gal-Ts (Roth and Berger 1982), sia-Ts (Roth et al.

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1985) and probably a number of terminating GTs which have not been mapped yet by immunoelectron microscopy. The first reaction occurring in trans-Golgi cisternae is galactosylation of GlcNAc residues. Most commonly, this reaction is catalyzed by the ubiquitous (though downregulated in neural tissue) b4gal-T1 (EC 2.4.1.22), probably the most thoroughly investigated mammalian GT (Berger and Rohrer 2003). Interest in this enzyme dates back to an intriguing finding of a high concentration of soluble gal-T in embryonic chicken brain later implicated in specific intercellular adhesion (Den et al. 1970). This line of research pertains to ectopic localization of Golgi GTs reviewed by Berger (2002) and is not addressed in this review. Galactosylation takes place by two classes of enzymes, the b1!4 and b1!3gal-T (for reviews see Amado et al. (1999) and Hennet (2002)). While the glycosidic linkage formed by these families is the same among their members, the acceptor substrates are different and define the biological role. Among all gal-Ts acting along this pathway, only gal-T1 has been deleted in mice (Asano et al. 1997; Lu et al. 1997) leading to postnatal lethality, endocrine defects and skin abnormalities. This enzyme also served as a unique paradigm for a protein modifier function: its binding of a-lactalbumin, a protein uniquely expressed in lactating mammary gland shifts recognition of GlcNAc-R as acceptor to glucose thereby forming lactose (Hill and Brew 1975). At present, interest in this enzyme focuses on trafficking, complex formation, shedding and possible regulation by phosphorylation. 10. 10a. Biosynthesis of N-glycan chains very often is terminated by sialylation which appears to immediately follow galactosylation. Although co-localization of endogenously expressed gal-Ts and siaTs has been confirmed at the level of light microscopy (Taatjes et al. 1987), evidence for co-localization at the ultrastructural level was only possible with transfected sia-T (Kweon et al. 2004). Out of the numerous different sia-Ts reviewed by Harduin-Lepers et al. (2005), ST6Gal1 has been investigated with respect to its targeting in some detail (see ‘Trafficking of Golgi glycosyltransferases’ section). Another paradigmatic feature of Golgi GTs was the identification of sequence motifs involved in donor or acceptor substrate binding, respectively. These were called sialyl motifs (Datta and Paulson 1995; Datta et al. 1998) and shown to be present also in polysiaTs (see below). Chain termination may involve other GTs as schematically shown on Fig. 7. An alternative to 2!6 is 2!3 sialylation (EC 2.4.99.6) depending on the relative expression of 2,3 versus 2,6 sia-Ts (Lee et al. 1989). 10b. Synthesis of the blood groups is catalyzed by allozymes encoded on the ABO locus; differences of four AAs between the blood group B (a3gal-Ts, EC 2.4.1.37) and blood group A forming enzyme (a-galNAc-T, EC 2.4.1.40) are known since their cloning (Yamamoto et al. 1990); however, the switch of their donor substrate specificity

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Figure 7. Biosynthesis of ABO(H) histo blood groups. Symbols are defined in the legend to Fig. 4

appears to rely on a single nucleotide exchange only leading to a P234S mutation. Thus, affinity changes from UDPGal to UDPGalNAc (Marcus et al. 2003). This type of inference on the basis of substrate specificity of a GT also lays the foundation for enzyme engineering projects as outlined in Hancock et al. (2006). The 0 enzyme most frequently is truncated at position 117 (instead of the full length 354 AAs) but a full length variant devoid of catalytic activity but Golgilocalized has been described (Amado et al. 2000). This raises the intriguing question whether the phenomenon of a correctly expressed yet inactive enzyme is an incidentally recognized tip of an iceberg. Blood group specifying GTs act only on fucosylated terminal galactoses as shown on Fig. 7. Thus, corresponding fuc-Ts precede their action to form the H epitope (more commonly designated 0 blood group) or Se (EC 2.4.1.69) on both type 1 and type 2 LacNAc’s. These two types of fuc-Ts are encoded by the FUT1 or the FUT2 gene, respectively. Together with fuc-Ts involved in biosynthesis of the Lewis structures (thoroughly reviewed by Lowe (1995)) a sizable diversity of structures can be made in the distal Golgi compartments as recently reviewed by Ma et al. (2006). A common theme of these structures is their involvement in cellular adhesion and recognition

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events both important in inflammation (Rosen 2004) and cancer metastasis (Borsig 2004). 10c. A variety of other chain termination reactions have been described such as polysialic acid, a polymer of a2!8 linked sialic acid residues. This structure extends a2,3 sialic acids of N-glycans preferably attached to the neural cell adhesion molecule (Bonfanti 2006) and bears important properties during embryonic neurogenesis. It is synthesized by two synergistic polysia-Ts designated ST8Sia II and ST8Sia IV (EC 2.4.), respectively (reviewed by Angata and Fukuda (2003)). Considering the conventional topology of these enzymes as luminally oriented catalytic sites, polysialic acid most likely occurs on the luminal side of trans-Golgi cisternae although a cytoplasmic orientation has been proposed (Bonfanti 2006). 10d. An important terminal biosynthetic reaction is sulfation catalyzed by a variety of sulfo-Ts with similar domain structure and topology as the GTs. The donor substrate is 30 -phosphoadenosine 50 -phosphosulfate (PAPS) synthesized in the cytoplasm and transported across the Golgi membrane alike the sugar nucleotides (Abeijon et al. 1997). Sulfo-Ts occur in the cytoplasm as well as in the GA both enzyme species belonging to a single gene superfamily with some common features of their enzymatic mechanisms (Negishi et al. 2001); those directed to the secretory pathway carry an internal signal sequence specifying their import into the endoplasmic reticulum; however, little is known concerning their further topogenesis within the Golgi cisternal stack. Golgi-associated sulfo-Ts either decorate in a highly specific manner glycosaminoglycans on their repetitive carbohydrate backbone or they add sulfate to terminal glycans or tyrosines on specific proteins as schematically shown on Fig. 8 (reviewed by Chapman et al. (2004). 11. The sulfated products exert highly specialized functions as also highlighted by an emerging group of genetic defects known as sulfation defects, in particular those caused by deficient 6-O-sulfotransferase-1 entailing spondyloepiphyseal dysplasia (OMIM 603799 (Thiele et al. 2004) and corneal GlcNAc-6-sulfo-T (C-GlcNAc6ST) causing macular corneal dystrophy (OMIM 217800) (Akama et al. 2000). An intriguing structure is the HNK-1 epitope almost exclusively found in neural tissue involved in neural development (Schachner et al. 1995); the role of the terminal sulfate, however, is not clear, since knockout-mice for the corresponding sulfo-T did not reveal any phenotype (Chou et al. 2002). The precursor structure, e.g. a glucuronic acid residue attached in b1!3 to LacNAc is formed by two specific glcA-Ts, GlcAT-P and GlcAT-S (EC not specified) of which GlcAT-P in fact seems to be involved in spatial memory formation (Yamamoto et al. 2002) and whose catalytic domain has recently been crystallized (Kakuda et al. 2004).

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Figure 8. Terminal glycans (including sulfates) expressed on a restricted set of protein carriers reflecting cell specificity of terminal GT and sulfotransferase expression.

O-glycosylation On Fig. 4 the currently known O-glycans are listed. Only the most abundant O-glycans initiated by GalNAc-Ts (EC 2.4.1.41) are exclusively synthesized in the GA whereas attachment of O-linked mannose and O-linked fucose as well as collagen glycosylation is ER-associated. The former may be extended by Golgi-associated GTs in analogy to O-linked GalNAc. O-linked GlcNAc is a special case as it is reversibly transferred to cytoplasmic and nuclear protein acceptors by a cytosolic Gn-T and appears not to be elongated by further carbohydrates (Hart et al. 2007). 1. The first step involves transfer of GalNAc to a Ser or Thr residue of a mucin-type domain of a glycoprotein catalyzed by one of the 18 different UDPGalNAc:polypeptide GalNAc-Ts (EC 2.4.1.41) hitherto listed on the CAZy database, enzyme family 27 (http://www.cazy.org/fam/GT27.html) (Ten Hagen et al. 2003). A general scheme of O-glycan biosynthesis is given on Fig. 9, again as a theme with many variations dependent on celltype, developmental stage and cellular differentiation. A few hallmarks distinguish the O- from the N-glycan biosynthetic pathway: (i) O-glycan biosynthesis is confined to the GA; (ii) the high diversity of genetic isoforms of polypeptide GalNAc-Ts reflects their restricted specificity to the different polypeptide backbones; (iii) the topography of O-glycosylation

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Figure 9. Simplified scheme of O-glycosylation initiated by GalNAc-Ts. GalNAc-Ts seem to be located throughout the Golgi stack (see text). Thus, truncated glycans may be expressed on the cell surface such as the TF (Thomsen-Friedenreich), Tn and sialosyl-Tn antigens. In many cells, only the glycophorin type of the O-linked tetrasaccharide is formed while in others (e.g. activated lymphocytes) a polylacNAc extension including a sialyl-Lex epitope is synthesized on a core 2 structure.

reactions is largely unknown, as the three different GalNAc-Ts localized by immuno electron microscopy did not reveal any cisternal preference for GalNAc-T1, whereas GalNAcT2 and T3 were preferentially labeled over the trans cisternae (Rottger et al. 1998); these results suggested that initiation of O-glycan biosynthesis may occur at any stage along cisternal progression and may partially explain the variable size of O-glycans on defined proteins; (iv) an interesting kinetic phenomenon has been observed in the case of GalNAc-T2 and -T4 in that a lectin domain of the enzyme specific for O-linked GalNAc-peptide promotes further transfer of GalNAc residues thereby increasing the density of O-glycans (Wandall et al. 2007). 2. The second step in O-Glycan biosynthesis may involve formation of the tumor-associated antigen sialyl-Tn by ST6GalNAc II (EC 2.4.99.3) (Sewell et al. 2006). A more common alternative is chain extension by b3gal-T also designated T-transferase (C1GalT1 or EC 2.4.1.122) as this enzyme is involved in the formation of the Thomsen–Friedenreich antigen and whose deficiency may cause the Tn-syndrome (Berger 1999). Cloning of this enzyme by Cummings and associates revealed a hitherto unique feature in GT enzymology as the activity in vivo depends on the co-

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expression of an X-linked chaperone designated COSMC or C1GalT2 (Ju and Cummings 2002) which may be mutated in the Tn-syndrome (Ju and Cummings 2005). Defects of both C1GalT1 or C1GalT2 have been associated also with IgA nephropathy (Barratt et al. 2007). 3. The structure galb1!4GalNAc-R (TF-antigen) is usually sialylated to form the glycophorin type tetrasaccharide depicted on Fig. 9. by ST3Gal I (EC 2.4.99.4) and ST6GalNAc IV (EC 2.4.99.7). 4. Formation of core 2 structures as depicted on Fig. 9 depends on expression of core 2 Gn-T (EC 2.4.1.102), which is up-regulated in T lymphocytes upon stimulation as initially shown by Piller et al. (1988). These may then be extended by repeating LacNAc’s and terminated by ABO(H) or sialylLewis blood groups. In fact, seven different core structures have been described which all may be elongated. A detailed description is given by Brockhausen (1995).

Glycolipids A huge diversity of glycolipids is synthesized in the Golgi stack essentially along a similar assembly-line like the mechanism described above for protein glycan chain elongation and termination. A few aspects specific to glycolipid biosynthesis are as follows: (i) the first glycosylation reaction, i.e. the formation of glucosylceramide, occurs on the cytoplasmic face (Coste et al. 1986) on the cis side of the GA where it can be bound by FAPP2 for transport to more distal sites of the GA (D’ Angelo et al. 2007). Very recent results also suggest that part of cytoplasmically oriented glucosylceramide is flipped to the luminal side in the ER whence elongation and termination of the glycolipids occurs along the transit through the GA (Halter et al. 2007). A detailed chart of biosynthetic pathways is available (Ichikawa and Hirabayashi 1998).

Proteoglycans The four main classes of proteoglycans comprise sulfated forms of keratan, chondroitin, dermatan and heparan. Biosynthesis of proteoglycans includes ER-associated steps such as translation and import of the core protein and glycan chain initiation by transfer of xylose. Further steps include core glycan elongation by two galactoses followed then by chain extension with alternating sugars. These occur in the GA. A common hallmark is their assembly in repeating disaccharides of variable length and the modifications imparted to the sugar chains in parallel to their assembly. An interesting and somewhat unique feature in glycan biosynthesis are the two established cases of families of bifunctional enzymes participating in the assembly of heparan sulfate: these are known as copolymerase EXT1 bearing two catalytic sites in tandem, one specific for the transfer of a4GlcNAc, the other for b4GlcA (EC 2.4.1.223). Concerted action of both catalytic sites produces repeating units of GlcNAca4GlcAb4. These structures

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are modified by N-deacetylase N-sulfotransferase (NDST1) (EC 2.8.2.8.) leading to sulfation of specific sites of GlcNAc. Little is known about the molecular interactions of the alternating GTs and their topography along the cisternal stack. An interesting view on their putative supramolecular structure is its designation as a “gagosome” (Esko and Selleck 2002): this would be a physical complex of enzymes involved in polymerizing the glycosaminoglycan backbone along with their modifications. Such a complex could explain the speed of the enzymatic reaction by substrate channelling and the formation of bifunctional enzymes during the evolution. It will be a challenging task to investigate how this concept fits to the notion of cisternal progression.

Comparative and evolutionary aspects Details of the glycosylation pathways described above refer to the human species; however, it is clear that all eukaryotes comprising as diverse species as yeasts, plants, worms, insects and mammals all synthesize their glycome by virtue of a GA. These aspects are dealt with in chapter 4.5. The basic principles of glycan assembly are common to all eukaryotes. The changes in evolution concern expression and specificities of the GT repertoire in a given organism (Varki 2006). This evolutionary change occurs over time by proneness to infections as a result of specific interactions between the host glycans and infectious agents. Alternatively, also sexual selection can lead to shifts in the glycan make-up as the example of the ABH blood groups show. Shifts of specificities of GTs by random genetic drift may always implicate pleiotropic changes with metabolic and morphogenetic consequences. This is exemplified by the genetic defects designated congenital disorders of glycosylation (Freeze 2006) which, in fact, cover the entire spectrum from embryonic lethality in case of complete enzymatic knock-out (for example b3galT, Xia et al. (2004)) to no apparent phenotypic changes in the case of absence of blood groups A or B. In between, GT polymorphisms can lead to all kinds of subtle phenotypic differences.

Trafficking of Golgi glycosyltransferases Sorting machineries required for the sorting of glycosyltransferases Following synthesis in the ER (see ‘The basics’ section), GTs are concentrated at ER exit sites and are packaged in transport vesicles that, in mammalian cells, seem to undergo homotypic fusion to form the ER–Golgi intermediate compartment before being delivered to the cis cisterna of the Golgi stack. Upon arrival in the GA the cell faces the difficult task to sort the individual GTs to specific cisternae of the GA (cis-, medial- or trans-Golgi) where at steady state the majority of a particular enzyme accumulates to perform its function within pathways outlined in ‘Glycosylation pathways’ section of this chapter. To determine how the cell can maintain this asymmetrical distribution of GTs in the GA while at the same time ensuring efficient passage of newly synthesized proteins through the secretory pathway is a major challenge for

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cell biologists. Chapters 2.6 and 3.2 address current models and the molecular machineries of intra Golgi trafficking. In the context of GTs, the conserved oligomeric Golgi (COG) complex appears on center stage. It consists of two lobes each comprising of four proteins. The precise role of the COG complex is not known so far but current evidence suggests that the COG complex is required for the retrograde trafficking of GTs (Chatterton et al. 1999; Kingsley et al. 1986; Ungar et al. 2002; Wu et al. 2004; Zolov and Lupashin 2005). Somatic mutations and siRNA experiments in mammalian cells as well as subtypes of CDG (Wu et al. 2004) showed global defects in protein glycosylation which are at least in part due to the mislocalization or instability of GTs. Despite intense research over the past decades it is still under debate whether the mechanism for the correct localization within the GA of different GTs is determined by the cytoplasmic-, transmembrane- and/or luminal domain (reviewed originally by Colley (1997)). Most likely this has to be assessed for each GT individually and possibly there are multiple signals/ mechanisms to localize the enzymes to their correct place within the GA.

Trafficking of galactosyltransferases At steady state the b4gal-T1 (EC 2.4.1.22) was found to be localized to the trans cisterna of the GA as determined by immuno electron microscopy (Roth and Berger 1982). Initial studies identified the transmembrane domain of b4gal-T1 as the important feature for the correct Golgi localization (Aoki et al. 1992; Nilsson et al. 1991; Teasdale et al. 1992). However, further data imply additional domains like the cytoplasmic domain to be required for efficient localization (Evans et al. 1993; Nilsson et al. 1991; Russo et al. 1992). Using a GFP-tagged form of the b4gal-T1 or the endogenous form, it was shown that the enzyme cycles between the trans-Golgi and the ER (Rhee et al. 2005; Zaal et al. 1999). Most recently (Schaub et al. 2006) could demonstrate that the cytoplasmic domain contains a specific signal that is required for the transport of b4gal-T1 from the trans-Golgi to the TGN. In contrast to the vesicular transport model, this result is unexpected in the view of a pure cisternal maturation model which would not require signals for proteins to be included in anterograde transport. In fact, mutating such a signal prevented delivery of b4gal-T1 to the TGN (Schaub et al. 2006, unpublished results). Either b4gal-T1 represents an exceptional case as it is also cycling back to the ER (Zaal et al. 1999) and that there are competing transport events which might be regulated by special signals or the cisternal maturation model has some unexpected requirements for the transport of Golgi enzymes between the trans-Golgi and the TGN. In any case, a leakage at the level of the recycling from the TGN back to the trans-Golgi could also explain the presence of a small fraction of b4gal-T1 at the surface in some cells (reviewed by Shur et al. (1998)). A soluble form of the enzyme was originally observed in bovine milk (Babad and Hassid 1966) and later purified from human milk (Gerber et al. 1979). However, it is not known if the soluble form is created by clipping the enzyme in the TGN with subsequent exocytosis (Strous and Berger 1982) or at the cell surface or both.

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For a3gal-T (EC 2.4.1.87) it was shown that overexpression of a soluble form of the cytoplasmic domain led to a mislocalization of the enzyme and that replacement of the TMD with a TMD from a PM protein did not affect the Golgi localization indicating that the cytoplasmic domain is a critical feature for its correct localization (Milland et al. 2002).

Trafficking of sialyltransferase ST6Gal1 (EC 2.4.99.1) seems to have a complicated mechanism or rather multiple mechanisms for its localization in the trans-Golgi cisterna as several reports using different experimental systems implicate different domains that are required for correct localization of the enzyme. Early on, Colley et al. (1989) found that sequences required for Golgi localization are within the CT, TMD and/or stem region. Later it was suggested that the TMD and flanking regions are sufficient for the Golgi localization with complete retention in COS and CHO cells only if CT and stem were present (Dahdal and Colley 1993; Munro 1991). Gradually increasing the length of the TMD resulted in the mislocalization of an ST6Gal1-lysozyme chimera to the plasma membrane (Munro 1995) but replacing the TMD of ST6Gal1 by the longer TMD of neuraminidase did not affect the localization (Dahdal and Colley 1993). Furthermore, the formation of disulfide bonded dimers of ST6Gal1 was assumed to promote retention of the enzyme in the GA as only monomers were found to be secreted (Chen et al. 2000; El-Battari et al. 2003) but Qian et al. (2001) identified Cys24 within the TMD as crucial for dimer formation explaining the absence of secreted dimers in the media. Recently, a very elegant study by Fenteany and Colley (2005) found that on one hand the CT is required for Golgi localization and on the other hand that oligomerization of the enzyme, mediated by luminal sequences, is required as well. This leads to a model where ST6Gal1 is concentrated in the Golgi due to its TMD and cytoplasmic domain which subsequently leads to the oligomerization mediated by the luminal sequences (Cys123 in particular).

Trafficking of N-acetylglucosaminyltransferase Early experiments using chimeric proteins between type II surface proteins (dipeptidyltransferase and transferrin receptor) and of Gn-T1 (EC 2.4.1.101) revealed that the TMD of Gn-T1 is required for localization of the enzyme in the medial/trans-Golgi (Tang et al. 1992). This was initially confirmed by Burke et al. (1992) using the TMD and immediate flanking regions to retain ovalbumin in the GA. However, a more detailed analysis later demonstrated that all three domains of the Gn-T1 contribute to the efficient localization within the GA (Burke et al. 1994). Further dissecting the role of the luminal domain of the GnT3 (EC 2.4.1.144) with a comprehensive examination of the three N-glycosylation sites of the enzyme revealed that not only the enzyme activity but also Golgi localization decreased by reducing the number of glycosylation sites (Nagai et al. 1997). Moreover, additional work from Taniguchi’s group showed that part of the stem region of the Gn-T5 (EC 2.4.1.155) participates in Golgi

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retention through disulfide bond mediated homo-oligomer formation without affecting the enzymatic activity (Sasai et al. 2001). In contrast, analysis of constructs containing domains of the human O-glycan core 2 Gn-T (C2Gn-T, EC 2.4.1.102) fused to GFP clearly identified the CT and TMD but not the stem region as the necessary and sufficient parts of the enzyme for Golgi localization (Zerfaoui et al. 2002). The authors could also show that the CT and TMD of C2Gn-T fused to the luminal domain of fuc-T7, (EC 2.4.1.-) displaced the chimera from a trans-Golgi cisterna to cis-/medial-cisterna as analyzed by changes of the cell surface glycosylation pattern. Taken all results into account it becomes obvious that the different Gn-T’s either require all three domains for efficient Golgi localization but each of the domains might mediate a different function within the scheme or that the individual Gn-T’s might use different mechanisms to achieve Golgi localization and therefore require just one or two domains for this task.

Trafficking of fucosyltransferase Early studies demonstrated that fuc-T6 (EC 2.4.1.65) co-localizes with b4gal-T1 in the GA and monensin-induced swollen vesicles (Borsig et al. 1999) indicating a cycling of the enzyme through the TGN (see section on b4gal-T1). Milland et al. (2001) analyzed the significance of the CT for the localization of the fuc-T1 (EC 2.4.1.69) and found that upon deletion of the CT more of the mutant fuc-T1 retained a perinuclear staining pattern after addition of BFA compared to the wt fuc-T1, indicating a mislocalization of the tailless enzyme to the TGN probably due to a missing retrieval/recycling signal. Mutations of individual AAs within the CT (MWVPSRRH) further revealed a role of residues 3–7 for correct localization with a particular role for Ser5, suggesting a potential role of phosphorylation for this process (Milland et al. 2001). In contrast to this, Sousa et al. (2004) reported data based on immunofluorescence that deletion of the CT of fuc-T3 (EC 2.4.1.65) led to a mislocalization of the enzyme to an earlier cisterna of the Golgi than the wt protein. This apparent contradiction between the two studies can hardly be explained by a different experimental set-up but seems rather due to different localization requirements of fuc-T1 and fuc-T3. Analyzing the TMD of fuc-T3 it was shown that mutation of four non-hydrophobic residues (C16, Q23, C29 and Y33) to leucines led to its surface accumulation (Sousa et al. 2003). Furthermore, C16 and C29 were shown to be required for dimer formation of fuc-T3 and all four residues seem to be important for the incorporation of the enzyme into COPI vesicles indicating that the TMD is also required for recycling of the enzyme from later compartments (Sousa et al. 2003).

Trafficking of N-acetylgalactosaminyltransferase Initial experiments with the b4GalNAc-T (EC 2.4.1.92) demonstrated that homodimers are formed as a result of intermolecular disulfide bonds creation in the ER which might be an important feature for the localization (see above) (Zhu et al. 1997). Recently, a study using chimeric molecules between GalNAc-

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T and Sia-T2, CT and TMD fused to fluorescent protein tags, revealed that the CTof the GalNAc-T is sufficient to cause accumulation of the chimera mostly in the TGN as indicated by resistance to BFA (Uliana et al. 2006). In these experiments the TMDs did not seem to have an effect for the correct localization of the enzymes.

General remarks on trafficking of glycosyltransferases Taken together, it becomes obvious that the individual GTs do not have one common signal or feature that determines the localization and trafficking pathways for all of them. The enzymes have to be analyzed individually and what has been discovered for one does not necessarily apply to another. It seems that the localization for many GTs is mediated by a mix of oligomerization which requires the TMD and sometimes the stem region, and specific transport signals localized in the CT. The individual requirements and pathways for the correct localization of the GTs within the GA are especially important if the enzymes are used as markers for the GA in general because experiments using one particular enzyme could yield different results if performed using another enzyme.

Summary and outlook There is considerable advance in knowledge on Golgi glycosylation enzymes since publication of the centennial book (Berger and Roth 1997), mainly with respect to 3D-structures, catalytic mechanisms and defects leading to type II of CDG. In some cases new genetic isoforms have been added to the list of known glycosyltransferases complementing our knowledge on cell specificity of glycosylation pathways. An emerging field addresses the still poorly understood topogenetic mechanisms, the definition of interacting partners and activity regulation by complex formation and/or phosphorylation. At present, however, we are still far away from a unifying concept regarding the fine distribution of glycosyltransferases along the Golgi cisternal stack.

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Warren L, Buck CA, Tuszynski GP (1978) Glycopeptide changes and malignant transformation. A possible role for carbohydrate in malignant behavior. Biochim Biophys Acta 516: 97–127 Weinstein J, Lee EU, McEntee K, Lai PH, Paulson JC (1987) Primary structure of betagalactoside alpha 2,6-sialyltransferase. Conversion of membrane-bound enzyme to soluble forms by cleavage of the NH2-terminal signal anchor. J Biol Chem 262: 17735–17743 Wu X, Steet RA, Bohorov O, Bakker J, Newell J, Krieger M, Spaapen L, Kornfeld S, Freeze HH (2004) Mutation of the COG complex subunit gene COG7 causes a lethal congenital disorder. Nat Med 10: 518–523 Xia L, Ju T, Westmuckett A, An G, Ivanciu L, McDaniel JM, Lupu F, Cummings RD, McEver RP (2004) Defective angiogenesis and fatal embryonic hemorrhage in mice lacking core 1-derived O-glycans. J Cell Biol 164: 451–459 Yamamoto F, Clausen H, White T, Marken J, Hakomori S (1990) Molecular genetic basis of the histo-blood group ABO system. Nature 345: 229–233 Yamamoto S, Oka S, Inoue M, Shimuta M, Manabe T, Takahashi M, Miyamoto M, Asano M, Sakagami J, Sudo K, Iwakura Y, Ono K, Kawasaki T (2002) Mice deficient in nervous system-specific carbohydrate epitope HNK-1 exhibit impaired synaptic plasticity and spatial learning. J Biol Chem 277: 27227–27231 Yoshida A, Minowa MT, Takamatsu S, Hara T, Ikenaga H, Takeuchi M (1998) A novel second isoenzyme of the human UDP-N-acetylglucosamine:alpha1,3-D-mannoside beta1,4-N-acetylglucosaminyltransferase family: cDNA cloning, expression, and chromosomal assignment. Glycoconj J 15: 1115–1123 Zaal KJ, Smith CL, Polishchuk RS, Altan N, Cole NB, Ellenberg J, Hirschberg K, Presley JF, Roberts TH, Siggia E, Phair RD, Lippincott-Schwartz J (1999) Golgi membranes are absorbed into and reemerge from the ER during mitosis. Cell 99: 589–601 Zerfaoui M, Fukuda M, Langlet C, Mathieu S, Suzuki M, Lombardo D, El-Battari A (2002) The cytosolic and transmembrane domains of the beta 1,6 N-acetylglucosaminyltransferase (C2GnT) function as a cis to medial/Golgi-targeting determinant. Glycobiology 12: 15–24 Zhang W, Betel D, Schachter H (2002) Cloning and expression of a novel UDP-GlcNAc: alpha-d-mannoside beta1,2-N-acetylglucosaminyltransferase homologous to UDPGlcNAc:alpha-3-d-mannoside beta1,2-N-acetylglucosaminyltransferase I. Biochem J 361: 153–162 Zhang W, Revers L, Pierce M, Schachter H (2000) Regulation of expression of the human beta-1,2-N-acetylglucosaminyltransferase II gene (MGAT2) by Ets transcription factors. Biochem J 347: 511–518 Zhao Y, Nakagawa T, Itoh S, Inamori K, Isaji T, Kariya Y, Kondo A, Miyoshi E, Miyazaki K, Kawasaki N, Taniguchi N, Gu J (2006) N-acetylglucosaminyltransferase III antagonizes the effect of N-acetylglucosaminyltransferase V on alpha3beta1 integrinmediated cell migration. J Biol Chem 281: 32122–32130 Zhu G, Jaskiewicz E, Bassi R, Darling DS, Young WW, Jr (1997) Beta 1,4 N-acetylgalactosaminyltransferase (GM2/GD2/GA2 synthase) forms homodimers in the endoplasmic reticulum: a strategy to test for dimerization of Golgi membrane proteins. Glycobiology 7: 987–996 Zolov SN, Lupashin VV (2005) Cog3p depletion blocks vesicle-mediated Golgi retrograde trafficking in HeLa cells. J Cell Biol 168: 747–759

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Nucleotide sugar transporters of the Golgi apparatus Weihan Zhao and Karen J. Colley

Introduction The Golgi apparatus is the major site of protein, lipid and proteoglycan glycosylation. The glycosylation enzymes, as well as kinases and sulfatases that catalyze phosphorylation and sulfation, are localized within the Golgi cisternae in characteristic distributions that frequently reflect their order in a particular pathway (Kornfeld and Kornfeld 1985; Colley 1997). The glycosyltransferases, sulfotransferases and kinases are “transferases” that require activated donor molecules for the reactions they catalyze. For eukaryotic, fungal and protozoan glycosyltransferases these are the nucleotide sugars UDP-N-acetylglucosamine (UDP-GlcNAc), UDP-galactose (UDP-Gal), GDPfucose (GDP-Fuc), CMP-sialic acid (CMP-Sia), UDP-glucuronic acid (UDP-GlcA), GDP-mannose (GDP-Man), and UDP-xylose (UDP-Xyl) (Hirschberg et al. 1998). For the kinases, ATP functions as the donor, while for the sulfotransferases, adenosine 30 -phosphate 50 -phosphate (PAPS) acts as the donor (Hirschberg et al. 1998). The active sites of all these enzymes are oriented towards the lumen of the Golgi cisternae. This necessitates the translocation of their donors from the cytosol into the lumenal Golgi compartments. In this chapter we will focus on the structure, function and localization of the Golgi nucleotide sugar transporters (NSTs), and highlight the diseases and developmental defects associated with defective transporters. We direct the reader to several excellent reviews on Golgi transporters for additional details and references (Hirschberg et al. 1998; Berninsone and Hirschberg 2000; Gerardy-Schahn et al. 2001; Handford et al. 2006; Caffaro and Hirschberg 2006).

The identification of NSTs and the diseases and defects in development caused by mutant transporters Abundant evidence now exists for the importance of glycoconjugates in both development and in fundamental processes in adult organisms (Varki 1993; Haltiwanger and Lowe 2004). The critical role of NSTs and the maintenance of nucleotide sugar levels in the glycosylation of proteins, lipids and proteoglycans has been highlighted by a number of transporter mutants that lead to developmental defects in model organisms such as C. elegans and Drosophila, to decreased virulence of parasites such as Leishmania, and to severe diseases in humans and cattle.

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In early studies, mutants exhibiting altered glycosylation in both mammalian cell lines and other organisms, such as yeast and the protozoan parasite Leishmania, were isolated and found defective in nucleotide sugar transport by biochemical analysis (Ballou et al. 1991; Descoteaux et al. 1995; Herman and Horvitz 1999; Patnaik and Stanley 2006). In these analyses, investigators quantified the transport of radiolabeled sugar nucleotides into sealed vesicles under different conditions using filtration or centrifugation to separate the vesicles from the assay medium (reviewed in Hirschberg et al. 1998). For example, Deutscher et al. (1984) demonstrated that Lec2 CHO cells possessed only 2% of the CMP-Sia transport activity of wild type CHO cells, while others demonstrated similar decreases in UDP-GlcNAc transport activity in the Kluyveromyces lactis mnn2-2 mutant (Abeijon et al. 1996a), in UDP-Gal transport activity in the MDCKII-RCAr mutant (Brandli et al. 1988), and in GDP-Man transport activity in both the L. donovani C3PO mutant and in the Saccharomyces cerevisiae vrg4 mutant (Ma et al. 1997; Dean et al. 1997). These studies demonstrated that nucleotide sugar transport was absolutely required for glycosylation in mammalian, yeast and protozoan cells and provided investigators with a way to clone the defective transporters by complementation. The cloning of NST coding sequences opened the way to further characterization of the structure and function of the transporters by expression in heterologous systems and reconstitution into proteoliposomes. This also allowed investigators to identify inactivating NST mutations leading to developmental defects and human disease.

Defects in the GDP-fucose and CMP-Sia transporters lead to two congenital disorders of glycosylation Leukocyte adhesion deficiency syndrome type II (LAD II), also called congenital disorder of glycosylation (CDG) IIc, is a rare autosomal recessive human syndrome characterized by a general reduction of fucose in glycoconjugates due to a deficiency in GDP-Fuc transport (reviewed in Hirschberg 2001; Becker and Lowe 1999). Patients have an abnormal facial appearance and exhibit severe psychomotor and growth retardation, recurrent infections, and periodontitis. Using patient fibroblasts and screening for recovery of glycocon€ bke et al. (2001) and Lu € hn et al. jugate fucosylation in transformants, both Lu (2001) cloned the human and C. elegans GDP-fucose transporters, respectively. These investigators and Helmus et al. (2006) have identified several specific mutations in LADII/CDG IIc patients that lead to disease. The murine CMP-Sia transporter was cloned by complementation of sialylation deficient Lec2 CHO cells and its activity verified by heterologous expression in S. cerevisiae (Eckhardt et al. 1996; Berninsone et al. 1997). Recently, inactivating mutations in this transporter were found to result in a new CDG type II that was diagnosed in a 4-month-old boy (Willig et al. 2001; Martinez-Duncker et al. 2005). Decreased sialylation in the patient led to macrothrombocytopenia, neutropenia, and complete lack of the sialyl Lewis X antigen on polymorphonuclear cells. The patient experienced progressive

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hemorrhaging, respiratory distress syndrome, and opportunistic infections. Ultimately, complications including pulmonary viral infection, massive pulmonary hemorrhage, and respiratory failure led to death at the age of 37 months (Willig et al. 2001).

Drosophila Fringe connection and C. elegans SQV7, two multi-substrate NSTs required for signaling and cell interactions during development The Drosophila Fringe connection transporter was cloned by Selva et al. (2001) and Goto et al. (2001) who identified mutants in Fringe connection in screens for segment polarity and limb defects. These mutants had defects in Fringe-dependent Notch signaling, and in the Wingless/Wnt, Hedgehog, and fibroblast growth factor signaling pathways that require heparan sulfate expression. Fringe is a GlcNAc transferase that modifies O-linked fucose residues on the Notch receptor’s epidermal growth factor repeats, and this modification differentially modulates the binding of Notch to receptors and its signaling pathways (Haltiwanger and Lowe 2004). Accordingly, Fringe connection is a Golgi localized multi-substrate nucleotide sugar transporter that transports UDP-GlcNAc, UDP-GlcA, and UDP-Xyl (Selva et al. 2001). Other groups cloned putative human orthologs of Fringe connection (Suda et al. 2004; Ishida et al. 2005), and over expression of one of these proteins in mammalian cells increased surface levels of heparan sulfate, consistent with the activity of Drosophila Fringe connection (Suda et al. 2004). C. elegans sqv mutants exhibit a squashed vulval phenotype and a reduction in hermaphrodite fertility (Herman et al. 1999). All eight of these mutant genes encode proteins involved in different aspects of proteoglycan biosynthesis (Herman and Horvitz 1999; Hwang and Horvitz 2002). The sqv7 gene encodes an NST that transports UDP-GlcA, UDP-GlcNAc, and UDP-Gal in a competitive and non-cooperative fashion (Berninsone et al. 2001). Surprisingly, two other C. elegans NSTs, the SRF-3 and CO3H5.2 proteins, are redundant with SQV-7 and each other, and they exhibit a dramatically different noncompetitive and simultaneous mechanism. The implications of this will be discussed below.

A defective UDP-GlcNAc transporter leads to complex vertebral malformation in cattle The yeast and canine UDP-GlcNAc transporters were cloned by complementation of the transporter defect the K. lactis mnn2-2 mutant (Abeijon et al. 1996b; Guillen et al. 1998). Interestingly, the sequence similarity of these two functionally equivalent transporters from different species is very low (22%), but not uncommon among transporters with the same specificity from different species. This and the high sequence similarities observed between transporters with different specificities, highlights the importance of biochemically verifying the true substrates of recombinant NSTs (Caffaro and Hirschberg 2006). Recently, a recessively inherited disease in cattle, complex vertebral malformation, was found to be the result of a missense mutation in

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the bovine UDP-GlcNAc transporter (Thomsen et al. 2006). The disease, which results in severe malformation of the vertebrae, abortion of fetuses, and perinatal death, has been reported in cattle all over the world. In fact, it was reported that approximately 30% of the elite sires in Japan and Denmark are carriers for this disease (Thomsen et al. 2006).

The GDP-mannose transporter is required for the virulence of parasites and is an essential protein in yeast Protozoans and yeast differ from vertebrates in that they require the translocation of GDP-Man into the Golgi lumen for extensive mannosylation of their glycoconjugates. GDP-Man transporters were cloned by complementation of the S. cerevisiae vrg4 mutant and the L. donvani C3PO mutant (Descoteaux et al. 1995; Ma et al. 1997; Poster and Dean 1996). The importance of this transporter in both organisms is underscored by the fact that vrg4 is an essential gene in yeast, and by the requirement for GDP-Man transport and the biosynthesis of mannose-containing surface glycoconjugates for Leishmania virulence. The latter observation identifies the Leishmania LPG2 GDP-Man transporter as a possible drug target for the treatment of Leishmaniasis.

Nucleotide sugar transporter specificity and mechanism In early studies, biochemical assays employing topologically correct membrane vesicles were used to identify and characterize NST activities. These initial studies showed that transport in most cases is organelle specific, is temperature sensitive, saturable (Kms of 1–10 mM), concentrates nucleotide sugars 50- to 100-fold in Golgi vesicles/liposomes relative to their concentration in the assay medium, does not require ATP, is not altered by ionophores, and that NSTs are antiporters (Hirschberg et al. 1998). These initial observations were verified by expression of recombinant NSTs in heterologous systems and the reconstitution of purified, recombinant transporters into proteoliposomes. Importantly, these more recent studies demonstrated that single transporter proteins were sufficient for transport activity and revealed surprising multi-substrate specificities and unique mechanisms for some NSTs.

Antiporter mechanism The ability of NSTs to function as antiporters, where the nucleotide sugar is stoichiometrically exchanged for the corresponding nucleoside monophosphate (see Fig. 1), was supported by early studies which showed that preloading nucleoside monophosphates into Golgi membrane vesicles, or proteoliposomes containing transporters, stimulated the transport of their respective nucleotide sugars into the lumen of these vesicles (Hirschberg et al. 1998). More recently both the recombinant L. donovani LPG2 GDP-Man transporter and the murine CMP-Sia transporter were reconstituted into

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A

B

Figure 1. The nucleotide sugar transport/antiport cycle in the Golgi apparatus. (A) GDP-Man, which is synthesized in the cytoplasm, is transported into the Golgi lumen by the GDP-Man transporter. In the Golgi lumen, GDP-Man is a substrate for mannosyltransferases (triangle), which transfer Man to glycoconjugate substrates (rectangle). GDP, the other product of the transfer reaction, is converted by a lumenal nucleoside diphosphatase (oval) to GMP. The export of GMP to the cytosol is coupled to the import of GDP-Man. (B) This type of antiport mechanism occurs for the exchange of CMP-Sia and CMP using a distinct CMP-Sia transporter. The major difference is that no diphosphatase activity is needed because CMP is released following the sialyltransferase reaction.

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phosphatidylcholine liposomes and their specificity and mechanism reevaluated (Segawa et al. 2005; Tiralongo et al. 2006). In both cases investigators observed that preloading the liposomes with the corresponding nucleotide monophosphate yielded a 3-fold higher initial rate of transport relative to liposomes that were not preloaded, again supporting an antiporter mechanism. Other work by Tiralongo et al. (2006) using the reconstituted murine CMP-Sia transporter showed that the rate of CMP-Sia transport was also stimulated under “equilibrium exchange conditions” and suggested that this transporter is a simple mobile carrier with a binding site that alternates between both sides of the membrane. Most biochemical evidence suggests that the nucleotide monophosphate is exchanged for the corresponding nucleotide sugar (reviewed in Hirschberg et al. 1998). This requires that the nucleoside diphosphates released after transfer of the sugar to the glycoconjugate substrate are converted to nucleoside monophosphates by diphosphatases (Fig. 1A). Of course, CMP-Sia is the one exception because CMP is directly released following sugar transfer (Fig. 1B). Additional evidence for the importance of disphosphatase activities in the nucleotide sugar antiporter mechanism came from the finding that deletion of the S. cerevisiae Golgi guanosine diphosphatase (Gda1) decreased GDP-Man transport into membrane vesicles and led to a partial defect in the addition of Man to both glycoproteins and glycolipids (Abeijon et al. 1993; Berninsone et al. 1994). Other work by D’Alessio et al. (2005) demonstrated that GDP-Man dependent glycosylation is reduced but not eliminated in nucleoside diphosphatase mutants in yeast, and suggested that other mechanisms may lead to nucleoside monophosphate translocation. Recent studies by Muraoka et al. (2007) in which the transporter mechanism was evaluated for the endoplasmic reticulum (ER) localized human UGTrel7 transporter, that is capable of transporting UDP-Gal, UDP-GlcA and UDPGlcNAc, and the Golgi localized Drosophila Fringe connection transporter, suggested that the former transporter may be a UDP-sugar/UDP-sugar antiporter, while the latter may use UDP as efficiently as UMP as an antiport substrate.

Multi-specificity and redundancy of NSTs Several NSTs have been identified as multi-substrate transporters including the C. elegans SQV-7 transporter and the Drosophila Fringe connection transporter described above. Other multi-substrate and redundant NSTs emerged as investigators searched the human, Drosophila and yeast genomes for putative nucleotide sugar transporters, and subjected these newly cloned putative transporters to extensive analyses using multiple nucleotide sugars (Segawa et al. 2002; Muraoka et al. 2001; Ashikov et al. 2005). For example, the UDP-Gal transporter was first cloned by complementation of mammalian and yeast mutants (Miura et al. 1996; Tabuchi et al. 1997). Later, Segawa et al. (2002) cloned the Drosophila UDP-Gal transporter and found that both the human UDP-Gal transporter 1 (hUGT1) and the Drosophila transporter were

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specific for both UDP-Gal and UDP-GalNAc. More recently, Capul et al. (2007) identified two genes (LPG5A and LPG5B) in Leishmania major that encode UDP-Gal transporters with partially overlapping activity. The multi-specificity and redundancy of NSTs raised new questions concerning mechanism and potential differences in the roles of NSTs with similar substrate specificities. Investigators have now begun to address some of these questions. The C. elegans genome encodes 18 putative NSTs based on sequence homology with transporters from other species, but the transport of only seven nucleotide sugars is required for glycosylation, and a similar situation exists in humans (Caffaro et al. 2007; Martinez-Duncker et al. 2003). Three redundant, multi-substrate specific C. elegans NSTs, SQV7, SRF-3, and CO3H5.2, have been characterized. As described above, the SQV-7 protein is a multi-substrate transporter that is specific for UDP-GlcA, UDP-GlcNAc, and UDP-Gal and transports these substrates in a competitive and non-cooperative fashion (Berninsone et al. 2001). In contrast to the sqv mutants, the C. elegans srf mutants exhibit no obvious behavioral or morphological changes, but do have defects in cell surface molecules that alter their binding to antibodies and lectins and block infection/colonization by parasites (reviewed in Hoflich et al. 2004). The srf-3 gene encodes a transporter that is specific for both UDP-Gal and UDP-GlcNAc (Hoflich et al. 2004), while the CO3H5.2 gene encodes a transporter specific for UDP-GlcNAc and UDPGalNAc (Caffaro et al. 2006). Surprisingly, the CO3H5.2 and SRF-3 transporters use a simultaneous and non-competitive substrate transport mechanism that differs from the competitive mechanism used by the SQV7 transporter (Caffaro et al. 2006, 2007). A deletion of 16 amino acids in the loop between transmembrane (TM) helices 2 and 3 of the CO3H5.2 protein preferentially decreased UDP-GalNAc transport by 85–90%, but did not impact UDP-GlcNAc transport, suggesting two independent translocation sites for these nucleotide sugars (Caffaro et al. 2006). The existence of these three C. elegans transporters that exhibit partially overlapping substrate specificity (UDP-GlcNAc) and expression patterns, led Caffaro et al. (2007) to investigate this redundancy. They used RNAi technology to knock down the CO3H5.2 gene in srf-3 mutants and found that a defect in both transporters led to developmental and morphological changes not observed in the srf-3 mutant alone. This strongly suggested that these two transporters are at least partially redundant and begged the question why redundancy was needed. One possibility, suggested by the investigators, is that certain nucleotide sugars need to be maintained at high levels so that processes requiring these molecules can proceed with high efficiency under a variety of circumstances. In addition, if nucleotide sugar levels drop, different glycosylation pathways can be differentially affected depending upon the affinity of the associated glycosyltransferases for the particular nucleotide sugar. For example, in the MDCKII-RCAr cell mutant where availability of UDP-Gal is limited, a decreased galactosylation is observed for glycoproteins, glycolipids and keratan sulfate proteoglycans,

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while the amounts of chondroitin and heparan sulfate proteoglycans remain relatively normal (Toma et al. 1996). Similarly, in LADII/CDG IIc patients who have a defect in the GDP-Fuc transporter, low dose oral Fuc supplementation therapy partially restores P-selectin-mediated, but not E-selectin-mediated binding of neutrophils (Marquardt et al. 1999a, b). Since the synthesis of P- and E-selectin glycan ligands depends upon different Fuc transferase activities (Huang et al. 2000), it is likely that the enzymes have different affinities for GDP-Fuc and that this leads to differences in glycoconjugate expression under limiting GDP-Fuc levels. Another possible reason that there may be genetic pressure to maintain the expression of rendundant multi-substrate transporters is so that the non-overlapping functions of these transporters can be maintained (Caffaro et al. 2006). A third possibility, is that different transporters with similar specificity function in conjunction with specific glycosylation pathways. This idea is supported by the work of Capul et al. (2007) who cloned two UDP-Gal transporters from L. major (LPG5A and LPG5B) and demonstrated that deficiencies in these transporters impacted the biosynthesis of the two predominant Leishmania surface glycoconjugates differently.

NST structure and sequence requirements for function, trafficking, and localization Topology and oligomerization Hydrophobicity plots and topology prediction algorithms based on the deduced amino acid sequences of NSTs suggest that these proteins are multispanning membrane proteins containing six to ten TM regions with both their amino- and carboxy-termini in the cytosol (Hirschberg et al. 1998) (Fig. 2). Initial studies to define the topology of the K. lactis UDP-GlcNAc transporter in Golgi vesicles have been performed and suggest either a six or eight TM helix topology (Berninsone and Hirschberg 2000). In contrast, Eckhardt et al. (1999) evaluated the membrane topology of the murine CMP-Sia transporter using immunofluorescence microscopy following epitope insertion and selective membrane permeablization, and obtained data supporting a 10 TM helix model for this transporter (Eckhardt et al. 1999). Further studies are needed to define the topological arrangement of other NSTs. Most NSTs are thought to exist as homodimers. For example, the rat liver UDP-GalNAc and GDP-Fuc transporters both migrate with molecular masses of approximately 40 kDa upon denaturing gel electrophoresis, but exhibit molecular masses of 80–90 kDa in native glycerol gradients (Puglielli et al. 1999; Puglielli and Hirschberg 1999). VRG4, the S. cerevisiae GDP-Man transporter has also been found to be a homodimer (Gao and Dean 2000). A carboxy-terminal region of this protein, which includes the last TM helix, is necessary for dimer formation, and truncated proteins lacking this sequence are unstable and rapidly degraded. Interestingly, overexpression of an aminoterminal truncated VRG4 protein in yeast causes a dominant negative growth

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Figure 2. Topology and functional regions of the Golgi NSTs. (A) Topology and TM helices required for transport activity. (i) CMP-Sia transporter: TM helix 7 (TM7) is required for the specificity of this transporter for CMP-Sia, whereas the TM helices 2 and 3 enhance the efficiency of CMP-Sia transport. (ii) UDP-Gal transporter: TM helices 1 (TM1) and 8 (TM8) are necessary but not sufficient for UDP-Gal transport, and other helices in different combinations (2, 3 and 7 OR 9 and 10) must be included with TM1 and TM8 for transport activity. (B) Sequences required for dimerization and ER export. (i) In the GDP-mannose transporter, a carboxyterminal sequence containing the last TM helix is involved in dimerization, whereas the amino-terminal 44 amino acids include an ER export signal. (ii) In the CMP-Sia transporter, a di-isoleucine motif and a terminal valine residue (boxed) at the very carboxy-terminus mediate its ER export.

phenotype. This is believed to be a consequence of the formation of inactive heterodimers of the truncated protein and the endogenous full length protein, and supports the notion that homodimerization of VRG4 is crucial for its function. In contrast LPG2, the Leishmania GDP-Man transporter, which migrates as a hexamer in native glycerol gradients and upon pore-limited native gel electrophoresis (Hong et al. 2000). Further functional studies will be needed to determine whether the hexamer form of this protein is the active state in the membrane.

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Sequence and structural requirements for substrate recognition and transport The identification of mutants in NSTs first gave hints to what regions and amino acid residues are critical for their function. Gao et al. (2001) identified a conserved region in the yeast Vrg4 GDP-Man transporter (amino acids 280–291) as the transporter’s GDP-Man binding site, because mutations in this sequence reduced binding to a photoaffinity substrate analog and led to decreased GDP-Man transport. Gerardy-Schahn and colleagues (Eckhardt et al. 1998; Oelmann et al. 2001) identified the various mutations in the Lec2 and Lec8 complementation groups that lead to defects in the CMP-Sia transporter and UDP-Gal transporter, respectively. While many of the identified mutants of the CMP-Sia transporter were deletions that led to mislocalization and low expression, the Gly189Glu mutant was localized in the Golgi and well-expressed, suggesting that this amino acid was critical for transporter activity per se (Eckhardt et al. 1998). Likewise, the DSer213 and Gly281Asp mutants of the UDP-Gal transporter were localized properly and expressed well, but still inactive. Interestingly, introducing these changes into the CMP-Sia transporter also led to its inactivation, suggesting that these conserved residues are important for general transporter mechanism (Oelmann et al. 2001). Aoki et al. (2001, 2003) identified critical TM helices in both the CMP-Sia and UDP-Gal transporters. These transporters are 43% identical and yet are absolutely specific for their respective substrates. The investigators created chimeras containing TM helices from both transporters and found that CMPSia transporter TM helix 7 was necessary and sufficient for transport of CMPSia when inserted into a UDP-Gal transporter background, while the inclusion of TM helices 2 and 3 enhanced efficiency of transport. In contrast, TM helices 1 and 8 of the UDP-Gal transporter were necessary but not sufficient for UDPGal transport in the context of the CMP-Sia transporter. Only the inclusion of either TM helices 9 and 10, or TM helices 2, 3 and 7 from the UDP-Gal transporter, in addition to TM helix 1 and 8, could generate a chimeric transporter competent to transport UDP-Gal (Aoki et al. 2003).

Sequence requirements for NST ER export and retrieval In eukaryotic secretory pathway, exit of secretory proteins from the ER relies on their sorting into ER-derived COPII-coated vesicles. Much of this sorting is mediated by specific, cytoplasmically exposed signals that can be recognized by subunits of the COPII coat (Barlowe 2003). ER export signals have been found in several NSTs. The amino-terminal 44 amino acids of VRG4, the yeast Golgi GDP-Man transporter, are likely to include an ER export signal because deletion of this region leads to ER accumulation, and fusion of these sequences to related ER proteins promotes their transport to the Golgi apparatus (Gao and Dean 2000). A di-isoleucine motif and a terminal valine in the last four amino acids of the carboxy-terminal cytoplasmic tail of the murine CMP-Sia transporter mediate the ER export of the protein (Zhao et al.

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2006). These signals are independent and both need to be deleted or replaced to abolish ER export. Signals predicted to allow the COPI-coated vesicle-mediated ER retrieval have been found in both ER NSTs and some Golgi NSTs (Martinez-Duncker et al. 2003). The cloning of the gene for a second functional isoform of the human UDP-Gal transporter (hUGT2) predicted that this protein is truncated at its carboxy-terminus (Ishida et al. 1996). Comparison of the localization of the two isoforms revealed that hUGT1 is localized in the Golgi, while hUGT2 is localized in the ER and Golgi (Kabuß et al. 2005). Kabuß et al. (2005) demonstrated that a dilysine motif (LysValLysGlySer) found in the carboxyterminal cytoplasmic tail of hUGT2 was responsible for its ER retrieval and its dual localization. Fusion of this motif is sufficient to redistribute the Golgi CMP-Sia transporter to the ER. A similar dilysine ER retrieval motif also is found in the K. lactis Golgi UDP-GlcNAc transporter, however it is unclear whether this motif actually functions as an ER retrieval signal (Abeijon et al. 1996b). Interestingly, work done by Abe et al. (2004) suggests that a COPImediated retrieval of the GDP-Man transporter to the ER is a critical step in the Golgi localization of this transporter and that lysine residues in its carboxylterminal cytoplasmic tail are necessary for COPI coat interaction and retrieval.

Golgi targeting of NSTs and the organization of glycosylation machinery Many studies on the signals and mechanisms of Golgi glycosylation enzyme localization have revealed that sequences mediating Golgi localization are complex and may reside in different domains of these type II membrane proteins (Colley 1997). Likewise, several redundant mechanisms including those involving lipid partitioning, oligomerization, and retrieval, may be used to maintain Golgi enzymes in their resident cisternae (Colley 1997; Mironov et al. 2005). When considering the organization of glycosylation pathways in the Golgi, it is tempting to speculate that NSTs are co-compartmentalized with the glycosyltransferases that use their nucleotide sugar substrates as donors and that they may even form functional complexes. For this reason we were surprised to find that the CMP-Sia transporter showed a more restricted medial–trans Golgi localization than might be indicated by the rather broad Golgi distribution of sialyltransferases involved in both the sialylation of glycoproteins and glycolipids (Zhao et al. 2006). This finding, together with others (D’Alessio et al. 2003; Kabuß et al. 2005), suggests that CMP-Sia as well as other nucleotide sugars move freely in the lumen of the Golgi apparatus. This is consistent with recent evidence that the Golgi cisternae are more interconnected than once believed (Mironov et al. 2005). The identity of the sequences that mediate the Golgi localization of the NSTs has not been widely investigated, however the sequence requirements for the Golgi localization of some viral multi-spanning membrane proteins have been determined. The first TM helix of the avian coronavirus E1 protein is sufficient to localize two cell surface proteins to the Golgi and is likely to

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play a role in mediating E1 Golgi localization (Machamer and Rose 1987). Later studies showed that uncharged polar residues that line one face of this TM helix are important for Golgi localization (Swift and Machamer 1991). Work by Locker et al. (1994) suggested that the cytoplasmic tail of the mouse hepatitis virus M protein plays a role in its Golgi localization. We have recently found that, while the CMP-Sia transporter cytoplasmic sequences have no direct role in Golgi localization, fusing the first TM helix of the transporter to the lumenal sequences of an inefficiently retained Golgi sialyltransferase can reduce the level of its Golgi exit (W. Zhao and K. J. Colley, unpublished data). This suggests that, like the Golgi localization of the infectious bronchitis virus E1 protein, the first TM helix of the CMP-Sia transporter may be involved in its Golgi localization. NST Golgi localization could also be mediated by interactions with glycosyltransferases. Complex formation between a transporter and its corresponding glycosyltransferase would presumably enhance the efficiency of the glycosylation reaction by facilitating the transfer of the nucleotide sugar to the glycosyltransferase. Along these lines, Sprong et al. (2003) found that a portion of the Golgi UDP-Gal transporter can associate with the ER localized UDP-galactose:ceramide galactosyltransferase to allow UDP-Gal import into this compartment. However, evidence for other functional NST-glycosyltransferase complexes is lacking. Radiation inactivation studies suggest that the galactosyl- and sialyltransferases are not in functional complexes with the corresponding transporters (Fleischer et al. 1993). Moreover, redistribution studies show that there is no complex formation between the CMP-Sia transporter and the corresponding sialyltransferases (Zhao et al. 2006). Although weak interactions between glycosyltransferases and their respective NSTs are still possible, these results suggest that NSTs may not rely on glycosyltransferases for their Golgi localization.

Conclusions Investigators have made great progress in identifying NSTs and defining their substrate specificity. However, many questions remain concerning the mechanism of nucleotide sugar transport, the roles and expression of multisubstrate and redundant NSTs, the potential connection of redundant transporters with specific glycosylation pathways, the mechanisms of NST Golgi localization, and how NSTs and glycosyltransferases are organized within the Golgi apparatus. For example, how is the developmental, cellular and tissue expression of redundant and multi-substrate NSTs controlled? Do differences in sub-Golgi localization of NSTs with similar substrate specificity allow these transporters to be compartmentalized with glycosylation enzymes in different pathways? How does the simultaneous transport of two substrates occur in one transporter? What are the precise interactions mediating nucleotide sugar recognition and transport? How are NSTs localized in the Golgi, and how are they organized vis a vis their respective glycosyltransferases to

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promote efficient glycosylation? The realization that mutant NSTs lead to human disease and developmental defects has and will continue to generate interest in these proteins and will hopefully stimulate additional research to answer these many questions.

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Swift AM, Machamer CE (1991) A Golgi retention signal in a membrane-spanning domain of coronavirus E1 protein. J Cell Biol 115: 19–30 Tabuchi M, Tanaka N, Iwahara S, Takegawa K (1997) The Schizosaccharomyces pombe gms1 þ gene encodes an UDP-galactose transporter homologue required for protein galactosylation. Biochem Biophys Res Commun 232: 121–125 Thomsen B, Horn P, Panitz F, Bendixen E, Petersen AH, Holm L-E, Nielsen VH, Agerholm JS, Arnbjerg J, Bendixen C (2006) A missense mutation in the bovine SLC35A3 gene, encoding a UDP-N-acetylglucosamine transporter, causes complex vertebral malformation. Genome Res 16: 97–105 Tiralongo J, Ashikov A, Routier F, Eckhardt M, Bakker H, Gerardy-Schahn R, Von Itzstein M (2006) Functional expression of the CMP-sialic acid transporter in Escherichia coli and its identification as a simple mobile carrier. Glycobiology 16: 73–81 Toma L, Pinhal MA, Dietrich CP, Nader HB, Hirschberg, CB (1996) Transport of UDPgalactose into the Golgi lumen regulates the biosynthesis of proteoglycans. J Biol Chem 271: 3897–3901 Varki A (1993) Biological roles of oligosaccharides: all of the theories are correct. Glycobiology 3: 97–130 Willig T-N, Breton-Gorius J, Elbim C, Mignotte V, Kaplan C, Mollicone R, Pasquier C, lot F, Cartron J-P, Gougerot-Pocidalo M-A, Debili N, Guichard, Filipe A, Mie Dommergues J-P, Mohandas N, Tchernia G (2001) Macrothrombocytopenia with abnormal demarcation membranes in megakaryocytes and neutropenia with a complete lack of sialyl-Lewis-X antigen in leukocytes – a new syndrome? Blood 97: 826–828 Zhao W, Chen TL, Vertel BM, Colley KJ (2006) The CMP-sialic acid transporter is localized in the medial–trans Golgi and possesses two specific endoplasmic reticulum export motifs in its carboxyl-terminal cytoplasmic tail. J Biol Chem 281: 31106–31118

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Luminal lectins Beat Nyfeler, Eva Koegler, Veronika Reiterer and Hans-Peter Hauri

Introduction Asparagine-linked glycosylation (N-glycosylation) is a major post-translational modification of secretory and membrane proteins and influences important physical protein properties such as conformation, stability and solubility (Helenius and Aebi 2001). The majority of proteins that enter the secretory pathway receives multiple N-linked glycans. N-glycosylation is initiated cotranslationally in the lumen of the endoplasmic reticulum (ER) by oligosaccharyltransferase. This multisubunit protein complex scans nascent proteins for N-glycosylation consensus sequences (Asn–X–Ser/Thr) and catalyzes the transfer of a 14-saccharide core glycan to the asparagine residue (Fig. 1). About two-thirds of all consensus sites are glycosylated. After conjugation to the protein, the 14-saccharide core is trimmed in ER and Golgi by glycosidases and extended in the Golgi by glycosyltransferases (Kornfeld and Kornfeld 1985). ER glucosidases I and II remove the three glucose (Glc) residues, whereas ER a1,2 mannosidase I and Golgi a1,2 mannosidases 1A, 1B and 1C trim the a1,2-linked mannoses (Man). In the Golgi, two additional Man residues are cleaved and the N-glycans undergo complex glycosylation by the addition of N-acetylglucosamine (GlcNAc), fucose, galactose and sialic acid residues. After traversing the Golgi, glycoproteins carry various N-linked glycans differing in composition and structure. This heterogeneity allows mature glycoproteins to fulfill a plethora of functions including the presentation of interaction sites for other molecules (Varki 1993). In contrast, nascent glycoproteins in the early secretory pathway, termed high-mannose glycoproteins, display only few but distinct oligosaccharide structures which function as recognition tags for different sugar-binding proteins (lectins). With their carbohydrate recognition domain (CRD), the lectins bind newly synthesized glycoproteins and control their folding, degradation, transport and sorting. Here, we provide an overview of the different animal lectins localized to the lumen of the secretory pathway (Table 1) and describe them grouped according to their proposed function.

Functions of luminal lectins Protein folding and quality control in the ER: calnexin, calreticulin The lumen of the ER features a specialized cellular environment for protein folding and modification. Multiprotein networks of general chaperones,

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Figure 1. The N-linked core glycan. The 14-saccharide core glycan contains two N-acetylglucosamine (GlcNAc, boxes), nine mannose (Man, circles) and three glucose (Glc, triangle) residues. The nomenclature of the different branches and the type of glycosidic linkage is indicated.

peptidyl prolyl isomerases and thiol oxidoreductases help newly synthesized proteins to gain their native conformation. Important factors in the folding process of glycoproteins are calnexin (CNX, Wada et al. 1991) and calreticulin (CRT, Fliegel et al. 1989), two homologous calcium-binding lectins which localize to the ER and function as molecular chaperones (Ou et al. 1993). CNX is a type I transmembrane protein of 90 kDa, whereas CRT is a 60-kDa soluble luminal protein that is retained in the ER by its C-terminal KDEL retention signal. Both proteins are monomeric and display a similar luminal fold consisting of a globular and an extended domain. The globular domain contains the CRD and has a b-sandwich fold related to plant legume lectins. The extended domain is proline-rich, hence termed P domain, and comprises two b-strands forming a long hairpin. Bound substrate glycoproteins are believed to localize to the space between the extended P domain and the globular lectin domain. How are glycoproteins bound? Although the contribution of protein–protein interactions to substrate capture is still a matter of debate (Williams 2006), binding of carbohydrate moieties by the CRDs of CNX and CRT is well documented and accepted. The in vitro established carbohydrate specificity for a single a1,3-linked Glc and three Man residues in the A branch is in line with the preferential binding of monoglucosylated glycoproteins observed in vivo. Hence, CNX and CRT bind glycoproteins immediately after addition of the N-linked core glycan and removal of the two outermost Glc residues by ER glycosidases I and II. The majority, if not all, glycoproteins bind to either CNX, CRT or both (Helenius et al. 1997). If the association with the two lectins is inhibited, for instance by the ER glucosidase inhibitor castanospermine, glycoprotein folding is more rapid but also more error-prone which results in more misfolded and degraded protein species. Therefore, CNX and CRT increase the efficiency of protein folding by (i) slowing down the folding reaction, (ii) preventing off-pathway folding reactions and aggregation, and (iii) favoring native disulfide-bridge forma-

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Table 1. Overview of luminal animal lectins Lectin

Intracellular localization

Carbohydrate specificity

Proposed function

Calnexin

ER

Calreticulin

ER

ER a1,2mannosidasea

ER

Protein folding Protein folding Protein degradation

EDEM1

ER

EDEM2

ER

EDEM3

ER

Yos9p(yeast)a

ER

Monoglucosylated A branch Monoglucosylated A branch Man9GlcNAc2: Man trimming to Man5–6GlcNAc2 High-mannose (Man8GlcNAc2) High-mannose (Man8GlcNAc2) High-mannose (Man8GlcNAc2) Mannose?

ERGIC-53

ERGIC

ERGLa VIP36

ER ERGIC, Golgi

VIPL

ER

CI-M6PR

Trans-Golgi, endosomes, plasma membrane

CD-M6PR

Trans-Golgi, endosomes, plasma membrane

a

High-mannose, broad specificity (Man9GlcNAc2 to (Man6GlcNAc2) ? De-glucosylated A branch De-glucosylated A branch Mannose 6-phosphatemonoester, Mannose 6-phosphatediester

Mannose 6-phosphatemonoester

Protein degradation Protein degradation Protein degradation Protein degradation ER-ERGIC transport

Protein sorting in ER? Golgi–ER transport? intra-Golgi transport? Protein sorting in ER? Transport of lysosomal proteins from trans-Golgi and plasma membrane to endosomes, IGF-II receptor Transport of lysosomal proteins from trans-Golgi to endosomes.

Lectin activity uncertain

tion. The latter is achieved by exposing the substrate glycoproteins to the oxidoreductase ERp57 which binds to the P domain of CNX and CRT and catalyzes the formation and isomerization of disulfide bonds. The association of glycoproteins with CNX and CRT is transient and terminated by the action of ER glucosidase II that removes the remaining single Glc residue of the A branch. The resulting deglucosylated high-mannose oligosaccharide structure shows low affinity toward CNX and CRT leading to the release of glycoproteins into the lumen of the ER. Correctly folded glycoproteins can

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now exit the ER in coat protein II (COPII)-coated vesicles. While a single association with CNX and CRT suffices for some glycoproteins to gain their native conformation, others are still misfolded. These misfolded proteins are recognized by UDP-glucose:glycoprotein glucosyltransferase (UGT1), a folding sensor which re-glucosylates nearly native glycoproteins. Terminally misfolded proteins are not recognized and are ultimately eliminated by ER-associated degradation (ERAD). Hence, UGT1 is the ER quality control component that monitors the conformational status of folding glycoproteins. Re-glucosylation of non-native glycoproteins allows multiple rounds of association with CNX and CRT, a process termed calnexin/calreticulin cycle (Parodi 2000). Having two lectins with different topologies broadens the scope of glycoproteins substrates that can be assisted in this cycle. In addition to their involvement in protein folding, CNX and CRT seem to have additional functions in development. Although CNX- and CRT-deficient cell lines can survive in culture, knockout mice show severe phenotypes. A knockout of CNX results in motor disorders associated with a dramatic loss of nerve fibers and in reduced survival. A CRT knockout is embryonically lethal.

ER-associated protein degradation: ER mannosidase, EDEM, Yos9p Terminally misfolded proteins need to be removed from the ER as their accumulation may compromise the folding and secretory capacity of the cell since misfolded proteins have a tendency to aggregate. If refolding by the CNX/CRT cycle is unsuccessful, terminally misfolded glycoproteins are retrotranslocated into the cytosol and degraded by the proteasome. This process, known as ERAD, is determined by mannose trimming as indicated by the fact that kifunensin, an inhibitor of ER mannosidase I (ERManI), or deoxymannojirimycin inhibit ERAD. Mannose trimming exposes sugars that do not allow reglucosylation by UGT1 and association with CNX/CRT. In yeast trimming of a single mannose residue from branch B suffices to divert a glycoprotein to the ERAD pathway. In contrast, mammalian glycoproteins require more extensive mannose trimming down to Man5–6 (Molinari 2007). In particular, removal of the terminal mannose of the A branch prevents reglucosylation by UGT1 and binding to CNX/CRT with additional contribution of mannose trimming from branches B and C. Overexpression and knockdown studies indicate that mannose trimming down to Man5–6 is catalyzed by ERManI (Avezov et al. 2008). Overexpressed ERManI localizes to a percentriolar ER-derived structure that may constitute a specific ER quality compartment but confirmation of this notion requires localization of endogenous ERManI. Moreover, it is presently unclear if ERManI also operates as a mannose lectin routing glycoproteins to ERAD. Additional factors in ERAD are the recently discovered putative mannosebinding proteins EDEM1, EDEM2 and EDEM3 which are homologous to ERManI (Olivari and Molinari 2007). EDEMs (for ER degradation-enhancing a-mannosidase-like proteins) have different tissue distributions and it is currently unclear if they are functionally redundant. Endogenous EDEM1 is concentrated in ER buds that give rise to 150 nm vesicles lacking COPII and

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ERGIC-53 but contain misfolded alpha1-antitrypsin and occasionally Derlin-2 that may constitute part of the retrotranslocation channel (Zuber et al. 2007). Whether these structures relate to the ERManI compartment, remains to be shown. Up- or downregulation of EDEMs modulates degradation of foldingdefective glycoproteins. For instance, EDEM1 enhances removal of the terminal branch A mannose (Ermonval et al. 2001; Olivari et al. 2006), but the mechanism of action is not fully understood. EDEMs may act as mannosidases that determine the rate of ERAD substrate demannosylation. Alternatively, they may act as classical chaperones by preventing aberrant oligomerization and aggregate formation (Hosokawa et al. 2006). Yet another possibility is a lectin function in bridging misfolded proteins to the retrotranslocation pore in view of EDEMs association with derlin 2 and 3. A second class of lectin-like ERAD factors is Yos9p in yeast (Kanehara et al. 2007). Although the name Yos9p is derived from the mammalian protein OS9, Yos9p's mammalian orthologue has not been characterized. Yos9 shares a lectin-like domain with the mannose 6-phosphate receptors (Whyte and Munro 2001) and is required for efficient ERAD. In yeast Yos9p binds to misfolded glyproteins, and misfolded glycoproteins degrade very poorly in the absence of Yos9p. Although the degradation of non-glycosylated variants shows no dependence on Yos9p, the role of glycans in this process has not been clarified. Like proposed for EDEMs, Yos9p may operate in guiding terminally misfolded glycoproteins to the retrotranslocation pore.

Protein transport: L-type lectins Upon correct folding, native proteins are exported from the ER in COPIIcoated vesicles that mediate transport to the ER Golgi intermediate compartment (ERGIC). In many cases protein export from the ER is selective. In this process transmembrane proteins can directly interact with the cytosolic COPII coat and thereby convey their selective incorporation into transport vesicles. In contrast, for soluble proteins, selective recruitment into COPII vesicles requires the assistance of transmembrane receptors, termed cargo receptors (Appenzeller et al. 1999; Baines and Zhang 2007). In mammalian cells, the best characterized cargo receptor is ERGIC-53 (Hauri et al. 2000b) that belongs to a family of four related lectins also comprising ERGL, VIP36 and VIPL. These four proteins are all type I membrane proteins, share a conserved luminal CRD that is homologous to leguminous plant lectins (L-type lectins), localize to the ER/ Golgi interface, and are believed to assist glycoprotein sorting and transport in the early secretory pathway.

ERGIC-53 ERGIC-53 (ER Golgi intermediate compartment protein of 53 kDa, gene name: LMAN1) is a dynamic protein which cycles between ER and ERGIC with a minor cycling route via the cis-Golgi (Appenzeller-Herzog and Hauri 2006; Schweizer et al. 1988). At steady state ERGIC-53 is mainly localized to the ERGIC. The

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cycling properties of ERGIC-53 are determined by sorting motifs in its carboxyl-terminus, which interact with cytosolic coat components. A di-phenylalanine motif interacts with the Sec24 subunit of the COPII coat and mediates ER export, and a di-lysine motif binds COPI and retrieves ERGIC-53 back from ERGIC and cis-Golgi. The luminal domain of ERGIC-53 contains the CRD and a coiled-coil stalk domain that allows oligomerization into homo-dimers and hexameres. The crystal structure of the CRD of ERGIC-53 uncovered two Ca2+binding sites and an overall b-sandwich structure composed of one concave and one convex b-sheet with the ligand-binding site located in a negatively charged cleft. ERGIC-53 binds high-mannose glycoproteins and its CRD shows in vitro a broad specificity but low affinity towards various high-mannose carbohydrate structures. What is the function of ERGIC-53’s lectin domain? By its CRD, ERGIC-53 can capture glycoproteins in the lumen of the ER and through its cytosolic di-phenylalanine motif bind to COPII, thereby recruiting soluble cargo glycoproteins into COPII vesicles. Hence, ERGIC-53 acts as a cargo transport receptor by mediating ER-to-ERGIC transport of some glycoproteins. In addition, ERGIC-53 facilitates the assembly of immunoglobulin M polymers in the lumen of the ER. The identified cargo glycoproteins of ERGIC-53 include blood coagulation factors V (FV) and VIII (FVIII), cathepsin C, and cathepsin Z. Loss-of-function mutations in human ERGIC-53 result in the clinical manifestation of combined FV and FVIII deficiency. Patients lacking ERGIC-53 show a bleeding phenotype due to reduced secretion of FV and FVIII into the blood plasma. Interestingly, efficient secretion of FV and FVIII requires an additional factor known as MCFD2 (Zhang et al. 2003). MCFD2 is a small soluble protein that interacts with ERGIC-53 in a calcium-dependent but lectin activity-independent manner and is believed to recruit FV and FVIII to ERGIC-53. In contrast, MCFD2 is dispensable for the interaction of ERGIC-53 and cathepsin Z. In this interaction ERGIC-53 recognizes a high-mannose N-glycan and a folded b-hairpin peptide structure in cathepsin Z. The requirement of a combined oligosaccharide/ peptide structure may limit the repertoire of possible ERGIC-53 cargo proteins and has led to the suggestion that ERGIC-53 might function in secondary quality control by capturing only native cargo proteins. Cargo protein release from ERGIC-53 occurs in the ERGIC probably due to a drop in pH which results in the protonation of His178, the loss of a calcium ion, and lowered affinity for glycans.

VIP36 The ERGIC-53-related lectin VIP36 (vesicular integral-membrane protein of 36 kDa, gene name: LMAN2, Fiedler et al. 1994) localizes to the early secretory pathway and cycles between ER, ERGIC, and cis-Golgi (Fullekrug et al. 1999). If highly overexpressed, VIP36 can be found in the Golgi, apical and basolateral vesicles, and the plasma membrane. In its cytoplasmic domain VIP36 carries a putative phenylalanine/tyrosine ER export motif and a di-basic motif (KR) that

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may operate in retrieval although it does not entirely conform to the established di-lysine ER retention/retrieval consensus signal. Less efficient retrieval than observed for ERGIC-53 may explain why VIP36 has access to the transGolgi and why there is some controversy as to whether VIP36 also operates in the late secretory pathway. In contrast to ERGIC-53, VIP36 is N-glycosylated but lacks the luminal coiled-coil domain required for oligomerization. Hence, VIP36 likely functions as a monomer. The luminal CRD was crystallized recently (Satoh et al. 2007). It exhibits a b-sandwich fold composed of two antiparallel b-sheets. Although not supported by all studies, VIP36 seems to bind carbohydrates in a calcium-dependent manner and the crystal structure identified Asp131, Asn166 and His190 as calcium-binding residues. It was postulated that VIP36 may function in post-ER quality control of glycoproteins in the Golgi by recycling inadequately trimmed glycoproteins (Hauri et al. 2000a). Recent biochemical data lend support to this hypothesis. VIP36 shows highest affinity for the deglucosylated high-mannose A branch as assessed by affinity chromatography (Kamiya et al. 2005). The binding strength changes with pH and shows a bell-shaped dependence with an optimum at pH 6.5. This optimum corresponds to the pH of the medial/trans-Golgi. Thus, glycoproteins with an untrimmed A branch that have inadvertently escaped the ER would be captured by VIP36 in the Golgi and retrieved back to the cis-Golgi or even the ER for another round of mannose trimming. The functional characterization of VIP36 suffers from the lack of characterized cargo glycoproteins and from the fact that overexpression of the protein considerably shifts its localization from the early to the late secretory pathway.

ERGL Human ERGL (ERGIC-53-like protein, gene name: LMAN1L), identified in a prostate-specific EST cluster, is homologous to ERGIC-53 (Yerushalmi et al. 2001). SLAMP (sublingual acinar membrane protein) likely represents the corresponding rat orthologue (Sakulsak et al. 2005). In contrast to the ubiquitously expressed ERGIC-53, ERGL mRNA is found only in a few tissues, including prostate, spleen, salivary gland, cardiac atrium and distinct cells of the central nervous system. The reason for this tissue-specific expression is currently unclear. The amino acid sequence of ERGL reveals a luminal coiledcoil domain which may allow oligomerization. ERGL has a longer C-terminal tail than the other animal L-type lectins and lacks typical transport motifs. No cell line has been found that expresses endogenous ERGL protein but transfection studies with human ERGL have provided some interesting information (Liang, L. and Hauri H.P., unpublished). ERGL is confined to the ER and has a short half life of about 30 min as opposed to ERGIC-53 with a half life of days. Surprisingly, overexpression of ERGL selectively retains ERGIC-53 in the ER by disulfide-bond-mediated interaction, and this interaction reduces mannosebinding activity of ERGIC-53 in an in vitro assay. Together these data suggest that ERGL may function as an ERGIC-53-regulating protein in the ER.

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VIPL VIPL (VIP36-like protein, gene name: LMAN2L) exhibits 43% and 68% sequence similarity to ERGIC-53 and VIP36, respectively. Like VIP36, VIPL is a monomeric and glycosylated membrane protein (Nufer et al. 2003). However, in contrast to VIP36 that has access to the Golgi and is complex glycosylated, VIPL is an ER resident protein that displays high-mannose-type glycans and remains endoglycosidase-H sensitive. The ER localization of VIPL is determined by a di-arginine-based ER retention motif in its cytosolic tail (RKR). In addition, VIPL contains a potential ER export motif (FY) and, if the ER retention motif is mutated, can leave the ER. The luminal CRD of VIPL shows the highest affinity for the three Mana1–2Mana1–2Man residues of the A branch (Kamiya et al. 2008). Glucosylation strongly reduces binding. Sugar-binding of VIPL is calciumdependent and most efficient at pH 7.5–8, which corresponds to the pH of the ER. This pH optimum, together with the carbohydrate specificity, suggests that VIPL binds glycoproteins in the ER. One scenario is that VIPL delivers highmannose glycoproteins, released from the CNX/CRT cycle, to ERGIC-53 for transport. Binding to VIPL might protect high-mannose glycans on native glycoproteins from extensive mannose trimming which would otherwise divert the glycoprotein to the ERAD pathway. Furthermore, VIPL may control the intracellular distribution of other lectins since its overexpression relocalizes ERGIC-53 to the ER. VIPL’s short half-life of 30 min is in line with a potential regulatory function. So far no direct cargo glycoproteins have been identified for VIPL, but a siRNA-based knockdown of VIPL results in reduced secretion of two glycoproteins of unknown identity with a molecular mass of 35 kDa and 250 kDa (Neve et al. 2003).

Lysosomal protein sorting in the Golgi: P-type lectins After export from the ER and transport through the ERGIC folded proteins reach the cis-Golgi. In the cis-Golgi newly synthesized proteins destined for lysosomes are modified by N-acetylglucosamine 1-phosphate at the 6th position of selected mannose residues by the action of GlcNAc-phosphotransferase (Kornfeld and Mellman 1989). From the resulting phosphodiester intermediate the N-acetylglucosamine residue is removed by uncovering enzyme in the TGN resulting in a phosphomonoester (Kornfeld 1987). The secondary or tertiary structure of the protein seems to be crucial for recognition by the N-glycan phosphorylation machinery. Obtaining this modification at one or two mannose residues, lysosomal enzymes can now be distinguished from other proteins traversing the Golgi apparatus. In the TGN the mannose 6-phosphate receptors (MPRs) bind about 50 different acid hydrolases, transport them to and release them in endosomes upon a drop in pH. Further transport to lysosomes occurs in a MPR-independent manner. For completeness it is worth mentioning that some lysosomal hydrolases are targeted to lysosomes in a MPR-independent manner.

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MPRs are the only members of the P-type lectin family and come in two flavours: cation-dependent MPR (CD-MPR; Mr 46 kDa) and cation-independent MPR (CI-MPR; Mr 300 kDa, also termed IGF-II/MPR since it also acts as insulin-like growth factor II receptor). Both MPRs cycle constitutively between TGN, early endosomes, recycling endosomes, late endosomes and plasma membrane, but they are absent from lysosomes. Trafficking is controlled by post-translational modifications such as phosphorylation, palmitoylation, and by transport signals present in the cytoplasmic tail of the MPRs. Export from the trans-Golgi is mediated by binding to the clathrin-adapter AP-1. GGA1, 2 and 3 bind the acidic cluster-dileucine in the cytoplasmic tail and mediate binding of the receptors to AP1, which then nucleates clathrin-coated vesicles (Puertollano et al. 2001; Zhu et al. 2001). After reaching endosomes, a phenylalanine–tryptophan motif ensures retention in late endosomes. Recycling back to the trans-Golgi network is mediated by a clathrin-independent pathway involving TIP47, Rab9, PACS-1, and AP-1. MPRs can reach the plasma membrane either by recycling from early or late endosomes or by missorting directly from the trans-Golgi. Subsequent internalization at the plasma membrane is accomplished by the YSKV motif of the CI-MPR whereas the CD-MPR has three separate internalization sequences (phenylalaninecontaining sequence, tyrosine-based motif and a dileucine motif). The luminal domain of the CD-MPR binds lysosomal enzymes. It consists of a single a helix near the N-terminus and nine primarily antiparallel b strands that form two b sheets (Olson et al. 2002). When ligand is bound, the binding site encompasses the phosphate group and the terminal three mannose rings of the cargo protein. Two amino acid side chains (Q66 and R111) constitute the binding specificity for mannose 6-phosphate rather than glucose 6phosphate. Additionally, the phosphate group itself is relevant for highaffinity ligand recognition by the receptor. A Mn2+ ion and water establish additional contacts with the phosphate. Structural analysis revealed that the carbohydrate recognition domain lies relatively deep inside the protein. This explains the high binding affinity of the CD-MPR. In comparison to many other lectins, where water fills the ligand-free cleft, the binding pocket of the CD-MPR is occupied by a loop in the absence of ligand. The role of the CD-MPR at the plasma membrane remains mysterious. At neutral pH (pH 7.4) the CD-MPR does not bind ligands efficiently, but it may regulate secretion of mannose 6-phosphate containing ligands into the extracellular milieu. The CI-MPR has a large extracytoplasmic domain composed of 15 repeating units. CI-MPR possesses two distinct carbohydrate recognition sites and a single IGF-II binding site. Structure-based sequence alignments with the carbohydrate recognition domain of the CD-MPR and mutagenesis experiments predict an arginine residue and several other amino acids in two repeating units to be important for ligand recognition. The two CRDs of the CI-MPR are structurally and mechanistically similar to each other, but there are also differences. First, the amino-terminal binding site shows efficient binding at higher pH and second, it is able to bind to larger M6P-OGlcNAc

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phosphodiesters, M6P-OCH3 phosphodiester and mannose 6-sulfate with only slightly lower affinities than M6P (Distler et al. 1991). CI-MPR is posttranslationally modified with N-glycans in the extracytoplasmic domain, although they are not required for ligand-binding. However, the formation of disulfide bonds is crucial for proper folding of the receptor. Furthermore, the receptor is palmitoylated but neither the attachment site nor the function

Figure 2. Localization and function of luminal lectins. Secretory and membrane proteins enter the ER co-translationally through the Sec61 translocation pore and get N-glycosylated by the oligosaccharyltransferase complex (OST). Upon trimming by ER glucosidase I and II, monoglucosylated proteins are bound by CNX and CRT which increases the efficiency of protein folding (2). After release from CNX/CRT by ER glucosidase II (3), glycoproteins can follow three different fates depending on their conformation. Nearly native glycoproteins will be re-glucosylated by UGT1 and re-enter the CNX/CRT cycle (4). Terminally misfolded glycoproteins will be subjected to extensive Man trimming by ER mannosidase I and re-translocated into the cytosol for proteasomal degradation (6). The EDEMs may act as mannosidases or as lectin chaperones. Native glycoproteins will be captured and packaged into COPII vesicles for ER exit. VIPL might bind native glycoproteins released from CNX/CRT (7) and pass the proteins on to ERGIC-53 (8). ERGIC-53 is a cargo receptor for a subset of glycoproteins and facilitates ER to ERGIC transport (9). In the ERGIC, cargo is released upon a drop in pH and perhaps calcium. The ERGIC-53-related lectin VIP36 functions at the Golgi interface and is proposed to capture proteins which have escaped the ER quality control machinery and recycle them back to the ER (10) or cis-Golgi (11). For sorting to lysosomes, GlcNAc 1-phosphate is transferred to high-mannose glycans of lysosomal enzymes in the cis-Golgi (12), uncovered in the trans-Golgi (13). Man6 phosphorylated proteins are captured in the TGN by MPRs and transported to endosomes (14).

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is known. CI-MPR can also endocytose various ligands including Man6Pmodified lysosomal enzymes from the plasma membrane.

Conclusions and perspectives Animal cells control the folding, degradation, sorting and transport of newly synthesized glycoprotein by expressing several luminal lectins which differ in their intracellular localization and function as illustrated in Fig. 2. Based on the characterization of the carbohydrate-specificities of the different lectins, the A branch of the core-glycan emerges as important determinant for the fate of a glycoprotein. While mono-glucosylation of the A branch retains a glycoprotein in the CNX/CRT cycle, extensive trimming of a1,2-linked Man residues in the A branch seems to mark terminally misfolded glycoproteins for ERAD. Moreover, the de-glucosylated A branch is the preferential binding substrate of VIPL, which might protect native glycoproteins from extensive Man trimming in the ER and degradation. The increasing number of CRD crystal structures allows us to understand the molecular determinants of carbohydrate-specificity. A case in point is the change of VIP36’s sugarspecificity based on a rational amino acid substitution in its CRD (Kamiya et al. 2008). Apart from binding to N-glycans of substrate glycoproteins, at least some of the lectins seem to recognize also protein determinants. While the notion of such a bi-partite interaction is still controversial for CNX and CRT, ERGIC-53 definitively recognizes a combined oligosaccharide/peptide structure (Appenzeller-Herzog et al. 2005). Future work is likely to identify the contribution of protein determinants to substrate binding and the binding specificity of the other lectins. Further unresolved questions concern the functions of the less-characterized lectins such as the EDEMs, VIP36, VIPL or ERGL. Are the EDEMs solely lectins or do the proteins possess mannosidase activity? Do the three EDEMs fulfill redundant functions? What are the cargo proteins of VIP36, VIPL and ERGL? Do the lectins relay substrate proteins as proposed for VIPL passing cargo on to ERGIC-53?

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Supplementary references Protein folding and quality control in the ER Denzel A, Molinari M, Trigueros C, Martin JE, Velmurgan S, Brown S, Stamp G, Owen MJ (2002) Early postnatal death and motor disorders in mice congenitally deficient in calnexin expression. Mol Cell Biol 22: 7398–7404 Ellgaard L, Riek R, Herrmann T, Guntert P, Braun D, Helenius A, Wuthrich K (2001) NMR structure of the calreticulin P-domain. Proc Natl Acad Sci USA 98: 3133–3138 Hammond C, Braakman I, Helenius A (1994) Role of N-linked oligosaccharide recognition, glucose trimming, and calnexin in glycoprotein folding and quality control. Proc Natl Acad Sci USA 91: 913–917 Jackson MR, Cohen-Doyle MF, Peterson PA, Williams DB (1994) Regulation of MHC class I transport by the molecular chaperone, calnexin (p88, IP90). Science 263: 384–387 Michalak M, Lynch J, Groenendyk J, Guo L, Robert Parker JM, Opas M (2002) Calreticulin in cardiac development and pathology. Biochim Biophys Acta 1600: 32–37

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ER-associated protein degradation Buschhorn BA, Kostova Z, Medicherla B, Wolf DH (2004) A genome-wide screen identifies Yos9p as essential for ER-associated degradation of glycoproteins. FEBS Lett 577: 422–426 Eriksson KK, Vago R, Calanca V, Galli C, Paganetti P, Molinari M (2004) EDEM contributes to maintenance of protein folding efficiency and secretory capacity. J Biol Chem 279: 44600–44605 Hirao K, Natsuka Y, Tamura T, Wada I, Morito D, Natsuka S, Romero P, Sleno B, Tremblay LO, Herscovics A, Nagata K, Hosokawa N (2006) EDEM3, a soluble EDEM homolog, enhances glycoprotein endoplasmic reticulum-associated degradation and mannose trimming. J Biol Chem 281: 9650–9658 Hosokawa N, Tremblay LO, You Z, Herscovics A, Wada I, Nagata K (2003) Enhancement of endoplasmic reticulum (ER) degradation of misfolded Null Hong Kong alpha1antitrypsin by human ER mannosidase I. J Biol Chem 278: 26287–26294 Hosokawa N, Wada I, Hasegawa K, Yorihuzi T, Tremblay LO, Herscovics A, Nagata K (2001) A novel ER alpha-mannosidase-like protein accelerates ER-associated degradation. EMBO Rep 2: 415–422 Mast SW, Diekman K, Karaveg K, Davis A, Sifers RN, Moremen KW (2005) Human EDEM2, a novel homolog of family 47 glycosidases, is involved in ER-associated degradation of glycoproteins. Glycobiology 15: 421–436 Molinari M, Calanca V, Galli C, Lucca P, Paganetti P (2003) Role of EDEM in the release of misfolded glycoproteins from the calnexin cycle. Science 299: 1397–1400 Oda Y, Hosokawa N, Wada I, Nagata K (2003) EDEM as an acceptor of terminally misfolded glycoproteins released from calnexin. Science 299: 1394–1397 Olivari S, Galli C, Alanen H, Ruddock L, Molinari M (2005) A novel stress-induced EDEM variant regulating endoplasmic reticulum-associated glycoprotein degradation. J Biol Chem 280: 2424–2428 Szathmary R, Bielmann R, Nita-Lazar M, Burda P, Jakob CA (2005) Yos9 protein is essential for degradation of misfolded glycoproteins and may function as lectin in ERAD. Mol Cell 19: 765–775

Protein transport Anelli T, Ceppi S, Bergamelli L, Cortini M, Masciarelli S, Valetti C, Sitia R (2007) Sequential steps and checkpoints in the early exocytic compartment during secretory IgM biogenesis. EMBO J 26: 4177–4188 Appenzeller-Herzog C, Roche AC, Nufer O, Hauri HP (2004) pH-induced conversion of the transport lectin ERGIC-53 triggers glycoprotein release. J Biol Chem 279: 12943–12950

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Dahm T, White J, Grill S, Fullekrug J, Stelzer EH (2001) Quantitative ER-Golgi transport kinetics and protein separation upon Golgi exit revealed by vesicular integral membrane protein 36 dynamics in live cells. Mol Biol Cell 12: 1481–1498 Hara-Kuge S, Ohkura T, Ideo H, Shimada O, Atsumi S, Yamashita K (2002) Involvement of VIP36 in intracellular transport and secretion of glycoproteins in polarized MadinDarby canine kidney (MDCK) cells. J Biol Chem 277: 16332–16339 Itin C, Roche AC, Monsigny M, Hauri HP (1996) ERGIC-53 is a functional mannoseselective and calcium-dependent human homologue of leguminous lectins. Mol Biol Cell 7: 483–493 Kappeler F, Klopfenstein DR, Foguet M, Paccaud JP, Hauri HP (1997) The recycling of ERGIC-53 in the early secretory pathway. ERGIC-53 carries a cytosolic endoplasmic reticulum-exit determinant interacting with COPII. J Biol Chem 272: 31801–31808 Klumperman J, Schweizer A, Clausen H, Tang BL, Hong W, Oorschot V, Hauri HP (1998) The recycling pathway of protein ERGIC-53 and dynamics of the ER-Golgi intermediate compartment. J Cell Sci 111(Pt 22): 3411–3425 Nichols WC, Seligsohn U, Zivelin A, Terry VH, Hertel CE, Wheatley MA, Moussalli MJ, Hauri HP, Ciavarella N, Kaufman RJ, Ginsburg D (1998) Mutations in the ER-Golgi intermediate compartment protein ERGIC-53 cause combined deficiency of coagulation factors V and VIII. Cell 93: 61–70 Nufer O, Kappeler F, Guldbrandsen S, Hauri HP (2003) ER export of ERGIC-53 is controlled by cooperation of targeting determinants in all three of its domains. J Cell Sci 116: 4429–4440 Nyfeler B, Michnick SW, Hauri HP (2005) Capturing protein interactions in the secretory pathway of living cells. Proc Natl Acad Sci USA 102: 6350–6355 Nyfeler B, Zhang B, Ginsburg D, Kaufman RJ, Hauri HP (2006) Cargo selectivity of the ERGIC-53/MCFD2 transport receptor complex. Traffic 7: 1473–1481 Teasdale RD, Jackson MR (1996) Signal-mediated sorting of membrane proteins between the endoplasmic reticulum and the Golgi apparatus. Annu Rev Cell Dev Biol 12: 27–54 Velloso LM, Svensson K, Pettersson RF, Lindqvist Y (2003) The crystal structure of the carbohydrate-recognition domain of the glycoprotein sorting receptor p58/ERGIC53 reveals an unpredicted metal-binding site and conformational changes associated with calcium ion binding. J Mol Biol 334: 845–851 Vollenweider F, Kappeler F, Itin C, Hauri HP (1998) Mistargeting of the lectin ERGIC-53 to the endoplasmic reticulum of HeLa cells impairs the secretion of a lysosomal enzyme. J Cell Biol 142: 377–389 Wendeler MW, Paccaud JP, Hauri HP (2007) Role of Sec24 isoforms in selective export of membrane proteins from the endoplasmic reticulum. EMBO Rep 8: 258–264 Yamaguchi D, Kawasaki N, Matsuo I, Totani K, Tozawa H, Matsumoto N, Ito Y, Yamamoto K (2007) VIPL has sugar-binding activity specific for high-mannose-type N-glycans, and glucosylation of the {alpha}1,2 mannotriosyl branch blocks its binding. Glycobiology 17: 1061–1069

Lysosomal protein sorting in the Golgi Breuer P, Korner C, Boker C, Herzog A, Pohlmann R, Braulke T (1997) Serine phosphorylation site of the 46-kDa mannose 6-phosphate receptor is required for transport to the plasma membrane in Madin-Darby canine kidney and mouse fibroblast cells. Mol Biol Cell 8: 567–576 Chao HH, Waheed A, Pohlmann R, Hille A, Von Figura K (1990) Mannose 6-phosphate receptor dependent secretion of lysosomal enzymes. EMBO J 9: 3507–3513 Dahms NM, Hancock MK (2002) P-type lectins. Biochim Biophys Acta 1572: 317–340

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Dahms NM, Rose PA, Molkentin JD, Zhang Y, Brzycki MA (1993) The bovine mannose 6phosphate/insulin-like growth factor II receptor. The role of arginine residues in mannose 6-phosphate binding. J Biol Chem 268: 5457–5463 Diaz E, Pfeffer SR (1998) TIP47: a cargo selection device for mannose 6-phosphate receptor trafficking. Cell 93: 433–443 Duncan JR, Kornfeld S (1988) Intracellular movement of two mannose 6-phosphate receptors: return to the Golgi apparatus. J Cell Biol 106: 617–628 Koster A, Von Figura K, Pohlmann R (1994) Mistargeting of lysosomal enzymes in M(r) 46,000 mannose 6-phosphate receptor-deficient mice is compensated by carbohydrate-specific endocytotic receptors. Eur J Biochem 224: 685–689 Meresse S, Hoflack B (1993) Phosphorylation of the cation-independent mannose 6phosphate receptor is closely associated with its exit from the trans-Golgi network. J Cell Biol 120: 67–75 Morgan DO, Edman JC, Standring DN, Fried VA, Smith MC, Roth RA, Rutter WJ (1987) Insulin-like growth factor II receptor as a multifunctional binding protein. Nature 329: 301–307 Olson LJ, Zhang J, Lee YC, Dahms NM, Kim JJ (1999) Structural basis for recognition of phosphorylated high mannose oligosaccharides by the cation-dependent mannose 6-phosphate receptor. J Biol Chem 274: 29889–29896 Reczek D, Schwake M, Schroder J, Hughes H, Blanz J, Jin X, Brondyk W, Van Patten S, Edmunds T, Saftig P (2007) LIMP-2 is a receptor for lysosomal mannose-6-phosphate-independent targeting of beta-glucocerebrosidase. Cell 131: 770–783 Riederer MA, Soldati T, Shapiro AD, Lin J, Pfeffer SR (1994) Lysosome biogenesis requires Rab9 function and receptor recycling from endosomes to the trans-Golgi network. J Cell Biol 125: 573–582 Roberts DL, Weix DJ, Dahms NM, Kim JJ (1998) Molecular basis of lysosomal enzyme recognition: three-dimensional structure of the cation-dependent mannose 6phosphate receptor. Cell 93: 639–648 Rohrer J, Kornfeld R (2001) Lysosomal hydrolase mannose 6-phosphate uncovering enzyme resides in the trans-Golgi network. Mol Biol Cell 12: 1623–1631 Sahagian GG, Distler J, Jourdian GW (1981) Characterization of a membrane-associated receptor from bovine liver that binds phosphomannosyl residues of bovine testicular beta-galactosidase. Proc Natl Acad Sci USA 78: 4289–4293 Schweizer A, Kornfeld S, Rohrer J (1996) Cysteine34 of the cytoplasmic tail of the cationdependent mannose 6-phosphate receptor is reversibly palmitoylated and required for normal trafficking and lysosomal enzyme sorting. J Cell Biol 132: 577–584 Schweizer A, Kornfeld S, Rohrer J (1997) Proper sorting of the cation-dependent mannose 6-phosphate receptor in endosomes depends on a pair of aromatic amino acids in its cytoplasmic tail. Proc Natl Acad Sci USA 94: 14471–14476 Steet R, Lee WS, Kornfeld S (2005) Identification of the minimal lysosomal enzyme recognition domain in cathepsin D. J Biol Chem 280: 33318–33323 Tong PY, Tollefsen SE, Kornfeld S (1988) The cation-independent mannose 6-phosphate receptor binds insulin-like growth factor II. J Biol Chem 263: 2585–2588 Wan L, Molloy SS, Thomas L, Liu G, Xiang Y, Rybak SL, Thomas G (1998) PACS-1 defines a novel gene family of cytosolic sorting proteins required for trans-Golgi network localization. Cell 94: 205–216

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The Golgi ribbon and the function of the golgins Maria A. De Matteis, Alexander A. Mironov and Galina V. Beznoussenko

Introduction The Golgi apparatus (GA) is present in different organisms in very different forms. When visualized by immunofluorescence in most mammalian cells, the Golgi ribbon appears as a lacy structure that occupies a volume of 5–7 mm in length, 1–2 mm in width, and 3–5 mm in depth (Storrie and Kreis 1996), and that surrounds the centrosome (or microtubule-organizing centre; MTOC) (see Chapter 2.14). The positions of the MTOC and the GA depend on cell polarity. In many polarized epithelial cells, the centrosome is positioned near the apical portion of the cell surface (Ojakian et al. 1997), where the GA also resides. From dozens to hundreds of Golgi stacks that act as a single organelle are linked together to form an interconnected, ribbon-like structure in the perinuclear area (Hidalgo Carcedo et al. 2004; Polishchuk and Mironov 2004; Mogelsvang et al. 2004). Completely isolated stacks are rare (Cole et al. 1996a,b). The connectivity between individual stacks is appreciable in living cells, where photobleaching of a fraction of the GA containing chimeras between resident Golgi proteins (i.e. enzymes) and green fluorescent protein (GFP) quickly induces a disappearance/decrease of fluorescence from other parts of the GA, suggesting rapid diffusional exchange of the Golgi enzymes (such as ManII and GalT, but not ManI, Marra et al. 2001) between the Golgi stacks (Cole et al. 1996a,b). Continuity of the stacks within the GA has been confirmed using scanning electron microscopy (SEM). The SEM complementary observation of the fractured samples of the GA has demonstrated that while almost all of the adjacent stacks appear to be separated from one another, they are not actually separated, but remain continuous with each other (Inoue 1992). The Golgi ribbon is, however, just one of the possible shapes of this organelle, as it is found in different organisms in very different forms: from scattered tubular networks (Saccharomyces cerevisiae) and isolated multiple stacks of cisternae (Pichia pastoris, Drosophila, plants), to the above-described continuous pericentrosomal ribbon of many mammalian cells. The overall structural organization of the GA can vary also within the same species between different cell types: in mammals, for instance, it is present as isolated stacks in oocytes and skeletal muscle cells, while it forms a continuous ribbon in the majority of other cell types. Finally, the Golgi ribbon can change its organization in the very same cell through the cell cycle (since it undergoes

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cycles of fragmentation and reformation during the mitotic process) and according to the state of activity of an organelle. Despite its very “conserved” functions, the reasons for such a variegated architecture of the GA in different organisms, or even in different cells of the same organism, are not yet really understood. A number of possibilities have been put forward, with the one that is more frequently considered being related to the GA and cell “motility”: directing the movement of a continuous organelle (such as the Golgi ribbon) within the cell would be easier and more effective than coordinately moving multiple unlinked organelles (isolated Golgi stacks). This appears to be required during cell migration, when the GA is reoriented towards the leading edge of the cell, to provide membranes for the advancing cell front. However, two levels of consideration challenge this assumption: the isolated Golgi stacks in plants are highly motile along actin filaments (see Chapter 4.1), and cells deprived of a central GA migrate apparently normally (Kondylis and Rabouille 2003). Perhaps a more likely explanation is that the integrity of the Golgi ribbon may represent a sort of “signal”, e.g. for a Golgibased mitotic checkpoint (Colanzi et al. 2003; Chapter 3.15). Finally, one should consider the “easiest” explanation, that the ribbon represents just one of the possible configurations of the organelle and it is linked to its functional state (i.e. transport activity, see below). In spite of our uncertainties regarding the reasoning behind a continuous organelle instead of isolated stacks or cisternae, we know a lot about the molecular mechanisms that are responsible for the building up and maintenance of the Golgi ribbon: these are the focus of this Chapter.

The role of microtubules A common feature of the cells that do not have a continuous Golgi ribbon is the absence of the radial organization of MTs around the MTOC. This feature is seen for mammalian oocytes and myotubes, which do not have a Golgi ribbon (Polishchuk et al. 1999; Trucco et al. 2004), in insect cells (Kondylis and Rabouille 2003), and in plants, yeast, and some protists. Protist cells that have a MTOC usually also have their GA organized into a ribbon, reinforcing the correlation that exists between the existence of the MTOC and the presence of a Golgi ribbon. Thus, a first prerequisite for the formation of the Golgi ribbon is the presence of a radial MT array centred on the MTOC. The second requirement for the formation of the Golgi ribbon is the presence of MTs per se. Depolymerization of MTs with specific agents (e.g. nocodazole) (Cole et al. 1996a,b; Thyberg and Moskalewski 1999; Polishchuk et al. 1999; Trucco et al. 2004) induces the fragmentation of the Golgi ribbon into many peripheral pieces. These Golgi fragments have a typical stacked organization (Polishchuk et al. 1999). In favourable EM sections, the Golgi ministacks formed after depletion of cells from MTs can be seen in close association with endoplasmic reticulum (ER) exit sites (Polishchuk et al. 1999; Storrie et al. 1998).

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Although the mechanisms underlying this Golgi fragmentation are beyond the scope of this Chapter (see some details in Chapter 2.14), mention should however be made that Golgi fragmentation occurs with the participation of the diffusion of Golgi enzymes through the ER (Cole et al. 1996a,b; Storrie et al. 1998), with this mechanism having been questioned more recently (Pecot and Malhotra 2006). Here it is important to note that these Golgi ministacks can reassemble into a continuous ribbon upon removal of the MT-disrupting drug and restoration of the MTOC and MT organization (Ho et al. 1989). The third requirement is that MTs maintain their dynamic instability. Indeed, taxol, which stabilizes MTs and induces the polymerization of MTs not connected with the MTOC and of MT bundles localized in the cell periphery (Wehland et al. 1983), also induces the fragmentation of the Golgi ribbon. Even when MT dynamics are lowered by addition of very low concentrations of nocodazole or of taxol, the Golgi ribbon undergoes fragmentation (Minin 1997). Finally, connection between the Golgi ribbon and MT organization emerges also from the observation that the Golgi ribbon is severed into isolated stacks during the cell cycle, in G2, when reorganization of the MT star could contribute to this peripheral Golgi fragmentation (Thyberg and Moskalewski 1999). G2-blocked cells do not show major differences either in the number of cisternae that comprise a single stack, or in the average diameter of the stacks; however, when compared with control cells, they show stacks that are isolated (i.e. not interconnected by membrane tubules) in most cases and are not longitudinally aligned. The stacks in G2 cells are either isolated or connected in small groups of two to four stacks; this Golgi fragmentation in G2 is mirrored by the much lower recovery rate of GalT–GFP in G2 cells compared to interphase cells in FRAP experiments (Hidaldo Carcedo et al. 2004). The requirement for MTs and the MTOC underlies the involvement of MT-based motors, and in particular of dynein, in building up and maintaining the Golgi ribbon. Dynein is a multisubunit minus-end-directed microtubule motor that is known to transport a variety of cargoes in animal cells (Vallee et al. 2004; Vale 2003). Cytoplasmic dynein cycles constitutively between the ER and the GA. It co-localizes partially with the intermediate compartment (Roghi and Allan 1999). Dynein function requires a series of “accessory proteins”, those that activate dynein, such as dynactin, and those involved in the recruitment of dynein–dynactin to membranes, such as Bicaudal-D (Matanis et al. 2002), Rab6 (Short et al. 2002), beta III spectrin (Holleran et al. 2001), and CLIPR-59 (Perez et al. 2002). Dynactin is concentrated at MT plus ends and can transiently capture Golgi membranes that are then transported towards the minus ends by dynein (Vaughan et al. 2002). These mechanisms are discussed in more detail in Chapter 2.14. Several studies have suggested that dynein can associate with the mammalian GA through Golgi-associated spectrin and dynactin, a protein complex

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that is required for the movement of dynein cargo (Kumar et al. 2004; Schroer sy-Theulaz et al. 1992; Harada et al. 1998). 2004; Burkhardt et al. 1997; Corthe Cells lacking dynein cannot concentrate organelles in the perinuclear region, such as the GA and lysosomes (Narada et al. 1998), while the over-expression of p50/dynamitin results in the dissociation of dynein heavy chain from the membrane of peripheral Golgi elements (Roghi and Allan 1999) and induces the dispersal of the GA from the perinuclear region (Burkhardt et al. 1997). The situation is different in polarized epithelial cells, where MTs are primarily oriented with their plus ends basally, near the GA, and their minus ends in the apical cytoplasm. Here, the distribution of MT motors is also different and a selected kinesin isoform (KIFC3) has been shown to have roles in MT-dependent centralization and positioning of the GA in some polarized epithelial cells (Xu et al. 2002). The balance between the activity of the minus-end motor dynein, and the plus-end motor kinesin will determine the final positioning of the GA. However, the directionality of membrane trafficking and cycling between the ER and the GA is unlikely to be achieved through the control of motor–membrane interactions; rather, the motors, and especially dynein, probably remain bound throughout the whole anterograde–retrograde cycle, with their activity (and not their membrane association) being modulated accordingly (Fritzler et al. 1993). This is suggested also by the finding that dynein heavy chain is found on brefeldin-A-induced tubules that are moving retrogradely towards microtubule plus ends (Roghi and Allan 1999). Although the Golgi ribbon is centred on the MTOC, it does not contact it, with most of the Golgi elements situated at a distance of 1–3 mm from the centrosome. The reason for this is not clear, although it is likely that the MTs growing from the centrosome at a high density “push” the Golgi membranes out, forcing them to stay at some distance from the cell centre (Polishchuk and Mironov 2004). Kinesin, the plus-end MT-based motor that serves for centrifugal movement of post-Golgi carriers and the ER might have a role in maintaining this MTOC–Golgi distance, as its inactivation results in the collapse of the Golgi around the centrosome. In cells without kinesin, the ER is also located near to the centrosome (Feiguin et al. 1994). Another mechanism preventing overlap of the GA with the MTOC might involve GMAP-210, a protein involved in the interaction of Golgi membranes with MTs (Chabin-Brion et al. 2001). GMAP-210 captures short MT seeds that are formed at the centrosome by their minus ends, and together with the Golgilocalized pool of the CLASPs that attach to and stabilize the MT plus ends, GMAP-210 generates a meshwork of short MTs that are associated with adjacent Golgi stacks and links them up to form a ribbon (Rios et al. 2004). Another possible mechanism might involve the actin/myosin machinery: inducing actin depolymerization induces a “collapse” of the GA on the MTOC (Valderrama et al. 1998). The reason for this is also not completely clear (see also Chapter 2.14). Isolated Golgi membranes from intestinal epithelial cells are enriched in myosin-I, dynein, and its in vitro motility activator dynactin

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(p150/Glued). Myosin-I is present on all membranes in the Golgi fraction; dynein is present only on a small membrane fraction (Fath et al. 1994).

The role of membrane input from the endoplasmic reticulum The presence of an intact MT system is not sufficient, per se, to guarantee the formation of the Golgi ribbon: the continuous input of ER-derived membranes moving centripetally along MTs towards the Golgi area is also a primary requirement, as it represents the source of the membranes that promote the formation of connections between stacks. This absolute requirement for membrane input is testified by the observation that the Golgi ribbon disconnects into isolated stacks as soon as the arrival of membranes from the ER is slowed down or interrupted (Marra et al. 2007). Indeed, Rambourg et al. (1993) also showed that the Golgi ribbon of prolactin cells is fragmented when the function of the GA is blocked (compare figs. 1 and 7 in Rambourg et al. 1993). The ribbon configuration of the GA in mammalian cells thus reflects an “active” state of the organelle, as it receives and absorbs membranous carriers from the ER (the ER-to-Golgi carriers; EGCs) (Marra et al. 2007). For ER-derived membranes to sustain the formation of the Golgi ribbon, they need to be completely integrated into the cisterna stacks; this implies that the highly pleiomorphic membranes coming out from the ER (i.e. the EGCs) need to be re-shaped, and transformed into flat discs (i.e. the Golgi cisternae), and integrated into the Golgi ribbon. Indeed, following their generation at the ER, EGCs undergo a series of modification steps with regard to their molecular composition, ultrastructure and dynamics (Marra et al. 2001), which prime them for entering the GA. Among these, a key step is the acquisition of a class of Golgi-associated proteins that are collectively known as the Golgi tethering factors, the golgins, due to their property of joining membranes together.

The golgins The golgins were originally identified as a family of Golgi-localized autoantigens using antibodies derived from the sera of patients with a variety of autoimmune disorders (Chan and Fritzler 1998). Most of these golgins were then cloned by screening of expression libraries with these autoantibodies, such as Golgin-160 (Fritzler et al. 1993), giantin (Seelig et al. 1994), GMAP-210 (Rios et al. 1994), Golgin-245/p230 (Fritzler et al. 1995), Golgin-97 (Griffith et al. 1997) and Golgin-67 (Eystathioy et al. 2000). Among the different golgins, GM130, giantin, and p115 have been the most studied to date. Most of the golgins have extensive coiled-coil regions throughout their entire polypeptide chain, a common protein motif that is known to form an extended rod-like structure (Kjer-Nielsen et al. 1999; Burkhard et al. 2001). Coiled-coil proteins have two a-helices that wrap around each other with a slight left-handed superhelical twist, forming the rod-like structure

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(Gillingham et al. 2003). These proteins have elongated shapes that can reach several dozens of nm in length. Most of the golgins are peripheral membrane proteins and they often have short non-coiled-coil regions at either end that mediate their targeting and other interactions. Thus, the C-terminus of GM130 binds to GRASP65, a lipid-anchored protein on the cis-Golgi network (Barr et al. 1998). Similarly, four of the golgins share a C-terminal GRIP domain that is sufficient to target them to the trans-side of the GA (see Chapter 3.3). In contrast, other golgins, such as giantin, Golgin-84 and CASP, are anchored in the bilayer via a C-terminal transmembrane domain (Fridmann-Sirkis et al. 2004; see below). Although the golgins from different species share some structural features, their sequences are not well conserved (Renna et al. 2005). In addition, the golgins are not uniformly distributed among all species. Some organisms and cells contain significant levels of the golgins, while others lack most of them. Indeed, only a few of the golgins are present in yeast: Coy1p (similar to CASP), and Grp1p (similar to Golgin-160/GCP170) (Barr and Short 2003; Gillingham and Munro 2003), and the single GRIP protein, Imh1p (Siniossoglou et al. 2000; Tsukada et al. 1999; Short et al. 2005; Luke et al. 2003). Apart from Golgi localization and the presence of coiled-coil motifs, the only other common feature so far identified for the golgin family is that many members interact with small GTPases. Depending on their localization (Table 1), the golgins can be divided into two subfamilies: the cis- and trans-golgins. The cis-golgins include p115, GM130/Golgin-95, GRASP65, GRASP55, CASP, Coy1p (similar to CASP), Golgin-45, Golgin-67 and Golgin-84. Some of these were previously considered to be tethering factors. The trans-golgins include Golgin-97, GCC88, Golgin-160/ MEA-2/GCP170, Golgin-245/p230/tGolgin-1, GCC185, GMAP-210, and Grp1p (similar to Golgin-160/GCP170) in yeast (Barr and Short 2003; Gillingham and Munro 2003), and the related group of proteins – possibly splice variants – of GCP372 and GCP364 (Barr 1999). Giantin/macrogolgin is not restricted to either the cis or the trans pole of the GA. The golgins are recruited to Golgi membranes in several ways. Some of them are targeted by the Rabs (p115 by Rab1; Bicaudal-D1 and Bicaudal-D2 by Rab6) in a nucleotide-dependent manner. The GRASP proteins themselves are targeted via an N-terminal myristoyl group. GM130 associates with the membrane through an interaction with the GRASPs. Golgin-245, Golgin97, GCC88, and GCC185 are targeted to the trans-Golgi network (TGN) membranes by their C-terminal GRIP domain, in a G-protein-dependent process involving Arl1. In turn, the Rabs are targeted to membranes via Cterminal geranylgeranyl modifications, whereas most of the Arls are targeted to membranes by N-terminal myristoylation (Short et al. 2002; Matanis et al. 2002). Giantin, Golgin-84, and CASP contain long N-terminal coiled-coil regions that protrude into the cytosol and C-terminal transmembrane domains with aminoacid residues that are highly conserved across species. Their transmembrane domains also show a high degree of sequence similarity.

Cytosolic; rod like; 54 nm in length; coiled-coil; homodimer GM130; giantin, syn5, sly1, membrin, Ykt6, Rab1 (GTP), GOS28, ARFGEF; IRAP; No binding to MTs VTC, late IC; CGN, ciscisterna

Structure

Unknown To the IC, then ER

To the IC (?)

Role in mitotic Golgi fragmentation Redistribution under the action of BFA

Attachment of the CGN to the medial Golgi Through phosphorylation

Role in stacking

Small

IC-Golgi; intra-Golgi in vitro; ManI (but not ManII) ! IC at 15 C in Lec1

Role in transport

By cytoplasmic domain of about 100 residues adjacent to the TMD Small

By Rab1 and other binding partners

Non-compact zones

C-terminus of p115 by N-terminus of giantin

Type II protein; rod like; coiled-coil; 400 kDa

Giantin

Targeting

Localization

Main binding partners

p115

Characteristics

Table 1. Characteristics of golgins

Minimal; W/o GM130 – BFA resistance; in Drosophila w/o GRASP65 no inhibition Attachment of the CGN to the medial GA Through phosphorylation To the IC

CGN and cisternal rims of the 2nd and the 3rd cisternae; co-localization with cargo By GRASP and other binding partners

C-terminus of p115 by N-terminus of GM130; Rab1 (GTP); GRASP65

Cytosolic; rod like; coiled-coil

GM130

To the IC

(Continued)

Small; deletion of GM130 does not induce transport inhibition in Drosophila cell. Attachment of the CGN to the medial GA Significant

By myristoylation

IC/CGN

GM130. p24

Cytosolic with myristoylation

GRASP65

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Role in mitosis Redistribution under the action of BFA Result of protein depletion or inhibition Result of protein over expression

Role in transport Role in stacking

Localization Targeting

IC/CGN By TMD

Unknown

Type II protein; rod like; coiled-coil

Unknown

Unknown

Cytosolic with myristoylation GM130; p24; Golgin-45; TGF a Cis; VTS; rims; tubules By myristoylation

Structure

CASP

Unknown

Central Golgi fragmentation Unknown

GRASP55

Characteristics

Unknown

Central Golgi fragmentation

Unknown Regulation of formation of the attached CGN Unknown To the ER

Peripheral Golgi fragmentation

Result of protein over expression

Central Golgi fragmentation

GM130

Unknown Attachment of the CGN to the medial GA Significant Unknown

Peripheral Golgi fragmentation

Result of protein depletion or inhibition

Giantin

Peripheral Golgi fragmentation

Unknown IC

CGN Cytoplasmic domain adjacent to the TMD Minimal Attachment of the CGN to medial Golgi

Rod; coiled-coil; type II protein (N-cytosolic) Rab1 (GTP), not matrix proteins

Golgin-84

Central Golgi fragmentation with augmentation of the cisterna length and decrease of number of cisternae Unknown

GRASP65

*

Main binding partners

p115

Characteristics

Table 1. (Continued) 230 M. A. De Matteis et al.

Rab2. GRASP55 CGN Unknown

ERES Attachment of the CGN to medial Golgi Unknown Unknown Redistribution to the ER (?)

TGN (?) Unknown

Unknown Unknown

Unknown Unknown

Unknown

Main binding partners Localization Targeting

Role in transport Role in stacking

Role in mitosis Redistribution by BFA Results of inhibition of function

Rod; coiled-coil; cytosolic

Rod; Coiled-coil; cytosolic; similar to GM130; TMD unclear Unknown

Structure

Golgin-45

Golgin-67

Characteristics

ES-TGN Attachment of the TGN to the medial Golgi Unknown Cytosolic and endosomal Central fragmentation; increased cisternal length

Trans/TGN Homodimers bind ARL1

ARL1

Rod; coiled-coil; cytosolic

Golgin-97

Peripheral Golgi fragmentation

Unknown Cytosolic

Trans/TGN Homodimers bind ARL1; MACF-1; no binding to dynactin/ dynein ES-TGN Unknown

ARL1

Rod; coiled-coil; cytosolic

Golgin-245

Unknown

Unknown Cytosolic (?)

Unknown Unknown

TGN Rab6 (?)

Rab6. Dynactin

Rod; coiled-coil; cytosolic

Bicaudal-D1/D2

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In particular, key tyrosine and histidine residues are completely conserved. When such a conserved tyrosine residue is mutated to leucine in CASP, this protein no longer localizes to the GA, and accumulates in the ER instead (Gillingham et al. 2002). Both the cis- and trans-golgins are required to maintain the Golgi ribbon. Indeed, inhibition of the function of p115 (by different means) induces Golgi fragmentation into peripheral stacked cisternae and vesicle-like structures (Schroer et al. 2004). Both the over-expression and depletion of Golgin-84 results in the peripheral fragmentation of the Golgi ribbon (Diao et al. 2003), while inhibition of GM130/GRASP65 function in any way induces the fragmentation of the central GA (Puthenveedu et al. 2006; Marra et al. 2007). HeLa cells depleted of GRASP55 show a fragmented GA (Feinstein and Linstedt 2008). Similarly, microinjection of anti-giantin antibodies induces the fragmentation of the central Golgi ribbon (Beznoussenko et al. submitted), and depletion of TMF/ARA160 by RNA interference (RNAi) in NRK cells results in a modest dispersal of the Golgi membranes (Fridmann-Sirkis et al. 2004). RNAi of Golgin-97 or microinjection of anti-Golgin-97 antibodies also induce the fragmentation of the Golgi ribbon, although the Golgi fragments are larger than those induced by the block of GM130, GRASP65, and giantin (Beznoussenko et al. submitted). Golgin-245 depletion leads to replacement of a centralized, ribbon-like pattern with a tiny spotty pattern throughout the cell body (Yoshino et al. 2005). The peripheral fragmentation of the GA induced by deletion of Golgin-245 results in the dispersal of the GA to peripheral ministacks that are well preserved ultrastructurally (Yoshino et al. 2005). Thus, impairment of the function of most of the Golgins can induce either peripheral (p115, Golgin-84, Golgin-245) or central (GM130, giantin, GRASP65, Golgin-97) fragmentation of the Golgi ribbon (Fig. 1).

Other participants in the formation of the Golgi ribbon Other factors that are required for the maintenance of the Golgi ribbon include the different members of the p23-protein family (their over-expression leads to fragmentation of the Golgi ribbon; Rojo et al. 2000) and the retromer components (Seaman 2004). Depletion of the Golgi-associated conserved oligomeric complex also leads to fragmentation of the mammalian GA (Shestakova et al. 2006). Interfering with Golgi-associated enzymes can also induce Golgi fragmentation. This occurs with inhibition of PLA2 activity or expression (R. Polishchuk, personal communication) or upon PKD over-expression (Diaz Anel and Malhotra 2005). PLA2 has been shown to be required to generate and maintain tubular membranes at the GA (Brown et al. 2003), thus including also those that connect neighbouring stacks. For PKD, this is a component of one of the fission machineries that operates at the GA (Diaz Anel and

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Figure 1. GM130 depletion induces fragmentation of the Golgi ribbon. CHO (upper panel) and ldlG cells not expressing GM130 (lower panel) were analyzed by EM tomography. The 3D models of Golgi stacks were superimposed over the corresponding virtual EM images. The 3D models contain cis-most (red) and trans-most (magenta) cisternae. Vesicles (blue, 50–60 nm; white, 80–90 nm) are represented by software-generated spheres, centred on the centre of the vesicles. Models were made by A. A. Mironov, Jr and E. Fontana.

Malhotra 2005). In this respect, the other fissioning factor at the GA, the protein CTBP1/BARS (Weigert et al. 1999; Yang et al. 2005, 2006), would be different since it induces fragmentation of the GA only in the presence of mitotic cytosol (previously depleted of BARS) and not of interphase cytosol (Hidalgo Carcedo et al. 2004). Fragmentation of the GA with the redistribution of various Golgi markers, including Mann II and GM130, has been seen after inhibition of the Golgiresident GPI-anchored protein (GREG) (Xueyi et al. 2007). Expression of 23TMGREG, a fusion protein composed of GREG and the transmembrane domain of p23, also generates a scattered Golgi structure (Xueyi et al. 2007). Similarly, fragmentation of the GA is also observed in PIG-L CHO cells (Abrami et al. 2001) that are deficient in the biosynthesis of the GPI-anchor (Xueyi et al. 2007). In these PIG-L cells labelled with NAGTI-GFP, many circular or ring-like

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structures are seen, which are reminiscent of those induced by expression of GPI-GREG (Xueyi et al. 2007). RNAi of RINT-1, a protein linker between ZW10 and the ER soluble Nethylmaleimide-sensitive factor attachment protein receptor, syntaxin 18, induces Golgi fragmentation (Sun et al. 2007). Depletion of ZW10, a mitotic checkpoint protein implicated in Golgi/ER trafficking/transport, results in a central, disconnected cluster of Golgi elements and inhibition of ERGIC53 and Golgi enzyme recycling to the ER (Sun et al. 2007). Finally, there are several observations in favour of the necessity for the fusion machinery for the transformation of centrally located Golgi stacks into a ribbon-like structure. In cells microinjected with an SNAP mutant, we have seen significant inhibition of the centralization of EGCs (Mironov et al. 2003). However, this microinjection of the SNAP mutant also induces not only central Golgi fragmentation, but also strong vesiculation of Golgi stacks (Kweon et al. 2004). It seems that the over-expression of the SNARE GS15 lacking its transmembrane domain also induces central fragmentation of ManII-positive Golgi structures (i.e. see fig. 8n in Xu et al. 2002), while siRNA-mediated knockdown of GS15 transforms central GalT into a diffuse patterns that is similar to the ER (Xu et al. 2002). An antibody against the R-SNARE Ykt6 (see Chapter 2.1) induces the central fragmentation of the Golgi ribbon within the Golgi area (Zhang and Hong 2001).

The functional role of the Golgi ribbon What is the main purpose for such a complex array of molecular machineries for the building up and maintaining of a ribbon-like GA? Surprisingly, the basic functions of the GA, i.e. membrane trafficking and glycosylation, are not significantly different if they are carried out by a Golgi ribbon or by isolated Golgi stacks. Intra-Golgi transport does not depend on the ribbonlike organization of the GA (Trucco et al. 2004) and its stacked structure. The transport functions of the GA can continue at the same level if the ribbonlike GA is fragmented by depolymerization of MTs (Trucco et al. 2004). Thus, the presence of these “ministacks” in vivo (Trucco et al. 2004; Ward and Brandizzi 2004) and in cell-free assays (Pullikith and Wiedman 2002) allows the GA to perform all of its functions. Indeed, for glycosylation and protein sorting, even a single cisterna is sufficient (Varki 1998). Studies of glycosylation in the GA have shown that sugar nucleotide transporters and many glycosyltransferases are located within a single cisterna (Opat et al. 2001; Young 2004). This suggests that either a stack or even a single cisterna could be considered as the minimal Golgi nano-unit. Indeed, in insects, a double depletion of dGRASP and dGM130 leads to the quantitative conversion of Golgi stacks into clusters of vesicles and tubules, often featuring single cisternae; at the same time, the transport function is not severely impaired (Kondylis et al. 2005).

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What is then the reason for having a Golgi ribbon? In theory, the ribbon could facilitate the even distribution of Golgi enzymes among the stacks, and consequently a more efficient glycosylation of secretory proteins (Puthenveedu et al. 2006). However, when specifically tested on selective secretory proteins, no differences in glycosylation specifically attributable to the presence of a Golgi ribbon can be detected (Marra et al. 2007). As noted above, the presence of a Golgi ribbon might serve more specialized functions, such as the polarized delivery of membranes (that is indeed lost in nocodazole-treated cells). Consistent with this possibility, it has been recently shown that the fragmentation of the Golgi ribbon induced by overexpression of GRASP65 impairs polarized dendrite outgrowth in hippocampal neurons (Horton et al. 2005).

The role of the golgins as tethering factors The functions of the golgins should be in agreement with the models of intracellular transport. Within the framework of the vesicular model, the most obvious function of the golgins would be their role as the tethering factors. The presence of extensive coiled-coil regions in the golgins suggests that they can also adopt long rod-like structures. Therefore, in the framework of the vesicular model, a tethering factor is considered as tethering ER- or Golgi-derived vesicles to the GA (Allan et al. 2000; Moyer et al. 2001; Sonnichsen et al. 1998). Thus, the golgins are believed to be involved in the tethering of vesicles. In the framework of the vesicular model of intracellular transport, these proteins were considered as the factor helping docking of transport-coat-dependent vesicles acting before the SNAREs. Tethering is defined as a “formation of physical links, between two membranes that are due to fuse, before the engagement of the SNAREs” (Whyte and Munro 2002). This process might represent the earliest stage at which specificity is conferred on a fusion reaction and it may involve multiple interactions. Two broad classes of molecules are proposed to have roles in tethering: a group of coiled-coil proteins (GM130 and p115, see below), and several large, multisubunit complexes (such as the TRAPP and COG complexes; see Chapters 2.7 and 2.6) (Whyte and Munro 2002). These long rod-like molecules, the golgins, are thus attractive candidates as factors that link the Golgi cisternae or capture the transport vesicular carriers in the proximity of the cisternae, prior to fusion (Gillingham and Munro 2003). The function of some of the golgins has been interpreted as an interaction between the cis-golgins, such as p115, and GM130, which tethers an incoming COPI or COPII vesicle to the cis-Golgi membrane, and that leads to the docking and subsequent membrane fusion mediated by the SNAREs (Shorter et al. 2002). P115 and molecules like giantin, which cooperate with it in tethering at the GA, have an extended conformation, a property that might enable them to initially bridge relatively large distances between membranes (Lesa et al. 2000; Seemann et al. 2000a,b; Sonnichsen et al. 1998).

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Two additional models are presently available for the role of GM130: one in which GM130 mediates homotypic tethering of neighbouring cisternae (Puthenveedu et al. 2006), and one in which GM130 maintains the ribbon structure through its mediation of the incorporation of EGCs (Marra et al. 2007), promoting homotypic fusion of the cisternal rims (Puthenveedu et al. 2006). However, these hypotheses do not explain the function of the other golgins, and especially those associated with the trans-side of the GA. If the vesicular model of transport is wrong, there is the need to explain golgin function within the framework of other transport models. Due to the problems that we have in the explanation of experimental data based on the vesicular model as the main model of intracellular transport (see Chapters 1.2 and 3.2), we need to consider other possibilities and to find the function of the golgins in the framework of other models of intra-Golgi transport. There is here some evidence in favour of the cisterna maturation model. Indeed, COPI vesicles dependent on the golgin CASP contain more concentrated Golgi enzymes than other COPI-dependent vesicles (Malsam et al. 2005).

The role of golgins in stacking GM130 and p115 are considered to form part of the Golgi matrix, which maintains the stacked cisterna architecture of the GA (Nakamura et al. 1995; Seeman et al. 2000a,b). GRASP65, and its related GRASP55, also have roles in Golgi stacking independent of GM130 (Barr et al. 1998; Shorter et al. 1999). P115, giantin, GM130 and GRASP65 have been shown to be required for both cisterna re-growth and cisterna stacking (two sub-reactions during in vitro Golgi reassembly after mitotic fragmentation) (Shorter and Warren 1999). The golgins could also provide the physical link between the Golgi cisternae, facilitating their stacking (Gillingham and Munro 2003). GM130/GRASP, giantin and p115 could thus be involved into the stacking of Golgi cisternae when the GA reforms following mitosis, a case of membrane tethering without subsequent fusion (Shorter and Warren 1999). Several lines of evidence suggest that the golgins could be important for stacking. In vitro, reconstruction of Golgi cisternae from mitotic fragments requires some tethering factors (Rabouille et al. 1995). Moreover, EM has identified proteinaceous bridges linking adjacent cisternae together (Franke et al. 1976; Cluett and Brown 1992). Finally, it has been shown that the HR2 domain of p115 has an essential role in the elongation of the Golgi cisternae (Sohda et al. 2007). For instance, p115 is required for the stacking of reassembling Golgi cisternae at an early stage in shack formation, before GRASP65 (Shorter and Warren 2002). Indeed, p115 is only required for cisterna stacking (Shorter and Warren 2002). Further, the acidic domain of p115 is not sufficient for the reassembly of Golgi cisternae, with p115 dimerization also reported to be needed (Dirac-Svejstrup et al. 2000). Some matrix proteins, such as the GRASPs, are considered as a “glue” to attach cisternae to each other.

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As originally proposed, this model has several weaknesses. First, most of the golgins are excluded from the space between the medial cisternae (Beznoussenko et al. submitted), which makes a role in stacking of the medial GA cisternae problematic. Then GRASP65 is not essential for € tterlin et al. 2005), as the GA shows a stacked organicisterna stacking (Su zation in GRASP65-depleted HeLa cells, where the number of cisternae per stack is reduced from a mean of 6 to a mean of 3 per stack. Precise analysis € tterlin et al. (2005) gives the impression that the cisternae of fig. 2B of Su became longer. However, as the Golgi localization of GM130 is not affected by GRASP65 depletion, its binding to GRASP65 cannot be the sole mechanism for its localization to Golgi membranes. In these GRASP65-depleted HeLa cells, VSV-G was transported to the cell surface with similar kinetics to the control, and after washout of nocodazole, Golgi membranes reassembled in the pericentriolar region of the cells, just as seen in the control cells. BFA-induced Golgi dynamics are also normal in the absence of GRASP65, and GRASP65 is important for bipolar spindle formation € tterlin et al. 2005). (Su

The role of the golgins in promoting attachment of CGN and TGN Thus, so far, three main factors have to co-exist to generate the Golgi ribbon: an intact MT star system, the input of membrane from the ER, and the activity of the different golgins that have important roles for the integration of the ER-derived membranes into the cisternae of neighbouring stacks, thus ‘glueing’ the separated stacks into a ribbon. The SNARE/Rab machineries are also important here. If these main preconditions are fulfilled, the concentration of the Golgi stacks within a narrow space (near the centrosome) inevitably leads to the generation of the ribbon. As such, the function of golgins might only be important for the generation of the Golgi ribbon and not for transport. However, the golgins are present not only in cells that can form a Golgi ribbon, but also in plant, yeast and insect cells, where the ribbon is absent. Similarly, p115 is found in all eukaryotes, and the GRASPs are found in all eukaryotes except plants, with Golgin-45 present only in vertebrates and GM130 present only in mammals. As with p115, Rab1 is found in all eukaryotes, while Rab2, as with its partner Golgin-45, is also only present in vertebrates (Short et al. 2005). Thus, in cells where the Golgi ribbon is not formed, some of the golgins are absent (plants, yeast). However, in general this machinery is present in all species. Importantly, cells for which the fragmentation of the ribbon during mitosis is more important have a more developed golgin machinery. The obvious question that arises is thus whether the function of the golgins is just for the generation of the ribbon? Detailed analysis of this aspect has revealed that in the absence of the golgins, and even when the Golgi ribbon is artificially broken by

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depolymerization of MTs, intracellular transport is impaired. Elimination of the function of p115 induces severe alterations in transport (see Table 1). Even GM130/GRASP65 depletion leads to a delay of transport and to less precise protein glycosylation (Marra et al. 2007). In turn, inhibition of Golgin-245 affects the function of the lysosomal system. After siRNAdepletion of Golgin-245, lysosomes show a more central pattern and there is accumulation of aberrant multivesicular structures (Yoshino et al. 2005). Finally, inhibition of the function of Golgin-97 slows down intra-Golgi transport, making protein sorting at the level of the trans-Golgi/TGN less precise (Beznoussenko et al. submitted) and affecting endosome-to-TGN transport (Lu et al. 2004). As depletion of each of the golgins has substantial functional effects (Tables 1 and 2), this suggests that the formation of the ribbon may not be the main function of the golgins, but actually represent a lateral effect of golgin function that appears when the Golgi stacks are centralized. In this case, the function of the golgins could be for the building of the functional Golgi stacks. Indeed, in cells lacking GM130 and devoid of MTs, the most cis cisterna of the stacks is absent both before and after the release of the transport block, whereas in control cells this cisterna is absent in the resting stacks but is visible in transporting stacks (Marra et al. 2007). Careful € tterlin et al. (2005) gives the impression that without analysis of fig. 2Bc in Su GRASP65, the attachment of the cis-most cisternae to the Golgi cisternae is affected. Moreover, examination of fig. 7 in Colanzi et al. (2000) suggests that Golgi fragments formed during incubation of permeabilized interphase cells with mitotic cytosol are not covered by the cis-most cisterna from the cis side and by the trans-most cisterna from the trans side. Instead, from the trans side they are often covered by the trans ER. Thus, within the framework of the non-vesicular models of intra-Golgi transport, and taking into account the hypothesis that the cis and trans golgins are responsible for the attachment of the CGN and the TGN (see Chapter 1.2), respectively, this means that in the absence of transport the medial GA is not covered by the attached CGN from the cis side and by the attached TGN from the trans side. The functional role of this Golgi configuration could be the regulation of the fusion–fission events that are important for protein and lipid sorting within, for example, the ‘kissand-run’ models of intra-Golgi transport. At least this hypothesis is in agreement with the finding that in the absence of GM130 the attached CGN is not formed even in transporting stacks (Marra et al. 2007), whereas after inhibition of Golgin-97 the attached TGN cannot replace the trans ER attached to the medial Golgi cisternae in resting stacks after restoration of intra-Golgi transport (Beznoussenko et al. submitted). This inability could affect protein sorting. Golgins could also be important for the correct sorting of post-Golgi carrier trafficking to the newly formed plasma membrane. For instance, the insect

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golgin Lava lamp (Lva) is required for cellularization, the rapid division of a large oocyte into small cells (Papoulas et al. 2005). During mitosis some golgins are phosphorylated. This phosphorylation can interfere with golgin function, and thus, during mitosis, the Golgi ribbon fragments and then more severe dispersion occurs due to loss of function of golgins, such as p115, GM130 and GRASPs (see Chapter 3.15), and maybe other golgins. One could imagine the following models for the function of golgins. At the ER exit sites, p115 and Golgin-84 regulate the maturation of the EGCs through their interactions with Rab1, SNAREs or other factors. The main function of p115 is its activity that is related to the centralization of EGCs. Golgin-245 has a similar function. Uso1/p115 can directly promote the formation of SNARE complexes, and thus of membrane fusion (Shorter et al. 2002). Additionally, p115 enhances the lipid activation of cytidylyltransferase (Feldman and Weinhold 1998), which also facilitates the maturation of EGCs. If the function of p115, Golgin-84 and Golgin-245 is normal and the carriers arrive at the central Golgi area, this event forces the cis-Golgi network to form the attached CGN, which facilitates the integration of the EGCs into the stack. Inhibition of golgin function would affect this maturation process, leading to a block in carrier centralization and peripheral fragmentation of the GA. CGN attachment is GM130 and GRASP65 dependent. GM130-containing EGCs develop the ability to undergo homotypic coalescence (Marra et al. 2001) and fusion, and as a possible consequence, they become larger and acquire a complex disc-like shape. The acquisition of GM130 and the development of this tendency to undergo homotypic fusion can be viewed as a sort of maturation process through which EGCs gradually acquire the properties and composition of the next compartment, i.e. the Golgi cisternae. Indeed, the arrival of a synchronized wave of EGCs into the stacks coincides with an immediate increase in the number of cisternae per stack and in the surface area of the cisternae (concomitant with a decrease in the surface area of the EGCs), thus indicating that the EGCs are integrally incorporated into the stacks. GM130 is a key player in this process (Marra et al. 2007). However, even for most studies of GM130, its ultimate mechanism of action remains to be defined (Puthenveedu et al. 2006; Marra et al. 2007). Further, an attachment of the cis-most cisterna of the CGN per se is not sufficient for the formation of the ribbon. To complete ribbon formation, it is necessary to have the attachment of the trans-most cisterna to the stack. The cisternal part of the TGN attaches to the membrane domain that contains mostly cargo. Thus, only the concentrated action of all of the golgins is sufficient for the generation of the Golgi ribbon and for the normal function of the Golgi stacks. The function of giantin is the most mysterious and needs additional study. Thus, the hypothesis about the role of golgins in the regulation of the structure of the cis and trans sides of the GA appear rather probable.

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Figure 2. Scheme showing the role of ER-to-Golgi transport, cis-Golgins and trans-Golgins for the formation of the Golgi ribbon. Two Golgi stacks are shown in the top panel, each comprising three cisternae and two ER exit sites (ERES). The second panel demonstrates the exit of cargo from the ER. The ER exit sites became larger. The third panel shows formation EGCs located between the ER exit sites and the stacks. The fourth panel shows the fusion of two EGCs. The bottom panels show the resulting structure of the GA when ER-to-Golgi transport is blocked (left-most panel), when ER-to-Golgi transport occurs in normal cells (central-left panel), when the ER-to-Golgi transport occurs in cells devoid of GM130 (central-right panel), and when ER-toGolgi transport occurs in cells where the function of Golgin-97 is inhibited (right-most panel). As a result, the Golgi ribbon is normal in the transporting cells expressing GM130, and fragmented in cells without ER-to-Golgi transport (small fragments), in transporting cells without GM130 (small fragments), and in transporting cells without Golgin-97 (larger and more variable fragments).

Conclusions The data presented here show that the organization of the Golgi ribbon depends on many factors, such as the ability of cells to induce the concentration of Golgi stacks within a restricted space, the normal functioning of the golgins and the SNARE/Rab machineries, and several other less studied factors (a possible scheme of the interactions of these factors is shown in Fig. 2). The main function of golgins seems to be the regulation of the Golgi ribbon. Characteristics of golgins are summarized in Table 1. To date though, experimental observations can, and should, still be interpreted within the framework of several models of intracellular transport.

Abbreviations GA GFP

Golgi apparatus green fluorescent protein

Golgi ribbon and the function of the golgins

GREG MT MTOC SEM TGN

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Golgi-resident GPI-anchored protein microtubule microtubule-organizing centre scanning electron microscopy trans-Golgi network

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resident GPI-anchored protein in maintenance of the Golgi structure. Mol Biol Cell 18: 1261–1271 Yang JS, Lee SY, Spanò S, Gad H, Zhang L, Nie Z, Bonazzi M, Corda D, Luini A, Hsu VW (2005) A role for BARS at the fission step of COPI vesicle formation from Golgi membrane. EMBO J 24: 4133–4143 Yang JS, Zhang L, Lee SY, Gad H, Luini A, Hsu VW (2006) Key components of the fission machinery are interchangeable. Nat Cell Biol 8: 1376–1382 Yoshino A, Setty SR, Poynton C, Whiteman EL, Saint-Pol A, Burd CG, Johannes L, Holzbaur EL, Koval M, McCaffery JM, Marks MS (2005) tGolgin-1 (p230, golgin245) modulates Shiga-toxin transport to the Golgi and Golgi motility towards the microtubule-organizing centre. J Cell Sci 118: 2279–2293 Young WW Jr (2004) Organization of Golgi glycosyltransferases in membranes: complexity via complexes. J Membrane Biol 198: 1–13 Zhang T, Hong W (2001) Ykt6 forms a SNARE complex with syntaxin 5, GS28, and Bet1 and participates in a late stage in endoplasmic reticulum–Golgi transport. J Biol Chem 276: 27480–27487

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Functional cross talk between membrane trafficking and cell signalling The Golgi complex as a signalling platform Michele Sallese

Introduction Eukaryotic cells are complex systems that are capable of fine-tuning their functioning following internal and external perturbations. Their plasma membranes bearseveraldifferentreceptorproteinsthatcansensetheexternal milieu and transduce information across the plasma membrane to cytosolic proteins. These, in turn, transmit the messages to other partners along signalling cascades that lead to the required cellularresponse. Such events are known as ‘cell signalling’, and these cascades have critical effects on the behaviour of a cell, since they can affect cell motility and growth, and indeed, apoptosis. According to some estimations (Venter 2001; Imanishi et al. 2004), approximately 10–12% of the genes in the human genome encode for signal transduction proteins, thus highlighting the relevance of these functions to the cell. Signal transduction is a highly dynamic and regulated process. The overstimulation of a cell is limited by multiple control mechanisms, which range from desensitization to endocytosis, and which can eventually lead to the down-regulation of a receptor. The desensitization process involves several molecular events that leads to the uncoupling of a receptor from its downstream effectors. In recent years, the extension of our understanding of signal transduction mechanisms has been accompanied by the emergence of an important role of endomembranes in these signalling processes (Sorkin and Von Zastrow 2002; Di Fiore and De Camilli 2001). Receptor internalization leads to a decrease in plasma-membrane signalling (as part of the desensitization process), although the endocytosed receptor can continue to engage in signalling pathways that are different from those activated when the same receptor was present at the plasma membrane. It is now clear that the endosomal membranes are not only a sorting compartment where plasma-membrane receptors are either committed towards the degradative lysosomal pathway or dephosphorylated and recycled back to the plasma membrane for a further round of signalling; but also provide a signalling membrane from where specific signals originate (Di Fiore and De Camilli 2001; Sorkin and Von Zastrow 2002). As well as the endosomal membranes, several studies have reported the presence of signalling proteins on different organelles, including the

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endoplasmic reticulum (ER), the Golgi complex, the mitochondria, and the nuclear membrane, although our knowledge of their roles in these locations remains vague (Sallese et al. 2006). The aim of this chapter is thus to bring together the available information relating to the presence and roles of signalling proteins on the membranes of the Golgi complex. The Golgi membranes host a variety of classical signalling molecules that are generally present and functional at the plasma membrane (Sallese et al. 2006). These range from kinases (Birkeli et al. 2003; Colanzi et al. 2003) to phosphatases (Lavoie et al. 2000), to phospholipases (Freyberg et al. 2003), heterotrimeric G-proteins (Wilson et al. 1994) and phosphodiesterases (Asirvatham et al. 2004), to name but a few (Donaldson and LippincottSchwartz 2000). The easiest explanation for their presence on the Golgi complex is that they can be visualized on endomembranes because they are in transit towards their final destination, the plasma membrane (Michaelson et al. 2002; Sallese et al. 2006). Indeed, ER and Golgi membranes function as a platform for the assembly of multiprotein complexes (e.g. heterotrimeric G proteins). In addition, these membranes host the palmitoyl transferases (Ohno et al. 2006), the function of which is an obligatory step in the acylation of many proteins before their transfer to their functional sites of action; again, generally the plasma membrane (Marrari et al. 2007). However, a growing body of evidence shows that these signalling proteins that are localized on endomembranes can be in their active state, thus suggesting that they can have a functional role in these locations (Bivona and Philips 2003). We envisage four different scenarios for the roles of these signalling proteins on the Golgi complex. First, the Golgi localized signalling proteins could participate into Golgi disassembly during the mitosis downstream to growth factor receptor activation on the plasma-membrane (Fig. 1). Second plasmamembrane receptors could signal to the Golgi, regulating its activity and its secretory functions (Buccione et al. 1996), (Fig. 1). Third, plasmamembrane receptors could use the Golgi membranes, exploiting them as a central location, or as a signalling platform or relay station for the integration of messages in their transfer to their final destinations, which could include the cytoskeleton, mitochondria, etc. (Bivona and Philips 2003) (Fig. 1). Fourth, the traffic itself could trigger a signalling cascade that is involved in the coordination of the various secretory compartments (Pulvirenti et al. 2008) (Fig. 1) or possibly could affect other organelles or other cellular functions besides membrane trafficking (Fig. 1). Of these four scenarios, to data, the last two have only been support by a few indications. We believe that they represent an important area of investigation that it will be worthwhile to explore in the future.

The phospholipases The lipid derivatives include a large array of molecules for which an involvement in key cell signalling functions is well established; however, more

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Figure 1. The Golgi is a signalling hub. This cartoon schematizes four possible signalling pathways involving the Golgi complex. First, the Golgi localized signalling proteins could participate into Golgi disassembly during the mitosis downstream to growth factor receptor activation on the plasma membrane (arrow 1). Second, plasma-membrane receptors could signal to the Golgi, regulating its activity and its secretory functions (arrow 2). Third, plasmamembrane receptors could use the Golgi membranes, exploiting them as a central location, or as a signalling platform or relay station for the integration of messages in their transfer to their final destinations, which could include the cytoskeleton, mitochondria, etc. (arrow 3). Fourth, the traffic itself could trigger a signalling cascade that is involved in the coordination of the various secretory compartments, or possibly could affect other organelles or other cellular functions besides membrane trafficking (arrow 4). G Golgi complex; IC intermediate compartment; ER endoplasmic reticulum; N nucleus.

recently, they have also emerged as regulators of membrane trafficking (De Matteis et al. 2005). It is still not clear if their roles in membrane trafficking impinge on their signalling properties or rely on changes in the physical properties of the membranes, and indeed the available data suggest that there are contributions from both sides. To accomplish these important tasks, their levels are tightly regulated by a wealth of lipid-modifying enzymes, including kinases, phosphatases and lipases. The phospholipases are lipid-hydrolysing enzymes that can use the phosphatidylinositols (PIs) and phosphatidylcholine (PtdCho) in cell membranes as substrates. They can be divided into three families based on the phospholipid bond that they hydrolyse: phospholipase A1 (PLA1) and PLA2 hydrolyse fatty acids esterified on the sn-1 and sn-2 positions, respectively generating free fatty acid and lysophospholipid (Fig. 2). PLC hydrolyses the bond between the

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Figure 2. The diacylglycerol is the centrepiece of a functional lipid network. The scheme represents the metabolic pathways of the main signalling lipids possibly involved in the regulation of the Golgi functions. The arrows indicate the transformation of one compound into another in virtue of the enzyme written on the arrow itself. The flash indicates activation of PKD. AA, arachidonic acid; LPA, lysophosphatidic acid; PA, phosphatidic acid, DAG, diacylglycerol; AcylCoa, acyl-coenzyme A; PtdIns(4,5)P2, phosphatidylinositol (4,5)-bisphosphate; IP3, inositol (1,4,5)-trisphosphate; PC, phosphatidylcholine; SM, sphingomyelin. PLA2, phospholipase A2, PLD, phospholipase D; PI-PLC, phosphatidylinositol-specific phospholipase C; PAP, phosphatidic acid phosphatase; DAGK, diacyglycerol kinase; PC-PLC, phosphatidylcholinespecific phospholipase C; CerS, ceramide synthase; CDase, Ceramidase; CERT, ceramide transporter; SMS, sphingomyelin synthase; BARS, brefeldin A ADP-ribosylated substrate; PKD, protein kinase D. Pyrrophenone, PLA2 inhibitor; D609, PC-PLC and SMS inhibitor; FB1, (fumonisin B1) CerS inhibitor; U73122, PI-PLC inhibitor.

glycerol and the phosphate, generating diacylglycerol and phosphoinositols (Fig. 2), whereas PLD hydrolyses the bond between the phosphate and the inositol moiety generating phosphatidic acid and inositols (Fig. 2). The activity of PLA1 is mainly concentrated in the lumen of the lysosomes and on extracellular membranes, and therefore they are outside the scope of this review.

Phospholipase A2 Four main subfamilies of PLA2 have been identified: secretory PLA2 (sPLA2); cytosolic, Ca2þ -dependent PLA2 (cPLA2); intracellular, Ca2þ -independent PLA2 (iPLA2); and platelet-activating factor acetylhydrolases (PAF-AHs) (Brown et al. 2003). cPLA2 contains a Ca2þ -dependent membrane-binding domain (C2 domain). The activation of plasma-membrane receptors coupled to intracellular Ca2þ stimulation promotes the association of cPLA2 with the ER and with Golgi membranes (Evans and Leslie 2004). The recruitment of

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cPLA2 to these membranes could simply suggest that they act as relay stations along a plasma-membrane-initiated signalling cascade. However, several lines of evidence indicate the involvement of cPLA2 in the regulation of membrane trafficking (Brown et al. 2003). Membrane trafficking involves the formation of tubular-shaped membranes that can act as carriers or as structural elements in specific compartments, including in the cis- and trans-Golgi networks (CGN and TGN) (Lippincott-Schwartz et al. 2001). Brown and co-workers (2000) have reported that when the enzymatic activity of PLA2 is inhibited, the BFA-induced tubules that emanate from the Golgi complex are impaired, together with the redistribution of Golgi proteins to the ER. Along the same lines, in vitro experiments performed on purified Golgi membranes have indicated that activators of PLA2 promote tubule formation (Polizotto et al. 1999). The possible mechanisms for these PLA2-mediated effects rely on the local production of inverted cone-shaped lysophospholipids that drive the formation of positive curvature, a process that is involved in tubule formation (de Figueiredo et al. 1998); however, a role for arachidonic acid as a signalling mechanism cannot be ruled out. It appears that PLA2 also has a role in membrane fusion, but unfortunately many reports are based on in vitro evidence, which makes it hard to assess the real contribution of PLA2 to fusion steps in membrane trafficking. Specifically, snake venom PLA2 increases the fusion of liposomes and isolated secretory granules (Blackwood et al. 1996). Moreover, the use of PLA2 inhibitors impairs endosomal fusion, a phenomenon that can be overcame by the addition of arachidonic acid (Mayorga et al. 1993). PLA2 inhibitors are also able to block endosomal fusion in vivo (de Figueiredo et al. 2000). Similar to that hypothesized for tubule formation, PLA2 could participate in membrane fusion by changing the local membrane composition through the production of lysolipids and free fatty acids, and/or via signalling cascades. From a physiology standpoint, PLA2 regulates the retrograde transport of proteins between the Golgi complex and the ER. The use of PLA2 inhibitors has shown an impairment of retrograde transport of a chimeric construct between the KDEL receptor and VSVG, the temperature-sensitive variant of the vesicular stomatitis virus G protein (de Figueiredo et al. 2000). Similar approaches have shown that PLA2 activity is important for the recycling of the transferrin receptor from the recycling endosomes towards the plasma membrane (de Figueiredo et al. 2001), and for the maintenance of the Golgi ribbon (de Figueiredo et al. 1999), with PLA2 inhibitors promoting the formation of separate Golgi stacks that remain in the perinuclear area (de Figueiredo et al. 1999).

Phospholipase D The human genome contains two phospholipase D (PLD) genes known as PLD1 and PLD2 (Freyberg et al. 2003). Both of these preferentially hydrolyze

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the PtdCho into phosphatidic acid (PA) and choline. The PLD1 protein is distributed throughout the cell, including on endomembranes, such as the Golgi complex, the ER, and the endosomes, with only a minor fraction at the plasma membrane (Freyberg et al. 2001, 2003). In contrast, PLD2 is mainly present on the plasma membrane, although one study has reported about 20% of PLD2 at the Golgi cisterna rims (Freyberg et al. 2003). Consistent with its intracellular localization, PLD is part of signalling pathways initiated by external stimuli via growth factors, cytokines, and G-protein-coupled receptors (GPCRs). PLD also participates in membrane trafficking events and maintenance of the Golgi structure, although it is not known whether the signalling and trafficking functions of PLD are linked; e.g., receptors activated at the plasma membrane could affect vesicular transport via the PLD on the Golgi complex. A few studies that have used isolated Golgi membranes and permeabilized cells have shown the involvement of PLD in the release of nascent secretory vesicles from the TGN (Ktistakis et al. 1996). PLD also shows transphosphatidylation activity, such that in the presence of a primary alcohol, the phosphatidyl group is transferred to the alcohol, forming a phosphatidyl-alcohol instead of PA (Freyberg et al. 2003). Indeed, the exploitation of this property of PLD has provided the main tool to interfere with PLD activity, and thus to improve our understanding of the role of PLD in membrane transport. Shields and co-workers (2000) used 1-butanol to demonstrate that the PLD product PA is required for ER-to-Golgi transport of VSVG, as well as for TGN-to-plasma-membrane transport. In addition, they showed that 1-butanol heavily alters the structure of the Golgi complex, raising doubts as to the real cause of this membrane transport inhibition. A recent report has also linked the transport of the cystic fibrosis transmembrane conductance regulator (CFTR) to PLD activity (Hashimoto et al. 2008). Specifically, treatment with 1-butanol impairs the exit of CFTR from the ER, a phenotype that can be rescued by exogenous addition of PA. CFTR transport defects have also been reported in cells knocked down for PLD1 using siRNAs (Hashimoto et al. 2008). In addition, the presence of high levels of PA impairs Golgi-to-plasma-membrane transport of CFTR. Altogether, these data indicate that PLD/PA is an important regulator of CFTR transport, and it would thus be worth exploring whether the pharmacological modulation of PA levels could help in the treatment of cystic fibrosis. PLD1 appears to be normally inactive, with its activity increasing through direct interactions with protein kinase C (PKC), and Rho and Arf family GTPases (Hammond et al. 1997). Phosphatidylinositol 4,5-bisphosphate (PtdIns45P2) can also promote PLD1 activity by interacting with its pleckstrin homology (PH) domain and KR motif (Sciorra et al. 1999; Brown et al. 1993). In contrast, PLD2 has a high basal activity (at least in vitro), and it also requires PtdIns45P2 although it responds poorly to PKC, Rho and Arf (Powner and Wakelam 2002; Freyberg et al. 2003).

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In molecular terms, PLD1-dependent membrane trafficking relies on Arf1 stimulation, and in turn, the PA produced promotes the membrane recruitment of the coatomer I (COPI) complex (by direct interaction with the b subunit), NSF, the kinesins, and Arf1 itself in a sort of positive feedback cycle aimed at the formation of membrane carriers (Ktistakis et al. 1996, 2003). The PLD product PA can also affect membrane trafficking by acting at the fusion step. Indeed, a recent study carried out with the yeast proteins indicated that PA has a dual role during membrane fusion: it is required for the correct localization of the SNAP25 family proteins, and for the stimulation of the fusion process, at least in vitro (Liu et al. 2007). PA is also a potent activator of PtdIns4P 5-kinase (PI4P5K), the final enzyme of the PtdIns45P2 synthetic pathway (Jenkins et al. 1994). Indeed PLD can stimulate PtdIns45P2 synthesis on isolated Golgi membranes in an Arf1dependent manner (Siddhanta et al. 2000). However, the presence of PtdIns45P2 on the Golgi complex remains a matter of dispute, despite some reports that have shown that Golgi PtdIns45P2 contributes to the recruitment of bIII spectrin and ankyrin, and that PI 5-phosphatase, the enzyme that catalyses the initial dephosphorylation of PtdIns45P2, is expressed on the Golgi membranes (Freyberg et al. 2003). On the whole, this evidence supports the presence of PtdIns45P2 on the Golgi complex. Along with its signalling role, PLD1 could participate in the structure and function of the Golgi complex by substituting PtdCho with PA inside the biological membranes. Thus the two leaflets of cell membranes lose their equilibrium through the change of the cylindrical lipid PtdCho into the conical lipid PA (PA has a smaller polar head), and so to cope with this, the system can establish a new equilibrium by membrane bending (Kooijman et al. 2005). In line with this hypothesis, there is the localized presence of PLD2 on the rims of the Golgi cisterna, where strong curvature is needed (Freyberg et al. 2002). Alternatively, the PA produced on the rims could be part of the recruitment process for proteins involved in membrane trafficking (see above), since the cisterna rims also represent a hot spot for post-Golgi carrier formation.

Phospholipase C The phospholipases C (PLCs) are a family of at least 13 enzymes that can be divided into six classes: PLC b, g, d, e, z, h (Rhee and Choi 1992). These preferentially hydrolyse PtdIns45P2 to generate two important second messengers: diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (Ins145P3) (Rhee and Bae 1997). These PLCs are generally part of the signalling pathways initiated by GPCRs and growth-factor receptors on the plasma membrane (Exton 1996). Intracellularly, the PLCs are localized in the cytosol, on the plasma membrane (where the major pool of PtdIns45P2 is located), in the nucleus and on endomembranes, including the endosomal compartment and the Golgi complex (Mazzoni et al. 1992; Bertagnolo et al. 1995; Blayney et al. 1998). The PLC product DAG is required for membrane recruitment of

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several signalling proteins, including protein kinases (e.g., PKC) and small Gprotein exchange factors (e.g., Ras–GRP1). Ins145P3 binds to and opens ERand Golgi-located Ca2þ channels, which leads to an increase in cytosolic Ca2þ levels and the consequent signalling promoted by this important second messenger. Interestingly, Bikle and co-workers (2007) have reported the presence of PLCg1 in a complex with the Ins145P3 receptor, and of secretory pathway Ca2þ -ATPase1 (SPCA1) on the trans-Golgi of keratinocytes. They thus hypothesized a role for PLCg1 downstream of the Ca2þ -sensor receptor (CaR) in the regulation of the Ca2þ stores of the Golgi complex (see Ca2þ section below). PLCb2 and PLCb3 are also present on the Golgi complex (Diaz Anel 2007). In contrast, the cytosolic PLCe translocates to Golgi membranes in response to Rap1 activation, as a downstream effector of GPCRs and growthfactor receptors (Jin et al. 2001). Rap1 is a small G protein of the Ras superfamily that is mainly localized on the Golgi complex. Originally it was discovered as an inhibitor of Ras functions (cell growth, adhesion; see below), as it competes with Ras for binding to Raf-1, but does not activate Raf-1. However, later it was shown that Rap1 can activate the ERK pathway via B-Raf, and it can even synergise with Ras. The PLCe recruited to Rap1-containing membranes (e.g., the Golgi complex) also acts as a Rap1 guanine nucleotide exchange factor (GEF) through its Cdc25-homology domain, generating a positive feedback loop that is responsible for the sustained activation of Rap1 at the Golgi complex (at least 20 min), as a downstream effect of EGF-receptor stimulation (Jin et al. 2001). We would also expect that a PLC contributing to DAG levels at the Golgi complex could regulate membrane trafficking. Indeed, reduction of DAG levels in the Golgi by down-regulation of the PItransfer protein Nir2 impairs TGN-to-plasma-membrane transport of VSVG while leaving unaltered the bidirectional ER-to-Golgi and intra-Golgi transport steps (Litvak et al. 2005). The importance of DAG in membrane trafficking has been emphasized by a series of studies that have shown that the TGN exit of carriers is dependent on the fissioning protein PKD (Baron and Malhotra 2002). This is also one of the most clear signalling cascades that has been identified on the Golgi complex to date. Briefly, the bg subunit of the heterotrimeric G proteins can activate PLCb3, which by generating DAG recruits PKCh and PKD to Golgi membranes. PKCh also phosphorylates and activates PKD, which in turn, phosphorylates and activates the PI 4-kinases (PI4Ks) (Diaz Anel and Malhotra 2005; Diaz Anel 2007). The PtdIns4P generated by this signalling cascade could lead to the recruitment of several adaptor proteins (four-phosphate-adaptor proteins 1 and 2, FAPP1 and 2; ceramide transfer protein, CERT; adaptor protein complex 1, AP-1,) that are necessary for the formation/fission of carriers at the TGN that are directed towards the plasma membrane (De Matteis et al. 2005). Moreover, in a possible negative feedback loop, PKD can also directly phosphorylate CERT (Fugmann et al. 2007). Ceramide belongs to the pathway that contributes to the maintenance of DAG levels on the Golgi complex via sphingomyelin synthase (Fig. 2). Phosphorylating CERT decreases

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the transport of ceramide to the Golgi complex, and as a consequence, the amount of DAG (Fig. 2). Finally, this CERT phosphorylation leads to the quenching of the PKD activating pathway. A recent study by Egea and co-workers (2007) has further emphasized the important role of DAG in membrane trafficking. Here they showed a specific role for DAG in the fissioning of COPI-coated buds during retrograde, Golgi-toER, transport. Furthermore, in investigating the potential pathways for DAG generation on the Golgi complex using chemical inhibitors, they showed that at least three DAG-generating pathways participate in the DAG levels at the Golgi complex: the classical PLC pathway; the PA phosphatase (PAP) pathway downstream of PLD; and the sphingomyelin synthase pathway, as a by-product of sphingomyelin (Fig. 2) (Fernandez-Ulibarri et al. 2007). However, the PAP substrate PA can also be derived from acylation of the PLA2 catabolite lysoPA (Fig. 2). This finding suggests that all of the phospholipases taking part in Golgi functioning show cross talk by converging on DAG (Fig. 2).

Calcium and Ras at the Golgi complex Ca2þ is perhaps the most ubiquitous of the second messengers in vertebrates, and even slight variations in its levels can greatly affect cell behaviour. Extracellular Ca2þ concentrations are usually in the low millimolar range, while resting free cytosolic Ca2þ concentrations are in the order of 100 nM. A large amount of intracellular Ca2þ is stored inside the ER and the Golgi complex; both of these organelles can accumulate millimolar levels of luminal Ca2þ (Montero et al. 1995; Pinton et al. 1998). These steep Ca2þ gradients between the organelle lumens and the cytosol, and between the cytosol and the extracellular space, are maintained by a series of ATP-dependent pumps and channels that fine-tune this homeostasis. According to the classical paradigm, receptor activation at the plasma membrane stimulates the release of Ca2þ from the ER stores into the cell cytosol, which then activates downstream signalling proteins. More recently, the Golgi complex has also been considered as part of the intracellular Ca2þ response that can be triggered by extracellular stimuli (Pinton et al. 1998). The ER and Golgi complex release Ca2þ upon activation of the Ins145P3 receptors in their membranes (Berridge 2002; Pinton et al. 1998), while Ca2þ uptake involves two classes of Ca2þ -ATPase pumps, the sarcoplasmic and ER Ca2þ ATPase (SERCA) in the ER and Golgi membranes, and the Golgi-specific Ca2þ ATPase SPCA1 (Missiaen et al. 2007). As well as pumping Ca2þ , SPCA1 also supplies Mn2þ as a cofactor for the Golgi complex glycosyltransferases. The importance of the Golgi complex in cellular Ca2þ homeostasis is highlighted by the skin disorder Hailey–Hailey disease, a keratinocyte disorder that is characterized by cell–cell adhesion and differentiation defects, and that is caused by an inactivating mutation in the SPCA1 Ca2þ -ATPase gene (Hu et al. 2000). The contributions of SERCA and SPCA1 to the homeostasis of Ca2þ in the Golgi complex is also cell-type dependent. Indeed, as keratino-

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cytes rely almost exclusively on SPCA1, this indicates why Hailey–Hailey disease is cell specific (Callewaert et al. 2003). Fluctuations in the Ca2þ concentrations in the extracellular environment are also monitored by the Ca2þ receptor (CaR). This is a low affinity, seventransmembrane-domain GPCR that is coupled to various heterotrimeric G proteins of the Gi, Gq and G12/13 classes (Ward 2004). In keratinocytes, CaR coupling to the PLC signalling pathway can trigger the release of Ca2þ from intracellular stores, and hence promote cell differentiation. Recent studies have shown that the CaR is also on the TGN, whereby it can sense the luminal Ca2þ concentrations and regulate the Ca2þ uptake into the Golgi complex accordingly, through acting in concert with PLCg1 and SPCA1 (Tu et al. 2007). If this finding can be confirmed in future studies, it represents the first evidence of a Golgi-initiated signalling circuit. Changes in Ca2þ homeostasis within the Golgi lumen or in the cytosol proximal to this organelle can affect the functions of the Golgi complex and cell signalling. Indeed, there is evidence that variations in the levels of cytoplasmic Ca2þ are involved in different transport steps (Chen et al. 2002). Specifically, a role for Ca2þ in endosomal fusion has been reported, as well as in homotypic vacuolar fusion in yeast (Peters and Mayer 1998; Pryor et al. 2000; Mayorga et al. 1993). In addition, Balch and Beckers (1989) identified a Ca2þ -dependent ER-to-Golgi transport step, while Elazar and Porat (2000) used a reconstituted intra-Golgi transport assay to demonstrate that Ca2þ released from the Golgi complex is required for intraGolgi transport of VSVG. These data would thus suggest that Golgi transport is controlled by a micro-signalling circuit that is triggered by the arrival of cargo at the Golgi complex and that leads to the regulation of transport flow (see Chapter 2.8). There is also clear evidence for a role for Ca2þ in constitutive exocytosis and endocytosis in whole cells, as provided by Stamnes and co-workers (2002), in agreement to previous studies performed with purified organelle membranes and semi-intact cell systems. Here they showed that in NRK cells, VSVG transport is impaired by the Ca2þ chelator BAPTA, for the intermediate compartment-to-Golgi and the Golgi-to-plasma-membrane transport steps (Chen et al. 2002). When probed with the Shiga toxin b fragment, the functioning of the endocytic/retrograde pathway was blocked by BAPTA treatment at the endosomal-to-Golgi and Golgi-to-ER interfaces. Ca2þ chelators also promoted the detachment of the COPI protein coat from membranes, providing the first mechanistic explanation of how Ca2þ regulates transport throughout the constitutive exocytic pathway (Chen et al. 2002). As well as these effects on the COPI machinery, Ca2þ can affect membrane trafficking by various means. In the first instance, an impairment of membrane trafficking could result from changes in the intraluminal Ca2þ concentrations. This hypothesis is supported by the presence of a number of Ca2þ -binding proteins (CALNUC, P54/NEFA and Cab45) that appear to be

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devoted to the control of the luminal Ca2þ concentrations of the Golgi complex (Morel-Huaux et al. 2002; Scherer et al. 1996; Lin et al. 1999; Kawano et al. 2000). Secondly, changes in peri-Golgi Ca2þ concentrations can affect the activities of Golgi-localized Ca2þ -dependent proteins and/or result in the recruitment of cytosolic signalling proteins that are involved in the initiation of signalling cascades similar to those seen at the plasma membrane. For example, Ca2þ increases in the proximity of the Golgi membranes can activate L-CaBP1, a Ca2þ -binding protein that negatively modulates Ca2þ release from the Golgi complex via the Ins145P3 receptor (Haynes et al. 2004). Recently, interactions have also been seen between L-CaBP1 and the AP-1 adaptor, suggesting a specific role of L-CaBP1 in the regulation of transport (Haynes et al. 2006). Furthermore, Ca2þ activates neuronal calcium sensor-1 (NCS-1), a protein that can stimulate PI4KIIIb at the TGN, leading to an increase in constitutive and stimulated transport from the TGN to the plasma membrane (Haynes et al. 2005). NCS-1 binding to Arf1 can also compete with Arf1 in the activation of PI4KIIIb, suggesting the presence of two mutually exclusive pathways acting upstream of PI4KIIIb in the regulation of exit from the TGN (Haynes et al. 2005). A peri-Golgi Ca2þ increase can also activate the cysteine protease calpain, which is involved in many different cellular processes, including cell adhesion and migration. However, a recent study showed that Golgi calpain proteolyses the b-subunit of the COPI coatomer (Hata et al. 2006). This suggests that the activation of calpain by Ca2þ , which could be released from the Golgi complex during the transport of cargo, modulates its own transport by promoting the disassembly of the COPI coatomer. There is a second class of proteins that although not generally localized on the Golgi membranes, they are recruited to the Golgi in response to a cytosolic Ca2þ increase. These proteins include other neuronal calcium-sensor family proteins (hippocalcin, Vilip-1 and neurocalcin-d), cPLA2, K-Ras and Ras-GRP (see below for further details relating to Ras-GRP) (O’Callaghan et al. 2002; Bivona et al. 2003; Evans and Leslie 2004; Lopez-Alcala et al. 2008). Hippocalcin has been recently reported to be involved in the activation of Rasmediated Raf1-activation along the MAPK pathway initiated by N-methyl-Daspartate (NMDA) and KCl (Noguchi et al. 2007). Since the Golgi complex hosts the Ras-dependent MAPK activation pathway, we can hypothesize that the translocation of hippocalcin to the Golgi complex upon increased cytosolic Ca2þ levels allows it to participate in this signalling cascade from this Golgi location. This scenario opens the possibility of cross talk between Ca2þ released from the Golgi complex during membrane trafficking (if any) and signalling coming from the plasma membrane.

Ras The Ras proteins are the founder members of a large family of small GTPbinding proteins, which now includes Arf, Rab and Rho. Ras itself comprises

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three proteins, N-Ras, H-Ras and K-Ras, which are involved in the regulation of cell proliferation and cell death promoted by plasma-membrane receptors. The Ras family members are involved in cell signalling via three main pathways: Raf1/Erk, PI3K/AKT and RalGDS (Rodriguez-Viciana and McCormick 2005; Rodriguez-Viciana et al. 2004). Mutations in these Ras proteins that impair their GTPase activities are responsible for several human diseases, including cancers. Recent studies have reported a substantial bidirectional trafficking of H-Ras and N-Ras between the Golgi complex and the plasma membrane (Quatela and Philips 2006). Indeed, using genetically encoded fluorescent probes that can sense Ras activation, Philips and co-workers (2002) have demonstrated that growthfactor stimulation transiently activates Ras on the plasma membrane, while its activation is sustained on the Golgi membranes, from where it can promote cell growth or differentiation. Ras activation on the Golgi complex could be due to the retrograde transport of Ras from the plasma membrane, or it could rely on a diffusible mediator that transduces the message from the plasma membrane to the Golgi complex. Although both hypotheses might be valid, the most convincing experimental data indicate that Ca2þ is the diffusible signal involved in the activation of Ras that is already on Golgi membranes. Specifically, an intracellular Ca2þ increase recruits the Ca2þ -dependent Ras exchange factor Ras-GRP1 to the Golgi complex and the RAS GTPase-activating protein (GAP) CAPRI to the plasma membrane, leading to Ras activation at the Golgi complex and its inactivation at the plasma membrane (Bivona and Philips 2003). Of particular importance here, further studies have also shown that the outcome of Ras activation (cell growth versus cell differentiation) depends on its subcellular location (Quatela and Philips 2006; Mor and Philips 2006; Mor et al. 2007a). In T lymphocytes, activation of the T-cell receptor (TCR) promotes cell growth, while that of the lymphocyte-function-associated antigen 1 (LFA-1) receptor regulates cell adhesion, with both pathways involving Ras signalling (Mor et al. 2007b). Thus TCR engagement induces activation of Ras specifically at the Golgi complex, while the co-stimulation of the TCR and the LFA-1 receptor leads to the activation of Ras at the Golgi complex and the plasma membrane (Mor et al. 2007b). Interestingly, both of these signals rely on Ras–GRP1, but the DAG that is produced via the TCR is provided by PLC, while the PLD2 and PAP pathway produces DAG downstream of the LFA-1 receptor. It is not clear whether this PLC activation that occurs downstream of the TCR is localized to the Golgi complex or the plasma membrane. From these examples, it is evident that Ca2þ released from the Golgi complex or the ER can activate the Ras–GRP1/Ras signalling pathway. It would therefore be of importance to investigate whether the Ca2þ present in the peri-Golgi area during membrane trafficking, could do this as well. In addition, understanding whether Ras activation can affect membrane trafficking is also an important aspect that needs further investigation.

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Additional Golgi signalling Other classes of signalling proteins are known to be present on the membranes of the Golgi complex. These are considered here only very briefly for the sake of space and because they have been recently reviewed by our group (Sallese et al. 2006). The kinases form the most important families of signalling protein, and they are also present on Golgi membranes, from where they can modulate membrane trafficking and cell growth (Sallese et al. 2006).

The cyclic adenosine monophosphate (cAMP)–PKA pathway Protein kinase A (PKA) is by far the most important cAMP effector. PKA is a tetrameric kinase that comprises two regulatory and two catalytic subunits. cAMP induces the dissociation of the inhibitory, regulatory, subunits, thus freeing the active catalytic subunits. cAMP signalling is also believed to rely on functional complexes in restricted areas, so as to limit and optimize their actions in space (Zaccolo et al. 2002). The cAMP ‘signalosome’ is comprised of a GPCR that can promote GTP loading on the alpha stimulatory subunit (Gas) of heterotrimeric G proteins via a conformational change, which in turn activates the enzyme adenylyl cylase for the production of cAMP. The cAMP formed stimulates the phosphorylating activity of PKA as the final effect. As well as the desensitization mechanisms that work at the level of the activated receptor, this pathway is counterbalanced by the action of the phosphodiesterases (PDEs), which catabolise the cAMP that is formed. This signalosome complex is maintained and organized through the actions of a scaffold protein known as AKAP (Beene and Scott 2007). Remarkably, all of these cAMP pathway modules are located on Golgi membranes, strongly suggesting a specific function at this location (Cheng and Farquhar 1976a,b; Maier et al. 1995; Pooley et al. 1997; Martin et al. 1999; Li et al. 2003). From the functional standpoint, cAMP accelerates ER-to-Golgi and Golgi-to-plasma-membrane transport, while PKA inhibitors block Golgito-plasma-membrane transport with minor effects, if any, on ER-to-Golgi transport (Muniz et al. 1996, 1997). Increases in cAMP also alter Golgi morphology, with the generation of tubular networks among the Golgi cisternae (Muniz et al. 1996). Finally, PKA activity is required for retrograde transport, at the endosome–Golgi and Golgi–ER interfaces (Birkeli et al. 2003; Cabrera et al. 2003). At the molecular level, PKA can affect multiple transport steps, since its activity promotes the recruitment of Arf1 to Golgi membranes, which can then be followed by the pleiotropic effects that are known to be downstream of this master regulator of membrane trafficking (Martin et al. 2000). PKA also phosphorylates the KDEL receptor, uncovering its COPIbinding motif, and some of the SNAREs, modulating their ability to support membrane fusion (Cabrera et al. 2003; Hong 2005). Clearly the cAMP–PKA system is involved in multiple trafficking stages. The question thus arises: is the cAMP generated specifically at its site of action, or as cAMP can freely diffuse for hundreds of microns, does it derive from a

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different cellular compartment? The experimental information available cannot at present distinguish between these two possibilities. However, it would appear more effective to control membrane trafficking in restricted sub-Golgi areas (e.g., CGN, CGN) using sudden local elevations of cAMP generation, in contrast to the need to establish a cAMP gradient that arises elsewhere and that would activate the whole perinuclear area.

Protein kinase C The PKCs are a class of second-messenger-dependent kinases that comprises 10 members that can be grouped into: the classical PKCs (PKCa, b and g) that are DAG and Ca2þ dependent; the novel PKCs (PKC d, e, h, m and q) that are DAG dependent and Ca2þ independent; and the atypical PKCs (PKCz and i/l) that are DAG and Ca2þ independent. Most of these are recruited to Golgi membranes through to their C1 domain, which can bind to DAG, ceramides and arachidonic acid (Schultz et al. 2004). Interestingly, exogenous addition of ceramides, or their generation through stimulation of the IFNg receptor, induces cell apoptosis via the translocation of PKCd and PKCe to the Golgi complex (Schultz et al. 2003; Kajimoto et al. 2001, 2004). The role of PKC in trafficking is, however, still not completely understood. Small chemical PKC inhibitors/activators affect intra-Golgi and TGN-to-plasma-membrane transport, as revealed by their effects on the prototypical cargoes VSVG and the glucosaminoglycans (GAGs) (De Matteis et al. 1993; Fabbri et al. 1994; Buccione et al. 1996). Similar effects have been seen by stimulating the plasma-membrane IgE receptor (Buccione et al. 1996). This work represents an example of the plasma-membrane receptors regulating the Golgi functions, as described in the introduction (second scenario). The regulation of membrane trafficking by PKC activation involves control of the recruitment of the Arf1–COPI machinery to Golgi membranes, although an alternative interpretation has proposed a catalytically independent action of PKC. (De Matteis et al. 1993; Fabbri et al. 1994). More recent work from Tisdale and co-workers has shown important roles for PKCi/l in the sorting of retrograde-directed material from the ER–Golgi compartment, through actions on COPI and GAPDH (Tisdale and Artalejo 2006; Tisdale 2000; Tisdale et al. 2004). In conclusion, the PKCs at the Golgi complex serve the dual roles of amembraneraffickingregulatorandanapoptosismediator.Nodataareavailable to support traffic-dependent activation of the PKCs.

Heterotrimeric G proteins: key regulators or simple passengers? The heterotrimeric G proteins are the family of GTP-binding proteins that transduce the downstream signalling from the seven transmembrane domain receptors (the GPCRs). They are formed from three polypeptides, the Gbg dimer and the Ga subunit, whereby the characteristics of this last define the

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group classification of the whole trimer, in terms of its specific functional subfamily. Members of the Gai, Gaq, Gas and Gaz subfamilies have been clearly demonstrated to be on the Golgi complex (Denker et al. 1996; Helms et al. 1998). Some studies have proposed that these G proteins can be visualized on the Golgi complex since the trimer, and probably the whole signalosome that also includes a GPCR, is assembled and post-translationally modified on the Golgi membranes (Marrari et al. 2007). In contrast to this view, manipulation of the expression levels or the activity states of the heterotrimeric G-proteins influences the functionality of the secretory system. Thus, addition of the Gai activator mastoparan or of an excess of Gbg (which will titrate out the Gai subunit) in permeabilized cells results in a block of VSVG exit from the ER, which would suggest that Gai is required for this transport step (Schwaninger et al. 1992). In polarized cells, membrane trafficking from the TGN to the plasma membrane follows two separate routes, which are directed toward the apical and the basolateral membranes (Lipschutz et al. 2001). It appears that Gai is specifically required only for the basolateral-directed cargoes, since pertussis toxin treatment (a Gai-inactivating toxin) blocks only this pathway (Pimplikar and Simons 1993a,b). In contrast, Gas is important for apical, and not basolateral, transport (Pimplikar and Simons 1993a,b). Gai2 and Gaz appear to be involved in the maintenance of the structure of the Golgi complex, since their over-expression can counteract the actions of the Golgi disrupting agent, nordihydroguaiaretic acid; in addition, Gaz inactivation induces the disassembly of the Golgi complex (Yamaguchi et al. 2000). The regulators of G-protein signalling (RGSs) are a relatively recently discovered family of multi-domain proteins that have been implicated in various signalling pathways (Willars 2006). They all share an RGS domain, which can stimulate the GTPase activities of the G proteins, and thus their inactivation. RGS proteins are present on the Golgi complex (RGS–GAIP) and on Golgi-derived carriers (RGS4) (Sullivan et al. 2000; Wylie et al. 1999, 2003). The specific function of GAIP remains to be defined, although RGS4 is known to directly interact with COPI to inhibit the transport of aquaporin and alkaline phosphatase, probably by sequestering COPI from the Golgi membranes (Sullivan et al. 2000). In conclusion, there are several lines of evidence that support roles for heterotrimeric G-protein signalling in secretory transport and maintenance of Golgi morphology, although a potential role of the Golgi complex in their transport and assembly cannot be ignored. What appears to be lacking, instead, is a coherent picture of the stimulus–receptor–signalling–effector pathways in which the heterotrimeric G proteins are involved.

Conclusions The relatively small number of signalling proteins that have been considered above represent only the proof of concept that there exist on the Golgi

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complex classical signalling pathways that can regulate its functioning. Numerous other signalling proteins have also been reported to be on Golgi membranes, although, as yet, without any apparent roles. In addition, it has been shown that alterations in the expression/function of signalling proteins can affect the behaviour of the Golgi complex, although these might still be the results of indirect effects exerted in another cellular compartment. All of these data firmly point to roles of the classical ‘plasma-membrane’ signalling pathways superimposed on the basic trafficking machinery. We can conclude that experimental studies strongly support the concept that signals originating from the plasma membrane converge on the Golgi and can have effects on membrane trafficking. However, these signals also use the Golgi membranes as a platform to affect functions other than secretion. In contrast, there remain very few indications for traffic-triggered signalling cascades. With many signalling proteins that have been shown to be present on the Golgi complex and to influence membrane trafficking, it is difficult to envisage their primary activation at the plasma membrane; their functioning would indeed fit better with a local (Golgi complex) activatory loop. The first clear evidence of this was provided by the unfolded protein response (UPR), whereby the presence of unfolded proteins inside the ER can stimulate three sensor receptors that are localized in the ER membranes, thus initiating the signalling pathways that lead to the removal of the problem. We therefore await the evidence for similar pathways that operate on the Golgi complex. We are confident that in the near future we will understand a lot more about the information flow and involvement of these Golgi-located signalling proteins. Thus, it would be important to understand what are the signals that trigger Golgi-based cascades? What is the sensor receptors involved? What are the molecular mechanisms in the amplification/integration of these cascades? What are the effectors of the primary signals to produce a specific phenotype?

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The role of the cytoskeleton in the structure and function of the Golgi apparatus Gustavo Egea and Rosa M. Ríos

Introduction Cellular organelles in mammalian cells are individualized membrane entities that often become spherical. The endoplasmic reticulum (ER) and the Golgi apparatus (GA) are exceptions to this rule, as they are respectively made up of a continuous tubular network and a pile of flat disks. Their unique shapes are regulated by molecular elements (Kepes et al. 2005; Levine and Rabouille 2005), including the cytoskeleton. All cytoskeletal elements, together with cytoskeleton-associated motors and non-motor proteins, have a role in the subcellular positioning, biogenesis and function of most organelles, being particularly relevant in the GA. The GA is the central organelle of the eukaryotic secretory pathway. While its basic function is highly conserved, the GA varies greatly in shape and number from one organism to another. In the simplest organisms like budding yeast Saccharomyces cerevisiae, the organelle takes the form of dispersed cisternae or isolated tubular networks (Preuss et al. 1992; Rambourg et al. 2001). Unicellular green alga (Henderson et al. 2007) and many protozoa like Toxoplasma gondii (Pelletier et al. 2002) and Trypanosoma brucei (He 2007; He et al. 2004) contain a single pile of flattened cisternae aligned in parallel. The organization of the GA in this manner is referred to as a Golgi stack, which usually contains two regions: one central and poorly fenestrated (compact) and other lateral and highly fenestrated (non-compact) (Kepes et al. 2005) (see the 3D modelling of a GA stack of a control NRK cell in Fig. 2). In fungi (Mogelsvang et al. 2003; Rossanese et al. 1999), plants (daSilva et al. 2004; Hawes and Satiat-Jeunemaitre 2005) or Drosophila (Kondylis and Rabouille 2003) many separate Golgi stacks are dispersed throughout the cytoplasm. In all these cases, each Golgi stack is associated with a single ER exit site (ERES), forming a secretory unit. In contrast, in most mammalian cells, the GA is a single-copy organelle shaped like a ribbon, containing numerous stacks joined by a tubular network and located near the nucleus (Ladinsky et al. 1999; Rambourg and Clermont 1986). In mammals, Golgi stacks are segregated from the ERES, and the Golgi ribbon is closely associated with the centrosome, the main organizing centre for cytoplasmic microtubules (MTOC) (Rios and Bornens 2003; Saraste and Goud 2007). The cytoskeleton imposes the localization of the GA. Depending on the cellular model, either microtubules (MTs) or actin filaments (AFs) have the

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greater influence (for instance, in mammalian and plant cells, respectively). Historically, the first cytoskeleton element to be linked to the GA and associated membrane trafficking was the MT (Thyberg and Moskalewski 1999). Some time later, both AFs and actin-associated proteins were clearly implicated (for recent reviews see Egea et al. (2006); Lanzetti (2007); Ridley (2006); Smythe and Ayscough (2006); Soldati and Schliwa (2006)), and more recently intermediate filaments (IFs) also appear to interact with Golgi membranes and participate in protein trafficking (Gao and Sztul 2001; Styers et al. 2005; Toivola et al. 2005). In this chapter, we provide a general overview of the structural and functional consequences of the coupling between the cytoskeleton and the GA in various cell models.

Microtubules and the structure and dynamics of the Golgi apparatus Microtubules and the structural integrity of the Golgi apparatus In non-polarized mammalian cells, the GA is closely associated with the centrosome, which is usually located near the nucleus at the cell centre (Rios and Bornens 2003) (Fig. 1). The spatial proximity of the GA and the centrosome has been known since Camilo Golgi’s time, but it has been confirmed by immunofluorescence studies in the last 20 years. The close association with the centrosome is maintained even under conditions where cellular architecture is undergoing major remodelling, which occurs during cell migration (Kupfer et al. 1982), fusion of myoblasts to form myotubes (Ralston 1993; Tassin et al. 1985b), the delivery of lytic granules at the immunological synapse (Stinchcombe et al. 2006), phagocytosis (Eng et al. 2007; Stinchcombe et al. 2006) or neuronal polarization (de Anda et al. 2005). However, this association is broken when MT dynamics is perturbed by drugs like nocodazole (NZ) or taxol (TX) (Sandoval et al. 1984; Wehland et al. 1983), which suggests that the main factor governing Golgi ribbon integrity and localization is the microtubular network (Thyberg and Moskalewski 1985, 1999). Pioneer studies of the role of MTs in the structural organization of the mammalian GA used drugs that favour MT disassembly (Robbins and Gonatas 1964). In the absence of MTs, the Golgi ribbon fragments, giving rise to discrete Golgi elements or mini-stacks, which are dispersed throughout the cell (Fig. 1). However, the GA does not need to be either intact or near the nucleus for protein transport or glycosylation (Rogalski et al. 1984). NZinduced Golgi mini-stacks localize at the peripheral ERES, thus enabling the cell to maintain secretory transport from the ER (Cole et al. 1996; Trucco et al. 2004). Therefore, the GA of mammalian cells lacking MTs resembles the normal state of affairs in plant cells and fungi, where Golgi architecture and function occur without centralization. Recent developments in microscope technology have advanced our understanding of the dynamics of MTs and GA interaction. For example, 3D electron microscope studies have allowed individual MTs to be modelled

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Figure 1. Microtubule–Golgi interaction. Immunofluorescence images of human retinal pigment epithelial cells fixed with methanol and double labelled for tubulin (green) and the Golgi protein GMAP210 (red). Cells were treated either with nocodazole (þNZ; 10 mM/3 h) to depolymerize MTs or with taxol (þTX; 10 mM/5 h) to induce the complete polymerization and stabilization of tubulin into MT bundles. Control, NZ- and TX-treated cells were also processed for electron microscopy analysis. Control cells show the characteristic ribbon-like arrangement of numerous adjacent Golgi stacks localised around centrioles (coloured in red). After NZ or TX treatments, numerous discrete Golgi elements appeared which maintained the characteristic stacked morphology (mini-stacks). Note that no significant differences in the ultrastructure of mini-stacks are seen between the two treatments. In TX-treated cells, Golgi mini-stacks are mostly localised to the cell periphery, whereas those in NZ-treated cells are uniformly distributed throughout the cytoplasm. Bars: 5 mm (immunofluorescence microscopy images) and 200 nm (electron microscopy images).

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and the relationships with cisternae to be analyzed in situ. MTs specifically associate with the first cis-cisterna over long distances. MTs also cross Golgi stacks at multiple points via non-compact regions and cisternal openings (Marsh et al. 2001). Time-lapse microscopy studies have revealed that the overall 3D arrangement of the GA near the centrosome is relatively stable (Presley et al. 1997; Scales et al. 1997; Sciaky et al. 1997) although thin tubules are constantly formed and detached from the lateral portions of the GA. After extending from the GA, the tubules break off and move along MTs to the cell periphery. Some move directly to the ERES and, once attached, they collapse into them delivering Golgi proteins to the ER (Mardones et al. 2006). The motion of membrane elements from the ER to the GA is also critically dependent on MTs. If MTs are depolymerized, Golgi proteins that have recycled back to the ER are exported into pre-Golgi intermediates, which then fail to move to the pericentrosomal region and consequently, remain stationary in the vicinity of ERES. Over long periods of time, these de novo structures acquire a normal Golgi stack morphology and become completely functional for secretion (Trucco et al. 2004). These studies indicate that the ability to form a Golgi stack is an intrinsic property of ER-derived membranes that does not require MTs. However, they are required to link stacks into a single organelle and to ensure its central location around the centrosome. Another interesting aspect is the relationship between the GA and stable MTs. These MTs are characterized by the presence of detyrosinated and/or acetylated tubulin. They have a longer half-life and are more resistant to NZinduced depolymerization (Schulze et al. 1987). Most stable MTs, which often appear short and convoluted under the fluorescence microscope, concentrate around the centrosome and colocalize with the GA (Burgess et al. 1991; Skoufias et al. 1990; Thyberg and Moskalewski 1993). Immunoelectron microscopy further demonstrated a close connection between detyrosinated MTs and vesicles transporting newly synthesized proteins from the ER to the Golgi (Mizuno and Singer 1994). Therefore, it has been proposed that there is a reciprocal relationship between MT stabilization and Golgi membrane dynamics. More than 10 years later, the molecular mechanisms mediating this relationship are now beginning to be unveiled (see below).

Microtubule-motor proteins In the current view, it is difficult to understand how MTs contribute to the Golgi structure without considering how MTs mediate in Golgi-associated transport functions. In non-polarized cells, MTs are organized in a characteristic radial pattern with minus-ends anchored at the centrosome and plusends extending toward the cell periphery. Since the GA localizes around the centrosome, the predominant-associated motor activity involved should be minus-end directed. In eukaryotic cells, the primary molecular motor for minus-end-directed movements is cytoplasmic dynein 1 (Hook and Vallee 2006). Movement of transport carriers from peripheral ERES to the GA in the cell centre along MTs is mediated by dynein (Corthesy-Theulaz et al. 1992;

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Harada et al. 1998). In contrast, movement of membranes from the GA back to the ER is plus-end-directed and mediated by kinesin-2 (Stauber et al. 2006). Kinesins are a large protein superfamily, most of whose members have plusend-directed activity (Miki et al. 2005). Dynein and kinesin are structurally similar, consisting of two functional parts: a motor domain that reversibly binds to the cytoskeleton and converts chemical energy into motion, and a tail that interacts with cargo either directly or through accessory chains (Caviston and Holzbaur 2006). The mechanisms by which motors interact with a diversity of cargoes and subcellular targeting sites are not completely understood. One candidate factor proposed to link dynein to endomembranes is the multiprotein complex dynactin, an essential activator for most cytoplasmic dynein functions. Dynactin contains 11 subunits organized into an elaborate structure. The best characterized subunits are the Arp1 filament, p150Glued and p50 dynamitin. p150Glued is a dimer that forms a coiled-coil and binds to dynein, the ARP1 filament and MTs. Dynamitin is required for maintaining the integrity of the complex (Schroer 2004). Expression of a dominant negative form of p150Glued or overexpression of dynamitin induces the fragmentation of the GA into multiple dispersed elements (Burkhardt et al. 1997; Quintyne et al. 1999). The movement of transport carriers from the ER to the GA along MT tracks is blocked under these conditions, as occurs in NZ-treated cells (Presley et al. 1997). Most likely, this blockade occurs at the earliest phases of ER protein export, since the Sec23p component the COPII complex interacts directly with dynactin. This interaction would facilitate the formation of transport carriers and their motion to the GA (Watson et al. 2005). It is therefore widely accepted that the dynein/dynactin motor is primarily responsible for ER-toGolgi transport and the localization of the GA in the cell centre. An alternative model postulates that dynactin plays a role in coordinating the activity of opposing MT-motors and in regulating their processivity (Berezuk and Schroer 2007; Deacon et al. 2003; Haghnia et al. 2007; Ross et al. 2006). Supporting this view, dynein and kinesin colocalize in the same membrane structures (Welte 2004). Even more relevant for Golgi dynamics, kinesin-2 interacts with dynactin (Deacon et al. 2003). This interaction appears to involve primarily the non-motor subunit of kinesin-2 (KAP3) and the p150Glued subunit of dynactin. Knockdown of KAP3 blocked the Golgi-to-ER pathway and disorganized Golgi membranes (Stauber et al. 2006). The observation that kinesin-2 binds the same dynactin subunit as dynein raises the possibility that it could act as a molecular switch that coordinates bidirectional trafficking. Recent data suggest that the scenario could actually be more complicated. Thus, dynein associates with Golgi membranes through several distinct mechanisms and dynactin, via p150Glued, also binds Golgi-associated nonmotor proteins. Dynein intermediate chain directly interacts with huntingtin, which has an important role in vesicle transport. Huntingtin silencing disrupts the GA in HeLa cells (Caviston et al. 2007). ZW10, a mitotic checkpoint protein

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that anchors dynein to kinetochores, also performs important functions in membrane traffic. These include dynein targeting to the GA and other membranes, but also SNARE-mediated ER–Golgi trafficking. ZW10 depletion provokes Golgi dispersal and decreases the frequency of minus-end-directed movements (Arasaki et al. 2006; Hirose et al. 2004; Vallee et al. 2006; Varma et al. 2006). Interestingly, the small GTPases Cdc42 and Rab6 also play a role in regulating motor recruitment to membranes (Chen et al. 2005a; Matanis et al. 2002). Coatomer-bound Cdc42 prevents dynein binding to COPI vesicles, and expression of constitutively active Cdc42 blocks translocation toward the cell centre of NZ-induced stacks and ER-to-Golgi carriers (Chen et al. 2005a). Rab6 family members are involved in some MT-dependent transport steps from the trans-Golgi network (TGN). When Rab6 is activated, BicaudalD1/2 is recruited to the TGN, which in turn recruits dynein–dynactin complexes (Hoogenraad et al. 2001; Matanis et al. 2002). It has been proposed that these complexes could participate in a recycling pathway that begins at the TGN and leads directly to the ER (Young et al. 2005). However, a later study showed that the major target for the fusion of Rab6-containing vesicles is the plasma membrane and that Rab6 regulates the transport and targeting of constitutive secretion vesicles. This study also reports an additional interaction of BicaudalD1/2 with kinesin-1. Therefore, the interactions between Rab6, BicaudalD1/2, kinesin-1 and dynein–dynactin complexes may contribute to the regulation of motor recruitment and co-ordination of their activities to specify directionality (Grigoriev et al. 2007; Saraste and Goud 2007). Defining the relative contributions of all of these mechanisms to the overall regulation of Golgi dynamics will require further investigation.

Role of the Golgi apparatus in microtubule dynamics A new concept is emerging concerning the role of the GA in MT dynamics: the GA acting as a secondary MTOC (Luders and Stearns 2007). The ability of Golgi membranes to assemble and stabilize MTs was first noticed in hepatic cells after NZ treatment (Chabin-Brion et al. 2001). During NZ recovery, short MTs were invariably seen to associate with Golgi mini-stacks. In addition, purified Golgi membranes were shown to contain a-, b- and g-tubulin and to support MT nucleation. However, this study did not resolve whether the MT nucleation was primarily carried out by the centrosome (the MTs then being released and anchored to Golgi membranes) or directly by the GA. Experimental support for the latter hypothesis came from the analysis of the mechanisms regulating the centering of a radial array of MTs in cells lacking centrosomes (Malikov et al. 2005). In stimulated cytoplasmic fragments of melanophores, pigment granules form a central aggregate that becomes the focal point from which MTs radiate. Radial MT arrays also form and become centralized in centrosome-free cytoplasts obtained from non-pigment cells. Strikingly, the GA appeared to be located in the centre of the cytoplasts, close to the MT aster (Malikov et al. 2004). Recently, the GA has been unambigu-

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ously identified as an MTOC by laser ablation of the centrosome (Efimov et al. 2007). MT-nucleation at the GA was shown to require g-tubulin complexes. To date, two g-tubulin-interacting proteins have been associated with the GA: GMAP210 and AKAP450/CG-NAP. The former is a cis-Golgi-associated protein that copurifies with MTs (Infante et al. 1999; Kim et al. 2007) and recruits g-tubulin complexes to the GA (Rios et al. 2004). GMAP210 depletion fragments the Golgi ribbon into elements that remain near the centrosome (Rios et al. 2004). AKAP450 localizes at both the centrosome and the GA (Keryer et al. 2003a, b; Larocca et al. 2004; Takahashi et al. 1999). It contains two MT-binding domains and interacts with g-tubulin complexes, thus providing MT-nucleating sites to the centrosome and, probably, to the GA as well (Kim et al. 2007; Takahashi et al. 2002). Interestingly, AKAP450 also interacts with p150Glued and the expression of a mutant that disrupts this interaction causes GA fragmentation and dispersion in a similar manner to that observed with the overexpression of dynamitin. In RPE-1 cells, many MTs are generated from the TGN, where microtubule plus-end-binding proteins CLASPs localize. These proteins stabilize pre-existing MTseeds by coating them, thus preventing their disassembly. In this regard, the centrosomal protein CAP350, which was originally believed to participate in MT-anchoring at the centrosome (Yan et al. 2006) actually stabilizes MTs enriched in the Golgi complex, and thus helps to maintain the integrity of the GA in the vicinity of the centrosome (Hoppeler-Lebel et al. 2007).

Relationship between the Golgi apparatus and microtubules in different cellular systems So far, we have focused on what we have learned in cells displaying a radial MT array with plus-ends facing toward the cell cortex and minus-ends anchored at the centrosome. Direct observation of MTs reveals that cell lines with well-defined radial MT arrays are really a minority. In contrast, most cell lines display a loosely organized MT array. These differences are due to variations in the number of centrosomal anchored MTs, which ranges from their totality (lymphocytes) to practically none (epithelia) (Bornens 2002). In parallel, the morphology of the GA varies from a fully compacted shape around the centrosome to a highly extended one (Rios and Bornens 2003). The differentiation of specialized cells types in multicellular organisms frequently leads to the generation of non-radial MT arrays, which better serve the specialized functions of these cells (Dammermann et al. 2003; Musch 2004). Two of the most representative examples of non-radial MT arrays are the polarized epithelial cell and skeletal muscle fiber. In polarized epithelial cells, MTs form an apico-basal array with their minus-ends concentrated near the apical surface and their plus-ends facing the basal domain (Mogensen et al. 2000). This array determines the membrane trafficking, which is central to the function of epithelia. In addition to vertically arranged MTs, cell lines derived from columnar epithelia (MDCK or Caco-2) also show networks of horizontal MTs both at the cell apex and the

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cell base. In hepatocytes, MTs converge underneath the bile-canalicular lumen at the apical surface (Dammermann et al. 2003; Musch 2004). The GA shows compact morphology and typically lies just apical to the nucleus, wellseparated from the microtubule minus-ends. In MDCK cells, the GA extends upwards from the nucleus to the apical portion of the cell. At the same time, in epithelial cells, membrane proteins are segregated into functionally and structurally different apical and basolateral domains. Whereas MTs seem to be important in the organization of apical exocytosis, the actin cytoskeleton seems to be the main organizer for basolateral secretion. However, neither the motors that participate in the exit of various classes of proteins from the TGN, nor the molecular interactions that allow MTs and actin filaments to modulate luminal and basolateral polarity are fully understood (RodriguezBoulan et al. 2005). Skeletal muscle fibers are multinucleate cells resulting from the fusion of mononucleate myoblasts in myogenesis. During this process, both pericentriolar proteins and MT nucleation sites redistribute from the centrosome to the nuclear periphery (Dammermann et al. 2003; Tassin et al. 1985a). In the same way, the GA is redistributed into smaller perinuclear elements that are formed by stacked cisternae (Tassin et al. 1985b). As in non-polarized cells, these perinuclear Golgi elements appear localized near the ERES and associated with stable MTs (Lu et al. 2001; Percival and Froehner 2007; Ralston et al. 1999, 2001).

The actin-based cytoskeleton and the Golgi apparatus The actin-based cytoskeleton and the structural organization of the Golgi apparatus in mammalian cells The first experimental evidence that AFs and the GA are linked was the observation that the GA invariably becomes compacted when a large variety of clonal cell lines are treated with actin toxins that either depolymerize (mainly cytochalasins and latrunculins) or stabilize and nucleate AFs (jasplakinolide) (di Campli et al. 1999; Lazaro-Dieguez et al. 2006; Valderrama et al. 1998, 2000, 2001) (Fig. 2). However, at the ultrastructural level, Golgi stacks from cells treated with actin-depolymerizing or -stabilizing toxins appeared different. Thus, the former mainly show swelled cisternae, while the latter have perforated/fragmented cisternae, which remain totally flat (Fig. 2). Supporting these observations, cisternae in NZtreated cells remain completely flat (Thyberg and Moskalewski 1999; Trucco et al. 2004), and the GA in cells treated with NZ plus actin toxins display the same ultrastructural alterations as those seen in cells treated with actin toxins alone (Lazaro-Dieguez et al. 2006). This indicates that there is no synergic cooperation between MTs and AFs controlling the cisterna morphology. Therefore, as a general rule, MTs determine the pericentriolar localization of the Golgi ribbon, whereas AFs maintain the shape and membrane integrity of cisternae.

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Figure 2. Actin filaments–Golgi interaction. NRK cells treated with the actin-depolymerizing toxin latrunculin B (þLtB; 500 nM/45 min) or the actin-stabilizing toxin jasplakinolide (þJpk; 500 nM/45 min) show a similar compaction of the Golgi complex when viewed under the fluorescence microscope. In the respective panels, we also show the resulting F- and G-actin pools. At ultrastructural level, LtB treatment mainly produced swelling of cisternae. In contrast, Jpk treatment only induced fragmentation of cisternae. We also display 3D reconstructions obtained from electron tomograms of the GA from control and actin toxin-treated cells. In control cells, both the central compact (c) and the lateral non-compact (nc) regions are seen. In LtB-treated cells, the predominant alteration is the swelling of cisternae (asterisks), but some cisternal perforation/fragmentation is also seen (arrowheads). Jpk treatment leave cisternae flat, but they show numerous perforations (arrows) and perforations/fragmentations (arrowheads). A perforation/fragmentation of a Golgi stack is indicated by the dashed red lines. We also indicate, in the respective 3D Golgi modelling panels, the intra-Golgi pH values obtained for each experimental condition (asterisk indicates a statistical significance of p  0.01 according to the Student’s t-test).

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Cisternae are always flat, despite the huge amount of cargo that is continuously crossing the Golgi stack. When the amount of cargo to be transported is much higher, then membrane continuities appear between cisternae (Trucco et al. 2004). The aforementioned ultrastructural changes induced by actin toxins indicate that AFs provide the necessary mechanical stability to cisternae to prevent their expected spontaneous swelling as a consequence of the hyperosmotic protein content in transit through the Golgi stack. By analogy with red blood cells, the unique flat morphology of cisternae could result from the structural organization of the spectrin-actinbased cytoskeleton present in the GA (Bennett and Baines 2001; De Matteis and Morrow 2000). In this respect, spectrin and ankyrin isoforms, actin, and an anion exchanger (AE2) are all present in Golgi membranes (Beck et al. 1997; Devarajan et al. 1996, 1997; Godi et al. 1998; Heimann et al. 1999; Holappa et al. 2001, 2004; Stankewich et al. 1998; Valderrama et al. 2000). Together with these, ion regulatory molecules such as vacuolar Hþ-ATPases (Moriyama and Nelson 1989) and cation (NHEs) exchangers (Nakamura et al. 2005) resident in the Golgi or in transit to the plasma membrane could contribute to this postulated actin/spectrin-based cisternal mechanical stability by providing the appropriate intra-Golgi ion and pH homeostasis. This is essential, on the one hand for Golgi-associated post-translational protein and lipid modifications (Axelsson et al. 2001) and on the other hand to keep the cisternae flat. Thus, bafilomycin A1, an inhibitor of vacuolar ATPases, both induces cisterna swelling and slows the Golgi-to-ER protein transport (Palokangas et al. 1998). Curiously, the cisterna swelling after AFs depolymerization is accompanied by an increase in the intra-Golgi pH (Fig. 2). The restitution of normal actin cytoskeleton organization after the removal of actin-depolymerizing toxins is followed by the normalization of cisterna morphology and the intra-Golgi pH (Lazaro-Dieguez et al. 2006). This correlation strongly suggests that AFs could interact and modulate the activity of (some) ionic regulatory proteins (vacuolar ATPases, anion and cation exchangers, ionic channels, pumps, and/ or transporters) present in Golgi membranes. This interaction would be highly similar to that observed for some of these proteins present at the plasma membrane. Therefore, we postulate that the equilibrium of osmotically active ions would maintain the flatness of Golgi cisternae in concert with the actin assembly state (Lazaro-Dieguez et al. 2006). At the same time, the Golgi-associated spectrin-actin cytoskeleton system could organize the secretory molecular machinery controlling the lateral distribution of the main Golgi membrane components (De Matteis and Morrow 2000). Thus, a physical barrier could be formed in the compact region of cisternae by the conjunction of the spectrin-actin-based cytoskeleton together with particular lipids (for example, cylindrical-shaped ones) and proteins (for example, glycosyltransferases). This would result in a permanent inhibitory membrane area for the biogenesis of transport carriers. Future research in the Golgi spectrin and ankyrin isoforms should provide significant insights into Golgi architecture.

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The actin-based cytoskeleton and the biogenesis and motion of Golgi-derived transport carriers The presence in Golgi membranes of molecular components that trigger actin polymerization (Arp2/3, Cdc42, cortactin, N-WASP, syndapin) and those that determine vesicular budding (coatomer and clathrin coats) and fission (dynamin) suggests their physiological coupling, as occurs at the plasma membrane during endocytosis. Actin assembly provides the necessary structural support that facilitates the formation of transport carriers in the lateral portions of Golgi membranes. This can be achieved by generating force through de novo actin polymerization, which in turn can be accompanied by the mechanical activity of actin motors (myosins). In respect to the former possibility, the actin nucleators Arp2/3 and Spir1 are present in the Golgi (Carreno et al. 2004; Chen et al. 2004; Kerkhoff et al. 2001; Matas et al. 2004). Their respective upstream regulators can be diverse. For the Arp2/3 complex, the more consistent are Cdc42-N-WASP and dynamin2-cortactin (Cao et al. 2005; Chen et al. 2004; Luna et al. 2002; Matas et al. 2004). At the trans-Golgi network (TGN), there is experimental evidence of the functional coupling between dynamin-mediated membrane fission and Arp2/3-mediated actinbased mechanisms (Cao et al. 2005; Carreno et al. 2004; Kerkhoff et al. 2001; Kessels and Qualmann 2004; Praefcke and McMahon 2004; Rozelle et al. 2000). Thus, the interference with dynamin2/cortactin or syndapin2/dynamin2 protein interactions blocks post-Golgi protein transport (Cao et al. 2005; Kessels et al. 2006). Fewer data are available on early Golgi compartments, but an interesting functional connection between actin polymerization governed by Cdc42, coatomer (COPI)-mediated transport carrier formation, and microtubule motor-mediated motion has been described (Chen et al. 2005a). Under the activation of the ADP-ribosylation factor 1 (ARF1), actin, coatomer and the Cdc42 are all recruited to Golgi membranes (Stamnes 2002). Cdc42 interacts with g-COP subunit of the COPI-coated transport carrier in a cargo receptor p23-sensitive manner such that coatomer cannot simultaneously bind to Cdc42 and p23 (Chen et al. 2005a, b). Interestingly, the activation of Cdc42 (Cdc42-GTP) inhibits the recruitment of dynein to COPI-coated transport carriers. In contrast, the prevention of the COPI–Cdc42 interaction by p23 stimulates dynein recruitment on Golgi-derived transport carriers, and hence their MT-based transport. Overall, this could provide a safe control mechanism by which a COPI-mediated transport carrier cannot be moved (through microtubule motors) until it is completely assembled (when Cdc42 does not bind to coatomer) (Hehnly and Stamnes 2007). Supporting this idea, the disruption of actin filaments as well as the activation of the Cdc42-N-WASP-Arp2/3 signaling pathway by the expression of the constitutively activated mutant of Cdc42 (GTP-bound) block the COPI-mediated Golgi-to-ER protein transport (Luna et al. 2002; Valderrama et al. 2001). Therefore, the local fine regulation of the actin dynamics state on the transport carrier assembly could represent an early step that precedes the scission of the transport carrier in the lateral portions of

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cisternae for its subsequent switching to MT tracks and motility (Egea et al. 2006). The coupling between actin polymerization and transport carrier biogenesis occurs both at the TGN and in early Golgi compartments, but some of the molecular mediators that regulate both processes are unevenly distributed in the GA. Thus, regarding actin polymerization, Cdc42 and Arp2/3 are present to varying degrees in the Golgi stack (cis/middle/trans-cisternae) and at the TGN, but N-WASP is absent from the trans/TGN (Matas et al. 2004). Cortactin, which like N-WASP also recruits Arp2/3, is visualized at the tips and buds at both the cis- and trans-cisternae (Cao et al. 2005). It is then reasonable to postulate that, at the TGN, cortactin substitutes N-WASP in the recruitment of the Arp2/3 that mediate in the post-Golgi protein transport. If so, the cis-totrans/TGN segregation of some fundamental components involved in actin polymerization would facilitate the targeting and assembly of the specific molecular machinery that participates in the membrane budding and fission occurring in Golgi compartments. Thus, the sequential protein interactions syndapin2- or dynamin2-cortactin-Arp2/3 and Cdc42-N-WASP-Arp2/3 are respectively restricted to the TGN and to early Golgi compartments, participating in this manner in the post-Golgi and in the ER/Golgi interface protein transport. A key aspect in the structure of polarized cells (epithelial and neuronal) is the maintenance of polarized organization based on highly specific sorting machinery for cargo destined to the apical or basolateral membrane domain at the exit of the TGN (Rodriguez-Boulan et al. 2005). In accordance with the localization of Cdc42 in the trans/TGN (Matas et al. 2004), the expression of constitutively active (GTP-bound) or inactive (GDP-bound) Cdc42 mutants slows the exit of basolateral protein markers and accelerates apical ones (Cohen et al. 2001; Kroschewski et al. 1999; Musch et al. 2001). The downstream effectors involved in the regulatory protein sorting induced by Cdc42 at the TGN are unknown. The integrity of AFs is necessary for efficient delivery of some (but not all) proteins to the apical domain. For example, in MDCK cells the apical delivery of sucrase-isomaltase, but not that of lactase-phlorizin hydrolase or gp80, occurs along AF tracks (Delacour and Jacob 2006; Jacob et al. 2003). Interference with actin dynamics using actin toxins variably affects the exit of apical- and basolateral-targeted cargo from the TGN. Moreover, actin does not participate in the TGN egress of lipid raft-associated GPI-anchored cargo (Lazaro-Dieguez et al. 2007). Actin nucleation/polymerization activity on Golgi membranes can also give rise to the formation of actin comet tails, which consist of filamentous actin and various actin-binding proteins that focally assemble and grow on a membrane surface (Welch and Mullins 2002). Actin tails have been observed in raft-enriched TGN-derived vesicles in certain experimental conditions (Rozelle et al. 2000), but this does not seem to be the most efficient mechanism to specify directionality to transport carriers. In this respect, MT and AF tracks are more suitable. However, analogously to what happens at the plasma

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membrane (Merrifield 2004; Merrifield et al. 2005; Perrais and Merrifield 2005), Golgi-associated Arp2/3-mediated actin polymerization generates a force. Depending on whether this acts on the lateral portion of the cisterna or on the transport carrier membrane, it could respectively facilitate the membrane elongation that precedes membrane scission or propel the transport carrier away from the cisterna. The presence of components of the Arp2/3 complex in cisternae and periGolgi transport carriers (Chen et al. 2005a; Matas et al. 2004) endorses both possibilities, but direct experimental evidence of either phenomenon is still lacking. However, an interesting in vitro approach has recently been reported (Heuvingh et al. 2007). These authors observed actin polymerization around liposomes composed of a specific lipid that facilitates the recruitment of the activated form of ARF1. This actin polymerization was dependent on Cdc42 and N-WASP present in HeLa cell extracts, and resulted in the formation of actin comets, which pushed the ARF1 liposomes forward. Tight control of the coupling between Golgi-associated actin polymerization and membrane elongation and fission reactions prevents the structural and functional collapse of the GA. Part of this control can be achieved by regulating the activation state of Cdc42 in Golgi membranes. The Cdc42 GAP (GTPase-activating protein) ARHGAP10 and GEFs (guanine nucleotide exchange factor) Fgd1 and Dbs are present in Golgi membranes (Dubois et al. 2005; Estrada et al. 2001; Kostenko et al. 2005). In addition, the low levels of phosphatidylinositol 4,5-biphosphate (PIP2) present in the Golgi (De Matteis et al. 2005) could also facilitate this control. Note that PIP2 synergizes with Cdc42-N-WASP and cortactin in Arp2/3-triggered actin nucleation/polymerization (Rohatgi et al. 2000; Schafer et al. 2002). A variety of independent experimental approaches show that Cdc42 is the only Rho GTPase that functions in the Golgi complex in mammalian cell lines (Fucini et al. 2000; Matas et al. 2005; Prigozhina and Waterman-Storer 2004; Valderrama et al. 2000). However, neurons seem to be an exception. Citron-N, a RhoA-binding protein and ROCKinase-II are both seen in the neuronal GA (Camera et al. 2003). Likewise, LIMK1, a kinase that specifically phosphorylates ADF-cofilin, localizes to Golgi membranes (Rosso et al. 2004). For a review of the role of actin and actin-binding/regulatory proteins in the GA of neuronal cells see (Bornens 2002). In addition to actin polymerization, myosins also generate a force, which can promote the formation of transport carriers and/or their movement away from Golgi membranes along AF tracks. Non-muscle myosin II mediates both Golgi-to-ER and post-Golgi protein transport (DePina et al. 2007; Duran et al. 2003; Musch et al. 1997; Stow et al. 1998). This myosin is a non-processive motor that directly interacts with Golgi membranes in vitro (Fath 2005). It is postulated that this motor is tethered to the cisterna by its tail and to actin filaments by its head. Its subsequent motion along actin filaments could provide the force needed to extend Golgi-derived membranes away from the cisterna. This would be similar to what happens to tubules originating

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from liposomes incubated with microtubular motors (kinesin) moving along microtubules (Roux et al. 2002). This cisterna-derived membrane extension could facilitate the subsequent functional coupling of membrane scission protein(s), leading to the complete release of the transport carrier. The binding of a tropomyosin isoform to Golgi-associated short actin filaments (Percival et al. 2004) could facilitate non-muscle myosin II recruitment and its interaction with them. Myosin VI is another myosin motor located in the GA (Buss et al. 2004; Warner et al. 2003). It differs from other processive myosins (for example, myosin V and X) as it only moves transport carriers towards the fast-depolymerizing minus-end pole of the microfilament. Therefore, myosin VI could provide the force and directionality for the transport carrier movement away from cisternae according to the expected fast-growing plus-end polarization of the actin filaments originated from Golgi membranes (Chen et al. 2004). The interaction between myosin VI and optineurin, a partner of Rab8 (Sahlender et al. 2005) extends to the Golgi–actin cytoskeleton interaction the known role of some Rab proteins as linkers of endomembrane systems to cytoskeletal motors (Jordens et al. 2005). Myosin VI, together with optineurin and Rab 8, operates in protein sorting and transport at the TGN in polarized cells (Au et al. 2007). The myosin VI–optineurin complex is required in the basolateral protein sorting pathway mediated by the Golgi-associated clathrin adaptor protein AP-1B, which in turn is specifically regulated by Rab 8 (Ang et al. 2003). The inhibition of myosin VI results in the incorporation of basolateral membrane proteins into apical transport carriers and their delivery to the apical plasma membrane domain. Sorting of other basolateral or apical cargo does not involve myosin VI. This result suggests that myosin motors could selectively couple protein sorting and transport carrier biogenesis and motility. Finally, class I myosins are also reported to associate with Golgi membranes and on apical Golgi-derived vesicles from polarized cells (Fath and Burgess 1993; Jacob et al. 2003; Montes de Oca et al. 1997). In myosin Ia knock-out mice, apical markers sucrase–isomaltase and galectin-4 are mislocalized to the basolateral surface in intestinal epithelial cells (Tyska et al. 2005). The sorting ability of myosin I could be linked to its capacity to interact with lipid raft-associated cargo as this monomeric, non-processive motor binds to phospholipid vesicles (Hayden et al. 1990).

The Golgi apparatus–actin interaction in other cellular models Yeast The use of a large number of mutants that produce alterations in intracellular traffic in the budding yeast Saccharomyces cerevisie has led to the identification of proteins involved in both membrane trafficking and actin organization (Kaksonen et al. 2006; Mulholland et al. 1997). Most components of the secretory pathway and many of the actin-based cytoskeleton are conserved between yeast and mammalian cells. The actin cytoskeleton in yeast consists primarily of cortical patches and cables (Adams and Pringle 1984; Kilmartin and

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Adams 1984; Moseley and Goode 2006). AFs, and not MTs, polarizes growth in yeast (Novick and Botstein 1985). In Saccharomyces, MTs have not been implicated in the dynamics of any organelle except the nucleus (Rossanese et al. 2001). Many actin mutants accumulate large secretory vesicles and exhibit phenotypes consistent with defects in polarized growth (Pruyne et al. 2004). This, together with the polarized organization of actin cytoskeleton, has suggested a role for actin in the polarized transport of late secretory vesicles to the plasma membrane (Finger and Novick 2000; Mulholland et al. 1997). Thus, a mutation of GRD20, a protein involved in sorting in the TGN/endosomal system, showed aberrant secretion of the vacuolar hydrolase carboxypeptidase Y, but not other TGN membrane proteins, as well as defects in the polarization of the actin cytoskeleton (Spelbrink and Nothwehr 1999). Recently, depletion of Av19p in a strain that also lacks Vps1 (dynamin) and Apl2 (adaptor–protein complex 1) proteins results in secretory defects, accumulation of Golgi-like membranes, and a non-polarized actin cytoskeleton organization (Harsay and Schekman 2007). Finally, concentration of late (but not early) Golgi elements in the sites of polarized growth (the bud) depends on actin, which is transported along actin cables by type V myosin Myo2p (Rossanese et al. 2001). With regard to the early secretory pathway, AFs depolymerization with actin toxins does not affect ER-to-Golgi (Brazer et al. 2000) or Golgi-to-ER (M. Muñiz, personal communication) protein transport. Taken together, these results demonstrate that in yeast, actin organization directly participates in post-Golgi vesicular transport and in the Golgi inheritance.

Drosophila In adherent S2 cells derived from mixed Drosophila melanogaster embryonic tissues, it has recently been reported that Golgi inheritance occurs by duplication to form a paired structure. This process requires an intact actin cytoskeleton and depends on Abi/Scar but not WASP (Kondylis et al. 2007). In another recent study, the analysis of a genome-wide RNA-mediated interference screen in these cells showed that the depletion of the tsr gene (which codifies for destrin, also known as ADF/cofilin) induces Golgi membranes to aggregate and swell, resulting in inhibition of the HRP secretion (Bard et al. 2006). Coronin proteins dpdo1 and coro regulate the actin cytoskeleton, but also govern biosynthetic and endocytic vesicular trafficking, as indicated by mutant phenotypes that show severe developmental defects ranging from abnormal cell division to aberrant formation of morphogen gradients (Rybakin and Clemen 2005).

Dictyostelium The slime mould Dictyostelium discoideum (like Drosophila) is widely studied in developmental and cell biology. Cells of this protist are easy to manipulate by genetic and biochemical means. They contain various types of vacuole, ER and very small Golgi stacks (Becker and Melkonian 1996). Comitin (p24) is a dimeric Dictyostelium actin-binding protein present in the GA and in vesicle

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membranes that contains sequence motifs homologous to lectins. It is postulated that this protein may bind Golgi-derived vesicles to the actin filaments via the cytoplasmically exposed mannosylated glycans (Jung et al. 1996; Weiner et al. 1993). Villidin is another actin-binding protein present in this organism that seems to be associated with secretory vesicular and Golgi membranes (Gloss et al. 2003). LIS1 (DdLIS1) is a centrosomal protein required for the link between MTs, the nucleus and the centrosome that also controls the GA morphology. Mutants of this protein lead to MT disruption, Golgi fragmentation and actin depolymerization (Rehberg et al. 2005).

Caenorhabditis elegans Very little is known about the GA and actin cytoskeleton interaction in this organism, but consistent with a possible role of coronin 7 in Golgi trafficking (Rybakin et al. 2004), depletion of POD-1 gene (a Coronin 7 homolog) using RNA interference leads to aberrant accumulation of vesicles in cells of the early embryo (Rappleye et al. 1999). Moreover, CRP-1, a Cdc42-related protein, localizes at the TGN and recycling endosomes. Alteration of CRP-1 expression in epithelial-like cells affected the apical but not the basolateral trafficking (Jenna et al. 2005).

Plant cells The structural organization of the GA in plants has many points in common with animal cells but there are important differences, which are largely dependent on the different cytoskeleton organization of plant cells. Thus, interphase higher plant cells (angiosperms and some gymnosperms without flagellate sperm) lack doublet and triplet MTs and a single MTOC. Instead, numerous MTOCs are aligned in the cortex, which assemble and form the transverse bands referred to as cortical MTs. These MTs are essential for the transport of Golgi-derived vesicles formed during metaphase (Segui-Simarro et al. 2004), which subsequently fuse to form the phragmoplast (Jurgens 2005), the equivalent to the contractile ring in animal cells. In contrast to MTs, stationary AFs are most prominent in plant cells (known as actin bundles) where they are all oriented with the same polarity and aligned along the plant cell. Attached to the actin bundles are the ER, vesicles and numerous discrete or a few clustered Golgi stack-TGN units (also named Golgi bodies or dictyosomes). Importantly, Golgi units are highly variable in number (a few tens to hundreds) depending on the plant type, cell type and the developmental stage of the cell (Hawes and Satiat-Jeunemaitre 2005; Kepes et al. 2005). In polarized root hairs and pollen tubes, the TGN is a vesicular-like non-tubular compartment morphologically segregated from Golgi stacks. It localizes to growing tips of these cells, where together with actin, plant Rho/Rac members (ROPs and Rac1, respectively), Rab (Rab4a and Rab11) and ARF (ARF1) small GTPases regulate vesicular secretory and endocytic trafficking (Samaj et al. 2006). In plants, most of the endomembrane compartments are in constant movement together with the cytoplasmic streaming whereby cellular metab-

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olites are distributed all over the cell (Shimmen and Yokota 2004). ER vesicles and the Golgi units show actin-dependent dispersal and spatial organization and are propelled by the plant-specific myosin XI (Boutte et al. 2007). Discrete Golgi units contains a fine fibrillar material enriched in actin, spectrin and myosin-like proteins (especially the former) (Mollenhauer and Morre 1976; Satiat-Jeunemaitre et al. 1996). The depolymerization of AFs with actin toxins uncouples the association between specific regions of cortical ER with individual Golgi bodies (Boevink et al. 1998; Brandizzi et al. 2003). Thus, cytochalasin or latrunculin treatments induce the aggregation of Golgi bodies and variably alter the Golgi morphology. However, the latter depends on the cell type examined and the period of treatment (Chen et al. 2006; Satiat-Jeunemaitre et al. 1996). Actin toxins also perturb the coordinated movement of Golgi bodies and ER tubules (daSilva et al. 2004; Yang et al. 2005). Actin does not participate in the ER/Golgi interface protein transport (Saint-Jore et al. 2002), but it does it in post-Golgi trafficking to the plasma membrane and the vacuole. Thus, in the tip of growing cells like pollen tubes, AFs are the tracks on which Golgi-derived secretory vesicles are transported (Picton and Steer 1981; Vidali et al. 2001). Cargoes containing polysaccharides and the enzymes necessary for cell-wall morphogenesis also require an intact actin–myosin system (Blancaflor 2002; Hu et al. 2003; Miller et al. 1995; Nebenfuhr et al. 1999). Therefore, post-Golgi trafficking and the organization of vacuoles in plant cells require an intact actin cytoskeleton (Uemura et al. 2002).

The Golgi apparatus-intermediate filaments interaction IFs are found in nearly all animal cells. They are classified according to their distribution in specific tissues. In contrast to MTs and AFs, IFs do not exhibit polarity or bind nucleotides, and they are considered a more stable structure. IFs are of intermediate size (8–12 nm) in comparison to MTs (23–25 nm) and to AFs (6–8 nm). IFs maintain cell and tissue integrity thanks to their mechanical properties, cellular distribution and, as far as we know, from disease-associated IFs phenotypes. IFs participate in the regulation of key signaling pathways that control cell survival and growth, and also in protein targeting and membrane trafficking (Coulombe and Wong 2004; Kim and Coulombe 2007; Omary et al. 2004; Oriolo et al. 2007; Styers et al. 2005; Toivola et al. 2005). IFs extend from the plasma membrane to the nucleus in close vicinity to some organelles such as mitochondria, endocytic compartments, and the GA (Fig. 3). The first Golgi–IF interaction was reported for vimentin filaments at ultrastructural level (Katsumoto et al. 1991), and later confirmed biochemically (Gao and Sztul 2001). The interaction is mediated by the Golgi membrane-associated protein formiminotransferase cyclodeaminase (FTCD), a metabolic enzyme involved in conversion of histidine to glutamic acid (Gao et al. 1998). Overexpression of FTCD resulted in the formation of extensive FTCD-containing fibres originating from the GA and inducing its fragmentation, and whose fragments remained tethered to these fibers (Gao and Sztul,

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Figure 3. Intermediate filaments–Golgi interaction. The upper panels show an NRK cell double stained to reveal the association of the GA (using anti-mannosidase II antibodies; in red) with a network of vimentin intermediate filaments (in green) seen by immunofluorescence. Bar: 2 mm. The bottom panels illustrate the severe Golgi fragmentation that occurred in transfected Huh7 cells expressing the GFP-cytokeratin18 R89C mutant. The GA was revealed with anti-galactosyltransferase (GalT) antibodies. Neighbouring non-transfected cells show a normal GA (Kumemura et al. 2004) (images used with permission of the authors and the publisher). Bar, 4 mm.

2001). However, the GA appears to be normal in vimentin–null cells (Gao et al. 2002; Styers et al. 2004). Thus, it is postulated that the vimentin–FTCD interaction at the GA is essential for FTCD functionality, but not linked to the maintenance of Golgi organization. Oxysterol-binding protein (OSBP) regulates lipid and cholesterol metabolism and interacts with the GA in the presence of oxysterol (Ridgway et al. 1992). A splice variant form of the OSBPrelated protein 4 in which the PH domain and part of the oxysterol-binding domain are deleted, colocalizes with vimentin IF in the Golgi region and inhibits the intracellular cholesterol transport pathway mediated by vimentin (Wang et al. 2002). Epithelial cells express cytokeratins, whose mutations are also associated with epidermal, oral and ocular diseases (Uitto et al. 2007). Arginine 89 of cytokeratin18 plays an important role in IF assembly. The expression of this mutant cytokeratin-induced aggregations, loss of the cytokeratin cytoskele-

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ton and the fragmentation of the GA (Fig. 3). Moreover, the Golgi reassembly occurring after NZ or brefeldin A treatments was perturbed only in cells expressing both cytokeratins and vimentin IFs (Huh7 and OUMS29 cells), but curiously not in cells that only possess vimentin IFs (HEK293 cells) (Kumemura et al. 2004). Whether cytokeratin or vimentin IFs are involved in Golgi-associated protein sorting, vesicular formation and/or transport remains to be established. However, recent evidence indicates that IFs could regulate some membrane protein targeting events, which take place at the GA level. Thus vimentin directly binds AP-3 and thus regulates protein sorting in endo-lysosomes (Styers et al. 2004). Polarized enterocytes and hepatocytes depleted of some keratins by antisense strategies and in cytokeratin 8-null mouse cells showed altered apical protein transport (Ameen et al. 2001; Rodriguez et al. 1994; Salas et al. 1997). Maturation of glycosphingolipids is also impaired in vimentin-deficient cells, but the defect seems to be localized to the Golgi/endosomal interface transport (Gillard et al. 1994, 1998). Since MT-motors kinesin and dynein also control the dynamics of IFs (Helfand et al. 2002, 2003; Prahlad et al. 1998), the reported Golgi/endo-lysosomal membrane trafficking mediated by IFs may be under the control of the dynamic IF–MT interaction, which can also be applied to AFs and motors. In this respect, the actin-based motor myosin Va has been identified as a neurofilament-associated protein (Rao et al. 2002). In summary, the structural and functional interaction of the GA with IFs is not yet firmly established. However, since IFs are well-integrated with both actin and microtubule cytoskeletons and their motors (Chang and Goldman 2004), the organization of the GA may also be influenced by the organization and dynamics of IFs.

Conclusion Both MTs and AFs are necessary for correct Golgi positioning, architecture and trafficking. Strong evidence in favour of this view now indicates that the GA functions as a microtubule and as an actin-nucleating organelle. In general terms, the relationships between each cytoskeleton network and Golgi dynamics are complementary. Thus, in animals cells, the actin-dependent cytoskeleton (AFs and actin-binding/regulatory proteins) plays an important role in early events of vesicular transport (sorting and/or membrane fission), and in the maintenance of the flattened morphology of cisternae. Furthermore, MTs and associated motors are directly involved in the motion of Golgiderived transport carriers to their final destinations and in the positioning and organization of the Golgi ribbon. In contrast, in plant cells, endomembrane organization and trafficking are almost exclusively mediated by AFs. Both ER and individual Golgi stacks are directly anchored to actin bundles, and transport between the ER and the GA is cytoskeleton-independent. In yeast, much less is known, but both cytoskeleton elements participate in post-Golgi protein transport and Golgi inheritance.

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At the apex of complexity, we envision the GA of mammalian cells to be assembled in three stages, in which the cytoskeleton participates in a variable extent: the first flattens cisternae (which is actin-dependent); the second maintains the discrete stack structure in the tight parallel arrangement of a variable number of cisternae (which at the moment seems to be independent of cytoskeleton proteins), and the third maintains stacks together to produce the classical single ribbon-like Golgi structure (which is fully dependent on MTs). Variations in the relative contribution of each of these three steps could generate the diversity of GA arrangements observed in different biological systems.  s Antón and members of our Acknowledgements. We thank Michel Bornens, Ine zarorespective labs for their comments, as well as Sabrina Rivero and Francisco La  guez for help with figures. The work carried out in our laboratories has been Die supported by grants from Ministerio de Educación y Ciencia (G.E. and R.M.R.), Junta de Andalucía (R.M.R.), and Distinció Award from the Generalitat de Catalunya (G.E).

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The dynamin–cortactin complex as a mediator of vesicle formation at the trans-Golgi network Shaun Weller, Hong Cao and Mark A. McNiven

Dynamin function The conventional dynamins represent a family of large GTPases that are encoded by at least three distinct genes in mammalian tissue and contain four conserved domains; an N-terminal highly conserved tripartite GTP-binding domain located within the first 300 amino acids, a pleckstrin homology (PH) domain of 100 amino acids, a coiled-coil (CC) region and a modestly conserved proline-rich domain (PRD) at the C terminus. Several in vitro and in vivo studies have demonstrated convincingly that dynamin binds to phosphoinositides via its PH domain (Salim et al. 1996; Zheng et al. 1996; Achiriloaie et al. 1999; Lee et al. 1999; Vallis et al. 1999), facilitating a direct interaction of dynamin with membranes. The CC domain has been characterized as a GTPase-effector domain (GED) (Sever et al. 1999), whereas the PRD has been shown to bind to multiple effector molecules (for a review of these see McNiven et al. (2000a). Substantial evidence supports the concept that dynamin is a mechanoenzyme with the ability to compress membranes into tubules and subsequently constrict these tubules into vesicles. Seminal in vitro studies have demonstrated that dynamin self-assembles to form helical ring structures (Hinshaw and Schmid 1995). Under low-salt conditions (20). Mono- or paraphyletic origin of Escavata was not confirmed in all cases, and this taxon might be polyphyletic (Arisue et al. 2005). Consequence of divergence events cannot be resolved by the methods used, but the most probable locations for the tree rout are branches leading to Parabasalia/Diplomonada clade and to Opisthokonta. Putative positions for the rout are marked with cartoon fur-trees. The taxons lacking Golgi dictiosomes, are placed in boxes limited by solid lines. Biochemical, structural or molecular evidence for presence of a functionally active Golgi complex have been obtained for representatives of all taxa except Retortamonada and Oxymonada, marked by the dashed boxes. It is presumed that the cell of the last common ancestor contained Golgi dictiosomas, and in the course of evolution Golgi complex has shifted its morphology beyond recognition at least five times (strikethrough paper stacks).

Golgi apparatus and its function in Trichomonas (Parabasalia) Flagellated parabasalids are known as simbionts and parasites of insects and mammals. Experiments with Tritrichomonas foetus and Trichomonas vaginalis infecting urogenital tracts of humans and cattle, provided the majority of

Structure and function of the Golgi organelle in parasitic protists

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data on biochemistry and cell biology of trichomonads. The representatives of this group were in the focus of attention of cell biologists mainly because they possess hydrogenosomes, the anaerobic precursors of mitochondria (Andersson and Kurland 1999), and also as the presumably most ancient group of protists (Cavalier-Smith and Chao 1996; Keeling et al. 2000). Golgi of Trichomonas spp. is large and consists of 8–12 prominent cisterns located in vicinity of the nucleus. Golgi is differentiated into four classical subcompartments (cis, intermediate, trans and trans-Golgi network). Cis-cistern is connected with the parabasal filament and located in close proximity to the ER (Benchimol et al. 2001). GA together with parabasal filament forms so called “parabasal body”, the conspicuous character of all parabasalids. The major GA function is glycosylation of adhesins, the surface proteins, which mediate the parasite adhesion to the surface of an epithelial cell, and therefore, are being responsible for pathogenesis (Benchimol et al. 2001). Three-dimensional reconstruction of serial sections and immunofluorescence demonstrate that trichomonad’s GA do not undergo disassembly during mitosis, like Golgi organelle in mammalian cells; it is not sensitive to nocodazole and other anti-tubulin agents, though tubulin components can be detected by immunofluorescence. Other than in mammalians tubulin isoforms and/or tubulin-associated proteins were assumed to interact with GA in Trichomonas (Benchimol et al. 2001). Interesting, just before mitotic division GA elongates and then each cistern divides in halves. The process starts with the surface (trans) cisterns and moves towards the deepest cis-cisterns. It was demonstrated that the Golgi complex, parabasal filament, flagella with their basal bodies, center of organization of microtubules, and axostyle replicate simultaneously with the nuclear genome in the interphase, and segregate in mitosis. Thus, trichomonad’s GA, follows the cytoskeleton elements in their duplication, segregation and migration to the daughter cells (Ribeiro et al. 2000). Similar behavior of GA in mitosis was described for Toxoplasma gondii zoites (Pelletier et al. 2002).

Organization of intracellular secretory traffic in Giardia (Diplomonada) Diplomonads, including Giardia spp, the parasites of humans and livestock, were also considered once as the ancient group of eukaryotes (Hashimoto et al. 1994, 1998; Sogin et al. 1989), a “missing link between pro- and eukaryotes” (Kabnick and Peattie 1991). Though many of their “ancient” characters were found later to be due to reduction in response to the parasitic lifestyle, diplomonads are still believed to belong to one of the earliest eukaryotic lineages (Roger 1999). Giardia lamblia is a unicellular intestinal parasite and a leading cause of diarrhea disease in humans worldwide (Adam 2001). The life cycle of Giardia consists of two distinct phases: a flagellate vegetative trophozoite and a cyst with a wall adapted to the survival in the environment. Synthesis and

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secretion of the protective cell wall are essential for transmission of Giardia infectious stages. Endomembrane system of the Giardia trophozoites is composed of the perinuclear envelope and rough ER, transitional elements, the ERGIC zone, putative tubular–vesicular elements, Golgi-like smooth perinuclear membrane stacks, small (50–80 nm) vesicles, many of which are coated, and lysosome-like peripheral granules containing acid phosphatases (Adam 2001; Elmendorf et al. 2003). Occasional flattened cisternae and clefts in EM sections of encysting cells likely represent organized smooth ER induced by weak homotypic interaction of ER membrane proteins (Snapp et al. 2003). Trophozoites lack mitochondria, peroxisomes, secretory granules, and conventional Golgi apparatus (Adam 2001; Marti et al. 2003a). During encystation Giardia trophozoites secrete a fibrillar extracellular matrix of glycans and cyst wall proteins on the cell surface. Secretory proteins contain signal sequences. The bulk of newly synthesized material is exported from the RER as a pulse during the first 5–8 h after induction, and accumulates in a set of approximately spherical encystation-specific vesicles (ESVs), the specialized Golgi-like compartments generated de novo before secretion (Hell and Marti 2004; Marti et al. 2003b). ESVs are coated with clathrin and are connected with the ER. These post-ER vesicles neither have morphological characteristics of Golgi cisternae nor sorting functions. Like conventional Golgi cisternae, ESVs are sensitive to brefeldin A and associate with two Golgi markers, ’COPI and GiYip1 (Marti et al. 2003a, 2003b). The generation of vesicular–tubular clusters, cis-Golgi compartments, and formation of ESVs seems to require the small GTPase Sar1p (Stefanic et al. 2006). There are indications for aggregation of cyst wall material in the enlarged ER cisternae, which could subsequently transform into large carrier compartments and nascent ESVs (Lanfredi-Rangel et al. 2003). This bulk transport of the cyst material is similar to the export of procollagen in mammalian cells (Mironov et al. 2003). The Golgi consists of 3–20 parallel cisterns, appeared only at the late encysting stage of trophozoites. At this period, expression of GA enzymes, such as galactosyl transferase and N-acetylgalactosamine transferase, dramatically increases (Lujan and Touz 2003). Recently several proteosome subunits and HSP70-BiP have been found In Giardia. BiP is exported to ESVs and retrieved via its C-terminal KDEL signal from ESVs (Stefanic et al. 2006). Transitional ER regions and early ESVs were co-localized with the coat protein COP II, and maturing ESVs—with COPI (Marti et al. 2003a). Two syntaxin homologues associated with intraGolgi membrane traffic, have been identified in trophozoites (Dacks and Doolittle 2002); ARF homologues have been localized in the vicinity of nuclei of the vegetative and encysting trophozoites; Rab 1 protein responsible for vesicle transport to the membrane target, was described in ER and peripheral granules. It was shown that ARF and coatomer proteins COPI and COPII were sensitive to brefeldin A (BFA). BFA treatment of both vegetative

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and encysting trophozoites blocked protein transport and induced disassembly of NBD-ceramide labeled structures (Lujan et al. 1995a, b, c; Lujan and Touz 2003). Thus, Giardia trophozoites exemplify reduction of the Golgi structure and function. However, GA emerges at the certain stage of the parasite life cycle when exocytosis of proteins and polysaccharides is vital. Noteworthy, the genome of Giardia contains almost all key proteins involved in the intracellular transport.

Secretory traffic in Entamoeba histolytica trophozoites Entamoebas are another group of parasitic protists of presumably ancient ancestry; they lack mitochondria and Golgi dictiosomes. The very presence of ER in Entamoeba histolytica was questioned (Ghosh et al. 1999). E. histolytica is a human pathogen; contamination occurs through ingestion of cysts. Ingested cysts are differentiated into trophozoites and invade intestinal epithelium and other organs, in particular liver. Entamoeba cells secrete numerous surface glycoproteins and lectins, which regulate adhesion and invasive properties. In addition, during the encystation entamoebas synthesize proteinases and other molecules directly connected with the virulence, and a chitinase. Secretory pathways of these compounds were studied in connection with surveys for the factors of pathogenicity and drug targets. Though structural organization of GA in Entamoeba is still obscure, conventional mechanisms were shown to be involved in secretory transport of most molecules. Series of biochemical data proved existence of the ER and GA in Entamoeba. Signal sequences, which direct the synthesized proteins into distinct cellular compartments of trophozoites (KDEL, N-terminal signal, etc.) appeared to be structurally similar to the signal peptides of mammalian cells (Ghosh et al. 1999). EM studies, using better preservation conditions to improve visualization of endomembranes, revealed as well the presence of ER and Golgi elements in Entamoeba spp. (Chavez-Munguia et al. 2000). Large spheres located in vicinity of the nucleus and connected by the membrane network, were identified as a part of Golgi complex basing on immunolocalization of ARF and others molecules associated with vesicle transport. Finally, Bredeston et al. (2005) experimentally proved that fraction of “large vesicles” functions as GA; ER and Golgi compartments share glycosylation functions; and that chemical structure of enzyme transporters in E.histalitica is similar to one of the higher eukaryotes. There are evidence for the presence of sugar transporters in Entamoeba cells (Bredeston et al. 2005). In E. histalitica, like in mammalian cells, GA and the transport of most molecules, is sensitive to BFA. On the other hand, traffic of several de novo synthesized proteinases induced by external stimuli, occurred to be insensitive to BFA. These data suggest that two distinct transport systems occur in E. histolytica, one similar to classical membrane protein transport and

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another – independent of BFA and inducible by external stimuli (ManningCela et al. 2003). The Entamoeba histolytica Genome Annotation Database (http://www. tigr.org) includes all major genes responsible for intracellular transport, processing and synthesis of the secretory pathway proteins.

Golgi apparatus and secretory transport systems in Apicomplexa Phyllum Apicomplexa is composed of exclusively obligate intracellular parasites. Representatives of the genera Toxoplasma, Plasmodium, Eimeria, Cryptosporidium, Sarcocystis, all belonging to the class Coccidia, are ubiquitous parasites of humans and livestock. Life cycle of all Apicomplexa includes a motile invasive stage, a zoite, which penetrates the target host cell and rapidly multiplies there. Intracelluar organization of zoites is similar in all coccidian genera. Ultrastructurally a zoite may be regarded as a simplified model of the eukaryotic cell. It contains one nucleus, one mitochondrion, one rudiment plastid (apicoplast), a compact network of endoplasmic reticulum, a single dictiosome composed of one (Plasmodium) or 3–5 (Toxoplasma, Eimeria) cisterns, and a complex of apical secretory organelles (Hager et al. 1999).

Secretory traffic in Toxoplasma gondii zoites Due to specific replication mechanisms (Hager et al. 1999; Hu et al. 2001) T. gondii zoites are strictly polarized; they contain an extensive microtubular network and multiple microtubule organizing centers (Morrissette and Sibley 2002). A centrally located nucleus divides the cell in two parts. ER is reduced and located posterior, behind the nucleus, and the perinuclear space makes an essential part of the total ER volume. Small coated vesicles budding off the apical side of the nuclear envelope move to the Golgi complex located in close vicinity. HDEL motive responsible for recycling of ER resident proteins from AG to ER, was localized by immunocytochemical and genetic markers just above the nucleus; thus it was proved that nucleus envelope served as ERGIC (ER–Golgi intermediate compartment ) (Hager et al. 1999). The presence of a single Golgi apparatus in T. gondii was noted in early EM studies and was recently confirmed by three-dimensional reconstruction of serial EM thin sections (Pelletier et al. 2002). In T. gondii, the single Golgi is located apical to the nucleus, adjacent to the centrosomes, and closely associated with a single ER exit site, which appears to be a specialized region of the nuclear envelope (Hager et al. 1999; Hartmann et al. 2006; He 2007). Each subsequent zoite division splits GA in halves. By video fluorescence microscopy and three-dimensional reconstructions of serial thin sections it was demonstrated that the new GA grows by autonomous duplication of the old one, which raises the possibility that the Golgi is a paired structure analogous to centrioles (Pelletier et al. 2002; see Chapter X).

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The GA has been shown to be essential for the formation of three specialized secretory organelles, found in zoites of all apicomlexans: rhoptries, micronemes and dense granules (Joiner and Roos 2002). Dense granules distributed uniformly all over the cytoplasm, rhoptries and micronemes located apically. Secretion from these organelles is critical for parasite invasion and establishment of intracellular infections. Morphologically and functionally dense granules resemble mature secretory granules in sectertory cells of mammals. Rhoptries display features of both endosomes and secretory granules (Bishop and Woodmane 2000; Foussard et al. 1991). T. gondii zoites were the first parasitic protists, in which tyrosine-dependent mechanisms of protein sorting was identified (Joiner et al. 1990; Mordue et al. 1999; Sibley et al. 1985). The Toxoplasma genome (http://toxoDB.org) possesses all seven of the predicted COPI subunit homologues, found in mammalian cells. BetaCOP has amino acid insertions specific to T. gondii and a C-terminal insertion that is unique to apicomplexan parasites (Smith et al. 2007). Apicoplast beta-COP changed at most, which lead to perhaps an overall alteration of function. Forty eight residues of the beta-COP are likely to be functionally important, because they exhibit subtle yet specific amino acid changes among apicomplexans, kinetoplastids, and fungi (Smith et al. 2007). Protein sorting and transport at early stages of the secretory pathway in Toxoplasma are regulated mainly by conservative mechanisms similar to those in yeast and mammalian cells: insert of hydrophobic motives in cytoplasmic domains of cargo proteins; COP2 coating; COP1-dependent recycling of resident proteins; formation of adaptin complexes, etc. (Ajioka et al. 1998; Liendo et al. 2001; Stedman et al. 2003). Signal sequences, such as the bipartite terminal NH2 domain, which mediates co-translational translocation of the plastid proteins into ER and subsequent post-translation translocation into the apicoplast, have been revealed by a series of molecular and genetic studies (Roos et al. 1999; Waller et al. 2000; Yung et al. 2001). In T. gondii protein transport via Golgi is inhibited by low temperatures, BFA and nocodazole, a ubiquitous inhibitor of microtubule assembly (Soldati et al. 1998; Stokkermans et al. 1996). Vesicles budding from distal parts of trans-cisterns are coated with clathrin (Liendo et al. 2001). Many surface antigens of T. gondii have a conservative transmembrane glycosylphosphatidyl inositol (GPI) motive which serves as a signal for building in into the plasma membrane (Karsten et al. 1998). It was demonstrated that in T. gondii Rab6 GTPase mediated the retrograde transport of proteins from dense granules to GA. T. gondii mutants with aberrant Rab6 expression demonstrated abnormal cytokinesis (multiple instead of binary fission) suggesting the role of Rab6 in coupling of mitosis and cytokinesis (Stedman et al. 2003). Figure 2 summarizes current interpretation of mechanisms underlying secretory traffic in zoites of Apicomplexa.

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Figure 2. Scheme of post-Golgi protein transport in zoites of Apicomplexa, summarizing the existing experimental data (according Joiner and Roos 2002). Protein traffic from ER to Golgi organelle and through Golgi cisternae occurs in vesicles coated with coatomer proteins COP1 and COP2. It is regulated by conservative signal sequences and by small GTPases of Rab family. Soluble proteins of dense granules (DG) are transported from Golgi to DG by default in a signal independent way, whereas transport of membrane proteins is regulated by length of the transmembrane domain. Proteins of rhoptries (ROP) and micronemes (MN) are transported from Golgi via the specialized rhoptries precursor compartment. Transmembrane proteins of ROP and MN contain signal motifs for tyrosin- and adaptin-dependent sorting. Targeting soluble MN proteins requires binding with transmembrane escort proteins. Proteins targeted to apicoplast, possess NH2-terminal domain, which directs protein transport first to AG, and then, after cleavage of the terminal peptide, into the apicoplast lumen, using transport peptide, homologous to one found in plants. It is not clear, whether all proteins synthesized in Golgi are transported through apicoplast (black dashed arrows). Direction of products processed in apicoplast is yet unknown. Solid gray lines indicate traffic routes proved by direct experiments, black dashed lines—hypothetical pathways.

Erythrocyte stage of Plasmodium and extracellular protein transport Structure and function of the Golgi in Plasmodium falciparum, the intraerythrocyte parasite that causes malaria in humans, depends on its life cycle. In human erythrocytes P. falciparum resides inside a parasitophorous vacuole (PV), and exports the synthesized proteins through the plasmalemma and the parasitophorous vacuole membrane (PVM) into erythrocyte cytoplasm. Previous results suggesting that this parasite exports its exoicytic system into the host cells (Banting et al. 1995; (Lauer et al. 1997) have been recently argued (Adisa et al. 2007; Struck et al. 2005). In the intraerythrocyte stage, the P. falciparum ER consists of the nuclear envelope with two small protrusions that develop into an extended reticular network as the parasite enlarges (Van Dooren et al. 2005). Neither structural,

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nor biochemical approaches detected presence of typical Golgi in the asexual erythrocyte stages of the plasmodium life cycle – trophozoites and the ring stage. As the ring stage develops, the ER expands from the nuclear membrane to form a reticulum throughout the cell. When parasite divides, the ER forms around the individual dividing nuclei but remains connected with the ER of other developing merozoites until very late in schizogony (Adisa et al. 2007; Van Dooren et al. 2005). In 3D, the Golgi-like compartment of P. falciparum is consisted of 1–3 tubular or flattened cisterns surrounded be vesicles of different size budding off the perinuclear space (Bannister et al. 2004). The Golgi is initially present at one or two foci that multiply as the parasite matures (Adisa et al. 2007). Many proteins involved in the exocytosis, such as: P. falciparum ERC (PfERC), PfBip (Kumar et al. 1988; La Greca et al. 1997; Van Dooren et al. 2005), the COPI protein, Pfbeta-COP (Adisa et al. 2001), PfERD2 – the retrieval receptor homologues to the signal recognition particle (Van Wye et al. 1996), and others (Gardner et al. 2002), have been identified, although the machinery for protein glycosylation is minimal. Coatomer proteins COPI, COPII, and other elements of the vesicle fusion machinery and intracellular traffic, such as Sar1p, Sec 31p, Sec 23, Pf NSF, etc. have been immunolocalized. The PfSar1p defines a network of membranes wrapped around parasite nuclei (Cooke et al. 2004; Hayashi et al. 2001). PfBet3p is largely present as a membrane- or cytoskeleton-bound pool (Adisa et al. 2007). PfGRASP colocalized with the cis Golgi marker ERD2 (Struck et al. 2005), but apart from the trans-Golgi marker PfRab6, suggesting that the cisand trans-Golgi compartments are spatially separated in P. falciparum cells (Adisa et al. 2007). Transport of most proteins is inhibited by BFA via interaction with ARF. Resident proteins of the parasitophorous vacuole (PV), as well as transit proteins directed to erythrocytes, are released into the PV lumen from the transport carriers. Electron microscopy revealed characteristic vesicles encircled by double membrane (DMV), budding off ER in the parasite cell. External membrane of vesicles fuses with the parasite plasmalemma and releases “daughter” internal vesicles which, in turn, fuse with PVM and discharge the content into the erythrocyte cytoplasm (Cooke et al. 2004; Olliaro and Castelli 1997). Application of green fluorescent protein and luciferase in combination with transfection of P. falcipatum erythrocyte stages revealed signals targeting parasite proteins to different membrane compartments (Klemba et al. 2004; Tabe et al. 1984). It was shown that translocation of soluble parasitic proteins through PVM requires energy in a form of ATP, and most likely the ATP-dependent translocators are involved (Ansorge et al. 1996). Plasmodium creates its own membrane system in the infected erythrocyte, but some important for the parasite pathogenesis proteins were detected free in erythrocyte cytoplasm, not surrounded by membrane envelopes.

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These proteins pass through PV as polypeptides lacking secondary structure (unfolded proteins). In the erythrocyte cytoplasm these proteins undergo folding; they accumulate in the specialized cell regions and interact with the erythrocyte resident proteins (Taraschi et al. 2001, 2003). Characteristic membrane compartments arise in the erythrocyte cytoplasm whilst parasite matures inside PV. In the ring stage PVM produces short finger-shaped protrusions. These protrusions bud off the PVM and give rise to small membrane carriers (Bannister et al. 2003, 2004). Elongated membrane cisterns with dense walls and electron transparent content known as Maurer’s Clefts (MCs), appear inside the erythrocyte. Role of MCs in transport of parasite antigens has been studied in connection with Pf EMP1 (P. falciparum erythrocyte memebrane protein). PfEMP 1 protein is a polymorphic integral protein, which mediates adhesive properties of infected erythrocytes. Adhesion of erythrocytes to the vascular surface causes the lethal syndromes in malaria patients. It is assumed that MCs serve as a transit depot while transporting of PfEMP1. This protein is anchored to the MC membrane by the C-terminal domain. Also it has been demonstrated that some cytosolic proteins are transported to the erythrocyte membrane in complexes adhered to the MC membrane (Cooke et al. 2004). Another membrane structure, a tubular–vesicular network (TVN) connected with PVM, was revealed in cytoplasm of infected erythrocytes. The membranes of TVN and PV lack electron dense coating typical for MCs, and they have similar antigen composition. TVN is assumed to take part in transporting the components of the erythrocyte PM from the environment to the parasite cell (Lauer et al. 1997). Many elements of vesicular transport machinery of the parasite origin have been immunolocalized in the cytoplasm of the infected erythrocytes, including proteins of coatomer complexes Sar1p, Sec 31p, Sec 23, Pf NSF, etc. (Cooke et al. 2004; Hayashi et al. 2001).

Golgi apparatus in Kinetoplastida Trypanosoma brucei, T. cruzi, and Leishmania spp. are the causative agents of devastating diseases—sleeping sickness, Chagas’ disease, and human visceral leishmaniasis correspondingly (Overath and Engstler 2004), and in these species the secretory apparatus was especially thoroughly studied. Basically, secretory apparatus in parasitic kinetoplastids, as well as in their close relatives, free-living Euglenoidea, is organized in a similar way as in mammalian cells. Golgi apparatus of kinetoplastids is composed of one dictiosome with 6–12 flat cisterns, divided in four standard compartments (cis, medial, trans, and trans-Golgi network), easily visualized by the marker proteins (Duszenko et al. 1988; McConville et al. 2002a, b; Weise et al. 2000). Pharmacological and cell biological studies suggest an intimate relationships between the Golgi and the basal bodies (Field et al. 2000; He et al. 2004). Recently it was shown that in dividing flagellates GA was formed de novo

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from ER in ERGIC compartment (Golgi export sites), which is located in trypanosomes close to the nucleus (He et al. 2004). Thus Golgi biogenesis in T. bruceri supports the de novo biogenesis model, versus the template model exemplified by Toxoplasma gondii (He 2007; Chapter X). Transport of endoproteases, protein-like phophatases, lipophosphoglucans and membrane glucose transporters in Leishmania spp.; L-mannosidase in T. cruzi, and variable surface glycoproteins (VSGs) in Trypanosoma spp. was in the focus of numerous studies in search for targets of potential vaccines (for review see Becker and Melkonian (1996)). Trypanosomatids are characterized by an extremely high velocity of endocytosis, exocytosis, sorting and concentration of the exported and recycled proteins. Every minute 107 surface glycoproteins undergo recycling from ER, the site of their synthesis, to PM and back. Ten percent of VSGs synthesized by T. brucei, contain the anchoring GPI motif, which provides building of these proteins into the membrane (versus only 0.5% of surface proteins in mammalian cells), and all of them are glycosylized (Overath and Engstler 2004). In trypanosomids N-glycosylation, insertion of GPI anchors, and other elements of protein processing usually (but not always) take place in GA and use mechanisms and enzyme systems described for higher eukaryotes (Parodi 1993; Rubotham et al. 2005). Immune co-localization of these proteins and GA markers demonstrated that T. brucei VSGs directed to the cell surface by the standard pathway, were revealed in the ER, GA, trans-Golgi network and, finally at the PM surface (Duszenko et al. 1988). Fifty-fold increase in VSGs concentration was recorded during their transportation to PM (Grunfelder et al. 2002). It remains unresolved why monezin, a ionophore of monovalent cations, which inhibits GA–PM transport in most cellular system (Mollenhauer et al. 1990), does not effect VSGs secretion, although it causes adequate alterations in GA morphology (swelling of trans-Golgi compartment), and blocks N-glycan synthesis (Bangs et al. 1986; Duszenko et al. 1988). It was suggested that additional pathway for VSGs synthesis and recycling may exist (Ferguson et al. 1986; Grunfelder et al. 2002). T. brucei Rab1, 2, 18 and X2 all localize to the Golgi (Ackers et al. 2005), but show different distribution patterns in bloodstream (BSF) versus insect (procyclic) forms of the parasite (Dhir et al. 2004; Field et al. 2000). For example, TbRab18 is expressed only in BSF (Jeffries et al. 2002). Depletion of ARL1, the Golgi-localized ARF-like protein, disrupts Golgi stacks and leads to cell death only in the BSF cells (Price et al. 2005). Developmentally regulated trafficking of the lysosomal membrane protein P67 (Alexander et al. 2002) and procyclin (Engstler and Boshart 2004), a major surface glycoprotein in procyclic cells, has also been reported. Furthermore, different effects of actin depletion on Golgi morphology have been noted in BSF, but not in procyclic cells (Garcıa-Salcedo et al. 2004). Sequencing of genes encoding some proteins of kinetoplastid secretory pathway revealed remarkable similarity to genes for analogous proteins in

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yeast and mammalian cells (Bangs et al. 1993; Descoteaux et al. 1995; Meissner et al. 2002; Ryan et al. 1993).

Structure and function of the Golgi-like organelle in Microsporidia Microsporidia are intracellular parasites that infect all taxa of animals with bilateral symmetry (Bilateria). Basing on ultrasrtuctural data which showed no mitochondria, hydrogenosomes or peroxisomes, no stacked Golgi dictyosomes, no 9 þ 2 structures, the 70S ribosomes of the prokaryote type and the ancient type of mitosis, and on early SSUrDNA-inferred phylogenies, Microsporidia were once believed to be an early diverged lineage of Eukaryotes evolved before acquisition of mitochondria (Vossbrinck et al. 1987; Vossbrinck and Woese 1986). The later research evidenced though that microsporidia are highly derived rather than primitively simple (Keeling and Slamovits 2004). Three groups of facts ruined the hypothesis of the ancient ancestry: (i) it was discovered that microsporidia possess genes for mitochondria-targeted proteins (Germot et al. 1996), and structurally recognizable mitochondria relicts, the mitosomes (Williams et al. 2002); (ii) molecular phylogenetic analyses inferred from several genes, showed that microsporidia do not evolve early in the eukaryotic evolution, but are either Fungi or their close relatives (Fischer and Palmer 2005; Keeling 2003; Keeling et al. 2000; Thomarat et al. 2004); (iii) information from completed (Encephalitozoan cuniculi (Katinka et al. 2001)), and oncoming Spraguea lophii and Nosema locustae genome projects revealed high homology with yeasts alongside with specific genome organization, which reflected reductive evolution of these organisms due to parasitic lifestyle (Fedorov and Hartman 2004; Katinka et al. 2001; Keeling 2001). Currently it is widely accepted that microsporidia (phylum Microsporidia Balbiani 1882) are highly specialized lineage of Fungi (Adl et al. 2005; Arisue et al. 2005; Keeling 2003; Richards and Cavalier-Smith 2005; Stechmann and Cavalier-Smith 2003). Mirosporidia life cycle consists of a proliferative stage (meronts and sporonts), sporogenic stage (sporoblasts), and spores, the infectious stage and the only one that can survive in the environment. Ultrastructurally meronts can be characterized by the absence of extracellular envelopes and a very few membrane structures in their cytoplasm. Sporonts are defined by appearance of the electron dense envelope outside the plasma membrane, the precursor of the exospore (the outer layer of the spore wall). GA is hardly visible in meronts and early sporonts. In the latter it appears as a conglomerate of membrane profiles of about 30 nm in diameter (Vavra and Larsson 1999) accumulated in the vicinity of ER cisterns deriving from the perinuclear space (Sokolova et al. 2001). At late proliferative stages the conglomerate grows in size, includes more elongated profiles, and

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forms a “tubular cluster”, which looses connection with the nucleus, migrates to the distal end of the cell and eventually transforms into trans-Golgi tubular network associated with polar filament proteins (PTP) pathway in prespore stages. Application of osmium impregnation techniques, which results in deposition of insoluble osmium specifically inside the lumens of cis-Golgi, suggested the portions of perinuclear space with associated elongated ER cisterns, conglomerates of 30 nm- membrane profiles and “tubular clusters” belong to the cis-Golgi compartment (Sokolova et al. 2001). At the sporoblast stage morphogenesis of the internal structures of the spore takes place. Sporoblasts possess a well-developed voluminous Golgi, portion of which function as a container for the polar filament proteins (PFP). The spore (1–25 mm in length depending on the species) is defined by appearance of the electron transparent endospore (the inner layer of the spore wall). Spores are equipped with a unique set of organelles – the extrusion apparatus, which functions when the spore content is injected into a host cell (Vavra and Larsson 1999). The central role in the process of infection belongs to the polar filament (PF). When the spore is activated to firing, PFPs undergo self- assemblage causing transformation of the filament into a tube (Keohane and Weiss 1999; Weidner 1982; Weidner and Byrd 1982). This tube

Figure 3. Sections through secretory compartment in microsporidia proliferative stages and spores. (a). Golgi organelle in meronts can be visualized as conglomerates of 20–40 nm membrane profiles (arrows) connected with the perinuclear space and ER cisterns. (b). In sporonts, conglomerates grow in size and transforms into a “tubular cluster” (arrow), which eventually transforms into trans-Golgi tubular network associated with polar filament pathway in pre-spore stages. (c). Late Golgi compartment in a developing spore is composed of tubular networks (TN1 and TN2) and membrane containers with polar filament elements (PF and APF). Polar filament protein-containing profiles are budding off the TN2 (arrow). APF – apical portion of the PF; ER – endoplasmic reticulum; N – parasite nuclei arranged in diplokaria; PF – polar filament; PP – primordial polaroplast (a part of the infection machinery, presumably ER derivate); SW – spore wall; TN 1 and TN2–trans Golgi tubular networks. Figure 3c is provided by E. Seliverstova

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delivers the sporoplasm into the target cell, like a syringe. In sporoblasts and mature spores PFPs reside in membrane-bound tubular compartment seen on cross-sections as rows of coils underlying the spore surface. PF coils are often associated with tubular networks, well seen in immature spores and sporoblasts (Fig. 3). Staining with thiamin pyrophosphatase (Takvorian and Cali 1994; YS, unpublished observations) proved the PFP-containing profiles and tubular networks to be homologous to trans-Golgi compartment of other eukaryotes. Actually the whole GA, which in spore is represented by the transGolgi compartment, is being transformed into the polar filament. Interestingly, the membrane contours surrounding PF coils remain inside the spore shell after the sporoplasm discharge; Ca2 þ influx seemingly plays a pivotal role in PF discharge which is inhibited by calcium channel antagonists and calmodulin inhibitors (Keohane and Weiss 1999; Pleshinger and Weidner 1985; Weidner 1982; Weidner and Byrd 1982), thus resembling a specialized version of exocytosis analogous to trichocyst discharge in ciliates (Plattner 1993; Plattner et al. 1991). Quick-freezing cryosubstitution and chemical fixation, followed by 3-D tomography demonstrated that the Golgi analogs of at least two microsporidia Paranosema grylli and P. locustae appeared as 300-nm networks of thin (25–40-nm in diameter), branching or varicose tubules, that displayed histochemical features of a Golgi (Beznoussenko et al. 2007). Interestingly, that Golgi-like structures of microsporidia never displayed vesicles, even when the membrane fusion was inhibited. These tubular networks were connected to the perinuclear space (Snigirevskaya et al. 2006), endoplasmic reticulum, the plasma membrane and the forming polar tube (Beznoussenko et al. 2007). They were positive for microsporidian Sec13, subunits of COP and analogs of giantin and GM130. The spore-wall and polar tube proteins were transported from the ER to the target membranes through these tubular networks, within which they underwent concentration and glycosylation (Beznoussenko et al. 2007). Importantly, the intracellular transport of secreted proteins in microsporidia occurs by a progression mechanism that does not involve the participation of vesicles generated by coat proteins I and II (Beznoussenko et al. 2007). We believe that the model of “avesicular” transport is not unique to microsporidia. Analyses of the minute (2.9 Mbp) and completely sequenced genome of the mammalian microsporidium E. cuniculi (Katinka et al. 2001) and partly sequenced genome of P. (Antonospora) locustae parasitizing insects (genome size 5.4 Mbp) (http://jbpc.mbl.edu/ Nosema/index.html) revealed all the most important protein machineries that are involved in co-translational translocation of polypeptide chains into the ER lumen, in intracellular transport, and exocytosis, well-characterized in yeast and mammalian cells; although some of these machineries lack non-essential components. E. cuniculi has a limited number of proteins involved in intracellular transport (Katinka et al. 2002), including two subunits of Sec61, the proteins

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responsible for the incorporation of polypeptide chains into the ER lumen, and seven enzymes involved in glycosylation. Among these, five are glycosyltransferases. Among six SNAREs, there are two R-SNAREs (SNC2 and synaptobrevin) and four Q-SNAREs (syntaxin 5, VAMP, Bos1 and Vti1). Importantly, Microsporidia and yeast SNAREs are very similar; and the latter, in turn, display high degree of sequence similarity to the mammalian SNAREs (Von Mollard et al. 1997). The machinery responsible for the dismantling of SNARE complexes is represented by only one protein, Sec18 (homologous to mammalian NSF), although there are no proteins that are homologous to SNAPs (Katinka et al. 2001). This means that the SNARE machinery in microsporidia might work slower than in mammalian cells. The minimal set of Rab proteins (Ypt1, Rab1b, Rab5, Ytp6 and Rab10) and the Rab-GDP dissociation inhibitor are present in E. cuniculi genome. Instead of the seven subunits of COPI typical for mammalian and plant cells, microsporidia have only six, with the epsilon-COPI missing (Katinka et al. 2001). In mammalian cells, epsilon-COPI mediates generation of COPI vesicles (Guo et al. 1994). The ARF machinery in microsporidia is also limited by only two ARFs and one exchange factor for ARF. Another important gene missing, besides epsilon-COPI, is ARF-GAP, instead E. cuniculi genome contains an ARFlike protein that can partially replace ARF-GAP (Lu et al. 2001). On the other hand, among four known subunits of the COPII machinery, only three, Sec13, Sec23, and Sec31, are found in microsporidia (Katinka et al. 2001). Although Sar1p is present, Sec12 that operates as a Sar1 exchange factor is absent. Finally, microsporidia lack clathrin, which can be explained by lack of lysosomes and absence of endocytosis. It can be concluded that microsporidia exploit the similar basic machinery for protein transport and secretion as mammalian and yeast cells marked by extreme reduction of non-functional components. Additionally, analysis of microsporidia cells provides evidence in favor of minimal role of the coated vesicles as transport carriers.

Golgi organelle on the phylogenetic tree of eukaryotes Once, dictiosomes were considered as an important argument in phylogenetic reconstructions of the Tree of Life, not less important than flagella or mitochondria. Basing on the presence or absence of the stacked Golgi cisterns the kingdom Protozoa was divided into two subkingdoms: Adictyiozoa and Dictyozoa (Cavalier-Smith 1993). It is just a curious episode of only historical value; such taxons do not exist anymore. Indeed, presence or absence of typical dictiosomes cannot serve as a reliable character to differentiate major taxons, because structure of Golgi organelle my vary not only between closely related taxons (stacks are absent in Oxymonad and present in Trymastix (Dacks and Doolittle 2001)) but within the life span of the same species (like in Giardia and Plasmodium).

Homo Saccharomyces Encephalitozoon Dictyostelium Entamoeba Mastigamoeba Arabidopsis Chlamydomonas Porphyra Phytophthora Giardia Trypanosoma Naegleria

Metazoa, Bilateria Fungi Microsporidia Eumycetozoa Entamoebidae Mastigamoebidae Embryophyta Chlorophyceae Rhodophyceae Stramenopiles Diplomonadida Kinetoplastida Heterolobosea

Snap 25b þ þ NR NR NR ? þ NR NR NR NR NR ?

Synt-axins þ þ þ þ þ ? þ þ þ þ þ þ ?

þ þ þ þ þ ? þ þ þ þ þ þ ?

Rab þ þ þ NR þ ? þ þ NR þ þ þ ?

Arf-gap þ þ þ NR þ NR þ þ þ þ þ þ þ

b-COP þ þ þ þ þ þ þ þ þ þ þ þ NR

Vps

Proteins involved in endomembrane traffica

þ þ NR þ þ þ þ þ þ þ þ þ NR

AP

þ þ þ þ þ ? þ þ NR þ þ þ ?

Sec1

a

Syntaxins – the proteins of SNARE family, encoded by genes homologous to sso, sed5, pep12 Saccharomyces); Snap25 – gene for SNAP (soluble NSFatachment protein, essential element of fusion machinery), homologous sec9 in Saccharomyces); rab–proteins of GTPase family, which participate in ER – Golgi transport; arf-gap – proteins of GTPase family, participate in intra-Golgi transport; beta-COP is a subunit of coatomer protein, participate in the retrograte ER – Golgi transport; Vps – complex of retromer proteins, encoded by Vps26 and ps35 genes, involved in protein recycling in trans-Golgi network; AP – family of adaptor proteins, involved in coating of clathrin vesicules; Sec1 – a protein, which mediates anchoring of SNARE (SNAP receptors) to the membrane. b Snap 25 was detected only in mammalian cells, higher plants and yeast and, therefore, can hardly regarded as a conservative protein. NR – not revealed; “?”– not examined.

Genus

*

Taxon

Table 1. Proteins associated with the Golgi apparatus in different groups of eukaryotes. (Based on data from genome projects (Dacks and Doolittle 2002; Dacks et al. 2003; Katinka et al. 2001; Fedorov and Hartmann 2004), and original papers on Naegleria and Mastigamoeba (Dacks et al. 2003))

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In three of six supertaxa defined by the new classification of protists (Adl et al. 2005) there are at minimum eight groups of species, in which GA does not display stack organization (Table 1, Fig. 1). However, molecular–genetic and/or biochemical data point to the presence of the organelle with Golgi functions in all lineages examined, with the exceptions of Oxymonada and Retortamonada. So far no proves has been obtained for these organisms, but in the closely related taxa GA was revealed either in hidden (Giardia) or apparent (Trimastix) form (Dacks et al. 2003). The fact that the organelle with Golgi functions presents in all contemporary groups of eukaryots suggests that a “common eukaryotic ancestor” possessed GA, as well as mitochondria, introns, splicing mechanisms, and all basic biological and molecular features of an eukaryotic cell (Dacks and Doolittle 2001). The simplicity of “early diverged lineages” most likely is a consequence of secondary loss of cell structures, for example, as result of switching to parasitism. The complexity of organization and conservatism of proteins and genes involved in secretory traffic in different eukaryotic groups (Table 1 and 2) strongly suggest that GA arose once in the evolution of the eukaryotic cell before acquisition of coat machinery.

Table 2. Some features of the Golgi in Protists Feature

Parabasalia Diplomonada Entamoebidae

Apicomplexa Kinetoplastida

Microsporidia

Segregation from the ER Periodical continuity with the ER Periodical continuity with the post-Golgi Network of smooth and varicose tubules Presence of more than two compartments Stacked disk-like cisternae COPI vesicles Clathrin vesicles Movement by actin (fragmented Golgi) Movement by microtubules H þ -ATP pump Golgi glycosidases Nucleotide transporters Matrix proteins Sar1/COPII ARF/COPI AP/clathrin SNAREs

þ þ

þ þ

þ þ

þ

þ

þ

þ

þ

þ

þ

þ

þ

þ þ þ 

þ þ þ 

   

þ þ þ þ þ þ ? þ þ

 þ þ þ  þ ? ? þ

 þ Reduced þ  Reduced Reduced  Reduced

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Evolution of the Golgi complex  spa  r Je kely Ga

Introduction The Golgi complex evolved very early during the origin of the eukaryotic cell. It is present in every eukaryote living today, including parasitic lineages formerly considered as Golgi-lacking (Bredeston et al. 2005; Dacks et al. 2003; Dacks and Doolittle 2002; Marti et al. 2003), and thus was present in the last common eukaryotic ancestor (LCEA). Its origin traces back to a protoeukaryotic stage and was a significant step in the prokaryote–eukaryote transition, one of the major transitions in the history of the biosphere ry 1995). The Golgi evolved into a compartment (Maynard Smith and Szathma with a central role in the modification and sorting of secreted and membrane proteins as well as proteins destined for other intracellular compartments (e.g. lysosomes). To understand the evolutionary history of the Golgi complex we have to analyse the history of its individual constituents. The availability of dozens of eukaryotic full genome sequences coupled to an understanding of eukaryote phylogeny allow us to reconstruct the ancestral state of the Golgi complex in the LCEA and to gain insights into the diversification of Golgi functions in different lineages.

Comparative genomics and Golgi evolution in crown eukaryotes The phylogenetic context of comparative genomic reconstructions The availability of full genome sequences from dozens of eukaryotic organisms allows us to reconstruct the gene inventory of the LCEA and the history of gene losses and duplications in descendants of the LCEA. These studies can give us an idea of the timing of events during the history of eukaryotic gene families in the eukaryotic crown group (i.e. the LCEA and all its descendants). The reconstruction is simple if we deal with genes that can be found in all eukaryotic genome sequences. In such cases we can be sure that this gene or gene family was also present in the LCEA. The inference of the presence of a gene in the LCEA can be more difficult if a gene is not present in all eukaryotic genomes. In such cases we have to know the phylogeny of eukaryotes and the position of the root of the eukaryotic tree. According to the current consensus on eukaryote phylogeny there are six major monophyletic eukaryotic superphyla (Adl et al. 2005; Keeling et al.

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Figure 1. The six major eukaryotic superphyla and the likely position of the root of the eukaryotic tree. Representative organisms for each superphylum are shown.

2005; Simpson and Roger 2004) that encompass most or all of the known eukaryotic diversity (Fig. 1). There is no widely held consensus regarding the rooting of the eukaryotic tree (Brinkmann and Philippe 2007), but the most prominent one, based on rare genomic rearrangements (Richards and Cavalier-Smith 2005; Stechmann and Cavalier-Smith 2002), places the root between unikonts (opisthokonts and amoebozoa, ancestrally with one cilium) and bikonts (plants, chromalveolates and excavates, ancestrally with two cilia). Assuming that this rooting is correct, we can infer the ancestral presence of a gene in the LCEA if we can identify its orthologues in both unikont and bikont taxa. However, if the phyletic distribution is not universal for a given gene, we always have to keep in mind that the reconstructions rest on the accepted rooting of the eukaryotic tree. There are other caveats that one has to keep in mind, such as the possibility of horizontal gene transfer (HGT) between distinct lineages. HGT is expected to have occurred for example during the evolution of several major algal groups (e.g. chromophyte algae) that derived from secondary endosymbiotic events between a photosynthetic eukaryotic alga and a non-photosynthetic eukaryotic host (Archibald et al. 2003; Deane et al. 2000; Gould et al. 2006).

Evolution of vesicle coating and tethering complexes The broad phyletic distribution of several proteins and protein complexes indispensable for Golgi function indicates their ancestral presence in the LCEA. These include all major vesicle coating complexes and their regulators as well as the Golgi tethering complexes and the Golgi fusion machinery.

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The major vesicle coating complexes, COPI, COPII, and clathrin, are all ancient, widely distributed, and were most likely present in the LCEA (Dacks and Doolittle 2001; Dacks and Field 2007; Field et al. 2007; Schledzewski et al. 1999). Besides, three subunits of the retromer complex (Vps26, Vps29, and Vps35), are also ancient (Dacks et al. 2003; Nakada-Tsukui et al. 2005; Oliviusson et al. 2006), as well as the four adaptor protein (AP) complexes (Boehm and Bonifacino 2001; Dacks and Field 2007; Field et al. 2007). In contrast, GGA proteins that contain a domain homologous to the ear domain of g-adaptin are evolutionarily younger, being only present in fungi and animals (Boehm and Bonifacino 2001; Field et al. 2007). The Golgi tethering complexes that contribute to the specificity of vesicle fusion events are also widespread and thus ancient. These include the TRAPPI complex, two subunits (Trs120p and Trs130p) of TRAPPII, the Dsl1 complex, the Vps52p/53p/54 subunits of the GARP complex, and the COG complex (Koumandou et al. 2007). Besides, the four SNARE families mediating membrane fusion events (including Golgi SNAREs), all major small GTPase families (Sar1, Arf, SRb, Rab, Ran, Ras, Rho/Rac) as well as regulators of their nucleotide cycle (GAPs and GEFs) were also present in the LCEA (Dacks and Doolittle 2001, kely 2003; Mouratou et al. 2005). 2002; Je Starting with a complex Golgi in the LCEA, an important factor that shaped the diversity of the organelle is gene loss and the simplification of trafficking pathways and compartment morphology. This trend can be observed in several independently evolving parasitic lineages. Microsporidia, a divergent parasitic fungal lineage, reduced or lost several gene families involved in membrane traffic (reduced Rab and SNARE families, no clathrin). As a consequence, they lack vesicular carriers altogether and only possess a tubular Golgi (Beznoussenko et al. 2007; Beznoussenko and Mironov 2002; Mironov et al. 2007). The intestinal parasite Giardia intestinalis also dramatically reduced its secretory apparatus. It lost several of the trafficking complexes (no COG, GARP, TRAPPII, reduced TRAPPI, and Dsl1 complexes) and lost Golgi stacking. The reduction of the trafficking machinery correlates with the extreme n et al. simplification of Golgi morphology also seen in this organism (Luja 1995; Marti et al. 2003). Quite contrary to the drastic reduction of Golgi trafficking and morphology in parasitic lineages Golgi complexity increased in some lineages, most prominently in parallel with the advent of multicellularity in animals and plants. The independent origin of multicellularity in both land plants and animals was paralleled with an expansion of the SNARE proteins of the secretory pathway (Dacks and Doolittle 2002; Sanderfoot 2007). The Rab small GTPase family also expanded from 5–20 Rab family members in protists to 25–60 members in multicellular eukaryotes independently in animals and plants (Lal et al. 2005). Interestingly, some amoeboid protists also largely extended their Rab repertoire. The excavate Trichomonas vaginalis, a sexually transmitted pathogen has 65 Rab proteins. A similar independent diversifi-

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cation of Rab families is observed in the enteric protozoan parasite Entamoeba histolytica (Saito-Nakano et al. 2005) and the soil amoeba Dictyostelium discoideum (Eichinger et al. 2005; Lal et al. 2005).

Evolution of ER and Golgi glycosylation The Golgi is the major site of protein glycosylation in eukaryotes. The known glycosyltransferases (GT) sequences from both eukaryotes and prokaryotes have been classified into 90 distinct GT families (GT1–GT90), each having up to several thousand members (Coutinho et al. 2003a) (http://afmb.cnrs-mrs.fr/ CAZY/index.html). Despite the close sequence similarity within these families, the enzymes belonging to one GT family can show up to 14 distinct experimentally demonstrated activities (Coutinho et al. 2003a). More sensitive sequence similarity searches using PSI-BLAST and structural comparisons uncovered a deep evolutionary relationship between several distinct GT families. About 75% of all GTs can be included into one of only three monophyletic GT superfamilies (Liu and Mushegian 2003) (GT-A, GT-B, GT-C), each including enzymes with dozens of distinct activities. Protein glycosylation is ubiquitous in eukaryotes (Becker and Melkonian 1996). Several GT families trace back to the LCEA and many have prokaryotic ancestors. The glycosylation machineries also diversified considerably in eukaryotes ranging from the extremes of complete loss of N-glycosylation (microsporidia) to gene family expansion into the hundreds (plants, animals). The distribution of individual Golgi GTs is very patchy and, assuming a unikont–bikont rooting, is replete with independent losses. Plants, animals, and choanoflagellates retained a largely overlapping set of GT families while fungi and several chromalveolate, excavate and amoebozoan protists lost them or modified them beyond recognition by non-iterated BLAST searches (Table 1). Mucin-type O-glycosylation that generates GalNAc-a-Ser/Thr O-glycans (Wilson 2002) is characteristic of animals (Spiro 2002). The polypeptide (pp) aGalNAc transferases (GT27 family) that initiate O-glycosylation are widespread in animals but can also be found in the choanoflagellate, Monosiga brevicollis, and in two apicomplexan animal parasites, Toxoplasma gondii (Stwora-Wojczyk et al. 2004) and Cryptosporidium sp. (Templeton et al. 2004), both belonging to the chromalveolates (Table 1). The isolated presence of aGalNAcT in these parasites suggests that they may have acquired it by HGT from their animal host (Templeton et al. 2004). Two other protists, Trypanosoma cruzi and Dictyostelium discoideum, have O-glycans that are attached as GlcNAc-a-Ser/Thr instead of GalNAc-a-Ser/Thr (Jung et al. 1998; Previato et al. 1998). These O-linked glycans are synthesized by pp aGlcNAc transferases that prefer UDP-GlcNAc over UDP-GalNAc as donor substrate (Ercan and West 2005), and that are distantly related to the animal pp aGalNAcT (West et al. 2004). It is likely that the common ancestor of these two types of enzymes traces back to the LCEA and it had GlcNAc transferase activity. GalNAc transferase activity presumably evolved later, sometime before the animal–choanoflagellate common ancestor.

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Table 1. Phyletic distribution of ER and Golgi glycosyltransferase families in five eukaryotic superphyla

The presence/absence of the various GT families was established either by reciprocal BLAST searches at the NCBI (e-value cut-off 0.02), species-specific databases (Paramecium: http://paramecium.cgm.cnrs-gif.fr/db/index; Tetrahymena: http://tigrblast.tigr. org/er-blast/index.cgi?project¼ttg; Ostreococcus, Monosiga: http://genome.jgi-psf. org) or is based on the CAZy database (http://www.cazy.org/). The upper part of the table shows the core N-glycan structure found in the respective species. The species name abbreviations used are: Gi – Giardia intestinalis; Lm – Leishmania major; Tv –Trichomonas vaginalis; Tb – Trypanosoma brucei; Pf – Plasmodium falciparum; Pt – Paramecium tetraurelia; Tg – Toxoplasma gondii; Tt – Tetrahymena thermophila; Cr – Chlamydomonas reinhardtii; At – Arabidopsis thaliana; Ol – Ostreococcus lucimarinus; Eh – Entamoeba histolytica; Dd – Dictyostelium discoideum; Ec – Encephalitozoon cuniculi; Sc – Saccharomyces cerevisiae; Hs – Homo sapiens; Nv – Nematostella vectensis; Mb – Monosiga brevicollis.

O-mannosylation forming a Man-a-Ser/Thr linkage is found in Opisthokonts (Table 1). It is initiated in the ER by protein O-mannosyltransferases (POMT1, POMT2 in human, PMT1-7 in yeast, GT39 family). In yeast O-mannosylated proteins are further processed in the Golgi by mannosyltransferases (fungal specific GT15 family, e.g. yeast MNT1) (Ernst and Prill 2001). O-mannosylation is a sorting determinant for cell surface delivery in yeast (Proszynski et al. 2004) and is also required for cell wall stability and viability (Goto 2007). Several POMT/PMT homologues can also be found in the anaerobic parasite, Trichomonas vaginalis a member of the Excavata. The isolated presence of these enzymes in this parasite may be due to HGT from its host. If it is the case, the pathway evolved in opisthokonts.

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O-fucosylation forming Fuc-a-Ser/Thr linkage on EGF repeats of various proteins has been described in animals (Harris and Spellman 1993). In Drosophila this modification occurs in the ER (Luo and Haltiwanger 2005). O-fucosyltransferases catalyzing the reaction can only be found in animals, including the cnidarian Nematostella vectensis, as well as choanoflagellates (Table 1). Proteoglycan biosynthesis is initiated by the formation of a Xyl-b-Ser bond by protein-O-xylosyltransferase (GT14). This enzyme is localised to the late ER and early Golgi compartments (Vertel et al. 1993). Xylosyltransferases can be found in animals, choanoflagellates, in Dictyostelium and plants, indicating their ancestral presence in the LCEA (Reiter 2002). One GT family (GT34) is only present in plants and fungi and contains plant Golgi xylosyltransferases (e.g. ATX1) that are involved in the biosynthesis of xyloglucan, a major hemicellulose in the cell wall of land plants (Faik et al. 2002; Reiter 2002), by forming a-1,6-linkages on a b-1,4-glycan backbone. The related fungal Golgi mannosyltransferases (yeast Mnn10p, Mnn11p) are involved in the biosynthesis of mannan, an N-glycan structure built of a backbone of about 50 mannoses and short side branches (Jungmann et al. 1999; Munro 2001). Closely related enzymes are lacking from animals. N-glycosylation with GlcNAc-b-Asn bonds is the most common form of protein modification. It is phyletically more widespread in eukaryotes and its core pathway is related to the N-glycosylation system creating Glc-b-Asn and GalNAc-b-Asn bonds in archaebacteria (Fig. 2). This core N-glycan synthesis pathway has already been present in the LCEA, as demonstrated by its wide phyletic distribution (Samuelson et al. 2005). Similarly to the vesicle trafficking complexes, gene loss and pathway simplification represent an important trend in glycan evolution. These secondary simplifications involve the loss of Golgi GT families and different degrees of shortening of the core N-glycan precursor due to the loss of the ER monosaccharyltransferases (Kelleher and Gilmore 2006; Samuelson et al. 2005). Fungi lost several ancestral GT families and rely on GT families that are either fungal specific (GT15), or are shared only by plants but not animals (GT34, GT71). This is reflected in the relative simplicity of the N-glycans of yeast that occur in two basic forms only, both generated by attaching mannose residues to the trimmed core N-glycan (Munro 2001). However, yeasts have retained the complete ER core N-glycan synthesis machinery, and the trimmed N-glycan core serves as the substrate for the Golgi mannosyltransferases. The microsporidian Encephalitozoon cuniculi is much more reduced, as it completely lacks the N-glycosylation machinery (including ALG7 and STT3) and the N-glycan processing apparatus (GTs and lectins) (Samuelson et al. 2005). Microsporidia only have O-glycosylation, and retained an ER dolichylphosphate-mannose-protein mannosyltransferase (GT39).

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Figure 2. Synthesis of the core glycan structure of N-linked glycoproteins in the ER. Synthesis of the dolichol-PP-GlcNAc2Man5 precursor starts on the cytoplasmic side of the ER. Dolichol-PPGlcNAc2Man5 is flipped into the ER lumen by Rft1. The subsequent action of 6-membrane monosaccharyltransferases (GT families 22, 57–59) makes dolichol-PP-GlcNAc2Man9Glc3. The oligosaccharide is transferred from the lipid carrier to the nascent polypeptides in an N-X-T/Smotif-dependent manner by the oligosaccharyltransferase (OST), a heterooligomeric transmembrane protein complex with one catalytic subunit, the 13-transmembrane STT3 protein (Helenius and Aebi 2004; Kelleher and Gilmore 2006).

G. intestinalis and the apicomplexan Plasmodium falciparum, the causative agent of malaria, also almost completely lost the N-glycosylation machinery. Both organisms have lost all ALG GTs except ALG7 and also lost the flippase that flips the half made precursor from the cytoplasmic to the lumenal side of the ER. They retained the STT3 subunit of the OST, indicating that the short dolichol-PP-GlcNAc2 precursor that is synthesized is transferred to proteins. Given the sequential nature of glycan synthesis, the severe truncation of the ER core glycan structure suggests that no further modification of N-glycans takes place in the Golgi of these organisms. Consistent with this, the enzymes for further N-glycan processing and the N-glycan binding lectins (ERGIC-53, calreticulin, calnexin) are missing from the genome of G. intestinalis and P. falciparum.

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At the other end of the spectrum lay vertebrates and land plants that greatly expanded some of their GT families. The core synthesis machinery in the ER is unaltered, but several glycosyltransferases evolved that decorate the core oligosaccharide in the Golgi complex. These expanded GT families in plants synthesize the complex polysaccharides of their cell walls. In Arabidopsis about 50 enzymes have been estimated to be involved in pectin synthesis alone (Coutinho et al. 2003b). Humans have 29 ppGaNTases forming the mucin-type GalNAc-a-Ser/Thr, many of which show tissue specific expression patterns (Ten Hagen et al. 2003). The elucidation of the role of these tissue specific activities in multicellular development and the generation of cell type diversity is an important challenge for the future.

Evolution of cargo sorting Some of the trafficking signals and receptors operating in the secretory pathway are ancient and their presence can be inferred in the LCEA. The KDEL receptor for retrieval of ER proteins from the Golgi is present in every eukaryote. Dileucin motifs that regulate the targeting of membrane proteins from the Golgi to the lysosomes in animals (Bonifacino and Traub 2003; Letourneur and Klausner 1992) are also conserved in Trypanosomes (Allen et al. 2007), indicating their potential ancient presence in eukaryotes. The C-terminal tyrosine-based sorting signals can be found in animals, plants and protists, indicating their ancestral nature. Some lectins recognising glycan structures in the ER and Golgi are also ancient. This includes the mannosebinding lectin ERGIC-53 found in the ER–Golgi intermediate compartment (Fiedler and Simons 1994). The retrieval of membrane proteins to the ER by KKxx-COOH motifs (Becker and Melkonian 1996; Teasdale and Jackson 1996) is also ancient. Such dileucine motif-based sorting signals are also present in the OSTcomplex in mammalian cells. It is not the catalytic OST subunit but three other subunits that are responsible for ER retrieval. One of these subunits (OST48) uses a dilysine signal formed by two lysines at positions  3 and  5 (Fu and Kreibich 2000). This motif is conserved in Plasmodium OST48 and a tandem dilysin motif can be found in the OST48 of fungi and the free-living ciliate, Tetrahymena thermophila, indicating the ancestral presence of this retrieval mechanism for TM proteins in the LCEA (Fig. 3). The sorting of acid hydrolases from the TGN to the vacuole/lysosome seems to be less conserved. The sorting receptors in animals and plants are nonhomologous, and employ different mechanisms of cargo recognition (Masclaux et al. 2005). In plants, the vacuolar targeting signals are short motives in the cargo peptide sequence. The most common is the NPIR motif usually located at the N terminus of cargo proteins that is recognised by members of the BP-80 family (Robinson et al. 2005) with no relatives in animals and fungi. In mammals, lysosomal sorting of hydrolases depends on mannose-6-phosphate residues in their oligosaccharides recognised by mannose-6-phosphate receptors (MPR). The evolution of this recognition system

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Figure 3. Conservation of the dilysine-based sorting motif in the very C-terminus of OST48. Alignments of the very C-termini are shown. GenBank IDs: Homo sapiens NP_005207, Tribolium castaneum XP_969003, Nematostella vectensis XP_001632938, Saccharomyces cerevisiae EDN62965, Schizosaccharomyces pombe NP_588153, Dictyostelium discoideum XP_629963, Entamoeba histolytica XP_648945, Trichomonas vaginalis XP_001316950, Plasmodium falciparum XP_001352067, Plasmodium chabaudi XP_740758, Paramecium tetraurelia XP_001425674, Tetrahymena thermophila XP_001013892, Cryptosporidium parvum XP_626385, Chlamydomonas reinhardtii XP_001699044, Oryza sativa NP_001059165, Arabidopsis thaliana AAK59450.

is relatively recent. It is absent from Drosophila (Dennes et al. 2005) and other insects, but its absence is due to secondary loss. MPR homologues showing sequence similarity across their entire length to both vertebrate cationdependent (CD-MPR, about 270 amino acids) and cation-independent (CI-MPR, about 2,300 amino acids) MPRs can be found in the cnidarian sea anemone, Nematostella vectensis (Putnam et al. 2007), and also in the choanoflagellate, Monosiga brevicollis. Choanoflagellates are the protist sister group to animals (King 2004; Lang et al. 2002), and the fact that they contain MPRs indicates that this sorting pathway evolved before the animal– choanoflagellate last common ancestor. Mannose-6-phosphate-dependent sorting may also function in yeast, where an MPR-like receptor has been identified with a role in lysosomal enzyme sorting. However, whether it is a bona fide cargo receptor is not known (Whyte and Munro 2001). In Dictyostelium discoideum, an amoebozoan protist, some lysosomal enzymes have N-linked oligosaccharides containing mannose-6-phosphate in a phosphomethyldiester linkage (Man-6-P-OCH3) that can also bind to mammalian MPR (Souza et al. 1997). There is, however, no clear MPR homolog in Dictyostelium. Mannose-6-phosphate-dependent lysosomal sorting therefore seems to have evolved somewhere in the Opistokhont lineage, before the animal–choanoflagellate split. The mechanisms of cytoplasmic sorting of MPR homologues is also likely conserved in Opisthokonts. MPRs in mammals are sorted by a C-terminal DXXLL-type dileucine motif that is flanked by an acidic cluster of residues and

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Figure 4. Conservation of sorting motifs in MPRs in Opisthokonta. The acidic cluster and DXXLL-type C-terminal sorting motif is partially conserved in the choanoflagellate (Monosiga brevicollis) and chidarian (Nematostella vectensis) cation-dependent (A) and cation-independent (B) MPRs. Alignments of the very C-termini are shown. GenBank IDs: CD-MPRs Xenopus laevis AAI57733, Homo sapiens BAD96512, Strongylocentrotus purpuratus XP_001201595, Takifugu rubripes AAZ08350, Nematostella vectensis EDO46637, Monosiga brevicollis EDQ87334; CI-MPRs Danio rerio NP_001034716, Homo sapiens AAF16870, Gallus gallus NP_990301, Monosiga brevicollis EDQ91968.

is recognised by GGA proteins, a class of Arf-dependent clathrin adaptors (see above). The acidic cluster and one of the leucines is also conserved in choanoflagellate, cnidarian, and sea urchin MPRs, indicating the probable conservation of the Arf, GGA, and clathrin-dependent sorting mechanism in these organisms (Fig. 4). The clathrin-dependent sorting of receptors for acid hydrolases is more conserved than the receptors themselves and their lumenal recognition mechanisms. Beside MPRs the plant BP-80-like receptors, as well as yeast Vps10p, an unrelated receptor involved in carboxypeptidase sorting, are also sorted in a clathrin-dependent manner. Clathrin coat assembly also generally involves an interaction between tyrosine or dileucine sorting motifs in the cytoplasmic tail of the receptors and adaptor complexes, as well as GGA proteins (the latter only in fungi and animals). In plants vacuolar sorting mechanisms further diversified since there are two distinct vacuolar compartments, a storage vacuole and a lytic vacuole. Both vacuole types employ a distinct targeting mechanism (Robinson et al. 2005).

Early evolution of the Golgi in the prokaryote–eukaryote transition Common origin of eukaryotic and archaebacterial N-glycosylation systems The membrane-bound machinery that synthesizes and attaches the core oligosaccharide of glycoproteins to nascent polypeptide chains in the endoplasmic reticulum (ER) of eukaryotes (see above) is related to the N-glycosyla-

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tion system found in archaebacteria (Eichler and Adams 2005). Archaebacteria have the catalytic STT3 subunit of the OST (COG1287) that catalyzes oligosaccharide transfer onto proteins in an N-X-T/S-motif-dependent manner (Igura et al. 2007). They also use a dolichol-phosphate or pyrophosphate carrier (Kelleher and Gilmore 2006; Lechner et al. 1985; Wieland et al. 1985) coupled to a large diversity of glycan structures not found in eukaryotes. Archaebacteria contain the closest prokaryotic relatives of the ALG7 glycosyltransferase (UDP-GlcNAc:dolichol-phosphate GlcNAc-1-phosphate transferase), the first enzyme in the synthesis of dolichol-PP-linked glycans (Samuelson et al. 2005). Besides, the Archaeoglobus fulgidus genome contains two gene clusters, putatively involved in protein glycosylation, containing STT3-related genes, several glycosyltransferases and transporters that are presumably involved in the flipping of the dolichol-P/PP-linked oligosaccharides (Eichler and Adams 2005). The incontestable homology between the archaebacterial and eukaryotic N-glycosylation systems and their lack from most eubacteria (except for Campylobacter sp. that acquired the pathway via HGT (Samuelson et al. 2005)) indicate that a similar system was present in the last common ancestor of archaebacteria and eukaryotes. In archaebacteria the first steps in the synthesis of the dolichol-P/PP-linked oligosaccharide occur in the cytoplasm. The half-made precursor is subsequently flipped to the extracellular face. The transfer of the oligosaccharide onto proteins occurs at the cell surface as the nascent polypeptide chain co-translationally translocates across the PM through the translocon (proteinconducting channel) (Eichler and Adams 2005). In eukaryotes oligosaccharide transfer occurs in the ER lumen that corresponds to the extracellular side of archaebacteria. This strongly suggests that the topology of eukaryotic endomembranes originated via a single inward budding step from a precursor state similar to the one in modern archaebacteria.

Origin of secretory endomembranes by invagination Secretory endomembranes probably evolved very early during the history of eukaryotes (Becker and Melkonian 1996; Cavalier-Smith 2002; Devos et al. kely 2003, 2007). The primitive endomembranes of the proto-eukary2004; Je ote must have initially been continuous with the PM and formed a network of invaginations or tubules. The sorting of membrane proteins could already have evolved at this stage as the translocon, the glycosylation machinery and other proteins were selectively enriched in these invaginations (Fig. 5). This system could have been the evolutionary precursor of the ancient and universal ER retention and retrieval systems for soluble (K/HDEL-COOH) and TM proteins (KKxx-COOH) (Becker and Melkonian 1996; Teasdale and Jackson 1996). Such mechanism could have retained and restricted the activity of the translocon and the OST complex in the primitive ER generating a membrane network with a distinct composition. ER retention of the originally monomeric STT3 could have evolved by the recruitment of additional subunits with the conserved KKxx motifs (see above).

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Figure 5. Hypothetical stages in the evolution of eukaryotic secretory endomembranes in the proto-eukaryotic (stem eukaryote) lineage.

The maintenance of this tubular reticulum was likely dependent on the developing eukaryotic cytoskeleton (Cavalier-Smith 2002). Cytoskeletal motors may also have had an active role in generating and maintaining membrane tubules as happens in modern eukaryotes. The engulfment and maintenance of the first secretory endomembranes was probably also facilitated by the common ancestor of the small GTPases Arf/Sar1/SRb, proteins with indispensable roles in ER and Golgi transport and co-translational kely 2003). A tubular network could targeting of ribosomes to the ER (Je have been able to sustain membrane transport functions even without vesicle budding (an analogous tubular endomembrane system evolved secondarily in microsporidia). Cargo sorting could have already evolved in such tubules and could have been under the control of coating complexes that bound and curved membranes without inducing vesicle budding (Mironov et al. 2007). At a later stage the evolving reticulum of secretory endomembranes topologically segregated from the PM. Before topological separation occurred the membrane re-fusion machinery had to evolve to maintain transport functions. The evolution of membrane fusion probably coincided with the origin of the SNARE family of membrane fusion proteins (Dacks and Doolittle 2002). The evolution of membrane fission evolved to separate membrane tubules, generate vesicles and to allow the equal distribution of the fragmented endomembranes during cell division. There are diverse mechanisms of membrane fission that operate in the Golgi. Most prominently it is the coat complexes that can pinch off vesicles from the growing membrane tubules. Another mechanism operating in the TGN relies on the localised synthesis of dyacylglycerol (DAG) that induces local curvature of

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the membrane. DAG can be synthesized by several pathways, most of which are ancient.

Evolution of Golgi identity In eukaryotes endomembrane compartments have distinct and well-defined contents (cargo) that have to be delivered from one membrane compartment to the other or to the extracellular surface. The membrane compartments have to be tagged to be identifiable for directional transport (via attachment of molecular motors) and for fusion with the correct target membrane. This necessitates a molecular barcoding system where a given molecular label on the cytoplasmic side corresponds to a given cargo composition inside the membrane carriers. This correlation between the outside and inside components gives every compartment a unique identity. The evolution of the eukaryotic cell is partly a history of the diversification of transport compartments with a unique identity. Most of the gene families and complexes that determine compartment identity evolved via paralogous gene duplications before the LCEA. This is evident from the homology between coat complexes (clathrin, adaptin, COPI, and most likely also COPII), syntaxins, syntaxin-binding proteins families (SM proteins), and small GTPases (but not between distinct tethering complexes) kely 2003; Je kely and (Dacks and Doolittle 2004; Devos et al. 2004, 2006; Je Arendt 2006; Koumandou et al. 2007; Schledzewski et al. 1999). The increase in the number of distinct compartments can thus be interpreted as a process of duplication events made possible by the duplication and divergence of the genes regulating membrane identity (Becker and Melkonian 1996; Cavalierkely 2003). The distinct Smith 2002; Dacks and Field 2007; Devos et al. 2004; Je identity of the ER and the Golgi complex probably evolved by the subdivision of the first secretory endomembrane domain by the evolution of distinct, but paralogous, molecular tags (Sar1–Arf, COPII–COPI). This subdivision could have occurred parallel with the increase in the complexity of glycosylation steps and the increase in cargo destinations (e.g. PM, phagosome, spore wall, cilium). Secretory membrane domains of distinct composition were established by the evolution of dynamic forward sorting of cargo into the trans compartment (towards the PM) and reverse sorting of the enzymes and fusion proteins into the cis compartment. The cargo and the corresponding molecular tags could have been segregated into cis and a trans compartments as the steady state of a dynamic process involving constant forward and reverse sorting of specific components. Besides the distribution of cargo other factors contribute to the generation of compartment identity. Distinct compartments are also labelled by their unique lipid composition. The Golgi is marked by the combined presence of PtdIns(4)P and Arf (Munro 2005). Such a system of cargo-independent identity-determination probably evolved after the cargo-dependent mechanism (but before the LCEA) and evolved sharper delineation of membrane domains and compartments.

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General conclusions

Concluding remarks, questions, and perspectives

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Concluding remarks, questions, and perspectives Alexander A. Mironov and Margit Pavelka

During these years, beginning from the discovery of the intracellular transport by Jamieson and Palade in 1967, a huge work has been accomplished in deciphering of the molecular mechanisms involved in the intracellular transport. Compared to what was known even 20 years ago, scientists have gained an extraordinary amount of information about the mechanisms of intracellular transport. A major part of the molecular mechanisms has already been unraveled. These insights have unified the several, previously independent subfields in trafficking area into a single coherent discipline – intracellular transport. We tried to summarize most of the known facts and overlap these facts with the main models of transport, and map all these functions, and possibly develop a universal idea about the mechanism of intracellular transport. The mechanisms proposed here can be generally applied to all the steps of the intracelluar transport, and are not restricted to one. This wholistic approach helps us to better comprehend the functioning and evolution of the intracellular transport systems. Now, it could be important to outline the main wholistic questions that remain within the area of intracellular transport. Within the ER-to-Golgi transport step, the main questions are the mechanism of COPII-dependent concentration of cargo, the precise role of COPII coat and COPII-dependent vesicles, mechanisms of the carrier formation at the ER exit sites. It is not clear whether the exit of cargo from the ER, maturation of the ER-to-Golgi carriers and their centralization need membrane fusion and how the COPII/COPI-system functions during the cargo exit from the ER. Important questions are related to the role of ER-to-Golgi connections and better characterization of the intermediate compartment. At the Golgi apparatus level, the main wholistic questions are the mechanisms of the function of the cis Golgi network and the Golgi exit sites; what could be the mechanism of intra-Golgi transport at steady state and whether membranes of the cargo domain undergo maturation during their progression from the cis to medial Golgi and then to the trans Golgi network. There remain many questions related to the problem of stacking. What is the role for Golgi matrix? Are there matrix receptors and if so, how do they recycle? Why are so-called Golgi matrix proteins not found between cisternae? If these matrix proteins have other functions, what glues Golgi cisternae? Why are cisternae flat and narrow? Why the number of medial cisternae is so constant in the same cell type? Why does Golgi cisternae have perforations, and why are the perforations not present in the center of cisternae?

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It is not clear what is the exact role of membrane fusion during transport, and whether membrane fusion is necessary to enter the medial Golgi, and required for the exit out of the Golgi apparatus, and whether the cargo domain changes its SNARE composition during progression through the Golgi stacks. How many fusion events occur during intra-Golgi transport? It remains to study how the SNARE machinery is organized within the secretory pathway. What is the role for Rabs? What is the role of Ca2þ flux for the transport and what is its source? The molecular mechanisms involved into Golgidependent signalling should be deciphered better. Many questions are linked with the intercisternal connections. What is the role of intercisternal connections, and how are connections formed? Can lipids diffuse along intercisternal connections, and are connections suitable for diffusion of soluble cargoes? Is there any pH gradient across the stack and if yes, how is this gradient preserved in the presence of intercisternal connections? What is the mechanism responsible for the break down of connections? The most enigmatic issue is the role of COPI-vesicles. One should prove or disprove, whether COPI-vesicles are carriers for anterograde or retrograde cargoes, or find other functions for these structures. The precise organization of the Golgi exit sites remains to be specified. What is the role for fission here? Has fission an important role for the departure of post-Golgi carriers? If yes, what are the molecular machines responsible for fission? The precursors of all carriers departing from the Golgi exit site should be identified together with the molecular mechanisms involved in these processes. The maturation of these carriers during their travelling towards the sites of their destinations needs additional examination. We hope that this book will be useful for scientists working in other fields of the cell biology, for students studying cell biology and for general readers giving them a possibility to have all necessary information in one place. Moreover, we think that some chapters of the book could be useful also for the researchers working in the field of intracellular transport. Sometimes, the overview of the field could help in the determination of the future research directions. The future stage of the development of the field could be the creation of the web site devoted to the intracellular trafficking, where scientists could have all important information already in the form of extended reviews on all proteins involved and all steps of transport. It could be done in the framework of Wikipedia or other platforms.

Contributors

Contributors in alphabetical order Arvan Peter Division of Metabolism, Endocrinology & Diabetes University of Michigan Medical Center 1150 W. Medical Center Drive Ann Arbor MI 48109 USA Banfield David Department of Biology The Hong Kong University of Science and Technology Clear Water Bay Knowloon Hong Kong, SAR of China Berger Eric G. Institute of Physiology University of Zürich Winterthurerstrasse 190 8057 Zürich Switzerland Beznoussenko Galina V. Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Bonifacino Juan S. Cell Biology and Metabolism Program National Institute of Child Health and Human Development National Institutes of Health Bethesda MD 20892 USA Cao Hong Center for Basic Research in Digestive Diseases and Department of Biochemistry and Molecular Biology Mayo Clinic College of Medicine 200 First Street Southwest Rochester MN 55905 USA Castino Roberta Department of Medical Sciences University “A. Avogadro” Via Solaroli 17 28100 Novara Italy

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Claas Christoph BIOQUANT University of Heidelberg Im Neuenheimer Feld 267 69120 Heidelberg Germany Colley Karen J. Department of Biochemistry and Molecular Genetics University of Illinois at Chicago College of Medicine 900 S. Ashland Avenue M/C 669 Chicago IL 60607 USA Daraspe Jean IBITEC-S CEA and CNRS URA2096 91191 Gif-sur-Yvette and LRA17V University Paris-Sud 11 91405 Orsay France De Matteis Maria A. Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Deborde Sylvie Margaret Dyson Vision Research Institute and Department of Cell and Developmental Biology Weill Medical College of Cornell University 1300 York Ave New York 10065 USA Derby Merran C. Department of Biochemistry and Molecular Biology and Bio21 Molecular Science and Biotechnology Institute The University of Melbourne Melbourne Victoria 3010 Australia Derganc Jure Institute of Biophysics, Faculty of Medicine University of Ljubljana Lipiceva 2 1000 Ljubljana Slovenia Egea Gustavo Department Biologia Cellular i Anatomia Patològica Facultat de Medicina Universitat de Barcelona C /Casanova, 143 08036 Barcelona Spain

Contributors Ellinger Adolf Department of Cell Biology and Ultrastructure Research Center for Anatomy and Cell Biology Medical University of Vienna Schwarzspanierstraße 17 1090 Vienna Austria Eskelinen Eeva-Liisa Department of Biological and Environmental Sciences Division of Biochemistry University of Helsinki Viikinkaari 5D, 00014 Helsinki Finland Gleeson Paul A. Department of Biochemistry and Molecular Biology and Bio21 Molecular Science and Biotechnology Institute The University of Melbourne Melbourne Victoria 3010 Australia Gravotta Diego Margaret Dyson Vision Research Institute and Department of Cell and Developmental Biology Weill Medical College of Cornell University 1300 York Ave New York 10065 USA Hauri Hans-Peter Biozentrum University of Basel Klingelbergstrasse 70 4056 Basel Switzerland Hawes Chris School of Life Sciences Oxford Brookes University Oxford, OX3 OBP UK Hong Wanjin Cancer and Developmental Cell Biology Division (CDCBD) Institute of Molecular and Cell Biology (IMCB), A*STAR 61 Biopolis Drive Singapore 138673 Singapore Hsu Victor W. Harvard Medical School Brigham and Women’s Hospital One Jimmy Fund Way, Smith 538 Boston MA 02115 USA

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Isidoro Ciro Department of Medical Sciences University “A. Avogadro” Via Solaroli 17 28100 Novara Italy

Jékely Gáspár Max Planck Institute for Developmental Biology Spemannstrase 35 72076 Tübingen Germany

Kahn Richard A. Department of Biochemistry Emory University School of Medicine 1510 Clifton Rd. Atlanta GA 30322 USA

Képès François Epigenomics Project, Genopole CNRS & University of Evry Tour Évry2 91034 ÉVRY cedex France

Koegler Eva Biozentrum University of Basel Klingelbergstrasse 70 4056 Basel Switzerland Lakkaraju Aparna Margaret Dyson Vision Research Institute and Department of Cell and Developmental Biology Weill Medical College of Cornell University 1300 York Ave New York 10065 USA Lee Stella Y. Division of Rheumatology, Immunology and Allergy Brigham and Women’s Hospital Harvard Medical School One Jimmy Fund Way, Smith 528 Boston MA 02115 USA

Contributors Lieu Zi Zhao Department of Biochemistry and Molecular Biology Bio21 Molecular Science and Biotechnology Institute The University of Melbourne Melbourne Victoria 3010 Australia Lu Lei Cancer and Developmental Cell Biology Division (CDCBD) Institute of Molecular and Cell Biology (IMCB), A*STAR 61 Biopolis Drive Singapore 138673 Singapore Luini Alberto Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 S. Maria Imbaro (Chieti) 66030 Italy Lupashin Vladimir Department of Physiology and Biophysics College of Medicine University of Arkansas for Medical Sciences Little Rock AR 72205 USA Mardones Gonzalo A. Cell Biology and Metabolism Program National Institute of Child Health and Human Development National Institutes of Health Bethesda MD 20892 USA McNiven Mark A. Center for Basic Research in Digestive Diseases and Department of Biochemistry and Molecular Biology Mayo Clinic College of Medicine 200 First Street Southwest Rochester MN 55905 USA Micaroni Massimo Laboratory of Intracellular Traffic Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy

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Mironov Alexander A. Laboratory of Intracellular Traffic Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 S. Maria Imbaro (Chieti) Italy Nakano Akihiko Department of Biological Sciences, Graduate School of Science University of Tokyo, 7-3-1 Hongo, Bunkyo-ku Tokyo 113-0033 Japan and Molecular Membrane Biology Laboratory RIKEN Discovery Research Institute, 2-1 Hirosawa, Wako Saitama 351-0198 Japan Neumüller Josef Department of Cell Biology and Ultrastructure Research Center for Anatomy and Cell Biology Medical University of Vienna Schwarzspanierstraße 17 1090 Vienna Austria Nyfeler Beat Biozentrum University of Basel Klingelbergstrasse 70 4056 Basel Switzerland Osterrieder Anne School of Life Sciences Oxford Brookes University Oxford, OX3 OBP UK Pavelka Margit Department of Cell Biology and Ultrastructure Research Center for Anatomy and Cell Biology Medical University of Vienna Schwarzspanierstrasse 17 1090 Vienna Austria Pepperkok Rainer Cell Biology Cell Biophysics Unit, EMBL Meyerhofstraße 1 69117 Heidelberg Germany

Contributors Perez-Vilar Juan Cystic Fibrosis/Pulmonary Research and Treatment Center University of North Carolina at Chapel Hill Chapel Hill NC 27599 USA Polishchuk Roman S. Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Rambourg Alain IBITEC-S CEA and CNRS URA2096 91191 Gif-sur-Yvette and LRA17V University Paris-Sud 11 91405 Orsay France Reiterer Veronika Biozentrum University of Basel Klingelbergstrasse 70 4056 Basel Switzerland Ríos Rosa M. Departamento de Señalización Celular CSIC-Centro Andaluz de Biomedicina y Medicina Regenerativa 41092 Sevilla Spain Rizzuto Rosario Department of Experimental and Diagnostic Medicine Section of General Pathology University of Ferrara Via Borsari 46 44100 Ferrara Italy Rodriguez-Boulan Enrique Margaret Dyson Vision Research Institute and Department of Cell and Developmental Biology Weill Medical College of Cornell University 1300 York Ave New York 10065 USA Rohrer Jack Institute of Physiology University of Zürich Winterthurerstrasse 190 8057 Zürich Switzerland

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Sallese Michele Unit of Genomic Approaches to Membrane Traffic Laboratory of Membrane Traffic Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Sato Ken Department of Life Sciences, Graduate School of Arts and Sciences University of Tokyo Komaba, Meguro-ku Tokyo 153-8902 Japan Scanu Tiziana Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Sokolova Yuliya Y. Tulane National Primate Research Center Division of Microbiology 18703 Three Rivers Road Covington LA 704333 USA and Laboratory of Microbiological Control All-Russian Institute for Plant Protection Sh. Podbelskogo, 3 189620 St. Petersburg – Pushkin Russia Sparkes Imogen School of Life Sciences Oxford Brookes University Oxford, OX3 OBP UK Starkuviene Vytante BIOQUANT University of Heidelberg Im Neuenheimer Feld 267 69120 Heidelberg Germany Svetina Saša Jozef Stefan Institute Jamova 39 and Institute of Biophysics, Faculty of Medicine University of Ljubljana Lipiceva 2 1000 Ljubljana Slovenia

Contributors Ungar Daniel Department of Biology University of York York, YO10 5YW UK Verbavatz Jean-Marc IBITEC-S CEA and CNRS URA2096 91191 Gif-sur-Yvette and LRA17V University Paris-Sud 11 91405 Orsay France Verissimo Fatima Cell Biology Cell Biophysics Unit, EMBL Meyerhofstraße 1 69117 Heidelberg Germany Wang Yanzhuang Department of Molecular, Cellular and Developmental Biology University of Michigan, 830 North University Avenue Ann Arbor MI 48109-1048 USA Weller Shaun Center for Basic Research in Digestive Diseases and Department of Biochemistry and Molecular Biology Mayo Clinic College of Medicine 200 First Street Southwest Rochester MN 55905 USA Wilson Cathal Department of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale 8 66030 Santa Maria Imbaro (Chieti) Italy Yang Jia-Shu Division of Rheumatology, Immunology and Allergy, Brigham and Women’s Hospital, and Department of Medicine Harvard Medical School One Jimmy Fund Way, Smith 528 Boston MA 02115 USA Zhao Weihan Department of Biochemistry and Molecular Genetics University of Illinois at Chicago College of Medicine 900 S. Ashland Avenue M/C 669 Chicago IL 60607 USA

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Index

Index A N-Acetylglucosaminyltransferases 169 Acid hydrolases 214, 388, 389, 397, 414, 682, 684 Acid phosphatase 250, 407, 409, 640, 650 Actin 72, 90, 91, 107, 224, 226, 270, 271, 277, 278–286, 288, 289, 307, 309–311, 325, 338, 383, 433, 502, 503, 536, 563, 572, 574, 589, 611–613, 657, 663 Adaptins 109, 111, 420, 436, 653, 654, 677, 687 Adaptors 23, 27, 72, 82, 87, 90, 93, 109–112, 254, 257, 283, 284, 302, 311, 360, 361, 377, 382, 388, 389, 391, 392, 395, 396, 417–421, 425, 432, 436, 437, 440, 501, 527, 563, 565, 567, 569, 570, 572, 574, 594, 596, 662, 677, 684 Adaptor protein complex1, AP-1 109, 111, 112, 215, 254, 257, 284, 311, 361, 362, 382, 389–393, 395–397, 417, 418, 420, 432, 460, 477, 501, 502, 525, 624 Adhesion 171–173, 191, 254, 255, 257, 258, 320, 326, 522, 525, 527, 529, 649, 651, 656 Antiporter 163, 193, 195 Apical 9, 18, 28, 146, 150, 212, 223, 226, 261, 276, 277, 281, 283, 285, 288, 305, 306, 322, 342, 345, 347, 359, 375, 378, 380, 382, 384, 459, 478, 489, 494, 536, 552, 563–574, 652, 653, 659 Apicomplexa 647, 652–654, 663, 678, 681

Arf GAPs 54, 108, 111, 112, 661, 662 Arf, ADP-ribosylation factor 54, 90, 106, 107–114, 252, 257, 280, 309, 361, 363, 436, 479, 501, 569, 581, 623, 662, 677, 684, 686, 687 Arf-dependent adaptors 110, 112 ARF1 54, 87–92, 95, 97–99, 106– 108, 253, 257, 259, 260, 280, 282, 285, 309, 310, 361, 362, 388, 392, 393, 432, 436, 440, 479, 480, 490, 581, 594, 614, 618, 624 ARFGAP1 89, 92, 93, 96, 98, 111 Arfrp1 107, 113, 362, 363, 439–441 Arls 3, 106, 107, 114, 228, 363, 365, 368, 440 Arl1s 107, 112–114, 228, 231, 362– 368, 439–442, 624, 657 Attached CGN 18, 21, 230, 238, 239 Avoidance of non-granule proteins 499

B BAPTA 144, 150–152, 256 BARS 89, 92–94, 233, 250, 305–307, 323, 352, 379, 572, 591, 593, 595, 596 Basolateral 18, 146, 150, 212, 261, 277, 281, 283, 285, 305, 306, 322, 347, 359, 362, 375, 377, 382–384, 459, 477, 552, 563–574 Bet3 131–137, 139, 624 Beta-glucocerebrosidase 409, 419 Bifunctional enzymes 176, 177 Blood groups 171, 172, 176, 177 Brefeldin A (BFA) 88, 110, 491, 581, 615, 641, 650

Index

C Ca2+ binding protein 145, 147–148, 256, 257 Ca2+ channel 149, 152, 254 Ca2+ pump 147, 148, 154 Calcium 3, 143, 144, 147, 149, 153, 208, 212–214, 216, 255, 257, 486, 497, 498, 660 Calmodulin 145, 148, 150–152, 660 CALNUC 147–149, 256 Camillo Golgi 7, 630 Cargo receptor 82, 211, 216, 280, 432, 493, 494, 624, 683 Carrier maturation–progression model 350 Cathepsin 212, 403–405, 407, 408, 446, 475 Caveolae 425, 520, 523 Cazy database 163, 174, 679 Cisterna maturation–progression model 10, 11, 353, 354, 347 Cis-Golgi network 21, 32, 228, 239, 321, 327, 338 Cis-, medial- and trans-Golgi 19 Cis-perforated cisternae of the intermediate compartment 21 Cisternal maturation 11, 94, 95, , 178, 352, 426, 491, 544, 554, 557, 581, 599, 617, 618, 625, 626 Classical cisterna maturation model 375 Clathrin 8, 10, 23, 26, 27, 32, 87, 90, 93, 110, 146, 154, 215, 280, 283, 301, 302, 309, 311, 325, 352, 361, 380, 384, 388–390, 392, 393, 395, 396, 414, 418, 420, 425, 426, 428, 430, 431, 436, 437, 440, 461, 462, 466, 467, 475, 477, 479, 488, 498, 501, 502, 525, 527, 563, 567–570, 574, 611, 618, 650, 653, 661, 662, 663, 677, 684, 687 Clathrin-coated vesicles (CCV) 8, 10, 23, 27, 215, 307, 388, 389, 392, 393, 395–397, 420, 467, 611

*

707

Coat protein complex II 78 Coatomer 44, 54, 87–92, 94–98, 152, 253, 257, 275, 280, 346, 348, 349, 436, 479, 480, 488, 581, 594, 595, 650, 654–656, 662 COG 3, 56, 120–126, 178, 439, 442, 443, 677 COG complex 56, 57, 120–127, 178, 443, 677 Combined models 352 Compartments along the secretory pathway 16, 30, 32, 347, 404 Compensatory endocytosis 475 Conference in Pavia 9 Constitutively secreted proteins 154, 477 Constitutive secretory (CS) pathway 485, 486, 499–501 COPI 3, 8, 10, 11, 16, 18, 21–24, 26, 29, 32, 54, 55, 58, 59, 87–99, 109, 110, 122, 123, 135, 144–146, 149, 151, 152, 200, 212, 235, 236, 253, 256, 257, 260, 261, 275, 280, 325, 326, 333, 334, 336, 338, 342–345, 347–349, 352, 353, 361, 364, 377, 383, 384, 432, 436, 440, 443, 479, 481, 489, 491, 583, 593–596, 617, 618, 624, 650, 653, 655, 661, 677, 687 COPI and endosomes 479 COPI vesicles 10, 18, 23, 24, 88, 89, 92, 94–98, 126, 180, 236, 275, 316, 317, 326, 334, 336, 338, 343–346, 348, 349, 352, 353, 491, 581, 582, 594, 595, 597, 617, 618, 626, 661, 663, 696 COPII 3, 7, 9, 16, 17, 18, 21, 23, 32, 46, 47, 59, 72, 79–83, 88, 97, 107, 112, 133, 152, 199–212, 216, 235, 325, 333–339, 353, 432, 444, 543, 613, 614, 617, 624, 626, 650, 655, 661, 663, 677, 687, 695 COPII-coated buds 16 Condensing vacuoles 486, 487, 489

708

*

Index

Correlative light-electron microscopy (CLEM) 21, 379, 393, 395 Cortactin 3, 280–282, 301–303, 306–307, 309–311 Cytoplasmic domain 53, 81, 90, 94, 97, 138, 165, 178, 179, 212, 229, 230, 432, 433, 437, 566, 571, 653 Cytoplasmic dynein 225, 273, 274, 310, 380, 382, 571 Cytoskeleton 3, 19, 30, 107, 248, 249, 270, 271, 274, 277, 279, 280, 283, 284–286, 288, 289, 326, 335, 338, 433, 440, 486, 502, 556, 572, 585, 613–615, 649, 655, 686

D Delivery of GPCs 380–383 Diffusible cargoes 11, 377 Dileucine motif 215, 567, 569, 682, 683 Diplomonads 647, 649, 648, 662, 663 Disassembly 55, 74, 132, 152, 248, 249, 257, 261, 271, 276, 334, 522, 581–583, 585, 590–599, 649, 651 Discoveries in the field of intracellular transport 7 Dynactin 225, 226, 231, 274, 275, 338, 415 Dynamin 3, 280, 284, 301–307, 309–311, 325, 352, 379, 430, 438, 475, 505, 572

E EGTA 143, 144, 150–152 Electron tomography 464, 467, 527, 529, 643 Elimination of Golgi enzymes from GPC 377 Endocytic system 7, 378, 459, 468, 477, 479 Endocytic TGN 9, 26, 459, 462, 463, 467, 468

Endocytosis 3, 8, 25, 66, 70, 107, 154, 247, 256, 280, 307, 310, 311, 393, 406, 408, 409, 417, 418, 425, 426, 428, 430, 437, 467, 468, 475, 476, 479, 480, 494, 520, 521, 570, 657, 661 Endophilin B 89, 92–94 Endoplasmic reticulum 611 Endosome 3, 9, 19, 23, 25, 26, 30, 51, 70, 71, 108, 109, 112, 114, 135, 144–146, 154, 209, 214–216, 238, 251, 252, 259, 285, 309, 351, 358– 363, 365, 367, 368, 375, 378, 382– 384, 388–397, 408, 409, 414, 415, 417–421, 425–431, 433–446, 460– 463, 466, 467, 476–481, 488, 489, 495, 501, 503, 521, 527, 556, 552, 563–566, 569, 572, 653 Endosomal compartments 144, 253, 360, 402, 460, 461, 477, 564 Endothelial cells 488, 520, 529, 551 Entamoebida 647, 662, 663 Epithelial cells 19, 21, 26, 223, 226, 272, 276, 277, 283, 287, 305, 342, 345, 380, 382, 425, 487, 494, 563–566, 569, 572, 574, 649 ER 1, 3, 7–9, 16–19, 21–23, 27, 29, 30, 32, 46, 49, 51, 54, 56–59, 70, 72, 78–83, 88, 94, 96–98, 106–108, 125, 133–139, 143–151, 153–155, 163, 165, 167, 174, 176, 178, 180, 195, 198, 200, 201, 207–214, 216, 217, 224–227, 229, 230, 232, 234, 235, 237–240, 248–252, 254–256, 258– 262, 270, 271, 273–275, 277, 279– 282, 284–286, 288, 306, 314, 321, 322, 324, 327, 333–339, 342, 344, 347, 430, 431, 435, 438, 443, 444, 459, 461–464, 467, 468, 476, 480, 481, 485–487, 489–491, 493, 506, 520, 521, 524, 525, 527, 529, 542– 545, 547, 549, 558, 565, 574, 580– 582, 585, 587–590, 595, 597, 611– 615, 617–619, 623, 626, 636–641, 643, 650–655, 657–663, 678, 679, 682, 684–687, 695

Index

ER-Golgi intermediate compartment (ERGIC) 51, 83, 108, 177, 209, 211–214, 216, 217, 388, 404, 652, 682 ER-Golgi transport 18, 46, 56, 136 ER–Golgi connections 17, 18 ER–Golgi interface 16, 18, 333, 336, 613, 615 ER-to-Golgi carriers 16, 18, 21, 23, 32, 145, 146, 151, 227, 275, 337, 344, 695 ER exit sites 9, 23, 29, 81, 106, 133, 177, 224, 239, 240, 324, 333, 480, 543, 587, 588, 613, 614, 695 ER export 21, 82, 198–200, 212, 214, 462, 615, 617 ER quality control 210, 213, 216 ERAD 210, 211, 214, 217 ERES 8, 21, 32, 81, 146, 231, 240, 270, 271, 273, 277, 333–337, 384, 480, 489, 587, 626 ERGIC 3, 51, 56, 209, 211, 212, 214, 216, 334, 336, 338, 339, 388, 650, 652, 657 Evidence against the cisterna maturation model 347 Evolution 3, 9, 73, 107, 138, 177, 535, 619, 647, 648, 658, 663, 675–678, 680, 682, 684, 686, 687, 695 Exocytosis 7, 25, 32, 144, 152, 154, 178, 256, 277, 475, 476, 480, 485– 487, 500, 502–505, 521, 529, 536, 538, 563, 651, 655, 657, 660 Exocytosis of SGs 504

F Fissioning of GPCs 378, 379 Flippases 324, 681 FRAP 225, 395, 547, 548, 550, 571, 614 Function of the Golgi apparatus 25, 270, 523, 580, 647

*

709

Function of the Golgi exit site 27 Fusion 1, 10, 24, 26, 30–32, 43, 49– 53, 55, 57, 58, 66, 72, 73, 83, 98, 130, 132–136, 138, 139, 143–146, 150, 152–154, 177, 199, 200, 233– 236, 238–240, 251, 253, 256, 259, 271, 275, 277, 318, 325, 326, 334, 336, 338, 339, 342, 344, 350–352, 358, 360, 361, 365, 367, 368, 376, 377, 378, 382–384, 392, 397, 419, 425, 432–437, 439, 441, 460, 475, 478, 481, 485, 487–489, 492, 493, 497, 498, 500, 502, 503, 505, 506, 526, 536, 547–549, 553–556, 573, 574, 582, 585, 588, 590, 591, 594–598, 612, 614, 631, 642, 655, 660, 662, 676, 677, 686, 687, 695, 696 Fusion with the PM 152, 382, 384

G Gagosome 177 Galactosylation 125, 171, 196 GDI displacement factor 67, 74 GDP–GTP exchange factors 67 Gel phase transitions 541 General principles of intracellular transport 29 GGA(s) (1-3) 27, 215, 361, 389–397, 436, 437, 440, 569, 624 GlcNAc2Man9Glc3 681 Glycosidases 1, 162, 207, 208, 347, 545, 663 Glycosylation 1, 8, 20, 22, 25, 87, 95, 124–127, 150, 153, 161, 162, 164–168, 170, 174, 176–181, 190, 191, 195–197, 200–202, 207, 234, 235, 238, 271, 350, 378, 403, 416, 443, 524, 525, 537, 538, 555, 581, 623, 630, 638, 641, 649, 651, 655, 657, 660, 661, 678, 685, 687 N-Glycosylation 8, 28, 125, 161, 167, 168, 179, 207, 403, 416, 657, 678, 680, 681, 684, 685

710

*

Index

Glycosyltransferases 27, 125, 127, 161–165, 167, 170, 171, 177, 181, 190, 196, 200–201, 207, 234, 255, 279, 358, 545, 558, 661, 678, 679, 682, 685 Goblet cells 19, 21, 26, 535, 536, 544, 545, 547, 549, 551, 556, 558 Golgi biogenesis 338–339, 584, 586, 588, 589, 599, 626, 657 Golgi cisternae 9, 11, 17, 18, 21–24, 26, 27, 29, 87, 94, 95, 120, 138, 143–146, 148, 149, 168, 170, 171, 173, 190, 200, 227, 235, 236, 238, 239, 259, 279, 305, 307, 316, 326, 327, 346–350, 358, 359, 376–378, 435, 459, 462, 466–468, 486, 487, 489, 493, 556, 580–584, 593, 594, 596–598, 623–625, 650, 654, 695 Golgi complex 3, 88, 91, 97, 132, 133, 136, 143, 155, 247–249, 251–262, 276, 278, 282, 307, 314, 315, 325, 333–338, 384, 388, 393, 395, 396, 402, 403, 409, 414, 426, 435, 440, 442, 475, 481, 488–491, 496, 500, 506, 536, 543, 545, 556, 568–569, 571, 581, 598, 648, 649, 651, 654, 675, 677, 682, 687 Golgi enzymes 3, 18–20, 53, 95, 126, 127, 148, 162, 178, 200, 223, 225, 235, 236, 344, 346–349, 353, 354, 376–378, 405, 427, 443, 478, 581, 589, 595, 626 Golgi lumen 8, 25, 145, 148, 149, 193, 194, 256, 500, 581, 590 Golgi ministacks 224, 225 Golgi ribbon 9, 24, 28, 126, 223–227, 229, 232–235, 237, 239, 240, 251, 270, 271, 276, 277, 288, 352, 369, 468, 582, 583, 585, 586, 590–595, 599, 625, 631, 632 Golgi stack 9, 11, 18, 19, 21, 22, 25, 26, 28, 95, 143, 147, 152, 175–177, 223, 224, 226, 233, 234, 236–240, 251, 270, 272, 273, 277–279, 281, 284, 285, 288, 305, 326, 333,

347–350, 354, 358, 363, 368, 375, 377, 405, 426, 463, 464, 467, 468, 487, 591, 522, 524, 581–596, 598, 599, 612–615, 617–619, 625, 630, 631, 635, 636, 638, 640, 657, 677, 696 Golgi vesicles 23, 193, 197, 302, 344, 348, 573, 591, 592, 617 Golgi-localized, gamma-ear containing, ADP ribosylation factor binding proteins, GGAs 389 Golgins 3, 22, 26, 56, 112, 113, 122, 223, 227–229, 231, 232, 235–240, 362–369, 439, 441, 442, 586, 594, 616 GPC precursors 376–379, 381 Gradients within a Golgi stack 19 Granule lumen 490, 536, 539, 541, 546–549, 551, 553, 558 Granule organization 547, 553 GRIP domain 11, 113, 228, 364, 365, 367–369, 439, 441, 442 GTPase activating proteins 67, 108, 438, 439

H Habc domain 44, 45, 59 Heterokaryon experiments 10, 345 Heterotrimeric G proteins 109, 248, 254, 256, 259–261 History 3, 9, 93, 675, 685, 687 History of models of intracellular transport 9 Hot spots on the PM 383

I Inheritance 3, 284, 288, 580, 581, 586, 587–589, 591, 598, 599 Intercisternal connections 9, 24, 29, 347, 349, 350, 352–354, 492, 696 Intermediate filaments 271, 286, 287

Index

Intracellular traffic 1, 2, 16, 143, 283, 475, 476, 535, 539, 541, 542, 553–555, 647, 655, 696, 701, 702 Intracellular transport 1–3, 7–9, 29–31, 66, 94, 130, 143, 145, 150, 154, 235, 236, 238, 240, 253, 466, 475, 477, 481, 493, 535, 537, 539, 509, 647, 651, 652, 660, 695, 696 Intra-Golgi transport 3, 8–11, 21, 28, 29, 70, 72, 88, 89, 94–96, 109, 144, 145, 150, 209, 234, 236, 238, 254, 256, 338, 342, 343, 346–348, 350–354, 435, 489, 490, 491, 493, 496, 544, 596, 662, 696 In vitro reconstitution experiments 1 IP3 145, 147, 149, 150, 250 Isolation of COPI vesicles 349

K KDEL receptor 96, 251, 259, 404, 682 Kinetoplastida 647, 656, 662, 663 Kiss-and-run model 25, 350–352, 354, 376, 477, 481, 491, 496, 506

L Labeled lipid analogues 521 Lateral diffusion model 31, 350, 375 Lateral segregation 326 LCEA (last common eukaryotic ancestor) 675–678, 680, 682, 687 Lectins 3, 7, 124, 196, 285, 461, 558, 651, 680–682 Live-cell imaging 379, 393, 395, 396, 614 Longin 44–46, 58 Lysosome 8, 32, 97, 146, 153, 167, 214–216, 226, 238, 250, 28, 351, 375, 383, 388, 389, 402, 406, 408, 409, 414, 415, 417–421, 425, 426– 428, 431, 436, 440, 444, 446, 460, 477, 480, 488, 495, 501, 504, 527,

*

711

536, 552, 564, 644, 650, 661, 675, 682 Lysosomal membrane protein 298, 414–420, 499, 501, 657 Lysosomal sorting 209, 682, 683

M Macromolecules 1, 535–537, 539, 541–545, 546, 556, 631, 648 Macular corneal dystrophy 173 Mannose 6-P receptor 402–404 Mannose 6-phosphate receptors, MPRs 211, 214, 216, 309, 360, 367, 382, 388–393, 395–397, 402, 406–409, 414, 421, 427, 429, 460, 466, 682 Mannosidases 19, 21, 126, 162, 167–170, 181, 207, 209–211, 216, 217, 287, 348, 657 Matrix proteins 3, 22, 72, 230, 236, 349, 521, 551, 585, 589, 590, 591, 595, 599, 613, 615, 616, 619, 663, 695 Membrane 425–427, 429–433, 435–437, 439–441, 444–446 Membrane attachment of Rabs 66 Membrane curvature 92, 108, 109, 310, 315–318, 321, 322, 324, 326, 445, 572 Membrane flow 163, 346, 427, 476, 559, 635, 642–644 Membrane input from the endoplasmic reticulum 227 Membrane shape 315, 318–321, 323, 327 Membrane spontaneous curvature 318–320, 322, 323, 327 Membrane traffic 66, 71, 73, 93, 106–110, 112, 130, 134, 138, 139, 144, 226, 234, 247–249, 251–262, 271, 275, 276, 283, 286, 288, 305, 363, 364, 366–368, 383, 406, 425, 433, 440, 444, 501, 562, 612, 623, 650, 662, 677, 704

712

*

Index

Microsporidia 253, 647, 658–663, 677, 678, 680, 686 Microtubules 18, 22, 224, 270, 271, 276, 283, 325, 336, 337, 338, 375, 379, 381, 383, 395, 415, 480, 302, 574, 585, 586, 612, 625, 649, 663 Microtubule-organizing centre 223, 241, 338 Mint(s) (1-3) 109 Mitosis 25, 112, 230, 231, 236, 237, 239, 248, 249, 334, 335, 581, 582, 584, 585, 588–596, 598, 599, 615, 649, 653, 658 Models of intra-Golgi transport 11, 29, 236, 238, 342, 343, 352, 353, 491 Molecular switches 67, 121, 274 Molecular tools 2, 580 Morphodynamics 630 Morphology of the Golgi apparatus 19 Motor proteins 26, 72, 73, 235, 270, 273, 274, 358, 360, 382, 432 Movement 10, 22, 25, 224, 226, 273–275, 282, 283, 285, 286, 334, 338, 342, 351, 379, 380–382, 384, 396, 415, 432, 478, 489, 502, 504, 540, 580, 611–614, 619, 663 Mucins 535–539, 541–547, 551–556, 558–560 Mucin gels 541, 551, 552, 556, 558, 559 Mucolipidosis 168, 408 Multimerization/condensation 486, 493, 499 Multi-spanning membrane protein 200 Myosin 72, 226, 227, 280, 282–284, 286, 288, 289, 310, 380, 382, 574, 589, 612

N Non-diffusible cargoes 11 Nucleotide sugar transporter 192, 193, 195

190,

O OST (oligosaccharyltransferase) 167, 207, 216, 681–683, 685

P Palmitoylation 55, 58 Parasitic proteists 647, 651, 653, 655 Phospholipase A2 250, 323 Phospholipase C 250, 253, 323 Phospholipase D 250, 251 Phosphorylation 22, 55, 57, 90, 171, 180, 181, 190, 214, 215, 229, 239, 253, 255, 311, 334, 390, 391–393, 403, 405, 437, 440, 466, 581, 583, 584, 592–596, 599 Phosphotransferase 168, 214, 403, 405, 408 Phyletic distribution 676, 679, 680 Physiology of traffic 1 Pichia pastoris 80, 223, 333, 587, 588, 624, 635, 642, 643 Plant 2, 3, 7, 18, 19, 120, 124, 177, 208, 211, 224, 237, 270, 271, 285, 286, 288, 333, 334, 338, 360, 364, 426, 429, 459, 461, 485, 580, 589, 599, 611, 612, 614–619, 631, 632, 635, 641, 654, 661, 662, 676, 677, 678, 680, 682, 684 Plant and bacterial toxins 459 Pleiomorphic transport carriers (PTCs) 388, 389, 393–397 Plus-end motor kinesin 226 Polarity 3, 192, 223, 277, 285, 286, 554, 563, 565, 567, 571, 574, 584, 599 Polysialic acid 173 Processing glycosidases 162 Progression model 9–11, 29, 31, 343, 346, 347, 350, 353, 354, 554 Prosaposin 407, 409 Protein folding 1, 96, 207–210, 216, 219

Index

Protein kinase A 57, 259 Protein kinase C 252, 260, 379 Protein traffic 211, 498, 654

R Rabs 3, 66–74, 121, 122, 228, 361, 435, 438, 503, 504, 574, 696 Rab cycle 73 Rab effectors 69, 71, 361 Rab escort protein 66, 74 Rabs family of proteins 66 Ras 25, 66–69, 79, 106, 254, 255, 257, 258, 438, 677 Reassembly 236, 288, 309, 339, 581–583, 590, 591, 594–599 Regulatory secretory proteins (RSPs) 9, 383, 485–487, 489–499, 502 Retrograde recycling routes 460 Retrograde routes 459, 460, 461, 468 Retrograde transport 26, 32, 94– 98, 251, 258, 259, 333, 336, 347, 351, 358, 360, 362, 364, 366–369, 396, 397, 405, 426, 427, 428, 436, 441, 443, 459, 460, 466–469, 501, 581, 595, 653 Role of cargoes 28 Role of COPI for endosome function 481 Role of COPI vesicles 24, 88, 94, 346, 353, 696 RSP polymerization 496

S Saccharomyces cerevisiae 22, 66, 79, 191, 223, 270, 233, 363, 430, 475, 580, 584, 588, 623, 632, 679, 683 Sar1 79–83, 106, 107, 112, 138, 337, 432, 613–615, 624, 650, 655, 656, 661, 663, 677, 686, 687 Sec12 79–82, 342, 614, 624, 661

*

713

Sec13/31 79–83, 337 SEC7 22, 91, 342, 623, 624, 632, 635–638, 640, 642 SEC14 623, 624, 638–640, 642 Sec16 80, 81, 334, 624, 626 Sec23/24 79–83 SERCA 147–149, 255 Secretory pathway 3, 16, 21, 23, 25, 30, 32, 59, 66, 70, 72, 127, 133, 143, 147, 154, 161, 163, 173, 177, 199, 207, 211–213, 254, 270, 283, 284, 306, 314, 315, 325, 333, 339, 346, 347, 363, 376, 402, 404, 405, 406, 409, 431, 436, 477, 485, 486, 499, 500, 501, 503, 504, 506, 524, 535, 536, 546, 558, 559, 632–636, 638, 643, 644, 651–653, 657, 677, 682, 696 Shiga toxin 68, 144, 155, 256, 362, 367, 368, 426, 428–430, 441, 442, 463, 466 Sialylation 20, 125, 171, 191, 200, 544, 549 Signalling platform 247–249 Small G proteins 360–363, 366, 369 SNARE 3, 7–9, 23, 24, 31, 43–59, 66, 72, 73, 98, 121, 122, 126, 130, 134– 139, 143, 151–153, 234, 235, 237, 239, 240, 259, 275, 326, 338, 339, 342, 346–348, 350, 351, 353, 354, 360, 361, 367, 368, 376, 378, 384, 432, 433, 435–439, 441–444, 460, 478, 487, 488, 503, 504, 529, 573, 574, 595–597, 615, 618, 623, 624, 626, 661–663, 677, 686, 696 SNARE-motif 44, 47, 50, 53, 56, 58, 59 i-SNAREs 53 Qa-SNAREs 43, 44, 47, 49, 52, 56 Qb-SNAREs 24, 44, 47 Qc-SNAREs 44, 47, 49 t-SNAREs 47, 50, 360, 460 v-SNAREs 43, 47, 49, 443, 488 Sorting 1, 2, 8, 25, 28, 47, 53, 55, 59, 67, 87, 90, 92, 95, 96, 99, 110–112,

714

*

Index

126, 153, 167, 177, 199, 207, 209, 211, 212, 214, 216, 217, 234, 238, 247, 260, 281, 283, 284, 288, 311, 321, 326, 336, 351, 359, 360, 375, 382, 388, 389, 390–392, 396, 397, 402, 406–408, 418, 420, 425, 427, 428, 431, 436, 437, 444, 445, 460, 461, 466, 481, 488, 493–499, 501, 502, 505, 506, 520, 521, 525, 535, 552, 553, 556, 558, 563–567, 569, 571, 572, 574, 598, 619, 650, 653, 654, 657, 675, 679, 682–687 Sorting of RSPs 493 SPCA 147–149, 151, 254, 255, 256 Spectrin 225, 253, 279, 286, 310, 325 Spondyloepiphyseal dysplasia 137, 173 Stacking 22, 229–231, 236, 237, 580, 582–585, 592–594, 596, 598, 625, 677, 695 Stacks in G2 cells 225 Stem domain 165 Stimulation of exocytosis 475 Stimulation of endocytosis 475 Storage/secretory granules (SGs) 22, 26, 351, 384, 485–491, 493, 495, 497–500, 502–505 Structure of GPCs 376, 478 Structure of Rabs 68 Structure of the Golgi exit site 21, 25, 384 Sulfotransferase 173, 174, 190

T Tethering 31, 44, 50, 55–57, 72, 83, 96, 120–122, 126, 130, 134–139, 227, 228, 235, 236, 338, 342, 344, 346, 352, 358, 360, 363, 368, 382, 432, 433, 435, 436, 439, 441–444, 522, 574, 580, 582, 586, 594–598, 612, 614, 625, 676, 677, 687 The “sorting by retention” model 499

Tn-syndrome 175, 176 Topography of glycosylation enzymes 166 Traffic 163, 164, 170, 171, 177–180, 215, 359, 459, 564, 565, 569, 571, 572, 574 Transbilayer area asymmetry 317, 327 TRANSCET 26, 27, 29, 32 Trans-Golgi 19, 26–28, 52, 53, 112, 136, 147, 150, 168, 170, 171, 173, 177–180, 209, 213, 215, 217, 228, 232, 238, 240, 241, 254, 280, 325, 347, 377, 404, 405, 433, 435, 441, 444, 460, 462–464, 467, 468, 486, 490, 502, 521, 523, 525, 527, 529, 543, 545, 546, 556, 616, 618, 630, 655, 657, 659, 660, 662 Trans-Golgi Network (TGN) 3, 18, 25, 26, 32, 71, 108, 144, 146, 155, 215, 228, 241, 251, 275, 280, 301, 302, 347, 354, 358, 388, 402, 405, 419, 420, 426, 429, 459, 460, 461, 464, 467, 475, 476, 481, 486, 487, 506, 529, 552, 564, 594, 618, 640, 649, 656, 657, 662 Transitional ER 81, 333, 588, 626, 650 Transport 1–3, 7–11, 17, 18, 21, 25, 26, 28–32, 43, 46, 47, 49–52, 55–58, 66, 68, 70–72, 74, 78, 81, 87–99, 109, 120, 122, 126, 130, 132–139, 143–150, 152–154, 164, 167, 168, 176–178, 181, 190–201, 207, 209, 211–217, 224, 225, 229–231, 234– 238, 240, 251, 252, 254, 255–261, 271, 273–275, 279–288, 302, 305– 307, 309–311, 324, 333–339, 342, 343, 346–354, 358–360, 362–364, 366–369, 375–377, 379, 381–384, 388, 389, 393, 394, 396, 397, 402, 403, 405, 406, 408, 409, 414, 417– 420, 425–428, 430–446, 459–461, 463, 466–469, 475–481, 485, 487, 489–491, 493, 494, 496, 500–503,

Index

505, 506, 520, 527, 535, 537, 539, 544, 554, 555, 558, 565, 565–567, 569, 573, 580, 581, 583, 585, 590, 595, 596, 599, 611, 614, 615, 617, 618, 625, 631, 632, 635, 641, 642, 647, 650–657, 660–662, 686, 687, 695, 696 Transport models 11, 32, 236, 334, 544 Transport of secretory proteins through endosomes 476 Transmembrane domain 1, 43, 50, 53, 56–58, 161, 165, 178, 228, 233, 234, 256, 260, 324, 327, 338, 390, 416, 418, 420, 571, 613, 654 TRAPP 3, 44, 130–139, 235, 436, 439, 442–444, 624, 677 Tubules 16–18, 21, 24–27, 30, 82, 93, 95, 154, 224–226, 230, 234, 251, 273, 282, 286, 301, 302, 305, 307, 309, 323, 324, 337, 338, 343, 348, 349, 352, 353, 359, 360, 367, 375, 377, 379, 380, 384, 431, 461, 478– 480, 487, 492, 526, 528, 554, 572, 593, 595, 611–613, 618, 624, 631– 643, 660, 663, 685, 686 Two transport steps during Golgi-toPM transport 478

U Ubiquitination 58, 594, 597, 598 Uncovering enzyme 168, 214, 404, 405 UDPGlcNAc: lysosomal enzyme-1phosphotransferase 168 UDPGalNAc: polypeptide GalNAc-Ts 174

*

715

144–146, 151–154, 177, 180, 191, 193, 195, 197, 199, 210, 211, 212, 215, 216, 233–236, 252, 273, 275, 281, 283–286, 301, 302, 304, 305, 307, 311, 314, 316, 317, 319, 325, 326, 333, 334–339, 342–354, 359, 360, 375–378, 380, 388, 389, 392, 395, 405, 408, 414, 148, 420, 425, 426, 431–433, 435, 439, 441–444, 461, 467, 475, 487, 491, 501–503, 505, 543, 552, 567, 572–574, 580– 583, 585, 586, 588, 590–592, 594, 595, 597, 598, 611, 614, 617, 618, 626, 629–632, 635, 642, 643, 650– 655, 660, 661, 663, 686, 695, 696 Vesicle fusion 55, 72, 132, 133, 139, 574, 594, 655, 677 Vesicle tethering 44, 55, 56, 121, 126, 134, 135, 138, 586, 596, 625 Vesicle scission 82, 307 Vesicle transport 74, 274, 650, 651 Vesicular model 8, 10, 30–32, 235, 236, 238, 342, 343, 344, 354, 375, 376, 544, 631 Vesicular–tubular clusters 16, 143, 650 Visualization of GPC 380 Von Willebrand factor 520, 524, 529, 551 VTCs 16, 143, 334, 336–338, 491

W Weibel-Palade bodies 488, 524, 527, 529, 551 Wheat germ agglutinin 461, 467, 529

Y V Vesicles 2, 8–11, 16, 18, 19, 21, 23, 24, 27, 29–32, 43, 46, 49, 52, 53, 56– 59, 67, 71, 72, 78–83, 87, 88, 89, 92– 98, 120, 122, 126, 127, 133–139,

Yeast 2, 3, 8, 10, 18, 19, 22, 28, 43, 45–47, 49, 51–58, 67, 72, 73, 78, 79, 81, 83, 91, 94, 97, 98, 107, 113, 120– 124, 126, 127, 130–139, 145, 148, 152, 177, 191–193, 195, 197, 199,

716

*

Index

209–211, 224, 228, 237, 253, 256, 270, 283, 284, 288, 302, 334, 336, 342, 346, 348, 363–365, 402, 427, 430, 431, 434, 435–446, 477, 479, 480, 485, 498, 502, 567, 574, 580, 588, 597, 619, 623–626, 629, 630,

632–637, 640–644, 647, 653, 658, 660–662, 679, 680, 683, 684

Z Zone of exclusion

19