Tobacco BY-2 Cells: From Cellular Dynamics to Omics (Biotechnology in Agriculture and Forestry)

Biotechnology in Agriculture and Forestry Edited by T. Nagata (Managing Editor) H. Lörz J. M. Widholm

Biotechnology in Agriculture and Forestry Volumes already published and in preparation are listed at the end of this book.

Biotechnology in Agriculture and Forestry 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics Edited by T. Nagata, K. Matsuoka, and D. Inzé

With 102 Figures, 16 in Color, and 7 Tables

123

Series Editors Professor Dr. Toshiyuki Nagata University of Tokyo Graduate School of Science Department of Biological Sciences 7-3-1 Hongo, Bunkyo-ku Tokyo 113-0033, Japan Professor Dr. Horst Lörz Universität Hamburg Institut für Allgemeine Botanik Angewandte Molekularbiologie der Pflanzen II Ohnhorststraße 18 22609 Hamburg, Germany

Professor Dr. Jack M. Widholm University of Illinois 285A E.R. Madigan Laboratory Department of Crop Sciences 1201 W. Gregory Urbana, IL 61801, USA

Volume Editors Professor Dr. Toshiyuki Nagata (address see above) Dr. Ken Matsuoka RIKEN Plant Science Center 1-7-22 Suehirocho, Tsurumi-ku Yokohama 230-0045, Japan

Professor Dr. Dirk Inzé VIB/Ghent University Department of Plant Systems Biology Technologiepark 927 9052 Ghent, Belgium

Library of Congress Control Number: 2006924827

ISSN 0934-943X ISBN-10 3-540-32673-1 Springer Berlin Heidelberg New York ISBN-13 978-3-540-32673-1 Springer Berlin Heidelberg New York This work is subject to copyright. All rights reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable for prosecution under the German Copyright Law. Springer is a part of Springer Science + Business Media springer.com © Springer-Verlag Berlin Heidelberg 2006 Printed in Germany The use of general descriptive names, 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. Editor: Dr. Dieter Czeschlik, Heidelberg, Germany Desk Editor: Dr. Andrea Schlitzberger, Heidelberg, Germany Cover design: design&production GmbH, Heidelberg, Germany Typesetting and production: LE-TEX Jelonek, Schmidt & Vöckler GbR, Leipzig, Germany Printed on acid-free paper 31/3100 5 4 3 2 1 0

We dedicate this volume to the late Professor Jeff Schell for his encouragement of the development of this subject.

Preface

It is our utmost pleasure to present a new book on tobacco BY-2 cells, Tobacco BY-2 Cells: From Cellular Dynamics to Omics, as the 58th volume in the book series Biotechnology in Agriculture and Forestry (BAF). It represents an extension of the previous book Tobacco BY-2 Cells, vol. 53 of the BAF. Moreover, the content is rather different from the latter and includes new topics, gleaned from the First International Symposium on Tobacco BY-2 Cells held at the Plant Science Center of the RIKEN, Yokohama, organized by Nagata, Matsuoka and Inze, in September 2004. To this symposium came more than 200 people from different parts of the world to discuss issues. Although most of the contributors to the previous volume of Tobacco BY-2 Cells gave talks on their subjects, there were many other speakers who presented new topics and approaches. So we enjoyed the symposium very much. Thus we decided to compile a new volume on tobacco BY-2 cells which includes these new topics. In addition, towards the end of the symposium, our common understanding was that the tobacco BY-2 cell system is still important in plant biology, in particular for studying the dynamic features of plant cells. We hope this volume is useful for plant biologists. Contents of the book are as follows: in Chapters I.1–I.6, various aspects of the cell cycle and cellular dynamics using BY-2 cells are described. In Chapters II.1–II.3, physiological and developmental aspects of BY-2 cells are discussed. In Chapters III.1–III.3, recent developments in the knowledge of intracellular traffic of BY-2 cells are described. In Chapters IV.1–IV.3, BY-2 cells as a host for infectious diseases are discussed. Chapter V.I describes the dynamic features of mitochondrial fusion and division, while Chapter V.2 discusses how BY-2 cells are useful also for elucidating the biosynthesis of isoprenoids. In Chapters VI.1–VI.3, recent developments in the omics of BY-2 cells are described. Finally, Chapters VII.1 and VII.2 include two technical advances in handling BY-2 cells. Tokyo, Yokohama, Ghent, May 2006

Toshiyuki Nagata, Ken Matsuoka, and Dirk Inzé

Contents

Section I Cell Cycle and Cellular Dynamics I.1

1 2 3 4 5 6 7 8 I.2

1 2 3 4 I.3

1 2 3 4

Novel Approaches for Cell Cycle Analysis in BY-2 . . . . . . . . . . . . . . O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Modified alc-Inducible System for Transgene Expression in BY-2 Use of the alc System to Define the Time of Action of Induced Protein An Example of Functional Analysis Using the AlcR-GR Gene Switch Transient Protein Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emerging Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Composition of Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dynamics and Structure of the Preprophase Band and the Phragmoplast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Geelen and D. Inzé Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Preprophase Band (PPB) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Phragmoplast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Formation of Cortical Microtubules in a Cell-Free System Prepared from Plasma Membrane Ghosts and a Cytosolic Extract of BY-2 Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T. Murata and M. Hasebe Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Microtubules Nucleate as Branches on Existing Microtubules in the Cortical Arrays of Plant Cells . . . . . . . . . . . . . . . . . . . . . . . . Analysis of the Molecular Mechanisms of Microtubule Nucleation in a BY-2 Cell-Free System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3

3 3 5 6 9 13 16 19 19 23 23 24 31 35 36

41 41 41 44 47 47

X

I.4 1 2 3 4 5 I.5 1 2 3 4 I.6

1 2 3 4 5 6

Contents

Chromosome Dynamics in Tobacco BY-2 Cultured Cells . . . . . . . . . S. Matsunaga, N. Ohmido, and K. Fukui Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dynamic Analysis of Condensin Complexes . . . . . . . . . . . . . . . . . . Dynamic Analysis of Heterochromatic Protein 1 . . . . . . . . . . . . . . . Dynamic Analysis of Aurora Kinases . . . . . . . . . . . . . . . . . . . . . . . Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ion Channels Meet Cell Cycle Control . . . . . . . . . . . . . . . . . . . . . . . R. Hedrich and D. Becker Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell Elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The NACK-PQR MAP Kinase Cascade Controls Plant Cytokinesis . M. Sasabe, Y. Takahashi, T. Soyano, H. Tanaka, K. Kousetsu, T. Suzuki, and Y. Machida Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Plant Cytokinesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The MAPK Cascade Involved in Cytokinesis . . . . . . . . . . . . . . . . . . Regulation of the NACK-PQR Pathway in the Cell Cycle Machinery Downstream Factors of the NACK-PQR Pathway . . . . . . . . . . . . . . Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

51 51 52 53 57 60 61 65 65 66 66 70 75 79

79 81 82 86 88 91 92

Section II Physiological and Developmental Aspects II.1

1 2 3 4 5 6

Characterization of a Cell Division Factor from Auxin-Autotrophic 2B-13 Cells Derived from the Tobacco BY-2 Cell Line . . . . . . . . . . . . . . . . . . . . . . . . . . . T. Shimizu, K. Eguchi, I. Nishida, K. Laukens, E. Witters, and T. Nagata Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Characterization of Cell Division-Inducing Activity in the Culture Filtrates of 2B-13 Cells . . . . . . . . . . . . . . . . . . . . . . . Purification of the CDF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Searches for CDFs in the Culture Filtrates of Tobacco BY-2 Cells . . . Concluding Remarks and Future Perspectives . . . . . . . . . . . . . . . . . Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

97

97 98 100 102 103 104 106

Contents

II.2 1 2 3 4 5 II.3

1 2 3 4

The BY-2 Cell Line as a Tool to Study Auxin Transport . . . . . . . . . . J. Petrášek and E. Zažímalová Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Present State of the Art of Cell-to-Cell Transport of Auxins . . . . . . . Auxin Transport Studies in Planta . . . . . . . . . . . . . . . . . . . . . . . . . . Auxin Transport Studies in Simplified Models . . . . . . . . . . . . . . . . Concluding Remarks and Future Prospects . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tobacco BY-2 Cells as a Model for Differentiation in Heterotrophic Plant Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Y. Miyazawa and A. Sakai Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hormonal Factors Affecting Starch-Storing Cell Differentiation in BY-2 Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulations of Gene Expressions Required for Differentiation and Dedifferentiation of Starch-Storing Cells . . . . . . . . . . . . . . . . . Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

XI

107 107 107 109 110 114 115 119 119 120 124 127 130

Section III Intracellular Traffic III.1 Imaging the Early Secretory Pathway in BY-2 Cells . . . . . . . . . . . . . D.G. Robinson and C. Ritzenthaler 1 The Early Secretory Pathway in Plants: A Brief Introduction . . . . . . 2 General Description of the BY-2 Endomembrane System . . . . . . . . 3 The Golgi Apparatus: Structure, Motility and Behaviour During Mitosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 The Endoplasmic Reticulum: Distribution and ER-Export Sites (ERES) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 BY-2 Cells: A Model System for Studying the Action of BFA . . . . . . 6 Appendix: Standard Fixation Protocols for Electron Microscopy and Immunostaining . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III.2 Molecular Study of Prevacuolar Compartments in Transgenic Tobacco BY-2 Cells . . . . . . . . . . . . . . . . . . . . . . . . . . S.W. Lo and L. Jiang 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

135 135 136 137 140 143 147 148 153 153 155 163 164 165

XII

Contents

III.3 Autophagy and Non-Classical Vacuolar Targeting in Tobacco BY-2 Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . K. Toyooka and K. Matsuoka 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Autophagy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Protein Aggregation and Degradation . . . . . . . . . . . . . . . . . . . . . . . 4 Direct Transport from the ER to the Vacuole . . . . . . . . . . . . . . . . . . 5 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

167 167 168 174 175 177 178

Section IV As a Host for Infectious Diseases IV.1 In Vitro Translation and Replication of Tobamovirus RNA in a Cell-Free Extract of Evacuolated Tobacco BY-2 Protoplasts . . . . K. Ishibashi, K. Komoda, and M. Ishikawa 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Commercial Wheat Germ Extract and Rabbit Reticulocyte Lysate Can Support the Translation but not the Replication of Tobamovirus RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Establishment of an in Vitro Translation–Replication System for Plant Viral RNA with an Extract of Evacuolated BY-2 Protoplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Conclusion and Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV.2 Using BY-2 Cells to Investigate Agrobacterium–Plant Interactions . S.B. Gelvin 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 In What Form is T-DNA Transferred from Agrobacterium to Plant Cells? . . . . . . . . . . . . . . . . . . . . . . . . . 3 How Soon after Agrobacterium Infection Can we Detect Transgene Expression? . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Using Tobacco BY-2 Cells to Follow the Journey of T-DNA Through the Plant Cell . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Using Tobacco BY-2 Cells to Investigate the Response of Plant Cells to Agrobacterium Infection . . . . . . . . . . . . . . . . . . . . 6 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

183 183 186 186 189 190 192 195 195 196 196 198 200 205 205

Contents

IV.3 Regulation of Elicitor-Induced Defense Responses by Ca2+ Channels and the Cell Cycle in Tobacco BY-2 Cells . . . . . . . Y. Kadota and K. Kuchitsu 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Elicitor-Induced Cytosolic Free Ca2+ Concentration ([Ca2+ ]cyt ) Change and its Regulatory Mechanisms . . . . . . . . . . . . . . . . . . . . . 3 Involvement of Putative Voltage-Dependent Ca2+ Permeable Channels, NtTPC1A/B, in Elicitor Signaling . . . . . . . . . . 4 Elicitor-Induced Cell Cycle Arrest . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Cell Cycle Dependence of Elicitor-Induced Defense Signaling . . . . . 6 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

XIII

207 207 208 210 213 214 217 218 218

Section V Other Cellular Functions V.1 1 2 3 4 V.2

1 2 3 4 5 6 7 8

Dynamic Mitochondria, their Fission and Fusion in Higher Plants . S. Arimura and N. Tsutsumi Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mitochondrial Fission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mitochondrial Fusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Use of Tobacco BY-2 Cells to Elucidate the Biosynthesis and Essential Functions of Isoprenoids . . . . . . . . . . . . . . . . . . . . . . A. Hemmerlin, E. Gerber, M.-A. Hartmann, D. Tritsch, D.N. Crowell, M. Rohmer, and T.J. Bach Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Use of Tobacco BY-2 Cells for Biosynthetic Studies . . . . . . . . . . Incorporation of Stably Labelled Glucose into Selected Isoprenoid End-Products . . . . . . . . . . . . . . . . . . . . . . Incorporation of Additional Pathway-Specific, Stably Labelled Precursors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Incorporation of Pathway-Specific Radiolabelled Precursors . . . . . . In Vivo Effects of Isoprenoid Precursors and Pathway Intermediates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isoprenylation of Proteins in BY-2 Cells . . . . . . . . . . . . . . . . . . . . . Conclusion and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

225 225 227 232 235 236 241

241 242 243 248 253 255 262 263 265

XIV

Contents

Section VI Omics VI.1 Tobacco BY-2 Proteome Display, Protein Profiling and Annotation Using Two-Dimensional Electrophoresis and Mass Spectrometry-Based Cross-Species Identification . . . . . . K. Laukens and E. Witters 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Gel-Based Analysis of the BY-2 Proteome . . . . . . . . . . . . . . . . . . . . 3 MS-Based Protein Structure Analysis . . . . . . . . . . . . . . . . . . . . . . . 4 Searching Sequence Databases with BY-2 Peptide Mass Spectra . . . 5 Proteomic Data Integration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI.2 EST and Microarray Analysis of Tobacco BY-2 Cells . . . . . . . . . . . . K. Matsuoka and I. Galis 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 The RIKEN Tobacco BY-2 EST and BY-2 Microarray . . . . . . . . . . . . 3 An Example of Microarray-Based Analysis: Jasmonate-Dependent Gene Expression and Metabolic Studies . . . . 4 Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI.3 Proteomics of Tobacco Bright Yellow-2 (BY-2) Cell Culture Plastids M.A. Siddique, W. Gruissem, and S. Baginsky 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 The Proteome of BY-2 Cell Culture Plastids . . . . . . . . . . . . . . . . . . . 3 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Protocols for BY-2 Plastid Isolation and Protein Fractionation . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 275 276 281 285 287 288 289 291 293 293 293 298 308 309 313 313 316 322 322 325

Contents

XV

Section VII Technical Advances VII.1 Cryopreservation of Tobacco BY-2 Suspension Cell Cultures Using an Encapsulation – Simple Prefreezing Method . . . . . . . . . . . . . . . 329 T. Kobayashi, T. Niino, and M. Kobayashi 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 2 Methods for Cryopreservation of Cultured Plant Cells . . . . . . . . . . 330 3 Conditions for Cryopreservation of Cell Suspension Cultures . . . . . 331 4 The Encapsulation–Simple Prefreezing Method for Cryopreservation of BY-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332 5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334 6 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336 VII.2 High Throughput Microinjection Technology for the Single-Cell Analysis of BY-2 in Vivo . . . . . . . . . . . . . . . . . . . H. Matsuoka, Y. Yamada, K. Matsuoka, and M. Saito 1 Microinjection in the Post-Genome Era . . . . . . . . . . . . . . . . . . . . . 2 Single-Cell Manipulation Supporting Robot (SMSR) . . . . . . . . . . . . 3 Microinjection of a Dominant-Negative Protein into BY-2 Cells . . . 4 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

339 339 339 343 345 346

Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 347

List of Contributors

S. Arimura Laboratory of Plant Molecular Genetics, Graduate School of Agricultural and Life Sciences, University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-8657, Japan, e-mail: [email protected] T.J. Bach Centre National de la Recherche Scientifique, Institut de Biologie Moléculaire des Plantes (UPR 2357), Département “Isoprénoïdes”, 28 rue Goethe, 67083 Strasbourg, France, e-mail: [email protected] S. Baginsky Institute of Plant Sciences and Functional Genomics Center Zürich, Swiss Federal Institute of Technology, ETH Zürich, 8092 Zürich, Switzerland, e-mail: [email protected] D. Becker Plant Molecular Physiology and Biophysics, Julius-von-Sachs-Institute for Biosciences, University of Würzburg, Julius-von-Sachs-Platz 2, 97082 Würzburg, Germany, e-mail: [email protected] D.N. Crowell Department of Biology, Indiana University–Purdue University Indianapolis, Indianapolis, Indiana 46202, USA J.H. Doonan John Innes Centre, Colney Lane, Norwich, NR4 7UH, UK, e-mail: [email protected] K. Eguchi Department of Biological Sciences, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan K. Fukui Department of Biotechnology, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita 565-0871, Osaka, Japan

XVIII

List of Contributors

I. Galis RIKEN Plant Science Center, 1-7-22 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan D. Geelen Department of Plant Production, Faculty of Bioscience Engineering, Ghent University, Coupure links 653, 9000 Gent, Belgium, e-mail: [email protected] S.B. Gelvin Department of Biological Sciences, Purdue University, West Lafayette, Indiana 47907-1392, USA, e-mail: [email protected] E. Gerber Centre National de la Recherche Scientifique, Institut de Biologie Moléculaire des Plantes (UPR 2357), Département “Isoprénoïdes”, 28 rue Goethe, 67083 Strasbourg, France W. Gruissem Institute of Plant Sciences and Functional Genomics Center Zürich, Swiss Federal Institute of Technology, ETH Zürich, 8092 Zürich, Switzerland M.-A. Hartmann Centre National de la Recherche Scientifique, Institut de Biologie Moléculaire des Plantes (UPR 2357), Département “Isoprénoïdes”, 28 rue Goethe, 67083 Strasbourg, France M. Hasebe National Institute for Basic Biology, Myodaiji-cho, Okazaki, Aichi, 444-8585 Japan R. Hedrich Plant Molecular Physiology and Biophysics, Julius-von-Sachs-Institute for Biosciences, University of Würzburg, Julius-von-Sachs-Platz 2, 97082 Würzburg, Germany A. Hemmerlin Centre National de la Recherche Scientifique, Institut de Biologie Moléculaire des Plantes (UPR 2357), Département “Isoprénoïdes”, 28 rue Goethe, 67083 Strasbourg, France D. Inzé Department of Plant Systems Biology, Flanders Interuniversity Institute for Biotechnology, Ghent University, Technologiepark 927, 9052 Gent, Belgium

List of Contributors

XIX

K. Ishibashi Graduate School of Agriculture, Hokkaido University, Sapporo 060-8589, Japan M. Ishikawa Plant-Microbe Interactions Research Unit, National Institute of Agrobiological Sciences, Tsukuba 305-8602, Japan, e-mail: [email protected] L. Jiang Department of Biology and Molecular Biotechnology Program, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China, e-mail: [email protected] Y. Kadota RIKEN Plant Science Center, 1-7-22 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan, e-mail: [email protected] M. Kobayashi BioResource Center, RIKEN Tsukuba Institute, Koyadai 3-1-1, Tsukuba, Ibaraki 305-0074, Japan T. Kobayashi BioResource Center, RIKEN Tsukuba Institute, Koyadai 3-1-1, Tsukuba, Ibaraki 305-0074, Japan, e-mail: [email protected] K. Komoda Graduate School of Agriculture, Hokkaido University, Sapporo 060-8589, Japan O.A. Koroleva John Innes Centre, Colney Lane, Norwich, NR4 7UH, UK K. Kousetsu Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan K. Kuchitsu Department of Applied Biological Science, Tokyo University of Science, 2641 Yamazaki, Noda, Chiba 278-8510, Japan K. Laukens Laboratory for Plant Biochemistry and Plant Physiology, Center for Proteome Analysis and Mass Spectrometry, Department of Biology, University of Antwerp, Groenenborgerlaan 171, 2020 Antwerp, Belgium, e-mail: [email protected]

XX

List of Contributors

S.W. Lo Department of Biology and Molecular Biotechnology Program, The Chinese University of Hong Kong, Shatin, New Territories, Hong Kong, China Y. Machida Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan, e-mail: [email protected] S. Matsunaga Department of Biotechnology, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita 565-0871, Osaka, Japan, e-mail: [email protected] H. Matsuoka Department of Biotechnology and Life Science, Tokyo University of Agriculture and Technology, Koganei, Tokyo 184-8588, Japan, e-mail: [email protected] K. Matsuoka RIKEN Plant Science Center, 1-7-22 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan, e-mail: [email protected] Y. Miyazawa Graduate School of Life Sciences, Tohoku University, Miyagi 980-8577, Japan, e-mail: [email protected] T. Murata National Institute for Basic Biology, Myodaiji-cho, Okazaki, Aichi, 444-8585 Japan, e-mail: [email protected] T. Nagata Department of Biological Sciences, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan T. Niino Genebank, National Institute of Agrobiological Sciences, Kannondai 2-1-2, Tsukuba, Ibaraki 305-8602, Japan I. Nishida Department of Biological Sciences, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan N. Ohmido Faculty of Human Development, Kobe University, 3-11 Tsurukabuto, Nada, Kobe 657-8501, Hyogo, Japan

List of Contributors

XXI

J. Petrášek Institute of Experimental Botany ASCR, Rozvojová 135, 16502 Prague 6, Czech Republic, e-mail: [email protected] C. Ritzenthaler IBMP CNRS, Institut de Biologie Moléculaire des Plantes, Strasbourg, France G.R. Roberts John Innes Centre, Colney Lane, Norwich, NR4 7UH, UK D.G. Robinson Heidelberg Institute of Plant Sciences-Cell Biology, University of Heidelberg, Im Neuenheimer Feld 230, 69120 Heidelberg, Germany, e-mail: [email protected] M. Rohmer Institut LeBel, Université Louis Pasteur/CNRS (UMR 7123), 4 rue Blaise Pascal, 67070 Strasbourg, France M. Saito Department of Biotechnology and Life Science, Tokyo University of Agriculture and Technology, Koganei, Tokyo 184-8588, Japan A. Sakai Department of Biological Sciences, Faculty of Science, Nara Women’s University, Nara 630-8506, Japan M. Sasabe Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan T. Shimizu Department of Biological Sciences, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan, e-mail: [email protected] M.A. Siddique Institute of Plant Sciences and Functional Genomics Center Zurich, Swiss Federal Institute of Technology, ETH Zurich, 8092 Zurich, Switzerland T. Soyano Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan

XXII

List of Contributors

T. Suzuki Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan Y. Takahashi Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan H. Tanaka Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan M.L. Tomlinson John Innes Centre, Colney Lane, Norwich, NR4 7UH, UK K. Toyooka RIKEN Plant Science Center, 1-7-22 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan, e-mail: [email protected] D. Tritsch Institut LeBel, Université Louis Pasteur/CNRS (UMR 7123), 4 rue Blaise Pascal, 67070 Strasbourg, France N. Tsutsumi Laboratory of Plant Molecular Genetics, Graduate School of Agricultural and Life Sciences, University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-8657, Japan E. Witters Laboratory for Plant Biochemistry and Plant Physiology, Center for Proteome Analysis and Mass Spectrometry, Department of Biology, University of Antwerp, Groenenborgerlaan 171, 2020 Antwerp, Belgium Y. Yamada CREST (Core Research for Evolutional Science and Technology), Japan Science and Technology Agency, Honcho 4-1-8, Kawaguchi, Saitama 332-0012, Japan E. Zažímalová Department of Plant Physiology, Faculty of Science, Charles University, Vinièˇcná 5, 12844 Prague 2, Czech Republic

Section I Cell Cycle and Cellular Dynamics

I.1 Novel Approaches for Cell Cycle Analysis in BY-2 O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan1

1 Introduction The BY-2 cell culture has proved a useful, and indeed indispensable, tool for the analysis of the plant cell cycle. This is, in large part, due to the degree to which the cells can be synchronised during the cell cycle (Nagata et al. 1982; Nagata 2004). The cell line has several other features that are advantageous for studies of basic plant cell biology. The cells are relatively large and grow as long chains. Combined with excellent optical clarity, this regular growth habit lends itself to direct microscopical observation of cellular processes and it has been used for this propose extensively. Finally, BY-2 can be easily transformed so that cellular processes can be manipulated at the molecular level, and the introduction of green fluorescent protein (GFP)-tagged constructs allows direct observation of cell division, protein localisation and organelle dynamics. In this chapter we concentrate on the production and use of transgenic BY-2. We describe and discuss the use of a chemically inducible gene switch which, when used in combination with synchronised cells, makes BY-2 an ideal system for detailed studies of the plant cell cycle. This approach allows the up- or down-regulation of gene function at precise times in the cell cycle. Potential applications include inducible overexpression, growth analysis, dual-fluorescent labelling for flow cytometric analysis, RNAi silencing and transient expression. We provide detailed protocols and discuss how to improve throughput for transformation and analysis of the cell culture.

2 A Modified alc-Inducible System for Transgene Expression in BY-2 Constitutive expression of cell cycle regulators can be difficult to interpret because compensatory changes in related processes may disguise (or confuse) changes in the actual target. Several studies indicate that constitutive overexpression of cyclins can affect the length of the S- and G2-phases, which could indirectly lead either to the observed shortening of the G1-phase or to increased length of the S- and G2-phases (Dewitte et al. 2003). Similarly, constitutive lossof-function mutations and dominant-negative constructs suffer from similar 1 John Innes Centre, Colney Lane, Norwich, NR4 7UH, UK, e-mail: [email protected]

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan

problems, the most severe being observed with essential genes where early lethality can limit phenotypic analysis. Moreover, in any transgenic experiment, there is high variability between parallel transgenic lines which can lead to difficulties in interpretation of the effect of the transgene. Variable levels of silencing and effects due to the genomic context of the integration site of the transgene can contribute to overall variability. In other model systems, such as yeast, conditional mutants and gene switches are widely used to overcome these problems. Although there are a few examples where temperature-sensitive mutants have been sought and isolated from Arabidopsis, this approach has not been widely applied in plant biology. Chemically inducible gene switches, however, have been quite widely applied in both whole plants and cell cultures, and have been useful in providing insights into essential biological processes. As compared to constitutive expression, conditional gene expression has several benefits. In the first place, it enables the primary effects of gene expression to be assayed. By application or, in some cases, withdrawal of the chemical, gene expression can be initiated at a time chosen by the experimenter. This has the advantage that the cell has less time to adapt to the presence of the gene product and one is more likely to observe the primary response. A second benefit, really an extreme example of the first, occurs if the gene product is deleterious to growth or viability. In this case, the effect of transient accumulation can be assayed without the selection against expression that would occur over successive cell generations. Finally, for complex processes such as the cell cycle or development, conditional expression allows conditional complementation of mutant phenotype at different stages. This can provide novel insights into a gene’s function, which may change during the different stages of development (Laufs et al. 2003) or the cell cycle. Progression through both development and the cell cycle involves a series of interdependent sequential functions where late functions can depend on the completion of early functions. Perturbation of early functions can indirectly modify late functions and constitutive upor down-regulation of gene function can be difficult to interpret. Work in whole plants indicated that the alcR/alcA ethanol-inducible regulon provided a switchable gene expression system suitable for both overexpression and complementation studies (Roslan et al. 2001; Deveaux et al. 2003; Laufs et al. 2003) and has been widely used in crop plants. The ethanol-inducible alcR/alcA (alc) gene expression system is widely used for conditional gene expression studies in both plants and fungi. It is a twocomponent chemically inducible gene expression system, originally developed as a gene switch in Aspergillus nidulans (Waring et al. 1989). The alcR encoded transcription factor ALCR and the alcA target promoter constitute the two components. The ALCR transcription factor drives gene expression from the alcA target promoter only in the presence of ethanol or acetaldehyde (Flipphi et al. 2001). It provides high speed of induction and the volatile nature of the inducer allows its removal from the system, resulting in a highly dynamic switch. However, for longer periods of induction ethanol needs to be supplied constantly. The ethanol switch has numerous advantages for a large-scale agri-

Novel Approaches for Cell Cycle Analysis in BY-2

5

cultural application, using a non-toxic, cheap, stable chemical. Although it would be very useful to be able to use the same system in both plants and cell cultures, the utility of the ethanol switch in suspension cells is severely limited as gene expression seems to be constitutive even in the absence of exogenous inducer. Under certain physiological conditions, notably anoxia, it is thought that substances are produced by the cells that mimic the effect of ethanol. This “leaky” expression is apparent when the alc system is used in plant suspension or callus cell cultures (Roberts et al. 2005) and, while it is possible to reduce the level of expression by additional aeration, this background expression is difficult to eliminate. A modified alc system, the alc-GR system, was generated by fusing the rat glucocorticoid receptor (GR) domain to the ALCR transcription factor (Roberts et al. 2005). Fusion of the GR domain to a protein restricts the protein to the cytoplasm until dexamethasone (dex) is applied. Then the fusion enters the nucleus. This is widely used in plants to produce dex-inducible transcription factors (Wagner et al. 1999; Gallois et al. 2002; Gomez-Mena et al. 2005). Tests on BY-2 have shown that the alc-GR system is tightly dex-inducible. Exogenously applied ethanol is not required for dex-inducible transgene expression and ethanol does not induce expression in the absence of dex. The alc-GR system has proven to be a good switch for cell synchrony studies. First, dex activation of the ALCR-GR transcription does not appear to perturb cell cycle progression. The inert nature of the ALCR-GR transcription factor in cell synchrony experiments provides a good basis for experiments where transgenes are expressed in cell cycle synchronised cultures. Therefore, using the alc-GR system, any changes to the duration of cell cycle phases can be directly apportioned to the induced transgene. Second, induction is very rapid – the expressed protein appears as soon as 1 h after induction (Koroleva et al. 2004; Roberts et al. 2005).

3 Use of the alc System to Define the Time of Action of Induced Protein We used the alc-GR expression system to monitor the consequences of D1 cyclin gene expression. D cyclins are known to be rate-limiting components for progression through G1, as association of these unstable proteins with the cyclin-dependent kinases (CDKs) leads to phosphorylation of the Rb protein, an event thought necessary for release of the E2F transcription factors and activation of the S-phase (Oakenfull et al. 2002). We addressed whether CycD1 could promote cell cycle progression in BY-2. We used timed CycD1 expression to define which stages of the cell cycle were affected. Cooper (1998) argued that it is not possible to identify G1 cyclins by the overexpression induced changes in the lengths of individual cell cycle phases without considering changes in the overall cellular growth rate. Any shortening of the G1-phase could be due

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O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan

to an increase in the rate of mass synthesis in all phases of the cell cycle. Our experimental approach allowed us to solve this logical problem by inducing extra D1 cyclin at specific times of the cell cycle and measuring the immediate effect. We were able to temporally dissect the function of D1 cyclin during the G1/S- and S/G2-phases of the cell cycle.

4 An Example of Functional Analysis Using the AlcR-GR Gene Switch Briefly, transgenic lines of BY-2 already carrying the gene switch cassette 35S-alcR-GR (Roberts et al. 2005) were retransformed with a second cassette containing the CycD1 gene or tagged HA:CycD1 under control of alc A promoter, using a hypervirulent strain of Agrobacterium tumefaciens (LBA4404.pBBR1MCSvirGN54D) (van der Fits et al. 2000) as described in Koroleva et al. (2004) and Protocol 1 (see Sect. 7.1). 4.1 D1 Cyclin Inducible Over expression in Synchronised Cells Suspension cultures can be synchronised at several different stages during the cell cycle using different methods. Stationary cells are arrested in the G0 phase by withdrawing nutrients from the media, such as sucrose (Menges and Murray 2002). Entry into S phase can be reversibly blocked by aphidicolin, a potent inhibitor of DNA polymerases α and δ (Sala et al. 1983; Nagata et al. 1992). Furthermore, the cells can be synchronised at the onset of mitosis by the microtubule-disrupting drug propyzamide (Nagata and Kumagai 1999). Using two types of synchrony, either G0 or S-phase arrest, we determined the effect of D1 cyclin expression at the different time points during the cell cycle (Koroleva et al. 2004). To test whether CycD1 had a specific effect on cell cycle progression, we first expressed CycD1 in cells released from stationary phase. Sucrose-starved or stationary cultures can be induced to re-enter the cell cycle by addition of fresh media and they proceed in a semi-synchronous manner into S-phase and mitosis. If CycD1 was induced at the same time as cells are sub-cultured into fresh media, entry into mitosis was accelerated, as judged by mitotic index (MI) measurements, by at least 2 h. CycD1 induction led to not only faster progression into mitosis but also a significantly increased MI. The induced culture entered mitosis and reached its peak value 2 h earlier and also achieved an approximately 4% higher peak value of MI compared to the control. Thus, overexpression of CycD1 from G0 significantly accelerated cell cycle progression through S-phase to mitosis. We used flow cytometric measurement of DNA content to follow the dynamics of S-phase entry, with or without expression of CycD1. Stationary BY-2 cells had a mainly G1 nuclearcontent. Even at 6 h after release from G0, the

Novel Approaches for Cell Cycle Analysis in BY-2

7

induced culture had a much higher proportion of cells in S-phase compared to non-induced culture. By 10 h the proportion of nuclei in the G2 peak exceeded the number in the G1 peak in induced culture while G1 nuclei still predominated in non-induced culture and much fewer cells contributed to the G2 peak (Koroleva et al. 2004). Screened at 8 h after induction, 8 out of 8 CycD1 lines and 4 out of 8 HA-CycD1 lines had a remarkable increase of cells in Sand G2-phases compared to non-induced controls. This marked acceleration of S-phase entry caused by overexpression of CycD1 represents a 2 h advance and, therefore, could fully account for the accelerated progression to mitosis. However, this does not rule out the possibility that S- and G2-phases were also affected by CycD1. To test whether the cell cycle acceleration was specific to G1 phase, we assessed the effect of CycD1 expression on S-phase cells by release from aphidicolin arrest. We induced expression of CycD1 during either early S-phase (dex added together with aphidicolin and then again after the removal of aphidicolin) or late S-phase/G2-phase (dex was added immediately after removal of aphidicolin). The cultures that were induced on release from the S-phase block progressed faster into mitosis and achieved a higher maximal value of MI than the non-induced control. The number of mitotic cells by 7−10 h was significantly increased and we estimated that induction of CycD1 expression at this stage accelerated entry into mitosis by about 1 h, suggesting an effect either in late S-phase or on the G2/M transition. We analysed the appearance of newly divided cells by flow cytometry to follow progression into and completion of mitosis. By 8−9 h after release, the number of cells that had passed through mitosis (as judged by the G1/G2 index) became significantly higher in cultures induced by dex, and the difference was maintained during several following hours. This effect could be due to the shortening of either the S- or G2-phase, as HA-CYCD1 protein could be detected as early as 1 h after induction, which should correspond to the peak in DNA synthesis in the S-phase after the release from aphidicolin (Sorrell et al. 1999). The observed effect of accelerated entry into mitosis could result in CycD1 effect in either late S- or G2-phase. Therefore, we tested the effect of earlier induction by inducing both during and after the aphidicolin block. In this case, the acceleration into mitosis was even greater, suggesting either that a long time was required for protein levels to accumulate or that CycD1 promotes progress through S-phase even in the presence of an aphidicolin block. The MI increased much earlier in cultures under continuous induction, with a significant increase in the number of cells in mitosis at 4 h after the aphidicolin block was released, indicating that progression into mitosis was faster (Koroleva et al. 2004). The initially higher speed of cell cycle progression in the cultures continuously induced by dex was most likely achieved due to accelerated progression through the S-phase. However, this acceleration does not speed up completion of the cell cycle as the number of cells that had passed through mitosis (as judged by the G1/G2 index) became equal in cultures induced by dex once or twice. By 12 h after release, the G1/G2 index became equal in control cultures and cultures induced

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O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan

by dex twice, but lower than the cultures induced once, indicating that cultures induced in early S-phase entered mitosis earlier but spent a prolonged time in mitosis. This was not the case when expression was induced at the time of aphidicolin release. The faster progression through mitosis in these cultures therefore could be attributed to late S- or G2-M-specific action of CycD1, and earlier expression during an S-phase block may lead to premature mitotic entry that then leads to a delay in mitotic progression. 4.2 The Activity and Potential Role of D1-Associated Kinase The BY-2 cell system is also amenable to biochemical studies, allowing protein complexes to be isolated from cells at defined stages of the cell cycle. To define the in vivo partners of D1 cyclin, we transformed BY-2 with epitope tagged versions of CycD1. Using an HA-tagged version, we immunoprecipitated proteins associated with CYCD1, from total soluble protein extracts. Western blot analysis with antibodies against the “PSTAIR” motif (specific for A-type CDKs) clearly demonstrated that CDKA had co-precipitated with CYCD1 (Koroleva et al. 2004). To assess whether CYCD1 bound to CDKB, the mitosis-specific CDKs, we probed immunoprecipitates with antibodies against CDKB1;1. Although, the antibody recognised the CDKB1 protein in whole cell extracts, we failed to detect it in the HA immunoprecipitates. However, data from yeast two hybrid analysis indicated that CYCD1 and CDKB2;1 can interact (O.A. Koroleva, M.L. Tomlinson, J.H. Doonan, unpublished). Therefore, it remains possible that CYCD1 does interact with both classes of CDK in vivo. CDKA remained at a constant level during the cell cycle, as reported previously by Sorrell et al. (2001), and this level was not changed significantly by CycD1 expression. CDKB1 protein was undetectable in stationary cells. After re-entering the cell cycle from a G0 release, CDKB1 could be first detected in late S-phase and then its levels increased gradually, reaching a peak during mitosis. Therefore, it is unlikely that CDKB would have mediated the CycD1-induced acceleration in the G0 synchrony experiment. To determine the substrate specificity of the CYCD1-associated kinase at different phases of the cell cycle during G1 synchrony experiment, we assayed HA-associated kinase activity towards two known substrates, histone H1 and Rb, and compared this with the total fraction of endogenous tobacco CDKs purified on p13Suc1 beads. We found that the expression of HA-CYCD1 led to an initial increase in HA-associated histone H1 kinase activity, followed by decline and second peak of activity at the time of mitosis. The non-induced control culture had a very low background level of HA-associated histone H1 kinase activity that did not change significantly over the time-course. HA-dependent phosphorylation of Rb protein increased steadily from G1 to mitosis in the induced culture. The Rb kinase activity in non-induced culture remained low. We measured total histone and Rb kinase activity to estimate whether ectopic expression of CycD1 led to global changes in CDK activity towards either of

Novel Approaches for Cell Cycle Analysis in BY-2

9

these substrates. The kinase activity associated with p13Suc1 beads was about an order of magnitude higher towards both substrates, compared to the HACYCD1-interacting fraction precipitated on HA beads. Total histone H1 kinase activity in the induced culture was dramatically increased compared to the noninduced control. The non-induced culture showed a steady increase in kinase activity from G1- through S-phase into mitosis. Kinase activity towards Rb protein followed a similar trend of steady increase through G1- and S-phase, and then a slight decrease at the time of mitosis in both induced and noninduced cultures, but the induced culture had higher levels of Rb kinase activity at each time point (Koroleva et al. 2004). These data indicate that the HA-associated CYCD1-dependent kinase makes a significant contribution to the increase in total CDK activity, and its contribution appears to be amplified, probably by activation of downstream kinases. 4.3 CYCD1 Protein Localisation Studies BY-2 cells also provide an excellent system by which to study protein localisation. Protein localisation can often provide clues as to its function or regulation. The living cell is a complex of morphologically distinct compartments with discrete and well-defined functions. At the intracellular level, compartmentation is achieved by the existence of multiple membrane-bound organelles, each containing a spectrum of biomolecules and set of proteins forming the organelle-specific “proteome”. An N-terminal fusion of GFP to CycD1 was made using the GATEWAY system (Invitrogen) and pGWB6 expression vector (a gift of Dr. T. Nakagawa, Shimane University, Japan) and expressed in lines of BY-2, using a hypervirulent strain of Agrobacterium (Koroleva et al. 2005). GFP-CYCD1 is primarily present in the nuclei of interphase cells but was much reduced or absent from the spindle domain during mitosis (Koroleva et al. 2004), where it may get degraded by a ubiquitin-dependent pathway: MG132 proteasome inhibitor treatment of cells led to increased levels of CycD1 on Western blots, which is consistent with CycD1 being an unstable regulator of CDK activity in the nucleus.

5 Transient Protein Expression 5.1 Comparative Analysis of Applications for Tobacco BY-2 vs Arabidopsis Col-0 Cell Cultures Stable transformation of BY-2 cells using inducible promoters to drive gene expression in a highly controlled manner, therefore, allows detailed functional analysis of genes involved in cell cycle regulation. Synchronised cell populations provide insight into the biochemistry, function and interactions of the transgenic protein. However, using current methodology, the number of genes

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O.A. Koroleva, G.R. Roberts, M.L. Tomlinson, and J.H. Doonan

that can be analysed in this way is limited. In part, this limitation is due to the high variability between independent transgenic cell lines carrying the same construct and makes necessary the rigorous statistical verification of the observed characteristics across several cell lines so that the phenotype can be correlated with the expression of the transgene. Thus, if we wish to use cell suspensions for systematic analysis of gene function on a whole genome scale, then new methods must be developed. In an effort to solve these problems, we have investigated the possibility of using transient transformation for understanding gene function. This approach is widely used in mammalian cells where viral vectors are used to deliver genes to whole populations of cultured cells. Unfortunately, most plant DNA viruses have limited tolerance in terms of the size of gene that they will accept and this has limited their development as vectors. Transient expression in plant cells, however, is achievable and has been widely used to dissect gene function. Perhaps the most commonly employed technique uses transformation of protoplasts with naked DNA and this is both effective and efficient in BY-2. This method can produce 25–30% of cells expressing protein within 18−24 h of transfection with GFP, and similar levels 36−40 h after transfection with the slow-maturating fluorescent protein dsRed (Bhat and Thompson 2004). The other method exploits Agrobacterium-mediated transformation and this has the advantages that whole cells can be transformed and the same plasmids and strains can be used subsequently to produce transgenic plants. However, we found that the efficiency of transient transformation of BY-2 cells by Agrobacteria is quite low: the percentage of cells that express the transgene is usually less than 0.01% of the cell input, which is more than adequate for the selection of stable transgenic lines but sub-optimal for use in transient transformation. However, the rate of transient transformation may vary depending on the culture conditions, length of co-cultivation with Agrobacteria and the nature of the introduced transgene. Agrobacterium strains are known to vary with respect to their ability to transfect particular hosts, so it is hoped that more efficient strains can be found. However, since transient expression using Agrobacterium offers so many advantages, in terms of both increased throughput and better integration into whole plant studies, we investigated other cell cultures as possible systems. Arabidopsis Col-0, originally derived in C. Koncz’s laboratory, had shown promise for transient expression of proteins for biochemical studies (Mathur et al. 1998). In contrast to BY-2, the Col-0 Arabidopsis cell culture has a much higher susceptibility to transformation by Agrobacteria and the proportion of cells transiently expressing the transgene can be up to 70% (Koroleva et al. 2005). We have developed a protocol (Protocol 3) that allows systematic protein localisation (Koroleva et al. 2005), which also has the potential for functional and biochemical analysis (Koroleva et al. 2004; Chan et al. 2005; Korolev et al. 2005; Mao et al. 2005). This method complements the traditional range of applications of the BY-2 cell culture: one strategy is to use Arabidopsis cell culture for fast screening of transient expression, and continue with detailed analysis

Novel Approaches for Cell Cycle Analysis in BY-2

11

Table 1. Comparative analysis of a range of applications for BY-2 vs Col-0 cell cultures, using transformation by Agrobacterium for both. The total number of asterisks indicates the relative merit of the different cell types

Option

Transformation by Agrobacterium Stable expression in BY-2 Transient in Col-0

Maintenance of the culture Transformations per week Time for expression Time for analysis GFP localisation Cell cycle analysis Growth analysis Inducible expression RNAi

Easy 36 2–4 weeks Unlimited Good resolution Yes Yes Yes Yes

Not easy > 96 3–4 days 3–5 days Brighter signal Yes for some No No Not yet

* – – * * * * * *

– * * – * * – – –

Problems

– High variability in cell size/growth – GFP fusions are not bright –

– Larger cell clusters

–

–

– Poor resolution of cell structures –

–

–

7

4

Score

Advantage BY-2 Col-0

of “interesting” genes in BY-2 later. Table 1 summarises the comparative characteristics and advantages of both cell cultures for analysis of transgene effect after transformation by Agrobacterium. Some particular types of analysis will be discussed below. 5.2 Use of Arabidopsis Cell Culture for Systematic Protein Localisation The function of a particular protein is best understood in the context of its micro-environment. Where the protein is located will often affect its function, by restricting access either to substrates or to interacting partners. Interacting partners will determine in part how multi-protein complexes are constructed and how they respond to extraneous signals from the environment or neighbouring cells. Information about a protein’s location or “compartmentation” is therefore essential to fully understand its function. Despite the existence of many software programs for prediction of putative protein localisation, it may not always be possible to predict localisation from the primary amino acid sequence because of the major role played by protein–protein interactions. A protein may be delivered to a given location as part of a multiprotein complex, and be anchored there by such interactions. Some motifs are relatively easy to recognise, but even they do not necessarily determine the ultimate location. For example, the presence of nuclear localisation signal (NLS) or nuclear export signal (NES) indicates probable nuclear or cytoplasmic location, but some NES sequences can have differential activity and some NES-like sequences are

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inactive (Henderson and Eleftheriou 2000). Studies aimed at defining specific nuclear localisation or nuclear export signals rely on experimental comparison of localisation of wild type with that of mutant proteins with introduced amino acid replacements in the tested motif (Henderson and Eleftheriou 2000). Moreover, many proteins change their location in response to signals or altered physiological states, so experimental verification will always be necessary. Previous approaches aimed at defining protein localisation have tended to be very laborious and time-consuming, allowing analysis of only a small number of proteins. Ideally, initial information on protein localisation could be obtained from a systematic screen that would utilise existing genomic resources and be cheap and convenient to implement. Excellent genomic resources are now available for Arabidopsis: the SSP ORF collection contains about 10,500 publicly available and verified Arabidopsis full-length ORFs, trimmed at the 5
translational start site (Yamada et al. 2003). We have taken advantage of this collection of clones and used GATEWAY technology for the fast and efficient production of GFP translational fusions with Arabidopsis proteins for localisation studies in model suspension culture cells via Agrobacterium-mediated transient transformation. For more detailed cell cycle studies some of these have been transferred in BY-2 cells. Related proteins often have different preferred subcellular localisation which reflect function: for example, mammalian B cyclins accumulate in cytoplasm in interphase and first appear on centrosomes in prophase (Jackman et al. 2003). We have shown that different CDKs (CDK C;2, CDK D;1, CDK D;2, CDK D;3 and CDKF;1) and cyclins (CycD3;1, CycA2;3, Cyc A3;2 and CycT1) have distinct patterns (Koroleva et al. 2005). Even closely related CDKs that are localised within the nucleus have subtly different GFP-fusion localisation patterns, although the functional significance of this is presently unclear. The use of EDE1-GFP fusions in transient assays has revealed novel insights into microtubule behaviour in the plant cell cortex (Chan et al. 2003) and in organisation of the mitotic spindle (Chan et al. 2005). Ideally, the GFP tag should be introduced into the native genomic copy of the gene. Comprehensive screens in yeast have already exploited site-specific integration to precisely place GFP in the correct genomic context, but this option is not yet possible in higher plants since site-specific integration is a very rare event (Lutz et al. 2004). Site-specific integration of GFP to give translational fusions in the correct genomic context has been reported in the moss Physcomitrella (Kiessling et al. 2000), but genome-wide surveys are not yet possible, due to the fragmentary nature of the cDNA collections available for this species (Reski and Cove 2004). Recently, Tian et al. (2004) demonstrated the use of multiple 35S enhancers in combination with native promoters to generate brighter fluorescent signals than provided by the native promoters alone. Apparently, the addition of the 35S enhancers and transformation into varied genomic contexts did not change either the tissue or cell type distribution or subcellular localisation and, in most cases, moderately increased levels of expression should provide

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valid localisation patterns. Other studies have compared the effect of placing GFP at the N terminus or at the C terminus of the ORF (Simpson et al. 2000; Escobar et al. 2003). The N-terminus strategy described here was dictated largely by the nature of the cDNA collection. While the entire collection was uniformly trimmed to an in-frame ATG at or near the N terminus, only a few were precisely trimmed at the C terminus and even these contained a stop codon that precluded C-terminal fusions. 5.3 Systematic Stop Codon Removal to Allow C-Terminal Protein Fusions Since many localisation signals are located at either the N or the C terminus, where proximal GFP fusion might interfere with correct protein localisation, it would be ideal to compare both types of fusions. We therefore devised a strategy for the systematic removal of stop codons from trimmed ORFs to allow in-frame fusion to GFP using the GW cloning system. There are at least two possible strategies to remove stop codons. One approach is to design a unique primer for each cDNA. Another approach, possible only where the cDNA has been already trimmed at the 3
end, is to design a primer that can mutate the stop codon for large subsets of the clones. This allows the use of a few primers to modify many genes. However, the latter strategy requires that the mutagenic nucleotides are close to the 3
end of the primer and, as such, are susceptible to repair during the PCR reaction. We designed a 3
oligonucleotide primer to anneal primarily to the vector sequence (in this case the SSP vector pUNI51) flanking the inserted ORFs but overlapping the stop codon and to contain a base pair mismatch to the stop codon. The targeted mismatch and the three surrounding residues of the mutagenic oligos were phosphorothioate residues. Phosphorothioate oligonucleotides prevent nuclease-mediated turnover of introduced synthetic nucleotide sequences (Kurreck 2003) and, therefore, should be resistant to the 3–5
endonuclease proof-reading activity present in most high-fidelity polymerases used for PCR. This method relies on conserved vector sequence being close to the 3
end of the ORF and is only applicable to the SSP clones originating from the PGEC lab, which were trimmed at both the 5
and 3
ends. This method presents an inexpensive and systematic way of analysing expression patterns for GFP fusions at both the C and N terminus of a protein of interest and we have found it to be scalable.

6 Emerging Techniques 6.1 RNAi as a New Tool for Gene Silencing in BY-2 Cells Increasing the level of a given gene product is straightforward and widely used. It would be immensely useful to reduce the level of a particular gene product.

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Fig. 1. RNAi silencing of previously introduced, stably expressed transgene (soluble GFP) in a BY-2 culture. Top panel shows calli viewed under blue light illumination; GFP-positive calli are fluorescent green. Bottom panel shows bright field photograph of the same area as shown in the panel above. A Control cells of the original GFP line plated on kanamycin-containing media. B A typical plate with media containing both hygromycin (for selection of GFP-RNAi transformed calli) and kanamycin (the selection marker of the original 35S:GFP line). Note that the fluorescent speckles represent non-transformed by RNAi vector cells of the original 35S:GFP line, proliferation of which is inhibited by hygromycin. C A rare partial silencing event: BY-2 calli transformed with a GFP RNAi construct and selected on hygromycin. One callus has moderate level of GFP expression. Note that neighbouring callus shows complete silencing

Silencing a particular gene using the RNAi technique is the standard approach, offering high precision in targeting selected sequences, and is widely exploited in whole plants. The phenomenon also occurs in BY-2 as demonstrated here by knockdown of GFP expression. Cells expressing GFP were secondarily transformed by a hairpin-loop RNAi construct containing GFP fragments. After antibiotic selection on plates, only one out of 100 resistant calli had detectable GFP fluorescence (Fig. 1B, C). In the same time, 100% of control calli had a very high level of fluorescence (Fig. 1A). The practical implementation of this approach relies on precise sequence match between the template introduced into the cell in the RNAi vector and the endogenous target. Heterologous sequences tend to be ineffective in triggering silencing and therefore the technique has limited application in species with not completely sequenced genomes. However, with the progress of BY-2 EST sequencing (http://mrg.psc.riken.go.jp/strc/index.htm), opportunities for studying the effects of gene silencing will increase. Most advantageous would be to use inducible elimination of a particular gene product at a specific cell cycle phase, thus complementing overexpression studies and providing in-depth analysis of the gene function.

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6.2 Growth Analysis Using Bioscreen C Measuring the growth rate of suspension cultures using the conventional technique of daily sampling and estimation of cell biomass or cell number is a very tedious and time-consuming analysis. We adapted a technique used by microbiologists for estimating growth parameters. A Bioscreen C plate reader shaker incubator and Growth Curves software was used to monitor optical density (OD600 ) of cell suspensions (Koroleva et al. 2004). The instrument allowed us to follow the rise in optical density (correlated with cell numbers in the sample) in up to 200 samples simultaneously, thus providing statistically proven results in a short period of time. 6.3 Two-Way Flow Cytometry Analysis As demonstrated in Fig. 2, flow cytometry can be used to distinguish cells expressing a specific nuclear-localised transgene fused to GFP, and simultaneously to estimate the ploidy of the nucleus stained by DAPI. Therefore, future applications might include an analysis of a changed rate of progression of cells

Fig. 2. Analysis of cell ploidy and proportion of GFP-positive cells in G1 (2C) and G2 (4C) phases of the cell cycle in cell culture transformed by GFP fusion of DNA-binding protein (At1g72740). A Cell ploidy in the culture 3 days after transformation. The x and y axes display the relative DNA content (linear scale) and number of nuclei respectively. Gates RN1 and RN2 were used to select nuclei in G1- and G2-phases for subsequent analysis. B GFP-positive cells in the G1 population, separated by using gate RN1. C GFP-positive cells in the G2 population, separated by using gate RN2. Proportion of GFP-positive cells was almost equal in G1 (41%) and G2 (34%) phases, so the expression of the transgene did not affect progression through the cell cycle

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through the DNA replication affected by a specific transgene. Separating of cells expressing and not expressing the GFP-tagged transgene on the cell sorting flow cytometer would make it possible to compare cDNA libraries and/or protein expression profiles and thus to determine a global effect of overexpression of a particular gene.

7 Protocols 7.1 Protocol 1. Method for Multiple BY-2 Transformation Day 1. Sub-culture BY-2 cells. Inoculate 2−5 ml of 1-week-old BY-2 cells in 100 ml of BY-2 medium. Stock BY-2 should be sub-cultured weekly. Day 2. Agrobacterium culture. Inoculate 0.7−1.5 ml of synthetic culture medium AB, containing appropriate antibiotics with Agrobacterium and grow to OD 0.5–0.8 for about 24 h at 25−28 ◦ C. Antibiotics used for all Agrobacterium strains: rifampicin 20 µg/ml (diluted from 20 mg/ml methanol stock); strain-specific: gentamycin 40 µg/ml (diluted from 50 mg/ml stock) or chloramphenicol 75 µg/ml (diluted from 50 mg/ml ethanol stock); plasmid selection marker: e. g. kanamycin 50 µg/ml (diluted from 100 mg/ml stock). Day 3. Co-culture Agrobacterium and BY-2 cells in deep Petri dishes (100 × 20 mm, Falcon), add 4 ml of 3-day-old BY-2 cells and 100 µl of Agrobacterium suspension and seal with Micropore tape. Co-culture for 2 days in the dark at 25 ◦ C without agitation. The Petri dishes must be horizontal. Day 5. Wash and spread BY-2 cells on selection plates, either using manifold or by centrifugation. Washing Using Manifold 1. Autoclave top parts of the Millipore filter manifold (cell harvester) and Miracloth filters (double-layered circles cut out by hot copper tube of appropriate diameter, wrapped in foil). Sterilise rubber seals in 5% peroxide for 15−20 min. Wipe bottom part of manifold with 70% ethanol. 2. Assemble manifold in a flow hood. Pour contents of each Petri dish (transformed cells) into a separate well of the manifold. Wash each well with sterile BY-2 medium to remove Agrobacteria. Repeat washing using approximately 80 ml of BY-2 medium per well. 3. Dismantle manifold. Using a sterile pipette, pour 1 ml of BY-2 medium onto a selection plate (40 ml of BY-2 media with 0.4% Phytagel and appropriate antibiotics, in a 100 × 20 mm Falcon Petri dish). Using sterile forceps, lift filter from manifold stage and carefully tip it over pool of liquid on the

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Phytagel plate. Close the plate and rotate with gentle shaking, spreading the cells evenly over the surface of Phytagel. 4. Seal the plate with Micropore tape and incubate in the horizontal position in the dark at 25 ◦ C. Transformed calli become visible after 2–3 weeks (2−5 mm in diameter). Alternative: Washing Using Centrifuge 1. Cell harvesting: add 8 ml of BY-2 medium and homogenise using a Pasteur pipette. 2. Centrifuge at 1,000 rpm for 1 min. Remove supernatant. Add fresh media and repeat washing step five times. 3. Finally resuspend in 1−2 ml and spread onto deep Petri dish plates with 0.4% Phytagel BY-2 medium containing appropriate antibiotics. 4. As step 4 above. Week 3–4. First sub-culture. Transfer spherical colony with a sterile colony picker from the transgenic selection plate onto a new Phytagel plate, again containing appropriate antibiotics. About nine colonies can be transferred to each new plate. Week 4–5. Sub-culture into liquid BY-2 medium. Transgene expression can be analysed as appropriate for each colony on the plates. A piece of each selected colony is removed from the plate and transferred to a small (50 ml) flask containing 20 ml of BY-2 liquid medium and antibiotics. Flasks are incubated at 25 ◦ C on a shaker. When the culture becomes thick with BY-2 cells, it can then be sub-cultured 1−10 ml in 100 ml. Carbenicillin selection should be maintained for at least three rounds of sub-culturing (weekly 1−2 ml in 100 ml) in liquid medium to ensure that Agrobacterium is completely eliminated. It is also sometimes necessary to repeat antibiotic selection on the transgenic line, in particular if the transgene is deleterious to the culture. 7.2 Protocol 2. Procedure for Using the alc-GR System in BY-2 1. Generate BY-2 primary transformants containing the 35S-alcR-GR cassette as described above. It is useful to select for these transformants on plates containing antibiotics and 1 µM dexamethasone. Inclusion of dexamethasone enables counter-selection of highly expressing ALCR-GR transformants. Highly expressing ALCR-GR transformants can display growth perturbation on induction due to “quenching”. 2. BY-2 ALCR-GR transformant lines must be transferred to liquid culture and propagated as a cell suspension. Lines expressing ALCR-GR can easily be selected for by RT-PCR.

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3. ALCR-GR transformant cell suspension lines are then secondarily transformed (as described above) with alcA-driven transgene constructs. Care should be taken to assay several (5–10) of the resultant secondary transformants for highly inducible expression of the alcA-driven transgene as some secondary transformants may display constitutive expression or no expression. 7.3 Protocol 3. High-Throughput Transformation of Arabidopsis Cell Culture 1. Dilute 3- to 7-day-old Arabidopsis Col-0 suspension culture 1:5 with fresh AT media. 2. Dispense 3−4 ml of the diluted suspension into six-well plates (Corning). 3. Add 1−3 mg of Agrobacteria from fresh selection plate by thick end of a sterile toothpick, in sterile conditions. 4. Seal the perimeter of the plate with Micropore tape (which allows aeration), and incubate in the dark at 25−27 ◦ C on a rotary shaker (150 rpm). 5. Observe expression 3–4 days after the transformation. 7.4 Protocol 4. Method for Systematic Removal of Stop Codons in GATEWAY Cloning 1. Design oligonucleotide primers to the vector sequences to amplify the gene of interest from the vector backbone, ensuring the C terminus of the protein is in frame with the GW reading frame. The 3
primer should be designed such that one of the base pairs in the stop codon of the gene of interest is mutated into a translated amino acid. This base pair and the surrounding three base pairs of the primer should be synthesised as phosphorothioate oligonucleotides. The 5
primer can be in any position close to the AUG translation start site. 2. Amplify gene in PCR with Platinum Pfx Hot-start polymerase (Invitrogen), using the following conditions: in a 50 µl reaction volume, use 1× buffer, 2× enhancer, 1 mM MgSO4 , 40 µM of each primer and 15 mM dNTP, and add 1 µl of Pfx polymerase. PCR regime: after initial step at 95 ◦ C for 5 min, do 35 cycles of 94 ◦ C for 15 s, 54 ◦ C for 30 s and 68 ◦ C for 1 min for each kilobase of template DNA, and the final extension step at 68 ◦ C for 10 min. 3. PEG precipitate the PCR product (Invitrogen, technical protocol) and resuspend in 10 µl of H2 O. Add 200−300 ng of purified PCR product (concentration determined by gel electrophoresis with appropriate quantitative DNA ladder) to 2µ of 5× BP buffer, 100 ng of entry vector and 1 µl of BP clonase (Invitrogen), and adjust total volume by H2 O to 10 µl. Leave reaction to proceed overnight at 20 ◦ C. Transform into DH5α strain of E. coli. 4. Assay colonies for the correct insert size by PCR amplification with the attL1 and attL2 primers. Inoculate correct colonies to liquid media and

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grow at 37 ◦ C overnight, then extract the plasmid DNA. Sequence the clones to ensure that the stop codon was mutated. 5. Transfer the insert from the entry clone via the LR GATEWAY recombination reaction to a suitable C-terminal fusion plant transformation vector. Use 100 ng of entry clone DNA, 100 ng of destination vector DNA, 1× LR buffer, 0.5 µl of Topoisomerase-1 and 1 µl of LR clonase (Invitrogen), and add TE buffer to 10 µl. Transform to a DH5α strain of E. coli. 6. Assay colonies with the attB1 and attB2 primers, by PCR amplification, to check for the correct insert size. Grow bacterial culture from the correct colony at 37 ◦ C overnight and extract the plasmid DNA. Transform to an appropriate Agrobacterium strain by electroporation.

8 Composition of Media 8.1 Liquid BY-2 Medium Composition: 4.3 g M&S salts media; 30 g sucrose; 100 mg inositol; 1 mg thiamine (1 ml, 1 mg/ml stock); 0.2 mg 2,4-D (20 µl, 10 mg/ml stock); 200 mg KH2 PO4 . Adjust to 1 litre with distilled water, pH 5.8 and dispense 100 ml per conical flask (volume 250 ml). Cover top and neck of the flask with thick aluminium foil, then autoclave. 8.2 Solid BY-2 Medium Composition: liquid BY-2 medium plus 0.4% Phytagel (Sigma); carbenicillin 500 µg/ml; Rovrol 20 µg/ml; and specific plasmid selection antibiotics, either kanamycin sulphate 200 µg/ml, hygromycin 41.6 µg/ml (Calbiochem) or spectinomycin added to the media before pouring 40 ml/plate. 8.3 AT Medium Composition: 4.4 g M&S salts media; 30 g sucrose; 0.05 mg kinetin; 0.5 mg NAA. Adjust to 1 litre with distilled water, pH 5.8 and dispense 50 ml per conical flask (volume 250 ml). Cover top and neck of the flask with thick aluminium foil, then autoclave.

References Bhat RA, Thompson RD (2004) The tobacco BY-2 cell line as a model system to understand in planta nuclear coactivator interactions. In: Nagata T, Hasezawa S, Inze D (eds) Tobacco BY-2

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Cells, Biotechnology in Agriculture and Forestry, vol 53. Springer, Berlin Heidelberg New York, pp 316–331 Chan J, Calder GM, Doonan JH, Lloyd CW (2003) EB1 reveals mobile microtubule nucleation sites in Arabidopsis. Nat Cell Biol 5:967–971 Chan J, Calder G, Fox S, Lloyd C (2005) Localization of the microtubule end binding protein EB1 reveals alternative pathways of spindle development in Arabidopsis suspension cells. Plant Cell 17:1737–1748 Cooper S (1998) On the interpretation of the shortening of the G1-phase by overexpression of cyclins in mammalian cells. Exp Cell Res 238:110–115 Deveaux Y, Peaucelle A, Roberts GR, Coen E, Simon R, Mizukami Y, Traas J, Murray JA, Doonan JH, Laufs P (2003) The ethanol switch: a tool for tissue-specific gene induction during plant development. Plant J 36:918–930 Dewitte W, Riou-Khamlichi C, Scofield S, Healy JM, Jacqmard A, Kilby NJ, Murray JA (2003) Altered cell cycle distribution, hyperplasia, and inhibited differentiation in Arabidopsis caused by the D-type cyclin CYCD3. Plant Cell 15:79–92 Escobar NM, Haupt S, Thow G, Boevink P, Chapman S, Oparka K (2003) High-throughput viral expression of cDNA-green fluorescent protein fusions reveals novel subcellular addresses and identifies unique proteins that interact with plasmodesmata. Plant Cell 15:1507–1523 Flipphi M, Mathieu M, Cirpus I, Panozzo C, Felenbok B (2001) Regulation of the aldehyde dehydrogenase gene (aldA) and its role in the control of the coinducer level necessary for induction of the ethanol utilization pathway in Aspergillus nidulans. J Biol Chem 276:6950– 6958 Gallois JL, Woodward C, Reddy GV, Sablowski R (2002) Combined SHOOT MERISTEMLESS and WUSCHEL trigger ectopic organogenesis in Arabidopsis. Development 129:3207–3217 Gomez-Mena C, de Folter S, Costa MM, Angenent GC, Sablowski R (2005) Transcriptional program controlled by the floral homeotic gene AGAMOUS during early organogenesis. Development 132:429–438 Henderson BR, Eleftheriou A (2000) A comparison of the activity, sequence specificity, and CRM1-dependence of different nuclear export signals. Exp Cell Res 256:213–224 Jackman M, Lindon C, Nigg EA, Pines J (2003) Active cyclin B1-Cdk1 first appears on centrosomes in prophase. Nat Cell Biol 5:143–148 Kiessling J, Kruse S, Rensing SA, Harter K, Decker EL, Reski R (2000) Visualization of a cytoskeleton-like FtsZ network in chloroplasts. J Cell Biol 151:945–950 Korolev AV, Chan J, Naldrett MJ, Doonan JH, Lloyd CW (2005) Identification of a novel family of 70 kDa microtubule-associated proteins in Arabidopsis cells. Plant J 42:547–555 Koroleva OA, Tomlinson M, Parinyapong P, Sakvarelidze L, Leader D, Shaw P, Doonan JH (2004) CycD1, a putative G1 cyclin from Antirrhinum majus, accelerates the cell cycle in cultured tobacco BY-2 cells by enhancing both G1/S entry and progression through S and G2 phases. Plant Cell 16:2364–2379 Koroleva OA, Tomlinson ML, Leader D, Shaw P, Doonan JH (2005) High-throughput protein localization in Arabidopsis using Agrobacterium-mediated transient expression of GFP-ORF fusions. Plant J 41:162–174 Kurreck J (2003) Antisense technologies. Improvement through novel chemical modifications. Eur J Biochem 270:1628–1644 Laufs P, Coen E, Kronenberger J, Traas J, Doonan J (2003) Separable roles of UFO during floral development revealed by conditional restoration of gene function. Development 130:785–796 Lutz KA, Corneille S, Azhagiri AK, Svab Z, Maliga P (2004) A novel approach to plastid transformation utilizes the phiC31 phage integrase. Plant J 37:906–913 Mao G, Chan J, Calder G, Doonan JH, Lloyd CW (2005) Modulated targeting of GFP-AtMAP65-1 to central spindle microtubules during division. Plant J 43:469–478 Mathur J, Szabados L, Schaefer S, Grunenberg B, Lossow A, Jonas-Straube E, Schell J, Koncz C, Koncz-Kalman Z (1998) Gene identification with sequenced T-DNA tags generated by transformation of Arabidopsis cell suspension. Plant J 13:707–716

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Menges M, Murray JA (2002) Synchronous Arabidopsis suspension cultures for analysis of cellcycle gene activity. Plant J 30:203–212 Nagata T (2004) When I encountered tobacco BY-2 cells! In: Nagata T, Hasezawa S, Inze D (eds) Tobacco BY-2 Cells, Biotechnology in Agriculture and Forestry, vol 53. Springer, Berlin Heidelberg New York, pp 1–5 Nagata T, Kumagai F (1999) Plant cell biology through the window of the highly synchronized tobacco BY-2 cell line. Methods Cell Sci 21:123–127 Nagata T, Oakada K, Takebe I (1982) Mitotic protoplasts and their interaction with tobacco mosaic virus RNA encapsulated in liposomes. Plant Cell Rep 1:250–252 Nagata T, Nemoto Y, Hasezawa S (1992) Tobacco BY-2 cell-line as the HeLa-cell in the cell biology of higher-plants. Int Rev Cytol 132:1–30 Oakenfull EA, Riou-Khamlichi C, Murray JA (2002) Plant D-type cyclins and the control of G1 progression. Philos Trans R Soc Lond B Biol Sci 357:749–760 Reski R, Cove DJ (2004) Physcomitrella patens. Curr Biol 14:R261–R262 Roberts GR, Garoosi GA, Koroleva O, Ito M, Laufs P, Leader DJ, Caddick MX, Doonan JH, Tomsett AB (2005) The alc-GR System. A modified alc gene switch designed for use in plant tissue culture. Plant Physiol 138:1259–1267 Roslan HA, Salter MG, Wood CD, White MR, Croft KP, Robson F, Coupland G, Doonan J, Laufs P, Tomsett AB, Caddick MX (2001) Characterization of the ethanol-inducible alc geneexpression system in Arabidopsis thaliana. Plant J 28:225–235 Sala F, Galli MG, Nielsen E, Magnien E, Devreux M, Pedrali-Noy G, Spadari S (1983) Synchronization of nuclear DNA synthesis in cultured Daucus carota L. cells by aphidicolin. FEBS Lett 153:204–208 Simpson JC, Wellenreuther R, Poustka A, Pepperkok R, Wiemann S (2000) Systematic subcellular localization of novel proteins identified by large-scale cDNA sequencing. EMBO Rep 1:287– 292 Sorrell DA, Combettes B, Chaubet-Gigot N, Gigot C, Murray JA (1999) Distinct cyclin D genes show mitotic accumulation or constant levels of transcripts in tobacco bright yellow-2 cells. Plant Physiol 119:343–352 Sorrell DA, Menges M, Healy JM, Deveaux Y, Amano C, Su Y, Nakagami H, Shinmyo A, Doonan JH, Sekine M, Murray JA (2001) Cell cycle regulation of cyclin-dependent kinases in tobacco cultivar Bright Yellow-2 cells. Plant Physiol 126:1214–1223 Tian GW, Mohanty A, Chary SN, Li S, Paap B, Drakakaki G, Kopec CD, Li J, Ehrhardt D, Jackson D, Rhee SY, Raikhel NV, Citovsky V (2004) High-throughput fluorescent tagging of full-length Arabidopsis gene products in planta. Plant Physiol 135:25–38 van der Fits L, Deakin EA, Hoge JH, Memelink J (2000) The ternary transformation system: constitutive virG on a compatible plasmid dramatically increases Agrobacterium-mediated plant transformation. Plant Mol Biol 43:495–502 Wagner D, Sablowski RW, Meyerowitz EM (1999) Transcriptional activation of APETALA1 by LEAFY. Science 285:582–584 Waring RB, May GS, Morris NR (1989) Characterization of an inducible expression system in Aspergillus nidulans using alcA and tubulin-coding genes. Gene 79:119–130 Yamada K, Lim J, Dale JM, Chen H, Shinn P, Palm CJ, Southwick AM, Wu HC, Kim C, Nguyen M, Pham P, Cheuk R, Karlin-Newmann G, Liu SX, Lam B, Sakano H, Wu T, Yu G, Miranda M, Quach HL, Tripp M, Chang CH, Lee JM, Toriumi M, Chan MM, Tang CC, Onodera CS, Deng JM, Akiyama K, Ansari Y, Arakawa T, Banh J, Banno F, Bowser L, Brooks S, Carninci P, Chao Q, Choy N, Enju A, Goldsmith AD, Gurjal M, Hansen NF, Hayashizaki Y, JohnsonHopson C, Hsuan VW, Iida K, Karnes M, Khan S, Koesema E, Ishida J, Jiang PX, Jones T, Kawai J, Kamiya A, Meyers C, Nakajima M, Narusaka M, Seki M, Sakurai T, Satou M, Tamse R, Vaysberg M, Wallender EK, Wong C, Yamamura Y, Yuan S, Shinozaki K, Davis RW, Theologis A, Ecker JR (2003) Empirical analysis of transcriptional activity in the Arabidopsis genome. Science 302:842–846

I.2 Dynamics and Structure of the Preprophase Band and the Phragmoplast D. Geelen1 and D. Inzé2

1 Introduction Plant cells feature unique cytoskeletal structures that contribute to their sessile lifestyle, the construction and maintenance of a rigid cell wall, and the structural organization of the plant body. A different cytoskeleton organization is adopted at different developmental or cell cycle stages. In particular microtubules are organized in a highly ordered fashion. The interphase cell carries a cortical array that looks like a spring captured inside the cell. The microtubules are predominantly arranged as parallel bundles that are perpendicular according to the longest axis of the cell. In small cells where cell polarity is poorly pronounced in terms of cell shape, the cortical microtubules and actin filaments are both randomly oriented. Small single cells and protoplasts in which the cortical array is disassembled are able to reconstitute a parallel arrangement of microtubules after passing through a phase of apparent disorganization. The information cues that are involved in arranging the cytoskeleton must therefore be embedded somewhere or somehow in each individual cell. Usually the cellulose microfibrils in the externally located cell wall are arranged in a fashion similar to that of the microtubules, and drugs that affect either one of them also change the organization of the other over time. This has led to the hypothesis that the orientation and organization of both structures are interrelated and somehow control one another. Because protoplasts have no cell wall, the reconstruction of the cytoskeleton cannot rely on the organization of fibrils outside the cell, suggesting that the microtubules perceive the primary information needed for their organization. External factors other than the cell wall influence the properties and organization of the cytoskeleton. In fact, the cytoskeleton responds to a wide diversity of biotic and abiotic factors that a plant encounters. In particular the plant hormones, auxins, gibberellins, and brassinosteroids are important because they complicate the analysis of cytoskeletonrelated phenomena when experiments are carried out in a whole plant context. The Bright Yellow-2 (BY-2) cells provide a suitable alternative system to study the relationship between microtubules and microfibrils. Virtually no BY-2 cells 1 Department of Plant Production, Faculty of Bioscience Engineering, Ghent University, Coupure

links 653, 9000 Gent, Belgium, e-mail: [email protected] of Plant Systems Biology, Flanders Interuniversity Institute for Biotechnology, Ghent University, Technologiepark 927, 9052 Gent, Belgium

2 Department

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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in the suspension show secondary wall thickenings, in contrast to Arabidopsis thaliana suspension cultures used in our laboratory that contain lignified cells with wall thickenings similar to those of xylem vessel cells. A thorough study of the structure of the cell wall in BY-2 cells, for example during different stages of development or throughout the cell cycle, is still lacking. The BY-2 cells are typically sausage shaped with a cortical microtubule array that is traverse to the longest axis, allowing predictions on the final orientation of the microtubules. Because BY-2 cells are very amenable to microscopic observation, over the last few years there has been an intensified use of BY-2 to study the structure and behaviour of microtubules. The BY-2 cells grow either individually or in different assemblies: some cells are grouped in files whereas others form small clumps of a handful of cells. Some variation in growth characteristics depending on the laboratory conditions or perhaps the origin of the culture has been noticed. The cells that are in files tend to be in a similar cell cycle phase and often nearly simultaneous cell divisions are observed, which is very handy when assessing the incidence of certain behavioural aspects of the cytoskeleton. Compared to cells that are in clumps, the cells that are in files offer good opportunities to find an area that is flat and large enough to monitor the dynamics of individual microtubules within the cortex. BY-2 cells are also approximately two-fold larger than, for instance, Arabidopsis suspension cells, allowing better observation of individual polymers both in interphase and in mitotis. The use of BY-2 cell cultures in combination with the application of green fluorescent protein (GFP) has greatly facilitated observation of the dynamic changes in the organization of the mitotic configurations. In this chapter we discuss the constituents and properties of two unique plant structures that have been intensely studied using BY-2 cells: the preprophase band (PPB) and the phragmoplast. Both structures are implicated in the final step of cell division, i. e. cytokinesis.

2 The Preprophase Band (PPB) Higher plant cells have a unique array of cortical microtubules that lie just underneath the plasma membrane (Hardham and Gunning 1978). In most differentiated cells that have expanded, the cortical microtubules are arranged as parallel bundles perpendicular to the longest axis of the cell. When somatic cells start to divide, the cortical array makes place for a band of microtubules that at first covers about two thirds of the peripheral area and gradually becomes more densely packed as the cell approaches the mitotic phase of the cell cycle. Before mitosis sets in, this PPB is destroyed in a matter of minutes. The orientation of the PPB usually parallels that of the cortical microtubules, so that cell division takes place in a plane perpendicular to the longest axis (Hofmeister 1863). Moreover, the cell divides in two halves of equal volume by

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splitting the mother cell across a plane with the smallest circumference (Errera 1888). Not every cell divides in the same way and the exceptions provide the means to create cellular differentiation and the development of different tissues. For example, the precursor of guard cells (the guard mother cell) separates across the longest axis so that two “elongated” cells emerge (Zhao and Sack 1999). Unequal wall thickenings lead to asymmetric expansion and formation of kidney-shaped guard cells with the stomatal pore in the centre. In earlier stages of stomata development meristemoid cells divide asymmetrically to produce subsidiary cells (Zhao and Sack 1999). In these instances, the PPB always forms at the position of the displaced nucleus and indicates the sites where the succeeding cell plate will join the parent walls (Pickett-Heaps 1969; Galatis and Mitrakos 1979). The PPB was visualized and described about 40 years ago by several groups (reviewed by Mineyuki 1999). Since then, based on correlative observations, many functions have been adhered to the PPB (Mineyuki 1999). Today the evidence that would substantiate the proposed functions is either lacking or scarce, leaving room for more exploration. Perhaps the most recurrent observation is that the position of the PPB corresponds to the division plane and the position where the cell plate inserts into the mother wall (Smith 2001). An important issue remains poorly understood: what mechanism controls the position of the PPB? A similar question is raised concerning the positioning of the acto-myosin ring. The correct placement of the contractile ring is mediated by the astral microtubules in animal cells and by the position of a previous division through the bud scar in budding yeast (reviewed by Guertin et al. 2002). The Mid1p protein is a key element involved in the central positioning of the cytokinetic ring in fission yeast (Paoletti and Chang 2000). This protein shuttles between the nucleus and the cortex to form a medial cortical band girdling the nucleus. The position of the nucleus is therefore instrumental to that of the cytokinetic ring. The situation in plants resembles that of fission yeast in that PPB positioning is also correlated with the position of the nucleus. The asymmetric division that preludes the subsidiary stomatal cells in the leaf epidermis (Geisler et al. 2003) or a more deeply positioned cell layer, such as the pericycle that has the capacity to produce side root primordia (Casimiro et al. 2003), involves nuclear movement toward the cell periphery. During standard divisions of vacuolated cells typically observed in BY-2 cultures, the nucleus moves from the periphery to the centre of the cell. Once settled in the centre it may provide the spatial cues for positioning of the PPB according to Errera’s rule. The movement of the nucleus to the cell centre is not inhibited by the DNA polymerase blocker aphidicolin, which halts cells in S-phase and suppresses PPB formation (Katsuta et al. 1990). Thus, the early migration of the nucleus does not require PPB microtubules. Once the PPB is established, the position of the nucleus is continuously adjusted to occupy the central spot of the PPB ring, which represents a second type of nuclear mobility. Nuclear movement can be an actin- or

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a microtubule-dependent process, indicating that different mobility processes occur independently from one another (Chytilova et al. 2000; Ketelaar et al. 2002; Van Bruaene et al. 2003). At this point it is not entirely clear how many mechanisms of nuclear migration exist and whether these depend on actin, microtubules or both. Because cytokinesis is temporally separate from the moment when the PPB occurs, it was postulated that a signpost is left behind that secures the cortical division site throughout mitosis. The identity of this molecule has remained enigmatic, and there is, for instance, no obvious homologue of the Mid1p found in plant sequence databases (Van Damme et al. 2004a). Interruption of PPB formation leads to misplaced cell plates, indicating that the PPB is prerequisite for the correct positioning of the division plane (Mineyuki et al. 1991a). The significance of the PPB for cell division was further substantiated by genetic evidence showing misplaced cell walls and aberrant cell shapes in tonneau Arabidopsis mutants (ton1 and ton2) that no longer produce a PPB (Traas et al. 1995). Because in tonneau and allelic mutants the organization of the cortical array is also disturbed, misplaced cell plates and abnormal cell shapes may not be a direct consequence of the lack of PPB formation (Torres-Ruiz and Jürgens 1994; McClinton and Sung 1997). Nevertheless, the PPB is not essential for mitosis as the ton/fass mutant cells maintain the capacity to divide. Division events that involve a PPB appear to be an adaptation for cells that are part of a complex multicellular architecture (Pickett-Heaps et al. 1999). PPBs are not characteristic for algae nor do they occur in sporophytic cells, or cells that are part of reproductive tissue such as the endosperm. In line with the idea that the PPB correlates with divisions in a tissue context it was noticed previously that some suspension-cultured cells divide without the need for a PPB (Mineyuki 1999). This has been substantiated for an Arabidopsis suspension culture that contains cells the majority of which do not form a PPB prior to division (Chan et al. 2005). The cells that divided without a PPB formed a bipolar spindle after nuclear envelope breakdown and showed variable spindle orientations and phragmoplast mobility. On the other hand, in those cells that did produce a PPB, a bipolar organization of the early spindle was established before nuclear envelope breakdown, with many more constraints for the positions taken by the spindle and the phragmoplast. This PPB-controlled restriction on the division plane may provide the means for neighbouring cells to control division planes in the context of a tissue or organ. Yet another deviation of the conventional organization of mitotic figures occurs in tobacco suspension cells that frequently produce abnormal PPBs (Hasezawa et al. 1994). The “conventional PPB” surrounds the nucleus which locates at the centre, flanked by stacked vacuoles at either end of the cell. In about one-fourth of the cells, either the PPB is displaced in relation to the nucleus or the PPB itself is aberrant. PPBs can be spiral shaped, partially split in two halves, or occurring as doublets (Granger and Cyr 2001). In cases where two PPBs are formed, either one of them is ignored during phragmoplast guidance, or alternatively some sort of combination of the two positions of

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either ring is used. Despite the fact that PPB may not be an essential structure for division, it does matter when it concerns cell plate positioning. This was elegantly demonstrated by centrifugation experiments of mitotic Tradescantia stamen hair cells that displaced the spindle away from its original position. The ensuing phragmoplast was shown to migrate back to the former place where the PPB was located. The process or mechanism by which PPB establishes the division site has hitherto remained elusive. Tangled is a candidate protein that may open up the quest for cortical division site markers (Smith et al. 2001). The tangled gene is required for correct cell plate alignment, but it is not needed for PPB formation (Cleary and Smith 1998). PPBs are still produced and the majority cross the cell at its shortest diameter. The misoriented cell plates found in tangled are mainly because spindle orientation is not controlled normally and the division plane and the spatial guidance of the phragmoplast are no longer adjusted according to the position formerly indicated by the PPB (Cleary and Smith 1998). To unveil some of the mysteries shrouding the origin and function of the PPB, attention has been given to the development of the PPB. For the time being, it is not known what triggers the onset of PPB formation. Normally, the PPB emerges during G2-phase when the cortical microtubules start to disappear. Taxol, a drug that impedes microtubule turnover, prevents PPB formation, indicating that the process depends on de novo microtubule synthesis in agreement with the incorporation of fluorescent tubulin (Cleary et al. 1992; Panteris et al. 1995). Protein synthesis is not required, as cycloheximide does not prevent the assembly of PPB microtubules (Mineyuki 1999), meaning that the degraded cortical microtubules are somehow recuperated for PPB microtubule growth. Translocation or sidewise sliding of microtubules has not been observed, suggesting that there must be an increase in turnover during cortical microtubule breakdown (Hush et al. 1994; Shaw et al. 2003; Vos et al. 2004). Dhonukshe and Gadella (2003) and Vos et al. (2004) have independently measured the dynamics of microtubules that locate at or near the equatorial plane prior to entry into mitosis, and compared that to microtubules elsewhere in the cortex. The data show changes in microtubule dynamics that would corroborate the transition from a cortical arrangement to a narrow band of microtubules. The growth and shrinkage rates of microtubules and the frequencies at which these two states alternate (catastrophe and rescue) determine the dynamics of the microtubules. During PPB formation, catastrophe frequency and the microtubule polymerization rate increase. Although the reported frequencies of catastrophe and rescue are controversial, both groups argue that the change in microtubule dynamics near the cortical area that surrounds the nucleus is responsible for PPB microtubule elongation and stabilization. The model proposed by Vos et al. (2004) also suggests an increase in the initiation of microtubules to account for the dramatic accumulation of microtubule bundles during the final stages of PPB formation. The de novo

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synthesis of microtubules was proposed to go hand in hand with a search and capture mechanism that stabilizes those microtubules contributing to the PPB. In plants, microtubules do not emanate from centrosomes and can indeed be initiated along existing microtubules at the cortex, although this is not a necessity (Shaw et al. 2003). Microtubule initiation has been observed using a GFP marker, microtubule end-binding protein 1 (EB1), that binds to both the plus and minus ends of microtubules (Chan et al. 2003, 2005). Despite the abscence of centrosomes in plants, some components such as γ -tubulin and the spindle pole body Spc98 from yeast (AtSpc98) are genetically conserved (Erhardt et al. 2002). The localization of γ -tubulin and AtSpc98 suggests that plant microtubules may be initiated not only at the cortex but also at the perinuclear area (Liu et al. 1993; Binarová et al. 2000; Erhardt et al. 2002). In fact, in vitro experiments have shown that microtubule initiation is highly abundant at the perinuclear area (Erhardt et al. 2002). The initiation of microtubules at the nuclear envelope, in particular during preprophase, produces microtubules that reach out to connect with the PPB, forming a structure reminiscent to spikes in the wheel of a bicycle. This suggests that a search and capture mechanism operates to stabilize those microtubules that bridge the nuclear envelope and the PPB. Microtubules that go “astray” may spend too much time reaching the end of a cell and therefore have an increased risk of undergoing catastrophy. Those that travel the shortest distance to the cell cortex survive and underpin the central position of the PPB microtubules or may even bend off to become inserted into the PPB itself. Microtubules initiated within the PPB could be similarly stabilized and thereby support an increase in microtubule density as PPB development progresses. Rather than contributing to formation of the PPB per se, the increase in catastrophe events in the area around the PPB could account for the disappearance of the cortical microtubules. The accumulation of PPB microtubules would be less affected because more microtubules are initiated at the position of the PPB and/or at the perinuclear area growing toward the cortical division site. Whether the change in microtubule dynamics suffices for the cortical array breakdown is not clear at this point. For instance it is possible that local activation of a microtubule-severing protein provoking internal cleavage of microtubules increases the number of unstable minus ends, with the net effect that microtubules depolymerize faster. The severing protein katanin is a heterodimer of an enzymatic p60 unit and a regulatory p80 unit (Stoppin-Mellet et al. 2002). A p60 homologue KSS1 has been identified in Arabidopsis that is involved in the destruction of perinuclear microtubules during the re-establishment of cortical microtubules directly after cell division (Burk et al. 2001). The parallel arrangement of the cortical network is lost in katanin mutants, but ultimately PPBs are still produced. Thus, KSS1 katanin is not essential for PPB formation. Katanin or other severing proteins may nevertheless influence the microtubule turnover rate to allow the liberation of sufficient tubulin molecules in due time, which are then

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incorporated into the newly synthesized microtubules of the PPB. To address this possibility, the dynamics of the cortical microtubules needs to be measured in the KSS1 mutants. Stabilizing proteins, such as MOR1, an ortholog of XMAP215, can also control the dynamic instability of microtubules (Whittington et al. 2001; Yasuhara et al. 2002; Hamada et al. 2004). At non-permissive temperature, the mor1 mutation causes shortening and randomization of the cortical microtubules (Whittington et al. 2001). Hence, destruction or modulation of MOR1 activity would promote the degradation of the cortical array. Xenopus XMAP215 and the human homologue TOG could be subject to phosphorylation by cyclindependent kinase CDK (Vasquez et al. 1999; Charrasse et al. 2000). The plant CDKA;1 concentrates at developing and at mature PPBs (Mineyuki et al. 1991b; Colasanti et al. 1993; Mews et al. 1997; Stals et al. 1997). The inhibition of kinases by broad-range effectors 6-dimethylaminopurin and staurosporin prevents PPB narrowing, suggesting that protein phosphorylation is required for microtubule bundling in the PPB (Nogami and Mineyuki 1999). MAP65 proteins are candidate microtubule-associated proteins (MAPs) that were shown to bundle microtubules in vitro and in vivo (Chan et al. 1999; Van Damme et al. 2004b). The MAP65-1 family members from tobacco and MAP65-1 and MAP65-5 from Arabidopsis bind the PPB (Smertenko et al. 2000; Van Damme et al. 2004b). In contrast, AtMAP65-3 and AtMap65-4 do not concentrate at the PPB microtubules, regardless of whether they are constitutively produced as GFP fusion proteins (Fig. 1). This indicates that secondary modifications or cell status-dependent protein stability controls their microtubule binding capacity. Potential phosphorylation sites occur in MAP65-1 and MAP65-5 and other members of the MAP65 protein family. These proteins are therefore putative candidates for bringing the PPB microtubules into closer proximity.

Fig. 1. Differential in vivo microtubule binding of AtMAP65-4-GFP. AtMAP65-4-GFP (left) concentrates specifically at the perinuclear microtubules at the beginning of prophase. As a general microtubule label, red fluorescent tubulin (TUA2-RFP) was co-expressed (middle). TUA2-RFP labels the perinuclear microtubules and the PPB and cortical microtubules in the neighbouring cell at the bottom of the image. Merged fluorescence is depicted at the right. Bar 10 µm

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A PPB matures when it reaches its most narrow width of approximately 2 µm, which is close to 10−20 min ahead of microtubule disassembly. GFP-tagged Medicago CDKA localizes in tobacco suspension cells to a very narrow band that may correspond to the matured PPB at the point just before breakdown (Weingartner et al. 2001). From earlier microinjection studies it had already been concluded that the CDK/cyclin B kinase complex causes disassembly of the PPB microtubules, while interphase cortical microtubules, spindle, and phragmoplast were left unaffected (Hush et al. 1996). Taken together, the data assign a dual function to CDKA activity in relation to PPB development: the maturation and compaction of the PPB microtubules and disintegration of the microtubules at the beginning of prophase. Thus, CDKA;1 may act on more than one PPB component. As MOR1 is associated with the PPB (Twell et al. 2002), it is a possible target for modulation by CKDA-mediated phosphorylation, and because of its microtubule-stabilizing function it may contribute to either PPB assembly or PPB disassembly. Pending the evidence for MOR1 and MAP65 phosphorylation, Aurora kinase may represent yet another player that adds to the complexity of microtubule dynamics in mitotic cells. Recently, three Aurora-like kinases were identified from Arabidopsis (Demidov et al. 2005; Kawabe et al. 2005). An antibody that recognizes the AtAurora1 and AtAurora2 homologues binds the PPB, and a similar conclusion was drawn from GFP-tagged AtAurora1 expressed in BY-2 cells (Fig. 2). Aurora kinases have essential functions in mitotic processes, such as chromosome condensation, spindle dynamics, kinetochore–microtubule interactions, and completion of cytokinesis (Carmena and Earnshaw 2003). There are three human Aurora kinases, classified as types A, B, and C (B and C types are actually very similar) according to their distribution and localization throughout mitosis (Adams et al. 2001). The localization patterns of plant Aurora kinases do not match those of Aurora A, B, or C. A new classification was proposed consisting of α and β classes, following the groups obtained by phylogenetic comparison (Demidov et al. 2005). BY-2 cells producing GFP-tagged Aurora1 show cytokinetic defects in line with the accumulation of the pro-

Fig. 2. AtAurora1-GFP produced in BY-2 cells. The collapsed three-dimensional image stack shows a cell prophase cell with a fluorescent band. AtAurora1-GFP accumulates in the ring and in the nucleus. Bar 10 µm

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tein in the cell plate (Van Damme et al. 2004a). Overexpression of GFP-tagged AtAurora3 leads to cell division defects more related to cell plate orientation and spindle defects, the latter in line with the localization of AtAurora3 at kinetochores (Kawabe et al. 2005). Actin also plays a role in division plane positioning, beginning at the transition from G1- to S-phase by mediating the migration of the nucleus from the periphery to the cell centre (Miyake et al. 1997). Later, actin filaments accumulate in the PPB and may help to narrow the ring of microtubules (Eleftheriou and Palevitz 1992). Whether actin directly affects the position where microtubules appear or whether the effect is indirect, via for instance the control of the position of the nucleus, is not clear. Actin filaments accumulate at the PPB and align with the microtubules (Kakimoto and Shibaoka 1987; Palevitz 1987; Traas et al. 1987; Schmit and Lambert 1990). When the PPB microtubules disintegrate, the actin band disappears in most systems but remains in others (Mineyuki 1999). In Tradescantia and BY-2 cells, upon breakdown of the PPB a region devoid of actin is formed called the actin-depleted zone (ADZ) (Cleary et al. 1992; Liu and Palevitz 1992). As the ADZ coincides with the PPB, it too marks the cortical division site. The ADZ is detected throughout metaphase and anaphase, suggesting that it contributes to a mechanism for positioning the new cell wall. However, the removal of the ADZ with actin drugs does not have a dramatic effect on the orientation of the cell plate, indicating that another signal is left behind that serves as a beacon for cell plate guidance (Hoshino et al. 2003). Our recent findings put forward the plasma membrane as a new player in marking the cortical division site (Vanstraelen et al. 2006). A GFP-tagged kinesin, GFP-KCA1, accumulates at the plasma membrane at the beginning of mitosis but is refrained from doing so at a narrow area surrounding the nucleus. At this position, the plasma membrane shows an actin depleted zone (Cleary et al. 1992; Liu and Palevitz 1992). Likewise, the plasma membrane does not accumulate GFP-KCA1 and hence it was called the KCA1 depleted zone (KDZ). The KDZ persists throughout mitosis and depends on the formation of a PPB. During mitosis, the KDZ is independent from the actin and microtubule cytoskeleton. Therefore it may provide spatial information to the phragmoplast and perhaps controls cell plate positioning independent from the actin network.

3 The Phragmoplast The dividing plant cell produces a new separating cell wall between the daughter nuclei in a way that looks very different from that which can be microscopically observed in animal or yeast cells. Plant cytokinesis, followed by differential interference contrast (DIC) microscopy, shows the deposition of refracting material at the equatorial plane, which gradually expands from

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the cell centre toward the cell periphery until it reaches the mother wall where it fuses. Not only does the process involve the formation of a new wall but also a new piece of plasma membrane is generated to seal off the two daughter cells. This implies that vesicular trafficking is a major constituent of cytokinesis just as in animal cytokinesis. In fact, plants have developed a mechanism for cytokinesis-specific vesicle fusion involving soluble N-ethylmaleimidesensitive fusion attachment protein receptor (SNARE) proteins. The system that accommodates for the homotypic fusion of vesicles at the equatorial plane is reviewed elsewhere (Jürgens 2005). Using improved preservation methods for electron microscopy, structural intermediates of the cell plate were resolved (Samuels et al. 1995). Golgi-derived vesicles are targeted to the equatorial plane, where they fuse into tube-like structures. The cell plate matures first in the middle, where vesicles and tubules fuse into a network (the tubulo-vesicular network or TNV), and gradually merges into a fenestrated sheet. The refractory material that we see in a DIC image contains callose detected in the TNV and fenestrated plate. The callose is not detected elsewhere and is produced after activation of callose synthase(s) in the maturing cell plate (Samuels et al. 1995; Hong et al. 2001). The callose synthase machinery is likely to originate from vesicles released by the Golgi, as blocking endoplasmic reticulum-Golgi transport, by means of brefeldin A, a potent inhibitor of ARF-GTPase that mediates vesicular coating and traffic, prevents cell plates from being completed in BY-2 (Yasuhara et al. 1995). A continuous supply of vesicles is therefore needed to finalize the plate. When the cell plate is completed, callose disappears gradually and is replaced by cellulose (Samuels et al. 1995). The precise timing of cellulose synthesis is not really clear but presumably occurs when finger-like protrusions of cell plate outer edge fuse to the plasma membrane so that cellulose synthase rosettes from the plasma membrane can migrate into the new plate. In live recordings, the emergence of cellulose may be announced by a clear stiffening of the cell plate. Cell plate development is accompanied by parallel-arranged microtubules and actin filaments formed in between the daughter nuclei. This mitotic configuration is called the phragmoplast. At its centre it carries the cell plate also referred to as the midline. The phragmoplast microtubules are two opposing bundles that point to each other with the plus end termini and that partially overlap (Euteneuer and McIntosh 1980). Remarkably, phragmoplast microtubules labelled with fluorescent tubulin and immunostained with antitubulin antibody or with GFP-tagged tubulin never show signal at the centre, as if no microtubules reach or are near the midline (Zhang et al. 1990; Smertenko et al. 2000; Granger and Cyr 2001; Hasezawa and Kumagai 2002). Detailed structural analysis of cryofixed phragmoplasts using electron tomography has revealed a cell plate assembly matrix (CPAM) (Segui-Simarro et al. 2004). The CPAM at the midline has previously been interpreted as dense material “hindering” the entry of fluorescent markers, leaving this region unstained. The digitalized and reconstructed tomographic images show no

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abundant overlap of the phragmoplast microtubules as is commonly accepted. Perhaps the overlapping of the microtubules is a temporal phenomenon and occurs during establishment of the bipolar organization of the phragmoplast. Later, most microtubules would then more likely contribute to the transport to and from the cell plate. A candidate for the transport of Golgi-derived vesicles is AtPAKRP2, a membrane-associated kinesin that accumulates at the midline (Lee et al. 2001). There are two distinct phases in the development of the cell plate that are clearly distinguished by the behaviour of the microtubules. The first step is the preparation of a bundle of microtubules that emerges at the mid zone of the spindle. In the beginning the microtubules are arranged as two discs and later new microtubules appear at the rim of the phragmoplast which outwardly expands. The second phase is marked by the depolymerization of the central microtubules which may supply the tubulin dimers needed for the microtubules initiated at the outer borders. The plate matures first at the centre by closing the “windows” in the fenestrated sheet. Some gaps remain through which microtubules pass for the formation of plasmodesmata. The phragmoplast-localized kinesin Hinkel/AtNack1 is required for the depolymerization of the microtubules in the centre and mutations in hinkel are cytokinesis defective due to an arrest in phragmoplast expansion (Strompen et al. 2002; Tanaka et al. 2004). As the central microtubules are no longer decomposed, the pool of free tubulin drops, limiting the supply for further outward growth of the phragmoplast. Phragmoplast expansion is controlled by a mitogen-activated protein kinase (MAPK) signalling cascade; MAPKKK binds the kinesin Hinkel/AtNack1 and is targeted by means of this interaction to the midline (Nishihama et al. 2002). The MAPK signalling pathway probably modifies microtubule-associated proteins that regulate microtubule stability. At the expanding edge the overlapping microtubules should be stabilized in order to maintain the integrity of phragmoplast bipolarity. Additional kinesins have been identified that associate with the phragmoplast. Carrot DcTKRP120 is a BimC-type kinesin that strongly accumulates at the midline (Barroso et al. 2000). It may have a function similar to TKRP125 from tobacco, the first BimC-type kinesin described in plants that is required for the outward translocation of phragmoplast microtubules (Asada et al. 1997). Although TKRP125 particularly concentrates at the spindle midzone, it was proposed that it contributes to the outward translocation process in the phragmoplast by sliding the antiparallel-arranged microtubules using its plus-end directional mobility (Asada et al. 1997). Both in the spindle and in the midbody, the cytokinetic apparatus in animal cells, bipolarity is maintained by BimC-type kinesins (reviewed by Liu and Lee 2001). A similar function is anticipated for BimC-type kinesins in plants, perhaps with TKRP125 more related to the anaphase spindle and DcTKRP120 to the phragmoplast. Besides kinesins, other MAPs contribute to the formation and/or stabilization of the phragmoplast (reviewed by Otegui et al. 2005). For instance, the MAP65 proteins are a small family of nine members. MAP65 proteins show

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similarity to PRC1/Ase1p/Feo/SPD-1 which localizes to the spindle midzone and is required for spindle integrity (Otegui et al. 2005). MAP65-3 is mutated in pleiade Arabidopsis mutants that show severe cytokinesis defects in roots (Müller et al. 2004). The roots cells affected become multinuclear presumably because phragmoplasts are not held properly together at the midline and the opposing microtubule discs appear to be further apart. The AtMAP65-3 protein has capacity to bind microtubules and strongly accumulates at the midline, where it presumably forms cross-bridges between the opposing phragmoplast microtubules (Smertenko et al. 2000; Van Damme et al. 2004a). MAP65 members bundle microtubules in vitro and in vivo (Chan et al. 1999; Van Damme et al. 2004b; Wicker-Planquart et al. 2004). AtMAP65-1, AtMAP65-5, and AtMAP65-8 associate with the phragmoplast microtubules as well (Van Damme et al. 2004a). These MAPs do not bind the same microtubule subdomains and concentrate along the entire length of the phragmoplast microtubules, or in the case of AtMAP65-8 near to the nucleus where supposedly the minus ends congregate. In addition to subregional specificity, the MAP65 proteins show a differential behaviour toward the different microtubules that occur throughout mitosis. The spindle, for instance, is devoid of AtMAP65-1-GFP whereas the same protein strongly binds the phragmoplast (Fig. 3). Because AtMAP65-1-GFP also binds the cortical array and is continuously present, some secondary modification must take place to modulate its microtubule-binding activity. A phragmoplast component that has received little attention is actin. Several drug studies have pointed to a role for actin in the organization, expansion, and integrity of the phragmoplast. In the presence of actin drugs such as latrunculin B and bistheonellide A, the phragmoplast does not disintegrate and there are no major effects on its position in the cell. However, the disturbance of actin slows down phragmoplast expansion and new microtubules assemble to initiate the formation of additional phragmoplasts (Yoneda et al. 2004). The actin filaments made during phragmoplast initiation are shorter and more densely packed than at the cortical network (Schmit and Lambert 1990; Endle et al. 1998). Formin stimulates the nucleation of actin and interacts with the barbed ends of the filaments. Intense nucleation of actin is anticipated, as two of the 20 Arabidopsis formin homologues AtFH5 and AtFH6 are abundant at the midline (Van Damme et al. 2004a; Ingouff et al. 2005). The inactivation of AtFH5 leads to a delayed cellularization of the endosperm presumably because phragmoplast formation is slowed down. Therefore actin and actin nucleation at the midzone seem to be important for early events during cytokinesis. Several actin-related protein (arp2/3) homologues have been cloned recently but so far their relationship to cytokinesis has not been reported (Mathur 2005).

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Fig. 3. Time-lapse recording of BY-2 cells expressing AtMAP65-1-GFP. Three cells that are dividing synchronously were recorded from metaphase until the end of cytokinesis. Spindle microtubules are not labelled. In contrast, AtMAP65-1-GFP strongly accumulates at phragmoplast microtubules. In addition, AtMAP65-1-GFP accumulates in dots, a phenomenon also observed in interphase cells (Van Damme et al. 2004b). Bar 10 µm

4 Future Perspectives The study of the plant cytoskeleton enters a new era, with advanced microscopy and the use of GFP markers as the most important leading technologies. Tomographic analysis of complex configurations such as the PPB and the phragmoplast provides insight into the fine structure and the three-dimensional organization of the different parts. The technology is not (yet) widely applied, mainly because of the substantial financial cost to run such an electron microscopy facility. An important advantage of the technology is that it can be applied to Arabidopsis wild type and mutants with various cell division defects.

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The use of GFP markers, on the other hand, resolves details of the dynamics of different structures such as microtubules, actin filaments, and vesicles or similar structures. During recent years, in addition to the classic microtubule (microtubulebinding domain (MBD)-GFP) and actin (Talin-GFP) markers, many more proteins have been introduced that specifically label different cell division structures. The future challenge is to combine the differential subcellular localization patterns of these proteins with the three-dimensional models built from structural analysis. Integration of these datasets will allow development of dynamic three-dimensional scenarios for plant mitosis.

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Katsuta J, Hashiguchi Y, Shibaoka H (1990) The role of the cytoskeleton in positioning of the nucleus in premitotic tobacco BY-2 cells. J Cell Sci 95:413–422 Kawabe A, Matsunaga S, Nakagawa K, Kurihara D, Yoneda A, Hasezawa S, Uchiyama S, Fukui K (2005) Characterization of plant Aurora kinases during mitosis. Plant Mol Biol 58:1–13 Ketelaar T, Faivre-Moskalenko C, Esseling JJ, de Ruijter NCA, Grierson CS, Dogterom M, Emons AMC (2002) Positioning of nuclei in Arabidopsis root hairs: an actin-regulated process of tip growth. Plant Cell 14:2941–2955 Lee Y-RJ, Giang HM, Liu B (2001) A novel plant kinesin-related protein specifically associates with the phragmoplast organelles. Plant Cell 13:2427–2439 Liu B, Lee YRJ (2001) Kinesin-related proteins in plant cytokinesis. J Plant Growth Regul 20:141–150 Liu B, Palevitz BA (1992) Organization of cortical microfilaments in dividing root cells. Cell Motil Cytoskeleton 23:252–264 Liu B, Marc J, Joshi HC, Palevitz BA (1993) A γ -tubulin-related protein associated with the microtubule arrays of higher plants in a cell cycle-dependent manner. J Cell Sci 104:1217– 1228 Mathur J (2005) The ARP2/3 complex: giving plant cells a leading edge. BioEssays 27:377–387 McClinton RS, Sung ZR (1997) Organization of cortical microtubules at the plasma membrane in Arabidopsis. Planta 201:252–260 Mews M, Sek FJ, Moore R, Volkmann D, Gunning BES, John PCL (1997) Mitotic cyclin distribution during maize cell division: implications for the sequence diversity and function of cyclins in plants. Protoplasma 200:128–145 Mineyuki Y (1999) The preprophase band of microtubules: its function as a cytokinetic apparatus in higher plants. Int Rev Cytol 187:1–50 Mineyuki Y, Murata T, Wada M (1991a) Experimental obliteration of the preprophase band alters the sites of cell division, cell plate orientation and phragmoplast expansion in Adiantum protonemata. J Cell Sci 100:551–557 Mineyuki Y, Yamashita M, Nagahama Y (1991b) p34cdc2 kinase homologue in the preprophase band. Protoplasma 162:182–186 Miyake T, Hasezawa S, Nagata T (1997) Role of cytoskeletal components in the migration of nuclei during the cell cycle transition from G1 phase of tobacco BY-2 cells. J Plant Physiol 150:528–536 Müller S, Smertenko A, Wagner V, Heinrich M, Hussey PJ, Hauser M-T (2004) The plant microtubule-associated protein AtMAP65-3/PLE is essential for cytokinetic phragmoplast function. Curr Biol 14:412–417 Nishihama R, Soyano T, Ishikawa M, Araki S, Tanaka H, Asada T, Irie K, Ito M, Terada M, Banno H, Yamazaki Y, Machida Y (2002) Expansion of the cell plate in plant cytokinesis requires a kinesin-like protein/MAPKKK complex. Cell 109:87–99 Nogami A, Mineyuki Y (1999) Loosening of a preprophase band of microtubules in onion (Allium cepa L.) root tip cells by kinase inhibitors. Cell Struct Funct 24:419–424 Otegui MS, Verbrugghe KJ, Skop AR (2005) Midbodies and phragmoplasts: analogous structures involved in cytokinesis. Trends Cell Biol 15(8) Palevitz BA (1987) Actin in the preprophase band of Allium cepa. J Cell Biol 104:1515–1519 Panteris E, Apostolakos P, Galatis B (1995) The effect of taxol on Triticum preprophase root cell: preprophase microtubule band organization seems to depend on new microtubule assembly. Protoplasma 186:72–78 Paoletti A, Chang F (2000) Analysis of mid1p, a protein required for placement of the cell division site, reveals a link between the nucleus and the cell surface in fission yeast. Mol Biol Cell 11:2757–2773 Pickett-Heaps JD (1969) Preprophase microtubules and stomatal differentiation; some effects of centrifugation on symmetrical and asymmetrical cell division. J Ultrastruct Res 27:24–44 Pickett-Heaps JD, Gunning BES, Brown RC, Lemmon BE, Cleary AL (1999) The cytoplast concept in dividing plant cells: cytoplasmic domains and the evolution of spatially organized cell division. Am J Bot 86:153–172

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Samuels AL, Giddings TH Jr, Staehelin LA (1995) Cytokinesis in tobacco BY-2 and root tip cells: a new model of cell plate formation in higher plants. J Cell Biol 130:1345–1357 Schmit A-C, Lambert A-M (1990) Microinjected fluorescent phalloidin in vivo reveals the F-actin dynamics and assembly in higher plant mitotic cells. Plant Cell 2:129–138 Segui-Simarro JM, Austin II JR, White EA, Staehelin LA (2004) Electron tomographic analysis of somatic cell plate formation in meristematic cells of Arabidopsis preserved by high-pressure freezing. Plant Cell 16:836–856 Shaw SL, Kamyar R, Ehrhardt DW (2003) Sustained microtubule treadmilling in Arabidopsis cortical arrays. Science 300:1715–1718 Smertenko A, Saleh N, Igarashi H, Mori H, Hauser-Hahn I, Jiang C-J, Sonobe S, Lloyd CW, Hussey PJ (2000) A new class of microtubule-associated proteins in plants. Nat Cell Biol 2:750–753 Smith LG (2001) Plant cell division: building walls in the right places. Nat Rev Mol Cell Biol 2:33–39 Smith LG, Gerttula SM, Han S, Levy J (2001) Tangled1: a microtubule binding protein required for the spatial control of cytokinesis in maize. J Cell Biol 152:231–236 Stals H, Bauwens S, Traas J, Van Montagu M, Engler G, Inzé D (1997) Plant CDC2 is not only targeted to the pre-prophase band, but also co-localizes with the spindle, phragmoplast, and chromosomes. FEBS Lett 418:229–234 Stoppin-Mellet V, Gaillard J, Vantard M (2002) Functional evidence for in vitro microtubule severing by the plant katanin homologue. Biochem J 365:337–342 Strompen G, El Kasmi F, Richter S, Lukowitz W, Assaad FF, Jürgens G, Mayer U (2002) The Arabidopsis HINKEL gene encodes a kinesin-related protein involved in cytokinesis and is expressed in a cell cycle-dependent manner. Curr Biol 12:153–158 Tanaka H, Ishikawa M, Kitamura S, Takahashi Y, Soyano T, Machida C, Machida Y (2004) The AtNACK1/HINKEL and STUD/TETRASPORE/AtNACK2 genes, which encode functionally redundant kinesins, are essential for cytokinesis in Arabidopsis. Genes Cells 9:1199–1211 Torres-Ruiz RA, Jürgens G (1994) Mutations in the FASS gene uncouple pattern formation and morphogenesis in Arabidopsis development. Development 120:2967–2978 Traas J, Bellini C, Nacry P, Kronenberger J, Bouchez D, Caboche M (1995) Normal differentiation patterns in plants lacking microtubular preprophase bands. Nature 375:676–677 Traas JA, Doonan JH, Rawlins DJ, Shaw PJ, Watts J, Lloyd CW (1987) An actin network is present in the cytoplasm throughout the cell cycle of carrot cells and associates with the dividing nucleus. J Cell Biol 105:387–395 Twell D, Park SK, Hawkins TJ, Schubert D, Schmidt R, Smertenko A, Hussey PJ (2002) MOR1/GEM1 has an essential role in the plant-specific cytokinetic phragmoplast. Nat Cell Biol 4:711–714 Van Bruaene N, Joss G, Thas O, Van Oostveldt P (2003) Four-dimensional imaging and computerassisted track analysis of nuclear migration in root hairs of Arabidopsis thaliana. J Microsc 211:167–178 Van Damme D, Bouget F-Y, Van Poucke K, Inzé D, Geelen D (2004a) Molecular dissection of plant cytokinesis and phragmoplast structure: a survey of GFP-tagged proteins. Plant J 40:386–398 Van Damme D, Van Poucke K, Boutant E, Ritzenthaler C, Inzé D, Geelen D (2004b) In vivo dynamics and differential microtubule-binding activities of MAP65 proteins. Plant Physiol 136:3956–3967 Vanstraelen M, Van Damme D, De Rycke R, Mylle E, Inzé D, Geelen D (2006) Cell cycle-dependent targeting of a kinesin at the plasma membrane demarcates the division site in plant. Curr Biol 16:308–314 Vasquez RJ, Gard DL, Cassimeris L (1999) Phosphorylation by CDK1 regulates XMAP215 function in vitro. Cell Motil Cytoskeleton 43:310–321 Vos JW, Dogterom M, Emons AMC (2004) Microtubules become more dynamic but not shorter during preprophase band formation: a possible “search-and-capture” mechanism for microtubule translocation. Cell Motil Cytoskeleton 57:246–258

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Weingartner M, Binarova P, Drykova D, Schweighofer A, David J-P, Heberle-Bors E, Doonan J, Bögre L (2001) Dynamic recruitment of cdc2 to specific microtubule structures during mitosis. Plant Cell 13:1929–1943 Whittington AT, Vugrek O, Wei KJ, Hasenbein NG, Sugimoto K, Rashbrooke MC, Wasteneys GO (2001) MOR1 is essential for organizing cortical microtubules in plants. Nature 411:610–613 Wicker-Planquart C, Stoppin-Mellet V, Blanchoin L, Vantard M (2004) Interactions of tobacco microtubule-associated protein MAP65-1b with microtubules. Plant J 39:126–134 Yasuhara H, Sonobe S, Shibaoka H (1995) Effects of brefeldin A on the formation of the cell plate in tobacco BY-2 cells. Eur J Cell Biol 66:274–281 Yasuhara H, Muraoka M, Shogaki H, Mori H, Sonobe S (2002) TMBP200, a microtubule bundling polypeptide isolated from telophase tobacco BY-2 cells is a MOR1 homologue. Plant Cell Physiol 43:595–603 Yoneda A, Akatsuka M, Kumagai F, Hasezawa S (2004) Disruption of actin microfilaments causes cortical microtubule disorganization and extra-phragmoplast formation at M/G1 interface in synchronized tobacco cells. Plant Cell Physiol 45:761–769 Zhang D, Wadsworth P, Hepler PK (1990) Microtubule dynamics in living dividing plant cells: confocal imaging of microinjected fluorescent brain tubulin. Proc Natl Acad Sci USA 87:8820– 8824 Zhao L, Sack FD (1999) Ultrastructure of stomatal development in Arabidopsis (Brassicaceae) leaves. Am J Bot 86:929–939

I.3 Formation of Cortical Microtubules in a Cell-Free System Prepared from Plasma Membrane Ghosts and a Cytosolic Extract of BY-2 Cells T. Murata and M. Hasebe1

1 Introduction Microtubules are involved in polarized cell expansion and cell division in green plants (Lloyd 1991). At interphase, a transverse cortical array of microtubules develops beneath the plasma membrane (Lloyd and Chan 2004). The microtubule array is widely believed to regulate the direction of cell expansion via the deposition of oriented cellulose microfibrils in the cell walls. During the initial stage of cell division, a preprophase band of microtubules develops at the site where the cell plate fuses to the parental cell wall at cytokinesis, and the band disappears during the development of a mitotic spindle, before the cell plate develops (for review see Mineyuki 1999). A perinuclear array of microtubules that radiates from the nuclear envelope is evident when the preprophase band forms, and this array is later reorganized into the mitotic spindle. During the succeeding course of cell division, the mitotic spindle segregates the chromosomes and the phragmoplast contributes to the construction of a cell plate that divides the two daughter cells. Microtubule nucleation is a prerequisite for the organization of microtubule arrays, and knowledge of this process is indispensable in understanding the molecular mechanisms of the organization of the subsequent array. A cell-free system prepared using tobacco BY-2 cells has greatly contributed to the elucidation of the nucleation mechanisms. In this chapter we review the mechanisms of microtubule nucleation, with special emphasis on this cell-free system.

2 Microtubules Nucleate as Branches on Existing Microtubules in the Cortical Arrays of Plant Cells 2.1 Microtubule Organizing Centers in Plant Cells Microtubules, which are polymers of αβ-tubulin heterodimers, are nucleated at centrosomes in animal cells and spindle pole bodies in fungal cells, sites known as microtubule organizing centers (MTOCs) (Job et al. 2003). γ -tubulin ring protein complexes (γ TuRCs), which contain γ -tubulin and Spc98 family 1 National

Institute for Basic Biology, Myodaiji-cho, Okazaki, Aichi, 444-8585 Japan, e-mail: [email protected]

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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proteins, localize predominantly to the MTOCs and are essential for microtubule formation. Biochemical and electron microscopic analyses of animal γ TuRCs have shown that one γ TuRC nucleates a single microtubule (Zheng et al. 1995) and is incorporated into the minus end of the newly formed microtubule (Moritz et al. 1995). Although the details of the molecular mechanisms of microtubule nucleation are still unclear, a γ TuRC clearly acts as a nucleus for αβ-tubulin heterodimer assembly (Job et al. 2003). Tubulin proteins are conserved among eukaryotes (Oakley 2000). In flowering plants, a fraction of γ -tubulin is a component of a large (> 1,500 kDa) protein complex (Stoppin-Mellet et al. 2000). The large complex has microtubule nucleation activity (Drykova et al. 2003), suggesting that it is a plant γ TuRC. Further characterization of the complex might reveal whether it contains Spc98 family proteins. A conserved function for γ -tubulins among plant, animal, and fungal cells has also been suggested by the expression of plant γ -tubulin in fission yeast cells, which revealed that Arabidopsis thaliana γ tubulin can nucleate microtubules at the spindle pole body in fission yeast (Horio and Oakley 2003). In flowering plants, microtubule arrays change during the progression of the cell cycle, and MTOCs are inconspicuous or absent. The microtubule nucleation sites in the microtubule arrays have been analyzed, and the role of γ -tubulin in microtubule formation in perinuclear arrays has been well characterized. The nuclear envelope has been proposed to be a microtubule nucleation site (Lambert 1993). In tobacco BY-2 cells, microtubules radiate from the nuclear envelope during the S and G2 phases (Hasezawa and Nagata 1991). Isolated nuclei are capable of microtubule nucleation, and both γ -tubulin and an ortholog of Spc98 localize at the nuclear envelope in isolated nuclei. Antibodies against γ -tubulin and the Spc98 ortholog inhibit microtubule nucleation, indicating that these proteins are essential for microtubule nucleation at the nuclear envelope (Erhardt et al. 2002). In contrast, the origins of microtubules in other microtubule arrays are not well known (Wasteneys 2002). Unexpectedly, γ -tubulin was found to localize along the length of microtubules, rather than at their ends in the spindle, the phragmoplast (Liu et al. 1993; Ovechkina and Oakley 2001), or the cortical array (Hoffman et al. 1994; Ovechkina and Oakley 2001). However, the γ -tubulin signal in the cortical array is unclear in some cell types (Liu et al. 1993; Dibbayawan et al. 2001), including tobacco BY-2 cells (Erhardt et al. 2002; Kumagai et al. 2003), probably because of interference from cytoplasmic signals. Clearly, γ -tubulin is not a good minus end marker of microtubules in these arrays, and GFP-tagged AtEB1a, a microtubule end-binding protein, has been used to label the minus ends instead of γ -tubulin (Chan et al. 2003). The role of γ -tubulin on the sides of microtubules was unknown. It was assumed that such γ -tubulin does not nucleate microtubules, and other roles, such as stabilization of microtubules, were expected (Joshi and Palevitz 1996; Shimamura et al. 2004). Alternatively, it was predicted that γ -tubulin nucleated on the sides of microtubules in the spindle (Smirnova and Bajer 1994) or the cortical

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arrays (Wasteneys 2002). Although this prediction is supported by the patterns of microtubule reassembly after drug-induced microtubule depolymerization of the cortical arrays (Falconer et al. 1988; Wasteneys and Williamson 1989), the idea of microtubule nucleation on existing microtubules was controversial until its direct visualization using live imaging. 2.2 Microtubule Nucleation Visualized by Live Imaging The live imaging of individual microtubules constituted a breakthrough in the study of the microtubule nucleation sites in cortical arrays. Shaw and coworkers visualized the initiation of individual microtubules in the cortical cytoplasm of Arabidopsis thaliana hypocotyl cells expressing YFP-α-tubulin (Shaw et al. 2003). Microtubules initiated as branches on pre-existing microtubules, although some microtubules initiated at sites without any related structures (Shaw et al. 2003). Later, Murata et al. (2005) showed that most of the latter microtubules also originate from pre-existing microtubules. Using quantitative analysis in tobacco BY-2 cells and Arabidopsis thaliana leaf cells expressing GFP-α-tubulin, they observed that most microtubules were nucleated as branches on pre-existing microtubules. Microtubules originating from pre-existing microtubules are likely released from each initiation site by microtubule-severing proteins (Shaw et al. 2003). The pre-existing microtubules shrink after the initiation of a new microtubule. Considering the depolymerization of the original microtubules, the microtubule-independent nucleation sites in A. thaliana are probably formed by the depolymerization of the original microtubules from microtubule-dependent nucleation sites. The microtubule-dependent microtubule nucleation may explain the reestablishment of cortical arrays following the completion of cytokinesis. In BY-2 cells, microtubules elongate from the nuclear envelope after the disappearance of the phragmoplast. The growing ends of the perinuclear microtubules attach to the cell cortex, and the first cortical microtubules appear at the attachment sites (Kumagai et al. 2001). The first cortical microtubules may be initiated from the nuclear microtubules at the attachment sites. Based on the localization of γ -tubulin in the cytoplasm and the detection at the cell surface of spot-like structures of AtSPC98-GFP expressed in tobacco BY-2 cells, Erhardt et al. (2002) proposed the recruitment of cytoplasmic γ TuRC into the cortical cytoplasm. The recruitment of the nucleation sites is supported by the movement of AtEB1-GFP, a protein used as a marker for microtubule nucleation, in the cortical cytoplasm of cultured A. thaliana cells (Chan et al. 2003). It is still unclear whether endogenous AtEB1 proteins localize in the microtubule nucleation sites of the cortical array. The biochemical characterization of microtubule nucleation components is needed to clarify the molecular mechanisms of the recruitment of microtubule nucleation sites.

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3 Analysis of the Molecular Mechanisms of Microtubule Nucleation in a BY-2 Cell-Free System 3.1 Development of a Cell-Free System for Microtubule Assembly Nucleation and the subsequent organization of microtubules in animal centrosomes have been studied using a system involving Xenopus laevis eggs. Cytoplasmic extracts from the eggs support the assembly of centrosomes that are capable of microtubule nucleation around sperm centrioles that cannot nucleate microtubules (Stearns and Kirschner 1994). The recruitment of γ -tubulin to the centriole is essential for microtubule nucleation in this system. When a mitotic extract is used, the nucleated microtubules reorganize into spindles. Thus, the system has been used for the analysis of spindle assembly, and several factors essential for spindle assembly have been characterized using immunodepletion techniques (Walczak et al. 1996; Tournebize et al. 2000). In green plants, Sonobe and Takahashi (1994) reported increases in microtubules on plasma membrane ghosts upon the addition of both the ghosts (Fig. 1) and an extract derived from tobacco BY-2 cells. We improved the experimental system (Fig. 2) and demonstrated that microtubule assembly depends on existing microtubules (Murata et al. 2005). Upon the addition of the extract to the ghosts, microtubule assembly on the ghosts begins within 2 min, and clusters of microtubules have formed after 10 min at room temperature (23–27 ◦ C). In living cells, newly assembled microtubules have been observed to initiate as branches on pre-existing microtubules. Newly polymerized microtubules can be distinguished from pre-existing microtubules by the incorporation of rhodamine-labeled tubulin. Although microtubule assembly in the extract during in vitro incubation has been reported, no microtubule formation has been detected in samples lacking a template plasma membrane.

Fig. 1. Plasma membrane ghosts isolated from tobacco BY-2 cells labeled with anti-α-tubulin. Scale bar 10 µm

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Fig. 2. Schematic representation of a cell-free system for assaying microtubule assembly on plasma membrane ghosts. Protoplasts isolated from BY-2 cells are evacuolated by centrifugation in Percoll solution, gently homogenized, and centrifuged at 39,000 × g for 20 min. High-speed supernatant by the centrifugation is referred as a cytosolic extract. Plasma membrane ghosts are prepared by attaching protoplasts to a poly-l-lysine-coated cover slip and shaking them in a buffer. The resulting plasma membrane ghosts are incubated in the high-speed supernatant, and the microtubules on the ghosts are assayed by immunofluorescence microscopy

3.2 Role of γ -Tubulin in Microtubule Assembly on Plasma Membrane Ghosts Immunodepletion, which involves the depletion of specific proteins from extracts using antibody beads, is a powerful tool in the functional analysis of proteins that are present in cytosolic extracts. Because cytosolic extracts obtained from evacuolated protoplasts (miniprotoplasts) of BY-2 cells contain little proteolytic activity (Sonobe 1996), the technique is applicable to the cell-free system prepared using the extracts. As described above, microtubule nucleation in the cell-free system requires the presence of microtubules. Because the role of microtubule-bound γ -tubulin in microtubule nucleation is still controversial (see Sect. 2.1), we examined the involvement of γ -tubulin in microtubule-dependent microtubule nucleation in the cell-free system using immunodepletion (Murata et al. 2005), as described below. Using an antibody raised against tobacco γ -tubulin, we observed the redistribution of γ -tubulin during the assembly of microtubules in the cell-free system. During microtubule assembly, microtubules formed at the sites of mi-

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crotubules containing γ -tubulin. Therefore, it is likely that γ -tubulin that is bound to the sides of existing microtubules nucleates new microtubules. Supporting this idea, the addition of the anti-tobacco-γ -tubulin antibody inhibited microtubule formation. To separate γ -tubulin recruitment and microtubule nucleation, oryzalin or propyzamide, an inhibitor of αβ-tubulin heterodimer assembly, is added to the extract. Under these conditions, γ -tubulin is recruited to microtubules on the ghosts, but no assembly of microtubules occurs. Microtubules can nucleate as branches after the removal of the extracts, followed by the addition of purified αβ-tubulin (Fig. 3). Because αβ-tubulin heterodimers cannot assemble in the presence of oryzalin or propyzamide, these results rule out the possibility that very short microtubules form in the extract lacking plasma membrane ghosts, and attach onto microtubules on ghost plasma membranes. Microtubule nucleation on the sides of existing microtubules is inhibited by the immunodepletion of γ -tubulin from the extracts (Fig. 3), indicating that γ -tubulin recruited from the extract to the sides of microtubules nucleates new microtubules. The roles of microtubule-bound γ -tubulin in the mitotic spindle and phragmoplast are still unknown. Further improvement of the system will be necessary for its use with spindles and phragmoplasts.

Fig. 3. Reconstitution of microtubule-branching activity on plasma membrane ghosts (Murata et al. 2005). Microtubules on buffer-treated ghosts cannot nucleate microtubules (top). Incubation of the ghost microtubules with a cytosolic extract that contains a tubulin polymerization inhibitor confers microtubule nucleation ability that does not require the polymerization of microtubules during the incubation. Many microtubules nucleate as branches on the extract-treated ghost microtubules upon incubation with purified tubulin (middle). The effect of the cytoplasmic extract is diminished by the depletion of γ -tubulin from the extract (bottom)

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4 Future Perspectives Microtubule nucleation on pre-existing microtubules in the form of branches is unique to plants, although microtubule nucleation through anti-parallel bundle formation has recently been observed in fission yeast (Janson et al. 2005). Characterization of the proteins involved in microtubule branching in both lineages will provide insight into the evolutionary aspects of microtubule nucleation, especially with respect to the origins of the very different nucleation systems in the animal and plant lineages. The characterization of the γ -tubulin binding proteins in green plants may be a good first step. Several homologues of Spc98 family proteins have been identified in plant genomes (T. Murata and M. Hasebe, unpublished data), and genetic, biochemical, and cell biological approaches will be needed for their characterization. Additional genome information and resources from tobacco BY-2 cells should facilitate the isolation and characterization of plant γ -tubulin binding proteins. The molecular mechanisms involved in the reorganization of microtubules after microtubule nucleation are mostly unknown (Lloyd and Chan 2004), and the BY-2 cell-free system will be useful when it comes to addressing the problem in a more sophisticated manner. ATP is necessary for the microtubule depolymerization performed by severing proteins and the microtubule reorientation performed by motor proteins (Hartman et al. 1998; Desai et al. 1999; Asbury 2005). The addition of cytoplasmic extracts containing ATP to the BY-2 ghosts led to microtubule depolymerization (Sonobe 1990), and further conditional adjustments will make possible the reorganization of newly formed microtubules, facilitating the isolation of the factors involved in these processes using immunodepletion experiments.

References Asbury CL (2005) Kinesin: world’s tiniest biped. Curr Opin Cell Biol 17:89–97 Chan J, Calder GM, Doonan JH, Lloyd CW (2003) EB1 reveals mobile microtubule nucleation sites in Arabidopsis. Nat Cell Biol 5:967–971 Desai A, Verma S, Mitchison TJ, Walczak CE (1999) Kin I kinesins are microtubule-destabilizing enzymes. Cell 96:69–78 Dibbayawan TP, Harper JDI, Marc J (2001) A γ -tubulin antibody against a plant peptide sequence localises to cell division-specific microtubule arrays and organelles in plants. Micron 32:671–678 Drykova D, Cenklova V, Sulimenko V, Volc J, Draber P, Binarova P (2003) Plant γ -tubulin interacts with αβ-tubulin dimers and forms membrane-associated complexes. Plant Cell 15:465–480 Erhardt M, Stoppin-Mellet V, Campagne S, Canaday J, Mutterer J, Fabian T, Sauter M, Muller T, Peter C, Lambert AM, Schmit AC (2002) The plant Spc98p homologue colocalizes with γ -tubulin at microtubule nucleation sites and is required for microtubule nucleation. J Cell Sci 115:2423–2431 Falconer MM, Donaldson G, Seagull RW (1988) MTOCs in higher-plant cells – an immunofluorescent study of microtubule assembly sites following depolymerization by APM. Protoplasma 144:46–55

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Hartman JJ, Mahr J, McNally K, Okawa K, Iwamatsu A, Thomas S, Cheesman S, Heuser J, Vale RD, McNally FJ (1998) Katanin, a microtubule-severing protein, is a novel AAA ATPase that targets to the centrosome using a WD40-containing subunit. Cell 93:277–287 Hasezawa S, Nagata T (1991) Dynamic organization of plant microtubules at the three distinct transition points during the cell cycle progression of synchronized tobacco BY-2 cells. Bot Acta 104:206–211 Hoffman JC, Vaughn KC, Joshi HC (1994) Structural and immunocytochemical characterization of microtubule-organizing centers in pteridophyte spermatogenous cells. Protoplasma 179:46–60 Horio T, Oakley BR (2003) Expression of Arabidopsis γ -tubulin in fission yeast reveals conserved and novel functions of γ -tubulin. Plant Physiol 133:1926–1934 Janson ME, Setty TG, Paoletti A, Tran PT (2005) Efficient formation of bipolar microtubule bundles requires microtubule-bound gamma-tubulin complexes. J Cell Biol 169:297–308 Job D, Valiron O, Oakley B (2003) Microtubule nucleation. Curr Opin Cell Biol 15:111–117 Joshi HC, Palevitz BA (1996) γ -Tubulin and microtubule organization in plants. Trends Cell Biol 6:41–44 Kumagai F, Yoneda A, Tomida T, Sano T, Nagata T, Hasezawa S (2001) Fate of nascent microtubules organized at the M/G1 interface, as visualized by synchronized tobacco BY-2 cells stably expressing GFP-tubulin: time-sequence observations of the reorganization of cortical microtubules in living plant cells. Plant Cell Physiol 42:723–732 Kumagai F, Nagata T, Yahara N, Moriyama Y, Horio T, Naoi K, Hashimoto T, Murata T, Hasezawa S (2003) γ -tubulin distribution during cortical microtubule reorganization at the M/G1 interface in tobacco BY-2 cells. Eur J Cell Biol 82:43–51 Lambert AM (1993) Microtubule-organizing centers in higher plants. Curr Opin Cell Biol 5:116– 122 Liu B, Marc J, Joshi HC, Palevitz BA (1993) A gamma-tubulin-related protein associated with the microtubule arrays of higher plants in a cell cycle-dependent manner. J Cell Sci 104:1217–1228 Lloyd C, Chan J (2004) Microtubules and the shape of plants to come. Nat Rev Mol Cell Biol 5:13–22 Lloyd CW (1991) The cytoskeletal basis of plant growth and form. Academic Press, London. Mineyuki Y (1999) The preprophase band of microtubules: its function as a cytokinetic apparatus in higher plants. Int Rev Cytol 187:1–49 Moritz M, Braunfeld MB, Sedat JW, Alberts B, Agard DA (1995) Microtubule nucleation by gamma-tubulin-containing rings in the centrosome. Nature 378:638–640 Murata T, Sonobe S, Baskin TI, Hyodo S, Hasezawa S, Nagata T, Horio T, Hasebe M (2005) Microtubule-dependent microtubule nucleation based on recruitment of γ -tubulin in higher plants. Nat Cell Biol 7:961–968 Oakley BR (2000) An abundance of tubulins. Trends Cell Biol 10:537–542 Ovechkina Y, Oakley BR (2001) Gamma tubulin in plant cells. Method Cell Biol 67:195–212 Shaw SL, Kamyar R, Ehrhardt DW (2003) Sustained microtubule treadmilling in Arabidopsis cortical arrays. Science 300:1715–1718 Shimamura M, Brown RC, Lemmon BE, Akashi T, Mizuno K, Nishihara N, Tomizawa K, Yoshimoto K, Deguchi H, Hosoya H, Horio T, Mineyuki Y (2004) Gamma-tubulin in basal land plants: characterization, localization, and implication in the evolution of acentriolar microtubule organizing centers. Plant Cell 16:45–59 Smirnova EA, Bajer AS (1994) Microtubule converging centers and reorganization of the interphase cytoskeleton and the mitotic spindle in higher plant Haemanthus. Cell Motil Cytoskel 27:219–233 Sonobe S (1990) ATP-dependent depolymerization of cortical microtubules by an extract in tobacco BY-2 Cells. Plant Cell Physiol 31:1147–1153 Sonobe S (1996) Studies on the plant cytoskeleton using miniprotoplasts of tobacco BY-2 cells. J Plant Res 109:437–448 Sonobe S, Takahashi S (1994) Association of microtubules with the plasma-membrane of tobacco BY-2 cells in vitro. Plant Cell Physiol 35:451–460

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Stearns T, Kirschner M (1994) In vitro reconstitution of centrosome assembly and function: the central role of gamma-tubulin. Cell 76:623–637 Stoppin-Mellet V, Peter C, Lambert AM (2000) Distribution of gamma-tubulin in higher plant cells: cytosolic gamma-tubulin is part of high molecular weight complexes. Plant Biol 2:290–296 Tournebize R, Popov A, Kinoshita K, Ashford AJ, Rybina S, Pozniakovsky A, Mayer TU, Walczak CE, Karsenti E, Hyman AA (2000) Control of microtubule dynamics by the antagonistic activities of XMAP215 and XKCM1 in Xenopus egg extracts. Nat Cell Biol 2:13–19 Walczak CE, Mitchison TJ, Desai A (1996) XKCM1: a Xenopus kinesin-related protein that regulates microtubule dynamics during mitotic spindle assembly. Cell 84:37–47 Wasteneys GO (2002) Microtubule organization in the green kingdom: chaos or self-order? J Cell Sci 115:1345–1354 Wasteneys GO, Williamson RE (1989) Reassembly of microtubules in Nitella tasmanica – assembly of cortical microtubules in branching clusters and its relevance to steady-state microtubule assembly. J Cell Sci 93:705–714 Zheng Y, Wong ML, Alberts B, Mitchison T (1995) Nucleation of microtubule assembly by a γ -tubulin-containing ring complex. Nature 378:578–583

I.4 Chromosome Dynamics in Tobacco BY-2 Cultured Cells S. Matsunaga1 , N. Ohmido2 , and K. Fukui1

1 Introduction During mitosis in eukaryotes, the nuclear genome is organized into highly condensed structures referred to as chromosomes. The dynamic processes of chromosome condensation and segregation play crucial roles in the equal separation of genetic information to both daughter cells. Since disruption of these processes is harmful to cells, causing, for example, chromosome aneuploidy, appropriate regulation is indispensable. Chromosome dynamics involves modification of the molecules that regulate these processes during mitosis. The usefulness of tobacco BY-2 (Nicotiana tabacum cv. Bright Yellow-2) cultured cells in analyses of chromosome dynamics has been recently demonstrated. Tobacco BY-2 cells possess the following advantages. Their mitotic chromosomes are not small and monocentric, which is advantageous because analyses of chromosome dynamics are generally performed under a microscope. The haploid genome size of the amphidiploid species Nicotiana tabacum (2n = 4x = 48) is about 4,500 Mb (Arumuganathan and Earle 1991) and the total nuclear genome is divided into 48 mitotic chromosomes. The length of these mitotic chromosomes varies from 2 to 6 µm (Kenton et al. 1993; Moscone et al. 1996), which is larger than that in Arabidopsis thaliana (c.a. 2 µm). Such chromosome size makes it possible to analyze the detailed morphology and structure of chromosomes. Although BY-2 cultured cells are derived from N. tabacum, it is probable that a certain level of ploidy change and chromosome abnormality occurs during cell culture. However, any problem encountered in studies of chromosome dynamics has not been reported to date. Another advantage of tobacco BY-2 cells is their ability for high synchronization. Because mitotic chromosomes only emerge in the short mitotic phase, synchronous cell systems are very useful for studying chromosome dynamics. A highly synchronous method has been established in tobacco BY-2 cells (Nagata et al. 1992) and the usefulness of synchronized BY-2 cells has been widely demonstrated in plant cell cycle studies (Ito 2000). Synchronized BY-2 cells provide the most efficient mitotic chromosomes because a mitotic index of more 1 Department

of Biotechnology, Graduate School of Engineering, Osaka University, 2-1 Yamadaoka, Suita 565-0871, Osaka, Japan, e-mail: [email protected] 2 Faculty of Human Development, Kobe University, 3-11 Tsurukabuto, Nada, Kobe 657-8501, Hyogo, Japan Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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than 90% is attained through sequential synchronization with aphidicolin and propyzamide (Nagata and Kumagai 1999). Recently, the green fluorescent protein (GFP) has become the most powerful and popular tool in plant cell biological analyses. Dynamic analyses of proteins fused with GFP allow direct in vivo observations of proteins or organelles in living plant cells. This application has also been used for imaging chromosomes and nuclei in living cells. We recently conducted analyses of GFP-fused proteins involved in chromosome dynamics using tobacco BY-2 cells.

2 Dynamic Analysis of Condensin Complexes Condensin complexes are highly conserved among eukaryotes (Hirano 2002). Condensin I is composed of five subunits: two core subunits (CAP-E/SMC2 and CAP-C/SMC4) (where CAP is chromosome associated protein and SMC is structural maintenance of chromosome) which belong to the structural maintenance of chromosomes (SMC) family, and three non-SMC subunits (CAP-D2, CAP-G, and CAP-H) (Losada and Hirano 2005; Table 1). SMC family proteins participate in a number of processes involved in chromosome dynamics (Hirano 2002; Jessberger 2002; Hagstrom and Meyer 2003). CAP-D2 and CAP-G share a highly degenerate, repeating motif known as the HEAT repeat (Neuwald and Hirano 2000) and CAP-H belongs to the kleisin superfamily (Schleiffer et al. 2003). Several studies have revealed that condensin subunits are essential for in vitro and in vivo chromosome condensation and segregation (Hirano and Mitchison 1994; Hirano et al. 1997; Hagstrom et al. 2002; Wignall et al. 2003). Another condensin complex (condensin II) has been identified in HeLa cells (Ono et al. 2003). Condensin II shares the same pair of SMC subunits with condensin I; however, it contains a different set of non-SMC subunits (CAP-D3, CAP-G2, and CAP-H2). Moreover, although condensin II is also localized on Table 1. Subunits of condensin complexes S. cerevisiae H. sapiens A. thaliana SMC2 (I and II) Smc4

hCAP-C

AtCAP-C

SMC4 (I and II) Smc2

hCAP-E

AtCAP-E1, E2

HEAT (IA)

Ycs4

hCAP-D2 CAB72176

HEAT (IB)

Ycs5/Ycg1

hCAP-G

BAB08309 AtCAP-H

Kleisin (IC)

Brn1

hCAP-H

HEAT (IIA)

–

hCAP-D3 At4g15890

HEAT (IIB)

–

hCAP-G2 At1g64960

Kleisin (IIC)

–

hCAP-H2 AtCAP-H2

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mitotic chromosomes, condensin I and II show distinct distributions along the axis of a single chromosome (Ono et al. 2004). Condensin II and I associate with chromosomes sequentially in this order and have different functions with respect to mitotic chromosome assembly (Hirota et al. 2004). A few plant-related studies have revealed that SMC2 subunits are involved in development in A. thaliana (Liu et al. 2002; Siddiqui et al. 2003). A. thaliana contains two SMC genes, AtCAP-E1 and AtCAP-E2, with AtCAP-E1 comprising more than 85% of the total SMC2 transcripts. Embryo lethality caused by double mutation of SMC genes suggests that AtCAP-E1 and AtCAP-E2 are involved in development in A. thaliana. Recently, dynamic analyses of the non-SMC subunits of Arabidopsis condensins, AtCAP-H and AtCAP-H2A, were performed using tobacco BY-2 cells (Fujimoto et al. 2005). Figure 1 shows the subcellular localization of GFPAtCAP-H in tobacco BY-2 cells at different stages. In interphase, GFP signals were predominantly localized in the cytoplasm of cells transformed with GFPAtCAP-H. GFP-AtCAP-H was found in the cytoplasm until the end of prophase. In prometaphase, some signals were detected on the chromosomes, while in metaphase, almost all signals moved to the chromosomes and the signal intensity rapidly increased. After chromosome segregation, a few signals diffused into the cytoplasm but the main signal remained on the chromosomes. The chromosome signals finally diffused to the cytoplasm after cytokinesis. The dynamics of GFP-AtCAP-H2 during mitosis of tobacco BY-2 cells is shown in Fig. 2. In interphase, GFP signals were mainly detected in the nucleolus and slightly in the nucleoplasm. Signals were localized in the nucleolus until the end of prophase, moving mainly to the entire chromosomes after disappearance of the nucleolus. The signal intensity on the chromosomes was weaker than that of AtCAP-H. During chromosome segregation, the signals were equally localized on both chromatids, and when the nucleolus reformed in the nucleus, signals once again appeared here. AtCAP-H and AtCAP-H2 are localized in mitotic chromosomes from prometaphase to telophase. However, their localizations during interphase are different; although AtCAP-H is localized in the cytoplasm, AtCAP-H2 is localized in the nucleus, particularly in the nucleolus. These different localizations are indicative of functional differentiation between condensin I and II in A. thaliana.

3 Dynamic Analysis of Heterochromatic Protein 1 Heterochromatin corresponds to the relatively gene-poor, late replication, repetitive sequences found near the centromere and telomere. In contrast, euchromatin replicates relatively early in the cell cycle and contains low-copy DNA sequences, including the majority of genes. Heterochromatin and euchromatin show differences in heterochromatin protein 1 (HP1), a highly conserved non-histone chromosomal protein, between yeast and plants and functionally

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Fig. 2. Subcellular localization pattern of GFP-AtCAP-H2 in tobacco BY-2 cells. Upper row shows DNA staining with DAPI and lower GFP fluorescence. Cells at a, h interphase, b, i prophase, c, j prometaphase, d, k metaphase, e, l anaphase, f, m telophase and g, n cytokinesis. Scale bar; 10 µm

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Fig. 3. Subcellular localization pattern of GFP-AtLHP1 in tobacco BY-2 cells. Upper row shows DNA staining with DAPI and lower GFP fluorescence. Cells at a, g interphase, b, h prophase, c, i prometaphase, d, j metaphase, e, k anaphase and f, l telophase. Scale bar; 10 µm

Chromosome Dynamics in Tobacco BY-2 Cultured Cells

Fig. 1. Subcellular localization pattern of GFP-AtCAP-H in tobacco BY-2 cells. The upper row shows DNA staining with DAPI (4
, 6-diamidino-2-phenylindole) and the lower shows GFP fluorescence. Cells at a, h interphase, b, i prophase, c, j prometaphase, d, k metaphase, e, l anaphase, f, m telophase and g, n cytokinesis. Scale bar; 10 µm

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are involved in chromatin packing and gene silencing (Li et al. 2002). HP1 interacts with the modifier protein of heterochromatin, inducing position effect variegation (PEV), which is encoded by the Su(var)2-5 gene in Drosophila (James and Elgin 1986). The HP1 family is composed of relatively small proteins (15−35 kDa). They have a chromodomain (CD) in the highly conserved N-terminal region, a chromo shadow domain (CSD) in the C-terminal region, and a variable length linker (hinge region) (Eissenberg and Elgin 2000). CENP-A, a centromeric-specific protein, together with HP1 is involved in the association and segregation of sister kinetochores during mitosis in animal cells (Sugimoto et al. 2001). In plants, the lhp-1 mutant in Arabidopsis exhibits alterations in leaf and flower organs, cell size, and flowering time transitions (Gaudin et al. 2001). To obtain insight into the formation and maintenance of heterochromatin, we investigated kinetic binding of GFP-tagged Arabidopsis Like Heterochromatin Protein 1 (AtLHP1 in tobacco BY-2 cells. AtLHP1 dynamically changed its localization during mitosis (Fig. 3). GFPAtLHP1 foci were detected in interphase nuclei. They then diffused in the cytoplasm during late prophase to anaphase and relocalized on chromatin in telophase. The molecular heterochromatic regions indicated by the localization of AtLHP1 were mostly limited to centromeric regions, although the heterochromatic region is not sharply delimited on tobacco metaphase chromosomes. Post-transcriptional histone tail modification including methylation and acetylation is also involved in heterochromatin formation (Jackson et al. 2002). Especially, methylation of lysine 9 on Histone 3 by Su(var)3-9 methyltransferase in Drosophila is linked to gene silencing and heterochromatin formation. Colocalization between HP1 and chromatin histone modification was also shown in plant cells (Yu et al. 2004). Chromosome preparations for immunostaining of Met-H3K9 (methylated lysine 9 of histone H3) in GFP-AtLHP1 transformed cells were carried out according to Hasezawa and Kumagai (2002). The antibody against tri-Met-H3K9 and the secondary antibody, anti-rabbit Cy3, were used for immunostaining detection in the BY-2 cells. As a result, the signals of GFP-AtLHP1 and Met-H3K9 were shown to be colocalized in interphase cells. However, AtLHP1 dynamic behavior during chromosome segregation revealed that the foci do not always localize in concert with Met-H3K9. GFP-AtLHP1 localized at specific foci in interphase nuclei, but most GFP-AtLHP1 diffused into the cytoplasm from prophase to anaphase and relocalized at the Met-H3K9 sites in telophase. These results are similar to those obtained with human and mouse HP1, with release from chromatin to the cytoplasm during prophase to anaphase (Sugimoto et al. 2001). With proceeding cytokinesis, HP1 was predominantly found in the newly formed daughter nuclei, again displaying a heterochromatinlike distribution. These results suggested that, although the majority of HP1 diffuses in the cytoplasm, some populations are retained in the centromeric region and are involved in the association and segregation of sister kinetochores during mitosis (Sugimoto et al. 2002). This is consistent with plant cells,

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in which a dynamic change of HP1 on chromosomes is essential for precise association and segregation of chromosomes. These regions are dominated by highly trimethylated histone K9 of histone H3, suggesting formation of the heterochromatin center. AtLHP1 is also thought to be necessary for plant cell division, and histone modification by methylation is believed to play an essential role as a landmark for heterochromatin formation in plant cells.

4 Dynamic Analysis of Aurora Kinases Aurora kinases belong to the cell-cycle-dependent serine/threonine protein kinase family that regulates several mitotic events (Carmena and Earnshaw 2003). The paralog numbers of Aurora kinases are different among organisms. Yeast has only one Aurora kinase gene in its genome, while animal species have two, identified as Aurora A and B. Mammalian species, such as humans and mice, have an additional Aurora kinase designated as Aurora C, which is specifically expressed in the testis. The Aurora kinase family shows a high sequence similarity of more than 60% at the amino acid level, particularly in the kinase domain; however, Aurora kinases differ in their localization and function. Aurora A is localized at the centrosomes during interphase and at the spindle poles and mitotic spindle during mitosis. It is essential for centrosome maturation, maintenance, duplication, and segregation, in addition to stabilization of spindle microtubules during mitosis. Moreover, Aurora A phosphorylates centrosome- and mitotic spindle-related proteins. Aurora B localizes at the centromeres from prophase to metaphase and relocates to the spindle midbody at cytokinesis. Aurora B is a chromosomal passenger protein that interacts with INCENP, Survivin, and Borealin/Dasra B to form a chromosomal passenger complex (Vagnarelli and Earnshaw 2004). These proteins are necessary for localization of Aurora B for regulation of kinetochore formation, chromosome segregation, and cytokinesis. Phosphorylation of histone H3 at Ser10 and Ser28 by Aurora B is considered to be the fundamental role of Aurora B in chromosome segregation and cytokinesis (Goto et al. 2002). Although direct phosphorylation was not reported, the condensin complexes, which are involved in chromosome condensation, could not be localized on chromosomes in Aurora B-depleted cells, resulting in incorrect chromosome condensation (Ono et al. 2003). Aurora B phosphorylates topoisomerase II and ISWI (MacCallum et al. 2002; Morrison et al. 2002), indicating that Aurora B has a role for chromatin organization. Recently, two studies identified plant Aurora kinases in A. thaliana (Demidov et al. 2005; Kawabe et al. 2005). In the A. thaliana genome, three deduced amino acid sequences showed high similarity to those of animal Aurora kinase genes. The kinase domain of these three proteins shows more than 60% similarity to those of animal and yeast

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Aurora kinases. These three genes were designated AtAUR1(A. thaliana Aurora kinase)/AtAurora1, AtAUR2/AtAurora2, and AtAUR3/AtAurora3 (Demidov et al. 2005; Kawabe et al. 2005). The AtAUR1 and AtAUR2 sequences shows high similarities with the amino acid sequences of their kinase domains, exhibiting 95% sequence identity. In contrast, the kinase domain of AtAUR3 shares a 65% amino acid sequence identity with the other two AtAURs. Similar to animal Aurora kinases, the N-terminal regions in plants are variable in both length and sequence. Recently, dynamic analysis of GFP-fused AtAUR proteins was performed using tobacco BY-2 cells (Kawabe et al. 2005). Figure 4 shows dynamic analysis of AtAUR1 during mitosis in tobacco BY-2 cells. AtAUR1 was located on the nuclear membrane at interphase. At prophase, it moved toward peripheral regions of the nucleus near the spindle poles, resembling a cap-like distribution. At prometaphase, microtubules began to project from both poles, forming a mitotic spindle, and the localization of AtAUR1 appeared as fibers. At metaphase, localization appeared on the spindle-like structure. From anaphase to telophase, AtAUR1 was located in the spindle halves moving toward the spindle poles. As the mitotic spindle moved to opposite sides, the signal accumulated in separated spindles. During telophase, AtAUR1 clearly localized in the midzone between the signals in the peripheral region of the two cell nuclei, suggesting localization on the synthesized cell plate. Although localization of AtAUR2 in tobacco BY-2 cells showed almost the same pattern as that of AtAUR1, no significant AtAUR2 signals could be detected around the cell plate in the midzone at telophase. Recently, localization of AtAUR1 in the equatorial plate was also reported (Van Damme et al. 2004). When the mitotic chromosomes began to decondense, AtAUR1 and AtAUR2 gradually returned to the peripheral region of the cell nuclei. Although the N-terminal regions of AtAUR1 and AtAUR2 show almost no homology, their subcellular localizations are similar, except during cell plate formation. Animal Aurora A kinases localize in the centrosome during interphase, showing strong signals at both spindle poles (Kimura et al. 1997; Sugimoto et al. 2002). Consistent with its localization, Aurora A regulates the stabilization and maturation of centrosome and spindle poles (Giet et al. 1999). Animal cells have centrosomes and fungi have a spindle pole body, while plants have no centrosomes but microtubule organizing centers (MTOCs) in their nuclear membrane during interphase (Staiger and Lloyd 1991). The localizations near the nuclear membrane at interphase, cap-like localization at prophase, and spindle-like localization at metaphase resemble the localizations of γ tubulin, a component of plant MTOCs. This fact suggests that AtAUR1 and AtAUR2 colocalize with γ -tubulin and function in MTOCs. Figure 5 shows dynamic analysis of AtAUR3 in tobacco BY-2 cells. AtAUR3 localized at the nuclear periphery during interphase. At prophase, dot-like signals appeared when the chromosomes began to condense. At prometaphase, the signals moved to the metaphase plates along with the condensed chromosomes. At metaphase, the signals aligned in the center of the metaphase

Fig. 5. Localization pattern of AtAUR3 during mitosis of tobacco BY-2 cells. AtAUR3-GFPoverexpressed BY-2 cells were fixed and stained with DAPI. a–f DAPI; g–l GFP. Cells at a, g interphase, b, h prophase, c, i prometaphase, d, j metaphase, e, k anaphase, f, l telophase. Scale bar; 10 µm

Chromosome Dynamics in Tobacco BY-2 Cultured Cells

Fig. 4. Localization pattern of AtAUR1 during mitosis of tobacco BY-2 cells. AtAUR1-GFPoverexpressed BY-2 cells were fixed and stained with DAPI. a–f DAPI; g–l GFP. Cells at a, g interphase, b, h prophase, c, i prometaphase, d, j metaphase, e, k anaphase, f, l telophase. CW Cell wall (a, b); MT microtubule (i); SP mitotic spindle (j, k); SC sister chromatid (e); CP cell plate (l). Scale bar; 10 µm

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plates. However, at anaphase, in accordance with chromosome segregation, AtAUR3 signals were almost evenly observed on the entire chromosome. After cell division, the AtAUR3 signals returned to the nuclear membrane and the cytoplasm around the nucleus. The major substrate of Aurora kinases is histone H3 (Adams et al. 2001). In the late G2 phase, phosphorylation of histone H3 at Ser10 is initiated in pericentromeric heterochromatin, spreading throughout the chromosomes by metaphase, with dephosphorylation occurring after anaphase (Hendzel et al. 1997). In plants, the distribution pattern of phosphorylated histone H3 at Ser10 differs from that of monocentric and polycentric chromosomes. In tobacco BY2 cells, when chromosomes began to condense at prophase, dot-like signals were first detected in the pericentric regions (Kawabe et al. 2005). The dotted signals on the mitotic chromosomes then moved to the metaphase plate and, after segregation to opposite poles was complete, they dispersed along the sister chromatids at late anaphase. At telophase, the signals drastically reduced and then disappeared. The localization of AtAUR3 during mitosis was the same as that of phosphorylated histone H3 at Ser10. All three recombinant AtAURs can phosphorylate histone H3 at Ser10 in vitro (Demidov et al. 2005; Kawabe et al. 2005), strongly suggesting that AtAUR3 plays a major role in phosphorylation of histone H3 at Ser10 in vivo. Dot-like signals on the chromosomes between prophase and metaphase have also been reported in the case of yeast Aurora kinases and animal Aurora B kinases (Adams et al. 2001; Giet and Glover 2001; Murata-Hori et al. 2002). These patterns were shown to be located on the centromeres, suggesting that Aurora B regulates the formation and cohesion of kinetochores in animals (Kaitna et al. 2002; Murata-Hori et al. 2002). AtAUR3 also located along the metaphase plate, strongly suggesting that AtAUR3 is localized on centromeres during prophase and metaphase. When AtAUR3 is limited at the early mitotic phase, its localization pattern is similar to that of Aurora B. However, after metaphase, AtAUR3 is evenly located on the chromosomes, while animal Aurora B remains at the metaphase plate to regulate cytokinesis (Kaitna et al. 2000; Adams et al. 2001).

5 Conclusion and Perspectives BY-2 cultured cells are suitable for analyses of chromosome dynamics during mitosis because of their chromosome size and short cell cycle. The growth rate and morphology of each transformed BY-2 cell line were almost the same as the original BY-2 cells. Specifically, BY-2 cells are one of the most suitable materials for dynamic analyses of chromosomal proteins fused with GFP. In fact, even when we analyzed chromosomal proteins of A. thaliana in tobacco BY-2 cells, the result was same or highly similar with that in cultured Arabidopsis cells (Fujimoto et al. 2005).

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Although GFP offers great advantages in plant dynamic analyses, we should pay much more attention to potential artifacts in dynamic analyses of chromosomal proteins with GFP. Dynamic analyses with GFP do not show the real dynamics of proteins in vivo. GFP fusions may lead misfolding of target proteins and masking of specific regions interacted with DNA or other proteins. It is probable that overexpression of GFP fusions may have ceratin effects on the real dynamics or functions of the target protein with its expression under the native promoter. Results of dynamic analyses of chromosomal proteins should be confirmed with immunostaining using an antibody against the target protein. Recently, the development of GFP variants and novel fluorescent proteins has resulted in dramatic progress in dynamic analyses of chromosomal proteins. Dynamic analyses using multiple fluorescent proteins can be applied to analyses of chromosome dynamics in tobacco BY-2 cells. Moreover, chromosome dynamic analyses using tobacco BY-2 cells is particularly well suited to applications of advanced imaging analyses including fluorescence recovery after photobleaching (FRAP), fluorescence correlation spectroscopy (FCS), and fluorescence resonance energy transfer (FRET). Acknowledgements. We are grateful to Daisuke No (Kobe University), Satoru Fujimoto and Daisuke Kurihara (Osaka University) for providing the images of dynamic analyses using BY-2 cultured cells.

References Adams RR, Maiato H, Earnshaw WC, Carmena M (2001) Essential roles of Drosophila inner centromere protein (INCENP) and aurora B in histone H3 phosphorylation, metaphase chromosome alignment, kinetochore disjunction, and chromosome segregation. J Cell Biol 153:865–880 Arumuganathan K, Earle ED (1991) Nuclear DNA content of some important plant species. Plant Mol Biol Rep 9:208–219 Carmena M, Earnshaw WC (2003) The cellular geography of aurora kinases. Nat Rev Mol Cell Biol 4:842–854 Demidov D, Van Damme D, Geelen D, Blattner FR, Houben A (2005) Identification and dynamics of two classes of aurora-like kinases in Arabidopsis and other plants. Plant Cell 17:836–848 Eissenberg JC, Elgin SC (2000) The HP1 protein family: getting a grip on chromatin. Curr Opin Genet Dev 10:204–210 Fujimoto S, Yonemura M, Matsunaga S, Nakagawa T, Uchiyama S, Fukui K (2005) Characterization and localization analysis of Arabidopsis condensin subunits, AtCAP-H and AtCAP-H2. Planta 222(2):293–300 Gaudin V, Libault M, Pouteau S, Juul T, Zhao G, Lefebvre D, Grandjean O (2001) Mutations in LIKE HETEROCHROMATIN PROTEIN 1 affect flowering time and plant architecture in Arabidopsis. Development 128:4847–4858 Giet R, Glover DM (2001) Drosophila aurora B kinase is required for histone H3 phosphorylation and condensin recruitment during chromosome condensation and to organize the central spindle during cytokinesis. J Cell Biol 152:669–682

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Giet R, Uzbekov R, Kireev I, Prigent C (1999) The Xenopus laevis centrosome aurora/Ipl1-related kinase. Biol Cell 91:461–470 Goto H, Yasui Y, Nigg EA, Inagaki M (2002) Aurora-B phosphorylates Histone H3 at serine28 with regard to the mitotic chromosome condensation. Genes Cells 7:11–17 Hagstrom KA, Meyer BJ (2003) Condensin and cohesin: more than chromosome compactor and glue. Nat Rev Genet 4:520–534 Hagstrom KA, Holmes VF, Cozzarelli NR, Meyer BJ (2002) C. elegans condensin promotes mitotic chromosome architecture, centromere organization, and sister chromatid segregation during mitosis and meiosis. Genes Dev 16:729–742 Hasezawa S, Kumagai F (2002) Dynamic changes and the role of the cytoskeleton during the cell cycle in higher plant cells. Int Rev Cytol 214:161–191 Hendzel MJ, Wei Y, Mancini MA, Van Hooser A, Ranalli T, Brinkley BR, Bazett-Jones DP, Allis CD (1997) Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106:348–360 Hirano T (2002) The ABCs of SMC proteins: two-armed ATPases for chromosome condensation, cohesion, and repair. Genes Dev 16:399–414 Hirano T, Mitchison TJ (1994) A heterodimeric coiled-coil protein required for mitotic chromosome condensation in vitro. Cell 79:449–458 Hirano T, Kobayashi R, Hirano M (1997) Condensins, chromosome condensation protein complexes containing XCAP-C, XCAP-E and a Xenopus homolog of the Drosophila Barren protein. Cell 89:511–521 Hirota T, Gerlich D, Koch B, Ellenberg J, Peters JM (2004) Distinct functions of condensin I and II in mitotic chromosome assembly. J Cell Sci 117:6435–6445 Ito M (2000) Factors controlling cyclin B expression. Plant Mol Biol 43:677–690 Jackson JP, Lindroth AM, Cao X, Jacobsen SE (2002) Control of CpNpG DNA methylation by the KRYPTONITE histone H3 methyltransferase. Nature 416:556–560 James TC, Elgin SC (1986) Identification of a nonhistone chromosomal protein associated with heterochromatin in Drosophila melanogaster and its gene. Mol Cell Biol 6:3862–3872 Jessberger R (2002) The many functions of SMC proteins in chromosome dynamics. Nat Rev Mol Cell Biol 3:767–778 Kaitna S, Mendoza M, Jantsch-Plunger V, Glotzer M (2000) INCENP and an aurora-like kinase form a complex essential for chromosome segregation and efficient completion of cytokinesis. Curr Biol 10:1172–1181 Kaitna S, Pasierbek P, Jantsch M, Loidl J, Glotzer M (2002) The aurora B kinase AIR-2 regulates kinetochores during mitosis and is required for separation of homologous chromosomes during meiosis. Curr Biol 12:798–812 Kawabe A, Matsunaga S, Nakagawa K, Kurihara D, Yoneda A, Hasezawa S, Uchiyama S, Fukui K (2005) Characterization of plant Aurora kinases during mitosis. Plant Mol Biol 58:1–13 Kenton A, Parokonny AS, Gleba YY, Bennett MD (1993) Characterization of the Nicotiana tabacum L. genome by molecular cytogenetics. Mol Gen Genet 240:159–169 Kimura M, Kotani S, Hattori T, Sumi N, Yoshioka T, Todokoro K, Okano Y (1997) Cell cycledependent expression and spindle pole localization of a novel human protein kinase, Aik, related to Aurora of Drosophila and yeast Ipl1. J Biol Chem 272:13766–13771 Li Y, Kirschmann DA, Wallrath LL (2002) Does heterochromatin protein 1 always follow code? Proc Natl Acad Sci USA 99:16462–16469 Liu CM, McElver J, Tzafrir I, Joosen R, Wittich P, Patton D, Van Lammeren AA, Meinke D (2002) Condensin and cohesin knockouts in Arabidopsis exhibit a titan seed phenotype. Plant J 29:405–415 Losada A, Hirano T (2005) Dynamic molecular linkers of the genome: the first decade of SMC proteins. Genes Dev 19:1269–1287 MacCallum DE, Losada A, Kobayashi R, Hirano T (2002) ISWI remodeling complexes in Xenopus egg extracts: identification as major chromosomal components that are regulated by INCENPaurora B. Mol Biol Cell 13:25–39

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Morrison C, Henzing AJ, Jensen ON, Osheroff N, Dodson H, Kandels-Lewis SE, Adams RR, Earnshaw WC (2002) Proteomic analysis of human metaphase chromosomes reveals topoisomerase II alpha as an Aurora B substrate. Nucleic Acids Res 30:5318–5327 Moscone EA, Matzke MA, Matzke AJM (1996) The use of combined FISH/GISH in conjunction with DAPI counterstaining to identify chromosomes containing transgene inserts in amphidiploid tobacco. Chromosoma 105:231–236 Murata-Hori M, Tatsuka M, Wang YL (2002) Probing the dynamics and functions of aurora B kinase in living cells during mitosis and cytokinesis. Mol Biol Cell 13:1099–1108 Nagata T, Kumagai F (1999) Plant cell biology through the window of the highly synchronized tobacco BY-2 cell line. Methods Cell Sci 21:123–127 Nagata T, Nemoto Y, Hasezawa S (1992) Tobacco BY-2 cell line as the “HeLa” cell in the cell biology of higher plants. Int Rev Cytol 132:1–30 Neuwald AF, Hirano T (2000) HEAT repeats associated with condensins, cohesins, and other complexes involved in chromosome-related functions. Genome Res 10:1445–1452 Ono T, Losada A, Hirano M, Myers MP, Neuwald AF, Hirano T (2003) Differential contributions of condensin I and condensin II to mitotic chromosome architecture in vertebrate cells. Cell 115:109–121 Ono T, Fang Y, Spector DL, Hirano T (2004) Spatial and temporal regulation of condensins I and II in mitotic chromosome assembly in human cells. Mol Biol Cell 15:3296–3308 Schleiffer A, Kaitna S, Maurer-Stroh S, Glotzer M, Nasmyth K, Eisenhaber F (2003) Kleisins: a superfamily of bacterial and eukaryotic SMC protein partners. Mol Cell 11:571–575 Siddiqui NU, Stronghill PE, Dengler RE, Hasenkampf CA, Riggs CD (2003) Mutations in Arabidopsis condensin genes disrupt embryogenesis, meristem organization and segregation of homologous chromosomes during meiosis. Development 130:3283–3295 Staiger CJ, Lloyd CW (1991) The plant cytoskeleton. Curr Opin Cell Biol 3:33–42 Sugimoto K, Tasaka H, Dotsu M (2001) Molecular behavior in living mitotic cells of human centromere heterochromatin protein HPLalpha ectopically expressed as a fusion to red fluorescent protein. Cell Struct Funct 26:705–718 Sugimoto K, Urano T, Zushi H, Inoue K, Tasaka H, Tachibana M, Dotsu M (2002) Molecular dynamics of Aurora-A kinase in living mitotic cells simultaneously visualized with histone H3 and nuclear membrane protein importinalpha. Cell Struct Funct 27:457–467 Vagnarelli P, Earnshaw WC (2004) Chromosomal passengers: the four-dimensional regulation of mitotic events. Chromosoma 113:211–222 Van Damme D, Bouget FY, Van Poucke K, Inze D, Geelen D (2004) Molecular dissection of plant cytokinesis and phragmoplast structure: a survey of GFP-tagged proteins. Plant J 40:386–398 Wignall SM, Deehan R, Maresca TJ, Heald R (2003) The condensin complex is required for proper spindle assembly and chromosome segregation in Xenopus egg extracts. J Cell Biol 161:1041–1051 Yu Y, Dong A, Shen WH (2004) Molecular characterization of the tobacco SET domain protein NtSET1 unravels its role in histone methylation, chromatin binding, and segregation. Plant J 40:699–711

I.5 Ion Channels Meet Cell Cycle Control R. Hedrich and D. Becker1

1 Introduction BY-2 cells have advanced to becoming a convenient model system to study the cell cycle control machinery in plants in that these suspension-cultured cells continuously cycle between division and grow without differentiating (Nagata et al. 2004b). Much of what is known about cell cycle control in plants was discovered in BY-2 cells and led to the conclusion that cell cycle control represents a remarkably conserved process between animals and plants that has separated early in evolution (for review see Inze 2005b and references therein; Stals and Inze 2001b). Since plants, however, are sessile and any process in their entire life cycle relies on osmotic phenomena, plants have evolved unique strategies for growth, pattern formation and thus development (Gutierrez 2005d). The plant cell wall is a prerequisite for the establishment of turgor pressure and concomitantly prevents cell migration, as seen in animals. Thus plant differentiation and pattern formation largely occur post-embryonically based on the activity of meristematic cells that continuously undergo cell division, growth/elongation and differentiation throughout a plant’s life. While cyclin-dependent protein kinases (CDKs) drive cell cycle progression in both, plants and animals, there is increasing evidence of a key role of ion channels in general and K+ channels in particular in cell cycle control of animal cells. In plants K+ channels comprise part of the osmotic motor that controls turgor and osmotic-driven processes; in this chapter we focus on the relationship of K+ channels, K+ homeostasis, turgor control and cell cycle progression. Using synchronized BY-2 cells as a model system, we have identified and characterized novel tobacco K+ channel genes and investigated their role throughout the cell cycle. We found that K+ -uptake channels are crucial for transition from G1 - to S-phase. Micro pressure-probe assisted turgor measurements revealed that expression and activity of K+in channels correlates with turgor formation and cell elongation. Our data suggest a critical turgor threshold that separates dividing from elongating cells, which is discussed in the context of coordinated cytoplasmic/vacuolar K+ fluxes and other cell cycle related model systems. This chapter will thus concentrate on how to step from cell division to cell elongation and vice versa. A key player in both processes is the phytohormone 1 Plant

Molecular Physiology and Biophysics, Julius-von-Sachs-Institute for Biosciences, University of Würzburg, Julius-von-Sachs-Platz 2, 97082 Würzburg, Germany, e-mail: [email protected]

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Auxin. In previous studies on maize coleoptiles we have shown that Auxin activates the transcription and finally the density of certain K+ -uptake channels. Since cell division is K+ dependent too, we here discuss the role of individual K+ channels in controlling cell cycle progression. Our recent findings on the molecular physiology and biophysics of BY-2 potassium channels will be put into the context of K+ homeostasis at the different cell cycle phases. Comprehensive reviews on plant ion transport and the role of ion channels in plants are found elsewhere (Roelfsema and Hedrich 2002, 2005; Very and Sentenac 2003; Cherel 2004).

2 Cell Elongation Using the classical model system for growth and tropisms, the gras (maize) coleoptile (Darwin 1880), we could show that the growth promoting hormone auxin mediates cell elongation via the stimulation of K+ uptake. Detailed studies by Philippar et al. (1999) and Fuchs et al. (2003) identified the K+ channel ZMK1 as an essential element in this process. Upon Auxin treatment of coleoptile segments ZMK1 transcripts as well as channel activity increase. ZMK1 is a voltage-dependent K+ channel. Upon hyperpolarization of the plasma membrane ZMK1 is activated and mediates the uptake of potassium ions by the actively growing coleoptile cells (Bauer et al. 2000). Besides voltage, ZMK1 is activated by apoplastic acidification as well. Auxin is known to activate the plasma membrane H+ -ATPase and thereby induces a membrane hyperpolarization (Felle et al. 1991; Lohse and Hedrich 1992) accompanied by a pH drop in the cell wall. Protons extrude into the apoplast gate ZMK1 in a cooperative manner with membrane voltage and thereby increase the probability of this K+ channel opening (Bauer et al. 2000). How timing of K+ channel activity and density relate to the kinetics of cell elongation is reviewed in Becker and Hedrich (2002), and recent findings on Auxin signaling and transport are summarized in Blakeslee et al. (2005), Leyser (2005), Nemhauser and Chory (2005) and Paponov et al. (2005).

3 Cell Cycle 3.1 Potassium Homeostasis Controls Cell Cycle Progression To study the role of ion channels in cell cycle control we took advantage of synchronized tobacco BY-2 suspension-cultured cells (Nagata et al. 2004; Sano et al. 2004). In the presence of Aphidicolin, an inhibitor of DNA polymerase α, cells are arrested at the border of G1 - to S-phase. Following removal of Aphidicolin, cell cycling progresses in a synchronized manner and cells enter

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S-phase, reaching the mitotic peak after 8−9 h. Our recent studies have shown that cell cycle progression in BY-2 cells is associated with pronounced changes in endogenous potassium concentrations. Cellular K+ content is high in G1 (following 24 h of Aphidicolin treatment), declines through S- and M-phases and reaches a maximum in late M-phase, at the border to G1 . In the presence of Cs+ , a well-known blocker of K+ -uptake channels, cell cycle re-entry is significantly delayed by about 2 h. Likewise, cell cycle progression is retarded and the typical 8 h mitotic peak observed in non-treated cells was not evident in a 12 h experiment. This is reflected by the fact that the rise in K+ content at late M- and G1 -phases is suppressed in the presence of Cs+ , suggesting that K+ supply represents a limiting factor for cell cycle progression in BY-2 cells. In line with this idea, a similar effect to Cs+ treatment is brought about by reducing in the culture medium the K+ content from 21 to 1 mM. Conversely, increasing K+ supply two-fold positively feeds back on cellular K+ level, accelerates cell cycle progression and reduces the time to peak in mitosis to 7 h. Encouraged by these findings a detailed survey of the K+ dependent cell cycle stage was initiated. Analysis by laser scanning cytometry (LSC) on the basis of cellular DNA content revealed a G1 arrest of BY-2 cells in the presence of either the

Fig. 1. Cell cycle-dependent changes in K+ cyt , turgor and channel expression. BY-2 cells were synchronized by aphidicolin treatment and cell cycle progression from G1 arrest (t = 0) was achieved by aphidicolin removal. Fluctuations of cellular potassium and cell turgor as well as K+ -channel gene expression were followed throughout the cell cycle (see text)

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K+ -channel blocker Cs+ or low external [K+ ]. This suggests that passing a [K+ ] threshold is necessary to proceed from G1 - to S-phase (see Fig. 1). 3.2 Nature of BY-2 Potassium Channels In order to identify the nature of the “K+ -threshold” in a similar manner as successfully applied to identify ZMK1, an Auxin-induced K+ channel important for cell elongation and tropisms (Philippar et al. 1999; Bennett et al. 2000; Fuchs et al. 2003), we isolated K+ channel genes expressed in BY-2 cells. Thereby four Shaker-like K+ channels were identified: the putative K+ -uptake channels NKT1 and NtKC1 as well as the two outward rectifiers NTORK1 and NTORK2 (Sano et al. 2004), the functional properties of which were accessed by growth rescue of a K+ -transport deficient E. coli mutant (Epstein and Kim 1971) and expression in Xenopus oocytes followed by electrophysiological characterisation (Sano et al. 2004). When following the expression of BY-2 K+ -channel genes by quantitative RT-PCR throughout the cell cycle, NKT1 expression increased together with the M/G1 markers E2F and CYCB1;2. Likewise NKT1 transcripts were highest when the cell cycle was arrested by phosphate starvation (Sano et al. 1999), again indicating expression in late M- to G1 -phase. In contrast, NtKC1 expression was highest in S-phase while NTORK1 transcript levels were shown to peak in G1 -phase. To elucidate the role of NKT1 in cell cycle progression we took advantage of inducible transgenic BY-2 cells expressing either NKT1 in antisense orientation or cells lines exhibiting NKT1 silencing due to overexpression. Both lines under inductive conditions are characterized by largely reduced NKT1 transcripts. As a matter of fact K+ content was reduced, cell cycle re-entry into S-phase was significantly delayed, and the peak of the mitotic index was retarded. 3.3 Physiology of BY-2 Potassium Channels Since K+ channels can control membrane voltage (charge balance) as well as mass flow of potassium, we studied the electrical properties of synchronized BY-2 cells along the different phases of the cell cycle. As predicted from our molecular and heterologous expression studies on NKT1 (Sano et al. 2004), a homologue of the Arabidopsis K+ -uptake channel AKT1 (Hirsch et al. 1998), BY-2 cells operate a hyperpolarization-activated K+in channel. This current was lowest in M-phase and highest in S-phase of the cell cycle (own unpublished results) and has been shown to be susceptible to blocking by Cs+ and Ba2+ . In view of the fact that microfilament-depolymerizing drugs affected the activity of the BY-2 K+in channel, microfilaments seem to participate in the regulation of K+ currents in this cell type (Hwang et al. 1997; Stoeckel and Takeda 2002). In root hairs of A. thaliana AKT1 activity is modulated by another K+ -channel α-subunit, AtKC1 (Reintanz et al. 2002). In BY-2 cells the corresponding homologue NtKC1 is expressed as well. This suggests that NKT1 and NtKC1,

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like AKT1 and AtKC1, form heteromeric channel complexes in BY-2 cells, too. In addition, the fact that neither KAT1- nor AKT2-like channels seem to be expressed in ‘Bright Yellow-2’ cells, NKT1/NtKC1 channels probably constitute the major molecular pathway for K+ -uptake in this cell type. Upon depolarization a K+ -release channel in the plasma membrane of BY-2 protoplasts activates, exhibiting the typical properties of plant outward rectifiers (Gaymard et al. 1998; Ache et al. 2000; Ivashikina et al. 2001; Hosy et al. 2003): activation at membrane voltages positive of the K+ equilibrium potential EK , sigmoidal, slow activation kinetics and K+ -dependent gating properties (c.f. Vanduijn et al. 1993). These currents, and thus K+ efflux, were maximal at G2 -phase, blocked by Ba2+ (Stoeckel and Takeda 2002) and resembled the functional properties of NTORK1 when expressed in Xenopus oocytes. Equipped with these channel components BY-2 cells are able to control K+ homeostasis during cell cycle progression. From the expression levels of NKT1 and NTORK1 as well as the fraction of inward and outward currents K+ uptake seems to dominate S-phase and K+ release has to take place at the onset of cell division. The direction of K+ flux is a function of the electrochemical potential which is determined by the membrane potential and the chemical potential for potassium ions. Comparing the membrane potential of S-phase (growing) with M-phase (dividing) cells revealed that the membrane potential of growing cells was about 40 mV more hyperpolarized. Dividing cells, however, exhibited membrane potentials around –65 mV, and thus appeared depolarized. Furthermore, we could show that plasma membrane H+ -ATPase activity and thus the driving force for K+ uptake was higher in elongating cells compared to dividing cells. This suggests a critical role for turgor formation and threshold for cell cycle re-entry and during cell cycle progression. Accordingly, in conjunction with changes in the electrical properties of the BY-2 plasma membrane, turgor pressure differences were recorded. Depolarized, dividing cells exhibited 50% of the turgor associated with hyperpolarized, growing cells, only. Since dividing and elongating cells differ by about 100 mosmol, in osmotic pressure turgor changes seem to follow the osmotic potential. These findings agree well with the observed changes in K+ content during cell cycle progression and suggest that potassium is the major osmolite that controls cell turgor and membrane potential in BY-2 cells. Interfering with K+ homeostasis, by means of K+in -channel block by Cs+ or low K+ supply, consequently feeds back on turgor and cell cycle progression. In the presence of Cs+ as well as under low K+ supply, cellular [K+ ] is constantly low throughout a 12 h period. BY-2 cells, however, still reach Sphase, though are delayed by about 2 h. This phenomenon might result from overcoming a turgor threshold rather than solely a [K+ ] threshold through adaptive processes. Hence, turgor formation, osmotic adjustment and cell cycle progression at low cellular [K+ ] in BY-2 cells could be achieved through de novo synthesis of osmotically active compatible solutes such as sugars, malate or mannitol (Peltier and Marigo 1999). In addition, a reduction in the volumetric

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elastic modulus (ε) by bonding rearrangements in the cell wall could aid this process (Kourie and Findlay 1991; Peltier and Marigo 1999; Cosgrove 2005).

4 Discussion Cell cycle progression is mainly controlled by evolutionary highly conserved serine/threonine kinases known as the cyclin-dependent kinases (CDKs) (Stals and Inze 2001). In parallel, plant hormones represent well-known regulators of the cell cycle, among them auxin, cytokinin and brassinosteroids which act as promoters of cell cycle progression, while the stress hormone abscisic acid inhibits cell division. Likewise, the nutritional status of plants such as sugar and phosphate availability has been shown to influence cell cycle progression (Sano et al. 1999; Nagata et al. 2002). Transcriptom analysis of synchronised Arabidopsis cell culture revealed that various genes encoding transporters for phosphate, sugars, nitrate and oligo-peptides as well as aquaporins, glutamate receptors and high-affinity K+ -transporters are regulated in a cell cycle specific manner (Menges et al. 2002). In a first genome-wide approach using BY-2 cells based on cDNA AFLP analysis Breyne et al. (2002) showed that transcripts of ATPases, potassium-, nitrate-, amino acid- or Ca2+ -transporters present clusters of cell cycle-modulated genes. Since potassium represents a macronutrient and an important osmoticum for plant cells we here address the question of how K+ -homeostasis feeds back on cell cycle progression in BY-2 cells. Likewise, K+ depletion, the K+ -channel blockers caesium and tetraethylammonium (TEA+ ) significantly delay cell cycle progression of aphidicolin synchronized cells. Cs+ as well as TEA+ represent K+ -channel blockers rather than inhibitors of high-affinity (micromolar range) K+ -transporters and the cell cycle responds to millimolar [K+ ] changes. Hence, these pharmacological studies point towards a role for voltage-dependent, hyperpolarization-activated K+ uptake channels in this process. BY-2 cells seem to express four Shaker K+ channels (Pilot et al. 2003), two K+ -uptake channels (NKT1 and NtKC1) and two K+ -release channels (NTORK1 and 2). NKT1 is an orthologue of the Arabidopsis AKT1 channel, the latter mediating K+ uptake from the soil into the growing root (Hirsch et al. 1998; Dennison et al. 2001). Interfering with NKT1 expression by means of anti-sense or co-suppression results in impaired K+ uptake in the respective cell lines and negatively feeds back on cell cycle progression and proliferation. Although the role of hyperpolarization-activated K+ channels in animals differs from that in plants (Craven and Zagotta 2005) a general role for potassium channels as regulators of growth factor-stimulated cell proliferation in animal cells has been proposed previously (Boonstra et al. 1981). Respective studies have shown that the human “ether-a-gogo” (h-eag) channel which is preferentially expressed in brain tissue is highly active in somatic cancer cell lines and overexpression of h-eag resulted in a transformed phenotype (Pardo et al. 1999; Pardo 2004). Likewise, in cultured cells, TEA+

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and related K+ -channel blockers prevented insulin-induced cell proliferation and arrested cells in G1 -phase (Guo et al. 2005). Overexpression of the voltagedependent K+ channels Kv1.3 or Kv1.4 increases cell proliferation in the absence of mitogens, whereas Kv1.6 overexpression inhibited mitogen-induced cell cycle progression (Vautier et al. 2004). Since animal K+ channels control cell cycle progression and exhibit an oncogenic potential it is tempting to speculate that plant outward rectifiers may impinge on cell cycle control too. In line with this hypothesis, an increase in transcripts of the BY2 channel NTORK1 and highest K+ -efflux channel activity was measured before and in G2 -phase, respectively. Whether overexpression or repression of NTORK affects cell cycle progression and entry into mitosis awaits further investigation. In contrast to NTORK1, NKT1 expression is highest in G1 and thus precedes maximal K+ -uptake channel activity during S-phase. In line with the proposed role of potassium salts as major osmolytes that drive turgor formation and thus plant cell growth/elongation we found that the osmotic pressure of elongating and dividing cells differs by at least 100 mosmol. Likewise, turgor pressure is more than two-fold higher in elongating cells compared to dividing cells. Thus, whenever cells start to elongate, NKT1/NtKC1 genes are turned on and channel activity increases. Channel-mediated K+ transport is favoured by increasing the driving force for uptake of potassium ions by activation of proton pumps, and hence membrane hyperpolarization and acidification of the apoplast. In contrast, entry into mitosis seems to require turgor reduction by means of potassium loss via K+ -efflux channels (Fig. 2). A turgor-dependent step for cytokinesis and thus cell division becomes evident during endosperm devel-

Fig. 2. Schematic representation of the balance between cell division and cell elongation. Cell division is favoured at low turgor pressure, achieved by suppression of K+ -uptake (NKT1) channels and activation of K+ -efflux (NTORK1). Conversely, cell elongation requires overcoming of a turgor threshold by means of activation of K+ uptake and shut down of K+ efflux

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opment. The syncytial phase of endosperm development is characterized by successive divisions of the triploid zygotic nucleus without cytokinesis (Berger 1999). Separation of nuclei occurs during later stages and is achieved by cell wall and thus turgor formation, initiating cytokinesis and the cellularization phase (Mayer and Jürgens 2004). Accordingly, mitosis and thus cell division and plate formation are accomplished at low turgor, while exit from mitosis, separation of daughter cells and growth are achieved by reactivation of K+ uptake. Cells impaired in K+ uptake exhibit delayed cell cycle progression suggestive of an active compensation of turgor formation by de novo synthesis of organic osmolytes or induced expression of high-affinity K+ transporters (Menges et al. 2002). In contrast to BY-2 cells, yeast seems to control cell cycle progression from G1 -phase via K+ -transporters (Yenush et al. 2002). Expression and/or activity of yeast high-affinity potassium transporters TRK1 and 2 and thus K+ homeostasis are affected in yeast mutants lacking the phosphatase Ppz (Merchan et al. 2004), implicating that post-translational mechanisms play an additional role in K+ -dependent control of cell cycle progression. Our results with BY-2 cells are well in agreement with the situation in a more complex system characterized by an additional differentiation step as a consequence of cell division and elongation. Leaves of tobacco and Arabidopsis seem to be a model system of higher complexity. Cells at the base of an expanding leaf undergo division and exhibit a gradual pattern of elongation and differentiation towards the leaf tip. Studies on leaf growth and development thus seem to provide answers to the controversially discussed question of whether cell division drives growth and differentiation or cell division is a consequence of growth (Beemster et al. 2005; Gutierrez 2005). Stiles et al. (2003) have shown that dividing and expanding cells exhibit different sensitivity to exogenous potassium supply. Light-stimulated growth in tobacco leaves peaks at the leaf base and is inhibited by the K+ channel blocker TEA+ , which reduces K+ uptake in these cells by about 60%. This points towards K+ -uptake channels as major pathways in elongating leaf cells. In contrast, inhibition of K+ uptake by TEA+ at the leaf tip was only 17%, suggesting that differentiated, fully developed leaf cells hardly express K+ -uptake channels. This observation is well in agreement with electrophysiological recordings on tobacco mesophyll cells, which have shown to express K+ -efflux channels while lacking hyperpolarization-activated K+ -uptake channels (Bei and Luan 1998). A direct correlation between K+ channel expression/activity in this model system, however, has not been established yet. Leaves of Arabidopsis cell cycle mutants constitutively overproducing CDK inhibitors (KRPs) consist of ten-fold fewer cells with an average size six-fold greater than that of control cells. On the contrary, overexpression of positive regulators of the cell cycle results in a massive increase in cell proliferation and more but significantly smaller cells (Gutierrez 2005, and references therein; Inze 2005). These data demonstrate tight feed back control mechanisms of cell division and elongation. No evidence, however, exists so far for an interrela-

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tionship between cell cycle control and differentiation. Thus, interference with cell cycle control mechanisms may cause morphological changes, but seems not to result in oncogenic transformation, as seen in animal cells (Farias et al. 2004; Gutierrez 2005). An exception is that of Agrobacterium-mediated cell proliferation and tumour formation in Arabidopsis. Deeken et al. (2003) have shown that suppression of shoot-specific K+ channels in favour of root-specific K+ channels provides for K+ supply and proliferation in rapidly growing tumour cells. Likewise, knock-outs of the potassium channels AKT2/3 or AKT1 fail to develop tumours, rendering the Arabidopsis K+ channels AKT1 and AtKC1 as valuable tumour markers. Thus it seems obvious to investigate K+ channel mutants with respect to cell cycle related phenotypes. This would include not only the well characterized Shaker-like K+ channels but also the TPK1-like potassium channels (Czempinski et al. 2002; Becker et al. 2004; Bihler et al. 2005) or SV-/TPC1-like Ca2+ -induced Ca2+ permeable channels (Hedrich and Neher 1987; Kadota et al. 2005; Peiter et al. 2005), known to be localized to the vacuolar membrane. In Arabidopsis TPK1 as well as TPC1 is expressed ubiquitously. Recent studies suggest that TPK1 constitutes a Ca2+ -activated vacuolar K+ channel (VK) channel, while TPC1 seems to account for a subunit of the prevalent slow-vacuolar (SV-) channel (Ivashikina and Hedrich 2005; Peiter et al. 2005). Vacuoles represent the largest plant organelles and may comprise up to 90% of cell volume in mature cells. An osmotic continuum between cytoplasm and vacuole is the basis for turgor formation and implicates that turgor formation is impossible without vacuoles. Vacuoles are highly dynamic structures that change morphology during cell cycle progression (Kutsuna and Hasezawa 2002; Kutsuna et al. 2003). Cell division is accompanied by fragmentation of the central vacuole into many small daughter vacuoles (Fig. 3A), a situation that is observed in dividing immature cells at the root tip. In contrast, elongating cells are characterized by fusion of daughter vacuoles to regenerate a large central vacuole, as seen in root cells of the elongation zone (Fig. 3B; Herman et al. 1994; Segui-Simarro and Staehelin 2005; Shimazaki et al. 2005). In addition to fusion events, vacuolar membrane including its transport proteins originates from ER (Herman et al. 1994). TPK1-like channels have been detected in BY-2 cells (T. Sano, pers. comm.) and it is thus tempting to consider that vacuolar ion channels undergo cell cycle-dependent fluctuations in expression and/or activity. In fact, BY-2 cells express two TPC1-like Ca2+ - and K+ -permeable channels NtTPC1A and B (Kadota et al. 2004), and both exhibit highest expression during M- and G1 -phase (Kadota et al. 2005). Both TPC1 and TPK1 channel proteins contain Ca2+ -binding EF hands and have been shown to be gated by cytoplasmic calcium (Bihler et al. 2005; Peiter et al. 2005). Thus an elevation of cytosolic [Ca2+ ] leading to activation of TPC1- and TPK-channels seems to accompany mitosis as well as cell cycle progression from G1 (Figs. 3A and B). Whether plant ion channels in general or K+ channels in particular control cell cycle progression or constitute targets of the RB/E2F/DP pathway known

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Fig. 3. Differential channel activities in elongating and dividing cells. A Elongating cells, characterized by large vacuoles, exhibit high NKT1/NtKC1 expression and H+ -ATPase activity. ATPase-mediated plasma membrane hyperpolarization results in activation of voltage-gated NKT1/NtKC1 K+ -uptake channels, potassium accumulation and generation of turgor. The Ca2+ activated BY-2 vacuolar channels NtTPK1 and NtTPC1A/B are highly expressed at G1 and mediate K+ - and Ca2+ -currents across the vacuolar membrane, respectively. B Dividing cells are characterized by small fragmented vacuoles. Low turgor observed in M-phase is accompanied by expression of the K+ -release channel NTORK1. Ca2+ -dependent activation of vacuolar NtTPC1A/B and NtTPK1 channels is suggested to participate in fusion of vacuolar vesicles and plasma membrane vesicles at the phragmoplast

to control the G1/S transition remains to be elucidated. Transgenic plants exhibiting impaired cell cycle control might serve as valuable tools to address this question. Clearly, turgor control via ion channel activity seems to determine cell cycle progression in plant cells. In addition, it may be considered that localised channel activity and thus local forces control cell plate formation and

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exit from mitosis. Considering the available model systems (BY-2, Arabidopsis, tobacco, rice) the combination of molecular genetics and electrophysiology will open the door for a deeper understanding of how ion channels communicate with cell cycle control machinery and thus control cell division, growth and differentiation. Acknowledgements. Our apologies to any colleagues whose work has not been cited here due to space limitations. We thank T. Nagata and T. Sano for stimulating discussions and a very successful collaboration. Our work was supported by grants from the Alexander-von-Humboldt Foundation and the Deutsche Forschungsgemeinschaft.

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I.6 The NACK-PQR MAP Kinase Cascade Controls Plant Cytokinesis M. Sasabe, Y. Takahashi, T. Soyano, H. Tanaka, K. Kousetsu, T. Suzuki, and Y. Machida1

1 Introduction Cell division involves three essential processes for the proper distribution of genetic information to daughter cells: replication of the sets of chromosomes, separation of the chromosomal pairs, and division of the cytoplasm. The final step is called cytokinesis, and it is the critical step for guaranteeing precise cell division and proper cell differentiation. In contrast to the first two steps of cell division, the mode of cytokinesis in plants appears to be different from the modes in yeast and animals. In animal cells, cytokinesis is achieved by constriction of the cell membrane from the outside to the inside (Field et al. 1999; Fig. 1A), whereas cytokinesis in plant cells occurs through the formation of new cell walls. This cell wall formation occurs from the interior to the periphery of the cell and is mediated by a plant-specific apparatus called the phragmoplast (Nishihama and Machida 2001; Fig. 1B). Regardless of the opposite directions of cytokinesis in plant and animal cells, the cytoskeletal structures are largely conserved. During late anaphase and telophase, animal cells develop a well-defined network of microtubules (MTs) between the two daughter nuclei as they migrate to the opposite poles of the cell. This MT-based architecture, called the central spindle or midbody, plays an essential role in cytokinesis. Plant cells construct a phragmoplast, which is also composed of MTs that lie mainly between the two daughter nuclei at late anaphase. The expansion of the phragmoplast is essential for the formation of new cell plates and for complete cytokinesis. The central spindle and the phragmoplast both contain two bundles of antiparallel MTs that are interdigitated at their plus ends. The minus ends of these MTs face the chromosomes separating to the opposite cell poles. The similarities of these cytokinetic structures imply that there are aspects of the regulation of cytokinesis that are conserved between animals and plants. Recently, XMAP215 and PRC1, which are involved in the regulation of MT structures during cell division in animal cells, were found to be conserved in higher plants and necessary for organizing MTs and for completing cytokinesis (Whittington et al. 2001; Twell et al. 2002; Müller et al. 2004). This suggests that, although the direction of cell division has diverged, similar mechanisms exist in different species for controlling cytokinesis via regulation of MT dynamics. Our 1 Division

of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan, e-mail: [email protected]

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Fig. 1. Cytokinesis in animals and plants. A Schematic representation of animal cytokinesis, which occurs by an outside-in process. B Schematic representation of plant cytokinesis, which occurs by an inside-out process. Spindle and phragmoplast MTs are shown in green, vesicles in red. C Phragmoplast expansion: disassembly of phragmoplast MTs at the inner edge of the phragmoplast and reassembly of MTs at the outer edge, which results in centrifugal expansion of the phragmoplast. The phragmoplast consists of antiparallel MTs that are interdigitated in the center at their plus ends (+ end) and in which MT turnover (depolymerization and repolymerization) occurs in parallel with cell plate formation

recent studies have revealed that a mitogen-activated protein kinase (MAPK) cascade is involved in plant cytokinesis, especially in the expansion of the phragmoplast, and it appears to be related to stability of phragmoplast MTs (Nishihama et al. 2001, 2002; Ishikawa et al. 2002; Soyano et al. 2003).

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2 Plant Cytokinesis Plant cytokinesis occurs in the phragmoplast, a structure composed mainly of MTs that are organized by tubulin dimers and actin filaments at the late anaphase step of cell division. At an early stage, the phragmoplast has a barrellike shape made up of anti-parallel MTs overlapping in the center at their plus ends. A new cell plate is generated at this equatorial zone. Once cell plate formation begins, the phragmoplast changes into a ring-like structure and expands centrifugally while maintaining its localization at the edge of the growing cell plates. Materials for the construction of the cell plate are supplied by the Golgi complex through the vesicle trafficking system (Yasuhara et al. 1995; Sonobe et al. 2000; Yokoyama and Nishitani 2001). It appears that the Golgi-derived vesicles are transported along the phragmoplast, continuously fused at the equatorial region, and the new cell plates expand as the cell walls mature. In other words, the formation of cell plates follows the expansion of phragmoplast MTs (Samuels et al. 1995; Nishihama and Machida 2001). The dynamics of MT structures, including the expansion of the phragmoplast, appears to be mediated by MT turnover, specifically, disassembly of MTs on the inside and reassembly of tubulin dimers on the outside of the phragmoplast (Nishihama and Machida 2001). The polymerization or depolymerization of MTs occurs on either the plus or minus ends, although it occurs much faster at the plus ends. This difference in rates is due to the polarity created by the head-to-tail association of α/β-tubulin heterodimers (Desai and Mitchison 1997). Therefore, the equator of the phragmoplast, where the plus ends of MTs are positioned, is both the starting point for MT turnover and the region at which the cell plate is formed (Fig. 1C). In conclusion, plant cytokinesis has mainly four processes: (1) the formation of the phragmoplast; (2) the transport, accumulation, and fusion of Golgi-derived vesicles; (3) the synthesis and maturation of cell walls; and (4) the expansion of the phragmoplast. Several factors involved in vesicle fusion and cell wall biosynthesis during cytokinesis in plants have been isolated from Arabidopsis (Nacry et al. 2000). KNOLLE, a plant-specific syntaxin, and KEULE, a Sec1 homologue, are key regulators of vesicle trafficking. These two proteins interact and participate in the fusion of Golgi-derived vesicles, which results in formation of the cell plate during cytokinesis (Lukowitz et al. 1996; Waizenegger et al. 2000; Assaad et al. 2001). Dynamin-related proteins (ADL1A and ADL1E) seem also to be involved in vesicle fusion during cytokinesis (Kang et al. 2003). Other factors including KORRIGAN and CYT1 encode the enzymes required for synthesis and/or maturation of a new cell plate (Zuo et al. 2000; Lukowitz et al. 2001). However, the mechanisms that are associated with MT dynamics promoting the formation and expansion of phragmoplast and those that couple cell plate formation to the turnover of phragmoplast MTs have not been well characterized. The results of recent studies suggest that a MAPK cascade is involved in these regulatory mechanisms.

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3 The MAPK Cascade Involved in Cytokinesis 3.1 NPK1: A MAP Kinase Kinase Kinase that Is a Positive Regulator of Cytokinesis NPK1 (nucleus- and phragmoplast-localized protein kinase 1) was identified as an M-phase-specific protein. The results of various types of experiments revealed the involvement of NPK1 in cytokinesis, especially in the expansion of cell plates (Nishihama et al. 2001). The gene for NPK1 from tobacco encodes a member of the mitogen-activated protein kinase kinase kinase (MAPKKK) family, which is expressed in BY-2 cells preferentially at the logarithmic phase of cell growth and in tissues that contain proliferating cells (Banno et al. 1993; Nishihama et al. 1997; Nakashima et al. 1998). In addition, the kinase activity of NPK1 also increases during late M phase (Nishihama et al. 2001). NPK1 consists of essentially two domains: a kinase domain similar to that of MAPKKKs in the amino-terminal half, and a regulatory domain in the carboxyterminal half (Fig. 2A). Deletion of the carboxy-terminal half increases kinase activity, indicating that this domain negatively regulates NPK1 (Banno et al. 1993; Machida et al. 1998). This domain also has an NLS (nuclear localization signal) motif, a coiled-coil structure, and consensus sequences for phosphorylation by cyclin-dependent protein kinases (CDKs) (Nishihama et al. 1997). Later, this region was identified as the binding site for the NACK1 kinesinlike protein, which regulates the activity and localization of NPK1 (Ishikawa et al. 2002; Nishihama et al. 2002). Thus, the regulatory domain seems to help activation of NPK1 through the interaction with other factors and/or the phosphorylation by CDKs. NPK1 is localized in the nucleus at interphase and prophase prior to breakdown of the nuclear envelope, whereas it is localized in the cytoplasm at metaphase. During cytokinesis, when the kinase activity of NPK1 increases (Fig. 3C), NPK1 shifts to the leading edge of the equatorial zone of the phragmoplast (Nishihama et al. 2001; Fig. 2B). To investigate the function of NPK1, we adopted a dominant-inhibitory strategy using a kinase-defective mutant (NPK1:KW) in tobacco BY-2 cells or tobacco plants. Overexpression of this mutant inhibits the lateral expansion of the phragmoplast and causes the generation of multinucleated cells with incomplete cell plates (Nishihama et al. 2001). These aberrant cell plates can be stained with calcofluor, which stains β-glucans in cell walls, or with aniline blue, which stains callose, a polysaccharide synthesized at the initial stage of vesicle fusion (Nishihama et al. 2001). Arabidopsis homologues of NPK1 (ANP1, 2, and 3) also appear to be involved in cytokinesis. Loss of function of two of the three homologues of NPK1 (ANP2 and ANP3) results in defects in cytokinesis, specifically, the formation of multinucleated cells with incomplete cell walls (Krysan et al. 2002). These results suggest that the activation of NPK1 is necessary for the progression of cytokinesis.

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Fig. 2. Colocalization of NPK1 and NACK1 at the equator of the phragmoplast in telophase. A Domain organization and critical regions in the NPK1 and NACK1 proteins. The coiled-coil structures (green) were predicted by the COILS program (Lupas et al. 1991), and the nuclear localization signal (red) and NACK1/NPK1-binding sites (bars) were determined in our previous studies (Ishikawa et al. 2002). B Subcellular localization of NPK1 (top) and NACK1 (bottom) at telophase in BY-2 cells. BY-2 cells were triple-stained with mouse antibodies against α-tubulin (green), rabbit antibodies against NPK1 or NACK1 (red), and 4
-6-diamidino-2-phenylindole (DAPI) for nuclei (blue). C Colocalization of NPK1 and NACK1 at the equator of the phragmoplast. BY-2 cells expressing GFP-NPK1 (green) were double-stained with NACK1-specific antibodies (red) and DAPI

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M. Sasabe et al. Fig. 3. Specific accumulation of NACK1 and activation of NPK1, NQK1, and NRK1 at late M phase during the cell cycle. A Plot of the mitotic indices of BY-2 cells synchronized at M phase. The cell cycle was arrested at prometaphase by propyzamide after release from an aphidicolin block. B Western blots with specific antibodies against NACK1, NPK1, NQK1, and NRK1. At the indicated times after the removal of propyzamide, the cells were harvested and proteins were extracted. C Protein kinase activities of NPK1, NQK1, and NRK1 determined by immunocomplex kinase assays using recombinant kinasenegative NQK1, kinase-negative NRK1, and myelin basic proteins as substrates, respectively

3.2 NACK1: A Kinesin-Like Protein (KLP) that Activates NPK1 The MAPK cascade is a signaling pathway conserved in all eukaryotes and consists of members of three protein kinase families: the MAPKKKs, the MAPKKs, and the MAPKs (English et al. 1999). The members of these families have highly conserved structures even though they participate in MAPK cascades with distinct physiological roles. Animals and yeast have some proteins that regulate the MAPKKKs via protein–protein interaction. For example, the small G protein Ras (Farrar et al. 1996; Luo et al. 1996) and the 14-3-3 protein regulate Raf MAPKKK (Irie et al. 1994); TAB1 regulates TAK1 MAPKKK (Shibuya et al. 1996); SSK1 regulates SSK2 MAPKKK (Posas and Saito 1998); and STE20 regulates STE11 MAPKKK (Wu et al. 1995). To isolate the activators of NPK1 MAPKKK, we used a functional yeast genetic system based on the mating pheromone-responsive MAPK cascade, which consists of STE11 MAPKKK, STE7 MAPKK, and FUS3 MAPK (Irie et al. 1994). We identified two KLPs in tobacco, which we designated NPK1-activating kinesins 1 and 2 (NACK1 and NACK2). Both of these proteins interact with NPK1 and increase its protein kinase activity (Nishihama et al. 2002). Messenger RNAs of NACK1 and NACK2 and NACK1 proteins accumulate specifically

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at the late M phase of the cell cycle, and the patterns of accumulation are consistent with the increase in the activity of NPK1 (Fig. 3B). The amino-terminal halves of NACK1 and NACK2 contain the MT-based motor domain, which is conserved among various KLPs, whereas the carboxyterminal halves have the typical stalk domains with coiled-coil structures (Fig. 2A). Yeast two-hybrid and biochemical analyses have shown that NPK1 is activated by direct association with NACK1, which is mediated by the interaction between predicted coiled-coil structures in the regulatory domain of NPK1 and in the stalk domain of NACK1 (Ishikawa et al. 2002; Nishihama et al. 2002). NACK1 is colocalized with NPK1 at the phragmoplast equator during cytokinesis in BY-2 cells (Fig. 2B and C), and it seems to regulate the subcellular localization as well as the activity of NPK1 via this direct interaction. We found that deletion of the regulatory domain of NPK1, which contains the NACK1-binding site, prevented the localization of NPK1 to the equator of phragmoplast. In addition, overexpression in tobacco cells of a mutant NACK1 protein that lacks the putative motor region (NACK1:ST) results in failure to accumulate NPK1 proteins at the phragmoplast equator and defects in cytokinesis (Nishihama et al. 2002; Fig. 4B). This suggests that NACK1 plays a role as a positive regulator in both the subcellular localization and the activation of NPK1 MAPKKK during cytokinesis. The homologues of the NACK1 and NACK2 genes in Arabidopsis are designated AtNACK1 and AtNACK2, respectively (Nishihama et al. 2002), and they are identical to HINKEL (HIK) and STUD (STD)/TETRASPORE (TES), respectively (Strompen et al. 2002; Yang et al. 2003). Loss-of-function mutations in the AtNACK1/HIK and STD/TES/AtNACK2 result in the occasional failure of somatic and male-meiotic cytokinesis, respectively (Hülskamp et al. 1997; Spielman et al. 1997; Nishihama et al. 2002; Strompen et al. 2002; Yang et al. 2003). Recently, we showed that these genes have redundant functions and are essential for cytokinesis during both male and female gametogenesis (Tanaka et al. 2004). Indeed, it seems likely that NACK1-related kinesin-like proteins in both tobacco and Arabidopsis are functionally conserved. 3.3 The NACK-PQR Pathway: A MAPK Cascade that Promotes Expansion of the Cell Plate We also identified the NQK1/NtMEK1 and NRK1 proteins of tobacco as a MAPKK and MAPK, respectively, which act downstream of NPK1 (Soyano et al. 2003). To isolate downstream factors of NPK1, we used a yeast genetic system that is based on the osmosensing MAPK cascade of yeast (Brewster et al. 1993; Maeda et al. 1994, 1995). In yeast, the osmosensing MAPK cascade includes Ssk2/22p and Ste11p MAPKKK, Pbs2p MAPKK, and Hog1p MAPK. We constructed a tobacco cDNA library and introduced it into yeast cells lacking the pbs2 gene but expressing NPK1 and NACK1 cDNAs. By screening this yeast cell’s library under high osmotic conditions, we isolated NQK1 cDNA. NRK1

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was then isolated as a binding partner of NQK1 using a yeast two-hybrid system. NPK1 phosphorylates and activates NQK1, which in turn phosphorylates and activates NRK1 (Soyano et al. 2003). Although NPK1 and NACK1 proteins rapidly disappear after M phase, NQK1 and NRK1 proteins accumulate throughout the cell cycle (Fig. 3B); however, the activities of NPK1, NQK1, and NRK1 in tobacco cells increase at the late M phase of the cell cycle and decrease after the M phase, and the patterns of activation of these protein kinases are similar to the pattern of NACK1 accumulation (Nishihama et al. 2001; Soyano et al. 2003; Fig. 3C). Moreover, studies on the subcellular localization of NQK1 and NRK1 revealed that they were also localized at the equator of the phragmoplast at least at telophase (our unpublished observations). These results suggest that activation of NPK1 MAPKKK by NACK1 binding causes the activation of NRK1 MAPK via the activation of NQK1 MAPKK at the equator of the phragmoplast during cytokinesis. The role of NQK1 in cell division was demonstrated by overexpressing a kinase-defective mutant form of NQK1 (NQK1:KW) in BY-2 cells and by mutation of ANQ1, the Arabidopsis homologue of NQK1. The NQK1:KW expressing cells and anq1 mutants were multinucleated and had incomplete cell plates (Fig. 4B). This indicated that NQK1 is required for expansion of the phragmoplast and for formation of the cell plates (Soyano et al. 2003). Although the activation of NRK1 is tightly coupled to the activation of NPK1 and NQK1, involvement of NRK1 in the formation of the cell plates has not yet been experimentally demonstrated. Recently, we identified a loss-of-function mutant of the Arabidopsis homologue of NRK1 (ANR1) that is defective in cytokinesis (our unpublished observations). These results suggest that the MAPK cascade composed of NPK1 MAPKKK, NQK1 MAPKK, and NRK1 MAPK is activated by binding of NACK1/2 KLP and promotes the formation of the cell plate. We designated this cascade the NACK-PQR pathway (Soyano et al. 2003; Fig. 4A).

4 Regulation of the NACK-PQR Pathway in the Cell Cycle Machinery How is the activation of the NACK-PQR pathway properly regulated? The progression of the cell cycle is strictly controlled by cyclin/CDK complexes. When cells are subjected to appropriate conditions, cyclin/CDK complexes are activated to phosphorylate various proteins that regulate phase-specific processes. Ito et al. (1998, 2001) revealed that the transcription of plant B-type cyclin genes, which are specifically expressed in the G2-M phase, is regulated by a specific cis-acting element in its promoter region named the M-specific activator (MSA) and by the trans-acting Myb-related proteins (NtMybA1, NtMybA2, and NtMybB1), which bind to this element. Vertebrates have three types of Myb proteins. All contain three Myb repeats, which are DNA-binding

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Fig. 4. Involvement of the NACK-PQR pathway in plant cytokinesis. A The NACK-PQR pathway in tobacco. B Generation of multinucleate cells with incomplete cell plates upon expression of a motor domain-less NACK1 (NACK1:ST; top panel), kinase-defective NPK1 (NPK1:KW; second panel from top), and kinase-defective NQK1 (NQK1:KW; third panel from top). Control BY-2 cells are shown in the bottom panel. Cells were stained with calcofluor (blue; cell wall) and propidium iodide (red; nucleus). Arrowheads Incomplete cell plates; N nucleus. Bars 20 µm

domains, and they are thought to have important roles in the cell cycle. In plants, most of the Myb proteins have only two Myb repeats, whereas NtmybA1, NtmybA2, and NtmybB have three imperfect repeats, and each repeat is closely related to the vertebrate Myb repeats (Ito et al. 2001). NACK1 mRNAs and NACK1 proteins accumulate specifically at the late M phase of the cell cycle (Nishihama et al. 2002). The promoter of the NACK1 gene, which has a similar pattern of expression as the B-type cyclin genes, has two MSA elements through which NtMybA1 and NtMybA2 promote its transcription at the G2-M phase (Ito et al. 2001). NtMybA1 and NtMybA2 are also transcriptionally controlled at this phase of the cell cycle. Recently, it has been reported that the abilities of these proteins to activate transcription is regulated by CDK-mediated phosphorylation (Araki et al. 2004). We propose that the initiation of cytokinesis can be transcriptionally controlled by a chain of M-phase-promoting mechanisms, which can be triggered by activation of CDKs. Because there are temporal differences between the pattern of coaccumulation and the activation of NACK1 and NPK1 proteins (i.e., both NACK1 and NPK1 proteins are accumulated at metaphase in M phase, whereas activation of NPK1 occurs after late anaphase in M phase), this model does not seem to be sufficient to explain the mechanisms by which the NACK-PQR pathway is activated only during cytokinesis.

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We have also shown that NPK1 proteins are phosphorylated until just before their activation, and, conversely, they are activated following their dephosphorylation (Nishihama et al. 2001), suggesting that posttranslational modifications such as phosphorylation are involved in the activation of NPK1. Although it is not clear which kinases are involved in the phosphorylation of NPK1 proteins, CDKs are candidates because the pattern of CDK activation coincides with the phosphorylation of NPK1 and because CDKs phosphorylate NPK1 in vitro (our unpublished observations). NACK1 proteins also have some CDK phosphorylation site sequences (Fig. 2A). During late anaphase and telophase, NACK1 is consistently localized at the equatorial zone of the phragmoplast, whereas NACK1 is dispersed in the cytoplasm from prometaphase to early anaphase, yielding patchy signals, which suggests that NACK1 binds MTs in the phragmoplast but not the spindle. Perhaps phosphorylation of NACK1 by CDKs and/or other protein kinases might change the ability of NACK1 to bind MTs. Recently, Weingartner et al. (2004) reported that overexpression of the constitutively active form cyclin B1 caused a failure in proper localization of NACK1 on phragmoplast MTs at anaphase or telophase; in other words, NACK1 proteins remained throughout the cytoplasm. This supports the hypothesis that cyclin/CDK complexes direct NACK1 to activate the NACK-PQR pathway. Which protein kinases phosphorylate NACK1 or NPK1 and how they regulate functions of these proteins remain to be determined.

5 Downstream Factors of the NACK-PQR Pathway In order to understand the mechanism that controls the progression of cytokinesis, we are currently searching for proteins that are downstream factors of the NACK-PQR pathway. We have found that, in BY-2 cells, overexpression of the dominant negative mutants NACK:ST, NPK1:KW, and NQK1: KW inhibits the expansion of the phragmoplast and the cell plates but does not affect phragmoplast formation (Nishihama et al. 2001, 2002; Soyano et al. 2003; Fig. 4). The phenotypes of these cells are similar to those of cells treated with taxol, a compound that blocks the depolymerization of MTs (Yasuhara et al. 1993). This suggests that MT disassembly is required for phragmoplast expansion. Strompen et al. (2002) also showed that loss-of-function mutations in AtNACK1/HIK result in the persistence of phragmoplast MTs in the center of the division plane from where they usually disappear after formation of the cell plate. Based on these results, we propose that activation of the NACKPQR pathway is required for reorganization of phragmoplast MTs, especially depolymerization of MTs inside the phragmoplast, during cell plate formation. Both genetic and physiological evidence suggests that appropriate organization and/or dynamics of MTs are critical for the progression of cytokinesis (Nishihama and Machida 2001; Mayer and Jürgens 2002). In animals, several components belonging to the KLP or microtubule-associated protein (MAP)

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families have been shown to regulate the formation or dynamics of the central spindle, which corresponds to the phragmoplast. In plants, however, few factors involved in phragmoplast dynamics have been identified. MAP65 (NtMAP65), an MT-binding protein identified in tobacco BY-2 cells, is one of the few factors involved in MT dynamics in plants (Chang-Jie and Sonobe 1993). This family of proteins is conserved among a variety organisms and includes Ase1p (anaphase spindle elongation factor) in yeast (Pellman et al. 1995), PRC1 (protein regulator of cytokinesis 1) in mammals (Jiang et al. 1998), SPD1 (spindle defective 1) in C. elegans (Verbrugghe and White 2004), and Feo (Fascetto) in Drosophila (Vernì et al. 2004). These MAPs localize to the cytokinetic apparatuses, and most of them are involved in cytokinesis. In Arabidopsis, The pleiade (ple) mutant was identified as a root-specific mutant defective in cytokinesis (Müller et al. 2002). More recently, the PLE gene was found to encode the AtMAP65-3 protein, which is a divergent member of the Arabidopsis MAP65 gene family (Müller et al. 2004). These results show that AtMAP65-3/PLE proteins also play an essential role in plant cytokinesis. Although members of the MAP65/Ase1p/PRC1 protein family have low amino acid sequence similarities, the secondary structures in their predicted coiled-coil regions are highly conserved (Schuyler et al. 2003). These proteins are enriched at the spindle midzone in yeast, at the central spindle midzone in animals, and at the phragmoplast in plants, and they may maintain the distribution of the spindle or phragmoplast MTs and their function to progress or complete cytokinesis (Jiang et al. 1998; Smertenko et al. 2000, 2004; Mollinari et al. 2002; Schuyler et al. 2003; Müller et al. 2004; Verbrugghe and White 2004; Vernì et al. 2004). Although the MAP65/Ase1p/PRC1 protein family has the ability to bind and bundle MTs, it is not yet clear how these proteins participate in the regulation of MT organization in vivo or how their functional activity is controlled. One exception to this is mammalian PRC1, which was identified as a CDK substrate. Its phosphorylation by CDK seems to be important for mitotic suppression of MT bundling (Mollinari et al. 2002). Recently, we have found that some MAPs were phosphorylated by NRK1 MAPK, suggesting that they are candidates of downstream factors of the NACK-PQR pathway (our unpublished observations). One of these proteins was identified as NtMAP65-1a, a protein belonging to the MAP65/Ase1p/PRC1 family. NRK1 was able to phosphorylate recombinant NtMAP65-1a to stoichiometric levels (our unpublished observations). In agreement with others’ observations (Smertenko et al. 2000; Fig. 5), we have found that a green fluorescent protein (GFP)-NtMAP65-1a fusion localizes on several MT structures, including cortical MTs, the preprophase band, and the phragmoplast. As shown in Fig. 5, the distribution of NtMAP65-1a in the phragmoplast appears wider than that of NACK1, NPK1, NQK1, and NRK1, which accumulate only at the equator of the phragmoplast. It will be important to characterize functions and the subcellular localization of NRK1-phosphorylated NtMAP65-1a in order to elucidate the relationship between the NACK-PQR pathway and NtMAP65-1a. Another gene that encodes MAP localized on the phragmoplast was iden-

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Fig. 5. Subcellular localization of NtMAP65-1a proteins at telophase in BY-2 cells. BY-2 cells expressing GFP-NtMAP65-1a (GFP; green) are shown in the left panels and Nomarski differential interference contrast (DIC) images are shown in the center panels. Merged images are shown in the right panels

tified from the gemini pollen1 (gem1) mutant of Arabidopsis, whose mutation causes various degrees of defects in cytokinesis during pollen mitosis I (Park et al. 1998; Twell et al. 2002). The GEM1 gene, which is identical to the MICROTUBULE ORGANIZATION 1 gene (MOR1) (Whittington et al. 2001), encodes a protein that is related to the MAP215 family of MAPs found in all eukaryotes. In mor1 mutants, the organization and the orientation of cortical MTs is disrupted (Whittington et al. 2001), suggesting that the loss of MOR1/GEM1 function causes an increase in MT instability. MOR1/GEM1 seems to have a similar role to that of XMAP215, the Xenopus homologue of MOR1/GEM1, which is known to stabilize MTs (Kinoshita et al. 2002). During cytokinesis, MOR1/GEM1 is also thought to regulate the organization of the phragmoplast MTs, and to be involved in cytokinesis (Twell et al. 2002). It will be interesting to examine whether NRK1 can phosphorylate MOR1/GEM1 proteins. Twenty-two proteins that are conserved in other organisms have been identified as the participants in animal cell cytokinesis. Eight of these, including PRC1, are found on the central spindle, and they appear to interact with each other and participate in some aspect of cytokinesis (Glotzer 2005). Although it is unknown whether MOR1/GEM1 is phosphorylated as part of the NACKPQR pathway, these proteins found on the phragmoplast might influence plant cytokinesis by directly or indirectly interacting with each other. Further studies and identification of proteins phosphorylated by NRK1 should help elucidate the mechanism by which the NACK-PQR pathway controls phragmoplast expansion.

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6 Future Perspectives Over the past decade, we have demonstrated that the MAPK cascade activated by the NACK1/NPK1 complex (the NACK-PQR pathway) plays an important role in phragmoplast expansion by regulating MT turnover during cytokinesis. The fact that MAPs are phosphorylated by NRK1 MAPK supports the involvement of this MAPK cascade in maintenance of the phragmoplast. This also suggests that several proteins and their phosphorylation, including MAP65 and MOR1/GEM1, regulate MT stability in both plants and animals. Many components involved in cytokinesis have recently been identified, and, intriguingly, despite different modes of cytokinesis in the various kingdoms, most of them are conserved. This indicates that the system for regulating the cytoskeletal structures involved in cytokinesis, such as the central spindle and phragmoplast, may be conserved among various organisms. Our and others’ results further suggest that control of balance of MT and actin stability and instability in these cytoskeletal structures is essential for their function in cytokinesis. Why the PQR MAPK cascade was adopted for the regulation of cytokinesis in plants is an intriguing question. There must be a tight relationship between activation and inactivation of the NACK-PQR pathway and MT turnover. An association of MAPKs with MT and actin cytoskeletons and/or MT- or actinassociated proteins has been found in animal, plant, and yeast cells. MAPK cascades are known to regulate a wide range of crucial cellular processes, including cell division, polarization, and biotic or abiotic stress responses. In addition, most of these responses are accompanied by dynamic changes in cytoskeletal distribution, which are likely to be regulated by MAPK-mediated phosphorylation of MT- or actin-associated proteins (Šamaj et al. 2004). On the other hand, many experiments using pharmacological inhibitors or disruptors of the cytoskeleton have shown that rearrangement of cytoskeleton also stimulates MAPK activity (Soyano et al. 2003; Šamaj et al. 2004), suggesting that there is a bidirectional signaling pathway between the MAPK pathways and cytoskeletal organization; in other words, both the MAPK pathways and the cytoskeleton may act as sensors and effectors (Soyano et al. 2003; Šamaj et al. 2004). The direct activation of NPK1 MAPKKK by NACK1 KLPs demonstrates that the MAPK cascade is activated by components of the cytoskeleton. Further identification and investigation of molecules that act upstream or downstream of the NACK-PQR pathway should help clarify the biological relevance of the MAPK cascade in cytokinesis. Acknowledgements. This work was supported in part by a grant from the Program for Promotion of Basic Research Activities for Innovative Biosciences, by a Grant-in-Aid for Scientific Research on Priority Areas (no. 14036216), and by a Grant-in-Aid for the 21st Century COE Program (System Bioscience) from the Ministry of Education, Culture, Sports, Science and Technology of Japan.

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Smertenko A, Saleh N, Igarashi H, Mori H, Hauser-Hahn I, Jiang CJ, Sonobe S, Lloyd CW, Hussey PJ (2000) A new class of microtubule-associated proteins in plants. Nat Cell Biol 2:750–753 Smertenko AP, Chang HY, Wagner V, Kaloriti D, Fenyk S, Sonobe S, Lloyd C, Hauser MT, Hussey PJ (2004) The Arabidopsis microtubule-associated protein AtMAP65-1: molecular analysis of its microtubule bundling activity. Plant Cell 16:2035–2047 Sonobe S, Nakayama N, Shimmen T, Sone Y (2000) Intracellular distribution of subcellular organelles revealed by antibody against xyloglucan during cell cycle in tobacco BY-2 cells. Protoplasma 213:218–227 Soyano T, Nishihama R, Morikiyo K, Ishikawa M, Machida Y (2003) NQK1/NtMEK1 is a MAPKK that acts in the NPK1 MAPKKK-mediated MAPK cascade and is required for plant cytokinesis. Genes Dev 17:1055–1067 Spielman M, Preuss D, Li FL, Browne WE, Scott RJ, Dickinson HG (1997) TETRASPORE is required for male meiotic cytokinesis in Arabidopsis thaliana. Development 124:2645–2657 Strompen G, El Kasmi F, Richter S, Lukowitz W, Assaad FF, Jürgens G, Mayer U (2002) The Arabidopsis HINKEL gene encodes a kinesin-related protein involved in cytokinesis and is expressed in a cell cycle-dependent manner. Curr. Biol 12:153–158 Tanaka H, Ishikawa M, Kitamura S, Takahashi Y, Soyano T, Machida C, Machida Y (2004) The AtNACK1/HINKEL and STUD/TETRASPORE/AtNACK2 genes, which encode functionally redundant kinesins, are essential for cytokinesis in Arabidopsis. Genes Cells 9:1199–1211 Twell D, Park SK, Hawkins TJ, Schubert D, Schmids R, Smertenko A, Hussey PJ (2002) MOR1/GEM1 has an essential role in the plant-specific cytokinetic phragmoplast. Nature Cell Biol 4:711–714 Verbrugghe KJ, White JG (2004) SPD-1 is required for the formation of the spindle midzone but is not essential for the completion of cytokinesis in C. elegans embryos. Curr Biol 14:1755–1760 Vernì F, Somma MP, Gunsalus KC, Bonaccorsi S, Belloni G, Goldberg ML, Gatti M (2004) Feo, the Drosophila homolog of PRC1, is required for central-spindle formation and cytokinesis. Curr Biol 14:1569–1575 Waizenegger I, Lukowitz W, Assaad F, Schwarz H, Jürgens G, Mayer U (2000) The Arabidopsis Knolle And KEULE genes interact to promote vesicle fusion during cytokinesis. Curr Biol 10:1371–1374 Weingartner M, Criqui MC, Meszaros T, Binarova P, Schmit AC, Helfer A, Derevier A, Erhardt M, Bogre L, Genschik P (2004) Expression of a nondegradable cyclin B1 affects plant development and leads to endomitosis by inhibiting the formation of a phragmoplast. Plant Cell 16:643–657 Whittington AT, Vugrek O, Wei KJ, Hasenbein NG, Sugimoto K, Rashubrooke MC, Wasteneys GO (2001) MOR1 is essential for organizing cortical microtubules in plants. Nature 411 610–613 Wu C, Whiteway M, Thomas DY, Leberer E (1995) Molecular characterization of Ste20p, a potential mitogen-activated protein or extracellular signal-regulated kinase kinase (MEK) kinase kinase from Saccharomyces cerevisiae. J Biol Chem 270:15984–15992 Yang CY, Spielman M, Coles JP, Li Y, Ghelani S, Bourdon V, Brown RC, Lemmon BE, Scott RJ, Dickinson HG (2003) TETRASPORE encodes a kinesin required for male meiotic cytokinesis in Arabidopsis. Plant J 34:229–240 Yasuhara H, Sonobe S, Shibaoka, H (1993) Effects of taxol on the development of the cell plate and of the phragmoplast in tobacco BY-2 cells. Plant Cell Physiol 34:21–29 Yasuhara H, Sonobe S, Shibaoka H (1995) Effects of brefeldin A on the formation of the cell plate in tobacco BY-2 cells. Eur J Cell Biol 66:274–281 Yokoyama R, Nishitani K (2001) Endoxyloglucan transferase is localized both in the cell plate and in the secretory pathway destined for the apoplast in tobacco cells. Plant Cell Physiol 42:292–300 Zuo J, Niu QW, Nishizawa N, Wu Y, Kost B, Chua NH (2000) KORRIGAN, an Arabidopsis endo1,4-beta-glucanase, localizes to the cell plate by polarized targeting and is essential for cytokinesis. Plant Cell 12:1137–1152

Section II Physiological and Developmental Aspects

II.1 Characterization of a Cell Division Factor from Auxin-Autotrophic 2B-13 Cells Derived from the Tobacco BY-2 Cell Line T. Shimizu1 , K. Eguchi1 , I. Nishida1 , K. Laukens2 , E. Witters2 , and T. Nagata1

1 Introduction The tobacco BY-2 cell line has been used as a model plant cell line in various aspects of plant science studies (Nagata et al. 1992, 2004) and is described further in other chapters of this book. In this chapter, however, we describe a cell line named 2B-13 derived from the BY-2 cell line. The 2B-13 cell line has been established as an auxin-autotrophic cell line from the BY-2 cell line by selection on auxin-free medium. Its growth rate has been significantly accelerated by UV light illumination (Noguchi et al. 1977). Thus the 2B-13 cell line can be considered as a habituated cell line according to the definition by Roger Gautheret (Meins 1982). Since 1988 we have compared the growth characteristics of the 2B-13 cell line with those of BY-2 in order to understand the auxin-autotrophic nature of 2B-13 cells. There are no apparent genetic differences between the two cell lines except for auxin requirements. Absolute auxin requirement for the proliferation of tobacco BY-2 cells has been well characterized. When auxin is depleted from the culture medium, cells cease to divide by the fourth day of culture. Upon the addition of auxin to the auxin-starved non-dividing BY-2 cells, semi-synchronous cell division is induced (Ishida et al. 1993). It has also been demonstrated that the auxin-starved cells are arrested at the G1 phase in the cell cycle (Nagata et al. 1999). Although clues to understanding the differences between the two cell lines regarding auxin requirements have not been easily obtained, it was recently found that the addition of culture filtrates of 2B-13 cells could induce cell division in the auxin-starved BY-2 cells, inducing semi-synchronous cell division. The time course of the induction of cell division by the addition of culture filtrates of 2B-13 cells was very similar to the time course observed for the addition of auxin to the auxin-starved tobacco BY-2 cells. This observation suggests that certain factors in the culture filtrates could be replaced by the action of auxin. Thus we conducted experiments on the characterization and purification of factors that could be replaced by auxin in the culture filtrates of 2B-13 cells. In this chapter, we describe the outcome of our trial for the purification of this factor. The significance of this factor to our understanding of auxin habituation 1 Department

of Biological Sciences, Graduate School of Science, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan, e-mail: [email protected] 2 Department of Biology, University of Antwerp (UA), Groenenborgerlaan 171, 2020, Antwerp, Belgium Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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is described. Further potential uses of this cell line in the plant sciences are also discussed.

2 Characterization of Cell Division-Inducing Activity in the Culture Filtrates of 2B-13 Cells When auxin is added to auxin-starved non-dividing BY-2 cells, induction of semi-synchronous cell division is observed (Ishida et al. 1993). Under this condition, when the culture filtrates of the 2B-13 cells were added to the auxinstarved cells, significant induction of cell division was observed (Fig. 1). This evidence seems to imply that the culture filtrates retain some factors that could be replaced by the effect of auxin. It should be noted in this context that some cases have been reported in which the amounts of detected plant growth regulators in cells and their milieu were higher than the control cells, as exemplified in the habituated cell line that was induced by irradiation of γ -rays (Campell and Town 1991) and by chemical mutagenesis (Frank et al. 2000). Therefore we examined whether the culture filtrates of 2B-13 cells retained higher amounts of auxin or cytokinins. As demonstrated in Table 1, auxin and cytokinin concentrations that were measured by GC-MS analysis in the culture filtrates were lower than or at least comparable to that of the BY-2 cell line. Furthermore, as the contents of auxin and cytokinins measured in the cells were also lower than those observed in the control BY-2 cells, increased biosynthetic activity of these two plant hormones could be excluded. This implies that the cell division-inducing activity observed in the culture filtrates of BY-2 cells may be attributed to factors other than plant growth regulators, such as auxin or cytokinins of low molecular masses. Thus, we sought for factors in the culture filtrate that were responsible for inducing cell division in the auxin-starved BY-2 cells. In order to examine this, the culture filtrates were treated with trypsin (Sigma, St. Louis, USA) or

Fig. 1. Cell division-inducing activity in the culture filtrates of 2B-13 cells. Auxinstarved tobacco BY-2 cells were prepared from the stationary phase cells that had been cultured for 3 days after washing with auxin-free medium. To these cells, 2,4-D (0.2 mg/l) or culture filtrates were added. As a control, water was added to these cells. The time course of induction of cell division was plotted

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Table 1. Quantitative measurements of auxin and cytokinins in culture filtrates and cells of tobacco 2B-13 cells and BY-2 cells A. Intracellular concentrations (pmol/g fresh wt.) IAA

Z

ZR

2B-13 cells 15

219

312

BY-2 cells

265

295

50

ZMP 27.5 218

B. Extracellular concentrations (pmol/g fresh wt.) IAA 2B-13 cells Less than 1 BY-2 cells

Z

ZR

ZMP

–

–

–

Less than 0.5 0.5–5.0 0.5–5.0 0.5–5.0

glycosidases to characterize structural features of cell division factors contained in the culture filtrates of 2B-13 cells. When the culture filtrates were treated with trypsin, cell division activity demonstrated by the culture filtrates was completely nullified, while that treated with heat-denatured trypsin was not (Fig. 2). This implies that the cell division factor contains peptide bonds. Moreover, when the cell culture filtrates were treated with highly purified Nglycopeptidase F (Takara Bio Inc., Shiga, Japan), cell division activity was also nullified, implying that the factor retained glycoside moiety in the protein for its biological activity (Fig. 2). Thus, the cell division factor (CDF) that causes cell division in auxin-starved tobacco BY-2 cells is likely to be a glycoprotein, which will be referred to as CDF hereafter.

Fig. 2. Treatment of culture filtrates with trypsin or glycopeptidase. Left To the auxin-starved tobacco BY-2 cells was added 2,4-D, water, culture filtrated treated with trypsin, buffer or denatured trypsin. Right To the auxin-starved tobacco BY-2 cells was added 2,4-D, water, culture filtrated treated with glycopeptidase F, buffer, or denatured glycopeptidase

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3 Purification of the CDF When we purified the CDF from the culture filtrates, CDF activities in the respective purification step were assessed, with the cell division-inducing activity observed upon addition to the auxin-starved tobacco BY-2 cells. First the culture filtrates of 2B-13 cells were applied to a DEAE-Sephacel ion exchange column and subsequently washed with the dissolved buffer. The CDF was bound to the column, while it was not found in the flowthrough fraction, as shown in Fig. 3. Washing the column with the buffer (pH 8.5) containing 1 M NaCl caused the release of the CDF fraction from the column (Fig. 3). However, at lower pH ranges of the buffer, this active fraction was not eluted, implying that the isoelectric point (pI) of the CDF was in the alkaline range and that CDF may retain certain basic amino acids in its structural components. This information instructed us in how to purify the CDF. In fact, we carried out the purification of the CDF by conducting consecutive chromatographies of hydroxyapatite, ConA affinity column and gel filtration with Superdex. After the culture filtrates were applied to the hydroxyapatite column, the CDF was recovered by elution with the buffer containing 1 M KCl. This procedure seemed to be important in order to remove contaminating polysaccharides from the culture filtrates, as the subsequent purification was significantly facilitated by this procedure. The CDF fraction was then applied to a ConA Sepharose 4B column. The CDF was eluted from the column with elution of buffer containing 0.5 M NaCl and 0.2 M methyl-α-d-glucoside. This result was consistent with the preliminary notion that there should be sugar moieties in its structure, whose presence is necessary for its biological functionality. When the CDF was further applied to the gel filtration using a Superdex 200 column, the active fraction was recovered in the elution volumes of 87−89 ml. Starting with 40 l of 3-day-old 2B-13 cells, the amounts of protein measured after the hydroxyapatite column elution, affinity purification and gel filtration were 4 mg, 20 µg and a few nanograms, respectively, according to

Fig. 3. Characterization of cell division factor (CDF). After culture filtrates of 2B-13 cells were applied to the DEAE-Sephacel column, the fractions that were obtained by the elution buffer (pH 8.5) containing 1 M NaCl displayed cell division activity, while the flowthrough fraction did not

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determination by the Bradford method. Thus, proteins were purified around 106 fold. As the extent of purification appeared to become significantly high by this sequential purification procedure, an aliquot of the purified fraction retaining the CDF activity was applied to SDS-PAGE. As shown in Fig. 4, bands in the SDS-PAGE detected by silver staining were few – one major band and a few other faint bands. The molecular size of this major band was estimated to be around 30 kDa (Fig. 4). This protein was trypsinized and the resulting peptides were analyzed by MALDI-TOF-MS. The resulting peptide mass fingerprint revealed a match with a probability of 84% against a P-glycoprotein (gi|6911161, MW 34565, pI 9.82, Gossypioides kirkii). Individual peptides were analyzed by MALDITOF/TOF-MS. Of the MSMS spectra, a peptide with 960 m/z gave a significant hit (p < 0.05) against a fragment of the glycoprotein EP1 gi|3434308, although a hit against a similar peptide of a P-glycoprotein (gi|51892223) was also obtained. Although the quality of the MSMS data was good, the low representation of tobacco and related species in public sequence databases (further discussed in Chapter VI.3, this vol.) resulted in a low matching probability, making it difficult to exactly identify the protein isolated thus far. Nevertheless, glycoproteins are found among the best hits and their physicochemical properties (MW, pI) correspond well with the observed values (Shimizu et al., 2006). Mass spectrometric analysis suggests that the CDF that could induce cell division in auxin-starved tobacco BY-2 cells could be some kind of P-glycoprotein.

Fig. 4. SDS-PAGE of the purified fraction from the culture filtrates of 2B-13 cells. Culture filtrates of 2B-13 cells were purified successively by column chromatographies of hydroxyapatite, ConA Sepharose, and Superdex in this sequence. Elute that was obtained by passage through Superdex was applied to SDS-PAGE. The electropherogram was observed after silver staining. A major band (indicated by the arrow) was estimated to be a molecular size of 30 kDa. Left lane shows molecular size markers

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P-glycoproteins belong to a gene family of the ATP-binding cassette (ABC) transporter. It should be noted in this context that some of the ABC-transporter genes in plants have been reported to have a certain link with auxin-signaling pathways; AtMDR1 interacts with PIN1 whose role is reported as a key function in polar auxin transport (Noh et al. 2001), while AtMDR5 is reported to have a role in sequestering auxin-conjugates in the vacuole (Gaedeke et al. 2001). Thus, it should be noted that the CDF was identified in the culture filtrates of 2B-13 cells, while MDRs are usually integrated in the cell membrane, implying that they retain transmembrane domains in their structure. It is intriguing to examine how the CDF is secreted into the medium. Regarding this, the eight peptide masses that were identified by MALDI-TOF-MS are located in a specific domain in a P-glycoprotein-like protein which is outside of the transmembrane domain. A possible explanation for this is that the specific domain could be cleaved by certain mechanisms, such as protein splicing, and secreted into the medium. Thus the CDF would have the ability to induce cell division in auxinstarved tobacco BY-2 cells. However, it remains to be determined whether the CDF has any relationship with either of these AtMDRs. It would be interesting to determine the role of the CDF in auxin-signaling pathways. Conversely, it is certain that the CDF that seems to be secreted from auxinautotrophic 2B-13 cells can induce cell division in auxin-starved tobacco BY-2 cells. The CDF is thought to have some relationship with auxin habituation. Auxin habituation was first discovered by Roger Gautheret in 1942; however, its molecular mechanism remains unclear. One of the reasons for this is that there is no molecular marker for habituation (Meins 1989). In this regard, the CDF identified in this study is a likely candidate for the molecular marking of the habituation. To prove this, however, further extensive works are required. Nonetheless, if this hypothesis is correct, elucidation of the molecular mechanism of habituation using this CDF as a molecular marker would be significantly advanced.

4 Searches for CDFs in the Culture Filtrates of Tobacco BY-2 Cells It is apparent that auxin-autotrophic 2B-13 cells secrete a glycoprotein that could induce cell division in auxin-starved tobacco BY-2 cells. We investigated further whether tobacco BY-2 cells could produce and secrete similar factors by themselves. To this end, however, we were not able to add the culture filtrates of tobacco BY-2 cells directly to the auxin-starved BY-2 cells, since the culture filtrates of BY-2 cells contain 2,4-D. Thus we conducted a partial purification of cultured filtrates of tobacco BY-2 cells by hydroxyapatite column chromatography, as described above. This procedure was performed basically to remove 2,4-D from the culture filtrates. When we added the partially purified fractions to the auxin-starved BY-2 cells, induction of cell division by this fraction was

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observed (Shimizu et al., in preparation). Thus it is certain that tobacco BY-2 cells secrete some factors that could be comparable to the 30 kDa protein from 2B-13 cells which induced cell division in auxin-starved BY-2 cells. Therefore, we applied the same purification procedure that was used for the purification of the 30 kDa protein mentioned above; namely, hydroxyapatite chromatography, ConA affinity chromatography and gel filtration with a Superdex column in this sequence. When we added the respective fractions eluted from the Superdex column to the auxin-starved tobacco BY-2 cells, we found two fractions in which cell division-inducing activities were detected (Shimizu et al., in preparation). Intriguingly, these two fractions independently brought about cell division activities in the auxin-starved tobacco BY-2 cells. These two fractions were named CDF-A and CDF-B, respectively. SDS-PAGE revealed that CDF-A and CDF-B display single bands in the respective SDS-PAGE. The molecular sizes of CDF-A and CDF-B were 25 and 40 kDa, respectively (Shimizu et al., in preparation). As the glycosidase treatment cancelled out the cell division-inducing activities of these two factors, these two proteins should retain sugar moieties in their structures and the presence of sugar moieties is required for their biological activity. The two glycoproteins from BY-2 cells and a 30 kDa glycoprotein from 2B-13 cells seem to have very similar biological characteristics; however, structural relationships among these three glycoproteins have not yet been determined.

5 Concluding Remarks and Future Perspectives The 2B-13 cell line has been established as an auxin-autotrophic cell line from the BY-2 cell line. The CDF identified as the 30 kDa glycoprotein secreted from 2B-13 cells induced cell division in auxin-starved tobacco BY-2 cells. Thus the CDF is thought to play a significant role in cell cycle progression from G1 phase to S phase. Characterization of the CDF, which may be structurally classified as an ABC transporter, should give clues to understanding the molecular basis of habituation. Although the molecular mechanism of habituation, which was discovered more than 60 years ago, has not yet been clarified, using the CDF as a molecular marker could facilitate the molecular studies of habituation. Thus the CDF may be considered as the first candidate for molecularly marking habituation. In order to validate this hypothesis, however, further extensive studies are required. Moreover, the 2B-13 cell line, which grows vigorously without the addition of auxin, may be useful for examining and understanding the effects of exogenously applied plant hormones and other factors upon the growth characteristics and biochemistry of plant cells. It is also intriguing to compare differences in EST information and protein profiling between 2B-13 cells and BY-2 cells, as both cell lines are genetically almost identical.

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6 Protocols 6.1 Culture of Tobacco BY-2 and 2B-13 Cells Tobacco BY-2 cells were cultured in the modified Linsmaier and Skoog medium (1965) as described by Nagata et al. (1992). The tobacco 2B-13 cells that were derived from the BY-2 cell line were basically cultured under the same conditions as that of BY-2 cells, except that they were cultured in the Murashige and Skoog medium (1962) supplemented with 3% sucrose, but without auxin. 6.2 Auxin Starvation Auxin starvation of tobacco BY-2 cells was attained after 3 days of cultivation in the medium without auxin, as described in Nagata and Kumagai (1999). Cessation of cell division of tobacco BY-2 cells by auxin starvation was confirmed under fluorescence microscopy after staining their nuclei with 4
,6-diamidino2-phenylindole (DAPI). When auxin was added to the auxin-starved BY-2 cells, regain of meristematic activity was determined by the increase in the mitotic index under fluorescence microscopy after staining with DAPI. Culture filtrates of 2B-13 cells (20 ml) recovered from the 3-day-old 2B-13 cells and respective purified fractions from the culture filtrates were added to the auxin-starved BY-2 cells to assess the cell division-inducing activity of respective fractions. Cell division-inducing activity was assessed with the auxin-starved BY-2 cells by counting mitotic indices. 6.3 Quantification of the Contents of Auxin and Cytokinins Purification of auxin and cytokinins was carried out essentially as described in Redig et al. (1996). In brief, frozen cells were extracted overnight at –20 ◦ C in CHCl3 /CH3 OH/H2 O/CH3 COOH (5:12:2:1). The amounts of cytokinins and auxin in the organic solvent fraction were determined by LC-MS/MS according to Witters et al. (1999) and by GC-MS according to Prinsen et al. (2000), respectively. 6.4 Enzyme Treatments of Culture Filtrates from 2B-13 Cells The culture filtrate (20 ml) from 2B-13 cells was dialyzed with a dialysis membrane (Spectra-Por MWCO 1000, Spectrum Lab., Houston, USA) in 2 l of water at 4 ◦ C for 12 h. The retentate that was lyophilized completely and then dissolved in 1 ml of water was added to auxin-starved BY-2 cells to assess its cell division activity. Trypsin (Sigma) was dissolved in 20 mM of phosphate buffered saline (PBS) (pH 7.0) containing 150 mM NaCl to make up to 1.0 mg/l. After 20 ml of the culture filtrates was dialyzed and lyophilized, it was dissolved in 1.5 ml of PBS,

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to which was added 0.05 ml of the trypsin solution, inactivated trypsin, or buffer alone, respectively. Incubation was carried out at 37 ◦ C for 3 h. Enzyme reaction was stopped by heat treatment at 100 ◦ C for 10 min. The filtrates were then dialyzed as described above and their cell division activities were assayed with auxin-starved BY-2 cells. Treatment of the culture filtrates with glycopeptidase F (Takara Bio Inc., Shiga, Japan) dissolved in 1 M Tris-HCl buffer (pH 8.6) was carried out basically as described for trypsin treatment. 6.5 Purification of CDF The culture filtrates that were lyophilized and dissolved in 10 mM K-phosphate buffer (pH 7.2) were reacted with 50 g of hydroxyapatite (Seikagaku-Kogyo, Tokyo, Japan) which had been equilibrated with 10 mM K-phosphate buffer (pH 7.2). After the column was washed with the equilibration buffer, fractions were eluted first with the buffer containing 1 M KCl and then with 0.3 M K-phosphate buffer (pH 7.2). The active fraction that was eluted by the buffer containing 1 M KCl was further applied to the column loaded with Con A Sepharose 4B (Amersham Pharmacia Biotech) at a flow rate of 0.2 ml/min. Con A Sepharose was equilibrated with 0.02 M Tri-HCl buffer (pH 7.4) containing 0.5 M NaCl. After the column had been washed with 5 ml of the equilibration buffer, fractions were eluted with 10 ml of the buffer containing 0.1 M glucose, 0.2 M methyl-α-d-glucoside, 0.5 M methyl-α-d-glucoside, and 0.2 M methylα-d-mannoside at a flow rate of 1 ml/min, being monitored with absorbance at 280 nm. Each fraction that was dialyzed was assayed for cell division activity with auxin-starved BY-2 cells. The biologically active fraction from the Con A Sepharose 4B column whose buffer was exchanged with 50 mM K-phosphate buffer (pH 7.2) containing 150 mM NaCl was concentrated to 1.2 ml with a concentrator (Vivaspin, 10,000 MWCO PES, Vivascience, Hannover). Subsequently after it was applied to HiLoad 16/60 Superdex 200 Prep Grad (Amersham Pharmacia Biotech), fractions were eluted with buffer at a flow rate of 1 ml/min, absorbance being monitored at 280 nm. Collected fractions (2 ml) that were dialyzed were assayed with cell division activity using auxin-starved BY-2 cells. An active peak eluted at 88 min was used for further amino acid sequence determination. Proteins purified as described above were separated by SDS-PAGE and visualized with silver staining. 6.6 MALDI-TOF/TOF MS The purified fraction was digested in solution using trypsin (Promega MS Gold). The resulting peptide mixture was further purified using Millipore ZipTips(C18) and data acquired by MALDI-TOF-MS/MS (4700 Proteomic Analyzer, Applied Biosystems). Processed data were compared with public databases using the Mascot matching algorithm as described by Laukens et al. (2004) and outlined in detail in Chapter VI.1 of this book.

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Acknowledgements. We wish to thank Somatech Center, House Food Co. (Yotsukaido-shi, Chiba, Japan) for their help in lyophilizing the culture filtrates of tobacco cells. This study was supported in part by a grant from the Japan Society for the Promotion of Science.

References Campell BC, Town CD (1991) Physiology of hormone autonomous tissue lines derived from radiation-induced tumors of Arabidopsis thaliana. Plant Physiol 97:1166–1173 Frank M, Rupp H-M, Prinsen E, Motyka V, Van Onckelen H, Schmülling T (2000) Hormone autotrophic growth and differentiation identifies mutant lines of Arabidopsis with altered cytokinin and auxin content or signaling. Plant Physiol 122:721–729 Gaedeke N, Klein M, Kolukisaoglu U, Forestier C, Mueller A, Ansorge M, Becker D, Magnum Y, Kuchler K, Schutz B, Mueller-Roeber B, Martinola E (2001) The Arabidopsis thaliana ABC transporter AtMDR5 controls root development and stomata movement. EMBO J 20:1875– 1887 Ishida S, Takahashi Y, Nagata T (1993) Isolation of an auxin-regulated gene encoding a G protein β subunit-like protein from tobacco BY-2 cells. Proc Natl Acad Sci USA 90:11152–11156 Laukens K, Deckers, P, Esmans E, Van Onckelen H, Witters E (2004) Construction of a twodimensional gel electrophoresis protein database for the Nicotiana tabacum cv. Bright Yellow2 cell suspension culture. Proteomics 4:720–727 Linsmaier L, Skoog F (1965) Organic growth factor requirements of tobacco tissue cultures. Physiol Plant 18:100–127 Meins F Jr (1982) Habituation of cultured plant cells. In: Schell J, Kahl G (eds) Molecular biology of plant tumors. Academic Press, New York, pp 3–31 Meins Jr F (1989) Habituation: heritable variation in the requirement of cultured plant cells for hormones. Annu Rev Genet 23:395–408 Murashige T, Skoog F (1962) A revised medium for rapid growth and bioassays with tobacco pith culture. Physiol Plant 15:473–497 Nagata T, Kumagai F (1999) Plant cell biology through the windows of highly synchronized tobacco BY-2 cell line. Methods Cell Sci 23:123–127 Nagata T, Nemoto Y, Hasezawa S (1992) Tobacco BY-2 cell line as the “HeLa” cell in the cell biology of higher plants. Int Rev Cytol 132:1–30 Nagata T, Ishida S, Nagata S, Takahashi Y (1999) Factors affecting cell division in plant cells. In: Altman A (ed) Plant biotechnology and in vitro biology in the 21st century. Kluwer, Dordrecht, pp 429–432 Nagata T, Hasezawa S, Inzé D (eds) (2004) Tobacco BY-2 cells. Biotechnology in agriculture and forestry, vol. 53. Springer, Berlin Heidelberg New York Noguchi M, Matsumoto T, Hirata Y, Yamamoto K, Katsuyama A, Kato A, Azechi S, Kato K (1977) Improvement of growth rates of plant cell culture. In: Barz W, Reinhard E, Zenk MH (eds) Plant tissue culture and its bio-technological application. Springer, Berlin Heidelberg New York, pp 85–94 Noh B, Murphy AS, Spalding EP (2001) Multidrug resistance-like genes of Arabidopsis required for auxin transport and auxin-mediated development. Plant Cell 13:2441–2454 Prinsen E, van Laer S, Öden S, van Onckelen H (2000) Auxin analysis. Meth Molec Biol 141:49–65 Redig P, Shaul O, Inze D, Van Montagu M, Van Onckelen H (1996) Levels of endogenous cytokinins, indole-3-acetic acid and abscisic acid during cell cycle of synchronized tobacco BY-2 cells. FEBS Lett 391:175–180 Shimizu T, Eguchi K, Nishida I, Laukens K, Witters E, Van Onckelen H, Nagata T (2006) A novel cell division factor from tobacco 2B-13 cells that induced cell division in auxin-starved tobacco BY-2 cells. Naturwissenschaften. in press Witters E, Vanhoutte K, Dewitte W, Machackova I, Benkova E, van Dangen W, Esmans E, van Onckelen H (1999) Analysis of cyclic nucleotides and cytokinins in minute plant samples using phase-system switching capillary ion spray LC-MSMS. Phytochem Anal 10:143–151

II.2 The BY-2 Cell Line as a Tool to Study Auxin Transport J. Petrášek and E. Zažímalová1,2

1 Introduction Auxin transport plays a key role in the regulation of plant growth and development. Either it runs in apoplast by mass flow in the phloem together with other metabolites and/or there exists a parallel, cell-to-cell, strictly directional, carrier-mediated active transport. The major contribution to our understanding of its physiology as well as molecular background comes from studies in planta. However, during the last ten years, tobacco cell lines such as BY-2 have provided a major impetus for precise analysis of the machinery performing auxin flow across cell membranes at the cellular level. In this chapter recent knowledge about the molecular mechanism of auxin transport is summarized, and the data are discussed in the context of recent findings concerning the role of directional cell-to-cell transport of auxin in plant development. The results obtained using BY-2 as well as other tobacco cell lines highlight the advantages of these models in studies of auxin-regulated processes such as cell division, elongation and establishment of cell polarity.

2 Present State of the Art of Cell-to-Cell Transport of Auxins Together with the processes of auxin biosynthesis, conjugation and degradation, unidirectional cell-to-cell transport of auxin (indole-3-acetic acid, IAA) plays a crucial role in the regulation of growth and development of plants (Blakeslee et al. 2005; Friml and Wisniewska 2005; Woodward and Bartel 2005). One of the most important features of auxin transport in the symplast is that it is unequivocally polar. The explanation of its polarity was simultaneously proposed by Rubery and Sheldrake (1974) and Raven (1975) and called the chemiosmotic polar diffusion model (Goldsmith 1977). According to this model, undissociated, a lipophilic form of native auxin molecule (IAA) can easily enter the cell cytoplasm from slightly acidic extracellular environment (pH 5.5) by passive diffusion. Since the pH of cytoplasm is more alkaline 1 Institute of Experimental Botany ASCR, Rozvojová 135, 16502 Prague 6, Czech Republic, e-mail:

[email protected] of Plant Physiology, Faculty of Science, Charles University, Vinièˇcná 5, 12844 Prague 2, Czech Republic

2 Department

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Fig. 1. Simplified scheme of auxin (IAA) transport at the cellular level. Both passive diffusion and specific auxin influx (uptake) and efflux carriers are involved in the transport of the IAA molecule and its dissociated form, auxin anion (IAA− ). In contrast to quite easy diffusion of relatively lipophilic molecules of IAA across the PM, hydrophilic IAA− anions can be transported only actively via carriers. One of the auxin transport inhibitors (ATIs), 1-naphthylphthalamic acid (NPA), has been proposed to act via binding to the complex of auxin efflux carrier. Moreover, ATIs may have broader effect on protein trafficking processes (Muday et al. 2003). The fungal toxin brefeldin A (BFA) influences polar auxin transport by disturbing Golgi-mediated and actindependent vesicle transport of putative auxin efflux carriers from the intracellular space to the PM (Geldner et al. 2001, 2003)

(pH 7), IAA molecules dissociate and the resulting hydrophilic auxin anions (IAA− ) are trapped in the cytosol. The exit or uptake of IAA− was proposed to be assisted by the auxin efflux and influx proteins, respectively, and the polarity of auxin transport was explained by their asymmetric distribution on the plasma membrane (PM) of the cell (Fig. 1). In the last decade, genes encoding putative auxin uptake (influx) and efflux carriers have been identified in Arabidopsis and other species (Morris 2000; Muday and DeLong 2001; Friml and Palme 2002; Morris et al. 2004; Paponov et al. 2005). Since the intracellular concentration of auxin is, among other things, critical for the setting of various developmental programmes, it is obvious that oriented transport of auxin plays an important role in the morphoregulatory and pattern formation processes such as embryogenesis (Friml et al. 2003), root development (Friml et al. 2002a; Blilou et al. 2005; reviewed in Teale et al. 2005), organ formation (Benková et al. 2003), vascular differentiation (Mattsson et al. 2003), tropisms (Friml et al. 2002b) and phyllotaxis (Reinhardt et al. 2003).

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3 Auxin Transport Studies in Planta 3.1 Auxin Transport Assays It has always been challenging to follow the auxin distribution in plants and to understand how it is established and maintained. Besides modern instrumental techniques for the quantification of endogenous auxin content (Ljung et al. 2005) and non-invasive tracking of the expression of auxin-responding reporter genes (Ulmasov et al. 1997), the direct measurement of radiolabelled auxin distribution has been one of the most frequently used approaches in studies of auxin transport (Goldsmith 1977). Upon application of labelled auxin to one end of a tissue segment, auxin movement is usually followed by measurement of the quantity of transported radiolabel. Although this approach was recently optimized for the whole Arabidopsis seedling (Geisler et al. 2003) it is not applicable for the measurement of auxin transport at the cellular level. 3.2 Inhibitors of Polar Auxin Transport One of the most fruitful approaches in studying the auxin transport machinery is the application of various inhibitors. The most widely used inhibitor of auxin efflux is 1-naphthylphthalamic acid (NPA), which belongs to a group of inhibitors known as phytotropins (Rubery 1990). The application of NPA to plant tissues results typically in an increase in auxin accumulation, presumably due to the inhibition of auxin efflux activity (Morris et al. 2004). Detailed knowledge about the mechanism, by which NPA and other phytotropins inhibit auxin efflux, is still lacking. NPA probably binds to a specific high affinity NPAbinding protein (NBP) located on the cytoplasmic face of the PM (Sussman and Gardner 1980), where it is associated with actin cytoskeleton (Cox and Muday 1994; Dixon et al. 1996; Butler et al. 1998). Recent evidence suggests that NBPs are aminopeptidases acting in cooperation with flavonoids, which are known as naturally occurring regulators of auxin efflux (Murphy et al. 2002). In addition, a more general, inhibitory effect of phytotropins on endocytotic processes was reported (Geldner et al. 2001). Another set of results that helped the understanding of auxin transport machinery comes from studies using fungal toxin brefeldin A (BFA), the inhibitor of Golgi-mediated vesicle trafficking and endosomal recycling. BFA inhibits auxin efflux activity in zucchini hypocotyls (Morris and Robinson 1998) and blocks auxin transport through the tissue (Robinson et al. 1999). Since BFA treatment effectively changes the proportion of proteins localized at the PM and in the endosomal space (reviewed in Geldner 2004), it is an ideal tool for studying constitutive cycling of both putative auxin efflux and uptake carriers.

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3.3 Genetic and Molecular Characterization of Components of the Auxin Transport Machinery Recent research using the model plant Arabidopsis thaliana led to the identification of proteins involved in the auxin transport machinery. These include promising candidates for both auxin uptake (influx) carrier, AUX1/LAXs (amino-acid permease-like proteins), and regulators of auxin efflux, PINs (PINformed, plant-specific PM proteins) and MDR/PGPs (multidrug-resistancelike/P-glycoproteins) (see Morris et al. 2004; Blakeslee et al. 2005; Friml and Wisniewska 2005). It seems that PIN proteins establish the direction of auxin flux by their asymmetric distribution at the PM. They may form complexes with other regulatory proteins and MDR/PGPs may stabilize these complexes (Blakeslee et al. 2005). The correct positioning of the auxin efflux complex seems to be assisted by the actin cytoskeleton. The application of actin drugs resulted in reduced polar auxin transport in maize coleoptiles (Cande et al. 1973) and in zucchini hypocotyls (Butler et al. 1998). Moreover, using BFA, it was shown that PIN proteins might undergo constitutive cycling between PM and the endosomal space, as indirectly suggested by Robinson et al. (1999), and that this process is actin-dependent (Geldner et al. 2001). It was shown that the mutation in Arabidopsis myosin VI led to the inhibition of basipetal auxin transport (Holweg and Nick 2004). All these observations strongly suggest that actin filaments are involved in both intracellular traffic of PINs and their correct targeting to the proper domains at the PM. The concept of trafficking and proper localization as well as function of components of the auxin efflux carrier complex has been proposed (summarized in Muday et al. 2003). In contrast to this, the mechanism(s) underlying the constitutive cycling of proteins in plants is still poorly understood (Murphy et al. 2005). Constitutive cycling of PINs is regulated by GNOM, one of the exchange factors for ADP-ribosylation factor-type GTPases (ARF-GEFs; Geldner et al. 2003, 2004) and other ARFs might also contribute to the trafficking of PINs (Xu and Scheres 2005). Interestingly, the regulation of endocytosis-dependent cycling of proteins in plant cells was shown to be regulated by auxin itself (Paciorek et al. 2005), suggesting a new mechanism of auxin action in plant cells.

4 Auxin Transport Studies in Simplified Models 4.1 Transport of Auxins in Suspension-Cultured Cells Since it is difficult to study the biochemical aspects of auxin transport by measurements at the whole-plant and organ level (see above), plant cells cultured in liquid medium represent a valuable alternative. The accumulation of auxin can be measured in time intervals after the direct addition of radiolabelled auxin

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Fig. 2. Differences in the translocation of IAA and two synthetic auxins NAA and 2,4-D in tobacco cell lines. A Auxins are translocated across PM according to their relative lipophilicity (NAA > IAA > 2, 4-D). As shown by careful measuring of diffusion and carrier-mediated transport of NAA, IAA and 2,4-D in tobacco suspension-cultured Xanthi XHFD8 cells by Delbarre et al. (1996), the accumulation level of NAA is controlled mainly by efflux carrier activity, while the accumulation of 2,4-D is determined by the activity of an uptake carrier. Both uptake and efflux carriers, as well as passive diffusion, contribute to the accumulation of IAA. Refer to Fig. 1 for symbol legends. B The accumulation of all three types of auxins (IAA, NAA, 2,4-D) is increased upon application of inhibitors NPA or BFA, thus reflecting disturbed auxin efflux machinery in BY-2 cells (Petrášek et al. 2005)

into the cell suspension. Indeed, the most important experiments, on which the chemiosmotic hypothesis of IAA transport was based, were performed using auxin-autonomous suspensions of crown gall cells of Boston Ivy (Rubery and Sheldrake 1974), where the time of addition as well as concentrations of auxin transport inhibitors or auxin itself could be readily controlled. The first characterization of auxin transport mechanisms at the cellular level was reported by Delbarre et al. (1994, 1996) in mesophyll protoplasts and cell suspension from tobacco cv. Xanthi XHFD8, respectively. As depicted on Fig. 2A, Delbarre et al. proposed a simple method to describe the role of passive diffusion and active carrier-mediated processes in the transport of three of the most common auxins, namely native IAA, synthetic naphthalene1-acetic acid (NAA) and 2,4-dichlorophenoxyacetic acid (2,4-D). The same cell suspension was used for the characterization of a new class of inhibitors of auxin influx (Imhoff et al. 2000) as well as for the study of phosphorylation/dephosphorylation of proteins of the auxin efflux complex (Delbarre et al. 1998). One such regulatory protein, PINOID kinase, was recently shown to regulate polar targeting of PIN proteins in Arabidopsis (Friml et al. 2004). In contrast to cell suspensions of Arabidopsis (which tend to form large cell clusters), the major advantage of intensively dividing cell cultures of tobacco

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is that the effects on cell morphology of various inhibitors as well as of auxin itself can be observed in parallel with the measurements of auxin accumulation. Since tobacco cell lines of a good quality are completely friable, it is possible to express all measured data as an equivalent of cell number. During the period of cell division, such high-quality tobacco cell lines usually form polar cell files instead of cell clusters. Based on the studies using tobacco cell line VBI-0 (Nicotiana tabacum L., cv. Virginia Bright Italia; Opatrný and Opatrná 1976), the inhibition of auxin transport by NPA and the consequent rise in the internal auxin level delayed the onset of cell division and disrupted its polarity (Petrášek et al. 2002). Mathematical modelling suggested that NPA possibly disturbs the gradient in auxin concentration along the cell file (Campanoni and Nick 2003). This effect might be mediated by the actin cytoskeleton, as shown by Holweg et al. (2003), using inhibitors of myosin action. In spite of the fact that the auxin-autonomous cell line VBI-I1 containing spherical cells was derived (Campanoni et al. 2001) from ‘parental’ VBI-0 line, VBI-0 itself is routinely maintained on both NAA and 2,4-D. These two auxins may control cell division and cell elongation by different pathways (Campanoni and Nick 2005); therefore, the BY-2 cell line is a better model in this respect, as it is only 2,4-D-dependent. 4.2 Transport of Auxins in BY-2 Cells The potential of simultaneous measurements of intracellular auxin accumulation and tracking of changes in various cell structures, together with directed transgenosis, makes tobacco BY-2 cells an invaluable tool for the study of auxin transport at the cellular level (Petrášek et al. 2003; Zažímalová et al. 2003) as well as for the characterization in vivo of functions of the proteins involved. BY-2 cells respond to the addition of inhibitors NPA or BFA by an immediate rise in the accumulation of IAA, NAA and 2,4-D (Fig. 2B). The transport of these auxins occurs in similar ways as suggested for tobacco Xanthi XHFD8 cells (Delbarre et al. 1996; Fig. 2A). Both the kinetics of NAA accumulation (Fig. 3A) and the arrangement of the cytoskeleton reflect differential sensitivity of BY-2 cells towards BFA and NPA (see Petrášek et al. 2003 for details). To test the proposed role of PIN proteins in the regulation of auxin transport, BY-2 cells were transformed with PIN1, PIN4, PIN6 and PIN7 genes from Arabidopsis thaliana. For all tested PINs, their strong inducible overexpression resulted in a decrease in auxin accumulation (Fig. 3B; Sk˚upa et al. 2005), followed by remarkable changes in cell morphology (Fig. 3D, E, F). Similar changes have been reported previously to be typical of the response to auxin depletion (Sakai et al. 2004). Moreover, the fact that NPA, an inhibitor of auxin efflux, was capable of preventing all observed “auxin-depletion”-induced changes (Fig. 3G) strongly points to the modification in auxin transport (Petrášek et al. 2005). In vivo studies using BY-2 cells expressing Arabidopsis PIN1 protein fused to GFP revealed its predominant localization at transversal PMs, although

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Fig. 3. Treatments with auxin efflux inhibitors or overexpression of putative auxin efflux carriers result in changed levels of endogenous auxin and distinct cell morphology in BY-2 cells. A Accumulation kinetics reflecting the increase in accumulation of auxin (NAA) upon application of inhibitors of auxin efflux, BFA and NPA (Petrášek et al. 2003). Error bars represent SEs of the mean (n = 4). B Decrease in accumulation of auxin upon PIN protein overexpression reflecting enhanced auxin efflux (Sk˚upa et al. 2005). Error bars represent SEs of the mean (n = 4). C Two-day-old BY-2 cells stably expressing Arabidopsis PIN1 protein in translation fusion with GFP. Merged five optical sections (each section 1 µm) through the cortical cytoplasm. The localization of PINs in BY-2 cells at transversal PMs suggests preferential direction of auxin flow. D–G Three-day-long overexpression of PIN proteins under strong promoter results in the cessation of cell division (E) and formation of starch-containing amyloplasts (Lugol staining) (F). All changes are rescued by simultaneous treatment with 10 µM NPA (G) to control-like situation (D), suggesting the inhibition of overexpressed PIN proteins (Petrášek et al. 2005). Scale bars 20 µm

a proportion of fusion protein was localized along longitudinal PMs and in the cortical cytoplasm (Fig. 3C). Thus, in the BY-2 cell line the distribution of PIN1-GFP in cell files resembles the PIN distribution pattern in the cells of the root elongation zone of Arabidopsis thaliana. It was recently shown (Paciorek et al. 2005), using Arabidopsis thaliana plants as well as both BY-2 and VBI-0 tobacco cells, that auxins can inhibit the endocytotic step of the constitutive cycling of PM proteins including PIP2 aquaporin, PM-ATPase and PINs. According to these results, auxin increases levels of PINs at the PM and concomitantly promotes its own efflux from cells. This finding implies a novel mode of auxin action, similar to some animal hormones and consistent with pleiotropic physiological effects of auxin on plant cells.

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Fig. 4. Schematic diagrams summarizing the effects of overexpression of the PIN proteins in BY-2 cells. A Control, exponentially growing cells expressing their endogenous tobacco efflux carriers. According to BLAST search in the EST database of Matsuoka et al. (2004), EST n. 3440f1 is the most promising candidate for an NtPIN gene. B Strong overexpression of Arabidopsis PIN protein stimulates auxin efflux. As a consequence, cells display typical auxin-starvation symptoms (Winicur et al. 1998; Sakai et al. 2004), such as inhibition of cell division, stimulation of amyloplast differentiation and marked cell elongation. C Upon inhibition of a proportion of endogenous as well as overexpressed auxin efflux carriers by NPA, cell division is restored to the control-like situation (Petrášek et al. 2005)

Altogether, our observations, summarized in Fig. 4, reveal that changes in BY-2 cell development are related to auxin transport and – from the point of view of molecular mechanisms of auxin transport – they strongly support the idea that PIN proteins are the rate-limiting components of the auxin efflux machinery, and probably the auxin efflux catalysts themselves.

5 Concluding Remarks and Future Prospects In conclusion, the BY-2 tobacco cell line together with a few other well-defined tobacco cell lines, can serve as an invaluable alternative experimental tool for auxin transport studies complementary to Arabidopsis plants. In our laboratory, the collection of BY-2 cells, transformed with components of both auxin influx (Laòˇnková et al. 2005) and efflux machinery (see above), is maintained and constantly extended. In future, these experimental materials may be useful for studies of the composition of the auxin transport machinery and quantitative aspects of its action, as well as for the characterization of the dynamics of proteins that play a role in the transport of auxin in vivo using fluorescent tags. It may also contribute to discovering the role of both auxin influx and efflux in the establishment of auxin levels inside the cell and their relation to the regulation of cell division, cell elongation and establishment and/or maintenance of cell polarity. Acknowledgements. This work was supported by the Grant Agency of the Academy of Sciences of the Czech Republic, project no. A6038303. PIN gene constructs were expressed in BY-2 cells in the framework of very stimulating, informal collaboration with Dr. Jiˇrí Friml (University of Tübingen, Germany). We also express our gratitude to Prof. Zdenˇek Opatrný (Charles University,

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Prague, Czech Republic) for a long-lasting fruitful collaboration on cell cultures and to Dr. David Morris (University of Southampton, UK) for collaboration and inspiring discussions on auxin transport.

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Paponov IA, Teale WD, Trebar M, Blilou I, Palme K (2005) The PIN auxin efflux facilitators: evolutionary and functional perspectives. Trends Plant Sci 10:170–177 Petrášek J, Elˇckner M, Morris DA, Zažímalová E (2002) Auxin efflux carrier activity and auxin accumulation regulate cell division and polarity in tobacco cells. Planta 216:302–308 ˇ Petrášek J, Cerná A, Schwarzerová K, Elˇckner M, Morris DA, Zažímalová E (2003) Do phytotropins inhibit auxin efflux by impairing vesicle traffic? Plant Physiol 131:254–263 ˇ Petrášek J, Seifertová D, Perry L, Sk˚upa P, Covanová M, Zažímalová E (2005) Expression of AtPINs in tobacco cells increases auxin efflux and induces phenotype changes typical for auxin depletion. Biol Plant 49 (Suppl):S10 Raven JA (1975) Transport of indoleacetic acid in plant cells in relation to pH and electrical potential gradients, and its significance for polar IAA transport. New Phytol 74:163–174 Reinhardt D, Pesce ER, Stieger P, Mandel T, Baltensperger K, Bennett M, Traas J, Friml J, Kuhlemeier C (2003) Regulation of phyllotaxis by polar auxin transport. Nature 426:255–260 Robinson JS, Albert AC, Morris DA (1999) Differential effects of brefeldin A and cycloheximide on the activity of auxin efflux carriers in Cucurbita pepo L. J Plant Physiol 155:678–684 Rubery PH (1990) Phytotropins: receptors and endogenous ligands. Symp Soc Exp Biol 44:119– 146 Rubery PH, Sheldrake AR (1974) Carrier-mediated auxin transport. Planta 118:101–121 Sakai A, Miyazawa Y, Kuroiwa T (2004) Studies on dynamic changes of organelles using tobacco BY-2 as the model plant cell line. In: Nagata T, Hasezawa S, Inzé D (eds) Biotechnology in agriculture and forestry, vol. 53. Tobacco BY-2 cells. Springer, Berlin Heidelberg New York, pp 192–216 ˇ Sk˚upa P, Cerná A, Petrášek J, Perry L, Seifertová D, Zažímalová E (2005) Characterisation of tobacco cell lines transformed with the AtPIN1, 4, 6, 7 genes from Arabidopsis. Biol Plant 49 (Suppl):S12 Sussman MR, Gardner G (1980) Solubilization of the receptor for N-1-naphthylphthalamic acid. Plant Physiol 66:1074–1078 Teale WD, Paponov IA, Ditengou F, Palme K (2005) Auxin and the developing root of Arabidopsis thaliana. Physiol Plant 123:130–138 Ulmasov T, Murfett J, Hagen G, Guilfoyle TJ (1997) Aux/IAA proteins repress expression of reporter genes containing natural and highly active synthetic auxin response elements. Plant Cell 9:1963–1971 Winicur ZM, Zhang GF, Staehelin LA (1998) Auxin deprivation induces synchronous Golgi differentiation in suspension-cultured tobacco BY-2 cells. Plant Physiol 117:501–513 Woodward AW, Bartel B (2005) Auxin: regulation, action, and interaction. Ann Bot 95:707–735 Xu J, Scheres, B (2005) Dissection of Arabidopsis ADP-RIBOSYLATION FACTOR 1 function in epidermal cell polarity. Plant Cell 17:525–536 Zažímalová E, Petrášek J, Morris DA (2003) The dynamics of auxin transport in tobacco cells. Bulgarian J Plant Physiol Spec Issue:207–224

II.3 Tobacco BY-2 Cells as a Model for Differentiation in Heterotrophic Plant Cells Y. Miyazawa1 and A. Sakai2

1 Introduction Starch is the principal form of carbon storage in many plants, and is synthesized in both photosynthetic and non-photosynthetic tissues. Starch found in chloroplasts is termed transitory starch; it is synthesized during the day from photosynthetically fixed carbon and mobilized at night, primarily in the form of sucrose or one of its derivates. It is also synthesized from sugars such as sucrose, in specialized synthesis/storage plastids termed amyloplasts (Kirk 1978). Due to the nutritional and economic importance of starch, many studies have been undertaken to investigate the enzymes involved in its synthesis, as well as the structure of the starch itself (reviewed in Ball and Morel 2003; Geigenberger et al. 2004; Jobling 2004). In addition, the patterns of mRNA expression involved in carbohydrate metabolism including starch synthesis during storage organ development have been analyzed in several plant species (Visser et al. 1994; Ainsworth et al. 1995; Burton et al. 1995; Bachem et al. 1996; Weber et al. 1997). These studies have shed light on the enzymatic properties and expression profiles of individual genes during storage organ development. Typically, storage organ development occurs as a series of specific temporal and spatial steps, involving cell division and subsequent differentiation. These steps often occur simultaneously, and thus are difficult to separate and characterize individually. Moreover, the complex structure of storage organs makes analysis of the cellular mechanisms of storage difficult. Even the rootcap cells, where amyloplast-containing columella cells can be distinguished from their original meristem, are not suitable for analyses on a molecular basis because these cells are too fragile, small and complex. In starch-accumulating tissues, carbohydrate metabolism begins with the cleavage of mobilized sucrose to produce hexoses for use in starch formation, cell wall construction, and energy release. However, pools of metabolic intermediates are shared by several pathways, with the direction of metabolite flow being determined by the needs of the cell. These metabolite fluxes, as well as the regulation of transcription and activity of carbohydrate metabolizing enzymes, vary according to both the 1 Graduate

School of Life Sciences, Tohoku University, Miyagi 980-8577, Japan, e-mail: [email protected] 2 Department of Biological Sciences, Faculty of Science, Nara Women’s University, Nara 630-8506, Japan Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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environmental and developmental state of the cell (reviewed in Geigenberger et al. 2004). Therefore, in order to investigate the mechanisms involved in the formation of starch storage cells, it is necessary to analyze them within a simplified system, in which the two developmental phases, cell proliferation and starch storage, can be detached. The Bright Yellow-2 (BY-2) cell line of tobacco (Nicotiana tabacum) is the model system of choice for these investigations. Exhibiting rapid growth rates and homogeneity, this cell line is widely used as a model system in studies of cell biology (for example, in studies of cell division, auxin action and the cytoskeleton), and has been referred to as “HeLa” cell in higher plants (Nagata et al. 1992; Geelen and Inzé 2001). There are several advantages in using the BY-2 tobacco cell line for such studies. First, the culture system is scalable, obviating the need for time-consuming preparation of large quantities of homogenous cells from whole plant material. Second, BY-2 cells in liquid culture medium form small cell clusters, each composed of several cylindrical cells connected in tandem, with each cell directly exposed to the culture medium. As a result, the effects of growth conditions (for example, nutrient, hormone or drug levels) can be assessed at the level of the individual cell, rather than the cell cluster. For instance, changes in cytoplasmic organelles such as plastids can easily be observed within individual BY-2 cells under the microscope, and recent research has shown that these organelles undergo morphological and functional changes, depending on culture conditions and developmental stage (Sakai et al. 2004). In this chapter, we describe how BY-2 cells are used to analyze both differentiation and dedifferentiation of starch-storing cells, via investigations into the dynamics of plastid morphology, cell shape and gene expressions, as well as the underlying hormonal control mechanisms.

2 Hormonal Factors Affecting Starch-Storing Cell Differentiation in BY-2 Cells 2.1 Auxin as the Primary Determinant of the Developmental State of BY-2 Cells Plant hormones are signal molecules that play a crucial role in growth and development. Since the discovery that auxin and cytokinin are required for the induction of cell division and growth (Miller et al. 1955), these hormones have been found to affect a number of developmental processes by either synergistic or antagonistic action. In tobacco BY-2 cells, exogenously supplied auxin (0.2 mg/L 2,4-dichlorophenoxyacetic acid; 2,4-D) alone is sufficient to maintain meristematic properties, for BY-2 cells are able to synthesize sufficient amount of cytokinin themselves. Although BY-2 cells have been shown to synthesize and maintain trace levels of auxin throughout their cell cycle, it is insufficient for the maintenance of meristematic potential. These properties make

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Fig. 1. Effect of hormonal conditions on BY-2 cells. Changes in cell number (left) and starch content per cell (right) during culture in D-medium (triangles), F-medium (circles) and Bmedium (squares). The values are the means of three independent experiments. Vertical bars represent the standard deviation

BY-2 cells ideal for the study of auxin activity and related gene expressions in plant cells (reviewed in Renaudin 2004). In a study of amyloplast development, Sakai et al. (1992) reported that BY-2 cells, cultured in a medium containing cytokinin (1 mg/L benzyladenine; BA) rather than 2,4-D (referred to as B-medium hereafter), began to accumulate starch within 3 days. Conversely, stationary-phase BY-2 cells cultured in conventional 2,4-D-supplied medium (referred to as D-medium hereafter) began to proliferate actively within 24 h, while their starch content remained low (Fig. 1). This effect is not limited to cells: under the same conditions, undifferentiated plastids (proplastids) are also induced to proliferate (Yasuda et al. 1988). In contrast, BY-2 cells grown in B-medium begin to accumulate starch within 24 h of transfer, and the rate of cell proliferation is reduced (Fig. 1). Furthermore, it has been demonstrated that 2,4-D depletion alone is sufficient to cause the observed differentiation (Sakai et al. 1996). Although the time course for starch accumulation is similar in B-medium and hormone-free medium (referred to as F-medium hereafter), starch accumulation is accelerated and cell proliferation slightly repressed in the former relative to the latter. Microscopic observations demonstrate that this differentiation step involves synchronous and drastic changes in cellular and plastid morphology; increase in cell volume and enlargement of plastids and starch granules (Sakai et al. 1996; Miyazawa et al. 1999, 2001). When 2,4-D is omitted from the culture medium, several changes occur: starch accumulation, cessation of cell proliferation at the G1 phase of cell cycle and cell elongation (Miyazawa et al. 1999). These changes are inconsistent with those observed during storage organ development in vivo. Since starch-storing cell differentiation in BY-2 cells is always accompanied by a reduction in cell proliferation and thus increased sucrose availability, one might suppose that this increase in sucrose availability, not 2,4-D depletion

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itself, may be the triggering factor for starch accumulation. In fact, several lines of evidence suggest that sucrose could induce the differentiation of storage cells (reviewed in Weber et al. 1997). However, in the presence of auxin, BY-2 cells cannot accumulate starch even if their cell proliferation is stopped with a cell cycle blocker aphidicolin (Sakai et al. 1996). Thus the accumulation of starch is not due to the increase in available sucrose resulting from the cessation of cell proliferation, but appears to be controlled by auxin. Further investigations were performed by adding auxin to BY-2 cells cultured in F-medium (Miyazawa et al. 1999, 2002b). The application of 2,4-D to cells grown in an F-medium always inhibited amyloplast development, regardless of when it was applied. Further, the addition of 2,4-D also allowed the amyloplast differentiating cells to regain their original meristematic properties, along with a marked decrease in starch content. Interestingly, when 2,4-D and aphidicolin were added together to starch-storing cells, starch content showed neither marked increase nor decrease (Miyazawa et al. 2002b), suggesting that the dedifferentiation of starch-storing cells comprises two separate processes: cessation of starch synthesis (triggered by 2,4-D addition) and starch degradation (linked with cell division). This supports the hypothesis that starch accumulation is directly controlled by auxin. The most likely reason for the simultaneous occurrence of starch degradation and cell division is that the cells break down starch to utilize it as a carbon source for cell division and/or alter the main routes of carbohydrate flux. For example, in potato, the predominant routes of carbohydrate mobilization change during tuber development, during which there is conversion of dividing cells to starch-storing cells (Fernie and Willmitzer 2001). A similar system may also operate in BY-2 cells. In fact, expression patterns of the genes responsible for starch synthesis and carbohydrate metabolism fluctuate between cell proliferation and starch storage, suggesting that carbohydrate flux might be under hormonal control (see Sect. 3.1 below). 2.2 The Role of Cytokinin in the Development of Starch-Storing Cells BY-2 cells have also played a pivotal role in the study of cytokinin kinetics during the plant cell cycle (reviewed in Roef and Van Onckelen 2004). Using BY-2 protoplasts, Hasezawa and Sy¯ono (1983) found that both cell division and cell elongation were controlled by changes in the relative levels of auxin and cytokinin, i. e. a high ratio of cytokinin to auxin is an important factor for BY-2 protoplasts to elongate. They also found that elongating cells contained many particles reported as starch-containing plastids. Sakai et al. (1992) found that BY-2 cells develop amyloplasts when cytokinin is added instead of auxin in the culture medium. Later studies revealed that in the presence of 2,4-D, cytokinin (BA) application caused neither amyloplast formation nor cessation of cell division (Sakai et al. 1996). Further, the addition of BA to hormone-free cultures always resulted in an increased starch accumulation and a decreased

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cell proliferation, irrespective of the timing of the hormone addition (Miyazawa et al. 1999). These observations suggest that depletion of 2,4-D plays a central role in the differentiation of starch-storing cells, while exogenously added cytokinin plays only a supplemental role. While the above interpretation may not necessarily be incorrect, recent studies have revealed that endogenous cytokinin does play an important role in cell differentiation. BY-2 cells can produce sufficient levels of cytokinin to maintain their meristematic state, with production peaking in three phases of the cell cycle: G1 , S and G2 /M (Redig et al. 1996). To date, no specific cytokinin biosynthesis inhibitor has been identified. However, lovastatin, a potent inhibitor of 3-hydroxy-3-methylglutaryl coenzyme A reductase, may be practically used for this purpose in BY-2 cells (Crowell and Salaz 1992). Indeed, lovastatin treatment reduced endogenous cytokinin content during cell cycle progression in BY-2 cells (Laureys et al. 1998, 1999). When lovastatin was added to cells cultured under 2,4-D-depleted conditions, the treatment inhibited starch accumulation, which was reversed by addition of either mevalonate or cytokinin (Miyazawa et al. 2002a). Analysis of cells grown in F-medium revealed a significant accumulation of trans-zeatin riboside, which was inhibited by lovastatin treatment. Since amyloplast formation occurs when 2,4-D-depleted cells cease cell proliferation at the G1 phase, the accumulation of cytokinin might correspond to the peak observed at G1 in 2,4-D-supplied synchronized cultures. In plant cells, mevalonate-derived metabolites have been shown to play critical roles in many cellular processes, such as phytosterol synthesis and protein prenylation (reviewed in Hemmerlin et al. 2004 and references therein), and cytokinins are synthesized not only via the mevalonate pathway but also via the methylerythritol phosphate pathway (Kasahara et al. 2004). Therefore, inhibition of amyloplast formation by lovastatin treatment cannot be explained entirely in terms of inhibition of cytokinin synthesis. However, the finding that 2,4-D deprivation induced a significant increase in cytokinin content, which correlates with starch accumulation, strongly suggests that the enhanced accumulation of endogenous cytokinin under auxindepleted condition is necessary for starch-storing cell differentiation in BY-2 cells. 2.3 Other Hormones that Affect Starch Accumulation in BY-2 Cells Other hormones have been reported to affect starch-storing cell differentiation. For example, gibberellins are shown to inhibit potato tuberization, and abscisic acid enhances starch accumulation in cultured rice cells (Jackson 1999; Akihiro et al. 2005). In BY-2 cells, addition of these hormones has no significant effect on starch accumulation. Similarly, addition of 1-aminocyclopropane-1-carboxylic acid, a precursor of ethylene, has a negligible effect on starch-storing cell differentiation (Miyazawa et al., unpublished results). As these results were observed in different species and different cultured cell lines, future efforts

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should focus on elucidating the common underlying mechanism of starchstoring cell differentiation. To the best of our knowledge, the only other phytohormone that affects starch accumulation in BY-2 cells is brassinolide. The addition of this hormone to cells cultured in F-medium results in a dose-dependent increase in cell number, reduction in cell length and decrease in starch content (Miyazawa et al. 2003), a mode of action similar to that of auxin. Indeed, a number of the physiological effects of brassinosteroids are analogous to those of auxin (reviewed in Clouse and Sasse 1998), although substitution of auxin with brassinolide in culture medium does not fully compensate for its absence. Moreover, conflicting results have been reported regarding the effects of brassinosteroids in their relationship with auxin and cell proliferation (reviewed in Clouse and Sasse 1998). Thus, the role of brassinosteroids in the control of cell proliferation and differentiation, as well as the possibility of crosstalk with auxin, remains to be determined.

3 Regulations of Gene Expressions Required for Differentiation and Dedifferentiation of Starch-Storing Cells 3.1 Hormonal Regulation of Starch Synthesis Gene Expression During Starch-Storing Cell Differentiation In heterotrophic cells, starch biosynthesis begins with mobilized sugar, primarily sucrose. The sugar is cleaved and enzymatically converted to ADP-glucose inside the plastid. Although the major pathways of starch synthesis and degradation have been described, how individual reactions are involved in starchstoring cell differentiation and dedifferentiation is less well understood. As mentioned previously, the BY-2 cell line provides a suitable model system for studying these processes. In BY-2 cells, starch-storing cell differentiation was suppressed by the addition of eukaryotic transcription/translation inhibitors, regardless of the timing of the addition (Sakai et al. 1997). This suggests that differentiation requires de novo expression of nuclear genes. Furthermore, pulse treatment of the cells with the nucleo-cytoplasmic gene expression inhibitors demonstrated that nuclear gene expression necessary for amyloplast development begins 6−12 h after transfer of the cells to 2,4-D-depleted medium (Sakai et al. 1999). In fact, expression of the genes encoding AgpS (a small subunit of ADP-glucose pyrophosphorylase, a key enzyme in starch synthesis), GBSS (granule-bound starch synthase) and SBE (starch branching enzyme) is up-regulated within 12 h after transfer to amyloplast-inducing medium and their gene expression is active throughout starch-storing cell differentiation (Miyazawa et al. 1999). The addition of 2,4-D or BA results in a decrease or increase in mRNA levels for the AgpS, respectively, regardless of the timing of addition. These

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changes correlate well with the changes in starch content following application of the hormones (Miyazawa et al. 1999). Additionally, reduction of endogenous cytokinins by lovastatin treatment results in a reduction in the AgpS transcript levels, which is reverted by the simultaneous addition of either mevalonate or cytokinins (Miyazawa et al. 2002a). Moreover, addition of cycloheximide to amyloplast differentiating cells promptly decreases mRNA levels for AgpS and GBSS. This indicates that continuous de novo synthesis of nuclear-encoded proteins is necessary to maintain expression of these genes (Miyazawa et al. 2001). In an effort to identify the de novo synthesized protein responsible for the expression of AgpS and GBSS, a differential display was performed (Miyazawa et al. 2002c). This resulted in the cloning of three starch-storing cell-specific (SCI) genes that are currently under investigation. In contrast to AgpS and GBSS, the expression of the gene encoding SBE was not affected by cycloheximide treatment, indicating that its gene expression is regulated in a different manner from that of the other two genes (Miyazawa et al. 2001). In addition, mRNA levels for the SBE-encoding gene decreased in cells cultured in the presence of 2,4-D (Miyazawa et al. 1999), indicating that the expression of at least one of the SBE genes is negatively regulated when BY-2 cells are not accumulating starch. These results suggest that expression of the starch synthesis genes is regulated antagonistically by auxin and cytokinin, either directly or indirectly, thereby controlling the rate of starch synthesis during amyloplast formation. Comparative analysis of the expression of carbohydrate-metabolizing genes between BY-2 meristematic and starch-storing cells has revealed that, to a certain extent, the carbohydrate-metabolizing gene expression is coordinately regulated by the developmental state of the cell. As shown in Fig. 2, mRNA levels for sucrose synthase, fructokinase and glucose-6-phosphate translocator are up-regulated by culturing under 2,4-D-depleted conditions. In contrast, transcript levels for invertase, UDP-glucose pyrophosphorylase, glucokinase and both cytosolic and plastidic phosphoglucomutase are largely unaffected. Moreover, mRNA levels for sucrose synthase, fructokinase and glucose-6-phosphate translocator are promptly down-regulated by 2,4-D addition (Fig. 3), indicating that at least some isoforms of these genes are coordinately expressed during the starch storage phase, probably producing the distinct carbohydrate metabolic pathway that occurs during the cell proliferation phase. Despite increasing knowledge of post-translational regulation of starch-synthesizing enzymes (reviewed in Tetlow et al. 2004), little is known about transcriptional regulation of these genes. Thus the findings that not only starch synthesis genes but also some of carbohydrate-metabolizing genes are transcriptionally regulated in a coordinate manner will open new avenues of research into the regulation of starch-storing cell differentiation.

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Y. Miyazawa and A. Sakai Fig. 2. RNA-gel blot comparisons of transcript levels between the starch-storing and cell-proliferation phases. Total RNA extracted from BY-2 cells cultured for 0, 24 and 48 h in F-medium (−2,4D) and D-medium (+2,4-D). Hybridization signals for sucrose synthase (SUSY), cell wall-bound invertase (cwInv), neutral invertase (nInv), alkaline/neutral invertase (anInv), UDP-glucose pyrophosphorylase (UGPase), cytosolic phosphoglucomutase (CPGM), hexokinase, fructokinase, glucose-6-phosphate translocator (GPT) and plastidic phosphoglucomutase (PPGM) are shown

Fig. 3. Response of transcript levels for sucrose synthase (SUSY), fructokinase and glucose-6phosphate translocator (GPT) to 2,4-D addition. 2,4-D was added to the culture 24 h after transfer to F-medium (left) and the transcript levels for SUSY, fructokinase and GPT were compared with cells cultured in F-medium (right)

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3.2 Organelle-Mediated Control of Starch-Storing Cell Differentiation Starch deposition in BY-2 cells is also inhibited by the addition of chloramphenicol (a prokaryotic translation inhibitor), regardless of the timing of the addition. However, in contrast to the effect of eukaryotic transcription/translation inhibitors, the inhibitory effect of chloramphenicol decreased as the timing of addition shifted to the late phase (Sakai et al. 1997; Miyazawa et al. 2000). Microscopic examination revealed the overall morphology of chloramphenicol-treated plastids to be similar to that of non-treated cells, except that starch granule enlargement was inhibited in the former (Miyazawa et al. 2000). Examination of AgpS mRNA accumulation revealed that inhibition of starch accumulation by chloramphenicol treatment did not affect mRNA accumulation. Therefore, the chloramphenicol-mediated reduction in starch accumulation is not due to a reduction in AgpS expression, but rather to unknown de novo-synthesized organellar proteins. This finding is in sharp contrast to the results of cycloheximide treatment, which showed that continuous nuclear gene expression was required for the expression of AgpS (Miyazawa et al. 2001). There are two possible means by which chloramphenicol could inhibit starch accumulation without affecting starch synthesis gene expressions. First, chloramphenicol might inhibit mitochondrial protein synthesis. If this were the case, the cell would only be able to generate energy from glycolysis (rather than via respiration), and once the carbohydrate source for synthesis was exhausted, starch accumulation would cease. Under this scenario, the inhibitory effect of chloramphenicol added at the later stages of amyloplast formation might be explained as follows: the cell accumulates precursors for starch synthesis (such as ADP-Glc) until the addition of inhibitor, and after its addition, starch continues to be accumulated until this pool is exhausted. Alternatively, chloramphenicol might inhibit protein synthesis from genes contained in plastid or mitochondrial genomes, which are required during the early stages of amyloplast formation. Such a protein might be encoded in one of the many open reading frames (ORFs) in the plastid or mitochondrial genomes that have not been as-yet defined a function. Assays of metabolites resulting from glycolysis and respiration, as well as investigation into specific ORFs in the plastid and mitochondrial genomes, will be required to elucidate the exact role of organellar protein synthesis in starch-storing cell differentiation.

4 Conclusions and Perspectives The BY-2 cell line is an ideal model for studying the nature of meristematic heterotrophic tissues. Because the complexity of highly organized storage tissues precludes analysis of the mechanisms controlling starch storage, we developed an in vitro starch-storing cell-inducing system using cultured tobacco BY-2 cells. In this chapter, we described the procedure for the phytohormone-

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mediated induction of starch-storing cell differentiation. To the best of our knowledge, this is the simplest and fastest system for inducing starch-storing cell development. Using this procedure, it was demonstrated that BY-2 cells differentiate into different cell types, a finding that opened up new approaches

Fig. 4. Proposed model of starch-storing cell differentiation and dedifferentiation in BY-2 cells. When BY-2 cells are actively proliferating under 2,4-D-supplied conditions, expression of at least some isoforms of sucrose synthase (SUSY), fructokinase, glucose-6-phosphate translocator (GPT) and starch synthesis genes is repressed. During starch-storing cell development, the expression of these genes is coordinately up-regulated and contributes to starch accumulation. Note that under this condition, expression of AgpS and GBSS are under the control of de novo synthesized cytosolic protein(s) (starch-storing cell induced proteins: SCIs). n Cell nucleus; mt mitochondrion; pp proplastid; ap amyloplast

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for investigating the developmental processes. As summarized in Fig. 4, this differentiation step is triggered primarily by both a depletion of auxin and increase of endogenous cytokinin. Coordinated changes in the levels of these hormones affect the levels of transcription of carbohydrate-metabolizing and starch synthesis genes, and expression of the genes encoding AgpS and GBSS are regulated by as-yet undefined nuclear-encoded proteins. Differential display analysis has been undertaken in an attempt to identify these factors and the cDNA candidates are currently being investigated. The finding that auxin addition causes starch-storing cells to dedifferentiate into meristematic cells will further extend the usefulness of the BY-2 culture system, since the mechanisms underlying dedifferentiation of starch-storing cells remain undefined. In order to understand the mechanisms controlling starch storage, identification of key molecules, such as the above-mentioned undefined protein(s), will be necessary. The homogenous and synchronous nature of the BY-2 starch-storing cell induction system lends itself easily to analysis using recently developed molecular biological techniques, such as cDNA arrays and proteomics (Baginsky et al. 2004; Laukens and Witters 2004; Matsuoka et al. 2004). Inhibition of organellar protein synthesis affects the starch accumulation, and so consideration of organellar function, in addition to starch synthesis gene expression, will be crucial to the understanding of starch storage in future studies. Undoubtedly, this simple culture system provides an ideal model for analyzing the processes of starch-storing cell differentiation and dedifferentiation; however, which state of development in the plant is represented by this system is not defined. There appear to be many similarities between the growth of BY-2 cells and the development of root cap cells; for example, in the development of amyloplasts, increase in cell volume and decrease in cell proliferation. In this context, meristematic BY-2 cells correspond to the initial cells located at the root apical meristem. Moreover, Winicur et al. (1998) reported that auxin deprivation also induces synchronous Golgi differentiation in BY-2 cells within 4 days, a process analogous to that observed during root cap development. Recently, Takahashi et al. (2003) reported that amyloplasts in root cap cells of Arabidopsis and radish seedlings were degraded by a hydrotropic stimulus, but restored once the stimulus was removed. That this reversal could be effected within 12 h suggests that starch degradation and restoration occur within the same columella cell: thus the root cap cells are capable of both differentiation and dedifferentiation. Further comparative analyses between BY-2 cells and plant tissues, together with detailed studies of starch-storing cell differentiation and dedifferentiation in BY-2 cells, will help to shed new light on the nature of heterotrophic plant cells. Acknowledgements. This work was supported by the Program for Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN), Grants-in-Aid for Young Scientists (B) (no. 15770039) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan, and for Scientific Research on Priority Areas (no. 170510003).

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Matsuoka K, Demura T, Galis I, Horiguchi T, Sasaki M, Tashiro T, Fukuda H (2004) A comprehensive gene expression analysis toward the understanding of growth and differentiation of tobacco BY-2 cells. Plant Cell Physiol 45:1280–1289 Miller CO, Skoog F, Saltza MH, Strong FM (1955) Kinetin, a cell division factor from deoxyribonucleic acid. J Am Chem Soc 77:1329–1334 Miyazawa Y, Sakai A, Miyagishima S, Takano H, Kawano S, Kuroiwa T (1999) Auxin and cytokinin have opposite effects on amyloplast development and the expression of starch synthesis genes in cultured Bright Yellow-2 tobacco cells. Plant Physiol 121:461–469 Miyazawa Y, Sakai A, Kawano S, Kuroiwa T (2000) Organellar protein synthesis controls amyloplast formation independent of starch synthesis gene expression. Cytologia 65:435–442 Miyazawa Y, Sakai A, Kawano S, Kuroiwa T (2001) Differential regulation of starch synthesis gene expression during amyloplast development in cultured tobacco BY-2 cells. J Plant Physiol 158:1077–1084 Miyazawa Y, Kato H, Muranaka T, Yoshida S (2002a) Amyloplast formation in cultured tobacco BY-2 cells requires a high cytokinin content. Plant Cell Physiol 43:1534–1541 Miyazawa Y, Kutsuna N, Inada N, Kuroiwa H, Kuroiwa T, Kawano S (2002b) Dedifferentiation of starch-storing cultured tobacco cells: effects of 2,4-dichlorophenoxy acetic acid on multiplication, starch content, organellar DNA content, and starch synthesis gene expression. Plant Cell Rep 21:289–295 Miyazawa Y, Sakai A, Matsunaga S, Asami T, Kawano S, Kuroiwa T, Yoshida S (2002c) Isolation and expression of a novel starch-storing cell-specific gene containing the KH RNA binding domain from tobacco-cultured cells BY-2. J Exp Bot 53:2451–2452 Miyazawa Y, Nakajima N, Abe T, Sakai A, Fujioka S, Kawano S, Kuroiwa T, Yoshida S (2003) Activation of cell proliferation by brassinolide application in tobacco BY-2 cells: effects of brassinolide on cell multiplication, cell-cycle-related gene expression, and organellar DNA contents. J Exp Bot 54:2669–2678 Nagata T, Nemoto Y, Hasezawa S (1992) Tobacco BY-2 cell line as the “HeLa” cell in the cell biology of higher plants. Int Rev Cytol 132:1–30 Redig P, Shaul O, Inzé D, Van Montague M, Van Onckelen H (1996) Levels of endogenous cytokinins, indole-3-acetic acid, and abscisic acid during the cell cycle of synchronized tobacco BY-2 cells. FEBS Lett 391:175–180 Renaudin JP (2004) Growth and physiology of suspension-cultured plant cells: the contribution of tobacco BY-2 cells to the study of auxin action. In: Nagata T, Hasezawa S, Inzé D (eds) Biotechnology in agriculture and forestry, vol. 53. Tobacco BY-2 cells. Springer, Berlin Heidelberg New York Roef L, Van Onckelen H (2004) Hormonal control of the plant cell cycle. In: Nagata T, Hasezawa S, Inzé D (eds) Biotechnology in agriculture and forestry, vol. 53. Tobacco BY-2 cells. Springer, Berlin Heidelberg New York Sakai A, Kawano S, Kuroiwa T (1992) Conversion of proplastids to amyloplasts in tobacco cultured cells is accompanied by changes in the transcriptional activities of plastid genes. Plant Physiol 100:1062–1066 Sakai A, Yashiro K, Kawano S, Kuroiwa T (1996) Amyloplast formation in cultured tobacco cells; effects of plant hormones on multiplication, size, and starch content. Plant Cell Rep 15:601–605 Sakai A, Miyazawa Y, Yashiro K, Suzuki T, Kawano S (1997) Amyloplast formation in cultured tobacco cells. II Effects of transcription/translation inhibitors on accumulation of starch. Cytologia 62:295–301 Sakai A, Miyazawa Y, Saito C, Nagata N, Takano H, Hirano H-Y, Kuroiwa T (1999) Amyloplast formation in cultured cells. III Determination of the timing of gene expression necessary for starch accumulation. Plant Cell Rep 18:589–594 Sakai A, Miyazawa Y, Kuroiwa T (2004) Studies on dynamic changes of organelles using tobacco BY-2 as the model plant cell line. In: Nagata T, Hasezawa S, Inzé D (eds) Biotechnology in agriculture and forestry, Vol. 53. Tobacco BY-2 cells. Springer, Berlin Heidelberg New York, pp192–216

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Takahashi N, Yamazaki Y, Kobayashi A, Higashitani A, Takahashi H (2003) Hydrotropism interacts with gravitropism by degrading amyloplasts in seedling roots of Arabidopsis and radish. Plant Physiol 132:805–810 Tetlow IJ, Morell MK, Emes MJ (2004) Recent advances in understanding the regulation of starch metabolism in higher plants. J Exp Bot 55:2131–2145 Visser RGF, Vreugdenhil D, Hendriks T, Jackobsen E (1994) Gene expression and carbohydrate content during stolon to tuber transition in potatoes (Solanum tuberosum). Physiol Plant 90:285–292 Weber H, Borisjuk L, Wobus U (1997) Sugar import and metabolism during seed development. Trends Plant Sci 2:169–174 Winicur ZM, Zhang GF, Stahelin LA (1998) Auxin deprivation induces synchronous Golgi differentiation in suspension-cultured tobacco BY-2 cells. Plant Physiol 117:501–513 Yasuda T, Kuroiwa T, Nagata T (1988) Preferential synthesis of plastid DNA and increased replication of plastids in cultured tobacco cells following medium renewal. Planta 174:235–241

Section III Intracellular Traffic

III.1 Imaging the Early Secretory Pathway in BY-2 Cells D.G. Robinson1 and C. Ritzenthaler2

1 The Early Secretory Pathway in Plants: A Brief Introduction Although the essential components of the COPI- and COPII-vesiculation machineries are expressed in plants (Andreeva et al. 1998; Jürgens 2004), the early secretory pathway in plants is organized in a much different way to that of animal and yeast cells (Nebenführ et al. 2002; Pavelka and Robinson 2003). This is undoubtedly a consequence of the different physiology of the plant cell, whereby the plant Golgi apparatus has been likened to a polysaccharide factory (Nebenführ and Staehelin 2001). Despite some controversy over several aspects of ER-to-Golgi transport (see below), it can be said that there is widespread agreement over the following key features:

• The plant Golgi apparatus is polydisperse, consisting of discrete Golgi stacks distributed throughout the cytoplasm (Staehelin and Moore 1995). • The plant Golgi apparatus itself is mobile, and this is microfilament-dependent (Staehelin and Moore 1995). • In higher plant cells, transitional ER (i. e. specific domains of ER export) does not seem to exist (Pavelka and Robinson 2003). • Transport between the ER and the Golgi apparatus does not involve a mobile, microtubule-dependent intermediate (ERGIC) compartment (Neumann et al. 2003). • COPI vesicles only form at the periphery of Golgi cisternae (Pimpl and Denecke 2000). • The plant Golgi apparatus does not fragment during mitosis (Nebenführ et al. 2000). • COPII proteins are present in plants and are essential for ER export (Phillipson et al. 2001). COPI and COPII vesicles were discovered on the basis of vesicle budding assays performed in vitro with subcellular fractions enriched in Golgi or ER membranes respectively (Schekman and Orci 1996; Balch 2004). Although the essentiality of the COPI and COPII coat protein recruiting machineries for successful protein transport through the early secretory pathway is generally accepted (see Bonifacino and Glick (2004) and Lee et al. (2004) for reviews), 1 Heidelberg

Institute of Plant Sciences-Cell Biology, University of Heidelberg, Im Neuenheimer Feld 230, 69120 Heidelberg, Germany, e-mail: [email protected] 2 IBMP CNRS, Institut de Biologie Moléculaire des Plantes, Strasbourg, France Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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the actual existence of these vesicles in vivo is still questioned by some researchers in the animal (e. g. Polishchuk et al. 2003) and plant (e. g. Hawes and Satiat-Jeunemaitre 2005) fields. Moreover, although implicit in the “secretory unit model” for ER-Golgi transport (Neumann et al. 2003), one is troubled by the low frequency in which direct tubular contacts between the ER and the Golgi apparatus have been reported in plant cells. It is also difficult to understand why only anterograde but not retrograde transport should be mediated by tubules.

2 General Description of the BY-2 Endomembrane System The function of the endomembrane system of tobacco BY-2 cells is essentially that of processing, sorting, and transport of glycoproteins and non-cellulose matrix polysaccharides to the cell wall and the vacuolar compartment(s). Correct targeting and the maintenance of the secretory pathway relies on different proteins, such as chaperones, components of the vesicular transport machinery and the cytoskeleton. The ER is the first compartment of the secretory pathway and also the largest, since it can represent up to 50% of the total cellular membrane surface area (Staehelin 1997; Hara-Nishimura et al. 2004). It is the most versatile organelle, as easily seen in BY-2 cells expressing ERtargeted GFP markers. Thus, the ER is very dense in actively growing cells (3to 4-day-old cultures), and adopts a very specific organization during mitosis and cytokinesis (Nebenführ et al. 2000; C. Ritzenthaler, unpublished results). As cells become older (6- to 7-day-old cultures) and progressively deprived of carbon sources and oxygen, the ER develops a much looser conformation (C. Ritzenthaler, unpublished results; see also Fig. 3A,B) similar to that observed in mature tobacco epidermal cells (e. g. Fig. 2 in Brandizzi et al. 2002). The other compartments, further downstream of the ER, are proportionally less affected by growth or environmental conditions, at least at the light microscopical level. Thus, Golgi fluorescent markers show little modification upon ageing of the cells (C. Ritzenthaler, unpublished results) or upon shifting to extreme temperatures such as 40 or 4 ◦ C (D. Robinson and C. Ritzenthaler, unpublished results). The recently characterized prevacuolar compartment (PVC) exhibits in fluorescence microscopy a mobile punctate pattern similar to those observed for Golgi stacks (Tse et al. 2004). By electron microscopy, the prevacuolar marker protein VSR (vacuolar sorting receptor) localizes to multivesicular bodies (Tse et al. 2004). More recently, evidence was provided suggesting that recycling of the VSR from the prevacuolar compartment to the Golgi apparatus is an essential process that is saturable and wortmannin sensitive (daSilva et al. 2005). The vacuoles and the cell wall constitute the final compartments of the secretory pathway. These compartments have been well characterized and BY-2 cells expressing specific markers of these compartments have been obtained (e. g. Mitsuhashi et al. 2000; Czempinski et al. 2002;

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Kutsuna and Hasezawa 2002; Kutsuna et al. 2003; Hoffmann and Nebenführ 2004; Yamada et al. 2005).

3 The Golgi Apparatus: Structure, Motility and Behaviour During Mitosis Logarithmically growing BY-2 cells have a Golgi apparatus that is typical for undifferentiated plant cells. It is difficult to ascertain the average number of Golgi stacks per interphase cell, but these were estimated to reach several hundred (Nebenführ et al. 1999). There is no preferential location: Golgi stacks are found in the cortical cytoplasm, in the cytoplasm around the nucleus, and

Fig. 1. Golgi visualization in tobacco BY-2 cells. A A typical Golgi stack, showing polarity parameters (lumen width, staining, intercisternal filaments) in a clear cis (c) to trans (t) gradient; chemical fixation. B The same as for A but from a high pressure frozen sample (courtesy of Dr. Andreas Nebenführ). C Single optical section through the cortex of a cell expressing GmManI-GFP. Arrows and arrowheads point to individual Golgi stacks in face and side-views, respectively. At high magnification, face-viewed Golgi stacks appear as disks (inset left), whereas those viewed from the side form a line (inset right). Bar 0.25 µm for A and B and 5 µm for C

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in the transvacuolar strands (Nebenführ et al. 1999). Each stack has between five and eight cisternae, which reveal nicely the classical polarity parameters (see Robinson and Kristen 1982 for details) of lumenal width (decreasing cis to trans), staining intensity (increasing cis to trans), and intercisternal filaments (medial and trans). These features can be seen in both chemically fixed and high pressure frozen fixed cells (Nebenführ et al. 1999; Ritzenthaler et al. 2002; Tse et al. 2004; see also Figs. 1A,B and 3C). It has been reported that Golgi stacks in transgenic cell lines expressing the processing enzyme GmManI (see below) are smaller in both cisternal number and cisternal diameter (Nebenführ et al. 1999). Our knowledge on the distribution and motility of BY-2 Golgi stacks has been dependent upon the availability of cell lines stably expressing fluorescent Golgi-localized markers. To date, only few (X)FP-labelled enzymes have been successfully used: (1) a GFP- or RFP-tagged class I α-1,2-mannosidase from Glycine max (Nebenführ et al. 1999; Ritzenthaler et al. 2002; Yang et al. 2005). This is an N-glycoprotein processing enzyme, which is located in the cis cisternae (Nebenführ et al. 1999); (2)fluorescent Golgi-localized marker a YFPtagged GONST1 from A. thaliana (Tse et al. 2004). This is a sugar nucleotide transporter located in the trans cisternae (Baldwin et al. 2001); (3) a GFP- or RFP-tagged sialyl transferase (Boevink et al. 1998; Saint-Jore et al. 2002). This is a terminal glycosyl transferase from mammalian sources, which correctly targets to the trans cisternae in plants (Boevink et al. 1998; Saint-Jore et al. 2002); (4) N-glycan GFP-tagged xylosyltransferase. This enzyme is preferentially located in medial cisternae (Follet-Gueye et al. 2003). Corresponding to the orientation of the stack being observed, all of these markers give rise to a fluorescent image that is either punctate (top-view) or rod-like (side view; see Fig. 1C). In addition to the expression of (X)FP-constructs as a means of visualizing the Golgi apparatus in living BY-2 cells, a specific labelling of Golgi stacks has

I Fig. 2. Confocal laser scanning micrographs showing the localization of COPI and COPII com-

ponents in GFP-HDEL and ManI:GFP transgenic tobacco BY-2 cells. A–C Optical section through the cortical cytoplasm of stably transformed cells expressing the Golgi marker GmMan1-GFP (green channel) after fixation and immunolocalization with anti-AtArf1 antibodies (red channel). As can be seen in the merged image, the COPI vesicle subunit co-localizes with GFP-fluorescence and is restricted to the margins of the Golgi stacks. Insets represent single immunolabelled Golgi stacks under face (left) and side (right) views at high magnification (five times compared to main image). D–F Optical section through the cortical cytoplasm of stably transformed cells expressing the ER marker GFP-HDEL (green channel) after fixation and immunolocalization with AtSec23 antibodies (red channel). As can be seen in the merged image and with more detail in the insets (magnified five times compared to main images), the COPII vesicle subunit labelling is located either directly on the membrane of the cortical ER or closely adjacent. G–I Optical section through the cortical cytoplasm of a live cell co-expressing the Golgi marker GmMan1-RFP (arbitrarily shown in green) and the COPII marker Sec13:GFP (red channel). As can be seen in the merged image and in more detail in the insets (magnified five times compared to main images), the COPII vesicle subunit marker is dispersed throughout the cortical cytoplasm, forming small punctae that greatly outnumber individual Golgi stacks. Remarkably, COPII labelling is frequently enriched around individual Golgi stacks. Bars 5 µm

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also been obtained by immunostaining with COPI antibodies (Ritzenthaler et al. 2002). Both AtArf1 and AtSec21 (γ −COP) antibodies bind to the rims of the cisternae, giving rise to a doughnut-like image (see Fig. 2A–C). When the immunostaining is performed on the ManI-GFP cell line, the punctate GFP signal is usually found in the middle of the red immunofluorescence signal (Fig. 2A–C). The immunostaining pattern reflects COPI-vesicle formation which takes place at the tubular periphery of the cisternae, a feature confirmed

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by immunogold labelling of cryosections, albeit on plants other than BY-2 (Pimpl et al. 2000). The first demonstration that plant Golgi stacks move along actin microfilaments which run parallel to the tubules of the cortical ER network was made in 1998 (Boevink et al. 1998). However, it would seem from subsequent work that ER-Golgi protein trafficking is not dependent on a functional actin cytoskeleton (Brandizzi et al. 2002; Saint-Jore et al. 2002). This work was performed on epidermal cells of tobacco leaves, so it is very likely to be valid for BY-2 cells as well. That this is indeed so is shown in the paper of Nebenführ et al. (1999). These authors established that BY-2 Golgi stacks are not in continuous motion, but that individual stacks could be seen moving and then stopping, whence they performed a kind of “wiggling” before recommencing movement. Directed movement was measured to occur at speeds of up to 2.4 µm/s in the cell cortex and 3.8 µm/s in transvacuolar strands. These velocities are one order of magnitude higher than those usually recorded in the leaf epidermis system (Boevink et al. 1998), and may reflect the higher metabolic activity of BY-2 cells. Treatment of BY-2 cells with actin inhibitors (cytochalasins, latrunculin B) and with 2,3-butanedione monoxime (BDM), an inhibitor of myosin ATPase, led to a cessation of Golgi stack movement (Nebenführ et al. 1999). Changes in Golgi stack distribution during the life cycle of BY-2 cells have been documented by Nebenführ and coworkers (2000). They showed that immediately prior to prophase, Golgi movement slows down, and about a third of the Golgi stacks previously situated in the cortical cytoplasm move into the perinuclear cytoplasm of the phragmosome. During metaphase around 40% of the Golgi stacks are clustered in the immediate vicinity of the spindle, with about half of these concentrated in an equatorial region, “the Golgi belt”, under the plasma membrane. During cytokinesis these Golgi stacks appear to remain concentrated adjacent to the phragmoplast. Interestingly, actin microfilaments do not seem to be absolutely required for the segregation and maintenance of the spindle/phragmoplast-associated Golgi stacks. Nebenführ et al. (2000) also addressed the question of Golgi inheritance in BY-2 cells. It had previously been reported that a doubling of Golgi stacks occurred during metaphase (GarciaHerdugo et al. 1988), but Nebenführ et al. (2000) were unable to identify any particular time point in the BY-2 life cycle where this might occur, nor was it possible for these authors to distinguish between de novo synthesis and fission as a means of Golgi multiplication.

4 The Endoplasmic Reticulum: Distribution and ER-Export Sites (ERES) The ER of BY-2 cells has a morphology that is typical for many plant cells, with small islands of cisternal membrane enmeshed in a network of tubules (Quader 1990; Staehelin 1997). This is most vividly seen in tangential optical

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sections through the cortex of cells expressing a fluorescent marker (Fig. 3A,B), either GFP-HDEL (the ER retention motif; Ritzenthaler et al. 2002; Yang et al. 2005) or Bip-DsRed (the ER chaperone; Yang et al. 2005). In contrast, the ramifications of the ER and its subdivision into cisternal and tubular regions are not readily appreciated in thin sections (Fig. 3C), nor is a particular relationship to Golgi stacks discernible. Specialized domains with a high density of vesicle budding profiles, characteristic of cells having transitional ER, e. g. Pichia pastoris (Rossanese et al. 1999; Mogelsvang et al. 2003) and Chlamydomonas reinhardii (Zhang and Robinson 1986), are not seen. Individual vesicle budding profiles are extremely rare in thin sections (Ritzenthaler et al. 2002), as noted elsewhere in the literature for other higher plant cells (e. g. Craig and Staehelin 1988; Staehelin 1997).

Fig. 3. ER visualization within tobacco BY-2 cells. Cortical ER of stably transformed cells expressing the ER marker GFP-HDEL as visualized by epifluorescence (A) and confocal microscopy (B). C Electron micrograph of a chemically fixed tobacco BY-2 cell, illustrating the ultrastructure of different organelles found within the cytoplasm. The endoplasmic reticulum (ER) appears as long tubular structures often coated with ribosomes. Golgi stacks (G) are composed of five to eight cisternae of lumenal width decreasing from cis to trans. The mitochondria (M) and plastids (P) are clearly surrounded by a double membrane and electron lucent starch bodies are often detected within amyloplast. The vacuole (V) delimited by the tonoplast appears empty. Bar 5 µm in A and B and 0.5 µm in C

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The question of morphology, density, and distribution of ER export sites (ERES) has only been possible to address after antibodies against plant COPII coat proteins became available (Movafeghi et al. 1999; Yang et al. 2005). These, together with the generation of a BY-2 cell line expressing a GFP-tagged COPII coat protein (LeSec13), have been of particular value in determining the relationship of ERES to Golgi stacks (Yang et al. 2005). The caveat that high levels of expression may act to perturb the system being investigated is always present when trying to elucidate the nature of cellular structures and their functional relationships to other organelles via GFP technology. Luckily, overexpression of the COPII-recruiting GTPase Sar1 and Sec13 are without effect on secretion when monitored in a protoplast enzyme secretion assay (daSilva et al. 2004; Yang et al. 2005). In addition, the credibility of GFP localization data is always enhanced when it correlates with the immunodetection of endogenous proteins; this is indeed the case with Sec13 in BY-2 cells (Yang et al. 2005). It is also most encouraging when the pattern of expression of a GFP construct in a plant cell shows the same behaviour as its counterpart in mammalian cells; again this has been demonstrated for LeSec13 in BY-2 cells (Yang et al. 2005). In marked contrast to daSilva et al. (2004), who have shown a coupling of images for Sar1-GFP and ST-YFP in tobacco leaf epidermal cells, and have therefore postulated a 1:1 relationship between ERES and Golgi stacks (“the secretory unit model”), the data of Yang et al. (2005) indicate that ERES greatly outnumber Golgi stacks in BY-2 cells. Similarly sized punctate fluorescent signals were observed after immunostaining with AtSar1, AtSec13, and AtSec23 antibodies (Fig. 2D–F; Yang et al. 2005). These punctae were located either directly on the membrane of the cortical ER or closely adjacent, giving rise to an image very reminiscent of the situation for COPII detection in animal cells (Presley et al. 1997; Stephens 2003). ER-bound Sec13 as visualized by the expression of LeSec13-GFP in dexamethasone-inducible BY-2 cells also shows a punctate distribution (Fig. 2G–I). Three features of this COPII labelling are indisputable: (1) it is very shortlived (having a half-life of less than 10 s), much more so than in mammalian cells (Stephens 2003); (2) the label is visible at the surface of the ER throughout interphase and mitosis, again different to the mammalian situation (Stephens 2003); (3) the label associates temporarily with the rims and less frequently with the faces of moving Golgi stacks, yet another characteristic not observed in animal cells, where COPII vesicles fuse homotypically to form ERGIC/VTCs (Stephens and Pepperkok 2001). Whether each punctate COPII signal represents an individual ERES is unclear, and a decision on this point must await the availability of appropriate cargo imaging data. However, the recruitment of COPII coat proteins to the surface of the ER is usually considered to reflect their interaction with export motifs in the cytoplasmic tails of membrane cargo molecules (Barlowe 2003; Duden 2003). If each visualized COPII binding site is indeed a bona fide ERES this would mean that either ERES detach from the surface of the ER and travel with Golgi stacks as a kind of COPII-coated prefusion complex or the ERES are dragged along in the plane of the mem-

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brane. In the first case, temporary contact with a moving stack would trigger the release and tethering of such complexes. For the second model to work, Golgi stacks and ERES must be directly connected via tubules or held together by scaffolding proteins. A decision on these two possible scenarios cannot, at the moment, be taken.

5 BY-2 Cells: A Model System for Studying the Action of BFA The macrocyclic fungal metabolite brefeldin A (BFA) is a most useful tool for analyzing protein trafficking in the secretory and endocytic pathways, both in animal and plant cells (Nebenführ et al. 2002). BFA binds to Sec7-type GEFs (guanidine exchange factors) which catalyze the GTPase Arf1 (Jackson and Casanova 2000). As a result, Arf1 is no longer able to recruit COPI proteins; COPI-vesicle production is prevented, and coatomer is released into the cytosol (Scheel et al. 1997). Since BFA-sensitive GEFs are often (but not exclusively) Golgi-localized it is this organelle that suffers the immediate consequences of Arf1 inactivation. The most frequently observed morphological response towards BFA treatment is therefore a disassembly of the Golgi apparatus and the incorporation of membrane and lumenal contents of the cisternae into the ER, a feature reported on numerous occasions for both animal (e. g. Sciaky et al. 1997) and plant (e. g. Satiat-Jeunemaitre and Hawes 1992; Satiat-Jeunemaitre et al. 1996; Boevink et al. 1998) cells. There are eight Arf-GEFs in the Arabidopsis database (Jürgens 2004), and some of the BFA-sensitive forms may localize to endocytic or other compartments (Steinmann et al. 1999; Baluška et al. 2002; Couchy et al. 2003; Xu and Scheres 2005). This is the probable explanation for the wide variation in morphological responses towards BFA which have been reported in the plant literature (Satiat-Jeunemaitre et al. 1996). As pointed out by Geldner et al. (2004), this variance in BFA effects is a manifestation of the “relative concentration of resistant and sensitive ARF/ARF-GEF complexes in the cell”. However, Arf1 in BY-2 cells is restricted to the Golgi apparatus (Ritzenthaler et al. 2002; Xu and Scheres 2005; see also Fig. 2A–C). Because of this, but also because effects of BFA on the endomembrane system are observable very quickly and at concentrations (10 µg/ml) comparable to those used in mammalian cell research, BY-2 cells can be considered to be a model system for demonstrating the effects of BFA on the early secretory pathway in plants. As reported by Ritzenthaler et al. (2002), application of BFA to BY-2 cells causes an almost instantaneous release of membrane-bound coatomer (as monitored by western blotting with AtSec21 antibodies) into the cytosol. Interestingly, and in contrast to the situation in animal cells (Donaldson et al. 1990), Arf1 is retained on the microsomal membranes for at least 30 min before showing signs of dissociation. These biochemical data were confirmed by immunostaining under confocal laser scanning microscope (CLSM). Changes

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in the architecture of the Golgi apparatus were recorded even after 5 min of BFA application: the cisternal number per Golgi stack decreased by two to three, with the remaining cisternae having a medial/trans-type morphology. Remarkably, although the cis-cisternae rapidly disappeared, the cis-located marker GmManI-GFP was still detectable in the CLSM, albeit as a doughnutlike image. This striking observation was interpreted by Ritzenthaler et al. (2002) as providing the first physiological evidence for cisternal movement (maturation) through the Golgi stack in a trans direction. More or less at the same time that the cis-cisternae are lost from the Golgi stacks, a second morphological change can be seen: the attachment of ER to the stack remnants, first on one side of the stack, then on both sides, giving rise to

Fig. 4. Hybrid ER-Golgi sandwich structures formed in BY-2 cells after short-term (< 15 min) treatment with 10 µm BFA. Single (A), double (B), and more seldom quintuple (C) Golgi cisternae of a trans-like character are wedged in-between ER cisternae. With increasing BFA treatment time the Golgi cisternae also fuse at the rims, with individual ER cisternae serving for direct continuities (D)

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a “sandwich” configuration. Images of such ER-Golgi sandwiches are presented in Fig. 4A–C. The ER in these sandwiches tends to be ribosome-free, and intercisternal filaments are often very prominent between the cisternae, pointing to the trans-like nature of the latter. ER-Golgi sandwiches were recorded in both the GmManI-GFP cell line and the wild type cell line, but especially in wild type cells these sandwich structures were seen to merge together into larger ER-Golgi hybrid structures after longer (30−60 min) BFA treatment times. In these hybrid structures the Golgi cisternae were now continuous with the ER, but their essential Golgi character can be deduced from the retention of islands of intercisternal filaments (Figs. 4D and 5) and from the presence of the ManI-GFP marker in the transgenic cell line (see Fig. 2E in Ritzenthaler et al. 2002). Such hybrids were frequently observed close to the nucleus and attained a considerable size (up to 10 µm; see Fig. 5; Yasuhara et al. 1995).

Fig. 5. Extreme examples of BFA-induced ER-Golgi hybrid structures in BY-2 wt cells. Intercisternal filaments remain recognizable at various positions, bearing witness to the Golgi-relevant parts of the structure

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Ritzenthaler et al. (2002) interpreted the rapid BFA-induced loss of ciscisternae and the formation of ER-Golgi hybrids in terms of the redistribution of SNARE molecules which are normally concentrated in transport vesicles (Elazar et al. 1994; Mossessova et al. 2003). The inhibition of COPI coat formation through BFA leaves the SNARES exposed all over the surface of the cisternae, especially those at the cis-face where the majority of COPI vesicles are normally formed. A spontaneous fusion with the nearest ER membrane would then occur. In some higher plant cells BFA application leads to the formation of a structure termed the “BFA-compartment”, which has the appearance of a bunch of

Fig. 6. Cytopathological effects induced by long-term (several hours) BFA treatment. A The ER has distended into a lacuna system. B Karyokinesis and cytokinesis are inhibited and the nuclei become highly convoluted, sometimes pinching off portions of the nucleus (DAPI-stained cells)

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grapes intermingled with individual Golgi cisternae (Satiat-Jeunemaitre et al. 1996). For reasons that remain unclear, such structures are seldom observed in BY-2 cells (Ritzenthaler et al. 2002), but as (Nebenführ et al. 2002) pointed out, they have been known in mammalian cells for some time and are considered to be an aggregate of trans-Golgi elements with endocytic organelles. BY-2 cells are also useful for demonstrating cytologically what can happen when BFA is applied for too long a time. Treatment periods of several hours or more lead to serious pathological anomalies. As depicted in Fig. 6A the endomembranes form a highly fenestrated continuum with large lacunae. Karyokinesis is perturbed and the nuclei are seriously deformed (Fig. 6B). Such data should convince physiologists to reconsider the usefulness of longterm experiments with this drug.

6 Appendix: Standard Fixation Protocols for Electron Microscopy and Immunostaining 6.1 Chemical Fixation for Electron Microscopy A pellet of BY-2 cells is resuspended in a primary fixative containing 2% (v:v) glutaraldehyde and 0.1 ml of saturated picric acid in 25 mM potassium phosphate buffer at pH 7.4. The fixation is performed initially for 15 min at room temperature before transferring to 4 ◦ C for 16 h. After four washes in 25 mM Pipes at pH 7.0 at room temperature, the cells are transferred to a secondary fixative containing 2% (w:v) osmium tetroxide and 0.5% (w:v) potassium ferrocyanide in 25 mM Pipes at pH 7.0 for 2 h at room temperature. After washing twice in Pipes buffer and twice in distilled water, the cells are transferred to 2% (w:v) aqueous uranyl acetate for 16 h at 4 ◦ C. After washing twice in water, the cells are dehydrated conventionally through an acetone series, and embedded in Spurr’s resin. 6.2 Chemical Fixation for Immunofluorescence BY-2 cells, 3 to 4 days after subculture, are fixed by directly adding one volume of 10% glutaraldehyde (EM grade, Electron Microscopy Sciences, Hatfield, Pennsylvania, USA) to 9 volumes of cultured cells. After 15 min at room temperature under gentle agitation, cells are washed by three cycles of centrifugation at 80 g for 2 min and replacement of supernatant with culture medium. Autofluorescence generated by glutaraldehyde fixation is quenched by adding 10 volumes of freshly prepared 0.1% (w/v) NaBH4 in saline phosphate buffer (PBS) to pelleted cells. NaBH4 treatment also presents the advantage of rendering aldehyde fixation irreversible and gently permeabilizes the plasma membrane. Cells are maintained for at least 2 h in NaBH4 before further processing, or can be stored in the dark at 4 ◦ C for up to 1 month.

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6.3 Immunofluorescence Labelling In this protocol, 0.5 ml of fixed cells are collected and allowed to settle on freshly prepared poly-l-lysine (MW > 300,000, Sigma) coated coverslips (22 × 40 mm) for 10 min. After removal of unbound cells by aspiration, the cell wall is partially digested by treatment with 0.1% (w/v) Pectolyase Y23 (Kikkoman Corp., Tokyo, Japan) and 1% (w/v) Cellulase RS (Onozuka; Yakult Honsha Corp., Tokyo, Japan) for 10−15 min at 28 ◦ C in BY-2 medium. Cells are washed three times with PBS before addition of blocking solution consisting of PBS, 5% (w/v) bovine serum albumin (BSA), 5% (v/v) normal goat serum, and 0.1% (v/v) cold water fish skin gelatin (Aurion, Wageningen, The Netherlands) for 1 h at room temperature. Cells are then incubated at 4 ◦ C overnight in appropriate dilution of primary antibody in PBS 0.1% (v/v) acetylated BSA (Aurion), washed four times in PBS, and incubated again in the dark for 4 h at room temperature with appropriate dilution of goat-anti-rabbit secondary antibodies in PBS, 0.1% (v/v) acetylated BSA. Cells are washed again four times before observation. Specificity of the labelling is controlled by omitting primary antibodies.

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daSilva LL, Snapp EL, Denecke J, Lippincott-Schwartz J, Hawes C, Brandizzi F (2004) Endoplasmic reticulum export sites and Golgi bodies behave as single mobile secretory units in plant cells. Plant Cell 16:1753–1771 daSilva LL, Taylor JP, Hadlington JL, Hanton SL, Snowden CJ, Fox SJ, Foresti O, Brandizzi F, Denecke J (2005) Receptor salvage from the prevacuolar compartment is essential for efficient vacuolar protein targeting. Plant Cell 17:132–148 Donaldson JG, Lippincott-Schwartz J, Bloom GS, Kreis TE, Klausner RD (1990) Dissociation of a 110-kD peripheral membrane protein from the Golgi apparatus is an early event in brefeldin A action. J Cell Biol 111:2295–2306 Duden R (2003) ER-to-Golgi transport: COP I and COP II function (review). Mol Membr Biol 20:197–207 Elazar Z, Orci L, Ostermann J, Amherdt M, Tanigawa G, Rothman JE (1994) ADP-ribosylation factor and coatomer couple fusion to vesicle budding. J Cell Biol 124:415–424 Follet-Gueye ML, Pagny S, Faye L, Gomord V, Driouich A (2003) An improved chemical fixation method suitable for immunogold localization of green fluorescent protein in the Golgi apparatus of tobacco Bright Yellow (BY-2) cells. J Histochem Cytochem 51:931–940 Garcia-Herdugo G, Gonzáles-Reyes JA, Gracia-Navarro F, Navas P (1988) Growth kinetics of the Golgi apparatus during the cell cycle in onion root meristems. Planta 175:305–312 Geldner N, Richter S, Vieten A, Marquardt S, Torres-Ruiz RA, Mayer U, Jürgens G (2004) Partial loss-of-function alleles reveal a role for GNOM in auxin transport-related, post-embryonic development of Arabidopsis. Development 131:389–400 Hara-Nishimura I, Matsushima R, Shimada T, Nishimura M (2004) Diversity and formation of endoplasmic reticulum-derived compartments in plants. Are these compartments specific to plant cells? Plant Physiol 136:3435–3439 Hawes C, Satiat-Jeunemaitre B (2005) The plant Golgi apparatus – going with the flow. Biochim Biophys Acta 1744:93–107 Hoffmann A, Nebenfuhr A (2004) Dynamic rearrangements of transvacuolar strands in BY-2 cells imply a role of myosin in remodeling the plant actin cytoskeleton. Protoplasma 224:201–210 Jackson CL, Casanova JE (2000) Turning on ARF: the Sec7 family of guanine-nucleotide-exchange factors. Trends Cell Biol 10:60–67 Jürgens G (2004) Membrane trafficking in plants. Annu Rev Cell Dev Biol 20:481–504 Kutsuna N, Hasezawa S (2002) Dynamic organization of vacuolar and microtubule structures during cell cycle progression in synchronized tobacco BY-2 cells. Plant Cell Physiol 43:965–973 Kutsuna N, Kumagai F, Sato MH, Hasezawa S (2003) Three-dimensional reconstruction of tubular structure of vacuolar membrane throughout mitosis in living tobacco cells. Plant Cell Physiol 44:1045–1054 Lee MC, Miller EA, Goldberg J, Orci L, Schekman R (2004) Bi-directional protein transport between the ER and Golgi. Annu Rev Cell Dev Biol 20:87–123 Mitsuhashi N, Shimada T, Mano S, Nishimura M, Hara-Nishimura I (2000) Characterization of organelles in the vacuolar-sorting pathway by visualization with GFP in tobacco BY-2 cells. Plant Cell Physiol 41:993–1001 Mogelsvang S, Gomez-Ospina N, Soderholm J, Glick BS, Staehelin LA (2003) Tomographic evidence for continuous turnover of Golgi cisternae in Pichia pastoris. Mol Biol Cell 14:2277–2291 Mossessova E, Bickford LC, Goldberg J (2003) SNARE selectivity of the COPII coat. Cell 114:483– 495 Movafeghi A, Happel N, Pimpl P, Tai GH, Robinson DG (1999) Arabidopsis Sec21p and Sec23p homologs. Probable coat proteins of plant COP-coated vesicles. Plant Physiol 119:1437–1446 Nebenführ A, Gallagher LA, Dunahay TG, Frohlick JA, Mazurkiewicz AM, Meehl JB, Staehelin LA (1999) Stop-and-go movements of plant Golgi stacks are mediated by the acto-myosin system. Plant Physiol 121:1127–1142 Nebenführ A, Frohlick JA, Staehelin LA (2000) Redistribution of Golgi stacks and other organelles during mitosis and cytokinesis in plant cells. Plant Physiol 124:135–151 Nebenführ A, Staehelin LA (2001) Mobile factories: Golgi dynamics in plant cells. Trends Plant Sci 6:160–167

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Nebenführ A, Ritzenthaler C, Robinson DG (2002) Brefeldin A: deciphering an enigmatic inhibitor of secretion. Plant Physiol 130:1102–1108 Neumann U, Brandizzi F, Hawes C (2003) Protein transport in plant cells: in and out of the Golgi. Ann Bot (Lond) 92:167–180 Pavelka M, Robinson DG (2003) The Golgi apparatus in mammalian and higher plant cells: a comparison. In: Robinson DG (ed) The Golgi apparatus and the plant secretory pathway. Ann Plant Rev 9:16–35 Phillipson BA, Pimpl P, daSilva LL, Crofts AJ, Taylor JP, Movafeghi A, Robinson DG, Denecke J (2001) Secretory bulk flow of soluble proteins is efficient and COPII dependent. Plant Cell 13:2005–2020 Pimpl P, Denecke J (2000) ER retention of soluble proteins: retrieval, retention, or both? Plant Cell 12:1517–1521 Pimpl P, Movafeghi A, Coughlan S, Denecke J, Hillmer S, Robinson DG (2000) In situ localization and in vitro induction of plant COPI-coated vesicles. Plant Cell 12:2219–2236 Polishchuk EV, Di Pentima A, Luini A, Polishchuk RS (2003) Mechanism of constitutive export from the Golgi: bulk flow via the formation, protrusion, and en bloc cleavage of large transGolgi network tubular domains. Mol Biol Cell 14:4470–4485 Presley JF, Cole NB, Schroer TA, Hirschberg K, Zaal KJ, Lippincott-Schwartz J (1997) ER-to-Golgi transport visualized in living cells. Nature 389:81–85 Quader H (1990) Formation and disintegration of cisternae of the endoplasmic reticulum visualized in live cells by conventional fluorescence and confocal laser scanning microscopy: role of calcium and the cytoskeleton. Protoplasma 151:167–170 Ritzenthaler C, Nebenführ A, Movafeghi A, Stussi-Garaud C, Behnia L, Pimpl P, Staehelin LA, Robinson DG (2002) Reevaluation of the effects of brefeldin A on plant cells using tobacco Bright Yellow 2 cells expressing Golgi-targeted green fluorescent protein and COPI antisera. Plant Cell 14:237–261 Robinson DG, Kristen U (1982) Membrane flow via the Golgi apparatus of higher plant cells. Int Rev Cytol 77:89–127 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 Saint-Jore CM, Evins J, Batoko H, Brandizzi F, Moore I, Hawes C (2002) Redistribution of membrane proteins between the Golgi apparatus and endoplasmic reticulum in plants is reversible and not dependent on cytoskeletal networks. Plant J 29:661–678 Satiat-Jeunemaitre B, Hawes C (1992) Redistribution of a glycoprotein in plant cells treated with brefeldin A. J Cell Sci 103:1153–1166 Satiat-Jeunemaitre B, Cole L, Bourett T, Howard R, Hawes C (1996) Brefeldin A effects in plant and fungal cells: something new about vesicle trafficking? J Microsc 181(Part 2):162–177 Scheel J, Pepperkok R, Lowe M, Griffiths G, Kreis TE (1997) Dissociation of coatomer from membranes is required for brefeldin A-induced transfer of Golgi enzymes to the endoplasmic reticulum. J Cell Biol 137:319–333 Schekman R, Orci L (1996) Coat proteins and vesicle budding. Science 271:1526–1533 Sciaky N, Presley J, Smith C, Zaal KJ, Cole N, Moreira JE, Terasaki M, Siggia E, LippincottSchwartz J (1997) Golgi tubule traffic and the effects of brefeldin A visualized in living cells. J Cell Biol 139:1137–1155 Staehelin LA (1997) The plant ER: a dynamic organelle composed of a large number of discrete functional domains. Plant J 11:1151–1165 Staehelin LA, Moore I (1995) The plant Golgi apparatus: structure, functional organization and trafficking mechanisms. Annu Rev Plant Physiol Plant Mol Biol 46:261–288 Steinmann T, Geldner N, Grebe M, Mangold S, Jackson CL, Paris S, Galweiler L, Palme K, Jurgens G (1999) Coordinated polar localization of auxin efflux carrier PIN1 by GNOM ARF GEF. Science 286:316–318 Stephens DJ (2003) De novo formation, fusion and fission of mammalian COPII-coated endoplasmic reticulum exit sites. EMBO Rep 4:210–217

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Stephens DJ, Pepperkok R (2001) Illuminating the secretory pathway: when do we need vesicles? J Cell Sci 114:1053–1059 Tse YC, Mo B, Hillmer S, Zhao M, Lo SW, Robinson DG, Jiang L (2004) Identification of multivesicular bodies as prevacuolar compartments in Nicotiana tabacum BY-2 cells. Plant Cell 16:672–693 Xu J, Scheres B (2005) Dissection of Arabidopsis ADP-ribosylation factor 1 function in epidermal cell polarity. Plant Cell 17:525–536 Yamada K, Fuji K, Shimada T, Nishimura M, Hara-Nishimura I (2005) Endosomal proteases facilitate the fusion of endosomes with vacuoles at the final step of the endocytotic pathway. Plant J 41:888–898 Yang YD, Elamawi R, Bubeck J, Pepperkok R, Ritzenthaler C, Robinson DG (2005) Dynamics of COPII vesicles and the Golgi apparatus in cultured Nicotiana tabacum BY-2 cells provides evidence for transient association of Golgi stacks with endoplasmic reticulum exit sites. Plant Cell 17:1513–1531 Yasuhara H, Sonobe S, Shibaoka H (1995) Effects of brefeldin A on the formation of the cell plate in tobacco BY-2 cells. Eur J Cell Biol 66:274–281 Zhang Y-H, Robinson DG (1986) The endomembranes of Chlamydomonas reinhardii. A comparison of the wild type with the wall mutants CW2 and CW15. Protoplasma 133:186–194

III.2 Molecular Study of Prevacuolar Compartments in Transgenic Tobacco BY-2 Cells S.W. Lo and L. Jiang1

1 Introduction 1.1 The Secretory Pathway Protein transport to the lysosome/vacuole is mediated by several membranebound compartments including the endoplasmic reticulum (ER), Golgi apparatus and endosomal/prevacuolar compartment (Lemmon and Traub 2000; Maxfield and McGraw 2004). Proteins reach the lysosome/vacuole because they contain sequence-specific vacuolar sorting determinants that are recognized by vacuolar sorting receptor (VSR) proteins (Neumann et al. 2003). The endosomal/prevacuolar compartments are membrane-bound organelles mediating protein traffic from both Golgi and the plasma membrane to vacuoles in eukaryotic cells (Lam et al. 2005). The plant secretory pathway is complicated due to the existence of at least two functionally and biochemically distinct vacuoles in plant cells: the lytic vacuole and protein storage vacuole (Jiang and Rogers 1998; Jauh et al. 1999; Vitale and Raikhel 1999). In addition, multiple vesicular transport pathways are responsible for transporting proteins to vacuoles in plant cells (Jiang and Rogers 2003). Recent studies have further demonstrated that prevacuolar compartments in plant cells are multivesicular bodies that merge secretory and endocytic pathways, leading to the lytic vacuole, a compartment thought to be equivalent to the mammalian lysosome or the yeast vacuole (Lam et al. 2005). 1.2 Plant Prevacuolar Compartments: Markers and Identification It has been challenging to characterize plant prevacuolar compartments (PVCs) due to the complexity of plant vacuolar systems, the existence of multiple pathways of vacuolar targeting and the lack of reliable markers. Several studies have led to the identification of lytic PVCs as multivesicular bodies in plant cells. In Arabidopsis, AtPep12p has been shown to localize to a late postGolgi compartment by immunoEM (da Silva et al. 1997). BP-80, a member of the vacuolar sorting receptor, has been localized to the Golgi apparatus and to a lytic PVC in pea root tip cells by immunoEM (Paris et al. 1997). 1 Department of Biology and Molecular Biotechnology Program, The Chinese University of Hong

Kong, Shatin, New Territories, Hong Kong, China, e-mail: [email protected] Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Similarly, a BP-80 homologue, AtELP from Arabidopsis, has been located to the Golgi apparatus and to a putative PVC characterized by ∼100 nm diameter tubules in Arabidopsis root tip cells (Sanderfoot et al. 1998). A recent study using confocal immunofluorescence microscopy with various VSR antibodies demonstrated that VSR proteins were predominantly concentrated on postGolgi, lytic prevacuolar compartments in various plant cells (Li et al. 2002). Since the BP-80 reporter (a fusion protein containing the transmembrane domain and cytoplasmic tail of BP-80) (Jiang and Rogers 1998) colocalized with VSR proteins, both VSR proteins and BP-80 reporter are thus markers for defining PVCs. Using these reliable PVC markers and transgenic BY-2 cells expressing the YFP-BP-80 reporter, the lytic PVCs were identified via immunoEM as multivesicular bodies in tobacco BY-2 cells (Tse et al. 2004). In addition, wortmannin induced the lytic PVCs to form small vacuoles in BY-2 cells (Tse et al. 2004). Compared to the lytic pathway, relatively little is known about the molecular components responsible for sorting proteins to the protein storage vacuole (Vitale and Hinz 2005). Several studies strongly indicate the existence of PVCs as a pathway leading to the protein storage vacuole. The α-TIP (tonoplast intrinsic protein) reporter (a reporter containing the BP-80 transmembrane domain and the cytoplasmic tail of α-TIP) (Jiang and Rogers 1998) trafficked directly from ER to PSV probably via a PVC termed DIP (dark intrinsic protein) organelles in tobacco (Jiang et al. 2000), while the precursor-accumulating (PAC) vesicles might serve as a PVC for delivering proteins to PSV directly from the ER in pumpkin seeds (Hara-Nishimura et al. 1998). Figure 1 shows a working model of the protein traffic and vesicular pathways leading to the lytic vacuole and protein storage vacuole in plant cells in which a hypothetical PVC for the protein storage vacuole might serve as an intermediate compartment merging traffic from Golgi and the ER directly. 1.3 Hypothesis and Tools Used in this Study Tobacco Bright Yellow 2 (BY-2) cells have served as useful tools in a wide range of physiological, developmental and cellular studies (Nagata et al. 2004) because of their excellent growth, synchronization and transformation properties (Geelen and Inze 2001). BY-2 cells have been particularly useful in studies of protein localization and organelle dynamics in the secretory pathway using green fluorescent protein (GFP) tagging (Mitsuhashi et al. 2000; Nebenfuhr et al. 2000; Kutsuna and Hasezawa 2002; Tse et al. 2004). Previously we successfully used transgenic BY-2 cells expressing GFP fusions to study protein trafficking and organelle dynamics. The generation of transgenic tobacco BY-2 cells expressing the GONST1-YFP and the YFP-BP-80 reporters has allowed us to characterize and identify the lytic PVC as a multivesicular body (Tse et al. 2004). Therefore, in this study, we test the hypothesis that BY-2 cells contain two distinct PVCs as pathways leading to LV and PSV,

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Fig. 1. Plant cells contain two prevacuolar compartments in the secretory pathways. Shown is a schematic diagram of three vesicular transport pathways leading to either the lytic vacuole (LV) or protein storage vacuole (PSV) from the endoplasmic reticulum (ER). The ER–Golgi– LV and ER–Golgi–PSV pathways involve clathrin-coated vesicle (CCV) and dense vesicle (DV) respectively (Jiang and Rogers 2003), while a direct ER–PSV pathway is mediated by precursoraccumulating (PAC) vesicle in pumpkin cotyledon or dark intrinsic protein (DIP) organelle in tobacco seed (Jiang et al. 2000). The prevacuolar compartment (PVC) for LV is a multivesicular body in tobacco BY-2 cells that is defined by vacuolar sorting receptor (VSR) protein and the YFPBP-80 reporter (Tse et al. 2004), while the hypothetical PVC for PSV is defined by the GFP-α-TIP reporter (Jiang and Rogers 1998)

where the lytic PVC and the PVC for the protein storage vacuole are marked by the BP-80 reporter and the α-TIP reporter respectively (Jiang and Rogers 1998; Tse et al. 2004). Towards this goal, we have generated transgenic BY-2 cell lines expressing these two GFP fusion reporters and preliminary results indicate that they may mark distinct PVC populations in BY-2 cells.

2 Results and Discussion 2.1 Reporters for Golgi and Distinct PVCs GFP localized to both nucleus and cytoplasm when it was expressed alone in BY-2 cells (Nakagami et al. 2002;Yamaguchi et al. 2002). However, when GFP is tagged with various targeting determinants, it was localized to different organelles and gave characteristic fluorescent patterns of destined organelles (Mitsuhashi et al. 2000). Therefore, GFP is a good reporter for studying protein subcellular localization and identifying organelles in transgenic BY-2 cells. Using YFP as a reporter, we had previously demonstrated that the YFP-BP-80 reporter and GONST1-YFP reporter located to lytic PVC and Golgi organelles

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Fig. 2. Chimeric constructs used in this study for the identification of distinct organelles in transgenic tobacco BY-2 cells. Three transgenic tobacco BY-2 cell lines expressing GONST1-YFP fusion, the GFP-BP-80 reporter or the GFP-α-TIP reporter were generated and used in this study. The predicted subcellular localization of these fusion proteins in transgenic BY-2 cells were also indicated. CaMV 35S pro Cauliflower mosaic virus 35S promoter; SP signal peptide; GFP green fluorescent protein; TMD transmembrane domain from BP-80; CT cytoplasmic tail from either BP-80 or α-TIP (tonoplastic intrinsic protein); NOS Nos terminator; PVC prevacuolar compartment; LV lytic vacuole; PSV protein storage vacuole

respectively when they were stably expressed in tobacco BY-2 cells (Tse et al. 2004). Therefore in this study, we first test the hypothesis that plant cells contain two distinct PVC populations leading to the lytic and protein storage vacuoles, in which the lytic PVC is marked by the GFP-BP-80 reporter (where the YFP was replaced by GFP in the YFP-BP-80 construct) while the PVC for the protein storage vacuole is represented by the GFP-α-TIP reporter (Jiang and Rogers 1998) when they are stably expressed in transgenic tobacco BY-2 cells. In addition, the transgenic tobacco BY-2 cell line expressing the Golgi marker GONST1-YFP (Tse et al. 2004) was used as a control. Figure 2 shows the key components of the three constructs used in this study and their predicted subcellular localization in transgenic tobacco BY-2 cells. 2.2 Expression of the Reporter Proteins in Transgenic Tobacco BY-2 Cells Upon Agrobacterium-mediated transformation and selection on MS media containing kanamycin, more than fifteen individual transgenic tobacco BY-2 cell lines were generated that expressed either the GFP-BP-80 reporter or the GFP-α-TIP reporter. Typical punctate fluorescent signals were readily detected from all transgenic cells expressing either reporter (Fig. 3A). Similarly, the Golgi marker GONST-YFP also exhibited typical punctate fluorescent signals in transgenic BY-2 cells (Fig. 3A). These results are consistent with our previous study in which both lytic PVC and Golgi showed punctate patterns in BY-2 cells (Tse et al. 2004). To further confirm the expression of these reporter proteins in newly generated transgenic cells, western blot analysis with GFP antibodies was carried out, where 3-day-old cultured cells were used for protein extraction. As shown

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Fig. 3. Expression of GFP or YFP reporter proteins in transgenic BY-2 cells. A Confocal fluorescent signals detected from 3-day-old living transgenic BY-2 cell lines expressing GONST1-YFP, GFPBP-80 and GFP-α-TIP reporter proteins. GFP Green fluorescent protein; YFP yellow fluorescent protein; DIC differential-interference-contrast. Bar 50 µm. B Western blot analysis of expressed proteins in 3-day-old transgenic BY-2 cell lines expressing either GFP-BP-80 or GFP-α-TIP reporter as detected by anti-GFP antibodies. WT Wild type BY-2 cell; CS cell soluble fraction; CM cell membrane fraction

in Fig. 3B, when GFP antibodies were used to detect the reporter, a strong band with expected molecular mass of 37.9 kDa was detected in the cell membrane (CM) fraction (lane 4) of transgenic cells expressing the GFP-BP-80 reporter, while an identical weak band was detected in the cell-soluble (CS) fraction (lane 3). Similarly, in transgenic cells expressing the GFP-α-TIP reporter, a strong band with correct size at 36.2 kDa was detected in the CM fraction (lane 6), while a similar weak band was detected in the CS fraction (lane 5). These bands must represent their corresponding expressed reporter proteins in transgenic BY-2 cells because no such bands were detected in either the CS or CM fraction of wild type (WT) untransformed control cells (Fig. 3B, lanes 1 and 2). These results indicate that both reporters remained as intact full-length integral membrane proteins because of their correct sizes and their being mainly present in the CM fraction. Thus, transgenic BY-2 cell lines expressing GFP-BP-80 and GFP-α-TIP reporters with expected expression patterns, molecular weight and membrane distribution were successfully generated.

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2.3 Reporter Profiles During Cell Culture Cycle Transgenic BY-2 cell lines expressing GFP fusions have been useful tools in the study of protein trafficking and subcellular localization (Neuhaus 2002). However, even in the same cell line expressing the same construct, the fluorescent signal patterns in transgenic BY-2 cells could be changed over the 7-day culture period due to the proteolytic degradation of cellular proteins at later stages after subculture (Mitsuhashi et al. 2000). Therefore, a study was carried out to determine the fluorescent signals and patterns during the course of the 7-day culture period so that the correct stages of cells can be used in subsequent immunolabeling studies for the identification of GFP-marked organelles. Transgenic BY-2 cells were subcultured every 7 days. During the 7-day culture period, cells were collected every day (from day 1 to day 7) either for direct detection of fluorescent signals via confocal imaging or for protein extraction to be used in western blot analysis. As shown in Fig. 4, upon subculture into fresh MS medium, typical punctate fluorescent signals were detected from day 1 to day 3 in transgenic BY-2 cells expressing either GFP-BP-80 or GFP-α-TIP reporter. However, such punctate signals gradually disappeared from day 4 to day 7, as determined by the increased detection of diffused fluorescent signals inside the vacuoles in cells expressing either reporter. Interestingly, some of the 7-day-old cells showed a sudden increase in diffuse signals. Such profiles of fluorescent signals were repeatable after the 7-day-old cells were subcultured into a fresh medium for both transgenic cell lines. To further investigate the changes of fluorescent signals detected in confocal imaging, western blot analysis was carried out using proteins extracted from collected cells from the 7-day period. In cells expressing the GFP-BP80 reporter, the full-length integral membrane reporter proteins with correct sizes at 38 kDa were detected by GFP antibodies in the CM fractions. However, the detected signals were strong during the first 3 days after subculture (i. e. from day 1 to day 3), decreased dramatically from day 4 and remained weak thereafter (Fig. 5A). Similarly, in cells expressing the GFP-α-TIP reporter, the amount of full-length reporter being detected in the CM fraction was highest from day 1 to day 3 upon subculture, but dramatically reduced from day 4 (Fig. 5B). Interestingly, in 7-day-old cells expressing either reporter, a strong band at 26 kDa which represents the free GFP was detected in the CS fraction, indicating that the GFP was cleaved and thus separated from the transmembrane domain of the original fusion protein upon reaching the vacuoles. The results obtained from the above confocal imaging demonstrate that both GFP-BP-80 and GFP-α-TIP reporter proteins exhibit punctate patterns during early stages of cultures but diffuse patterns during later stages of cultures, thus indicating different subcellular localization of the same reporter at different culture stages. Such a notion was also supported by the western blot analysis where soluble GFP was highly detected in the CS fraction of 7-day-old cells. We therefore hypothesize that the punctate patterns from day 1 to day 3 represent PVC localization of the full-length reporter proteins, while the diffused

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Fig. 4. Profiles of fluorescent signals in living transgenic BY-2 cells expressing PVC markers during the 7-day culture period. Transgenic BY-2 cells expressing either GFP-BP-80 reporter (A) or GFPα-TIP reporter (B) were subcultured every 7 days. During this 7-day culture period, living cells were collected at indicated times (D1 to D7) for observation of fluorescent signals under confocal microscopy. DIC Differential-interference-contrast. Bar 50 µm

patterns thereafter indicate vacuolar localization of cleaved free GFP. Such an hypothesis is also consistent with the possibility that the reporters move from PVC to vacuole for degradation during the later stages of the culture period. In addition, the changes in fluorescent signals could represent the normal

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Fig. 5. Western blot analysis of expressed proteins in transgenic BY-2 cells expressing PVC markers during the 7-day culture period. Transgenic BY-2 cells expressing either GFP-BP-80 or GFP-α-TIP reporter were subcultured every 7 days. During this 7-day culture period, cells were collected at indicated times (D1 to D7) for protein extraction into cell soluble (CS) and cell membrane (CM) fractions before being used in western blot analysis using anti-GFP antibodies

turnover of organelles marked by the GFP reporters. Moreover, autophage can be induced by nutrient starvation which causes protein degradation (Moriyasu et al. 2003). The absence of diffuse signals from 2- to 3-day-old cells expressing the GFP-α-TIP reporter could be due to light-dependent degradation of GFP, where light could induce conformational change in GFP for easier degradation by vacuolar proteinases in an acidic pH environment (Tamura et al. 2003). 2.4 Wortmannin-Induced GFP-Marked Organelles to Vacuolate in Transgenic BY-2 Cells As discussed previously, transgenic cells expressing either reporter showed typical punctate signals during the first 3 days after subculture, and these punctate signals might represent PVC localization. Therefore, either day 2 or day 3 cells were used for subsequent confocal immunofluorescence microscopy study and drug treatment study to identify GFP-marked organelles in transgenic BY-2 cells. Wortmannin inhibits phosphatidylinositol 3-kinase [PI(3)P], an enzyme that plays a crucial role in clathrin-coated vesicles budding in yeast and mammals (Thelen et al. 1994). In plant cells, wortmannin also inhibited PI(3)P production in Arabidopsis protoplasts (Kim et al. 2001) and prevented vacuolar sorting during autophage (Takatsuka et al. 2004), and blocked a fluorescent dye FM1-43 from reaching the vacuole during endocytosis (Emans et al. 2002). In addition, the recycling of VSR from PVC back to Golgi was also wortmannin sensitive, which resulted in leakage of BP-80-ligand to the vacuole (daSilva et al. 2005). Wortmannin also induced morphological changes of PVCs. When normal rat kidney cells were treated with wortmannin, extensive vacuolation and swelling of late endosomes were observed (Bright et al. 2001). Similarly, the

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Fig. 6. Wortmannin-induced organelles marked by both GFP-BP-80 and GFP-α-TIP reporters vacuolating in transgenic BY-2 cells. Transgenic BY-2 cells expressing GONST1-YFP, GFP-BP-80 and GFP-α-TIP reporters were treated with wortmannin at 16.5 µM for 1.5 h and then subjected to confocal imaging. Enlarged organelles and small vacuoles were observed in cell lines expressing either GFP-BP-80 or GFP-α-TIP but not in GONST1-YFP cells. The insets are enlarged images of vacuoles indicated by arrowheads. DIC Differential-interference-contrast. Bar 50 µm

lytic PVCs marked by the YFP-BP-80 reporter were induced by wortmannin (at 15 or 33 µM) to form small vacuoles in transgenic tobacco BY-2 cells (Tse et al. 2004). However, the same concentrations of wortmannin did not cause any visible changes in Golgi marked by GONST1-YFP in transgenic BY-2 cells. Thus, wortmannin treatment can be used as a tool to identify PVC as well as to distinguish the lytic PVC from Golgi in BY-2 cells. In order to determine whether organelles marked by the GFP-α-TIP reporter were PVCs, wortmannin treatment study was carried out using 2- to 3-day-old transgenic cells expressing GFP-α-TIP reporter, GFP-BP-80 reporter and GONST1-YFP, where the later two cell lines were used as controls. As shown in Fig. 6, upon wortmannin treatment at 16.5 µM for 1.5 h, GONST1YFP-marked Golgi remained unchanged as punctate patterns. However, in cells expressing either GFP-α-TIP or GFP-BP-80 reporter, small vacuoles were observed, indicating that organelles marked by either reporter might represent PVCs in transgenic BY-2 cells. These results are consistent with our previous study in which the lytic PVC marked by the YFP-BP-80 reporter formed small vacuoles in response to wortmannin treatment (Tse et al. 2004). In addition, it seems that organelles marked by GFP-α-TIP and GFP-BP-80 reporters had different degrees of response to wortmannin. Among 30 cells investigated, almost all organelles marked by the GFP-BP-80 reporter were induced to form small vacuoles upon wortmannin treatment, but some organelles marked by the GFP-α-TIP reporter remained unchanged (data not shown; see also Fig. 7 below). Taken together, these results demonstrate that organelles marked by GFP-α-TIP or GFP-BP-80 represent PVCs in BY-2 cells and that the lytic PVCs marked by GFP-BP-80 might be different from the PVCs marked by GFP-α-TIP.

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2.5 Organelles Marked by GFP-BP-80 and GFP-α-TIP Are Distinct PVCs in Transgenic BY-2 Cells Since wortmannin induced organelles marked by either GFP-BP-80 or GFPα-TIP reporter to form small vacuoles, organelles marked by both reporters are thus PVCs. To further confirm the identity of PVCs marked by the GFP-αTIP reporter and determine whether these PVCs are distinct from the PVCs marked by the GFP-BP-80 reporter, an immunolabeling study using VSR antibodies was carried out. We had previously demonstrated that VSR proteins were predominantly concentrated on PVCs and thus can be used as markers to define PVCs (Li et al. 2002). The PVC localization of the YFP-BP-80 reporter was further confirmed by confocal immunofluorescent study where the reporter was largely colocalized with VSR antibodies that also identified PVCs as multivesicular bodies in BY-2 cells (Tse et al. 2004). To determine whether GFP-α-TIP and GFP-BP-80 reporters mark the same PVC populations, a confocal immunofluorescence study with VSR antibodies was carried out. Transgenic cells were first treated with wortmannin at 16.5 µM for 1.5 h, followed by fixation and immunolabeling with VSR antibodies. As depicted in Fig. 7, in cells expressing the GFP-BP-80 reporter, most GFP-marked organelles (shown in green) were induced to form small vacuoles that colocalized with VSR (shown in red). This result is consistent with our previous

Fig. 7. Anti-VSR colocalized differently with GFP-BP-80 reporter and GFP-α-TIP reporter in wortmannin-treated transgenic cells. Transgenic BY-2 cells expressing these two reporters were fixed before (A) or after (B) treatment with wortmannin at 16.5 µM for 1.5 h, followed by labeling with VSR antibodies and detection by rhodamine-conjugated secondary antibodies (red), while the GFP signals were readily detected (green). Colocalization of GFP reporter (green) with VSR antibodies (red) is indicated by yellow in the merged images. Arrowheads and arrows indicate examples of colocalization and separation respectively. Enlarged images are examples of small vacuoles. DIC Differential-interference-contrast. Bar 50 µm

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Table 1. Quantification of antibody colocalization in confocal immunofluorescence images performed on GFP-BP-80 and GFP-α-TIP cells after wortmannin treatment at 16.5 µM for 1.5 h. The extent of colocalization was calculated in the direction from left to right. A t-test was performed in order to compare the results from individual paired assays of the two reporters. The resulting P values were statistically significant (P 100 queries) it is useful to get an idea about the random false positive hit rate by querying a random database. Use a large random database, select the proper proteolytic agent as trypsin and do not allow for species restrictions, MW restrictions or pI restrictions. • Allow for one missed cleavage as this frequently occurs and improves the matching; allowing for more missed cleavages typically reduces the score. • Allow all fixed modifications (e. g. carbamidomethyl cysteine if iodo acetamide was used as the alkylating agent). Optionally, depending on the quality of the alkylation procedure, the cysteine modification might be detailed in the variable modification table. • Allow for oxidized methionine as a frequent observed variable modification. Only when searching for a specific PTM should the respective modification be allowed for; it should never be set as a default. • Use a peptide mass tolerance that is roughly twice the accuracy value with which the spectra were acquired (typically around 30 ppm for internal calibrated PMF spectra or 0.5 Da for peptide sequence analyses). • Select the proper mass value denominators (typically MH+ and monoisotopic; ask the MS operator if you are not sure). • In general there should not be many scores exceeding the threshold value. If a high score for a search against this database is obtained, the first thing to investigate is whether the experimental mass values are independent. For example, do any pairs of mass values differ by less than the specified mass tolerance? Also, if a variable modification has been specified, and the difference between pairs of mass values equals the mass difference due to the modification, this can lead to high scores. 7.2.2 Second Submission to the Mascot Search Engine

• Select a large database such as NCBInr, and use the same conditions as for the first search. • The obtained results table will already list true positives but also will reveal experimental artifacts originating from human, animal, fungal and bacterial contamination, along with other false positives. Individual sample queries indicating false positives in this first submission should not be omitted from the second submission but should be examined with extra care. In case of an obvious contamination, the set of peptides matching with the false positive protein can be omitted in the second search. In case there is a possible ortholog match of a species belonging to another kingdom, the list containing the protein matches often contains a member of the same protein family belonging to the plant kingdom.

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7.2.3 Third Submission to the Mascot Search Engine

• Use Viridiplantae as the taxonomic restriction, use the same conditions as for the first pass and search against all available databases to get the best result. In case of peptide sequence data, include the EST databases. • The obtained results should contain more positives; however, the additional positives should be individually checked with data obtained in the previous submissions and verified as true positives. In order to gather more protein information from the EST-based results, a BLAST has to be performed. 7.2.4 Fourth Submission to the Mascot Search Engine (Peptide Mass Sequence Data only)

• A selection of queries can be resubmitted with a wider range of modifications and relaxed enzyme specificity. • The obtained results might reveal unexpected PTMs. Acknowledgements. The authors wish to thank Professor Henri Van Onckelen and Professor Edgard Esmans for their efforts in creating the Center for Proteome Analysis and Mass Spectrometry in Antwerp. K.L. and E.W. are postdoctoral fellows of the Fund for Scientific Research – Flanders (Belgium) (F.W.O. – Vlaanderen).

References Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215:403–410 Appel RD, Bairoch A, Sanchez JC, Vargas JR, Golaz O, Pasquali C, Hochstrasser DF (1996) Federated two-dimensional electrophoresis database: a simple means of publishing twodimensional electrophoresis data. Electrophoresis 17:540–546 Baginsky S, Siddique A, Gruissem W (2004) Proteome analysis of tobacco bright yellow-2 (BY-2) cell culture plastids as a model for undifferentiated heterotrophic plastids. J Proteome Res 3:1128–1137 Berggren K, Chernokalskaya E, Steinberg TH, Kemper C, Lopez MF, Diwu Z, Haugland RP, Patton WF (2000) Background-free, high sensitivity staining of proteins in one- and twodimensional sodium dodecyl sulfate-polyacrylamide gels using a luminescent ruthenium complex. Electrophoresis 21:2509–2521 Craig R, Beavis RC (2004) TANDEM: matching proteins with tandem mass spectra. Bioinformatics 20:1466–1467 Görg A, Weiss W, Dunn MJ (2004) Current two-dimensional electrophoresis technology for proteomics. Proteomics 4:3665–3685 Griffin TJ, Gygi SP, Ideker T, Rist B, Eng J, Hood L, Aebersold R (2002) Complementary profiling of gene expression at the transcriptome and proteome levels in Saccharomyces cerevisiae. Mol Cell Proteomics 1:323–333 Habermann B, Oegema J, Sunyaev S, Shevchenko A (2004) The power and the limitations of crossspecies protein identification by mass spectrometry-driven sequence similarity searches. Mol Cell Proteomics 3:238–249

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Laukens K, Lenobel R, Strnad M, Van Onckelen H, Witters E (2003) Cytokinin affinity purification and identification of a tobacco BY-2 adenosine kinase. FEBS Lett 533:63–66 Laukens K, Deckers P, Esmans E, Van Onckelen H, Witters E (2004) Construction of a twodimensional gel electrophoresis protein database for the Nicotiana tabacum cv. bright yellow2 cell suspension culture. Proteomics 4:720–727 Matsuoka K, Demura T, Galis I, Horiguchi T, Sasaki M, Tashiro G, Fukuda H (2004) A comprehensive gene expression analysis toward the understanding of growth and differentiation of tobacco BY-2 cells. Plant Cell Physiol 45:1280–1289 Matthiesen R, Bunkenborg J, Stensballe A, Jensen ON, Welinder KG, Bauw G (2004) Databaseindependent, database-dependent, and extended interpretation of peptide mass spectra in VEMS V2.0. Proteomics 4:2583–2593 Neverova I, Van Eyk JE (2005) Role of chromatographic techniques in proteomic analysis. J Chromatogr B Analyt Technol Biomed Life Sci 815:51–63 Perkins DN, Pappin DJ, Creasy DM, Cottrell JS (1999) Probability-based protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20:3551–3567 Rabilloud T, Strub JM, Luche S, van Dorsselaer A, Lunardi J (2001) A comparison between Sypro Ruby and ruthenium II tris (bathophenanthroline disulfonate) as fluorescent stains for protein detection in gels. Proteomics 1:699–704 Shevchenko A, Sunyaev S, Loboda A, Shevchenko A, Bork P, Ens W, Standing KG (2001) Charting the proteomes of organisms with unsequenced genomes by MALDI-quadrupole time-offlight mass spectrometry and BLAST homology searching. Anal Chem 73:1917–1926 Shinozaki K, Ohme M, Tanaka M, Wakasugi T, Hayashida N, Matsubayashi T, Zaita N, Chunwongse J, Obokata J, Yamaguchi-Shinozaki K, Ohto C, Torazawa K, Meng BY, Sugita M, Deno H, Kamogashira T, Yamada K, Kusuda J, Takaiwa F, Kato A, Tohdoh N, Shimada H, Sugiura M (1986) The complete nucleotide sequence of the tobacco chloroplast genome: its gene organization and expression. EMBO J 5:2043–2049 Stasyk T, Huber LA (2004) Zooming in: fractionation strategies in proteomics. Proteomics 4:3704– 3716 Sugiyama Y, Watase Y, Nagase M, Makita N, Yagura S, Hirai A, Sugiura M (2005) The complete nucleotide sequence and multipartite organization of the tobacco mitochondrial genome: comparative analysis of mitochondrial genomes in higher plants. Mol Genet Genomics 272:603–615 The Gene Ontology Consortium (2000) Gene ontology: tool for the unification of biology. Nature Genet 25:25–29 Tian Q, Stepaniants SB, Mao M, Weng L, Feetham MC, Doyle MJ, Yi EC, Dai H, Thorsson V, Eng J, Goodlett D, Berger JP, Gunter B, Linseley PS, Stoughton RB, Aebersold R, Collins SJ, Hanlon WA, Hood LE (2004) Integrated genomic and proteomic analyses of gene expression in Mammalian cells. Mol Cell Proteomics 3:960–969 Wilm M, Shevchenko A, Houthaeve T, Breit S, Schweigerer L, Fotsis T, Mann M (1996) Femtomole sequencing of proteins from polyacrylamide gels by nano-electrospray mass spectrometry. Nature 379:466–469 Witters E, Laukens K, Deckers P, Van Dongen W, Esmans E, Van Onckelen H (2003) Fast liquid chromatography coupled to electrospray tandem mass spectrometry peptide sequencing for cross-species protein identification. Rapid Commun Mass Spectrom 17:2188–2194 Yu H, Luscombe NM, Lu HX, Zhu X, Xia Y, Han JD, Bertin N, Chung S, Vidal M, Gerstein M (2004) Annotation transfer between genomes: protein–protein interologs and protein–DNA regulogs. Genome Res 14:1107–1118

VI.2 EST and Microarray Analysis of Tobacco BY-2 Cells K. Matsuoka and I. Galis1

1 Introduction Tobacco BY-2 cells as well as tobacco plants are widely used as model species in plant biology, although genome-related analysis has not yet been carried out. While working extensively on protein trafficking in the secretory pathway using this tobacco cell line as a model (see references in Matsuoka 2004; Toyooka and Matsuoka, this vol.), we decided to collect transcriptomic information in order to improve the usage of this cell line in plant cell biology when one of us started work at RIKEN. The first phase of this work, the transcriptome analysis of BY-2 (TAB) project to find and characterize novel and useful genes using tobacco BY-2 information (Fig. 1; http://mrg.psc.riken.go.jp/strc/index.htm), was funded by the RIKEN Plant Science Center between October 2000 and March 2005. In this chapter, we summarize the outcome of the TAB project, especially the expression sequence tag (EST) and microarray analyses, and discuss examples of gene characterization in combination with tobacco microarray and reverse genetics.

2 The RIKEN Tobacco BY-2 EST and BY-2 Microarray 2.1 EST One of the main focuses of our transcriptomic work is to understand how plant cells change gene expression during cell division and after termination of cell division. We therefore started to collect ESTs from lag, log and stationary phase BY-2 cells. To decrease the redundancy of the library contents we normalized by PCR before analyzing EST sequences of the resultant library MAT001. The average length of the EST inserts is approximately 1 kb. After analyzing the sequences of about 10,000 clones from one end we obtained 9,190 ESTs of approximately 500 bp. The 9,190 sequences grouped into 7,976 unique sequences. This EST library contained many cDNA fragments with no homology to any genes at both the nucleotide and protein level as well as genes that can encode proteins for transposons (Fig. 2). Using this EST set, we generated a cDNA 1 RIKEN Plant Science Center, 1-7-22 Suehirocho, Tsurumi-ku, Yokohama 230-0045, Japan, e-mail:

[email protected] Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Fig. 1. The RIKEN TAB project and its focus

Fig. 2. Gene distribution of tobacco BY-2 ESTs from two different libraries. Possible proteins that could be encoded by the BY-2 ESTs were analyzed with a BLASTX search. Left graphs Percentage of genes that showed homology to genes with known/predicted functions, for unknown proteins and no homology to known genes. Right graphs Distribution of genes among the EST-encoded genes with known/predicted functions

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microarray and analyzed the global changes of gene expression during cell growth (Matsuoka et al. 2004). Unfortunately, however, this EST collection contained only a few cDNA fragments for genes for respiration and other highly expressed genes including genes for ribosomal proteins. In addition to this uneven distribution of cDNAs, we did not find most of the cDNAs for secondary metabolism including methyl jasmonate (MJ)-regulated genes for alkaloid synthesis. Sequencing of another approximately 4,000 clones from the same library did not solve these problems. We speculated two reasons for such uneven clone distribution: too much normalization of cDNA and no expression of some classes of genes in the BY-2 cells grown under standard culture conditions. As BY-2 cells are a good model for the analysis of secondary metabolites (Nagata et al. 2004), we thought that such a biased distribution of clones on microarray reduced the usefulness of our cDNA microarray. We therefore prepared mRNAs from BY-2 cells that have been incubated with medium without sugars, or without auxin but with one of the following phytohormones: abscisic acid, brassinolide, 6-benzyladenine, GA3 , methyl jasmonate or salicylic acid. After recovering mRNAs from these cells and pooling these mRNAs we made a cDNA library and subjected it to a single-round normalization. Thereafter 5,512 ESTs were obtained from this second library (MAT005). These clones (our reference numbers BY20001–BY35384; DDBJ accession nos. BP530024– BP535535) contained several cDNA fragments for secondary metabolism including known jasmonate-inducible transcripts in BY-2 cells (e. g. BY22111, BY30333 and BY33224 for spermidine synthase). In addition, a significant proportion of clones in this library encode genes for respiration and for protein synthesis including ribosomal proteins (Fig. 2). These data indicate that the second library covers a higher degree of genes with housekeeping function and genes that are not expressed in cells grown under normal condition. Therefore we concluded that the combination of the first and second EST clones covers a wide range of transcript information in tobacco BY-2 cells. 2.2 Tobacco cDNA Microarray Using the EST clones, we generated a cDNA microarray to monitor the global change of gene expression in BY-2 cells under several different conditions. We first made a microarray using the ESTs that had been sequenced at the beginning and used to analyze gene expression during growth (Matsuoka et al. 2004). Usually, microarrays using cDNA as a target are not quantitative because the amount of target DNA spotted to the array surface varies significantly. The target gene fragments that we used for microarray contained vector fragments at the ends of the DNA because we amplified the insert of EST clones by PCR using universal primers that anneal to the vector part of the clones. Using the advantage that the vector fragments are attached to the ends of the spotted DNA on microarray, we developed a normalization method to ignore the difference

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Fig. 3. Comparison of microarray data and RT-PCR. To obtain evidence that our microarray data were quantitative, six genes with different expression patterns were selected and quantified by quantitative RT-PCR. The average of three experiments is shown

in the amount of spotted DNA on the array. The method that we employed is a version of the two-color method using Cy5-labeled cDNA probes and Cy3-labeled reference oligonucleotide probes that can hybridize to the vector part of the spotted DNA. We calculated the ratio of Cy5 and Cy3 signals and used this value as a normalized intensity of the signal of a particular gene on the spot. Another normalization method that we employed was to spot the same DNA on a different position of the glass slide and to select data that show that the differences in Cy3-normalized intensities of these two spots are less than threefold. We found that this data processing method yielded highly quantitative data; almost all the six example genes showed an almost identical pattern and level of gene expression by microarray and quantitative PCR (Fig. 3). Thus we concluded that the data processing method that we employed is suitable to detect quantitative differences in gene expression in tobacco BY-2 cells. Using the data collected by this method we first compared the difference in gene expression during the growth of tobacco BY-2 cells (Matsuoka et al. 2004). We found that the stationary-phase cells express higher levels of genes for signal perception as well as transporters for the absorption of nutrients from the medium. In contrast, many genes that showed higher levels of expression at the log phase are genes related to cytokinesis. Most of these genes were suppressed in cells incubated with medium without auxin and either with other hormones or without hormones (Matsuoka et al. 2004). Based on these findings

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Fig. 4. In situ localization of two representative log-high transcripts at root meristem. Two gene probes that showed highest expression at log phase and a constantly expressed control were hybridized to root sections of tobacco. After hybridization and detection of DIG-labeled probes with anti-DIG-AP and BCIP/NBT, sections were stained with DAPI to localize nucleus and dividing cells (right). Left Sense probe controls; center antisense probe to detect localization. Right bottom graphs Expression pattern of these three genes

we speculated that the combination of the filtering of genes using our growthdependent comprehensive transcriptomic information and microarray data on the spatial difference in gene expression in Arabidopsis roots will allow us to identify potential genes for cell cycle progression. This assumption is based on the fact that root meristems are the only sites where cytokinesis takes place in roots. Actually two of the genes that showed a high level of expression at the log phase showed specific expression at the root meristemic region in tobacco (Fig. 4). Therefore, using gene filtration methods we searched for novel genes that are likely be involved in plant cell division and found 21 such genes (Matsuoka et al. 2004). Future characterization of the products of these genes will reveal the biological function of these genes as well as the feasibility of the data filtration method. The result of the growth-dependent microarray analysis indicated that a global change of gene expression takes place during the growth of BY-2 cells. This suggests that even in cultured plant cells the gene expression is dynamically changed during changes in cellular conditions. Thus it is interesting to analyze the relationship between phytohormones and differentiation

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of tobacco cells. Unfortunately, however, few secondary metabolism gene fragments were present in our first EST collection. Thus we made a second library from hormone-treated cells, as described above, and used this to prepare a second set of microarray. In this array, we used 16,224 selected cDNA inserts from both libraries and several specific, EST-non-represented tobacco genes. In the next section we describe our detailed analysis of the methyl jasmonate response of tobacco BY-2 cells based on this microarray analysis.

3 An Example of Microarray-Based Analysis: Jasmonate-Dependent Gene Expression and Metabolic Studies 3.1 Background BY-2 cells lost their totipotentiality, i. e. the ability to dedifferentiate from and back into specialized tissues which can be achieved in plant cells by manipulation of the hormonal content in the cultivation medium. In this regard, BY-2 cells appear to have less potential in comparative morphogenetic studies. On the other hand, the cells repeatedly showed their usefulness in cellular studies, including the dissection of cell division machinery (see Sect. I in Nagata et al. 2004). In addition, the cells retained their ability for what is described here as “metabolic differentiation”. For example, auxin deficiency together with increased cytokinin content leads to amyloplast formation in the cells (Sakai et al. 2004). In the following text, we will show another example of metabolic differentiation in BY-2 cells that was examined by comprehensive BY-2 microarray. The plant stress hormone, methyl jasmonate (MJ), was used to induce accumulation of natural bioproducts (secondary metabolites) in BY-2 cells. It is shown that isolated BY-2 cells can normally receive the MJ hormonal signal and provide a useful model for investigation of secondary metabolism as well as MJ signal transduction pathways. The data may also be compared with two other recent MJ large-scale profiling results using AFLP for BY-2 cells (Goossens et al. 2003) and monocot grass sorghum cDNA microarray (Salzman et al. 2005). 3.2 Response of BY-2 Cells to Methyl Jasmonate Application of MJ substantially promotes natural bioproduct metabolism in a variety of plant species (Goossens et al. 2003; Oksman-Caldentey and Inze 2004; Oksman-Caldentey and Saito 2005; Salzman et al. 2005; Suzuki et al. 2005). Specifically, in tobacco BY-2 cells, the MJ treatment resulted in accumulation of nicotine-type alkaloids and promotion of the phenylproanoid pathway (Hakkinen and Oksman-Caldentey 2004; Galis et al. 2006). Furthermore, the initially yellow cells developed a bright orange color which is associated with the strong increase in intracellular, vacuolar-localized fluorescence (Fig. 5B).

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Fig. 5. Morphology and mortality of MJ-treated BY-2 cells. A Bright-field microscopy image of BY-2 cells in normal (2,4-D), hormone-free (HF) or HF plus 20 µM MJ (HF/MJ) medium. Arrows mark dead cells. B Autofluorescence of the same cells under fluorescence microscopy (λex 520−550 nm, λem > 580 nm). C Time course of cell death rate in normal (2,4-D), HF or HF/MJ medium. Values are the means of three replicates of the experiment ± SE

This fluorescence, observable under microscope filters for both fluorescein isothiocyanate (FITC) and rhodamine, most probably originates from some of the newly accumulated secondary metabolites that have yet to be characterized. It has also been reported that MJ caused arrest of the cell cycle at both G1/S and G2/M transition points in BY-2 cells (Swiatek et al. 2002, 2004). Furthermore, the cells cultivated with MJ show reduced expansion, which normally occurs in auxin-deficient (HF) medium (Fig. 5A). In addition, cell death rate in HF medium was significantly reduced by MJ (Fig. 5C), suggesting that MJ has an unspecified protective role in auxin-starved cells. It can be speculated that the accumulation of secondary metabolites, particularly phenolics, provide oxidative stress protection to the cells due to their well-known radical scavenging properties (reviewed by Blokhina et al. 2003). Furthermore, as some of the simple, exogenously applied, phenolic acids (cinnamic and ferulic acid) have strong inhibitory effects on germination and subsequent growth of the tobacco seedlings (I. Galis, unpublished data), accumulation of phenolics may provide an alternative explanation for the lack of cell division after application of MJ. In the following text, we summarize the effects of 20 µM of MJ on gene expression in BY-2 cells using the 16K cDNA microarray described above. Compared to other plant microarrays, the BY-2 16K array is enriched for MJ and other plant hormone transcripts, making it a more suitable tool for hormone-related studies. 3.3 MJ-Regulated Genes In order to examine the MJ-responsive genes, the expression profiles of BY-2 cells cultivated in HF (control) and HF/MJ (treated) media were compared. The

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Fig. 6. MJ treatment causes gradual change in global gene expression in BY-2 cells compared to cells cultivated in the medium without hormones. Seven-day-old stationary cells were washed in hormone-free (HF) medium and treated with MJ. Global gene expression at respectively 3, 6, 24 and 48 h time points after addition of 20 µM MJ was plotted against gene expression data from HF cultivated cells at corresponding time points

use of HF medium shows an advantage over the standard cultivation medium at the point when the cells at the 48 h time point are about to be compared. If one uses the standard, 2,4-D-containing medium for MJ treatments, the cells with MJ would display cell cycle arrest, while the control cells would extensively divide, especially at the later time points (24 and 48 h). In contrast, the cells cultivated in HF medium are kept at mostly non-dividing status in both treatments, resulting in the more comparable morphological conditions and thus the gene expression profile backgrounds of the cells. In general, MJ had an expectedly profound effect on gene expression of BY-2 cells. The number of genes that showed differential expression compared to HF-condition-grown cells gradually increased with time after addition of MJ, having a cascade-like effect on gene expression (Fig. 6). In summary, we found 828 genes that showed an increased level of expression in MJ medium to at least 2.5-fold above that of the HF control at any point used in the analysis (3, 6, 24, or 48 h). Similarly, 938 genes were down-regulated by at least 2.5-fold in the presence of MJ. Compared to AFLP data (Goossens et al. 2003), the down-regulated genes were especially overrepresented in our experiment as they found 376 up-regulated and only 83 individual repressed MJ-AFLP tags over the 48 h period (with approximately 20,000 transcript tags analyzed). Although the data are not treated for redundancy, it may appear that it is more difficult to isolate the repressed transcripts using the AFLP method in comparison to cDNA microarrays. In support of this, sorghum cDNA microarray (Salzman et al. 2005) data, available for individual plant tissues, show the following numbers of genes: (i) 1,316 and 918 genes were at least 2.5-times up-regulated at 27 h in the shoot and root, respectively; and (ii) 597 and 565 genes were at least 2.5-times repressed at 27 h in the shoot and root, respectively. Based on these three independent gene profiling experiments it can be estimated that, at genome-wide level, the expression of at least several hundred to thousand genes may be altered by exogenous application of MJ in plants. It is important, though, to distinguish between genes that are directly responsive to MJ and genes that respond in adaptation to the changing endogenous environment in the cells. To this end, detailed

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time course microarray data can be used to identify the direct and indirect effects of MJ in a global perspective. 3.4 Temporal Pattern Analysis of MJ-Induced Genes In our experiment, six individual groups of MJ-inducible genes with a distinctive expression profile during the time course of the experiment (−1 h = stationary, 0 h = washed, 3 h, 6 h, 24 h, 48 h) could be established using the Self-Organizing Map Clustering function. For this analysis, we used a distinct subset of 439 genes (Fig. 7) that showed absolute two-fold or higher increase of expression upon MJ treatment in comparison to the cells at the beginning of the experiment (i. e. cells washed in HF medium just before addition of MJ). Such analysis did not include MJ up-regulated genes with a nearly constant or declining expression level but still less repressed in comparison to cells in HF medium. Groups 1–3 contained genes with different amplitudes of constitutive induction over the cultivation period of the experiment (3−48 h), group 4 contained early but transiently expressed genes (3−24 h), group 5 included genes with a late response to MJ (24−48 h), and most of the genes in group 6 had less regular patterns of weaker induction over the later period of the experiment (24−48 h). In this analysis, groups 1–4 mostly represent primary response genes, while groups 5 and 6 contain late(r)-responding transcripts. In BY-2 cells, two major metabolic groups are generally induced by MJ: (i) nicotine-related compounds in alkaloid biosynthesis and (ii) phenylpropanoid accumulation which originates in the shikimate pathway and is driven by phenylalanine ammonia-lyase (PAL) activity. Both types of natural bioproducts accumulate in MJ-treated tobacco BY-2 cells at early stages and over the period of at least one week (Hakkinen et al. 2004; Galis et al. 2006). It is reasonable that the expression of the genes that are involved in their biosynthesis follows this pattern. In support, groups 1, 2 and 3 of early and constitutively up-regulated genes contained a number of genes that can be directly associated with phenylpropanoid and alkaloid metabolism. For example, several of the genes found in the backbone phenylpropanoid metabolism, including PAL-B (BP133607), C4H (AB236952) and 4CL-2 (U50846), were present in group 1, contributing to the early and constitutive enhancement of phenylpropanoid biosynthesis (Fig. 8B). Interestingly, the induction of phenylpropanoid metabolism by MJ can be tracked to the very upstream level of 2-dehydro-3-deoxyphosphoheptonate aldolase (DAHP synthase) in the shikimate pathway (Fig. 8B). This enzyme catalyzes the first condensation reaction in the shikimate pathway to produce cyclohexane skeleton (Strack 1997). Previously shown elicitor and woundrelated expression of DAHP synthase (Keith et al. 1991; Henstrand et al. 1992; Jones et al. 1995; Suzuki et al. 1995) and the early response (3 h) of the DAHP synthase gene on BY-2 microarray both point to the direct regulation of this gene by MJ. At this point, however, the possibility that this induction results from the positive metabolic drive in the cells cannot be completely excluded.

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Fig. 7. Classification of MJ-induced genes. The group of 439 MJ positively induced genes was classified by the Self-Organizing Map Clustering method into six clusters (Groups 1–6) on the basis of their expression profiles during the time course of the experiment. Details concerning individual MJ-induced groups of genes are given in Galis et al. (2006). The average for gene expression in each group is displayed on the right (note scale difference in group 1–3 graphs). Squares HF medium; triangles HF/MJ medium

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The time course data also provided a potentially new viewpoint on the role of two isoforms of ornithine decarboxylase (ODC) in tobacco phenylpropanoid and alkaloid biosyntheses, respectively. The ornithine decarboxylase (D89984, NtODC-2; Xu et al. 2004) from expression group 1 was co-lineated with PAL, C4H and especially with CCoA-MT expression (Fig. 8C). This suggests that these genes may be under common transcriptional control, leading to the synthesis of hydroxycinnamoyl-putrescine conjugates that preferentially accumulate in the MJ-treated BY-2 cells (Galis et al. 2006). In contrast, the expression of the dominant nicotine-related biosynthesis gene putrescine N-methyltransferase (PMT; BP532193) closely correlated with expression of another ODC-like gene (BP526041) in group 3. This might suggest that this ODC gene is controlled by a common transcription factor that is involved in alkaloid biosynthesis in tobacco (Fig. 8C). Interestingly, this ODC gene encodes a rather novel protein dissimilar to database-reported tobacco ODC proteins, although, in the EST search, several sequences with close similarity to the tobacco BP526041 can be found in Solanaceae species, including tomato (AI487426), potato (CV505729) and Capsicum (CA513990). Group 3 of MJ-induced genes, with overall the strongest amplitude of induction, also included the key JA biosynthetic gene, allene oxide synthase (BP131753, BP534611 and BP535068). The self-amplification ability of the JA signal in plant cells is discussed later when we characterize genes based on their functional assignment. The detoxification-related ESTs encoding glutathione S-transferases (7/76) were present in group 4 of early/transiently induced genes. In addition, genes involved in lipid metabolism, represented by the genes for lipase (BP529892), 12-oxophytodienoate reductase (BP531249) and omega-3 fatty acid desaturase (BP526848), belonged to this category, pointing at the putative function of these genes in transient release of lipids from the membrane and de novo synthesis of JA (Creelman and Mullet 1997). Group 5 (late/strong induction) included genes that may not be the primary targets of MJ as their expression only increased at 24 h or later after MJ application. Interestingly, four P450 homologues of unknown function belong to this group. This suggests that at least some of the genes in group 5 may be good candidates for metabolic enzymes involved in advanced/late phenolic metabolism (flavonoids, lignin) that are substrate-driven by gradually increasing amounts of PAL-derived phenolic substrates. In support of this, several other phenylpropanoid genes, such as those encoding caffeic acid 3-O-methyltransferases (BP531923 and BP532243), cinnamoyl alcohol dehydrogenase (X62343), or suberization and lignin-forming anionic peroxidase (BP128865, BP530376 and BP534423) homologues, seem to be similarly controlled. Specific attention should also be given to group 6 of MJ-induced genes (late/moderately induced). This group contained an extremely large proportion of genes without homology to known genes (105/156) and a significant number of transposon-related sequences among the remaining clones (12/52). The activation of transpositional events by MJ is most probably behind this atypical pattern in group 6. From the microarray data, we conclude that jasmonates

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J Fig. 8. Microarray-based reconstruction of metabolic flows in three main MJ-induced pathways

in tobacco BY-2 cells. Representative expression profiles for each gene found on the array are displayed in the graphs. Squares HF medium; triangles HF/MJ medium. A Self-induction loop of jasmonic acid (JA) biosynthesis: Aa BP529892; Ab BP527325; Ac BP535068; Ad BP131753; Ae BP532751; Af BP531249; Ag acyl-CoA oxidase BP535048. B Shikimate/phenylpropanoid pathway: Ba phospho-2-dehydro-3-deoxyheptonate aldolase 1 BP530031; Bb chorismate mutase BP533842; Bc phenylalanine ammonia-lyase BP133607; Bd cinnamate 4-hydroxylase; Be p-coumarate 3-hydroxylase BP525801; Bf caffeic acid 3-Omethyltransferase BP532243; Bg caffeoyl-CoA O-methyltransferase-like protein BP133524; Bh 4coumarate:coenzyme A ligase NTU50846; Bi BP532586; Bj X62343; Bk BP128865; Bl BP529067; Bm AF311783. C Nicotine alkaloid and polyamine biosynthesis: Ca ornithine decarboxylase D89984; Cb ornithine decarboxylase BP526041; Cc putrescine methyl transferase BP532193; Cd quinolinate phosphoribosyltransferase BP532561

may be one of the major contributors to the somaclonal variability observed in tissue cultures. In support of this, specific mobile element Tto1 from tobacco has been shown to be controlled by MJ at the transcriptional level (Takeda et al. 1998, 1999). The evolutionary role of jasmonates in metabolic diversification of plants in response to pathogenesis should also be considered on the basis of our data. 3.5 Functional Classification of MJ-Regulated Genes in BY-2 Cells In the previous text we mostly concentrated on the kinetic fashion and mutual relationship of MJ-responsive transcripts using the time course data. Now we will examine the MJ-transcriptome from three main functional viewpoints: (i) the previously mentioned ability to self-amplify the initial MJ/stress signal and interaction with other hormones, (ii) defense-related gene expression and (iii) phytoalexin accumulation, with special focus on phenylpropanoids. It is known that activation of JA synthesis is a natural part of the JA positive feedback loop (Goossens et al. 2003; Kubigsteltig and Weiler 2003). On the BY-2 microarray, we observed strong induction of the transcripts involved in JA synthesis after exogenous addition of 20 µM MJ (Fig. 8A). This induction was especially evident for the transcripts encoding allene oxide synthase, a ratelimiting step in JA synthesis (Fig. 8A). Simultaneously, other components of the JA-biosynthetic pathway, including allene oxide cyclase (BP532751) and 12-oxophytodienoate reductase (BP531249), together with several other genes putatively involved in JA synthesis (lipases, lipoxygenases and β-oxidase) were all up-regulated by MJ (Fig. 8A). De novo production of another stress hormone ethylene is also triggered during wound stress or with exogenous application of MJ (Kim et al. 1998; Hudgins and Franceschi 2004). From the 16K BY-2 microarray, homologues of ACC-oxidase (BP131676 and BP529287) were induced by MJ treatment, together with two ethylene-responsive element binding proteins (BP130171 and BP532067). This observation further confirms the previously reported cooperative action of jasmonates and ethylene in stress response (De

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Bruxelles and Roberts 2001; Kunkel and Brooks 2002; Devoto and Turner 2005). Another new putative hormone–hormone interaction could be deduced from the microarray data: a close homologue of IAA-Ala hydrolase (IAR3; BP532160 and BP533378) was induced by MJ with a strong group 1 profile, resulting in a possible release of free (active) IAA from its conjugated (inactive) form in the presence of MJ. Defense/pathogen-related transcripts comprise one of the major functional groups of MJ-regulated genes. We have already mentioned the transiently induced GSTs found in group 4; these stress-inducible proteins play an important role in catalyzing the conjugation of xenobiotics with glutathione, which leads to their detoxification. In addition to GSTs, the list of defense-related transcripts regulated by MJ in BY-2 cells further included cysteine and serine proteases, proteinase inhibitors, elicitor-induced genes, PR proteins, ABC transporters and other genes reportedly involved in plant defense. The third important function of jasmonates in defense response comprises accumulation of phytoalexins in the cells. In tobacco, two main classes of low molecular weight metabolites that contribute to the stress-induced phytoalexins, phenylpropanoids and alkaloids have already been mentioned. More specifically, treatment of BY-2 cells with MJ induced accumulation of several nicotine-related alkaloids (Hakkinen and Oksman-Caldentey 2004) and the application of 20 µM MJ to hormone-free medium-cultivated BY-2 cells resulted in increased contents of soluble, phenylpropanoid pathway-derived hydroxycinnamoyl-putrescine conjugates in the cells (Keinanen et al. 2001; Galis et al. 2006). The strong induction of both these pathways by MJ application in BY-2 cells, which was generously covered by genes found on the 16K BY-2 microarray, is summarized in Fig. 8B,C. These and other end-products may provide direct protection to the plants due to their insecticidal and antimicrobial properties, but also due to the antioxidant character of many phenylpropanoid derivatives, including coumarins and flavonoids. While the responsiveness of these pathways to stress is well known, the actual signaling pathways that coordinate the expression of genes remain largely unknown (Endt et al. 2002; Oksman-Caldentey and Inze 2004). Here, microarray analysis can provide useful clues to the MJ-signal transduction elements. 3.6 Crossover of Temporal Expression Patterns and Functional Classification: Identification of Regulatory Elements Large-scale microarray technology together with time course data allow efficient targeted identification of regulatory elements (transcription factors) that control gene expression in response to environmental signals. The strategy that we use is based on BY-2 microarray analysis of the closely co-regulated putative regulators and target metabolic genes in BY-2 cells upon elicitation with MJ. Using this strategy, we could identify the NtMYBJS1 (for jasmonate-specific MYB) gene that was closely co-expressed with the phenylpropanoid pathway

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Fig. 9. Expression of MJ-regulated NtMYBJS1 gene and related transcriptional targets. NtMYBJS1 gene was induced early by MJ and the transcript levels remain elevated, though at least 72 h after treatment (verified by RT-PCR analysis). Similarly, phenylpropanoid genes were induced within 3 h after MJ application, suggesting that the NtMYBJS1 gene may directly regulate these genes via its promoter DNA-binding transcription factor activity. The protein/DNA interactions, ectopic expression and metabolic composition of the transgenic cells are described in detail in Galis et al. (2006)

genes, PAL and 4CL (Fig. 9). Plant MYB proteins have been shown to regulate diverse developmental processes (e. g. Stracke et al. 2001), but the most established role of plant MYB proteins is in the control of the phenylpropanoid biosynthetic pathway (reviewed by Jin and Martin 1999; Stracke et al. 2001, Endt et al. 2002). Further analysis revealed that this MJ-controlled MYB protein, when ectopically expressed in BY-2 cells, results in increased amounts of phenylpropanoid conjugates, namely caffeoyl- and feruloylputrescine (CP and FP, respectively) in BY-2 cells. Particularly, CP is a typical product in MJ-treated BY-2 cells (Galis et al. 2006). Second round microarray analysis, involving differential comparison of the NtMYBJS1 transgenic lines and appropriate controls, further helped us to pinpoint the prospective transcriptional targets of the NtMYBJS1, tobacco PAL A and 4CL2 genes. In order to confirm the microarray data, the direct interaction of the recombinant MYB protein with promoter of PAL A and PAL B genes has been demonstrated by electromobility shift assay (Galis et al. 2006). Importantly, this gene also showed MJ- and wound-dependent expression in planta (our unpublished data), suggesting that the exclusive BY-2 data can be successfully interpolated to the whole plant level. Using an identical approach and herein described 16K BY-2 microarray, a glucan elicitor-inducible MYB protein was identified recently (Shinya et al., in prep.). It is interesting to note that although both of the proteins NtMYBJS1 and NtMYBGR1 induce the same phenylpropanoid genes in BY-2 cells, they are not cross-induced by MJ and elic-

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itor, respectively. This suggests that both wounding/herbivore (JA-dependent) and glucan elicitor regulate phenylpropanoid defense-related genes through the action of differential MYB genes as stimulus-specific molecular switches. A similar strategy may be employed in the future to functionally characterize more MJ-inducible genes, including the still in-seek receptor of MJ signal and other signal transduction components. For example, it has been shown that ubiquitin-associated F-box proteins, such as COI1, are involved in jasmonate signaling (Xie et al. 1998; Xu et al. 2002; Ren et al. 2005). In our recent work (Galis et al. 2006) we present several novel MJ-inducible F-box proteins that could be involved in jasmonate-related signaling pathways. These intriguing proteins and their function will be analyzed in the near future. In conclusion, the microarray approach has provided comprehensive data on changes in gene expression in various metabolic pathways and, together with progressing metabolomic approach, will allow efficient identification of metabolic processes and their regulation. The combination of microarray and time course data appeared to be a very powerful tool when the regulatory elements were targeted in the analysis. Also, once the candidate regulator is selected, microarray analysis of the overexpressor lines provides the most efficient tool for identification of downstream genes and pathways. In the meantime, we are collecting basic data using the current system which will be published as data files in series of separate papers. This will include, for example, regulation of gene expression by other hormones, abscisic and salicylic acids, cytokinin, nitric oxide and others, thus providing broad public access to the current data.

4 Future Perspectives In this chapter we summarized the current status of our analysis on tobacco BY-2 cell transcriptome using EST and microarray analyses. However, genome sequences of several model plant species are already available and many cDNAs of complete length have already been analyzed in these species. The next challenge of tobacco and BY-2 transcriptomic work will therefore be the collection of complete length cDNA sets of enough number. Actually we are currently analyzing 5
end sequences of some complete-length clones. We anticipate being able to release this information as well as the cDNA clones to the public within a couple of years from now. The genome sequence of tobacco and the wild-type species is another challenge; however, current development of the sequencing technology of complex genome (Rogers and Venter 2005) will allow us to analyze the whole tobacco genome within the near future. Thus obtaining the genome sequence information will not be a major bottleneck in biological research. In contrast, more comprehensive transcriptomic information under different conditions is essential in order to understand how genome information is used to reg-

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ulate living organisms. In this regard, our cDNA microarray has a limitation because we need large investment to prepare target DNA for the array and this causes the restriction in supply of microarray to the scientific community. Currently, uniform oligonucleotide-based microarrays/gene chips have been generated in several model organisms including model plants. These current microarrays/gene chips are made with the synthesis of DNA on the surface of chips; thus there is no limitation in supply of the array. Future systemic development of the microarray system of tobacco BY-2 cells will involve the development of oligo array using the current unigene information of tobacco (http://www.tigr.org/tigr-scripts/tgi/T index.cgi?species=tobacco) in combination with the sequence information of full-length cDNAs.

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VI.3 Proteomics of Tobacco Bright Yellow-2 (BY-2) Cell Culture Plastids M.A. Siddique, W. Gruissem, and S. Baginsky1

1 Introduction 1.1 Plastid Proteomics: An Active Research Field Plastids distinguish plant cells from cells of other eukaryotes. Depending on cell and tissue type, plastids develop and differentiate from progenitor plastids into different plastid types that are responsible for essential biosynthetic and metabolic activities. Among these activities are photosynthetic carbon fixation and the synthesis of fatty acids, pigments, starch and amino acids. Plastids are classified into different categories according to their structure (morphology), pigment composition (color), and other developmental aspects. Proplastids, which are found in meristematic cells of roots and shoots, are the progenitors of all other plastid types. Amyloplasts are mainly found in root cells, and etioplasts develop in dark-grown photosynthetic tissues where they can rapidly differentiate into photosynthetically active chloroplasts after illumination (Neuhaus and Emes 2000). The most fundamental distinction between plastid types is based on their primary energy metabolism, i. e. heterotrophy versus autotrophy. Plastids are excellent candidates for organelle proteomics because of the physiological processes they support and the basic research interests they have spawned for many years. Another important aspect is the evolutionary origin of plastids. It is now accepted that plastids originated from cyanobacteria through endosymbiosis (Goksøyr 1967). During co-evolution with the plant cell, most plastid genes were transferred to the nucleus. Plastids became semi-autonomous organelles that depend on nuclear-encoded proteins that are translated in the cytoplasm and subsequently imported into the plastid. Proteomics generates novel information on nuclear-encoded proteins that are transported into the plastid and thus provides insights into all plastid metabolic functions. Additionally, there is a substantial biotechnological interest in plastids as a production site for pharmaceuticals and other compounds. Plastids fix nitrogen and synthesize amino acids, and they are active in sulfur metabolism and isoprenoid biosynthesis (for review see Staehelin and Newcomb 2000). Full knowledge of all plastid functions will therefore facilitate biotechnology approaches (reviewed in Gewolb 2002). 1 Institute

of Plant Sciences and Functional Genomics Center Zurich, Swiss Federal Institute of Technology, ETH Zurich, 8092 Zurich, Switzerland, e-mail: [email protected]

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Despite the enormous interest in plastid biology, our current understanding of the metabolic functions and capacities of different plastid types is limited. Proteomics, the systematic analysis of all proteins from different plastid types, is one promising way towards a better understanding of plastid biology. Several proteomics studies have been conducted with chloroplasts in recent years and have provided valuable information about their metabolic capacities (Ferro et al. 2002; Peltier et al. 2002; Schubert et al. 2002; Ferro et al. 2003; Froehlich et al. 2003; Friso et al. 2004; Kleffmann et al. 2004). It has become clear, however, that chloroplast proteome analyses have reached saturation since the detection of new proteins is hampered by highly abundant photosynthetic proteins that dominate the proteome of fully developed, photosynthetically active chloroplasts. A valid strategy to circumvent this constraint and to increase proteome coverage is the use of different plastid types for high-throughput protein identification. Heterotrophic plastids, for example, do not contain highly abundant photosynthetic proteins and therefore allow the detection of other metabolic activities and regulatory factors. At the same time, the proteomes of different plastid types contain valuable information about plastid type-specific functions. Most of the studies reported to date have been conducted with fully functional chloroplasts and only three proteomics approaches have used heterotrophic plastid types. These include rice etioplasts (von Zychlinski et al. 2005), wheat amyloplasts (Andon et al. 2002) and BY-2 cell culture plastids (Baginsky et al. 2004). A comparison of the proteome data from the different plastid types confirms that the proteomes of heterotrophic and autotrophic plastids differ considerably, which is especially apparent with regard to their distinct energy metabolism. Heterotrophic plastids import metabolites such as ATP and glucose 6-phosphate for essential biosynthetic activities, for example the synthesis of starch from ADP glucose, fatty acids from acetate and amino acids from inorganic nitrogen (Weber et al. 2005). These pathways place a high demand for energy and reducing equivalents on the heterotrophic plastid. Substantial evidence has accumulated that heterotrophic plastids generate reducing equivalents by the oxidative branch of the pentose phosphate pathway that is initiated by glucose 6-phosphate. Glucose 6-phosphate as well as ATP are imported from the cytosol by well-characterized plastidic glucose 6-phosphate/phosphate translocator and ATP/ADP transporters (Weber et al. 2005). 1.2 Tobacco Bright Yellow -2 (BY-2) Cell Culture Plastids The BY-2 cell culture is a well-defined and commonly used model system for studies of cell cycle regulation and the structure of the cytoskeleton (Geelen and Inze 2001). BY-2 cells are non-green, rapidly growing cells. These cells can multiply up to 100 times within a week under conventional cell culture conditions (Nagata et al. 1992). Thus, BY-2 cells can be easily grown in large quantity in the laboratory.

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BY-2 cell culture plastids have many properties in common with undifferentiated proplastids. These include the following:

• The nucleoids of BY-2 plastids are similar to those of proplastids (Kirk and Tilney-Bassett 1978; Sakai 2001; Phillips et al. 2002). • BY-2 plastids reveal a DNA synthesis pattern that is similar to that of proliferating plant cells (reviewed in Sakai et al. 2004). • The plastid RNA polymerase from BY-2 plastids predominately transcribes plastid-encoded genes from NEP-promoter elements (Kapoor and Sugiura 1999), which is characteristic of undifferentiated plastids. • BY-2 plastids have retained the ability to develop and differentiate in a hormone-dependent manner (Miyazawa et al. 1999). These properties taken together suggest that the plastids from tobacco BY-2 cells represent an interesting undifferentiated plastid type with several features that characterize true proplastids. Therefore we decided to analyze the proteome of BY-2 plastids in order to obtain initial insight into the protein complement and metabolic capacities of undifferentiated plastids. In this chapter we report our proteomics strategy, the protein fractionation and identification from isolated BY-2 plastids, together with a detailed functional assignment of the identified proteins. We discuss potential metabolic networks that are active in the BY-2 plastid and compare our data with another heterotrophic plastid type, the etioplast. 1.3 Proteomics Strategies: Definition and Concepts Proteomics defines an approach for the systematic analysis of all proteins expressed in a cell. The progress of proteomics and proteomics-related technologies over the last decade is mainly based on two parallel developments. First, the wealth of genome information that has become available has paved the way for the large-scale analysis of proteins, for which amino acid sequences were deposited into databases (e. g. the Arabidopsis Genome Initiative 2000). Second, technological improvements in mass spectrometry, especially the development of soft ionization techniques for peptide analysis, have allowed the high-throughput identification of proteins from small amounts of samples (reviewed in Pandey and Mann 2000; Aebersold and Mann 2003). Although it is accepted that the proteome is dynamic and difficult to define, scientists aim at the most complete analysis of the protein complement of a cell or a tissue type under certain, well-defined conditions. In principle, two basic proteomics approaches can be distinguished. “Protein profiling” attempts the identification of all proteins that are present in a sample and results in a list of proteins. Combined with sophisticated protein or peptide fractionation strategies, protein profiling is a technically relatively simple approach for high-throughput analyses of the proteome of an organelle or a cell type and provides a snapshot of the major protein constituents (Washburn et al.

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2001). “Functional proteomics” concentrates on the identification of specific proteins related to specific biological processes. Although the identification concentrates on those proteins that are altered upon a stimulus or signal, the original analysis takes place at the level of the complete proteome. This approach often involves two-dimensional (2-D) gel electrophoresis. Proteins that differ in abundance are present or absent from one of the differently treated samples and are specifically analyzed and identified. In addition to 2-D PAGE, such changes can be revealed by other differential display techniques such as isotope-coded affinity tagging (ICAT) (Gygi et al. 1999, 2000) or isobaric tagging for relative and absolute quantitation (iTRAQ). Another example of functional proteomics is the identification of proteins isolated by affinity separation methods such as antibody affinity precipitation, native purification of protein complexes, or affinity ligand binding. All these approaches are to some extent designed to solve a specific biological question and the experimental design is based on a hypothesis. In the present study we used protein profiling for the analysis of the BY-2 plastid proteome (Fig. 1). Here, we employed a common shotgun proteomics approach with isolated organelles. So far, we have identified 168 proteins from BY-2 plastid preparations (Baginsky et al. 2004).

2 The Proteome of BY-2 Cell Culture Plastids 2.1 Protein Detection from Different Fractions With the isolated BY-2 plastids, we devised a multidimensional protein fractionation strategy to increase the dynamic range of our proteomics analysis (Figs. 1 and 2). Proteins were first separated by their different solubility, a procedure that is referred to as “serial extraction”. During this procedure, proteins are solubilized from membranes, with buffers of increasing solubilization capacity providing information of membrane association for every identified protein. Here, the proteins were fractionated into soluble proteins (Osmo), peripheral (8 M urea) and two integral membrane protein fractions, CHAPS and 5% SDS. The soluble proteins make up 17% of the total BY-2 plastid proteins (Fig. 2). This value indicates that the isolated BY-2 plastids were largely intact since the soluble proteins would have been lost from defective plastids. Peripheral membrane proteins (8 M Urea) make up 47% of the total BY-2 plastid proteins (Fig. 2). The integral membrane protein fraction contains 36% of all BY-2 plastid proteins. Almost all integral membrane proteins were already solubilized with the buffer containing 7 M urea, 2 M thiourea, 1% Brij 35 and 2% CHAPS. No additional proteins were solubilized with 5% SDS (Figs. 1 and 2). Figure 3 shows the distribution of protein identifications from the different serial extraction steps and provides a few examples for proteins that were identified from the osmotic shock (O), the urea (U) and the CHAPS (C) fraction.

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Fig. 1. Strategy for the BY-2 plastid proteome analysis. Starting from a serial extraction of proteins based on their different solubility (stained gel) we devised a multidimensional protein fractionation scheme (O Osmo, U urea, C CHAPS, S SDS). Proteins in the Osmo and urea fractions were further fractionated by ion exchange chromatography (IEX) or Blue Sepharose to subtract highly abundant purine nucleotide-binding proteins. All protein fractions were further subjected to SDS-PAGE, the gel was cut into ten pieces and each piece was subjected to in-gel tryptic digest. Peptide mixtures were further fractionated by reversed phase chromatography on C18 columns that were coupled online to an LCQ ion trap mass spectrometer. Mass spectrometric data were searched against the Viridiplantae protein database which was downloaded as a subsection of the National Center of Biotechnology Information (NCBI) non-redundant protein database (DB) (1.). Since large databases increase false positive discovery rates, we individually analyzed each peptide for protein inference (2.). ESI Electrospray ionization

Most proteins were identified from the osmotic shock and the urea fraction (52 and 55, respectively) and a considerably large number (38) from both the CHAPS and the urea fraction. This may be attributed to the fact that urea is unable to completely solubilize some proteins that require higher stringency for their solubilization. Notably, only seven proteins were identified in the Osmo and the urea fractions, suggesting that the separation of proteins into soluble and peripheral membrane proteins is an efficient strategy. Most of the proteins that were identified in the soluble fraction are active in amino acid and carbohydrate metabolism. The 8 M urea and CHAPS fractions contain envelope membrane proteins and some potential transporters, suggesting their function at the plastid outer membrane system. All data on the BY-2 plastid proteome were assembled in a proteome database for different plastid types which is freely available (see www.plprot.ethz.ch). For every plastid type, its interaction with the cytosol and the import of cytosolic compounds are of pivotal importance for their metabolism and func-

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Fig. 2. A Quantitative distribution of BY-2 plastid proteins among the different fractions of the serial extraction procedure. Values were determined by Bradford analyses. B Example of the subsequent multidimensional protein fractionation of the proteins contained in the Osmo fraction. Each fraction was analyzed by SDS-PAGE and subsequent silver staining. IEX 50–500 and 2000 indicate the concentration of KCl that was used for the elution of the bound proteins. Values are 50 mM, 100 mM, 200 mM, 500 mM and 2 M. The flowthrough fractions derived from the Blue Sepharose (BS-FT) and the ion exchange (IEX-FT) chromatography were also analyzed. Proteins bound to Blue Sepharose were eluted in a single step

Fig. 3. Number of proteins identified from each fraction and individual examples. O Osmo; U urea; C CHAPS

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tional maintenance. We identified a set of envelope transporters that import metabolites to support the plastid heterotrophic metabolism (Fig. 4). We identified the ATP/ADP transporter, the oxoglutarate/malate translocator and the outer envelope protein (OEP) 75 homologue. The ATP/ADP transporter and the oxoglutarate/malate translocator are involved in the metabolite exchange between the plastid and the cytosol (Neuhaus and Wagner 2000). The ATP/ADP transporter imports cytosolic ATP to feed energy into the heterotrophic plastid metabolism (reviewed in Neuhaus and Emes 2000). Oxoglutarate is the precursor of ammonia assimilation, and recent reports have suggested that the oxoglutarate/malate translocator imports carbon skeletons for net glutamate synthesis (Weber and Flugge 2002). In addition to the above mentioned transporters, we have identified the glucose 6-phosphate/phosphate translocator. The possible role of this translocator is the import of glucose 6-phosphate to initiate the oxidative pentose phosphate pathway which provides reducing equivalents for a diverse number of plastid metabolic activities (Schnarrenberger et al. 1995). The glucose 6-phosphate/phosphate translocator was not detected in autotrophic chloroplasts (Kleffmann et al. 2004). Our data are summarized in Fig. 4 which shows a comprehensive sketch of the identified proteins from characteristic heterotrophic metabolic pathways. Taken as a whole, most of the BY-2 plastid proteins have a function in the cellular metabolism, especially in carbohydrate and amino acid metabolism. We have identified glutamine synthetase, glutamate synthase and glutamate dehydrogenase, suggesting an active primary nitrogen fixation pathway (Fig. 4). From here, the reduced nitrogen can enter into several other amino acid synthesizing pathways. Several proteins from the arginine, the branched-chain amino acid and the aromatic amino acid biosynthesis were identified which utilize the reduced nitrogen. It appears that the BY-2 plastids provide the rapidly growing and dividing cell with the amino acids required for protein biosynthesis and other cellular functions. 2.2 Comparison Between BY-2 Cell Culture Plastids and Etioplast Since a shotgun proteomics approach as reported here is biased towards abundant proteins (Baginsky et al. 2004), it provides some limited quantitative information and allows a plastid type comparison for prevalent and dominant metabolic pathways. On the basis of BLAST searches, we identified 83 proteins that are common to etioplasts and BY-2 plastids, two heterotrophic plastid types (Fig. 5). The majority of these proteins were also detected in the chloroplast proteome, suggesting that they represent a housekeeping “core proteome” that is common to most, if not all, plastid types. The majority of all proteins in this “core proteome” have a function in carbohydrate and amino acid metabolism, two essential functions of plastids that are important for the plant cell. A number of proteins characteristic of BY-2 plastids were identified as well. Interestingly, protein products from the two large plastid encoded open

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Fig. 4. Schematic illustration of metabolic pathways in heterotrophic plastids. The number of identified peptides for each identified protein is given in parentheses ( first number) together with the staining intensity on a 2D-PAGE-based BY-2 plastid protein map (second number). DH Dehydrogenase; E-4-P erythrose-4-phosphate; G-6-P glucose 6-phosphate; GOGAT glutamate synthase; MEX maltose exporter; MIP major intrinsic protein; OPPP oxidative pentose phosphate pathway; PDC pyruvate dehydrogenase complex; PEP phosphoenol pyruvate; R-5-P ribulose5-phosphate

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Fig. 5. Comparison of proteins identified from BY-2 plastids and etioplasts on the basis of BLAST searches. An e-value of e-20 was used as a cut-off. The number in the intersection represents those proteins that were found in both plastid types

Fig. 6. Characteristic plastid transporters that were identified from BY-2 plastids and etioplasts (ET). Those that are common to the two plastid types are shown in grey; those that are plastid-type specific are shown in white

reading frames ycf1 and ycf2 were only identified in BY-2 plastids (Baginsky et al. 2004). Provided that BY-2 plastids represent an undifferentiated plastid type, this suggests that YCF1 and YCF2 perform important functions early in plastid development and differentiation. Their exact function is currently unknown, but recent data suggest that they are indispensable for plastid development (Drescher et al. 2000). All proteomics data from the different plastid types including the BY-2 plastid proteome are available in the plprot database which is accessible at www.plprot.ethz.ch (Kleffmann et al. 2006). Since heterotrophic plastids depend to a significant extent on the import of cytosolic compounds for their energy metabolism, the plastid envelope membrane transporters are of particular importance for plastid metabolism. To investigate whether differences exist between heterotrophic BY-2 plastids and etioplasts, we have depicted all identified transporters from these two plastid types in Fig. 6. Etioplast envelope membranes contain high amounts of a phosphoenolpyruvate/phosphate translocator (PPT) and a triosephosphate/phosphate translocator (TPT) whereas no hexose-phosphate transporter was identified (Fig. 6). In contrast, we did not identify PPT or TPT from BY-2 plastids but did find large amounts of a glucose 6-phosphate/phosphate translocator. This suggests that BY-2 plastids preferentially import hexose-phosphates while etioplasts rather import triose-phosphates for their energy metabolism. Our results therefore support the view that the heterotrophic metabolism of

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BY-2 plastids and etioplasts differs. This could be a consequence of the longterm adaptation of BY-2 plastids to the cell culture conditions and the external supply of glucose. Glucose feeding was shown to decrease the expression of TPT in tobacco seedlings (Knight and Gray 1995), supporting the view that the expression of plastid transporters is fine-tuned in response to prevailing metabolic conditions.

3 Concluding Remarks Our results show that the chosen strategy for the analysis of BY-2 plastid from tobacco bright yellow cell culture is feasible, although the tobacco genome is not yet completely sequenced. The proteomics community is currently witnessing an enormous effort towards the development of software tools that allow the database-independent identification of peptides, exclusively on the basis of mass spectrometric data. As more of these tools become available, the analysis of proteins from organisms that are not fully sequenced will become easier. At present, the BY-2 plastid proteome as reported here provides useful insights into the proteome of this plastid type and, in comparison with other proteome analyses, also prevalent metabolic activities.

4 Protocols for BY-2 Plastid Isolation and Protein Fractionation 4.1 Isolation of BY-2 Plastids and Purity Assessment Tobacco (Nicotiana tabacum) BY-2 cells were grown in modified Murashige and Skoog medium as previously described (Fan and Sugiura 1995) and cells were collected after 80−86 h (∼250 g fresh weight). Fresh cells were washed two times with 0.4 M mannitol, pH 5.0 and digested in three volumes of enzyme solution (1% Onozuka RS cellulose and 0.1% pectolyase Y-23 containing 0.4 M mannitol, pH 5.6) at 30 ◦ C for 1.5 h. Protoplasts were subsequently harvested by centrifugation at 1,500 × g at 4 ◦ C for 5 min. Three washing steps followed, using five volumes of ice-cold 0.4 M mannitol, pH 5.0. The pellet was resuspended in plastid isolation buffer [0.4 M mannitol, 20 mM Tris-HCL, pH 7.6, 0.5 mM EDTA, 1.2 mM spermidine, 7 mM 2-mercapoethanol, 0.6 (w/v) polyvinylpyrrolidone (PVP) and 0.1% (w/v) BSA] and protoplasts were broken by passing the suspension several times through layers of a 40 µm mesh under high pressure. The broken protoplasts were centrifuged at 1,500× g for 10 min at 4 ◦ C to pellet the cell debris and nuclei. The supernatant was then filtered through two layers of 20 µm mesh under low pressure. Finally Percoll was added to the filtrate to a final concentration of 15% and the whole suspension was centrifuged at 15,000 × g for 20 min at 4 ◦ C. The pellet was subsequently

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resuspended in plastid isolation buffer and loaded onto a sucrose density gradient (30–50–70% sucrose in 20 mM Tris-HCL, pH 7.6, 0.5 mM EDTA, 1.2 mM spermidine, 7 mM 2-mercapoethanol) for further purification from mitochondria, nuclei and cellular debris. A yellow band of BY-2 plastids was collected at the 50–70% sucrose interface and washed two times with plastid isolation buffer without PVP and BSA. After washing, the BY-2 plastids were loaded onto a linear sucrose gradient from 30–70% sucrose for further purification. Pure proplastids were washed two times with plastid isolation buffer and spun down by centrifugation before the pellet was stored at −80 ◦ C for further studies. The purity of the BY-2 plastid preparation was assessed by enzymatic activity assays for characteristic marker proteins of other cell organelles (fumarase and catalase measurements), combined with antibody detection of plastid marker proteins (data not shown). After the first sucrose density gradient centrifugation, both fumarase and catalase activities showed one major peak at the interphase between 30 and 50% sucrose, while the plastid marker protein was enriched at the interphase between 50 and 70% sucrose (band B). This suggests that peroxisomes and mitochondria can be efficiently separated from BY-2 plastids by sucrose density gradient centrifugation (Baginsky et al. 2004). For further plastid purification band B was collected, washed with plastid isolation buffer and loaded onto a linear sucrose density gradient ranging from 30–70% sucrose (data not shown). Plastids after the second density gradient were recovered, washed and used for further protein fractionation and proteome research. 4.2 BY-2 Plastid Protein Fractionation We first separated proteins on the basis of their different solubilities, a procedure that is also referred to as serial extraction (Fig. 1). Here, the proteins were fractionated into soluble proteins (Osmo), peripheral (8 M urea) and two integral membrane protein fractions, CHAPS and 5% SDS. We determined the total amount of proteins in each fraction by Bradford analyses (Bradford 1976). Soluble and peripheral membrane proteins (from the fractions Osmo and 8 M urea) were further fractionated by liquid chromatography using ion exchange chromatography (IEX) on MonoQ (Bio Rad, Hercules, USA) or affinity chromatography on Cibacron Blue Sepharose 3 GA (Sigma, Buchs, Switzerland) in order to subtract highly abundant purine nucleotide binding proteins (see example for the Osmo fraction in Fig. 2B). Proteins bound to MonoQ were eluted in five steps ranging from 50 mM to 2 M KCl in a buffer containing 20 mM HEPES/KOH, pH 7.9, 8 M urea, 100 mM NaCl, 5 mM MgCl2 , 10 mM DTT and 2 × protease inhibitor. Proteins bound to Blue Sepharose were separated from the flow-through by a single step elution in the above buffer containing 1.5 M KCl. Proteins from each fraction were further separated by their molecular mass by SDS-PAGE. Integral membrane proteins (2% CHAPS and 5% SDS) were directly subjected to high resolution SDS-PAGE. The SDS gels were cut

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into ten pieces and the proteins in each gel slice were immediately subjected to in-gel tryptic digest. Tryptic digest was performed according to Shevchenko and colleagues (Shevchenko et al. 1996) using sequencing grade modified trypsin (Promega, USA) in a ratio of one part trypsin to ten parts protein to be digested. Proteins were digested overnight at 26 ◦ C. After digestion and peptide elution from the gel slices peptides were lyophilized to dryness and stored at −80 ◦ C until further analysis. 4.3 Mass Spectrometric Protein Identification Prior to mass spectrometry, tryptic peptides were separated by reverse phaseliquid chromatography (RP-LC) on C18 material (Fig. 1). Subsequently, they were analyzed by electrospray ionization mass spectrometry (LC-ESI-MS/MS) with an ion trap (LCQDecaXP, Finnigan, San Jose). MS/MS data were analyzed with the SEQUEST software using the Viridiplantae subsection of the National Center of Biotechnology Information (NCBI) non-redundant protein database (http://www.ncbi.nlm.nih.gov/). We manually interpreted each SEQUEST output by using the following criteria:

• A cross-correlation score (Xcorr) above 2.5. • An ion coverage (ratio between detected and expected y- or b-ions) of more than 40%. • Only fragment spectra from doubly charged parent ions were considered. • Long peptides with more than 50 theoretical fragment ions (b and y ions) were not taken into account. • Grouping of at least four peptides with an Xcorr value of at least 2.5 to the same protein was rated as significant protein identification when at least one of them exhibited a ∆CN value (normalized difference in correlation score, giving the difference between the front ranking hit and the subsequent hits) higher than 0.1. • All MS/MS spectra that were not grouped as described above but matched the other criteria were visually examined for a correct peak assignment and evaluated using the following criteria: a gapless assignment of a series of y- or b-ions to peaks of high intensity; a y-ion with an N-terminal proline corresponding to a high peak; a neutral loss of 18 Da (loss of water) from yor b-ions carrying the amino acids serine or threonine. All proteins that were identified on the basis of the above criteria were assembled into a database in FASTA format and a BLAST search (Altschul et al. 1997) was performed against itself to eliminate redundant protein entries, i. e. identical proteins from different organisms (all entries giving an E-value of zero were deleted). Acknowledgements. We would like to thank the Functional Genomics Center Zurich for technical help and maintenance of infrastructure. M.A.S. is currently supported by generous support from

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the VELUX and EMDO foundations. Work in the authors’ laboratory is supported by the ETH and funds from the SEP initiative of the ETH Zurich to W.G. and S.B.

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Section VII Technical Advances

VII.1 Cryopreservation of Tobacco BY-2 Suspension Cell Cultures Using an Encapsulation – Simple Prefreezing Method T. Kobayashi1 , T. Niino2 , and M. Kobayashi1

1 Introduction Cultured plant cells, including the tobacco BY-2 suspension cell line, are important genetic resources used extensively in cytological and molecular biological studies. The maintenance of cell cultures by successive subculture is cumbersome, however, and entails the risk of losing the cell line due to disease, contamination, somaclonal variation, or technical errors. Cryopreservation of cell cultures in liquid nitrogen can overcome these problems and provides a reliable and cost-effective method for the long-term storage of plant genetic resources (Engelmann 2000; Sakai et al. 2002). Since the first report of cryopreservation of plant cells in liquid nitrogen (Quatrano 1968), additional cryogenic techniques have extended the applicability to a wide array of plant materials from various species, including undifferentiated cell suspensions (Joshi and Teng 2000), embryogenic cultures (Sakai et al. 1990), apical tips (Takagi et al. 1997), hairy roots (Yoshimatsu et al. 1996), and somatic embryos (Fang et al. 2004). Despite these recent advances in cryopreservation techniques, cryostorage of cell suspension cultures is not yet practical for laboratory use, mainly due to the need for a programmable freezer and complex procedures. In addition, undifferentiated suspension cells are generally difficult to cryopreserve due to their sensitivity to environmental stresses. Such undifferentiated cells, which consist of large vacuolated cells, are more prone to severe freezing injury compared with embryogenic cultures and apical organs, which contain small, cytoplasmic-rich meristematic cells (Wang et al. 2002). We recently established a simple and effective cryopreservation protocol that is suitable for routine laboratory use and that does not require specialized equipment (Kobayashi et al. 2005). In this chapter, we describe the methods for cryopreservation of cultured plant cells and the optimum conditions for cryopreservation of BY-2 cells. 1 BioResource Center, RIKEN Tsukuba Institute, Koyadai 3-1-1, Tsukuba, Ibaraki 305-0074, Japan,

e-mail: [email protected] National Institute of Agrobiological Sciences, Kannondai 2-1-2, Tsukuba, Ibaraki 305-8602, Japan

2 Genebank,

Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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2 Methods for Cryopreservation of Cultured Plant Cells Successful cryopreservation must prevent the formation of intracellular ice crystals during cooling and recrystallization during warming, which destroy cell structures and cause lethal damage to cells (Sakai et al. 2002). Plant cells, which contain large amounts of cellular water, are prone to severe freezing injury when exposed to liquid nitrogen. The avoidance of crystallization of cellular water is related to vitrification, the phase transition of an aqueous solution from a liquid to a noncrystalline solid (glass) at the glass transition temperature (Fahy et al. 1984). A viscous solution easily supercools to subzero temperatures and becomes amorphous glass (that is, it is vitrified) at around −110 ◦ C without the formation of ice crystals. A vitrified material can be preserved safely at the temperature of liquid nitrogen (−196 ◦ C) for an indefinite length of time, because the glass is stable below the glass transition temperature. Methods for cryopreservation of plant cells require two critical steps: dehydration and rapid cooling. Cell dehydration involves reducing cellular water and thus condensing the cytoplasm. The dehydrated cells are then rapidly cooled by being plunged directly into liquid nitrogen, which results in complete vitrification of the cell contents. Three methods are used for cryopreservation of plant cell and organ cultures: slow prefreezing, vitrification, and air dehydration. These methods mainly differ in the dehydration procedure. The slow prefreezing method is based on freeze-induced dehydration, which occurs during cooling at a slow, controlled rate (0.5 to 2 ◦ Cmin−1 ) to a definite temperature (−30 to − 80 ◦ C) with a programmable freezer in the presence of cryoprotectant solution (Schrijnemakers and Van Iren 1995). Slow cooling causes the development of ice crystals in the extracellular space and thus condenses the cryoprotectant solution, resulting in osmotic dehydration of cells. In this method, the contents of the optimally dehydrated cells are converted into glass after being plunged into liquid nitrogen, while the extracellular space is filled with crystalline ice. The vitrification method uses highly concentrated cryoprotectant solutions, ranging from 5 to 8 M, by which cells are osmotically dehydrated at a nonfreezing temperature (Sakai et al. 1990). Both cell contents and the surrounding cryoprotectant solution are then vitrified by immersion in liquid nitrogen. This complete vitrification method simplifies the cryogenic process and has extended the applicability of cryopreservation to a broad range of plant genetic resources (Sakai et al. 2002). Air dehydration is a convenient method that does not require a programmable freezer or cryoprotectants (Bachiri et al. 2000). Plant materials are precultured in sucrose-enriched medium for several days, then dried by silica gel or airflow in a flow cabinet to optimal water content, and finally cryopreserved in liquid nitrogen. Cryopreservation procedures for most plant materials require preculture and cryoprotectant treatment in advance of the severe dehydration step. Precul-

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turing cells in medium enriched with sugars or sugar alcohols for several days enhances their capacity to tolerate environmental stresses, such as dehydration, high osmotic pressures, and low temperatures, by inducing an accumulation of osmoprotective compounds in the cells (Bachiri et al. 2000). Cryoprotectant solutions consist of various combinations of cryoprotective chemicals, such as dimethylsulfoxide, glycerol, ethylene glycol, propylene glycol, and sugars. The cryoprotectant pretreatment induces stress tolerance and facilitates vitrification of plant materials by moderately dehydrating the cells, reducing intracellular ice crystallization, and allowing stable supercooling (Reinhoud et al. 2000; Menges and Murray 2004). Furthermore, cryoprotectants stabilize cellular membranes and proteins during dehydration and cryopreservation (Turner et al. 2001).

3 Conditions for Cryopreservation of Cell Suspension Cultures Cryopreservation of cell suspension cultures ultimately requires the regrowth of cell cultures that are equivalent to the original cell lines. The viability rate of cryopreserved suspension cells must be high to ensure the rapid regrowth of cell suspension cultures, as proliferation of suspension cells strictly depends on the initial density of cells in the culture and is markedly suppressed in cultures with low cell density (Stuart and Street 1969). However, cell suspension lines differ in their capacity to survive cryopreservation, so each cell line must be cryopreserved by the most appropriate procedure under optimized conditions. Maintaining high cell viability during cryopreservation can minimize the risk of morphological, biochemical, and physiological changes of cell characteristics caused by genetic variations or undesirable selection of particular types of cells. Many reports have indicated that cell cultures regrown after cryopreservation retain the same characteristics as the original cell lines based on DNA polymorphism (Hao et al. 2002; Liu et al. 2004), gene expression (Ryynänen et al. 2002), and biochemical, cytological, and physiological properties (Yoshimatsu et al. 1996; Urbanová et al. 2002; Menges and Murray 2004). Our knowledge of the influence of cryopreservation on plant cells is still limited. However, we know that cultured suspension cells frequently suffer from cumulative genetic or epigenetic changes during long-term maintenance by repeated subculture. Most cell suspension lines have not been evaluated genetically by molecular analyses during the sequential subculturing. Further genome-wide expression studies are necessary to assess whether cultured cells retain their characteristics during maintenance by either cryopreservation or successive subculture.

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4 The Encapsulation–Simple Prefreezing Method for Cryopreservation of BY-2 BY-2 cells have been cryopreserved successfully using an encapsulation technique combined with a simplified slow prefreezing method that requires no specialized equipment (Fig. 1). The encapsulation, which involves immobilizing the cells in hydrophilic alginate gel, has little effect on the viability and proliferation of cells (Bachiri et al. 2000) and allows replacement of the medium without centrifugation and easy handling of a large number of the materials in a short time (Hirai and Sakai 2003). Furthermore, the alginate gel protects the embedded cells from mechanical damage due to manipulation, direct exposure to cryoprotectant solution, and, by modulating the diffusion of the cryoprotectants, severe changes in osmotic pressure. The freeze-induced dehydration of cells in the slow prefreezing step can be achieved with simple cooling in a laboratory freezer rather than with controlled-rate cooling in a programmable freezer (Sakai et al. 1991; Menges and Murray 2004). The cryogenic vials are simply placed in a freezer at −30 ◦ C; the cells are cooled to −30 ◦ C at a certain rate (approximately −2 ◦ C min−1 ; Sakai et al. 1991) in the presence of cryoprotectant solution, and then held for an adequate time (1 to 1.5 h) at −30 ◦ C. The cryopreservation efficiency of BY-2 cells primarily depends on the physiological condition and growth phase of the starting materials. Many studies have demonstrated that cells at the exponential growth phase are preferable for cryopreservation (Bachiri et al. 2000; Menges and Murray 2004), because they have many characteristics that help to reduce freezing injury, such as small size, small vacuoles, dense cytoplasm, and low water content, as compared to cells at the lag or stationary phases. Pretreatment of BY-2 cells with cryoprotectant solution containing 2 M glycerol and 0.4 M sucrose for 45−60 min significantly promotes tolerance to cooling to −30 ◦ C and subsequent exposure to liquid nitrogen (Kobayashi et al. 2005). In contrast, prolonged preculture of BY-2 cells in medium containing 0.3 M sucrose, mannitol, or sorbitol for 1 day results in a marked reduction in cell viability. Therefore, short-term treatment with the cryoprotectant solution appears to be appropriate for the cryoprotection of BY-2 cells. The regrowth procedure is crucial for the recovery of cell growth capacity and the efficient reestablishment of cell suspension lines. To prevent the recrystallization of cellular water, cryopreserved BY-2 cells should be warmed rapidly by transferring cryogenic vials from liquid nitrogen to a water bath at 40 ◦ C (Engelmann 2000). Stepwise dilution of the cryoprotectant solution with medium containing a high concentration of sucrose can mitigate rehydration injury caused by a rapid reduction in osmotic pressure. Because immediate dispersion of the cells into a liquid culture results in loss of cell viability and arrest of cell division, intact beads formed from the encapsulated cells are cultured in liquid medium with gentle shaking for several days. BY-2 cells

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Fig. 1. The cryopreservation procedure for tobacco BY-2 suspension cells by the encapsulation– simple prefreezing method. mLS Linsmair and Skoog medium

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fully resume their capacity for growth within 1 day and rapidly proliferate in the beads (Kobayashi et al. 2005). Then the embedded cells are released into the culture medium by crushing the beads. The BY-2 suspension cell line can be regrown within 1 week. The cells of this cryopreserved line did not show obvious morphological differences or changes in the ability to grow compared with cells of an untreated control line (Kobayashi et al. 2005).

5 Conclusion The encapsulation–simple prefreezing method appears to be a good technique for reliable and routine cryopreservation of the BY-2 suspension cell line. The use of this technique may help to secure against the loss of BY-2 cell lines and enable maintenance of large numbers of transformed BY-2 derivatives for postgenomic studies. Additional studies are necessary to examine the effect of long-term cryopreservation in liquid nitrogen on the genetic stability of cultured cells and the applicability of this method to a wide range of plant cell lines.

6 Protocol 6.1 Materials

• Basal medium: modified Linsmair and Skoog (mLS) medium containing 30 g l−1 (0.09 M) sucrose and 0.2 mg l−1 2,4-dichlorophenoxyacetic acid. Adjust pH of media to 5.8 and autoclave for 20 min at 120 ◦ C • Encapsulation solution: mLS medium containing 2% (w/v) sodium alginate (300–400 cP; Wako Pure Chemical Industries, Osaka, Japan). Dissolve sodium alginate by stirring under gentle heating (see Note 1 below) • Gelation medium: mLS medium containing 0.1 M calcium chloride • Cryoprotectant solution: mLS medium containing 2 M glycerol and 0.4 M sucrose • Dilution media: mLS media containing 0.5 and 1.2 M sucrose, respectively • Cryovials (round bottom, 2.0 ml) • Cryovial rack (do not use a rack that covers the bottom of the vials) • Conical plastic tubes (15 and 50 ml) • Flasks (200 ml) • Pipettes • Pasteur pipettes • Forceps • Spatulas • Twelve-well microplates

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Low-speed centrifuge Rotary shaker Water bath Liquid nitrogen in a Dewar flask

6.2 Methods 6.2.1 Encapsulation and Cryopreservation Procedure 1. Transfer BY-2 cell suspension on day 3 of subculture to 15-ml conical tubes (see Note 2) and centrifuge the tubes at 100 × g for 5 min. 2. Remove the supernatant and gently resuspend the pelleted cells in four volumes of encapsulation solution. 3. Drip the mixture of cells and encapsulation solution into 50 ml of gelation medium in a 200-ml flask with a Pasteur pipette, which causes the gelation of the alginate and formation of beads approximately 4 mm in diameter. Keep the beads formed from the encapsulated cells in the gelation medium with gentle shaking on a rotary shaker for 5 min. 4. Remove the medium with a pipette, wash the beads with basal mLS medium, and then incubate the beads in 50 ml of basal mLS medium with gentle shaking for 10 min. 5. Replace the medium with cryoprotectant solution. Suspend the beads in at least 1 ml of the cryoprotectant solution per bead and incubate with gentle shaking for 45−60 min at 25 ◦ C. 6. Using forceps, transfer three beads into each cryovial with 300 µl of cryoprotectant solution (see Note 3). 7. Place the cryovials in a rack and store them in a laboratory freezer at −30 ◦ C for 2 h (see Note 3). 8. After removing the cryovials from the freezer, immediately immerse the cryovials in liquid nitrogen. 6.2.2 Regrowth Procedure 1. Warm each cryovial in a water bath at 40 ◦ C with gentle agitation. After thawing, immediately remove the cryovials from the bath. 2. Transfer the three beads to 20 ml of 1.2 M dilution medium in a 50-ml conical tube. Set the tube horizontally on a shaker and incubate the beads with gentle shaking for 15 min at 25 ◦ C. 3. Replace the medium with 0.5 M dilution medium and incubate for 15 min. 4. Replace the medium with basal mLS medium and incubate for 15 min. 5. Suspend three beads in 3 ml of fresh mLS medium in each well of a 12-well microplate and culture the beads with shaking (130 rpm) in the dark for 3 days at 27 ◦ C (see Note 4).

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6. Gently crush the beads with a spatula to release the embedded cells into the culture medium. Culture the cell suspension for an additional 4 days. 7. Transfer the cell suspension to 95 ml of fresh basal mLS medium in a 300-ml flask (see Note 5). 6.3 Notes 1. Sodium alginate is usually dissolved in calcium-free medium. We were able to dissolve sodium alginate in common mLS medium, because the calcium chloride concentration of the medium (3 mM) does not induce gelation of alginate. 2. Good physiological condition of BY-2 cells is essential for successful cryopreservation. The cell suspension at the exponential growth phase should consist of small cytoplasmic-rich cells and retain a high viability rate. Measure the cell growth in each laboratory and use each cell suspension at the exponential growth phase. 3. The freezing step is crucial for this cryopreservation protocol. To achieve cooling of the BY-2 cells to −30 ◦ C at a slow certain rate, keep the volume of the cryoprotectant solution plus beads in each cryovial around 400 µl. Make sure there is enough space in the freezer so the vials are not too close to each other. Do not cover the vials. 4. The beads must be cultured until the embedded BY-2 cells proliferate enough to induce cell suspension culture. Therefore, the duration of culture depends on the viability rate of the cryopreserved cells. When the cell viability is below 50%, the cell proliferation may take more than 3 days. 5. This protocol achieved 60–70% cell viability on day 1 of culture (three independent experiments) and 100% regrowth efficiency of cell suspension culture within 7 days (seven independent experiments with a total of 33 vials; Kobayashi et al. 2005).

References Bachiri Y, Bajon C, Sauvanet A, Gazeau C, Morisset C (2000) Effect of osmotic stress on tolerance of air-drying and cryopreservation of Arabidopsis thaliana suspension cells. Protoplasma 214:227–243 Engelmann F (2000) Importance of cryopreservation for the conservation of plant genetic resources. In: Engelmann F, Takagi H (eds) Cryopreservation of tropical plant germplasm. JIRCAS Agricultural Series no. 8. Japan International Research Center for Agricultural Sciences, Tsukuba, pp 8–20 Fahy GM, MacFarlane DR, Angell CA, Meryman HT (1984) Vitrification as an approach to cryopreservation. Cryobiology 21:407–426 Fang JY, Wetten A, Hadley P (2004) Cryopreservation of cocoa (Theobroma cacao L.) somatic embryos for long-term germplasm storage. Plant Sci 166:669–675

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Hao YJ, You CX, Deng XX (2002) Analysis of ploidy and the patterns of amplified fragment length polymorphism and methylation sensitive amplified polymorphism in strawberry plants recovered from cryopreservation. Cryo-Lett 23:37–46 Hirai D, Sakai A (2003) Simplified cryopreservation of sweet potato (Ipomoea batatas [L.] Lam.) by optimizing conditions for osmoprotection. Plant Cell Rep 21:961–966 Joshi A, Teng WL (2000) Cryopreservation of Panax ginseng cells. Plant Cell Rep 19:971–977 Kobayashi T, Niino T, Kobayashi M (2005) Simple cryopreservation protocol with an encapsulation technique for tobacco BY-2 suspension cell cultures. Plant Biotech 22:105–112 Liu Y, Wang X, Liu L (2004) Analysis of genetic variation in surviving apple shoots following cryopreservation by vitrification. Plant Sci 166:677–685 Menges M, Murray JAH (2004) Cryopreservation of transformed and wild-type Arabidopsis and tobacco cell suspension cultures. Plant J 37:635–644 Quatrano RS (1968) Freeze-preservation of cultured flax cells utilizing dimethylsulfoxide. Plant Physiol 43:2057–2061 Reinhoud P, Van Iren F, Kijne JW (2000) Cryopreservation of undifferentiated plant cells. In: Engelmann F, Takagi H (eds) Cryopreservation of tropical plant germplasm. JIRCAS Agricultural Series no. 8. Japan International Research Center for Agricultural Sciences, Tsukuba, pp 91–102 Ryynänen L, Sillanpää M, Kontunen-Soppela S, Tiimonen H, Kangasjärvi J, Vapaavuori E, Häggmen H (2002) Preservation of transgenic silver birch (Betula pendula Roth) lines by means of cryopreservation. Mol Breed 10:143–152 Sakai A, Kobayashi S, Oiyama I (1990) Cryopreservation of nucellar cells of navel orange (Citrus cinensis Osb. var. brasiliensis Tanaka) by vitrification. Plant Cell Rep 9:30–33 Sakai A, Kobayashi S, Oiyama I (1991) Cryopreservation of nucellar cells of navel orange (Citrus cinensis Osb.) by simple freezing method. Plant Sci 74:243–248 Sakai A, Matsumoto T, Hirai D, Charoensub R (2002) Survival of tropical apices cooled to −196 ◦ C by vitrification: development of a potential cryogenic protocol of tropical plants by vitrification. In: Li PH, Palva ET (eds) Plant cold hardness: gene regulation and genetic engineering. Kluwer/Plenum, New York, pp 109–119 Schrijnemakers EWM, Van Iren F (1995) A two-step or equilibrium freezing procedure for the cryopreservation of plant cell suspensions. In: Day JG, McLellan MR (eds) Cryopreservation and freeze-drying protocols. Methods in molecular biology, vol 38. Humana Press, Totowa, New Jersey, pp 103–111 Stuart R, Street HE (1969) Studies on the growth in culture of plant cells. IV. The initiation of division in suspensions of stationary phase cells of Acer pseudoplatanus D. J Exp Bot 20:556–571 Takagi H, Thinh NT, Islam OM, Senboku T, Sakai A (1997) Cryopreservation of in vitro-grown shoot tips of taro (Colocasia esculenta (L.) Schott) by vitrification. 1. Investigation of basic conditions of the vitrification procedures. Plant Cell Rep 16:594–599 Turner S, Senaratna T, Touchell D, Bunn E, Dixon K, Tan B (2001) Stereochemical arrangement of hydroxyl groups in sugar and polyalcohol molecules as an important factor in effective cryopreservation. Plant Sci 160:489–497 Urbanová M, Èellárová E, Kimáková K (2002) Chromosome number stability and mitotic activity of cryopreserved Hypericum perforatum L. meristems. Plant Cell Rep 20:1082–1086 Wang Q, Gafny R, Sahar N, Sela I, Mawassi M, Tanne E, Perl A (2002) Cryopreservation of grapevine (Vitis vinifera L.) embryogenic cell suspensions by encapsulation-dehydration and subsequent plant regeneration. Plant Sci 162:551–558 Yoshimatsu K, Yamaguchi H, Shimomura K (1996) Traits of Panax ginseng hairy roots after cold storage and cryopreservation. Plant Cell Rep 15:555–560

VII.2 High Throughput Microinjection Technology for the Single-Cell Analysis of BY-2 in Vivo H. Matsuoka1,2 , Y. Yamada2,3 , K. Matsuoka4 , and M. Saito1,2

1 Microinjection in the Post-Genome Era Since half a century ago, microinjection has been performed in experimental cell biology, but until recently its most frequent application has been the introduction of isolated nuclei into large cells such as egg cells 100−200 µm in diameter. Microinjection into smaller cells has been performed only by experts in research laboratories involved in specific single-cell studies (Erwee et al. 1985; Wolf et al. 1989; Saito and Matsuoka 1996; Zhao et al. 2001; Newmark et al. 2003; Saito et al. 2003; Tran et al. 2003; Yokota et al. 2003; King 2004). If the cell size is less than 50 µm, its difficulty increases extraordinarily (Fig. 1). Therefore microinjection has been recognized as a time consuming and laborintensive method. In today’s post-genome era, an intense need for high throughput microinjection has arisen. Based on cDNA information, various protein molecules such as gene expression products are now available and the structural analysis of these molecules is ongoing in various research subjects. The functional analysis of respective proteins in living cells is regarded as the next subject to be tackled. To carry this out, microinjection is necessary to introduce these molecules directly into living cells.

2 Single-Cell Manipulation Supporting Robot (SMSR) 2.1 The Concept of SMSR The most difficult step in microinjection is the insertion of a micropipette into a target cell. This requires careful and delicate manipulation of a micromanipulator. Further difficulty exists in many associated actions such as 1 Department of Biotechnology and Life Science, Tokyo University of Agriculture and Technology,

Koganei, Tokyo 184-8588, Japan, e-mail: [email protected] 2 CREST (Core Research for Evolutional Science and Technology), Japan Science and Technology

Agency, Honcho 4-1-8, Kawaguchi, Saitama 332-0012, Japan 3 Chuo Precision Industrial Co. Ltd., Oikawa Building, Kanda Awaji-cho 1-5, Chiyoda-ku, Tokyo

101-0063, Japan 4 Plant Science Center, RIKEN (Institute of Physical and Chemical Research), 1-7-22 Suehirocho,

Tsurumi-ku, Yokohama 230-0045, Japan Biotechnology in Agriculture and Forestry, Vol. 58 Tobacco BY-2 Cells: From Cellular Dynamics to Omics (ed. by T. Nagata, K. Matsuoka, and D. Inzé) © Springer-Verlag Berlin Heidelberg 2006

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Fig. 1. Cell size and microinjection difficulty. ES Embryonic stem

XY-stage manipulation, adjustment of microscope focus, the selection of a cell, transportation of the cell, exchanging a micropipette, and recording the microscopic image of the cell. These actions are split into two categories: actions performed by microscopic observation (on-microscope actions) and actions performed by taking the eye away from the microscope (off-microscope actions) (Fig. 2). Frequent changes between these on-microscope and off-microscope actions are practically very time-consuming and laborious. To perform these actions smoothly, we have developed a single-cell manipulation supporting robot (SMSR) (Matsuoka et al. 2005). The concept of SMSR is to allow an operator to concentrate only on the microinjection by avoiding perturbations caused by other associated actions. To realize this concept, a pair of XYZmicromanipulators and an XY-automatic stage were attached to a microscope and their drive controllers were centralized on a pair of joystick handles (Fig. 3). 2.2 XY-Address Registration We developed an XY-coordinate chip which was attached on the bottom of each dish (Fig. 4). By referring to these coordinates, the XY-address of each adhesive cell was registered. As for non-adhesive cells, a multimicrowell was devised in which one cell was stored per well. The XY-address of the center of each well was registered. There are two modes of registration for adhesive cells. One is direct discriminating registration. An operator checks each cell by microscopic observation during automatic scanning. The operator can register only cells with special morphological properties. The other mode is the batch processing registration. This mode utilizes a commercially available imaging system. Once the XY-addresses of respective cells are registered, microinjection can be performed at high speed, because each cell can be transported quickly to the center of the microscope view one by one on clicking the switch.

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Fig. 2. Consecutive actions involved in microinjection into many cells. The actions are divided into on-microscope and off-microscope actions

2.3 Multifunctional Micropipette Several types of micropipette are used for microinjection (Fig. 5). A singlechannel pipette is most frequently used for the introduction into cells as well as for cell support, while a two- or three-channel micropipette is used for electrophoretic introduction, electric or electrochemical measurement, and a combination of these (Matsuoka and Saito 2000; Saito and Matsuoka 2000). The tip diameter of a pipette varies according to the purpose of the pipette. For introduction into small cells with a diameter of 50 µm or smaller, the tip

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Fig. 3. A centralized control system for XYZ-micromanipulators and an XY-stage

Fig. 4. XY-coordinate chip for culture dish. This chip is applicable to any type of culture dish

diameter needs to be smaller than 1 µm. This is an empirical criterion based on our own experiences. Usually the introduction is performed by pressure application. Optimum pressure is determined by the tip diameter and the intracellular osmotic pressure of the target cell. Ionic compounds can be introduced also by electrophoresis.

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Fig. 5. Various types of capillaries used for microinjection, electrophoretic introduction, micro electrochemical measurement, and micro electric stimulation

3 Microinjection of a Dominant-Negative Protein into BY-2 Cells 3.1 Evaluation of Membrane Permeability at the Cell–Cell Junction BY-2 cells have a unique configuration of a linear chain of several cells. This is useful for the analysis of membrane permeability at the cell–cell junction. In order to assess its permeability, the electric impedance of the intercellular membrane (ZINT ) was measured and compared with that of the cell membrane contacting directly with external medium (ZEXT ) (Sotoyama et al. 1998). ZINT was unstable and varied from 4 to 800 MΩ, while ZEXT was between 1.7 and 2.3 GΩ (Fig. 6). This suggests that the intercellular membrane is more permeable to ions. This does not necessarily support the presence of the open passage of plasmodesmata. In fact, an anionic fluorescent dye, Lucifer yellow, did not diffuse to neighboring cells. The intensity of an electrical signal applied to the above-mentioned measurement was no higher than 5 mV. It is necessary to investigate whether such a small electrical signal would modify the membrane permeability. The dependence of the cross-membrane potential (VCMP ) on cell membrane permeability was investigated by Matsuoka et al. (1999). When VCMP was smaller than

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Fig. 6. Electrical impedance of BY-2 cell membrane. A, C Phase contrast photographs; B, D fluorescent photographs, with Lucifer yellow introduced as a diffusion marker; A, B measurement of the cell membrane (cell a) contacting with external medium; C, D measurement of intercellular cell membrane (between b and c cells)

250 mV, membrane permeability was not influenced by the electrical signal application. Further increase in VCMP made the cell membrane permeable. This is owing to the terminal cell membrane. Although we have not yet investigated the intercellular membrane, we suspect that it is not permeable to Lucifer yellow and consequently to larger protein molecules without any stimulating signal. 3.2 Protocol for Consecutive Microinjection and Image Recording of Many Single Cells In BY-2 cells, prolyl hydroxylase (PH) is synthesized in the endoplasmic reticulum (ER) and transported to the Golgi body. The ER-to-Golgi transport is mediated by coat protein complex II (COPII) (Antonny and Schekman 2001). At the same time, reverse transport from the Golgi to ER occurs. Therefore the PH is localized in both the ER and Golgi in dynamic equilibrium (Yuasa et al. 2005). The formation of COPII requires a specific protein, Sar1p. If a dominant-negative form of Sar1p is introduced in the cytosol, the formation of COPII would be inhibited and the ER-to-Golgi transport will be inhibited. Consequently the Golgi-to-ER transport would predominate and PH would be preferentially localized in ER after all. In order to demonstrate such a role of Sar1, we performed a microinjection experiment using a strain of BY-2 that could express PH fused with GFP (PH-GFP). BY-2 cells were adsorbed on the bottom of a culture dish that had been treated with poly-l-lysine beforehand. The dominant-negative protein was microinjected into BY-2 single cells. As microinjection takes 20−30 s per cell, it will take 33−50 min to perform microinjection into 100 cells. Moreover,

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Fig. 7. The complicated situation whereby time to take a photograph comes before finishing microinjection into all cells. In this figure, it is necessary to take a photograph of cell 1 after the microinjection into cell n

it is necessary to take photographs of fluorescent images of each cell at 30 min, 1 h, and 2 h after microinjection. Therefore the operator has to stop the microinjection to take the photograph of the first cell (cell 1) (Fig. 7). Even in such a case, the XY-stage can instantly be driven so that cell 1 is situated at the center of microscope view. In this way, the frequent interchanges of microinjection and image recording can be performed smoothly. 3.3 Effect of a Dominant-Negative Protein on ER-to-Golgi Trafficking Before the introduction of the dominant-negative protein, there are small fluorescent dots as well as a faint ring-like perinuclear fluorescence in a BY2 cell. Dynamic movement of fluorescent dots was observed at real time. When a dominant-negative form of Sar1p was microinjected into the cell, these fluorescent small dots disappeared by 2 h (K. Matsuoka et al., unpublished result), implying inhibition of ER-to-Golgi trafficking.

4 Concluding Remarks Single-cell analysis of BY-2 that can solely be performed with microinjection has been demonstrated. This shows that microinjection is feasible as a practical method even for small cells no greater than 50 µm in diameter. It is expected that the spatio-temporal precision of microinjection will become more advanced owing to progress in nanotechnology. Then microinjection would be referred to as nanoinjection. Such an advancement of microinjection will surely support the functional analysis of many proteins of gene expression products as well as other associated bioactive compounds.

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Acknowledgements. H.M. acknowledges support from CREST of the Japan Science and Technology Agency in the research subject ‘The High Throughput Creation of Disease Model Cells and the Analysis of their Function’.

References Antonny B, Schekman R (2001) ER export: public transportation by the COPII coach. Curr Opin Cell Biol 13:438–443 Erwee MG, Goodwin PB, Van Bel AJE (1985) Cell–cell communication in the leaves of Commelina cyanea and other plants. Plant Cell Environ 8:173–178 King R (2004) Gene delivery to mammalian cells by microinjection. Methods Mol Biol 245:167–174 Matsuoka H, Saito M (2000) Microbioelectronics for single-cell experiment. Electrochemistry 68:314–320 Matsuoka H, Sotoyama H, Saito M, Oh K-B, Horikiri S (1999) Effects of a pulsing electric signal on the cross membrane potential and cell division potentiality of a single cell of tobacco. Bioelectrochem Bioenerg 49:65–72 Matsuoka H, Komazaki T, Mukai Y, Shibusawa M, Akane H, Chaki A, Uetake N, Saito M (2005) High throughput easy microinjection with a single-cell manipulation supporting robot. J Biotechnol 116:185–194 Newmark PA, Reddien PW, Cebria F, Alvarado AS (2003) Ingestion of bacterially expressed double-stranded RNA inhibits gene expression in planarians. Proc Natl Acad Sci USA100 (Suppl 1):11861–11865 Saito M, Matsuoka H (1996) Application of multifunctional microelectrode to the in vivo analysis of the K+ response to CO2 stress. Bioelectrochem Bioenerg 39:115–118 Saito M, Matsuoka H (2000) pH–microelectrode–micropipette system for the measurement of enzyme reactions in picoliter space of a living plant cell. Anal Chim Acta 404:223–229 Saito M, Mukai Y, Komazaki T, Oh K-B, Nishizawa Y, Tomiyama M, Shibuya N, Matsuoka H (2003) Expression of rice chitinase gene triggered by the direct injection of Ca2+ . J Biotechnol 105:41–49 Sotoyama H, Saito M, Oh K-B, Nemoto Y, Matsuoka H (1998) In vivo measurement of the electrical impedance of cell membranes of tobacco cultured cells with a multifunctional microelectrode system. Bioelectrochem Bioenerg 45:83–92 Tran ND, Liu X, Yan Z, Abbote D, Jiang Q, Kmiec EB, Sigmund CD, Engelhardt JF (2003) Efficiency of chimeraplast gene targeting by direct nuclear injection using a GFP recovery assay. Mol Ther 7:248–253 Wolf S, Deom CM, Beachy RN, Lucas WJ (1989) Movement protein of tobacco mosaic virus modifies plasmodesmatal size exclusion limit. Science 246:377–379 Yokota E, Vidali L, Tominaga M, Tahara H, Orii H, Morizane Y, Hepler PK, Shimmen T (2003) Plant 115-kDa actin-filament bundling protein, P-115-ABP, is a homologue of plant villin and is widely distributed in cells. Plant Cell Physiol 44:1088–1099 Yuasa K, Toyook K, Fukuda H, Matsuoka K (2005) Membrane-anchored prolyl hydroxylase with an export signal from the endoplasmic reticulum. Plant J 41:81–94 Zhao Z, Cao Y, Li M, Meng A (2001) Double-stranded RNA injection produces nonspecific defects in zebrafish. Dev Biol 229:215–223

Subject Index

ABC transporter 102 abscisic acid 123 ACC-oxidase 305 acetosyringone 196, 197 actin 31 actin inhibitor 140 actin microfilament 140 ADP-glucose pyrophosphorylase 124 ADP-ribosylation factor-type GTPases (ARF-GEFs) 110 affinity ligands 281 AFLP 300 Agrobacterium 195–205 Agrobacterium tumefaciens 6 air dehydration 330 alc-GR System 17 alkalinisation of cytosolic pH 252 alkaloid 295, 298 – nicotine 298 alkylation 290 allene oxide synthase 303, 305 amyloplast differentiation 114 amyloplasts 119, 129 Aphidicolin 66 aphidicolin 7 Arabidopsis 108–114, 195, 198, 202, 205 Arabidopsis cell culture 10, 18 Aspergillus nidulans 4 ATP-binding cassette transporter 102 Aurora kinase 30, 57 – AtAUR1 58 – AtAUR2 58 – AtAUR3 58, 60 – Aurora A 57 – Aurora B 57, 60 – Aurora C 57 autolysosome 174, 178 autophage 160 autophagosome 168, 170, 174, 178 autophagy 167, 168

autophagy-related genes (ATGs) 168 autophagy-related proteins 170 AUX1/LAXs 110 auxin 107, 112, 114, 120 – accumulation 111–113 – 2,4-D 111, 112 – depletion 112 – 2,4-dichlorophenoxyacetic acid (2,4-D) 111, 112 – efflux carrier 113, 114 – IAA 107, 111, 112 – indole-3-acetic acid (IAA) 107, 111, 112 – influx (uptake) and efflux carriers 108 – influx and efflux 114 – influx inhibitors 111 – level 114 – NAA 111, 112 – naphthalene-1-acetic acid (NAA) 111, 112 – pleiotropic physiological effects 113 – radiolabelled 109, 110 – transport 107–112, 114 – transport inhibitor (ATIs) 108 auxin starvation 98, 104 auxin-autotrophy 97 2B-13 cell 97 benzyladenine 121 BFA 109, 111–113, 143 BFA-compartment 146 BFA-sensitive GEFs 143 BiFC 198 bimolecular fluorescence complementation 198, 200 bioinformatics 276 Bip-DsRed (ER-chaperone) 141 BLAST search 291 branching 248 – DMAPP branch 249

348

– IPP branch 249 brassinolide 124 brefeldin A (BFA) 109, 111–113, 143 Brome mosaic virus (BMV) 188 BY-2 Golgi stack 138 – classical polarity parameter 138 Ca2+ 208–212, 215, 217, 218 Ca2+ channels 210–212, 217 – AtTPC1 210 – CNGC 212 – TPC 210 – TPC1 210–212 callose 32 CAP 52 – CAP-C 52 – CAP-D2 52 – CAP-E 52 – CAP-G 52 – CAP-H 52 carbamidomethyl cysteine 290 β-carotene 246 – phytoene 246 CDKA 8 CDKs 12 cell cycle 85–87, 250, 297, 299 – cell cycle progression 261 – cytometric profile 250 – DNA synthesis 250 – G1–S control point 250 – synchronised BY-2 cells 250 cell cycle arrest 213–215 cell death 250 cell division 81, 114, 298 cell division factor (CDF) 99 cell elongation 114 cell morphology 112 cell plate 27, 79, 81, 82, 86, 88 cell polarity 114 cell suspension culture 329, 331 cell wall 79, 81, 82 cell-free system 44, 45 Chemical Complementation Assays 250 chemical fixation – electron microscopy 147 – immunofluorescence 147 chemiosmotic hypothesis 111 chemiosmotic polar diffusion model 107 chloramphenicol 127

Subject Index

chromatography 275, 277 – liquid 275, 277 chromosome 51 – chromosome condensation 51 – chromosome dynamics 51, 52 – mitotic chromosome 51 coat protein complex II 344 ConA 100 condensin 52 – condensin I 52 – condensin II 52 – condensin subunit 52 connectivity 279 – peptide 279 constitutive cycling of proteins 110 COPI antibody 139 – AtArf1 139 – AtSec21 (γ −COP) 139 COPI vesicle 135, 143 COPII protein 135 COPII vesicle 135, 142 cortical cytoplasm 113 cortical microtubules 24 covalent isoprenylation of proteins 253 – CaaX tetrapeptide motif 253 – farnesylated proteins 254 – geranylgeranylation of proteins 254 – GTP binding proteins 253 – prenyl cysteine lyase 254 – protein prenyl transferase 253 – thioether bond 253 cross-membrane potential 343 crown gall – Boston Ivy 111 cryopreservation 329, 330 cryoprotectant solution 330–332 CYCD1 8 cyclin 12 cyclin-dependent kinases (CDKs) 5 cytochrome b5 175 cytokinesis 79–82, 85–91, 296 cytokinin 98, 120 cytokinins 252 cytoskeletal structure 23 cytoskeleton 112, 198, 200 2,4-D 121, 122 D1 cyclin gene expression

5

Subject Index

databases 275, 280, 283–288 – 2DGE 287 – 2DGE reference 280 – EST 285 – protein 285 – proteome 284 – random database 286 – sequence 275, 285, 288 data-exchange 287 2DDiGE 280 dephosphorylation 88 differentiation and dedifferentiation in BY-2 cells 129 DiGE 281 digest 283 – in silico 283 division site 27 dominant-negative protein 343 DXP synthase 245 dynamin-related protein 228 – ADL1 229 – chloroplast 229 – conventional dynamin 228 – Dnm1p 228 – DRP3 228, 229 – peroxisomal division 229 E-64 170 efflux carriers 113 electric impedance 343 electrophoresis 275–277, 279 – gel 276 – two-dimensional gel 275, 277 elicitor 207, 208, 211, 212, 307 – cryptogein 207 embryogenesis 108 encapsulation 332 end-binding protein 1 28 ER 135, 136, 140, 167 – morphology 140 ER export sites (ERES) 140, 142 ER retention signal 176 – HDEL sequence 176 – K/HDEL sequence 176 – KDEL sequence 176 ER-bound Sec13 142 ER-derived transport vesicle 176 – HDEL 177 – K/HDEL signal 177

349

ERGIC 142 ER-Golgi hybrid 146 ER-Golgi sandwich 145 ER-Golgi transport 136 ER-to-Golgi transport 135 ESI 281, 282 EST database 170 ethanol-inducible gene expression system 4 expression sequence tag (EST) 170, 287, 291, 293 farnesol 254, 255 – apoptosis 255 – cell death 255 – co-localisation studies 256 – conjugation 256 – E, E-[1-3 H] farnesol 255 – farnesal 254, 256 – farnesoic acid 256 – farnesol phosphorylation 254 – farnesol toxicity 255, 258 – fluorescent analogue 256 – terpene synthases 254 farnesyl diphosphate 245, 254 – chaetomellic acid 245 – cytosolic FPP 255 – dolichol 245 – farnesyl diphosphate synthase 247, 259 – farnesylated proteins 245 – FPP synthase (FDS) 245 – FPP synthesis 255 – protein farnesyl transferase 245 farnesyl diphosphate (synthase) – mitochondrial transit peptide 247 farnesylation 285 fatty acid desaturase 303 flow cytometric 6 flow cytometry 15 fluorescent Golgi-localized marker 138 – GFP- or RFP-tagged class I α-1,2-mannosidase 138 – GFP- or RFP-tagged sialyl transferase 138 – GFP-tagged xylosyltransferase 138 – YFP-tagged GONST1 138 fluorescent tags 114 fluorochrome 280

350

FM4-64 256 formin 34 GATEWAY cloning 18 GATEWAY system 9 genetic transformation 195 genetic variation 331 genome 308 geranyl diphosphate 245 geranylgeranyl diphosphate 245 GFP 52, 112, 113, 198, 199 GFP-α-TIP 158 GFP-BP-80 158, 161 GFP-HDEL 141 GFP-tagged COPII coat protein (LeSec13) 142 gibberellins 123 glucocorticoid receptor (GR) system 5 13 C-labelled glucose isotopomers 243 glutathione S-transferase (GST) 306 glycopeptidase F 104 glycoprotein 281 glycoprotein specific dye 281 glycostaining 279 GNOM 110 Golgi apparatus 135, 137, 143, 167 Golgi belt 140 Golgi fluorescent marker 136 Golgi inheritance 140 Golgi marker 156 Golgi visualization 137 GONST1-YFP 155, 161 green fluorescent protein (GFP) 3, 52, 112, 113 – GFP-fused protein 52 GUS 197–201 habituation 103 heterochromatin 53 – Arabidopsis-like heterochromatin protein 1(AtLHP1) 56 – heterochromatic region 56 – heterochromatin formation 56 – heterochromatin protein 1 (HP1) 53 – HP1 family 56 hexoses 285 high throughput microinjection 339 histone H1 kinase 9 homology 288

Subject Index

HR 207, 212 hydroxyapatite 100 hypersensitive response 207 – cell death 207 – defense responses 207 IAA 99 IAA-Ala hydrolase 306 identification – cross-species 275 immunofluorescence labelling 148 importin 199, 200 inducible overexpression 112 intermediate (ERGIC) compartment 135 IPP isomerase 251 isoelectric focusing 281 isoelectric point 277, 286 isopentenyladenine 252 isoprenoid biosynthesis 257 – C-24 methyltransferases 258 – cytoplasmic isoprenoid biosynthesis 257 – non-sterol derivatives of MVA 261 – obtusifoliol 14-demethylase 258 – overproduction of sterols 257 – sterol homeostasis 258 – sterol intermediates 257, 258 – sterol methyltransferases 259 – sterol pathway 257 – steryl esters 258 isoprenoids 241 – campesterol 251 – diterpenes 241 – isofucosterol 251 – isoprene 241 – material for membrane biogenesis 246 – monoterpenes 241 – phytosterols 241 – plastidial pigments 241 – sesquiterpenes 241 – sitosterol 251 – stigmasterol 251 isotopic enrichment 246 K+ channels 65 K+ -transport 68 katanin 28 KDEL vesicle (KV)

176

Subject Index

351

kinesin 82, 84 – AtNACK1 85 – AtNACK2 85 – AtNACK1 88 – NACK1 84–89, 91 – NACK2 84, 85 labelling pattern 249 laser scanning cytometry lignin 303 lipase 303 lovastatin 123, 245 Lucifer yellow 344

67

M phase 82, 85–87 macroautophagy 168 MALDI 105, 281, 282, 284, 289 MALDI-TOF/TOF MS 105 MALDI-TOF/TOF-MS 101 MAP 89–91 – Ase1p 89 – Feo 89 – GEM1 90, 91 – MAP65 89, 91 – MOR1 90, 91 – PRC1 79, 89 – SPD1 89 – XMAP215 79, 90 MAP kinase cascade 79 MAP65 proteins 29 MAPK 84–86, 91, 214–217 – NRK1 85, 86, 89–91 – SIPK 215 – WIPK 215 MAPK cascade 80, 81, 84–86, 91 MAPKK 84–86 – ANQ1 86 – NQK1 85, 86, 89 MAPKKK 82, 84–86, 91 – ANP1 82 – ANP2 82 – ANP3 82 – NPK1 82, 85–89, 91 – NPK2 85 mass spectrometry 275, 285 mass spectrum 282 matrix-associated laser desorption ionization (MALDI) 281 MDR/PGPs 110

membrane permeability 343 MEP (pathway) 251, 252 – [1,1,1,4-2 H4 ]DX 251 – [2-14 C]-1-deoxy-d-xylulose 253 – alternative pathway 242 – 1-deoxy-d-xylulose 5-phosphate synthase 253 – fosmidomycin 252, 263, 265 – 1-hydroxy-2-methyl-2-(E)-butenyl 4-diphosphate 252 – ketoclomazone 263 mesophyll protoplast 111 metabolic cross-talk 250 – cytoplasmic and plastidial pathways 250 – exchange of intermediates 252 – intracellular complementation mechanism 250 metabolic differentiation 298 methionine 290 2-C-methyl-d-erythritol 4-phosphate (MEP) 242 3-methyladenine 170 methylation 56 – methyltransferase 56 methylerythritol phosphate (MEP) 246 – 1-deoxy-d-xylulos 5-phosphate 245 – DX reductoisomerase 245 – fosmidomycin 245, 251 – glyceraldehyde 3-phosphate 245 – MEP pathway 247 – MEP synthase 245 – phytoene 249 – pyruvate 245 mevalonic acid (MVA) – acetoacetyl-CoA thiolase 245, 261 – acetyl-CoA 245 – acetyl-CoA precursors 248 – citrate 248 – DMAPP 241 – HMG-CoA reductase 242, 245 – HMG-CoA synthase 245 – house-keeping HMGR 257 – 3-Hydroxy-3-methylglutaryl-coenzyme A (HMG-CoA) reductase 257 – inhibitors of MVA biosynthesis 242 – IPP 241 – IPP isomerase 245, 251, 252

352

– mevinolin 242, 245, 250, 251, 257, 263, 265 – MVA derivative 262 – MVA pathway 242, 251 – MVA pool 250 – MVA-derived precursor pool 246 – sterol biosynthesis 241, 242 – stress-induced HMGR 257 mevinolin 255 MG132 proteasome inhibitor 9 microarray 202, 293, 295, 298 microautophagy 168 microtubule 41, 42, 44 – nucleation 41–46 – organizing centers 41 microtubule dynamics 27 microtubules 79 miniprotoplast 45 mitochondrial dynamics 225, 226 – BY-2 cells 225 – in yeasts 225 mitochondrial fission 227 – DNA synthesis 227 – mitochondrial fission cycle 227 mitochondrial fission factor 230 – Fis1p 230 – FtsZ 231 – MD rings 230 – Mdv1 230 – Min 232 mitochondrial fusion 232, 233 – Fzo1 233 – in animals and yeast 232 – in plants 233 – Kaede 235 – Mgm1p 233 – Ugo1p 233 mitosis 51 – mitotic phase 51 modification 275, 276, 290 – post-translational 275, 276 molecular marker for habituation 103 MS analysis 251 – isotopomer distribution 251 MT bundling 89 multidimensional separation 276 multifunctional micropipette 341 MYB 307

Subject Index

NACK-PQR pathway 85–89, 91 1-naphthylphthalamic acid (NPA) 108, 109, 111–114 natural bioproducts 298 NMR analysis 246 – glucose isotopomers 247 – 1 H- and 13 C-NMR spectroscopy 248 – isotopic enrichment 247 NPA 108, 109, 111–114 NtTPC1 210 nuclear export signal (NES) 11 nuclear localisation signal (NLS) 11 off-microscope action 340 omics 288 on-microscope action 340 organ formation 108 ornithine decarboxylase 303 ortholog 283, 290 β-oxidation of fatty acids 261 – peroxisomal acetoacetyl-CoA thiolase 261 pentoses 285 peptide mass fingerprint 282, 284 peptide mass fingerprinting 281 peptide sequence analysis 281 peroxidase 303 P-glycoprotein 101 phenylalanine ammonia-lyase 301 phenylproanoid 298 phosphate 285 phosphoprotein 281 phosphoprotein specific dye 281 phosphorylated histone 60 phosphorylation 57, 82, 87–89, 91 phosphostaining 279 phosphotyrosine 285 phragmoplast 23, 79–82, 85, 86, 88–91 phyllotaxis 108 phytoene 246 phytohormone 295, 298 – abscisic acid 295 – auxin 296 – 6-benzyladenine 295 – brassinolide 295 – cytokinin 298 – GA3 295

Subject Index

– methyl jasmonate 295 – salicylic acid 295 phytosterols – C-24 methyltransferases 258 – campesterol 246 – cycloartenol 259 – 24-ethylidenelophenol 259 – Lab 170250F 261 – 24-methylenelophenol 259 – obtusifoliol 14-demethylase 261 – sitosterol 246 – sterol methyltransferases 259 – stigmasterol 246 phytotropins 109 PIN 110–114 PINOID 111 PIP2 113 plasma membrane (PM) 108, 109, 112, 113 plasma membrane ghost 44 plastids 119 plastoquinone 243 – all-trans-polyprenols 245 – 13 C enrichment 245 – Golgi apparatus 243 – 2-methylquinol 243 – protein-based transport system 243 PM-ATPase 113 PMF 282–284, 289, 290 post-translational modifications 281, 284 potassium 67 PR proteins 306 precipitation 277 – acetone/trichloro-acetic acid 277 pre-fractionation 279 preprophase band 23 pressure-probe 65 prevacuolar compartment (PVC) 136, 153 prolyl hydroxylase 344 protein 275 – abundance 275 – degradation 277 – interactions 275 – low abundant 279 – modifications 277 – modified 281 – orthologs 283 – profiles 275

353

– protein 285 – quantitation 277, 280 – visualization 280 protein aggregate 174, 175 protein degradation 174 protein identification 275, 282, 286 – cross-species 282 protein isoprenylation 262, 263 – [14 C]MVA-labelling of proteins 262 – CaaX isoprenylation motif 265 – chaetomellic acid 262 – CVIL geranylgeranylation motif 263 – GFP fusion proteins 265 – GTP binding proteins 262 – phytohormone signalling 262 – prenylated GFP fusion proteins 263 – protein farnesyltransferase 262 – protein prenyl transferase 265 protein localisation 11 protein purification 105 protein–protein interaction 275 proteome 275, 287 protoplast 195, 196, 198, 199 PSTAIR motif 8 PTMs 291 PVC 162, 163 Rb protein 5 reactive oxygen species (ROS) RFP 175 RNAi 3, 14 root development 108 ROS 208, 209, 214–217 – H2 O2 217 – O2 218 – cryptogein 217 RT-PCR 197 sample preparation 277 secondary metabolite – caffeic acid 303 – caffeoyl-putrescine 307 – cinnamoyl alcohol 303 – feruloylputrescine 307 secondary metabolites 295 secretory pathway 136 secretory unit model 142 shikimate pathway 301 sialyc acid 285

208

354

single-cell manipulation 339 slow prefreezing method 330, 332 SMC 52 – non-SMC subunit 52 – SMC2 52 – SMC4 52 – SMC family protein 52 – SMC subunit 52 SNARE molecule 146 spectrum 282 spermidine synthase 295 spindle 58 – mitotic spindle 58 – spindle pole 58 – spindle pole body 58 squalene epoxidase 245, 258, 259 – (3S)-2,3-oxidosqualene 259 – terbinafine 259, 260 squalene synthase 245, 258 – accumulation of squalene 260 – presqualene diphosphate 259 – SQS activity 260 – squalestatin 245, 259, 260 – squalestatin-1 258 – terbinafine 245 squalene synthase (SQS) 245 staining 280 – coomassie 280 – fluorescent 280 starvation 170, 172, 174 sterol biosynthesis 251 stop codon removal 13 sucrose starvation 170 systems biology 287 T-complex 196, 198 T-DNA 196–198, 200–202 Tobacco BY-2 cells 242 – amyloplasts 242 – high metabolic activity 242 – leucoplasts 242 – proplastids 242 tobacco cell line VBI-0 (Nicotiana tabacum L., cv. Virginia Bright Italia) 112 tobacco suspension-cultured Xanthi XHFD8 cells 111 tobacco Xanthi XHFD8 cells 112 tonoplast intrinsic protein (α-TIP) 154 transcriptome 287

Subject Index

transcriptomic information 293 transform 201 transformation 16, 195, 197–201, 205 transgenosis 112 transitional ER 135, 141 translation 183 transport of isoprenoid intermediates 246 triacylglycerol synthesis 259 – lipid particles 259 triglycerol synthesis – triglyceride-rich lipid droplets 260 tropisms 108 trypsin 104, 289 T-strand 196–199 tubulin 81 γ -tubulin 41, 42, 45, 46 turgor 65 ubiquinone 243 – 3,4-dihydroxy-5-hexaprenylbenzoate 247 – all-trans-polyprenol 254 – all-trans-polyprenols 245 – 13 C enrichment 245 – 4-hydroxybenzoate 243 – mitochondrial membrane 243 – mitochondrial respiration chain 245 – polyprenyl phosphates 243 – prenyl side chains 243 – prenyl transferase 243 – protein-based transport system 243 – solanesyl diphosphate synthase 247 – transport of IPP 243 – transport system 247 – UQ-10 243, 254, 255 – UQ-9 243 UV-spectroscopy 248 vacuolar sorting receptor (VSR) 136, 153 vacuole 167 – lytic vacuole 167 – storage vacuole 167 vascular differentiation 108 vir gene 196, 197 virulence genes 196

Subject Index

355

virulence protein 200–202 virus 183 – poliovirus 186 – Tobacco mosaic virus 183 – tobamovirus 183 – Tomato mosaic virus 183 – Turnip crinkle virus 188 vitrification 330, 331 VSR 162, 163 wortmannin

160, 164

XY-address registration 340 XY-automatic stage 340 XY-coordinate chip 340 XYZ-micromanipulators 340 yeast two-hybrid system YFP 199, 200 YFP-BP-80 155 cis-zeatin 252 trans-zeatin 123, 252

198