Advances in Botanical Research, Volume 48

Advances in

BOTANICAL RESEARCH Series Editors JEAN-CLAUDE KADER

Laboratoire Physiologie Cellulaire et Mole´culaire des Plantes, CNRS, Universite´ de Paris, Paris, France

MICHEL DELSENY

Laboratoire Ge´nome et De´veloppement des Plantes, CNRS IRD UP, Universite´ de Perpignan, Perpignan, France

CONTRIBUTORS TO VOLUME 48

ANTOINE BAUDRY Section of Cell and Developmental Biology, Division of Biological Sciences, University of California, San Diego, La Jolla, California 92093 ELS J. M. VAN DAMME Department of Molecular Biotechnology, Laboratory of Biochemistry and Glycobiology, Ghent University, 9000 Gent, Belgium FOLKERT A. HOEKSTRA Laboratory of Plant Physiology, School of Experimental Plant Sciences, Wageningen University, Wageningen NL‐6703 BD, The Netherlands YUE‐IE C. HSING Institute of Plant and Microbial Biology, Academia Sinica, Taipei 11529, Taiwan JAE‐HOON JUNG Molecular Signaling Laboratory, Department of Chemistry, Seoul National University, Seoul 151‐742, Korea STEVE KAY Section of Cell and Developmental Biology, Division of Biological Sciences, University of California, San Diego, La Jolla, California 92093 SANG‐GYU KIM Molecular Signaling Laboratory, Department of Chemistry, Seoul National University, Seoul 151‐742, Korea NAUSICAA LANNOO Department of Molecular Biotechnology, Laboratory of Biochemistry and Glycobiology, Ghent University, 9000 Gent, Belgium CHUNG‐MO PARK Molecular Signaling Laboratory, Department of Chemistry, Seoul National University, Seoul 151‐742, Korea WILLY J. PEUMANS Department of Molecular Biotechnology, Laboratory of Biochemistry and Glycobiology, Ghent University, 9000 Gent, Belgium PIL JOON SEO Molecular Signaling Laboratory, Department of Chemistry, Seoul National University, Seoul 151‐742, Korea MING‐DER SHIH Institute of Plant and Microbial Biology, Academia Sinica, Taipei 11529, Taiwan

CONTENTS OF VOLUMES 35–47 Series Editor (Volumes 35–44) J.A. CALLOW School of Biosciences, University of Birmingham, Birmingham, United Kingdom

Contents of Volume 35 Recent Advances in the Cell Biology of Chlorophyll Catabolism H. THOMAS, H. OUGHAM and S. HORTENSTEINER The Microspore: A Haploid Multipurpose Cell A. TOURAEV, M. PFOSSER and E. HEBERLE-BORS The Seed Oleosins: Structure Properties and Biological Role J. NAPIER, F. BEAUDOIN, A. TATHAM and P. SHEWRY Compartmentation of Proteins in the Protein Storage Vacuole: A Compound Organelle in Plant Cells L. JIANG and J. ROGERS Intraspecific Variation in Seaweeds: The Application of New Tools and Approaches C. MAGGS and R. WATTIER Glucosinolates and Their Degradation Products R. F. MITHEN

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Contents of Volume 36 PLANT VIRUS VECTOR INTERACTIONS Edited by R. Plumb Aphids: Non-Persistent Transmission T. P. PIRONE and K. L. PERRY Persistent Transmission of Luteoviruses by Aphids B. REAVY and M. A. MAYO Fungi M. J. ADAMS Whitefly Transmission of Plant Viruses J. K. BROWN and H. CZOSNEK Beetles R. C. GERGERICH Thrips as Vectors of Tospoviruses D. E. ULLMAN, R. MEIDEROS, L. R. CAMPBELL, A. E. WHITFIELD, J. L. SHERWOOD and T. L. GERMAN Virus Transmission by Leafhoppers, Planthoppers and Treehoppers (Auchenorrhyncha, Homoptera) E. AMMAR and L. R. NAULT Nematodes S. A. MacFARLANE, R. NEILSON and D. J. F. BROWN Other Vectors R. T. PLUMB

CONTENTS OF VOLUMES 35–47

Contents of Volume 37 ANTHOCYANINS IN LEAVES Edited by K. S. Gould and D. W. Lee Anthocyanins in Leaves and Other Vegetative Organs: An Introduction D. W. LEE and K. S. GOULD Le Rouge et le Noir: Are Anthocyanins Plant Melanins? G. S. TIMMINS, N. M. HOLBROOK and T. S. FEILD Anthocyanins in Leaves: History, Phylogeny and Development D. W. LEE The Final Steps in Anthocyanin Formation: A Story of Modification and Sequestration C. S. WINEFIELD Molecular Genetics and Control of Anthocyanin Expression B. WINKEL-SHIRLEY Differential Expression and Functional Significance of Anthocyanins in Relation to Phasic Development in Hedera helix L. W. P. HACKETT Do Anthocyanins Function as Osmoregulators in Leaf Tissues? L. CHALKER-SCOTT The Role of Anthocyanins for Photosynthesis of Alaskan Arctic Evergreens During Snowmelt S. F. OBERBAUER and G. STARR Anthocyanins in Autumn Leaf Senescence D. W. LEE A Unified Explanation for Anthocyanins in Leaves? K. S. GOULD, S. O. NEILL and T. C. VOGELMANN

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Contents of Volume 38 An Epidemiological Framework for Disease Management C. A. GILLIGAN Golgi-independent Trafficking of Macromolecules to the Plant Vacuole D. C. BASSHAM Phosphoenolpyruvate Carboxykinase: Structure, Function and Regulation R. P. WALKER and Z.-H. CHEN Developmental Genetics of the Angiosperm Leaf C. A. KIDNER, M. C. P. TIMMERMANS, M. E. BYRNE and R. A. MARTIENSSEN A Model for the Evolution and Genesis of the Pseudotetraploid Arabidopsis thaliana Genome Y. HENRY, A. CHAMPION, I. GY, A. PICAUD, A. LECHARNY and M. KREIS

Contents of Volume 39 Cumulative Subject Index Volumes 1–38

Contents of Volume 40 Starch Synthesis in Cereal Grains K. TOMLINSON and K. DENYER The Hyperaccumulation of Metals by Plants M. R. MACNAIR Plant Chromatin — Learning from Similarities and Differences J. BRZESKI, J. DYCZKOWSKI, S. KACZANOWSKI, P. ZIELENKIEWICZ and A. JERZMANOWSKI

CONTENTS OF VOLUMES 35–47

The Interface Between the Cell Cycle and Programmed Cell Death in Higher Plants: From Division unto Death D. FRANCIS The Importance of Extracellular Carbohydrate Production by Marine Epipelic Diatoms G. J. C. UNDERWOOD and D. M. PATERSON Fungal Pathogens of Insects: Cuticle Degrading Enzymes and Toxins A. K. CHARNLEY

Contents of Volume 41 Multiple Responses of Rhizobia to Flavonoids During Legume Root Infection JAMES E. COOPER Investigating and Manipulating Lignin Biosynthesis in the Postgenomic Era CLAIRE HALPIN Application of Thermal Imaging and Infrared Sensing in Plant Physiology and Ecophysiology HAMLYN G. JONES Sequences and Phylogenies of Plant Pararetroviruses, Viruses, and Transposable Elements CELIA HANSEN and J. S. HESLOP-HARRISON

Role of Plasmodesmata Regulation in Plant Development ARNAUD COMPLAINVILLE and MARTIN CRESPI

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Contents of Volume 42 Chemical Manipulation of Antioxidant Defences in Plants ROBERT EDWARDS, MELISSA BRAZIER-HICKS, DAVID P. DIXON and IAN CUMMINS The Impact of Molecular Data in Fungal Systematics P. D. BRIDGE, B. M. SPOONER and P. J. ROBERTS Cytoskeletal Regulation of the Plane of Cell Division: An Essential Component of Plant Development and Reproduction HILARY J. ROGERS Nitrogen and Carbon Metabolism in Plastids: Evolution, Integration, and Coordination with Reactions in the Cytosol ALYSON K. TOBIN and CAROLINE G. BOWSHER

Contents of Volume 43 Defensive and Sensory Chemical Ecology of Brown Algae CHARLES D. AMSLER and VICTORIA A. FAIRHEAD Regulation of Carbon and Amino Acid Metabolism: Roles of Sucrose Nonfermenting-1-Related Protein Kinase-1 and General Control Nonderepressible-2-Related Protein Kinase NIGEL G. HALFORD Opportunities for the Control of Brassicaceous Weeds of Cropping Systems Using Mycoherbicides AARON MAXWELL and JOHN K. SCOTT Stress Resistance and Disease Resistance in Seaweeds: The Role of Reactive Oxygen Metabolism MATTHEW J. DRING Nutrient Sensing and Signalling in Plants: Potassium and Phosphorus ANNA AMTMANN, JOHN P. HAMMOND, PATRICK ARMENGAUD and PHILIP J. WHITE

CONTENTS OF VOLUMES 35–47

Contents of Volume 44 Angiosperm Floral Evolution: Morphological Developmental Framework PETER K. ENDRESS Recent Developments Regarding the Evolutionary Origin of Flowers MICHAEL W. FROHLICH Duplication, Diversification, and Comparative Genetics of Angiosperm MADS-Box Genes VIVIAN F. IRISH Beyond the ABC-Model: Regulation of Floral Homeotic Genes LAURA M. ZAHN, BAOMIN FENG and HONG MA Missing Links: DNA-Binding and Target Gene Specificity of Floral Homeotic Proteins RAINER MELZER, KERSTIN KAUFMANN ¨ NTER THEIßEN and GU Genetics of Floral Development in Petunia ANNEKE RIJPKEMA, TOM GERATS and MICHIEL VANDENBUSSCHE Flower Development: The Antirrhinum Perspective BRENDAN DAVIES, MARIA CARTOLANO and ZSUZSANNA SCHWARZ-SOMMER Floral Developmental Genetics of Gerbera (Asteraceae) TEEMU H. TEERI, MIKA KOTILAINEN, ANNE UIMARI, SATU RUOKOLAINEN, YAN PENG NG, URSULA MALM, ¨ NEN, SUVI BROHOLM, ROOSA LAITINEN, ¨ LLA EIJA PO PAULA ELOMAA and VICTOR A. ALBERT Gene Duplication and Floral Developmental Genetics of Basal Eudicots ELENA M. KRAMER and ELIZABETH A. ZIMMER

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Genetics of Grass Flower Development CLINTON J. WHIPPLE and ROBERT J. SCHMIDT Developmental Gene Evolution and the Origin of Grass Inflorescence Diversity SIMON T. MALCOMBER, JILL C. PRESTON, RENATA REINHEIMER, JESSIE KOSSUTH and ELIZABETH A. KELLOGG Expression of Floral Regulators in Basal Angiosperms and the Origin and Evolution of ABC-Function PAMELA S. SOLTIS, DOUGLAS E. SOLTIS, SANGTAE KIM, ANDRE CHANDERBALI and MATYAS BUZGO The Molecular Evolutionary Ecology of Plant Development: Flowering Time in Arabidopsis thaliana KATHLEEN ENGELMANN and MICHAEL PURUGGANAN A Genomics Approach to the Study of Ancient Polyploidy and Floral Developmental Genetics JAMES H. LEEBENS-MACK, KERR WALL, JILL DUARTE, ZHENGUI ZHENG, DAVID OPPENHEIMER and CLAUDE DEPAMPHILIS Series Editors (Volume 45– ) JEAN-CLAUDE KADER Laboratoire Physiologie Cellulaire et Mole´culaire des Plantes, CNRS, Universite´ de Paris, Paris, France MICHEL DELSENY Laboratoire Ge´nome et De´veloppement des Plantes, CNRS IRD UP, Universite´ de Perpignan, Perpignan, France

Contents of Volume 45 RAPESEED BREEDING History, Origin and Evolution S. K. GUPTA and ADITYA PRATAP

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Breeding Methods B. RAI, S. K. GUPTA and ADITYA PRATAP The Chronicles of Oil and Meal Quality Improvement in Oilseed Rape ABHA AGNIHOTRI, DEEPAK PREM and KADAMBARI GUPTA Development and Practical Use of DNA Markers KATARZYNA MIKOLAJCZYK Self-Incompatibility RYO FUJIMOTO and TAKESHI NISHIO Fingerprinting of Oilseed Rape Cultivars ´ ˇ URN and JANA ZˇALUDOVA VLADISLAV C Haploid and Doubled Haploid Technology L. XU, U. NAJEEB, G. X. TANG, H. H. GU, G. Q. ZHANG, Y. HE and W. J. ZHOU Breeding for Apetalous Rape: Inheritance and Yield Physiology LIXI JIANG Breeding Herbicide-Tolerant Oilseed Rape Cultivars PETER B. E. MCVETTY and CARLA D. ZELMER Breeding for Blackleg Resistance: The Biology and Epidemiology W. G. DILANTHA FERNANDO, YU CHEN and KAVEH GHANBARNIA Development of Alloplasmic Rape MICHAL STARZYCKI, ELIGIA STARZYCKI and JAN PSZCZOLA Honeybees and Rapeseed: A Pollinator–Plant Interaction D. P. ABROL

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Genetic Variation and Metabolism of Glucosinolates NATALIA BELLOSTAS, ANNE DORTHE SØRENSEN, JENS CHRISTIAN SØRENSEN and HILMER SØRENSEN Mutagenesis: Generation and Evaluation of Induced Mutations SANJAY J. JAMBHULKAR Rapeseed Biotechnology VINITHA CARDOZA and C. NEAL STEWART, JR. Oilseed Rape: Co-existence and Gene Flow from Wild Species RIKKE BAGGER JØRGENSEN Evaluation, Maintenance, and Conservation of Germplasm RANBIR SINGH and S. K. SHARMA Oil Technology ¨ US BERTRAND MATTHA

Contents of Volume 46 INCORPORATING ADVANCES IN PLANT PATHOLOGY Nitric Oxide and Plant Growth Promoting Rhizobacteria: Common Features Influencing Root Growth and Development ´ NICA CREUS, MARI´A CELESTE MOLINA-FAVERO, CECILIA MO LUCIANA LANTERI, NATALIA CORREA-ARAGUNDE, MARI´A CRISTINA LOMBARDO, CARLOS ALBERTO BARASSI and LORENZO LAMATTINA How the Environment Regulates Root Architecture in Dicots ´ RIE LEFEBVRE, PHILIPPE MARIANA JOVANOVIC, VALE LAPORTE, SILVINA GONZALEZ-RIZZO, CHRISTINE LELANDAIS-BRIE`RE, FLORIAN FRUGIER, CAROLINE HARTMANN and MARTIN CRESPI

CONTENTS OF VOLUMES 35–47

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Aquaporins in Plants: From Molecular Structure to Integrated Functions OLIVIER POSTAIRE, LIONEL VERDOUCQ and CHRISTOPHE MAUREL Iron Dynamics in Plants JEAN-FRANC ¸ OIS BRIAT Plants and Arbuscular Mycorrhizal Fungi: Cues and Communication in the Early Steps of Symbiotic Interactions VIVIENNE GIANINAZZI-PEARSON, NATHALIE SE´JALON-DELMAS, ANDREA GENRE, SYLVAIN JEANDROZ and PAOLA BONFANTE Dynamic Defense of Marine Macroalgae Against Pathogens: From Early Activated to Gene-Regulated Responses AUDREY COSSE, CATHERINE LEBLANC and PHILIPPE POTIN

Contents of Volume 47 INCORPORATING ADVANCES IN PLANT PATHOLOGY The Plant Nucleolus ´ EZ-VA ´ SQUEZ AND FRANCISCO JAVIER MEDINA JULIO SA Expansins in Plant Development DONGSU CHOI, JEONG HOE KIM AND YI LEE Molecular Biology of Orchid Flowers: With Emphasis on Phalaenopsis WEN-CHIEH TSAI, YU-YUN HSIAO, ZHAO-JUN PAN, CHIACHI HSU, YA-PING YANG, WEN-HUEI CHEN AND HONG-HWA CHEN

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Molecular Physiology of Development and Quality of Citrus ´ S, JOSE´ M. FRANCISCO R. TADEO, MANUEL CERCO COLMENERO-FLORES, DOMINGO J. IGLESIAS, MIGUEL A. NARANJO, GABINO RI´OS, ESTHER CARRERA, OMAR RUIZ-RIVERO, IGNACIO LLISO, RAPHAE¨ L MORILLON, PATRICK OLLITRAULT AND MANUEL TALON Bamboo Taxonomy and Diversity in the Era of Molecular Markers MALAY DAS, SAMIK BHATTACHARYA, PARAMJIT SINGH, TARCISO S. FILGUEIRAS AND AMITA PAL

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32, Jamestown Road, London NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands First edition 2008 Copyright ß 2008, Elsevier Ltd. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@ elsevier.com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN: 978-0-12-374600-9 ISSN: 0065-2296 For information on all Academic Press publications visit our Web site at elsevierdirect.com Printed and bound in USA 08 09 10 11 12 10 9 8 7 6 5 4 3 2 1

Molecular Mechanisms Underlying Vascular Development

JAE‐HOON JUNG, SANG‐GYU KIM, PIL JOON SEO AND CHUNG‐MO PARK

Molecular Signaling Laboratory, Department of Chemistry, Seoul National University, Seoul 151‐742, Korea

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Vascular Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Xylem........................................................................... B. Phloem ......................................................................... III. Growth in Vascular Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Shoot Growth ................................................................. B. Comparison Between Dicots and Monocots ............................. C. Root Growth .................................................................. IV. Model Systems for Studying Vascular Development . . . . . . . . . . . . . . . . . . . . . A. Arabidopsis .................................................................... B. Populus ......................................................................... C. Zinnia........................................................................... V. Vascular Continuity and Regulatory Signals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Polar Auxin Transport ...................................................... B. Models for Vascular Strand Formation................................... VI. Regulation of Radial Patterning of Vascular Bundles . . . . . . . . . . . . . . . . . . . . A. Opposite Roles of HD‐ZIP IIIs and Kanadis ............................ B. Regulation of HD‐ZIP III Activities by Competitive Inhibitors ...... C. Other Adaxial–abaxial Axis Determinants ............................... VII. Identification of Vascular Development‐related Factors . . . . . . . . . . . . . . . . . A. HD‐ZIP IIIs and miRNA ................................................... B. Cytokinin ...................................................................... C. Brassinosteroid................................................................ D. Xylogen ........................................................................ E. Other Genes Involved in Vascular Differentiation ...................... Advances in Botanical Research, Vol. 48 Incorporating Advances in Plant Pathology Copyright 2008, Elsevier Ltd. All rights reserved.

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0065-2296/08 $35.00 DOI: 10.1016/S0065-2296(08)00401-1

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VIII. Tracheary Element Differentiation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Secondary Wall Formation ................................................. B. Programmed Cell Death..................................................... IX. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABSTRACT The plant vascular system is a complicated network of conducting tissues that interconnects all organs and transports water, minerals, nutrients, organic compounds, and various signaling molecules throughout the plant body. It is composed of two major tissues, xylem and phloem, that are diVerentiated from the meristemic tissue, procambium. Although physiological functions of the vascular tissues are well documented, genetic networks that regulate the process of vascular development are still poorly understood. Thorough researches on several model plants, such as Arabidopsis, Populus, and Zinnia, molecular genetic, genomic, and biochemical approaches have provided significant insights into the molecular mechanisms underlying vascular development. An array of signals and signaling molecules, including receptor molecules, ligands, vesicle‐traYcking components, and transcription factors, has been shown to mediate the specification and diVerentiation of vascular tissues and the vascular bundle patterning. Accordingly, future challenge in vascular biology is to understand how these signals are coordinately integrated to achieve optimized vascular development under a given growth condition.

ABBREVIATIONS APL ATHB BR CC CESA GT GX HD‐ZIP III HK IAA KAN miRNA PAT PCD PGP PINSV RAM

ALTERED PHLOEM DEVELOPMENT Arabidopsis thaliana homeobox brassinosteroid companion cell cellulose synthase glycosyltransferase glucuronoxylan homeodomain‐leucine zipper III histidine kinase indole‐3‐acetic acid KANADI microRNA polar auxin transport programmed cell death P‐glycoprotein PIN1‐specific vesicle root apical meristem

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RR SAM SCW SE START ta‐siRNAs TE ZeHB ZPR

response regulator shoot apical meristem secondary cell wall sieve element steroidogenic acute regulatory protein‐related lipid transfer trans‐acting short‐interfering RNAs tracheary element Zinnia elegans HB litter zipper

I. INTRODUCTION Land plants should develop ways of supporting their massive bodies, transporting water and inorganic material from the soil to plant organs, and relocating assimilates from the source to the sink. They have resolved most of these problems by evolving vascular system. Vascular plants are illustrated primarily by having the vascular tissues, consisting of highly diVerentiated conductive cells that function as tubes or vessels through which water and nutrients can be translocated throughout the plant body. These vessels connect all plant organs and serve as eYcient long‐distance transportation of water and diverse material. The vascular system consists of two major conductive tissues, xylem and phloem, that are diVerentiated from the meristematic tissues, such as procambium and vascular cambium (Aloni, 1987) (Fig. 1).

Vascular cambium Apical meristem

Fascicular cambium

Primary meristems

Interfascicular cambium

Protoderm Ground meristem Procambium Pith

Primary tissues Epidermis Ground tissue Vascular bundle Primary phloem Primary xylem Procambium

Primary growth

Fig. 1.

Stem developmental stages in angiosperms.

Rays Periderm Cortex Primary phloem Secondary phloem Vascular cambium Secondary xylem Primary xylem

Secondary growth

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The vascular system appeared early in the history of terrestrial plant life. Plant colonization on land may have occurred as early as in the Silurian period more than 400 million years ago (Scarpella and Meijer, 2004; Ye, 2002). To achieve successful transition from aquatic to terrestrial habitats, plants invoked mechanisms for protection against water loss, absorption of water and nutrients from the soil, transfer of assimilates throughout the entire organism, and mechanical support. The vascular tissues provide an eYcient way of resolving the problems of long‐distance transport of water and nutrients as well as providing suYcient rigidity for mechanical support. In the plant kingdom, the earliest example of an organized vascular system was encountered in the brown algae, which has a discernible phloem but no xylem (Aloni, 1987). The water‐conducting system evolved much later. It seems that lignin, which is indispensable for conducting water in plant stems, is apparently absent from the algae. However, lignin‐like compounds have been detected in Coleochaete, an algal model for land plant ancestry (Delwiche et al., 1989). Whereas the polysaccharide components of plant cell walls are highly hydrophilic and thus permeable to water, lignin is hydrophobic. The cross‐linking of polysaccharides by lignin is thus an obstacle for water absorption to the cell wall (Boerjan et al., 2003). Because of this reason, the incorporation of lignin became mandatory for the development of plants on land, because the lignified conducting and supporting cells in the xylem provide long‐distance water conduction and mechanical support necessary for the terrestrial habitat. Vascular plants can be grouped into three major classes: ferns, gymnosperms, and angiosperms. Ferns have the least evolved vascular system. They have vascular systems with specialized leaf and root structures but are still dependent on moist environments for reproduction. Gymnosperms or coniferous plants and angiosperms or flowering plants, known together as seed plants, have evolved reproductive processes that are independent of water. In addition, their embryos are encapsulated in tough coats, which prevent desiccation in the terrestrial environment and protect the seeds until environmental conditions are favorable for germination and growth. Angiosperms are further classified into two subgroups: monocots and dicots. Vascular bundles of dicots are arranged concentrically, while those of monocots are typically not arranged in a single circular array and lack secondary thickening growth from the cambium (Sachs, 1981). Vascular development has conventionally been studied through physiological, biochemical, and molecular approaches (Ye, 2002). Early physiologists have established that plant hormones, such as auxin and cytokinin, are essential for vascular tissue diVerentiation (Aloni, 1987; Sachs, 1991). A number of proteins and genes regulating diVerent stages of vascular development,

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such as early xylem diVerentiation, secondary cell wall (SCW) thickening, and programmed cell death (PCD), have been characterized through biochemical and subtractive hybridization approaches (Fukuda, 1997; Turner et al., 2007). Since vascular tissues develop internally and identification of vascular cell types are technically diYcult, cell type‐specific molecular markers and tissue‐ specific reporter genes are useful to visualize the diVerentiation process of vascular tissues. Two of the most useful reporter constructs are derived from the ARABIDOPSIS THALIANA HOMEOBOX 8 (ATHB8) and TED3 genes (Turner and Sieburth, 2002). The promoter of ATHB8, which is highly expressed in procambial cells, can be used as a marker for early stages of vascular diVerentiation (Baima et al., 1995; Elge et al., 2001). The promoter of TED3, which is specifically expressed in xylem cells, can be used for the study of xylogenesis (Igarashi et al., 1998). Histological staining methods with dyes, such as toluidine blue and phloroglucinol‐HCl, can visualize tracheary elements (TEs), main components of the xylem tissue (O’Brien and McCully, 1981). While these stains only indicate a lignified cell wall, Maule’s reagent can be used to discriminate between xylem and interfascicular cells, depending on their lignin compositions (Chapple et al., 1992). In addition, Aniline blue staining causes the phloem to fluoresce yellow under UV because of dye binding to callose deposited on the sieve plates. Free‐hand sections stained with these dyes give better anatomical images for the vasculature of stems and leaves (O’Brien and McCully, 1981). For high‐ quality images, confocal microscopy is employed to visualize the vascular tissues in the leaves and roots. During the last two decades, a significant progress in understanding the molecular mechanisms underlying vascular tissue diVerentiation has been achieved using diverse genetic and molecular biological approaches. Arabidopsis genetics is a powerful approach to isolate mutants with abnormal phenotypes in vascular development. Several key genes controlling vascular tissue diVerentiation have been identified through these approaches (Baucher et al., 2007; Fukuda, 2004; Ye, 2002). The genes characterized so far include PINFORMED1 (PIN1) (Ga¨lweiler et al., 1998) and GNOM (Steinmann et al., 1999), which are closely related to auxin transport and signaling, CONSTITUTIVE PHOTOMORPHOGENESIS AND DWARFISM (CPD) (Mathur et al., 1998), DWARF4 (DWF4) (Choe et al., 1998), and BRI1‐LIKEs (BRLs) (Can˜o‐Delgado et al., 2004; Zhou et al., 2004) involved in brassinosteroid (BR) biosynthesis and signaling, and xylogen genes modulating continuity of the vascular strands (Motose et al., 2004). The HD‐ZIP III and KANADI (KAN) genes regulate vascular pattern formation (Emery et al., 2003; Ohashi‐Ito and Fukuda, 2003). Furthermore, while ALTERED PHLOEM DEVELOPMENT

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(APL) is related to the xylem‐phloem switching (Bonke et al., 2003), VASCULAR‐RELATED NAC‐DOMAIN6 (VND6) and VND7 (Kubo et al., 2005) regulate xylem vessel transdiVerentiation. Using these genes, upstream and downstream genes can easily be isolated by molecular and genomic approaches, such as direct target screenings, microarrays, and yeast two‐hybrid analysis. In this chapter, we briefly describe the structure and function of the plant vascular system and summarize recent findings on vascular tissue diVerentiation. We focus mainly on data collected from Arabidopsis, which is widely used as a model plant for the study of primary growth, and Populus, a model plant frequently used to study secondary growth in wood plants. This chapter also includes molecular components and mechanisms aVecting vascular continuity and vascular cell development factors. The factors mediating xylem tissue diVerentiation are also discussed. For more in‐depth understanding of other aspects of this topic, other recent reviews will be helpful to the readers (Baucher et al., 2007; Carlsbecker and Helariutta, 2005; Fukuda, 2004; Helariutta, 2007; Scarpella and Meijer, 2004; Sieburth and Deyholos, 2006; Turner et al., 2007; Ye, 2002).

II. VASCULAR TISSUES The vascular tissues play essential roles in a range of physiological (transport of water and nutrients), developmental (transfer of signaling molecules), and architectural (physical support) aspects of plant growth. Basic units of the vascular tissue are xylem and phloem. The xylem transports and stores water, nutrients, and various plant hormones. It also lends a major mechanical support. The phloem provides passageways for the distribution of photosynthetic products, such as sucrose, and for translocation of the proteins and mRNAs involved in plant growth and development. Both xylem and phloem usually comprise more than one cell types, including conducting elements (TEs in the xylem and SEs in the phloem) and parenchyma and sclerenchyma cells (Aloni, 1987; Fukuda, 2004; Ye, 2002). Cells in the diVerentiated vascular tissue are typically long and slender. Since xylem and phloem function in the internal conduction of fluid and nutrients, they are structurally similar to pipelines. As plants grow, new vascular tissues diVerentiate in the growing tips of the plant. The new tissues are aligned with existing vascular tissues, forming the pathways connecting plant organs throughout the plant parts. The vascular tissues are arranged in long, discrete strands called vascular bundles, which are also referred to as vascular strands. These bundles also include supporting and protective cells

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7

in addition to xylem and phloem. In the stems and roots, the xylem typically lies closer to the interior of the stem with the phloem toward the exterior of the stem. A. XYLEM

Xylem formation is one of the most intensive fields of cell diVerentiation in plants. The study of xylem tissue diVerentiation has been driven by the economic usefulness of xylem as a major constituent of wood. The word ‘‘xylem’’ is derived from the Greek xylon, meaning wood (Tyree and Zimmermann, 2003). The xylem consists of a number of cell types, such as TEs, xylem parenchyma cells, and xylem fibers. The TEs are highly specialized and easily traceable water‐conducting cells. At maturity, they are emptied by the loss of all intracellular contents, including nucleus and cytoplasm, to form hollow tunnels (Fig. 6). The remaining cell walls are very thick and provide mechanical support for the plant to prevent them from collapsing under the high pressure resulting from fluid uptake, and the cavities inside provide a passage through which fluids can move. Moreover, SCW thickening provides an additional strength and rigidity. The presence of TEs is a defining characteristic of vascular plants to diVerentiate them from non‐vascular plants (Turner et al., 2007; Tyree and Zimmermann, 2003; Ye, 2002). There are two essential types of TEs in the xylem, tracheids and vessel elements. Tracheids are elongated, spindle‐shaped cells that communicate with adjoining tracheids by means of the numerous pits in their lateral walls. These pits are microscopic regions where the SCW is absent and the primary cell wall is thin and porous. The pits of one tracheid are typically located opposite to those of an adjoining tracheid, forming pit pairs. The pit pairs constitute a low‐resistance path for water movement between tracheids. The porous layer between the pit pairs, consisting of two primary walls and a middle lamella, is called pit membrane. Because tracheids have a much higher surface to volume ratio compared to vessel elements, the ability of high adhesion serves to hold water against gravity when transpiration is not occurring. This is likely an adaptation mechanism that helps plants prevent air embolisms (Turner et al., 2007; Tyree and Zimmermann, 2003). Vessel elements tend to be shorter and wider than tracheids and have perforated end walls that form perforation plate at each end of the cell. Like tracheids, vessel elements also have pits on their lateral walls. However, unlike tracheids, which are arranged in overlapping vertical files, vessel members are stacked end to end to form a larger unit called vessel. Because of the open cross walls in the vessels, they provide an eYcient way of conducting water. Vessel elements are found only in angiosperms and a

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small crop of gymnosperms called the Gnetales. In contrast, tracheids are present in both angiosperms and gymnosperms (Turner et al., 2007; Tyree and Zimmermann, 2003). B. PHLOEM

Unlike the xylem, which is composed primarily of dead cells, the phloem consists of live cells that transport sap both up and down through the plant body. The sap is a water‐based solution but rich in sugars synthesized by the photosynthetic regions. The sugars are transported to non‐photosynthetic parts, such as roots, or into storage structures, such as tubers or bulbs. The phloem contains several diVerent types of cells or structures, such as sieve elements (SEs), companion cells (CCs), phloem parenchyma cells, and phloem fibers. Among these, only the SEs are directly involved in translocation of sugars and other organic materials (Taiz and Zeiger, 2006; Thompson, 2006). Mature SEs are unique among living plant cells in that they lack many subcellular structures normally found in living cells. The SEs lack the nuclei and have very few vacuoles. Microfilaments, microtubules, Golgi bodies, and ribosomes are also generally absent from the mature cells. The walls are nonlignified, although they are secondarily thickened in some cases. There are two diVerent types, the highly diVerentiated sieve tube elements typical of the angiosperms and the relatively unspecialized sieve cells of gymnosperms (Taiz and Zeiger, 2006; van Bel et al., 2002). The SEs are further characterized by sieve areas, portions of the cell wall where pores interconnect the conducting cells. The sieve areas of sieve tube elements in angiosperm are more specialized than those in gymnosperms. For example, some of the sieve areas of sieve tube elements are diVerentiated in sieve plates, but sieve cells have no sieve plates, and all sieve areas are similar. Sieve plates have larger pores than other sieve areas in the cell and are generally found on the end walls of sieve tube elements, where the individual cells are joined together to form a longitudinal series called a sieve tube. The sieve plate pores of sieve tube elements are open channels that allow transport between cells. In the sieve cells, on the other hand, the pores of sieve cells are relatively unspecialized and appear to be blocked with numerous membranes. Each sieve tube element is usually associated with one or more CCs. The sieve tube element and the CC are formed by division of a single mother cell. The plasmodesmata, cytoplasmic strands that connect these two types of cells, penetrate the walls between sieve tube elements and their CCs, suggesting a close functional relationship and ease of transport between the two cells (Cilia and Jackson, 2004; Taiz and Zeiger, 2006). CCs play a role in the transport of photosynthetic products from producing cells in the mature

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leaves to the SEs in minor veins of the leaf. They also take over some of the critical metabolic functions, such as protein synthesis, that are reduced or lost during diVerentiation of the SEs. In addition, numerous mitochondria in CCs may supply energy in the form of ATP to the SEs. There are three types of CCs. Ordinary CCs have smooth walls and few or no plasmodesmata connections to cells other than the sieve tube. Transfer cells have much folded walls that are adjacent to non‐sieve cells, allowing for larger areas of transfer. Intermediary cells have smooth walls and numerous plasmodesmata connecting them to adjacent cells. The first two types of cells collect solutes through apoplastic transfers, while the third type can collect solutes symplastically through the plasmodesmata connections.

III. GROWTH IN VASCULAR PLANTS Plant growth occurs predominantly in the highly localized regions of cell division called meristems. Nearly all nuclear division (mitosis) and cell divisions (cytokinesis) take place in the meristemic regions. DiVerentiated plant cells generally cannot divide or produce cells of a diVerent type. Therefore, cell division in the meristems is required to provide new cells for expansion and diVerentiation of all tissues and initiation of new organs. Accordingly, maintenance of the cells requires a balance between two antagonistic processes: organ initiation and stem cell population renewal. The meristematic tissues are located at the tips of the shoot and root and are called shoot apical meristem (SAM) and root apical meristem (RAM), respectively (Figs. 1 and 3). Apical meristems may diVerentiate into three types of primary meristems, protoderm, procambium, and ground meristem. Protoderm lies around the outside of the stem and develops into the epidermis. Procambium lies just inside of the protoderm and develops into primary xylem and primary phloem. It also produces the vascular cambium, a secondary meristem. Ground meristem develops into the pith. It additionally produces the cork cambium, another secondary meristem. These meristems are responsible for primary growth and increase in length or height (Figs. 1 and 3). Secondary meristems are called lateral meristems because they surround the established stem of a plant and cause it to grow larger in diameter. Vascular cambium produces secondary xylem and secondary phloem, and this process continues throughout the plant life. This is what gives rise to wood in plants. Cork cambium gives rise to the bark of tree. In Arabidopsis, mature embryos contain procambial cells that will diVerentiate into xylem and phloem following germination, indicating that the vascular pattern is established during embryogenesis (Turner and Sieburth, 2002).

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The first indication of procambial cell formation becomes apparent during the transition from the globular to heart stages of embryogenesis, when the isodiametric cells of the lower tier of the embryo elongate in the apical–basal axis. As embryo completes diVerentiation, the procambium forms the apical–basal axis defined by a hypocotyl that contains a variable number of procambium cell files bounded by a single layer of pericycle. Finally, the procambium is connected with other procambial cells. This process occurs within the cotyledons, forming a continuous network of procambial cells (Busse and Evert, 1999). In most non‐embryonic tissues, vascular development is a continuous process in which procambium development is followed by diVerentiation of the xylem and phloem. In embryos, however, further diVerentiation of the procambium is halted during embryo maturation and further proceeds only when growth resumes following seed germination. After seed germination and during primary stem growth, most of the procambial cells diVerentiate into xylem centripetally and phloem centrifugally, leading to the formation of vascular bundles that are arranged through the ground tissues (Fukuda, 2004; Ye, 2002). Root growth continues to produce the narrow cylinder shape of vascular tissues present in the center of root, while shoot growth forms a ring of vascular bundles in a circular pattern around the stem (Fig. 3). Vascular cambium, a lateral meristem, is derived from procambial cells within the vascular bundles and from other parenchyma cells in the interfascicular region (Fig. 1). The transition from primary to secondary growth is intimately associated with the formation and functioning of vascular cambium. Vascular cambium divides to produce secondary xylem cells on the inside and secondary phloem cells on the outside. This growth increases the girth of the plant root or stem rather than length and triggers secondary thickening. As long as the vascular cambium continues to produce new cells, the stem or root will continue to grow in diameter. In woody plants, this process produces wood (Turner et al., 2007). A. SHOOT GROWTH

Primary growth of the shoots results from the activity of SAM, in which cell division is followed by progressive cell enlargement, typically elongation (Fig. 1). The initial step in the primary growth is the formation of procambium precursor cells. These cells can be visible during embryogenesis stage (Busse and Evert, 1999). Procambial cells give rise to xylem and phloem precursor cells. The final step in the primary growth is the specification into distinct types of vascular cells from the xylem and phloem precursor cells. Phloem precursor cells subsequently diVerentiate into various phloem cells,

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such as SEs, CCs, phloem parenchyma cells, and phloem fibers. Xylem precursor cells diVerentiate into TEs, xylem parenchyma cells, and xylem fibers. Xylem and phloem formed during primary growth are called primary xylem and primary phloem, respectively. In particular, primary xylem is divided into two types, protoxylem and metaxylem. Metaxylem develops following the protoxylem but before secondary xylem. It is distinguished by wider vessels and tracheids. Primary xylem, primary phloem, and procambium together form vascular bundles (Scarpella and Meijer, 2004; Taiz and Zeiger, 2006; Ye, 2002). Secondary growth is a growth of thickness. In many vascular plants, secondary growth is caused by the activity of the vascular cambium (Fig. 1). Vascular cambium exists between xylem and phloem; on its inside the cambium produces secondary xylem, and on its outside it forms secondary phloem. The primary xylem and phloem are pushed further inward and outward, respectively. The cells of the vascular cambium are often termed initials, since they initiate the formation of specialized precursor cells after division. The vascular cambium usually has two morphologically distinct types of initials, fusiform initials and ray initials. Fusiform initials are the mother cells of all secondary xylem and phloem cells. The axillary (parallel to the organ axis)‐elongated fusiform initials lead to the formation of the axial system, including tracheids, vessel elements, fibers, axial parenchyma cells, SEs, and CCs. They are highly vacuolated and have spindle‐like shapes. Ray initials, on the other hand, are nearly isodiametric, with smaller cells than fusiform initials. They generate the radially orientated rays in wooden plant (Iqbal and Ghouse, 1990). Because this growth usually ruptures the epidermis of the stem or roots, plants with secondary growth simultaneously develop a cork cambium, which is also referred to as phyllogen. The cork cambium is a secondary lateral meristem that produces the secondary outer surface, the bark, which replaces the epidermis. B. COMPARISON BETWEEN DICOTS AND MONOCOTS

In dicots, the vascular bundle consists essentially of four layers (Fig. 2). Toward the cortex, the vascular bundle may show a layer of fibers, which are occasionally called phloem fibers. A group of fibers frequently forms the bundle cap of the primary vascular bundle. The secondary xylem and phloem, which are produced by the vascular cambium, often crush the primary phloem cells, so that the primary phloem fibers may be the only recognizable evidence of the primary phloem in mature stems. Moving interior to the fibers, there is a functional primary phloem, including SEs and CCs.

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Dicot vascular bundle Cortex

Monocot vascular bundle Bundle sheath

Epidermis

Primary phloem Bundle cap fibers Functional phloem Vascular cambium

Pith

Fig. 2.

Primary xylem Metaxylem Protoxylem

Center

Comparison of vascular bundles in dicots and monocots.

The presence of two cell types with diVerent appearances gives the functional phloem area a superficially messy look. The next layer toward the pith is the vascular cambium. These cells are meristematic and divide mitotically to produce additional secondary xylem and secondary phloem in woody plants. The innermost layer of the vascular bundle is the primary xylem. Primary growths in the stems of monocots and dicots diVer in the arrangement of vascular bundles. In monocots, the vascular bundles have a random appearance scattered throughout the stem. In contrast, they are arranged in a circle near the outer edge just below the epidermis in dicots (Fig. 2). In addition, monocots usually do not have a vascular cambium, and thus they are unable to undergo secondary growth. The dicots and gymnosperms have vascular cambium. However, some woody monocots, such as agave, aloe, palm, and yucca, can develop substantial stems because of thickening growth by cell division and elongation of parenchyma cells in the ground tissue (Sachs, 1981; Scarpella and Meijer, 2004). C. ROOT GROWTH

In vascular plants, the root is the organ that typically lies below the surface of soil. Consequently, the major function of the root is absorption of water and minerals from the soil and anchoring of the plant body to the ground. They also function often as a storage organ for food (de Dorlodot et al., 2007; Schenk and Jackson, 2002). The Arabidopsis root is composed of four concentric cell layers, epidermis, cortex, endodermis, and stele (Fig. 3) (Taiz and Zeiger, 2006; Turner and Sieburth, 2002). The epidermis is the outside surface of the primary root and the extension of epidermal cells from the root hair. The cortex lies interior to the epidermis and comprises the bulk of the primary root. It functions

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Cell division region

Elongation region

Differentiation region

Primary tissues Epidermis Cortex Endodermis Pericycle Phloem pole Procambium Primary xylem Primary meristems Protoderm

Stele (vascular cylinder)

Ground meristem Procambium

Apical meristem Root tip

Fig. 3.

Root developmental stages in angiosperms.

primarily in storage of starch. The innermost one‐cell thick layer of the cortex is the endodermis, which surrounds the stele. The stele, which is also referred to as vascular cylinder, contains the vascular elements of xylem and phloem, procambium, and pericycle. The pericycle lines the outer edge of the stele and can produce lateral roots. Unlike monocot stems, the radial pattern of vascular tissues in monocot roots is arranged in a ring similar to that in dicot roots. Moreover, monocot roots are similar to dicot roots in that they contain pericycle, endodermis, cortex, and epidermis. However, the xylem and phloem cells in monocot roots are arranged in a circle around the pith, the central region of the stele. In dicot roots, however, the xylem cells form a central hub with lobes, and phloem cells fill in the spaces between the lobes. There are three areas of cells responsible for successful root growth, and such cell areas are directly related with cell growth stages: division, elongation, and cell diVerentiation (Fig. 3). Because root vascular tissues are derived from the activity of the RAM, root growth initiates at the bottom of the root. At the tip of every growing root, the root cap provides a mechanical protection to the meristemic tissue as the root advances through the soil. Behind the area of cell division, cells are elongated by
10 times compared to the original cell length. During this stage, the cells begin to diVerentiate into the protoderm, vascular tissue, and ground tissue. Finally, cells finish specializing into the dermis, vascular tissue, and cortex at the maturation zone. Root hairs appear first in this region.

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Roots of many vascular plants, especially dicots and gymnosperms, often undergo secondary growth, an increase of its diameter. A vascular cambium forms in the stele to produce secondary phloem and xylem. The epidermis is replaced by the periderm. As the stele increases in diameter, the cortex, pericycle, and endodermis disappear. Even nonwoody roots often undergo secondary growth, including those of tomato and alfalfa (de Dorlodot et al., 2007; Schenk and Jackson, 2002).

IV. MODEL SYSTEMS FOR STUDYING VASCULAR DEVELOPMENT It is generally accepted that biological functions are highly conserved in all living organisms. For example, genes that are involved in human heritable diseases have close homologues in the Arabidopsis genome (Arabidopsis Genome Initiative, 2000). Nevertheless, it is obvious that individual species have distinct features that can be discriminated physiologically and genetically. One example is vascular development, which is unique to plants and regulated a group of genes, if not all, present only in plant genomes. Our understanding of vascular tissue diVerentiation has been greatly enhanced in recent years using several model systems, such as Arabidopsis thaliana, Populus, and Zinnia elegans (Jansson and Douglas, 2007; Taylor, 2002; Turner et al., 2007). A. ARABIDOPSIS

Most of our current knowledge on plant regulatory mechanisms is based on the model dicot plant Arabidopsis thaliana. Molecular genetic studies on Arabidopsis have been employed to isolate numerous mutants in which diverse aspects of plant growth and development are disrupted. Arabidopsis is widely used as a model species for a series of obvious reasons: small physical size, short generation time, straightforward genetics, large number of oVspring, and small genome size (Goodman et al., 1995). In addition, its full genomic sequence has been fully determined, and numerous mutants have been reported. Arabidopsis is a powerful system for genetic analysis of vascular diVerentiation and pattern formation. It can be used to study not only the diVerentiation of multiple cell types in the vascular tissues but also the pattern formation at the organ level (Ye, 2002). Furthermore, TE cell culture systems with cell suspension are well established in this model plant. Although not as eYcient as the Zinnia system (see below), Arabidopsis cell suspensions can be

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induced to form up to 30% TEs within a 96‐h cell culture period under inductive conditions (Kubo et al., 2005; Oda et al., 2005). A large number of Arabidopsis mutants have been described with phenotypic abnormality in vascular tissue diVerentiation. These mutants exhibit various malformations in diverse aspects of vascular development, such as in the alignment of vascular bundles (Can˜o‐Delgado et al., 2000), formation of a network of veins in the leaves (Carland et al., 2002), division and diVerentiation of the procambial cells (Ma¨ho¨nen et al., 2000; Scheres et al., 1995), diVerentiation of primary xylem (Bonke et al., 2003; Can˜o‐Delgado et al., 2004; Kubo et al., 2005), and in the organization of vascular tissues within the bundles of the leaves and stems (Emery et al., 2003; Eshed et al., 2004; Green et al., 2005; Kidner and Martienssen, 2004; Kim et al., 2005; Li et al., 2005; McConnell et al., 2001; Prigge et al., 2005; Williams et al., 2005b; Zhong and Ye, 1999). However, the Arabidopsis system has some weak points to be used as a model system for vascular system research. Although it makes functional studies very easy, most plant organisms have diVerent reproductive strategies, making it a genetic extreme. The very short life cycle of Arabidopsis also makes many traits that are essential in most plants insignificant in this plant species. Seasonal growth and wood formation are obvious examples. Nevertheless, the Arabidopsis system is still useful, and a battery of mutants aVecting various aspects of vascular diVerentiation can easily and eYciently be isolated through numerous screening approaches (Jansson and Douglas, 2007; Taylor, 2002; Turner et al., 2007; Ye, 2002). B. POPULUS

Considering tree‐unique traits, such as wood formation, perennial growth, and seasonality, there is a need for a model tree to study specific aspects of tree biology and to understand regulatory mechanisms causing those unique traits. Xylem diVerentiation during secondary growth is the most obvious feature of trees. The Populus system oVers many potentials to study tree‐ specific traits that cannot easily be addressed in Arabidopsis and rice (Jansson and Douglas, 2007; Taylor, 2002). It is satisfying the ‘‘model’’ role for a number of reasons: fast growing, easy propagation, wide genetic diversity, ease of genetic transformation, establishment of molecular genetic maps, and tree genomics (Taylor, 2002). As a model system for tree biology, the development of Populus has been largely driven by the establishment of genomic and molecular biology resources for this plant species (Tuskan et al., 2006). Interestingly, in angiosperm phylogeny showing the eurosid clade containing Populus and

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Arabidopsis, Arabidopsis is more related to Populus than to the vast majority of other dicot taxa, such as gymnosperm trees (Jansson and Douglas, 2007). This information will facilitate comparative studies on Arabidopsis and Populus, enabling discovery of mechanisms conserved among eudicots. For example, the CONSTANS/FLOWERING LOCUS T module in Arabidopsis regulates the photoperiodic induction of flowering. In Populus, however, the same module regulates not only flowering initiation but also bud set in the late season, a process apparently absent in Arabidopsis (Bohlenius et al., 2006). Connection between the outputs from the studies of Populus and Arabidopsis are likely to be fruitful for the study on the possible overlapping regulatory mechanisms of vascular development within plant parts. Populus is particularly a powerful model system for the study of the highly organized nature of xylem diVerentiation during secondary growth. Although Arabidopsis can form secondary xylem tissues under inductive conditions (ChaVey et al., 2002), Populus provides better opportunities to study important plant processes absent or poorly developed in Arabidopsis, such as secondary xylem development (Tuskan et al., 2006; van Raemdonck et al., 2005) and functioning of the vascular cambium (Iliev and Savidge, 1999; Tuskan et al., 2006). A suite of critical genomic and molecular tools, such as genome sequences, EST collections, DNA microarrays, and transformation protocols, have been developed for Populus. It is thus expected that the study of secondary vascular tissue development in Populus will progress rapidly in the future. Recent genome‐wide expression analyses have identified candidate genes for secondary growth, and several other genes known to regulate the SAM activity have also been found to be expressed in the vascular cambium (Baucher et al., 2007).

C. ZINNIA

The formation of TEs from isolated leaf mesophyll cells in Zinnia elegans provides an excellent in vitro system for studying xylem diVerentiation (Fukuda, 1997; Fukuda and Komamine, 1980). When the mesophyll cells isolated from young Zinnia leaves are cultured in vitro in the presence of auxin and cytokinin, they transdiVerentiate into TEs within 72 h. This in vitro transdiVerentiation process mimics the in vivo xylem cell development. The advantage of this system is that the induction rate of TEs can reach up to 80% and isolated mesophyll cells are almost homogeneous. Consequently, the Zinnia cell culture system is useful for monitoring the series of events occurring during xylem diVerentiation because the process can be followed in

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single cells and the interference by other cell types is minimal (Fukuda, 2004; Helariutta, 2007; Turner et al., 2007; Ye, 2002). The transdiVerentiation of isolated Zinnia mesophyll cells proceeds through three stages (Fukuda, 1997, 2004). At stage 1, isolated mesophyll cells are induced to dediVerentiate by wounding with a combination of auxin and cytokinin. This process corresponds to the functional dediVerentiation process but occurs without cell division. At stage 2, dediVerentiated cells diVerentiate into procambial cells and subsequently into xylem cell precursors. At stage 3, TEs and xylem parenchyma cells are produced through the diVerentiation of xylem cell precursors. Additionally, a series of TE diVerentiation process involves patterned secondary‐cell‐wall deposition and PCD. Especially, BRs are actively synthesized during stage 2, and this is frequently used as a marker for the diVerentiation of procambial cell and xylem cell precursors (Yamamoto et al., 1997). By using the cultured Zinnia cell system in combination with genomic‐wide gene expression analysis and biochemical fractionation of influencing agents on diVerentiation, a number of genes associated with TE diVerentiation have been characterized (Demura et al., 2002; Ito et al., 2006; Milioni et al., 2002; Motose et al., 2004).

V. VASCULAR CONTINUITY AND REGULATORY SIGNALS The vascular tissues in the shoot (rosette and inflorescence) are present as a ring of separate veins. This pattern of stem veins serves two essential functions. It provides an apical–basal axis for material transport and serves as a link that connects the veins of lateral organs. A major issue in stem vein patterning is how the vascular tissues of lateral branches and organs are connected with the veins of the stem. Since the rosette leaves are highly compressed, Arabidopsis is not an appropriate plant for this study (Busse and Evert, 1999). The vascular tissues usually diVerentiate at predictable positions of the procambium. However, their diVerentiation also exhibits a high level of flexibility. For example, vascular patterns along the main axis of the plant is largely maintained through the elaboration and elongation of existing strand patterns, rather than being newly generated, but the arrangement in the leaf vascular strands is created de novo during the development of leaf primordium. Additional connections with existing vascular strands within fully expanding leave, emerging adventitious organs, and wounded tissues suggest the existence of partial self‐organizing mechanisms. Therefore, identification

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of the positional signals controlling vascular cell diVerentiation are crucial in understanding the self‐organized vascular patterning (Sachs, 1981; Scarpella and Meijer, 2004; Turner and Sieburth, 2002). It has long been proposed that auxin, which is basipetally transported from the SAM and young leaves, induces formation of procambial cells (Aloni, 1987; Sachs, 1991). Indole‐3‐acetic acid (IAA), a natural auxin species, can replace the inductive eVects of developing leaves on the vascular tissue formation, suggesting that auxin may act as a prominent signal controlling vascular development (Jacobs, 1952). It has also been suggested that basipetal regeneration of both xylem and phloem could be quantitatively induced by increasing concentrations of auxin. This hypothesis has been verified by the observations that regeneration of the vascular tissues could be reduced by the application of auxin polar transport inhibitors, such as TIBA (Thompson and Jacobs, 1966). In addition to exploring the inductive eVects of auxin, Sachs has demonstrated that auxin inhibits regeneration of vascular tissues around wounded spots (Sachs, 1969). Following application of auxin to a partially separated section of a pea epicotyl, a new vascular bundle was induced to join it up with the main vascular bundle. However, a newly formed vascular bundle no longer joins with a main vascular bundle when the main vascular bundle is deprived of auxin. These observations support that polar auxin flow is essential for continuous vascular development. A. POLAR AUXIN TRANSPORT

Plant architecture is largely dependent on the establishment of a range of polarities. Biochemical and physiological evidence suggests that the polarities of plant growth are regulated at the cellular level. Plant cells must therefore possess mechanisms that direct proteins or regulatory molecules asymmetrically to specific cell surfaces. Polar auxin transport (PAT) plays a key role in this process. The currently accepted model is based on the basipetal transport of auxin in living plant cells. It is now apparent that PAT is essential for the patterning of procambium and vascular tissues (Mattsson et al., 2003; Sachs, 1981). With the pKa of 4.75, a physiologically active auxin IAA acts as a weak organic acid that undergoes reversible dissociations, the equilibrium of which is pH‐dependent (Fig. 4A). In general, the nondissociated IAA forms can freely penetrate the plasma membrane, while its anion forms resulting from the dissociation process can be transported into the cytoplasm via an active transporter. Outside cells, at pH 5.5, there is always a significant percentage of IAA molecules that are not dissociated (Kramer and Bennett, 2006),

19

VASCULAR DEVELOPMENT

A

O

O O−

OH

B

H+

H+

N

N

IAA

+

IAA- + H+ pH ~5.5

pH ~7

IAA- + H+

IAA

Apical side IAA

PM

Endosomal compartment

PINs

H+

H+

?

Procambial cells

GNOM-dependent endosomes PINOID ? Phosphorylation BFA

Auxins

Basal side AtPINSV

H+

Carrier-driven auxin transport

PINs

Diffusion of auxin molecules

AUX1

PM H+-ATPase (proton pump)

PGPs

Fig. 4. Molecular mechanism of polar auxin transport. (A) Reversible dissociation of IAA. (B) Asymmetric localization of auxin eZux/influx carriers.

which can enter the cells through passive diVusion driven by concentration gradients. However, inside cells, where the cytoplasmic pH is
7.0, IAA dissociation is nearly complete, and the dissociated IAA molecules are present in the form of anions. As such, they can only be excreted actively via an active transporter. If the auxin eZux carriers responsible for IAA anion

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excretion from cells are localized asymmetrically, they would provide the auxin flow with a particular direction (Morris et al., 2004; Swarup et al., 2001; Zazˇ´ımalova´ et al., 2007). The influx carrier proteins, located on the top side of cells, uptake free IAA into the plant cell by both passive and active transports, while the eZux carrier proteins, located on the base, transport IAA anions out of the cell (Fig. 4B). Influx and eZux proteins include a set of plasma‐membrane‐localized proteins, such as PIN, P‐glycoprotein (PGP), and AUX members. PINs and PGP1/PGP19 export auxin from cells, while AUX1 and PGP4 function in auxin influx. Polar localization of these carrier proteins is mediated by vesicle traYcking (Steinmann et al., 1999) and by determinants of polarity and cell fate (Friml et al., 2004; Treml et al., 2005; Xu et al., 2006). Auxin seems to regulate the PIN gene transcription and cellular traYcking and localization of the PIN proteins (Blilou et al., 2005; Vieten et al., 2005). Researches of PAT support a central role for the PIN proteins as auxin eZux carriers. Sequence analysis of the PIN proteins showed that they are transmembrane proteins and may function as transporters. AtPINs also share a limited sequence similarity with some prokaryotic and eukaryotic transporters (Ga¨lweiler et al., 1998; Palme and Ga¨lweiler, 1999). The crucial role of the PIN proteins has been extensively studied in the polar auxin eZux machinery (Friml and Palme, 2002; Morris et al., 2004; Palme and Ga¨lweiler, 1999). Some pin mutants show serious defects in PAT (Okada et al., 1991; Rashotte et al., 2000), and the PIN proteins are localized in cells in a polar manner corresponding to the direction of auxin flow (Friml et al., 2002a,b; Ga¨lweiler et al., 1998; Mu¨ller et al., 1998; Palme and Ga¨lweiler, 1999). Moreover, PAT inhibitors phenocopy the loss‐of‐function pin mutations (Friml et al., 2002a,b; Maher and Martindale, 1980; Okada et al., 1991), and expression of AtPIN2 in yeast cells results in lower accumulation of auxin and structurally related compounds (Chen et al., 1998; Luschnig et al., 1998). All these observations indicate that PINs are potential candidates for auxin eZux carriers. However, the underlying molecular mechanisms remained mostly unknown until recently because of the technical inability to quantitatively assess auxin flow across the plasma membrane in multicellular systems (Zazˇ´ımalova´ et al., 2007). To dissect the molecular function of PINs, cell culture systems derived from the tobacco BY‐2 cell line (Nagata et al., 1992) and Arabidopsis cells were successfully employed (Petra´sek et al., 2006). Heterologous expression of the AtPIN1 protein in a translational fusion with green fluorescent protein (GFP) demonstrated its preferential localization at the transversal plasma membrane (Boutte´ et al., 2006; Petra´sˇek and Zazˇ´ımalova´, 2006). Auxin eZux was shown to be directly proportional to the degree of PIN expression, and

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AtPIN1‐related proteins were sensitive to PAT inhibitors, 1‐naphthylphthalamic acid (NPA) (Petra´sek et al., 2006). In both yeast cells and mammalian HeLa cells, which contain no PIN‐related systems, heterologous overexpression of the AtPIN proteins also resulted in an increase of auxin eZux. All these findings support the direct involvement of PIN proteins in catalyzing the eZux of active auxins and that PINs function as auxin eZux carriers. Although it was known that the polar PIN localization at the plasma membrane well corresponds to the direction of auxin flow (Benkova´ et al., 2003; Friml et al., 2002b, 2003), there was still no clear evidence showing that PINs determine the direction of auxin flow. To investigate this question, variants of PIN1 and PIN2 genes were tagged with haemagglutinin (HA) and/or GFP and expressed under the control of the PIN2 promoter (Wisniewska et al., 2006). The proPIN2::PIN2‐HA exhibited a normal PIN2‐like polar localization in root cortex and epidermal cells, while the proPIN2::PIN1‐HA was detected in epidermal root cells at the side of cells opposite to that of PIN2. The PIN2::PIN1‐GFP constructs also showed opposite localizations within epidermal cells. Only the PIN1‐GFP protein expressed in the correct cellular localization was able to mediate auxin translocation at the lower side of the root after gravity stimulation. These results showed that the PIN polarity is a primary factor determining the direction of auxin flow (Wisniewska et al., 2006), sustaining a central role for PINs in the PAT machinery. However, other experiments suggest that there is a wide functional redundancy among diVerent PIN proteins. For example, most single pin mutants except for pin1 show weak phenotypes, and ectopic expression of PIN proteins was observed in various mutant combinations (Blilou et al., 2005). Some quadruple pin mutations were embryo‐lethal (Friml et al., 2003). In addition, auxin itself positively feeds back on the PIN gene expression in a tissue‐specific manner through an AUX/IAA‐ dependent signaling pathway (Vieten et al., 2005). These results indicate a mechanism by which the loss of a specific PIN protein can be compensated for by auxin‐dependent ectopic expression of its homologues. The functional redundancy of the PIN‐dependent transport network might stabilize the auxin gradients and potentially contribute to the robustness of plant adaptive development. The PGPs are integral membrane ABC transporters that consist of two homologous halves, each containing a nucleotide‐binding fold/ATP‐ hydrolysis site and six transmembrane helices (Ambudkar et al., 1999). Plant PGPs have recently been proposed as probable candidates for the cellular and long‐distance transport of auxin (Geisler et al., 2003; Luschnig, 2002). AtPGP1 catalyzes auxin eZux, while AtPGP4 appears to function in auxin influx. In a recent study utilizing whole plants, protoplasts, and

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heterologous expression systems, AtPGP1 was proven to directly catalyze primary active eZux of IAA, a synthetic auxin 1‐naphthalene acetic acid (1‐NAA), and oxidative IAA breakdown products (Geisler et al., 2005) and that this AtPGP1‐meditated transport was sensitive to inhibitors of auxin eZux and ABC transporters. AtPGP1 was found to be localized in a polar, predominantly basal fashion (Balusˇka et al., 2005) (Fig. 4B). The function of AtPGP19, an auxin‐inducible close homologue of AtPGP1, is similar to that of AtPGP1. The atpgp19 mutant consequently exhibits a dwarfed phenotype and reduced auxin transport. Furthermore, reduction in basipetal auxin transport was observed in the atpgp1 and atpgp19 hypocotyls (Geisler et al., 2003), and the atpgp19 roots exhibited decreased NPA‐sensitivity (Lin and Wang, 2005). The auxin transport defects and dwarf phenotypes were more severe in the atpgp1 atpgp19 double mutants, suggesting an overlapped function between them (Geisler et al., 2003, 2005). AtPGP4 shares a 60% amino acid sequence identity with AtPGP1 (Geisler and Murphy, 2006). The Atpgp4 mutant exhibits reduced basipetal root transport and auxin influx as well as altered free auxin levels (Mancuso et al., 2005; Santelia et al., 2005; Terasaka et al., 2005). Analysis of auxin transport and contents in plants ectopically overexpressing AtPGP4 further suggests that AtPGP4 functions as an auxin uptake sink in the root cap (Terasaka et al., 2005). The coincidence of AtPGP4 and AtPGP1 localizations at opposite ends of cells, even when the orientations are inverted in specific tissues, suggests a pairing of both influx and eZux activities mediated by AtPGP4 and AtPGP1. The AUX family of auxin permeases functions in high‐aYnity cellular auxin uptake. AUX1, the best‐characterized family member, functions in auxin influx and redirection at the root tip (Sidler et al., 1998) (Fig. 4B). Although auxin can be transported inside the cell without a transporter, auxin‐influx carriers support the rapid influx of auxin into cells. AUX1 is asymmetrically localized on one side of the cell and facilitates directional auxin flow (Bennett et al., 1996; Swarup et al., 2001). It has also been shown that AUX1 can act as a bona fide auxin influx carrier in heterologous expression approaches using Xenopus oocytes (Yang et al., 2006) and human HeLa cells (Blakeslee et al., 2007). Genetic and pharmacological studies indicate that reversible phosphorylation is involved in the regulation of PIN‐dependent auxin transport (Benjamins et al., 2001; Friml et al., 2004; Garbers et al., 1996; Shin et al., 2005). The protein serine/threonine (Ser/Thr) kinase PINOID (PID) is the only as yet identified molecular component directly involved in the regulation of polar delivery of the PIN proteins (Benjamins et al., 2001; Christensen et al., 2000). Since AtPIN1 is recycled in cells by endosome‐like AtPIN1‐specific vesicles

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(AtPINSVs), PID might be involved in the asymmetrical transport of the AtPINSVs (Fig. 4B). The phenotype of the pid mutants is similar to that of the pin1 mutant (Bennett et al., 1995). PIN1 localizes to incorrect cellular positions in the pid mutants and in plants overexpressing PID (Friml et al., 2004). These results indicate that PID‐dependent phosphorylation leads to correct PIN localization. In such a scenario, in which the PID kinase is the only known molecular component, some phosphatase(s) could be involved in the reversible phosphorylation of the PIN polar targeting. Genetic analysis, localization assays, and phosphorylation studies demonstrated that PP2A, a protein phosphatase, and PINOID both partially colocalize with PINs and act antagonistically on their phosphorylation during the apical–basal polar targeting (Michniewicz et al., 2007). The GNOM protein plays a fundamental role in regulating endosome‐to‐ plasma membrane traYcking required for polar localization of PIN1. Targeting of PIN1 to the basal plasma membrane requires PIN1 recycling from the plasma membrane to endosomes, which is mediated by GNOM (Geldner et al., 2003; Mayer et al., 1993; Shevell et al., 1994; Steinmann et al., 1999). In the globular embryo of the gnom mutant, individual cells display disrupted polar distribution of PIN1 (Steinmann et al., 1999) and an excess of fragmented vascular structures in the leaves and cotyledons (Shevell et al., 1994), indicating that both basally localized PIN proteins and activity of GNOM are required for the apical‐to‐basal auxin flow. The mislocalization of PIN causes the defect of PAT and thus results in the discontinuity of vascular strands. Inhibitors of protein secretion from eukaryotic cells, such as brefeldin A (BFA), are powerful tools for the PAT studies. Treatments with low concentrations of BFA resulted in defects in growth and development because of altered PAT. Accordingly, PIN1 localization and PAT in the transgenic plants carrying a fully functional but BFA‐resistant GNOM mutant gene were insensitive to BFA (Geldner et al., 2003). These findings provide direct evidence for the specific role of GNOM in traYcking of PIN1 from the endosome to the plasma membrane. GNOM is a member of the large ARF guanine nucleotide exchange factor (ARF‐GEF) family, which regulate vesicle formation by activating ARF‐GTPases on specific membranes (Anders et al., 2008; Bonifacino and Jackson, 2003; Busch et al., 1996; Donaldson and Jackson, 2000; Geldner et al., 2003; Grebe et al., 2000; Kleine‐Vehn et al., 2008; Koizumi et al., 2005; Steinmann et al., 1999). ARF‐GTP is an active form that participates in the formation of transport vesicles. Conversion of ARF‐GTP to ARF‐GDP is necessary for vesicle uncoating, an overture of vesicle fusion with specific target membranes. ARF‐GEFs reversibly associate with specific membranes and thus control the ARF‐GTPase activity spatially and temporarily.

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The vascular tissues are anatomically unique among plant tissues in that the cells need to be precisely connected in order for the tissues to carry out their functions. The uniqueness of the vascular tissue organization is particularly obvious in the highly ordered pattern of veins in the leaves. Arabidopsis leaves develop a hierarchical reticulate venation pattern with formation of a central primary vein, followed by successive basipetal addition of secondary veins and finally higher‐order veins (Kang and Dengler, 2002; Mattsson et al., 1999). Of 266 Arabidopsis accessions, only two exhibited marked diVerences in the vein patterns. This well‐conserved and relatively easily visualized pattern of the veins in Arabidopsis has facilitated the identification of a number of mutants useful for the study of vascular tissue formation (Candela et al., 1999). However, the molecular process by which vein patterns are formed in a developing leaf remains largely unresolved. Sachs (1969) proposed that vein formation is governed by auxin as it flows through the vein. This concept led to the formulation of the canalization hypothesis (Sachs, 1981) (Fig. 4B). The model involves a feedback mechanism, in which diVerentiation increases the ability of cells to transport the signal that induces this diVerentiation. Consequently, the signal inducing vein diVerentiation is transported in a polar manner and is ‘‘canalized’’ into narrow strands (Sachs, 2003), in which at least part of the signals would be auxin (Sachs, 1981). The canalization hypothesis is further supported by experiments showing that leaf vein formation depends on auxin and its polarized transport (Mattsson et al., 1999; Sieburth, 1999). The canalization concept was initially developed through modeling by Mitchison (Mitchison, 1980, 1981). Since the model involves a feedback mechanism, the parameter of the transportation process would vary as a function of auxin flux. The strands that drain auxin from the tissue have high flux but with low concentration of auxin. Given an initial uniform auxin flow, development of these strands can be triggered by small random variations in auxin concentrations or by localized application of auxin at selected points. Strands of increased auxin flux are subsequently diVerentiated into the veins. Mitchison further suggested that there is a spatial pattern of localized sources in the leaf and that this pattern is continually changing as the leaf grows (Mitchison, 1980). This assumption made it possible, in particular, to simulate the formation of vein loops. Recent evidence using molecular markers also supports the localization of auxin sources (Aloni et al., 2003). Despite the experimental evidence and theoretical modeling supporting the canalization hypothesis, it still remains unclear whether canalization can fully

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explain the formation of vein patterns in the leaves. For example, while the canalization hypothesis could account for open branching patterns, it might not be able to account for the network of minor veins (Nelson and Dengler, 1997). Moreover, at the earliest stage of vein patterning, midvein formation becomes acropetal (Scarpella et al., 2004). The acropetal pattern of midvein formation seems in apparent conflict with the direction of auxin flow, which is directed from the distal part of a leaf toward its base (Dengler, 2001). The discovery of mutants with discontinuous venation but with relatively normal vein architecture further expanded the doubt about the canalization (Koizumi et al., 2000). As a result, a reaction–diVusion model has been proposed as an alternative mechanism (Koch and Meinhardt, 1994; Koizumi et al., 2000; Meinhardt, 1976). The reaction–diVusion model is based on the interaction of two or more diVusing substances (Koch and Meinhardt, 1994). Local fluxes of an activator, such as auxin, may trigger a positive feedback loop, in which cells with a high activator concentration would undergo vascular diVerentiation. The diVerentiated cells remove the substrate on which the production of activator depends. As a consequence, diVerentiation shifts the activator maximum to a neighboring cell, which will subsequently diVerentiate. The reaction– diVusion model is also often considered, in a more general sense, as a process in which two or more diVusing substances interact to generate a pattern (Koizumi et al., 2000; Nelson and Dengler, 1997). However, the limitation of the reaction–diVusion hypothesis is that it assumes diVusive transport of activator, whereas activator transport, such as auxin, is polar (Sachs, 1981). To refer to another exception, the PIN1 protein has a polar location at the plasma membrane (Ga¨lweiler et al., 1998; Steinmann et al., 1999). The altered vein patterns observed in the presence of auxin transport inhibitors or in the pin1 mutants suggest that polar transport is essential for the vein pattern formation (Mattsson et al., 1999; Sieburth, 1999). Mutants have been identified in which vein strands are discontinuous. In addition, mutants, whose vein strands are discontinuous but architecture of venation is preserved as in wild type, also provide an argument against the canalization hypothesis. This is because the discontinuity seen in the mutants seems to result from defects in patterning at an early stage rather than from defects in diVerentiation (Carland et al., 1999; Koizumi et al., 2000). Meanwhile, two studies using computer modeling (Rolland‐Lagan and Prusinkiewicz, 2005) or showing localization of PIN1 in mutants with discontinuous but with apparently normal vein patterning support the existence of the canalization mechanism in the mutants with discontinuous vein strands (Scarpella et al., 2006).

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VI. REGULATION OF RADIAL PATTERNING OF VASCULAR BUNDLES The xylem, procambium, and phloem tissues exhibit a distinct dorsoventral organization within the vascular bundles in Arabidopsis. While xylem is localized on the dorsal (adaxial, away from the axis) side, phloem is on the ventral (abaxial, toward the axis) side. The procambium tissue is located between the xylem and phloem tissues (Fig. 5A). Vascular pattern formation is largely conserved within species. Although collateral bundles are most

A

B

Stem

Xy Wild type

Central

Peripheral

Ph Collateral Radialized and abaxialized ex) phb phv rev phan

Adaxial Leaf

Amphicribral Radialized and adaxialized ex) GOF mutants:HD-ZIP IIIs kan1 kan2 kan3 35S::AS2 Amphivasal

Abaxial

C Adaxial miR390

HD-ZIP III

TAS3

SGS3/RDR6/ Translation DCL4 HD-ZIP III ta-siRNA

ZPRs

ZPRs

AGO1

AGO7 miR165/166 Auxin

KAN

AS1/AS2

ETT/ARF4

YAB Abaxial

Fig. 5. Adaxial and abaxial identities and radial patterns of vascular tissues. (A,B) Organization of vascular tissues into collateral bundles in Arabidopsis. (C) Genes involved in the specification of organ polarity interactions. It was modified from Kidner and Timmermans (2007).

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common in seed plants, a number of monocot species have amphivasal bundles, in which the phloem is surrounded by the xylem, or have amphicribral ones in which the phloem surrounds the xylem (Fig. 5B). Patterning of the vascular bundles in the shoot is closely associated with the adaxial– abaxial patterning of lateral organs and with the establishment of central versus peripheral identities within the stem (Fukuda, 2004; Kidner and Timmermans, 2007). Several determinants of the patterning of vascular bundles have been identified, including transcription factors and growth hormones. Among them, mechanisms of the antagonistic regulation of the HD‐ZIP III and KAN transcription factors have been most extensively studied (Fig. 5C). The radial pattern of the root is determined by the action of two transcription factors, SHORT ROOT (SHR) and SCARECROW (SCR) (Di Laurenzio et al., 1996; Helariutta et al., 2000). SHR and SCR are members of the GRAS family of transcriptional regulators (Pysh et al., 1999), and they together regulate the asymmetric cell divisions that take place during the formation of the two cell layers, endodermis and cortex. Asymmetric cell division is the primary mechanism that leads to the diversity of cell types by creating two cell types from one (Ten Hove and Heidstra, 2008). In this section, we mainly describe the vascular patterning associated with intracellular polarity. For in‐ depth analysis of the radial organization that is regulated by asymmetric cell division, we refer the readers to other reviews (Gallagher and Smith, 1997; Ten Hove and Heidstra, 2008). A. OPPOSITE ROLES OF HD‐ZIP IIIs AND KANADIS

Barton group has reported that the Arabidopsis phabulosa‐1d (phb‐1d) mutant induced by EMS causes conversion of the abaxial–adaxial leaf polarity (McConnell and Barton, 1998). This mutant shows diverse phenotypes, such as radialized leaves, trumpet‐shaped leaves, ectopic axillary meristem, amphivasal bundles, and filamentous sepals and petals. The phavoluta‐1d (phv‐1d) mutant is phenotypically similar to the phb‐1d mutant. The PHB and PHV genes encode proteins containing a DNA‐binding homeodomain/ leucine zipper domain (HD‐ZIP) followed by a putative sterol/lipid binding site, designated START (steroidogenic acute regulatory protein related lipid transfer) domain (Ponting and Aravind, 1999). Both the phb‐1d and phv‐1d mutants contain a single nucleotide change within the START domains. These observations suggest that the semidominant gain‐of‐function mutations would disturb steroid‐like ligand binding or abolish the need of such binding, which makes the transcription factors constitutively active (McConnell et al., 2001). It has been found that a miRNA binding sequence is located in the START

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domain and all known mutation sites in the phb and phv dominant mutants reside in the miRNA binding sequence, suggesting that these genes are regulated by miRNA (Rhoades et al., 2002). In Arabidopsis, there are three more genes, such as REVOLUTA/IFL1 (REV), ATHB8, and ATHB15/CORONA (CNA), in the HD‐ZIP III subfamily, which are regulated by miRNA. Single knock‐out mutants of these genes, except for REV, have no visible phenotypes. To avoid the problem related with genetic redundancy, several groups isolated gain‐of‐function alleles and multiple loss‐of‐function mutants. A gain‐of‐function rev‐10d mutant exhibits abnormal vascular patterning with the xylem surrounding the phloem (amphivasal bundle). Unlike the phb and phv dominant alleles, the polarity of leaf and floral organs is unaltered in the mutant (Emery et al., 2003). A loss‐ of‐function rev mutant shows abnormal lateral shoot meristem and flower meristem formation (Otsuga et al., 2001; Zhong and Ye, 1999). They also have defects in vascular development. The rev phb phv triple mutant does not have a functional embryonic shoot apical meristem (SAM) and produces abaxialized cotyledons. The triple mutant also displays amphicribal vascular bundles with the phloem surrounding the xylem, suggesting that these three genes play overlapping roles in regulating SAM formation, leaf polarity, and radial patterning. The cna phb phv triple mutant produces enlarged SAMs and fasciated meristem because of WUS suppression and ectopic amphivasal bundles. Interestingly, the two triple mutants, rev phb phv and cna phb phv, have mutually opposite phenotypes in SAM formation and vascular development. It is now evident that the HD‐ZIP IIIs have common as well as unique functions in controlling meristem identity, lateral organ polarity, and vascular development. The HD‐ZIP III genes regulate the expression domains of the KAN genes, which also regulate the HD‐ZIP III genes. KAN is a putative transcription factor with a GARP domain. In Arabidopsis, there are four KAN genes (KAN1, KAN2, KAN3, and KAN4). By in situ hybridization assays, the KAN expression was detected throughout the early globular embryo but limited to the abaxial side of the cotyledon primordia in the heart stage. After germination, localization of the KAN gene expression is restricted to the phloem of root and abaxial side of young leaf (Emery et al., 2003; Kerstetter et al., 2001). The expression of KAN2 and KAN3 is restricted to the phloem throughout the plant life. Notably, these expression patterns are complementary to those of the HD‐ZIP III genes. The loss‐of‐function kan mutant produces trichomes on the abaxial side of first two leaves. Abaxial mesophyll cells are moderately altered and have morphology similar to the adaxial mesophyll cells in the mutant. The kan1 kan2 double mutant exhibits conversion of the abaxial–adaxial polarity, and

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the expression domains of the HD‐ZIP III genes are expanded to the abaxial side in the mutant (Eshed et al., 2001). The kan1 kan2 kan3 triple mutant shows amphivasal vascular bundle which is indistinguishable from the rev‐ 10d vascular bundle (Emery et al., 2003). Ectopic expression of the KAN genes results in narrow cotyledons, arrested meristems, and abaxialized leaves. This mutant has no vascular bundles within the cotyledons and hypocotyl, while the gain‐of‐function mutants of PHB, PHV, and REV have amphivasal bundles. Furthermore, PHB is not expressed in the adaxial region of the KAN‐overproducing mutant leaves (Eshed et al., 2004). Conversely, KAN expression is greatly reduced in the phb‐1d mutant. Together, these observations propose that the HD‐ZIP III and KAN genes act antagonistically to regulate vascular polarity and other related plant organ development (Fig. 5B and C).

B. REGULATION OF HD‐ZIP III ACTIVITIES BY COMPETITIVE INHIBITORS

Recently, two research groups independently have reported a novel gene family, collectively designated LITTLE ZIPPERs (ZPRs) (Kim et al., 2008; Wenkel et al., 2007). This gene family includes four members, such as ZPR1, ZPR2, ZPR3, and ZPR4, which encode small proteins, consisting of 136, 105, 67, and 72 amino acids, respectively. The ZPR proteins are structurally unique in that they contain a leucine zipper (ZIP) motif similar to those found in the HD‐ZIP III proteins. The HD domain mediates DNA binding, and the ZIP motif participates in the dimerization of the HD‐ZIP III proteins (Ariel et al., 2007). Based on the structural similarity of the ZPR and HD‐ZIP proteins, it has been predicted that they might physically interact with one another. As predicted, it has been proven in vivo and in vitro that the ZPR proteins bind to the HD‐ZIP III proteins via the ZIP motifs, forming nonfunctional heterodimers. In contrast, they do not interact with other HD‐ZIP proteins (Kim et al., 2008). It has been shown that this interaction prevents the DNA binding activity of the REV protein (Wenkel et al., 2007) and the transcription activation activity of the PHB protein (Kim et al., 2008), providing a regulatory mechanism, which we call small interfering peptide (siPEP). A similar mechanism has been previously reported in both animal and plant cells. The inhibitor of DNA binding (ID) protein contains a helix‐loop‐helix (HLH) domain mediating protein–protein interaction but lacks a basic domain involved in DNA binding (Benezra et al., 1990). The ID protein binds to several bHLH transcription factors and inhibits their DNA binding by forming nonfunctional heterodimers. In addition, a plant HLH‐containing

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protein, KIDARI (KDR), has been proposed to regulate a set of plant bHLH transcription factors in a similar manner (Hyun and Lee, 2006). Notably, the ZPR3‐overproducing mutant, zpr3‐1d, is phenotypically similar to the mutants in which HD‐ZIP III expression is reduced. The zpr3‐1d mutant exhibits partially abaxialized leaves, fewer secondary shoots, and severely arrested SAM. In addition, the phb‐1d phenotype is recovered at least in part in the zpr3‐1d background, indicating that the ZPR3 protein negatively regulates the PHB activity in planta (Fig. 5C). The loss‐of‐function zpr3‐1 zpr4‐2 double mutant exhibits an array of phenotypic changes, including extra cotyledons, extra leaf formation at random positions, and SAM malformation. In this mutant, expression of the CLV3 gene increases by more than 20 times. The phenotypes of the double mutant are similar to those of phb‐1d and moderately recovered in the zpr3‐1 zpr4‐2 phv triple mutant (Kim et al., 2008). All these observations strongly support that the ZPR protein negatively regulates PHV and other HD‐ZIP III members via siPEP. Interestingly, REV and PHB positively regulate ZPR3 expression (Kim et al., 2008; Wenkel et al., 2007). It has thus been proposed that the HD‐ZIP III activity is coordinately modulated by a feedback loop involving competitive inhibitors ZPR3 and its functional orthologues in regulating SAM development in conjunction with miR165/166, demonstrating a novel way of regulating SAM development (see below). C. OTHER ADAXIAL–ABAXIAL AXIS DETERMINANTS

Mutations in ETTIN/AUXIN RESPONSE FACTORS3 (ETT/ARF3) have been isolated from a genetic analysis using the APETALA3 promoter‐ mediated ectopic expression of KAN in search of factors involved in the KAN‐mediated signaling (Pekker et al., 2005). Double mutants with ETT and its closely related gene ARF4 exhibit transformation of abaxial tissues into adaxial ones in all aerial plant parts, resembling mutations in the kan1 kan2 double mutant. In addition, both ETT and ARF4 are abaxially expressed (Fig. 5C). However, KAN is not required for both ETT and ARF4 transcriptions, and overexpression of ETT or ARF4 cannot rescue the kan mutants (Pekker et al., 2005). Therefore, KAN and ETT‐ARF4 seem to function cooperatively in abaxial cell specification and in mediatin101uxin signaling during the adaxial–abaxial patterning but act independently in diVerent signaling pathway (Fig. 5C). The function of the abaxial determinants ETT and ARF4, as well as the KAN genes, is opposed by adaxial determinants, such as ASYMMETRIC LEAVES1 (AS1) and AS2. The AS1 and AS2 genes act antagonistically to ETT and ARF4 in the regulation of downstream polarity genes (Garcia et al., 2006;

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Li et al., 2005; Xu et al., 2006). AS1 is an Arabidopsis orthologue of PHANTASTICA (PHAN) from Antirrhinum, the first gene demonstrated to be involved in the control of leaf polarity (Waites and Hudson, 1995; Waites et al., 1998). The as1 mutants show no clear polarity defects, neither do the as1 as2 double mutations. However, 35S::AS2 transgenic lines show defects in leaf polarity; the leaves of the transgenic plants are adaxialized (Lin et al., 2003) (Fig. 5B). AS1 and AS2 transcripts were detected in the adaxial domain and in the inner domain between the adaxial and abaxial domains of the leaves, respectively (Iwakawa et al., 2007). Ectopic expression of AS2 to a low level under the control of the AS1 promoter in the as2 mutant plants restored an almost normal phenotype, while strong expression of AS2 in wild‐type and in the as2 mutant, but not in the as1 mutant, showed narrow, upwardly curled leaves. These results indicate that AS2 represses cell proliferation in the adaxial domain in the presence of AS1 and that the appropriate expression of AS2 in adaxial part is critical for the symmetric development. Moreover, mutations of either AS2 or AS1 resulted in an increase in the transcriptional level of ETT/ARF3 and KAN2, which promote abaxial identity, and YABBY5 (YAB5). Thus, AS2 and AS1 might negatively regulate the expression of these genes in the adaxial domain, which might be related to the development of flat and expanded leaves (Iwakawa et al., 2007) (Fig. 5C). The antagonistic interactions between the adaxial and abaxial determinants might be important in maintaining the leaf polarity after growth away from the meristem. Intriguingly, the ETT‐ARF4, AS1‐AS2, and the HD‐ZIPIII‐KAN signals all converge to regulate the YAB genes (Kidner and Timmermans, 2007). The YAB gene family encodes six closely related transcription factors, three of which, including FILAMENTOUS FLOWER (FIL), YAB2, and YAB3, are expressed during vegetative development (Siegfried et al., 1999). The YAB genes contribute to abaxial cell identity, and the polarized expression directs lateral outgrowth at the adaxial–abaxial boundary. All the KAN, ETT, and ARF4 genes positively regulate YAB expression in the abaxial domain of the leaf, whereas AS1 and AS2 repress the YAB genes in the adaxial domain (Eshed et al., 2004; Garcia et al., 2006; Iwakawa et al., 2007; Li et al., 2005; Lin et al., 2003; Xu et al., 2006). Small interfering RNAs also play important roles in the adaxial–abaxial patterning (Fig. 5C). Both ETT/ARF3 and ARF4 mRNAs are targets of RNA interference (RNAi). It has been shown that ETT/ARF3 and ARF4 mRNAs are regulated by trans‐acting short‐interfering RNAs (ta‐siRNAs) (Allen et al., 2005; Williams et al., 2005a). There are three ta‐siRNAs in Arabidopsis: TAS1, TAS2, and TAS3. TAS transcripts are in turn regulated by miRNA‐directed cleavage, resulting in ta‐siRNAs of 21 base pairs (Allen et al., 2005). For example, ETT and ARF4 mRNAs are targets of

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TAS3‐derived ta‐siRNAs (Allen et al., 2005; Williams et al., 2005a). The leaf defects observed in the mutants overexpressing ta‐siRNA‐resistant ETT alleles are similar to the leaf phenotypes caused by mutations in the genes mediating ta‐siRNA biogenesis (Adenot et al., 2006; Fahlgren et al., 2006; Hunter et al., 2006; Peragine et al., 2004).

VII. IDENTIFICATION OF VASCULAR DEVELOPMENT‐RELATED FACTORS A. HD‐ZIP IIIs AND miRNA

In Arabidopsis, procambium precursor cells can be noticeable during the transition from the globular to heart stages (Baucher et al., 2007). Transcripts of PHB, PHV, REV, and ATHB15/CNA are detected in the adaxial regions of embryos, including procambial cells which appear from the heart/torpedo stages. ATHB8 is the only HD‐ZIP III gene that is not expressed in the apical half of the globular embryo. Expression pattern of PHV is similar to those of PHB and REV, but the expression level is relatively lower in the pre‐ procabium cells (Emery et al., 2003). PHB and PHV gain‐of‐function mutants show amphivasal bundles in the leaves, with the xylem surrounding the phloem, like rev‐10d (Baucher et al., 2007). The REV expression is detected within the developing pre‐procambial cells in the hypocotyls and developing vascular tissues (Otsuga et al., 2001). The rev‐6 mutation aVects interfascicular fiber diVerentiation. While the mutant is characterized by having less interfascicular fibers in the Col‐0 background, formation of the interfascicular fibers is largely unaltered in the Ler background (Otsuga et al., 2001). Vascular defects of rev‐6 are exaggerated by the phb‐13 and phv‐11 mutations but suppressed in the cna‐2 athb8‐11 double mutant. Histological analysis has shown that ATHB8 is expressed in the procambial cells located between the xylem and the phloem components. The expression of ATHB8 promoter‐GUS fusion (pATHB8::GUS) was also similarly localized in the procambium of cotyledons and roots (Baima et al., 1995). The gain‐of‐function ATHB8 mutation induces more xylem formation and lignin synthesis than wild‐type plants because of the increased activity of the fascicular and interfascicular cells (Baima et al., 2001). Procambial cells are diVerentiated to xylem and phloem components. However, ectopic expression of ATHB8 promotes only xylem development. Meanwhile, ZeHB10, an ATHB8 gene homologue in Zinnia, is expressed more strongly in the xylem precursor cells rather than in the procambial

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cells, showing that ATHB8 regulates xylem diVerentiation rather than procambium development. ATHB8 and its Zinnia homologue ZeHB10 are expressed in young leaves and roots. ATHB15/CNA and ZeHB13 also exhibit similar expression domains, but they are expressed more specifically in the procambium (Ohashi‐Ito and Fukuda, 2003). The athb15/cna single mutant shows slightly irregular vascular bundles (Prigge et al., 2005). Overexpression of a miRNA‐resistant ATHB15 gene (mATHB15) reduces vascular cell diVerentiation. Based on these observations, it has been proposed that ATHB15/CNA and ATHB8 may have antagonistic roles in vascular development (Kim et al., 2005). However, the cna phb phv and rev phb phv triple mutants exhibit mutually opposed vascular patterns, suggesting that the signaling networks governing the HD‐ZIP activities in vascular development would be more complicated than previously expected. The miR165/166 binding site is located within the START domain of the HD‐ZIP III genes (Rhoades et al., 2002; Tang et al., 2003). All the gain‐of‐ function phb and phv mutants identified so far contain single nucleotide substitutions in the microRNA (miRNA) binding sites. In these mutants, PHB and PHV are expressed on both adaxial and abaxial sides of the leaves (McConnell et al., 2001). An Arabidopsis mutant overproducing miR166 shows severe reduction of ATHB15/CNA and PHV genes (Kim et al., 2005). The expression levels of PHB and PHV are also dramatically reduced in a similar miR166‐overproducing mutant (Williams et al., 2005b). The distribution of miR166/165 is restricted to the embryonic meristem and the abaxial side of the leaf (Fig. 5C). It is therefore likely that miR165/166 regulates the expression domains of the HD‐ZIP III genes (Rhoades et al., 2002). In other words, miR165/166 may cleave the HD‐ZIP III mRNAs in the abaxial side. However, this notion is not suYcient to explain all experiment data (Bao et al., 2004). Especially, promoter analysis of the HD‐ZIP III genes using the GUS reporter system reveals that these genes are expressed where their transcript levels are detected (Baima et al., 1995; Kang et al., 2002). In addition, their expression patterns are not identical, even though their mRNAs all contain the miRNA binding sequences. Bao et al. proposed a modified model in which PHB mRNA level is reduced by miR165/166 not only in the abaxial region but also in the entire leaf. Extensive analyses of the expression patterns and domains of the HD‐ZIP III, MIR165, and MIR166 genes would be useful to confirm the model. The Arabidopsis genome contains two MIR165 genes and seven MIR166 genes. Notably, the expression patterns of the MIR166/165 genes are diverse, and they are regulated spatially and temporally (Jung and Park, 2007), strongly supporting the model.

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J.‐H. JUNG ET AL. B. CYTOKININ

Cytokinin is essential for the formation and maintenance of procambial cells (Aloni, 1987). The Arabidopsis wooden leg (wol) mutant was originally defined as a recessive mutation that showed reduced cell division in the procambium and abnormal diVerentiation of all vascular cells into protoxylem within the root (Scheres et al., 1995). The absence of phloem in the primary root of the wol mutant was apparently due to reduced division of procambial cells during embryogenesis (Ma¨ho¨nen et al., 2000; Scheres et al., 1995). The vascular cylinder of Arabidopsis primary root consists of both protoxylem and metaxylem. In the wol mutant, however, it is exclusively made of protoxylem (Can˜o‐Delgado et al., 2000). It was later shown that WOL is identical to CYTOKININ RESPONSE1 (CRE1) and ARABIDOPSIS HISTIDIN KINASE4 (AHK4) that encode a putative histidine kinase (HK), which is believed to function as a cytokinin receptor (Inoue et al., 2001; Ma¨ho¨nen et al., 2000; Suzuki et al., 2001). WOL/CRE1/AHK4 works via the phosphorylation cascade upon ligand binding (Inoue et al., 2001). Cytokinin is bound to the receptor, and the perceived signals are transferred to downstream genes through the phosphorylation cascade and finally regulate procambial cell proliferation. The WOL gene is expressed preferentially in the precursors of the procambium and in the procambium (Ma¨ho¨nen et al., 2000). These observations indicate that cytokinin plays a role in maintaining the procambial activity via the WOL/CRE1/AtHK4 cytokinin receptor. There are two additional cytokinin receptor genes, AHK2 and AHK3, in Arabidopsis (Inoue et al., 2001; Ma¨ho¨nen et al., 2000). The cre1 ahk2 ahk3 triple mutant, as well as the mutant ectopically expressing cytokinin‐ degrading CYTOKININ OXIDASE (CKX) under the pCRE1 promoter, phenocopies the wol phenotype (Ma¨ho¨nen et al., 2006b). It seems likely that cytokinin signaling is necessary for vascular diVerentiation. The exclusive phenotype of protoxylem diVerentiation might be induced simply by up‐ regulating the CKX expression during the post‐embryonic development, as has been observed in the wol mutant, even when cell proliferation in the vascular cylinder is nearly normal. This result suggests that cytokinin plays a role in cell specification; while protoxylem is the default cell fate in the absence of cytokinin signaling, and cytokinin signals are required to change this default identity (Ma¨ho¨nen et al., 2006b). Whereas the cre1 ahk2 ahk3 triple mutant phenocopies the wol phenotype, all the single and double mutants show essentially normal vascular patterning (Higuchi et al., 2004; Ma¨ho¨nen et al., 2006b). By searching for suppressors of the determinate root growth habit of wol, several intragenic suppressors were identified, including those encoding premature nonsense proteins (Ma¨ho¨nen

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et al., 2006b). Vascular organization of these mutants homozygous for a putative truncated CRE1 gene was essentially normal, indicating that root vascular morphology of the wol mutant reverts to wild type (Ma¨ho¨nen et al., 2006b). Similarly, the vascular morphology of the cre1‐2 and cre1‐1 mutants was normal (Higuchi et al., 2004; Inoue et al., 2001; Ma¨ho¨nen et al., 2006b). In addition, the various nonsense suppressor mutations eliminate a dose‐ dependent negative activity on vascular cell proliferation during root development. These observations indicate that wol seems to be an exceptional type of mutation that can overcome the eVect of the two antagonistically acting loci. The molecular nature of this negative activity was shown to be a phosphatase activity. When the wol cre1 mutant was complemented with a modified CRE1 gene (T278I) under the control of its own promoter, which lacks cytokinin binding activity and is thus locked in a phosphatase form, higher cytokinin responsiveness was achieved (Ma¨ho¨nen et al., 2006b). Therefore, it is evident that WOL is a bifunctional kinase/phosphatase; WOL is not only a kinase that phosphorylates phosphotransfer intermediate substrates in the presence of cytokinin but also a phosphatase that dephosphorylates the phosphotransfer intermediate substrates in the absence of cytokinin (Ma¨ho¨nen et al., 2006b). In Arabidopsis, immediate early responses of plants to cytokinin signals have been formulated as a multistep process; the binding of cytokinins induces autophosphorylation of the receptor HKs and subsequent transfer of the phosphoryl group to a histidine‐containing phosphotransfer protein (HPt) and to a response regulator (RR), ultimately regulating downstream signaling events (Hwang and Sheen, 2001; Kakimoto, 2003). The cytokinin signal is mediated by a His–Asp phosphorelay mechanism, which is exerted through sequential phosphotransfer between His and Asp residues in each component. Arabidopsis has five genes that encode histidine‐containing phosphotransfer proteins (AHP1–AHP5) (Suzuki et al., 2002). Single ahp mutants did not display any altered cytokinin sensitivity or any obvious eVects on plant growth and development. However, some double and higher‐order ahp mutants showed increasing resistance to cytokinin in various assays, indicating that there is functional overlapping between the members of the family and that these AHPs act as positive regulators of cytokinin signaling (Hutchison et al., 2006). It is now apparent that AHP1, AHP2, AHP3, and AHP5 each act as a positive regulator of cytokinin signaling in a partially redundant manner, while AHP4 may act as a negative regulator. The ahp1,2,3,4,5 quintuple mutant showed abnormal phenotypes in vascular development by having short primary root and reduced vascular cylinder, as observed in the wol mutant (de Leon et al., 2004; Hutchison et al., 2006;

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Ma¨ho¨nen et al., 2000; Scheres et al., 1995). In addition, xylem vessels, but no phloem vessels, were visible in the transverse sections of the ahp1,2,3,4,5 primary roots. However, the exclusive protoxylem development, another striking aspect of the wol mutant phenotype, was not observed in the quintuple mutant. On the contrary, while asymmetry is observed in the wild‐type vasculature, the protoxylem that was visible in the primary root of the ahp1,2,3,4,5 mutants formed a central cylinder of vessels that displayed radial symmetry (Hutchison et al., 2006). Two extragenic second‐site suppressor mutations at the AHP6 locus resulted in reduction of the exclusive protoxylem fate, characteristic of the wol mutant (Ma¨ho¨nen et al., 2006a). Single ahp6 mutant shows a similar phenotype, which sporadically lacks protoxylem specification. The AHP6 locus encodes a ‘‘pseudo’’ phosphotransfer intermediate protein, which lacks the conserved histidine residue critical for phosphorelay. Consequently, AHP6 acts as an inhibitor rather than as a positive regulator for the phosphotransfer. This indicates that the presence of AHP6 enables protoxylem specification by down‐regulating cytokinin signaling. Accordingly, AHP6 is expressed at a spatially specific manner in protoxylem and adjacent pericycle cell lineages (Ma¨ho¨nen et al., 2006a). This expression pattern during protoxylem development is also consistent with the inhibitory role of AHP6 in cytokinin signaling. There are 22 ARRs (ARR1–ARR22) in Arabidopsis, which are classified as two distinct types: type A, including 10 members and type B, which has 11 members. The type‐B ARRs function as transcriptional activators. In contrast, the type‐A ARR genes are induced rapidly by cytokinin, but the corresponding proteins do not have DNA‐binding activities and thus does not function as transcriptional regulators (Hwang and Sheen, 2001; Sakai et al., 2001). In the cre1 roots, cytokinin induces all the type‐A ARR genes, except for ARR15 and ARR16 (Kiba et al., 2002). In wild‐type roots, cytokinin induces ARR15 in the procambial cells but not in the xylem and phloem cells. ARR16 is induced in the pericycle. In addition, ARR15 was found to function as a repressor of the type‐B ARRs (Kiba et al., 2003). Because a loss‐of‐function mutant of ARR15 shows no apparent phenotype, ARR genes might be functionally redundant. Moreover, the arr1 arr10 arr12 triple mutant showed phenotypes displaying a very poor growth, quite similar to those of the wol mutant that virtually lacks cytokinin receptor activities. In this triple mutant, specification of root vascular (i.e., protoxylem specification) tissues is also aVected as severely as in wol. ARR10 and ARR12, together with ARR1, redundantly play pivotal roles in the AHK‐dependent phosphorelay signaling in response to cytokinin in roots (Yokoyama et al., 2007).

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C. BRASSINOSTEROID

Critical roles of BRs have been confirmed by observations of reduced xylem development and increased phloem development in a series of Arabidopsis mutants in which several BR receptor kinase genes, including BRASSINOSTEROID INSENSITIVE1 (BRI1), BRL1, and BRL3, are disrupted (Can˜o‐ Delgado et al., 2004). BRL1 and BRL3 were identified based on their sequence similarities to that of the previously identified BRI1 receptor, a leucine‐rich repeat receptor kinase that directly binds BR (Kinoshita et al., 2005; Wang et al., 2001). The BRI1 gene is ubiquitously expressed in dividing and elongating cells, whereas the BRL1 and BRL3 genes are expressed predominantly in the vascular tissues (Can˜o‐Delgado et al., 2004). BRL1 expression is intimately associated with the procambial cells of vascular bundles, whereas BRL3 expression is specifically localized in the phloem of cotyledons and leaves but not in stems. The brl1 mutant has an increased number of phloem cells and a decreased xylem diVerentiation in the vascular bundles. Furthermore, the bri1 brl1 brl3 triple mutants enhance the bri1 dwarfish appearance. They also exhibit abnormal vascular diVerentiation. These results suggest that signals through the BR receptors in the procambium induce xylem proliferation while simultaneously repress phloem proliferation. BR deficient mutants show abnormal vascular patterning, including increased amounts of phloem and decreased amounts of xylem (Choe et al. 1998, 1999; Mathur et al., 1998; Szekeres et al., 1996). Application of brassinazole, a specific BR biosynthesis inhibitor, further supports the role of BR in vascular development. Brassinazole induces excess formation of phloem and reduced formation of xylem in Arabidopsis (Nagata et al., 2001). The Zinnia system greatly facilitated analysis at a cellular level of the mechanisms by which BR regulates TE diVerentiation (Fukuda, 1997; Fukuda and Komamine, 1980). Inhibition of BR biosynthesis using uniconazole, another BR biosynthesis inhibitor, prevents the diVerentiation of procambium‐like cells into TEs without inhibiting cell division in the Zinnia culture system (Iwasaki and Shibaoka, 1991; Yamamoto et al., 1997). Subsequent application of exogenous brassinolide rescued its inhibitory eVect. Uniconazole appears to block the later stages of TE diVerentiation and prevents the expression of genes associated SCW deposition and PCD (Yamamoto et al., 1997). In the Zinnia system, such BR biosynthesis inhibitors do not suppress expression of the genes functioning during stages 1 and 2 of xylem cell diVerentiation but suppress the expression of most genes that function in stage 3. This suggests that endogenous BR might be necessary during the transition from stage 2 to stage 3 (Fukuda, 2004). Indeed, BR biosynthesis

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increases significantly at stage 2 when procambium‐like cells are produced by diVerentiation (Yamamoto et al., 1997, 2001). These observations strongly support that BR is involved in diVerentiation of parenchyma cells into TEs. BR up‐regulates the expression of ZeHB12, a Zinnia REV homologue. In addition, an additional HD‐ZIP III gene ZeHB10, an Arabidopsis ATHB8 gene homologue, is mostly expressed in the xylem cells and induced by BR (Ohashi‐Ito et al., 2002). In accordance with their BR induction, the accumulation of transcripts for ZeHB10, ZeHB11, and ZeHB12 was almost completely suppressed by uniconazole, and the suppressive eVects were eYciently reversed by addition of brassinolide, a biologically active BR form (Ohashi‐Ito et al., 2002). Notably, the expression of ZeHB13, an ATHB15 gene homolog, was restricted to the procambium and was not severely suppressed by brassinazole, unlike ZeHB10, 11, and 12 (Ohashi‐Ito and Fukuda, 2003). A histochemical promoter analysis using the pATHB15:: GUS transgenic Arabidopsis plants indicated that ATHB15 was also active specifically in the procambium. These observations strongly support that ZeHB13 and ATHB15 are pivotal transcriptional regulators responsible for early stages of vascular development (Ohashi‐Ito and Fukuda, 2003). It is also apparent that BR might regulate procambial cell diVerentiation to xylem by modulating the HD‐ZIP‐III gene expression. Transgenic Arabidopsis plants harboring CaMV 35S promoter‐driven expression of ZeHB10 and ZeHB12 with mutations in the START domains (mtZeHB10 and mtZeHB12, respectively) showed a higher‐level production of TEs and xylem precursor cells, respectively. In addition, the expression of BRL3 and BRI‐ASSOCIATED RECEPTOR KINASE 1 (BAK1) was up‐regulated in the Arabidopsis plants overexpressing ZeHB12. These observations further support the role of BR in the regulation of xylem diVerentiation (Ohashi‐Ito et al., 2005). D. XYLOGEN

Formation of continuous vascular strands through vascular cell diVerentiation has raised the possibility that some inductive signals would function in guiding vascular continuity. Such inductive signals have not yet been defined, mainly because of the diYculties in analyzing cell–cell interactions in planta. The in vitro Zinnia culture system provides an excellent experimental tool for directly analyzing cell–cell interactions and for isolating inductive signals (Fukuda, 1997; Fukuda and Komamine, 1980). Previous studies with this Zinnia culture system revealed that diVerentiation of TEs tends to promote diVerentiation of neighboring cells (Motose et al., 2001a). This inductive cell‐ to‐cell interaction was found to be mediated by a secretary factor acting

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locally, which was named xylogen. The fraction of arabinogalactan proteins (AGPs), containing a group of plant proteoglycans was found to contain a key component of xylogen that mediates local intercellular communication in TE diVerentiation of the cultured Zinnia cells (Motose et al., 2001b). The purified xylogen from the AGP fraction increased the frequency of TE diVerentiation, whereas chemically deglycosylated xylogen did not, indicating the requirement of glycosyl chains for the xylogen activity (Motose et al., 2004). Notably, xylogen derived from the dicot Zinnia could also promote the TE diVerentiation from culture cells of the monocot Asparagus. Xylogen synthesized heterologously in tobacco cells also promoted TE diVerentiation in the Zinnia cell system. These results suggest that xylogen is not species specific but common to diverse angiosperms. SDS‐PAGE analysis of the purified xylogen yielded a broad band of molecular mass in a range of 50–100 kDa, whereas deglycosylated xylogen showed a sharp band of 16 kDa (Motose et al., 2004). The purified protein (ZeXYP1) contains an N‐glycosylation site, a signal peptide, and a putative glycosylphosphatidylinositol (GPI) anchor, which are all characteristics of arabinogalactan proteins (AGPs). Analysis in silico indicated that ZeXYP1 is a hybrid‐type molecule with properties of both AGPs and nonspecific lipid‐transfer proteins (Kader, 1996; Motose et al., 2004). Two homologous Arabidopsis genes, At5g64080 and At2g13820, were identified by a BLAST search using the ZeXYP1 as a query and designated A. thaliana xylogen protein 1 (AtXYP1) and AtXYP2, respectively. Neither of the single mutants showed a distinct phenotype, but the double mutant has serious defects in the continuity of the vascular strands. Intriguingly, xylogen preferentially accumulates at only one end of the cell. The xylogen genes are abundantly expressed in procambium and immature xylem cells. The corresponding proteins were immunohistochemically detected in the apical side of the cell walls of diVerentiating xylem cells, suggesting that the immature cells may secrete the gene product directionally to the neighboring cells to regulate xylem diVerentiation. Taken together, these data suggest that xylogen is necessary for vascular development as a mediator of inducing signals and thus its polar secretion contributes to the continuity of the vascular network by promoting diVerentiation of the neighbor cells (Motose et al., 2004).

E. OTHER GENES INVOLVED IN VASCULAR DIFFERENTIATION

Bonke et al. (2003) has identified a gene, APL, encoding a member of the MYB coiled‐coil‐type transcription factor family. This gene is necessary for the determination of phloem identity in Arabidopsis. Most of the work was

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done with the root, which in Arabidopsis develops a very typical cellular organization with a central xylem axis and two phloem poles. In the apl mutant, cells at the phloem poles develop characteristics of xylem cells like TEs (Bonke et al., 2003). Using the markers for SEs and CCs, it was verified that phloem‐specific cell types are absent in the apl mutant. To examine whether the defects in cell diVerentiation are dependent on defects in the phloem‐related divisions, a fass ( fs) mutation was introduced into the apl background. The fs mutant undergoes excessive numbers of cell divisions in all tissues and rescues the wol mutant (Torres‐Ruiz and Jurgens, 1994). The fs apl double mutant phenotype was additive. More vascular cells are present, but TE‐like cells are still found in the position of phloem (Bonke et al., 2003). These observations suggest that APL is required for the asymmetric cell divisions as well as for SE and CC diVerentiation during phloem development and that other unknown factors promoting these divisions would be present in addition to APL. APL is expressed in the protophloem and, subsequently, in CCs and metaphloem. Interestingly, APL expression initiates after asymmetric divisions that lead to the specific cell types of the phloem poles. In mature embryos, its expression is detected in the prospective cells that are later diVerentiated into phloem tissues in cotyledons and hypocotyls during post‐embryogenesis. At later stages of plant development, APL is expressed in the phloem of all organs (Bonke et al., 2003). When APL was expressed ectopically throughout the procambium under the control of the WOL promoter, cells close to the root tip that normally diVerentiates into protoxylem remained undiVerentiated. The contrasting nature of the apl phenotype with TE‐like cells in positions normally occupied by phloem and the proWOL::APL phenotype with undiVerentiated cells in the protoxylem position together suggest that, in addition to specifying the phloem identity, APL may also be required for inhibiting xylem diVerentiation in the phloem poles during vascular development (Bonke et al., 2003). Vascular meristems participate in a highly ordered developmental process with a very prominent polarity. This polarity results in precisely orientated divisions of meristematic initials that produce highly specialized and spatially separated xylem and phloem cells. The factors that are necessary to establish and maintain this polarity remain unclear (Traas and Bohn‐Courseau, 2005; Williams and Fletcher, 2005). Insights into vascular bundle patterning have come from the study on the phloem intercalated with xylem ( pxy) mutant in Arabidopsis (Fisher and Turner, 2007). In the pxy mutant, phloem appears in ectopic positions and intermixed with xylem, and thus the shape of vascular bundles is flat instead of having the triangular shape. The mutant vascular bundles include similar

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cell numbers as in those of wild type, and they retain procambium. On the contrary, at least one cell type, metaxylem, is reduced, and veins appear to be poorly associated along the length of the stem in the pxy mutant. These results imply that there is a primary defect in the process of vascular cell diVerentiation (Fisher and Turner, 2007). More detailed analysis showed that procambial cell division patterns are disrupted in the pxy mutant. Since procambial cells are located tightly between phloem and xylem in the vascular bundles, procambial cell divisions along the phloem‐xylem axis cause linear cell files separating xylem and phloem. Instead, the procambial cells in the pxy mutants are located adjacent to the phloem and within a deeper region of the xylem. These pxy phenotypes suggest that the primary role of PXY is to maintain the appropriate orientation of procambial cell divisions (Fisher and Turner, 2007; Sieburth, 2007). PXY encodes a leucine‐rich repeat (LRR) receptor‐like kinase (Fisher and Turner, 2007). There is a large family consisting of such kinases in Arabidopsis, with more than 200 members in 15 diVerent subfamilies, yet their functions are known for only a small fraction of them (Shiu and Bleecker, 2001). The PXY‐like genes, PXL1 and PXL2, appear to function redundantly with PXY. Single pxl1 or pxl2 mutant does not exhibit an obvious phenotype in the stem. However, double mutants generate a more severe vascular phenotype than the pxy mutant, suggesting that PXL1 and PXL2 function redundantly with PXY. Furthermore, PXY and its functional orthologues are also closely related to CLAVATA1 (CLV1), BAM1, BAM2, and BAM3, which encode LRR receptor‐like kinases (DeYoung et al., 2006; Clark et al., 1997). CLV1 limits the number of cell divisions in the SAM, whereas BAM1, BAM2, and BAM3 appear to promote cell proliferation in the SAM. If PXY functions as an LRR receptor‐like kinase, there would be ligands that directly bind to PXY. Attractive candidates are the CLE peptides (Cock and McCormick, 2001). The Arabidopsis genome contains 31 CLE genes that encode 26 diVerent CLE peptides, including the putative CLV1 ligand, CLV3 (Cock and McCormick, 2001; Fletcher et al., 1999; Ito et al., 2006; Kondo et al., 2006; Sharma et al., 2003). Other classes of peptide hormones also bind LRR receptor‐like kinases and thus are also candidates for being the PXY ligand (Matsubayashi and Sakagami, 2006). Several genes have been identified to function in xylem tissue diVerentiation by using the microarray analysis on the Zinnia culture cells (Demura et al., 2002; Milioni et al., 2002). These microarray data include the genes encoding NAC transcription factors, which are up‐regulated within 30 min after induction of Zinnia cells. To gain an expression profile of xylem cell diVerentiation‐related genes in Arabidopsis, Kubo et al. (2005) established an

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in vitro xylem vessel element inducible system from Arabidopsis suspension cells. They monitored gene expression in a global manner and observed that several NAC transcription factors are highly expressed during xylogenesis. The corresponding proteins have been termed vascular‐related NAC‐domain (VND) proteins. There are seven VND proteins, VND1 to VND7, up‐regulated during the in vitro formation of xylem vessel elements. When ectopically overexpressed in Arabidopsis or Populus under the control of the CaMV 35S promoter (35S), only VND6 and VND7 could induce transdiVerentiation of various types of cells into xylem vessel elements, without changing cell shapes (Kubo et al., 2005). Surprisingly, the 35S::VND6 and 35S::VND7 plants showed clearly a diVerent morphology in the transdiVerentiated xylem vessel elements of the roots. VND6 overexpression induced metaxylem‐like vessel elements with reticulate and pitted thickenings of the secondary wall, whereas VND7 overexpression induced protoxylem‐like vessel elements with annular and spiral thickenings (Kubo et al., 2005). Accordingly, overexpression of the translational fusion of VND7 and VND6 to the SRDX, a strong repression domain, inhibited formation of metaxylem and protoxylem, respectively. However, there were no phenotypes for the vnd6 and vnd7 loss‐of‐function mutants or for transgenic plants carrying antisense VND6 and VND7. These results suggest that VND6 and VND7 are key regulators of xylem development with some functional redundancy.

VIII. TRACHEARY ELEMENT DIFFERENTIATION Formation of xylem cells is one of the most intensely studied aspects of cell diVerentiation in vascular plants (Aloni, 1987; Steeves and Sussex, 1989). The xylem is composed of a number of cell types, among which the highly specialized and easily recognizable water‐conducting cells, known as tracheids or vessels, have been most intensively studied. These two cell types are collectively referred to as TEs. The TEs undergo a very well‐defined process of diVerentiation that involves specification, elongation, SCW thickening, PCD, and cell wall removal (Fig. 6). At maturity, the TEs lose their nuclei and other cellular contents by leaving hollow tube through which water and nutrients pass. This process is coordinated such that neighboring TEs are joined together to form an interconnected network. Recent progress in studies of xylem diVerentiation has come from experiments both in vivo and in vitro expression studies on model systems as diverse as trees and cell cultures. These approaches have contributed to identifying a large number of candidate genes with potential roles in TE diVerentiation.

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Xylem parenchyma fibers, etc. Phloem tissue

Xylem tissue

Procambial Procambial cell identity

Cytoplasm

Cell elongation

Nucleus

Secondary Programmed cell wall cell death deposition

Chloroplast

Vacuole

Mature tracheary element Secondary cell wall

Fig. 6. Tracheary element (TE) diVerentiation occurring through identification of procambial cells, initiation of xylem diVerentiation, cell elongation, SCW thickening, and PCD.

Analysis of these genes has yielded important information on the diVerentiation processes, such as patterned SCW and PCD, which are two main morphological events occurring in TE diVerentiation (Fukuda, 2004; Turner et al., 2007; Ye, 2002). A. SECONDARY WALL FORMATION

Cell walls are frequently classified into two major types, primary cell walls and SCWs. The primary cell wall is generally a thin, flexible, and extensible layer formed in the growing cells, while the SCW is a thick layer formed inside the primary cell wall after cell growth is terminated. The SCW consists mainly of cellulose microfibrils. It also contains other cementing substances, such as lignin, hemicellulose, pectin, and proteins. The combinations of these substances add strength and rigidity to the wall (Fukuda, 1997; Turner et al., 2007). In developing TEs, SCW formation proceeds through cellulose microfibril deposition, deposition of other SCW components along the cellulose microfibrils, degradation and modification of primary cell walls, and lignification (Fukuda, 2004). Cellulose microfibrils are deposited in a transverse direction along the axis of the developing TEs, which have long been proposed to be controlled by the transversely oriented cortical microtubules (MTs) lying underneath the plasma membrane. Cortical MTs appear to function in the deposition of cellulose microfibrils, since both SCW pattern and cellulose microfibril orientation are altered by microtubule inhibitors. Adding taxol, an MT‐stabilizing drug,

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results in the stabilization of MT patterns and the coalignment of cellulose microfibrils. In addition, cellulose microfibrils are consistently parallel to cortical microtubules in developing TEs of some other cell types (Baskin, 2001). However, the relationship between cellulose microfibrils and cortical microtubules in regulating the pattern of SCW deposition may be more complicated than previously thought (Roberts et al., 2004). Through interference of cellulose crystallization with dyes, such as Congo Red and Evans Blue, Roberts et al. (2004) have examined the role of cellulose microfibrils in determining the localized deposition and continuity of SCW formation. When dyes were added to xylogenic cultures of the Zinnia mesophyll cells just before the onset of diVerentiation, cellulose microfibril deposition was disrupted during SCW formation and the pattern of SCW deposition was more irregular at localized sites that do not correspond to bands of cortical MTs. Furthermore, the application of microtubule inhibitors at diVerent stages of SCW deposition revealed that microtubules were lost before SCW deposition. These observations indicate that cortical microtubules may only be required at the early stages of SCW deposition but may not play a significant role in maintaining the pattern of SCW deposition (Roberts et al., 2004). Use of mutants with alterations in cortical MTs or cellulose microfibril patterns could eliminate possible nonspecific eVects that may occur with the use of pharmacological inhibitor. Two fragile fiber1 (fra1) and fra2 mutants have provided some insights into the relationship between cortical MT patterns and orientation of cellulose deposition. The fra2 mutant exhibits a reduction in fiber cell length and wall thickness, a decrease in cellulose and hemicellulose contents, and an increase in lignin condensation, indicating that the fra2 phenotype is a result of alterations in fiber cell elongation and cell wall biosynthesis. The FRA2 gene encodes a protein with a high similarity to katanin, a protein shown to be involved in regulating MT disassembly (Burk et al., 2001). In the SCW of fiber cells, the orientation of cellulose microfibrils is clearly altered, which reflects altered MT organization in the mutant (Burk and Ye, 2002; Burk et al., 2001). In contrast, the fra1 mutant shows alteration of cellulose microfibril orientation but without any defects in cell wall composition and cortical microtubule organization patterns. The fra1 phenotype is caused by a mutation in a kinesin‐like protein, and it remains unclear how this alters cellulose deposition (Zhong et al., 2002). SCWs in wood are mainly composed of three major components, cellulose, hemicellulose, and lignin (Turner et al., 2007). There has been a significant progress in the identification of genes involved in the synthesis of SCW components. Studies using Arabidopsis mutants have demonstrated that IRREGULAR XYLEM1 (IRX1), IRX3, and IRX5 interact with the

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cellulose synthase (CESA) complex in SCW formation (Doblin et al., 2002; Gardiner et al., 2003; Taylor et al., 2003). They initially localize within the cytoplasm and afterward colocalize at the plasma membrane with bands of cortical microtubules (Gardiner et al., 2003). In the absence of IRX1, IRX3, and IRX5, however, the remaining two subunits are retained in the cytoplasm, demonstrating that all three CESA proteins are required for the formation of the intact CESA complex. Actin, other component forming microtubules in the cell, did not colocalize with IRX3 in the developing xylem vessels, indicating that arrays of cortical microtubules, but not those of actin filaments, are necessary in maintaining normal CESA localization (Gardiner et al., 2003). Previous studies using microarray data and reverse genetics have identified several genes likely to be involved in the cellulose biosynthesis (e.g., KORRIGAN, COBRA‐LIKE4), however, their precise roles in the cellulose biosynthetic process remains largely unclear (Brown et al., 2005; Persson et al., 2005; Szyjanowicz et al., 2004). Hemicellulose includes several heteropolymers present in almost all plant cell walls. While cellulose is a 14‐ ‐linked polyglucan, hemicellulose can include various structures of polysaccharides, such as glucose, xylose, mannose, galactose, rhamnose, arabinose, fucose, glucuronic acid, and galacturonic acid. Based on the nature of substitutions, xylan is named methyl glucuronoxylan (GX), which is the most common hemicellulose found in dicot woods (Ebringerova and Heinze, 2000). The IRX9 gene, which encodes a putative glycosyltransferase (GT) (Coutinho et al., 2003), has been found to be essential for normal elongation of the xylan chain. It is expressed in cells undergoing SCW biosynthesis (Pen˜a et al., 2007). The irx9 knockout mutant exhibits a reduction in the GX level as well as a significant decrease in GX chain length (Brown et al., 2005; Lee et al., 2007a; Pen˜a et al., 2007). These results suggest that IRX9 is a putative xylan synthase required for GX backbone elongation (Lee et al., 2007a; Pen˜a et al., 2007). Two other GTs, FRA8 and IRX8, have been shown to be required for the biosynthesis of the tetrasaccharide primer sequence at the reducing end of GX (Pen˜a et al., 2007; Persson et al., 2007). Recently, IRX14 and PARVUS have been added to GTs functioning in the GX biosynthesis. (Brown et al., 2007; Lee et al., 2007b). The irx14 and parvus mutants exhibit large decreases in xylan, similar to the fra8, irx8, and irx9 mutants. Analysis of xylosyltransferase activity is consistent with a role for PARVUS in the synthesis of a primer, with IRX9 and IRX14 being involved in elongation of the xylan backbone (Brown et al., 2007). In the past two years, several transcription factors belonging to the NAC and MYB families have been shown to be key proteins in activation of SCW biosynthesis. Two NAC transcription factors, VND6 and VND7 (Kubo

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et al., 2005), regulate the specificity of metaxylem and protoxylem in the primary roots, respectively. Overexpression of the VND6 and VND7 induce SCW deposition in the respective xylem tissues. Two additional NAC transcription factors, NAC SECONDARY WALL THICKENING PROMOTING FACTOR1 (NST1) and NST2, function redundantly in regulating SCW deposition in the endothelium of anthers (Mitsuda et al., 2005). In the epidermis, NST1 overexpression causes cell wall patterning, reminiscent of TEs, whereas it does not cause patterning of ectopic SCWs in mesophyll and other cell types. It seems that NST1 regulates SCW deposition independent of cell wall patterning. A similar phenotype was also described for mutations in a MYB26 gene (Steiner‐Lange et al., 2003). Two research groups independently discovered important roles of Arabidopsis NAC proteins, SECONDARY WALL ASSOCIATED NAC DOMAIN PROTEIN1 (SND1) in SCW biosynthesis (Mitsuda et al., 2007; Zhong et al., 2007). The SND1 gene is specifically expressed in the interfascicular fibers. While suppression of SND1 function results in a severe reduction in SCW thickening in fibers, its overexpression activates the expression of genes involved in the biosynthesis of cellulose, xylan, and lignin, leading to ectopic deposition of SCWs. It is likely that SND1 and NST1 act redundantly in the regulation of SCW thickening in fibers (Mitsuda et al., 2007; Zhong et al., 2007). Lignin is the second most abundant terrestrial biopolymer. Lignin fills the spaces in the cell wall between cellulose, hemicellulose, and other component. It provides additional mechanical strength to the SCW and the SCW waterproof ability owing to its hydrophobic nature (Boerjan et al., 2003). Lignin biosynthesis begins in the cytosol with the synthesis of glycosylated monolignols from phenylalanine. The attached glucose renders monolignols water soluble and less toxic. Once transported to the cell wall, glucose is removed and lignin is formed by polymerization of the monolignols. The polymerization step is a radical–radical coupling, so the dehydrogenation to monolignol precedes polymer formation. This process is catalyzed by oxidative enzyme, such as peroxidases, laccases, and polyphenol oxidases. Oxidants of low molecular weight might also be involved in this step ¨ nnerud et al., 2002). By polymerization, lignin is first deposited in the (O middle lamella and in the cell corners of the primary wall after SCW formation has started, and the bulk of lignin is deposited after cellulose and hemicellulose have been deposited in the outer layer of SCW. Lignin deposition is influenced by the orientation of the cellulose microfibrils (Donaldson, 2001; Roussel and Lim, 1995). Several MYB transcriptional factors regulate the phenylpropanoid biosynthetic pathway leading to the synthesis of monolignols (Borevitz et al., 2000;

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Tamagnone et al., 1998). However, only a few MYB members, including PtMYB4 from pine (PatzlaV et al., 2003) and EgMYB2 from Eucalyptus (Goicoechea et al., 2005) have been shown to be directly associated with lignin biosynthesis. Both the PtMYB4 and EgMYB2 gene are expressed in secondary xylem and bind to the promoters of several lignin biosynthetic genes. Besides, overexpression of PtMYB4 results in ectopic deposition of lignin, while overexpression of EgMYB2 causes SCW thickness in fibers and vessels, suggesting that EgMYB2 regulates many aspects of SCW biosynthesis, not just lignification. Moreover, a LIM transcription factor can bind to the PAL box cis‐ element in the promoters of monolignol biosynthetic genes, and repression of the LIM gene leads to inhibition of lignin biosynthesis (Kawaoka et al., 2000). It will be interesting to investigate how these transcription factors act together to regulate lignin biosynthesis. B. PROGRAMMED CELL DEATH

The TE PCD has long been recognized as a primary example of developmental PCD in plants (Fukuda, 1997). The TEs reach their maturity after loss of cellular contents. At the final stage of TE maturation, the digested cellular contents are released out, usually into a neighboring hollow TE. A most striking feature of xylem cell death is the collapse of the vacuole that coincides with the digestion of the nucleus (Fukuda, 1997, 2004; Turner et al., 2007). The degradation of nuclear and chloroplast DNA is triggered by vacuole collapse and is completed within only 15 min after the vacuole collapse (Obara et al., 2001). The rupture of vacuole releases hydrolytic enzymes, including proteases, DNAses, and RNAses, into the cytoplasm (Funk et al., 2002; Ito and Fukuda, 2002; Lehmann et al., 2001), some of which may cause destruction of organelles, such as nucleus and chloroplast. In the Zinnia culture cells, vacuolar collapse is not suYcient for PCD as it does not induce digestion of the nucleus due to the lack of accumulation of hydrolytic enzymes (Obara et al., 2001). An S1 type nuclease, Zinnia endonuclease 1 (ZEN1), has been shown to function in the nuclear DNA degradation during TE PCD (Ito and Fukuda, 2002), indicating that the accumulation of hydrolytic enzymes is important for TE PCD. Moreover, after the accumulation step in the vacuole, the enzymatic activity of these enzymes strikingly increases at the time of vacuolar collapse. The study of the vacuolar localization of the XCP protease, a xylem‐specific papain‐like cysteine peptidase, produced more information on the accumulation of hydrolytic enzymes in the vacuole (Funk et al., 2002). It was localized in the vacuole and ectopic expression of XCP1 resulted in a reduction in plant size and early leaf senescence, as indicated by early loss of

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leaf chlorophylls. In contrast, there is at least one RNAse that accumulates in the ER of TEs (Lehmann et al., 2001). This enzyme may be activated by the acidification of the cytosol that results from vacuolar collapse. There has been a remarkable accumulation of data in recent years indicating that caspase inhibitors can suppress cell death in plants and that caspase‐ like protease activities are important in this process (Rotari et al., 2005). It is very intriguing that a similar proteolytic cascade may exist in plant and animal PCD, despite very little sequence conservation at the molecular level. In animals, caspases are specifically activated during PCD. In particular, caspases initiate cell death by degrading several proteins essential for cell integrity, such as poly(ADP‐ribose) polymerase, lamins, and gelsolin. In support of a role for these caspase‐like activities in plant PCD, experiments in tobacco have shown that treatment with menadione, a caspase inhibitor, can block the induction of DNA fragmentation and of poly(ADP‐ribose) polymerase cleavage during PCD (Sun et al., 1999). Other caspase inhibitors, such as acetyl‐Asp‐Glu‐Val‐Asp‐aldehyde (Ac‐DEVD‐CHO) and acetyl‐ Tyr‐Val‐Ala‐Asp‐aldehyde (Ac‐YVAD‐CHO), have also been shown to block PCD after pathogen induction (del Pozo and Lam, 1998). In addition, expression of p35, a caspase inhibitor, has been reported to reduce the onset of apoptosis in embryonic callus in maize (Hansen, 2000). However, these observations provide only indirect evidences suggesting that caspases are present in plants. Therefore, the identification of a protease exhibiting caspase activity is essential in elucidating the molecular mechanism that operates PCD in plants. The protease VACUOLAR PROCESSING ENZYME (VPE), which possesses caspase 1 activity, has been identified as a key component of a virus‐induced hypersensitive response that involves PCD (Hatsugai et al., 2004). Intriguingly, VPE is structurally unrelated to caspases, although it has a caspase‐1 activity. Silencing of VPEs suppresses vacuole collapse in the TMV‐infected leaves. VPEs may therefore be required to activate various vacuolar proteins involved in the collapse of vacuoles that occurs during plant PCD. One member of the VPE family in Arabidopsis is up‐regulated during TE formation and could therefore be involved in vacuole collapse. To clarify the association of caspase‐like protease with TE PCD, a study using the full range of caspase inhibitors available would prove very informative. PCD is recognized as a common process in eukaryotes during development and in response to pathogen infection and abiotic stress signals (Turner et al., 2007). Morphologically and biochemically, PCD in plants and yeast shares numerous common features with metazoans. These include chromatin condensation and nuclear DNA fragmentation, involvement of reactive oxygen species, and participation of caspase‐like proteases. Cell death

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signaling molecules, such as ceramides, can also regulate cell death in both animal and plant cells (Liang et al., 2003). These observations thus argue for conserved cell death signaling mechanisms in plants and animals. A few plant genes have been identified as orthologues of mammalian genes involved in apoptosis, including those encoding cytochrome C, At‐DAD1 (Arabidopsis thaliana Defender against Apoptotic Death‐1), and At‐BI1 (Arabidopsis thaliana BAX Inhibitor‐1). The release of cytochrome C from mitochondria is known to be an early marker of cell death (Turner et al., 2007). One study has shown that TE mitochondria are in connection with the process of TE PCD and that cytochrome C is released before the vacuole collapse (Yu et al., 2002). This result implies that vacuole rupture is not a trigger of TE PCD but more likely a result of TE PCD and that earlier PCD events remain to be characterized. The DAD1 gene has originally been discovered in hamster cells, where the cell line carrying the dad1 mutation dies via apoptosis (Nakashima et al., 1993). In Arabidopsis, there are two homologues of DAD1: AtDAD1 and AtDAD2. Transient expression assays have shown that overexpression of AtDAD1 or AtDAD2 could protect cells from PCD. This result indicates that the plant DAD proteins have the ability to suppress or significantly delay the induction of DNA fragmentation (Danon et al., 2004). In addition, AtDAD1–EGFP fusion has the same subcellular localization pattern as the ER‐targeted EGFP, indicating that the plant DAD1 protein is located in the ER. Interestingly, the ER has been proposed to be a new gateway to PCD in animal cells, with the implication of ER calcium (Scorrano et al., 2003). BAX inhibitor‐1 (BI‐1) has originally been identified as a cell death suppressor activated by BAX in yeast or mammalian cells (Xu and Reed, 1998). BI‐1 is evolutionarily conserved and predicted to be a transmembrane protein that localizes predominantly to the ER. Expression of plant BI‐1 mRNA is detected in various tissues, and its expression level is enhanced during senescence and under several types of biotic and abiotic stresses. Overexpression of BI‐1 from various plant species suppresses BAX‐, pathogen‐, or abiotic stress‐induced cell death in a variety of cells from yeast, plants, and mammalian origins. These observations support the idea that BI‐1 could have conserved function in diverse organisms. The atbi1‐1 and atbi1‐2 mutants exhibit accelerated progression of cell death upon infiltration of leaf tissues with a PCD‐inducing fungal toxin fumonisin B1 (FB1) and increased sensitivity to heat shock‐induced cell death (Watanabe and Lam, 2006). Overexpression of an AtBI1 transgene in the two homozygous mutant backgrounds rescued the accelerated cell death phenotypes. Recently, it has been demonstrated that AtBI1 plays a pivotal role as a highly conserved survival factor during ER stress (Watanabe and Lam, 2008). Plant BI‐1 is thus likely to play an important role as an

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attenuator for cell death progression under multiple stress conditions that could trigger PCD. Despite a large amount of information on TE PCD, virtually nothing is known of the induction pathway for the TE PCD at the biochemical or molecular level. Unidentified toxins, high concentrations of salts, or some chemicals may induce the apoptosis‐like death of particular cells (Katsuhara and Kawasaki, 1996; Wang et al., 1996). Calcium and nitric oxide may also be involved in the PCD process (Gabaldon et al., 2005; Groover and Jones, 1999). In addition, little is known about the genes and molecules that regulate the process. The PCD process that takes place at the last stage of TE diVerentiation is coupled tightly to the formation of SCWs. Indeed, it is not easy to separate experimentally PCD from SCW formation during TE diVerentiation. Identification of the genes and related mutants would represent an important breakthrough and may require specific genetic screens (Fukuda, 2004; Turner et al., 2007).

IX. CONCLUDING REMARKS Arabidopsis and Populus mutants that show defects in vascular development and cellular studies with the Zinnia xylogenic culture cells have provided insights into the formation of vascular tissues. Formation of the vascular system is a complex developmental process that must be regulated via the interactions of diverse genetic components and growth hormones. Patterning and organization of vascular system are quite variable among diVerent species. However, the molecular mechanisms underlying their spatial and temporal regulation are likely shared by virtually all vascular plants. For example, the HD‐ZIP III genes and its molecular regulator miR165/166 are present in diverse plant species (Floyd and Bowman, 2004), strongly supporting that key molecules and mechanisms regulating vascular development are conserved in all vascular plants. In addition, recent genome‐wide expression analyses in secondary vascular tissues have produced interesting findings. Several genes known to function in the SAM have also been found expressed in the vascular cambium (Schrader et al., 2004; Zhao et al., 2005). Although the transcripts for some important SAM identity genes, such as CLV3 and WUSCHEL (WUS), were not detected, CLAVATA3‐LIKE (CLE) family members were found in the SAM and vascular cambium tissues. Moreover, WUS‐related PttHB2 and PttHB3 and CLV1‐related PttRLK3 were detected in Populus. The coregulation of the SAM and vascular cambium by HD‐ZIP IIIs further support the shared functional mechanisms in various plant species.

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In recent years, critical mechanisms controlling the process of vascular development with their associated genes have been identified, and principles of phytohormone functioning in vascular continuity and diVerentiation of various cell types have been established. It is important to note, however, that we are far from having a full understanding of the molecules and mechanistic schemes that govern vascular development. Especially, we still know only a little about pre‐procambial cells. Pre‐procambial cells are hard to visualize and follow through developmental stages, so they have been the focus of few investigators. However, as a wider selection of marker genes has been described, research on pre‐procambial cell specification and function should become more tractable. This area is likely to be of critical importance for understanding vein patterning. Therefore, our next target should be to dissect the precursor stage of procambial and xylem cells on the basis of their cellular and molecular functions, which will allow us to detect new intercellular and intracellular signals that initiate each step. Another challenging issue for the future is the establishment and maintenance of the polarity of vascular cells for continuous vascular bundles. Certain levels of vascular tissue organization, such as the development of major and minor veins within the leaves, cannot be easily applied to the canalization concept. To understand the molecular mechanism of vascular cell polarity, we have to identify the asymmetrical intracellular signaling pathways that control the polarity. Introduction of new concept controlling the polarity might be good solution. For example, the concept of feedback module regulated by competitive inhibitors of small peptides has provided a stimulating clue as to how the HD‐ZIP III transcription factors, in conjunction with miR166/165, regulate both the SAM and vascular development (Kim et al., 2008; Wenkel et al., 2007). Furthermore, both the complexity of overexpression phenotypes and loss‐of‐function analyses suggest that we must identify their direct targets in order to understand the roles of genes comprehensively. Analysis of the target genes of the various transcription factors might provide the hint for the regulation of intracellular polarity. With the availability of various mutants showing defects in vascular patterning and diVerentiation and characterization of their corresponding genes using genomic tools, it will soon be possible to solve the multiple questions concerning vascular development.

ACKNOWLEDGMENTS This work was supported by the Brain Korea 21 (BK21), Biogreen 21 (20080401034001), and National Research Laboratory Programs and by

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grants from the Plant Signaling Network Research Center and Korea Science and Engineering Foundation (2007‐03415).

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Plant Lectins

ELS J. M. VAN DAMME, NAUSICAA LANNOO AND WILLY J. PEUMANS

Department of Molecular Biotechnology, Laboratory of Biochemistry and Glycobiology, Ghent University, 9000 Gent, Belgium

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. From the Discovery of Ricin to Modern Lectinology .................. B. Definition, Nomenclature, and Terminology............................. C. Old and New Paradigms in Plant Lectin Research...................... II. Rethinking Lectins in Terms of Carbohydrate Binding Domains . . . . . . . . III. Overview of Carbohydrate Binding Domains Identified in Plants. . . . . . . . A. Brief Historical Overview ................................................... B. Description of the Different Carbohydrate Binding Domains/Families ............................................................ IV. Sugar Binding Activity and Specificity of Plant Lectins . . . . . . . . . . . . . . . . . . A. Plant Lectins Possess Extended Binding Sites That Preferentially Interact with Complex Glycans ............................................ B. High Performing Analysis Techniques Urge to Revise the Old Paradigms............................................................ V. Biosynthesis, Topogenesis, and Subcellular Location of Plant Lectins . . . A. An Old Paradigm: Plant Lectins Follow the Secretory Pathway ... B. Experimental Evidence for the Occurrence of Nonsecretory Plant Lectins ................................................. C. Genome/Transcriptome Data Indicate That the Majority of all Plant Lectins Is Synthesized on Free Ribosomes and Resides in the Nucleocytoplasmic Compartment.................................. VI. Expression of Lectins in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. General Aspects............................................................... B. Highly Expressed ‘‘Classical’’ Lectins ..................................... Advances in Botanical Research, Vol. 48 Incorporating Advances in Plant Pathology Copyright 2008, Elsevier Ltd. All rights reserved.

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0065-2296/08 $35.00 DOI: 10.1016/S0065-2296(08)00403-5

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C. Inducible Low Expressed ‘‘Novel’’ Lectins ............................... VII. Why do Plants Express Lectins? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Classical Concepts: Many Hypotheses but Poor Experimental Evidence ...................................................... B. Novel Concepts ............................................................... C. Developing Research Areas................................................. VIII. The Plant Lectinome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABSTRACT Plant lectins are being studied for over a century. Until a decade ago, most information was obtained from biochemical, molecular, and structural studies of a reasonably high but still limited number of abundant lectins from seeds and vegetative storage organs. Though the results of these studies are still valid, the recent progress made in several areas of plant lectin research urge to profoundly revise the prevailing concepts. This contribution aims to give a comprehensive overview of the occurrence, taxonomic distribution, molecular evolution, and physiological role of plant lectins with the emphasis on relevant novel developments. The overview comprises a description of the 12 diVerent carbohydrate binding domains and annex lectin families that hitherto have been identified in plants. For each lectin family, a fairly detailed summary is given of the biochemical properties, taxonomical distribution, and evolutionary origin. Novel insights in some specific aspects of plant lectins like their biosynthesis and topogenesis, carbohydrate binding specificity, and regulation of expression are discussed. The last major section deals with an updated critical discussion of the physiological role of plant lectins with an emphasis on specific functions within the plant cell. Finally, the concept of ‘‘lectinome’’ is introduced.

ABBREVIATIONS ABA Calsepa CEBiP ConA CRA CV-N DPBB EAA EGFP ER ERAD EUL Gal GalNAc GJRL

Agaricus bisporus agglutinin Calystegia sepium agglutinin Chitin Elicitor Binding Protein Concanavalin A chitinase-related agglutinin cyanovirin double-psi
-barrel Euonymus europaeus agglutinin enhanced green fluorescent protein endoplasmic reticulum ER-associated degradation Euonymus lectin domain galactose N-acetylgalactosamine galactose-binding jacalin-related lectins

PLANT LECTINS

Glc GlcNAc GNA Hfr HIV JRL LecRK LegL LegLu LysM Man MarpoABA MJRL NFR Nictaba NLS Orysata PAG PDB PHA PP2 PR RIP RLK RlpA RobpsCRA SNA SNRLP TMD UDA WGA

109

glucose N-acetylglucosamine Galanthus nivalis agglutinin Hessian fly responsive human immodeficiency virus jacalin-related lectins lectin receptor kinase legume lectin ubiquitous legume lectin Lysin domain mannose ABA homolog from Marchantia polymorpha mannose-binding jacalin-related lectins Nod factor receptor Nicotiana tabacum agglutinin nuclear localization signal Oryza sativum agglutinin polynucleotide:adenosine glycosidase Protein DataBase Phaseolus vulgaris agglutinin phloem protein 2 pathogenesis related ribosome-inactivating protein receptor-like kinase Rare lipoprotein A CRA homolog from Robinia pseudoacacia Sambucus nigra agglutinin Sambucus nigra lectin-related protein transmembrane domain Urtica dioica agglutinin wheat germ agglutinin

I. INTRODUCTION A. FROM THE DISCOVERY OF RICIN TO MODERN LECTINOLOGY

The onset of the scientific discipline called ‘‘lectinology’’ dates back to the observation by Stillmark in 1888 that the toxicity of castor bean extracts is linked to the presence of a proteinaceous hemagglutinating factor called ‘‘ricin.’’

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Soon after, similar toxins were also reported in other plants. In 1898, Elfstrand introduced the term ‘‘hemagglutinin’’ as a common name for all plant proteins that clump blood cells (Elfstrand, 1898). In the following decade, the idea that toxicity is an intrinsic property of hemagglutinins was abandoned after Landsteiner and Raubitschek (1907) reported for the first time the presence of nontoxic lectins in the legumes Phaseolus vulgaris, Pisum sativum, Lens culinaris, and Vicia sativa. In the subsequent years, many nontoxic lectins were discovered and it became clear that plant lectins are widespread in the plant kingdom. The discovery that some hemagglutinins selectively agglutinated erythrocytes of a particular human blood group within the ABO system can be considered a milestone in the history of plant lectins. This finding led to the introduction of the term ‘‘lectin’’ which is derived from ‘‘legere,’’ the Latin verb for ‘‘to select’’ (Boyd and Reguera, 1949; Renkonen, 1948). However, since most hemagglutinins are also capable of agglutinating other cells, they were also referred to as agglutinins. Though the term lectin is actually most commonly used, the terms agglutinin and hemagglutinin still persist as synonyms. Once the blood group specificity of lectins was established, research aiming at the elucidation of the underlying mechanism for this biological activity was started. Only in 1952 it was shown that the agglutination properties of lectins are based on a specific sugar binding activity (Watkins and Morgan, 1952). From then onwards, lectins were no longer regarded as agglutinating factors but rather as sugar binding proteins that could be distinguished from other proteins on the basis of a well‐defined functional criterion. Classical methods to study the interaction between lectins and carbohydrates often relied on the agglutination of erythrocytes or other cell types. However, later on, the carbohydrate specificity of lectins was explored by indirect methods, such as for example, inhibition of precipitation or cellular agglutination by hapten sugars or glycoconjugates. More recently, the introduction of high performing techniques such as frontal aYnity chromatography and glycan microarrays enabled high throughput screening of large collections of carbohydrates and more complex glycans with only small amounts of a purified lectin.

B. DEFINITION, NOMENCLATURE, AND TERMINOLOGY

The first definition of lectins was based primarily on the sugar specificity and inhibition of the agglutination reaction. According to this definition, lectins are carbohydrate binding proteins of nonimmune origin which agglutinate cells and/or precipitate glycoconjugates (Goldstein et al., 1980). Although this definition was approved with minor changes by the Nomenclature

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Committee of the International Union of Biochemistry in 1981, there was a lot of debate, mainly because this definition was restricted to multivalent carbohydrate binding proteins (Dixon, 1981), and did not take into account those lectins that are poorly active agglutinins or the chimeric lectins that possess, in addition to their sugar binding domain, a second domain with catalytic activity. Several modifications to the definition were proposed, but all of them had their shortcomings. Finally to conclude the debate, a definition was introduced that is based on a single and simple criterion, namely the presence of at least one noncatalytic domain that binds reversibly to a specific carbohydrate. Accordingly, plant lectins are defined as ‘‘all plant proteins that possess at least one noncatalytic domain that binds reversibly to a specific mono‐ or oligosaccharide’’ (Peumans and Van Damme, 1995). Until today, this definition is still accepted by the scientific community. On the basis of the overall structure of lectins, a distinction is made between so‐ called hololectins and chimerolectins. In contrast to hololectins that consist solely of carbohydrate binding domains, chimerolectins also contain an unrelated domain that acts independently of the lectin domain. If the hololectin is composed of a single carbohydrate binding domain, it is referred to as a merolectin, whereas hololectins with two diVerent carbohydrate binding domains are also referred to as superlectins. The nomenclature of plant lectins is very diverse. Since the early discovery plant lectins were usually designated by a trivial name, often a composite of an abbreviation of the (scientific) name of the source plant followed by the suYx ‐in (e.g., ricin from Ricinus communis). Other plant lectins were designated by three‐letter abbreviations derived from the plant or plant material in which the protein was found (e.g., WGA, wheat germ agglutinin; PNA, peanut agglutinin). In an attempt to make lectin nomenclature more uniform, the three‐letter code was composed by using the first letter of the genus and species name of the plant followed by L (for lectin) or A (for agglutinin), as for example, in PSA (Pisum sativum agglutinin). To distinguish between diVerent plant lectins isolated from the same plant species, roman characters were often added after the three‐letter code. The elderberry (Sambucus nigra) lectins for instance, are referred to as SNA‐I to SNA‐V, reflecting the diVerent lectins in chronological order of their discovery. Unfortunately, this lectin nomenclature is impractical and over the years has lost its transparency since several three‐letter codes refer to two or more lectins from diVerent species (e.g., ACA stands for Amaranthus caudatus agglutinin but also refers to Allium cepa agglutinin). Therefore, we recently proposed to introduce abbreviations comprising the first three letters of the genus and species names followed by ‐a (e.g., Nictaba refers to Nicotiana tabacum agglutinin).

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ELS J. M. VAN DAMME ET AL. C. OLD AND NEW PARADIGMS IN PLANT LECTIN RESEARCH

For a long time, plant lectin research concentrated on (i) the isolation and characterization of new carbohydrate binding proteins with interesting sugar binding and biological activities and (ii) the elucidation of the physiological role of diVerent types of lectins from diVerent species/taxa. Since for practical reasons most scientists selected abundant lectins and especially those found in seeds as model proteins, the whole research on plant lectins was to a certain extent biased. Though this bias did not aVect the pioneering biochemical, structural, and activity studies, it had a negative impact on the development of a correct global view on what lectins are doing inside or outside the plant. As a result, persistent misconceptions gradually grew and eventually were taken for granted. Most unfortunately, the tenacious belief in (partly) incorrect but presumed canonical concepts seriously hampered the development and acceptance of novel ideas about the occurrence and function of lectins in plants. A first old paradigm deals with the occurrence and taxonomical distribution of plant lectins. On the basis of the documented occurrence of (abundant) forms, it was more or less generally believed that lectins are confined to a limited set of taxonomic groups comprising one or a few families (e.g., legume lectins), genera (e.g., Solanaceae lectins), or even species (e.g., jacalin). Moreover, there was also a tendency to assume that most plants express only one or a few closely related lectins. Though searches for weakly expressed homologs of the abundant lectins considerably broadened the documented taxonomic distribution, it was only until the advent of genome and transcriptome sequence programs that a fairly accurate global view could be developed of the overall phylogenetic distribution of each individual lectin family and the lectin complement of individual species. As will be discussed below in detail, all evidence suggests (i) that most lectin families are ubiquitous in land plants and (ii) that all these plants possess a very complex complement of lectin genes. A second paradigm concerns the domain architecture of plant lectins. For a long time, all documented plant lectins except the type 2 ribosome‐ inactivating proteins (RIPs) were hololectins. Accordingly, hololectins were considered the rule and chimerolectins the exception. Even after the discovery of novel chimeric forms (e.g., class I chitinases), the chimerolectins remained a minor group until sequencing of the Arabidopsis genome revealed the occurrence of an extended family of genes encoding receptor kinases with an extracellular domain equivalent to the legume lectins. At present, the huge amount of data generated by genome/transcriptome programs clearly indicates that the majority of all plant lectin genes have a chimeric domain

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architecture. This finding is of paramount importance in view of the function of plant lectins because it emphasizes the fact that most of them are not just sugar binding proteins but bi‐ or multi‐functional proteins. The next paradigm deals with the reigning ideas about the ‘‘general biology’’ of plant lectins. Because of the preferential use of abundant lectins and especially these from legume seeds as model systems, plant lectins are classically considered proteins that are highly expressed—mostly in a developmentally regulated manner—in seeds and/or vegetative storage tissues, where they are synthesized with a signal peptide and targeted via the secretory pathway into the vacuolar or extracellular compartment. Though this scenario holds true for most lectins studied in the past century, evidence is accumulating for about a decade (i) that plants also synthesize lectins in response to specific stress factors and (ii) that these inducible lectins are synthesized in the cytoplasm and reside in the cytoplasmic/nuclear compartment of the cell. Hitherto, only a limited number of low expressed inducible lectins have been described (e.g., jasmonate‐inducible lectins in tobacco and cereals). However, genome/transcriptome analyses not only confirmed the presence and expression of genes encoding cytoplasmic lectins in many species but also revealed that these genes are (nearly) ubiquitous in Embryophyta. It appears, therefore, that (low expressed) cytoplasmic/nuclear lectins are the rule and (highly expressed) vacuolar homologs the exception. The diVerential subcellular location of the ‘‘vacuolar’’ and ‘‘cytoplasmic’’ forms urges to revise another old paradigm concerning the physiological role of plant lectins. In principle, ‘‘vacuolar’’ lectins can only interact with either endogenous glycans present in the same compartment or when secreted with exogenous glycans from foreign organisms. This assumption combined with the high expression level and preferential interaction with foreign glycans eventually led to the concept that plant lectins do not fulfill a specific role within the plant but should be considered defense‐related proteins. In contrast, the ‘‘cytoplasmic’’ lectins are capable of interacting with endogenous glycans present in those compartments where signals are transduced and most cellular processes are regulated. Accordingly, it is believed that at least some cytoplasmic plant lectins play a specific endogenous role. A final paradigm that needs a thorough revision concerns the carbohydrate binding specificity of plant lectins. In the early days of lectinology, specificity was often discussed in terms of the recognition of mono‐ and/or oligosaccharides. Later, it became increasingly evident that most plant lectins preferentially interact with complex glycans from especially animal origin. Though most of the early data (which were obtained by assays based on competition) are still valid, the introduction of new technologies that measure direct interactions (surface plasmon resonance, frontal aYnity

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chromatography, and especially glycan microarrays) shed a new light on the carbohydrate binding properties of plant lectins. The most important conclusion is that—apart from a few exceptions—plant lectins exhibit a general preference for N‐glycans. Even lectins which were classically considered chitin specific (e.g., the tomato lectin and the tobacco lectin) appear to have a much higher aYnity for N‐glycans.

II. RETHINKING LECTINS IN TERMS OF CARBOHYDRATE BINDING DOMAINS Pioneering work with animal and especially vertebrate lectins revealed that the carbohydrate binding activity often resides in a small part of a complex protein and that proteins with a totally diVerent overall structure share the same or a very similar sugar binding motif. To cope with the apparent complexity of the animal lectin families, the composing carbohydrate binding domain(s) had to be regarded as discrete structural/functional units embedded in a complex multi‐domain protein. Accordingly, animal lectins are now routinely discussed in terms of carbohydrate binding domain(s) (Taylor and Drickamer, 2003). Since the great majority of all previously purified and characterized plant lectins are hololectins, there was less need to introduce a classification on the basis of the carbohydrate binding domain(s). Ironically the very first plant lectin that was identified, namely ricin, is a typical chimeric protein. However, ricin is such a notorious toxin that its classification as a so‐called type 2 RIP is self‐evident. The same applies to for example, class I chitinases, which are primarily considered chitinases rather than chimeric lectins with a chitinase domain. During the past few years, several examples were reported of genes in which a domain corresponding to a well‐defined plant lectin is fused to an unrelated domain. For example, upon infestation with Hessian fly larvae, wheat plants express a gene comprising an N‐terminal domain resembling amaranthin fused to a C‐terminal lytic toxin domain as well as a gene in which a C‐terminal domain equivalent to jacalin is linked to an N‐terminal dirigent domain. Moreover, genome and transcriptome analyses provide ample evidence for the occurrence in all land plants of numerous genes with one or more lectin domain(s) embedded in a more or less complex multi‐domain architecture. Because of this obvious complexity, it is also for plant lectins preferable to think in terms of carbohydrate domains. Therefore, the present chapter adopts a classification based on the identity of the sugar binding motif(s). This chapter aims to present a comprehensive up‐to‐date overview of the most recent ideas with regard to the occurrence, taxonomic distribution,

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molecular evolution, and physiological role of plant lectins. In principle, the overview is confined to lectins and lectin genes found in land plants (Embryophyta). Lectins from other Viridiplantae are not included but whenever relevant, the presence/absence of a given carbohydrate binding domain in the sequenced genomes of green algae will be discussed. To place the evolutionary origin of the modern plant carbohydrate binding domains in a proper perspective, the possible presence of homologous domains in other eukaryotes as well as in prokaryotes will be mentioned briefly. In the first part of the text, a comprehensive overview is given of the diVerent carbohydrate binding domains that hitherto have been identified in plants. For each carbohydrate binding domain/lectin family, a fairly detailed summary is given of the biochemical properties, taxonomical distribution, and evolutionary origin. Thereby, the emphasis is put on novel developments during the past decades. Moreover, whenever appropriate, reference is made to the (preliminary) unpublished results of a comprehensive database screening (done by WP and EVD). Issues that were extensively reviewed in previous papers are only briefly mentioned. The second part focuses on recently developed novel concepts with regard to the carbohydrate specificity, general physiology, and the phylogenetic distribution of the diVerent plant lectin families, and the conclusions based thereon for what concerns the importance of protein–carbohydrate interactions in growth and development of plants. In the third part, the concept of ‘‘lectinome’’ is introduced.

III. OVERVIEW OF CARBOHYDRATE BINDING DOMAINS IDENTIFIED IN PLANTS A. BRIEF HISTORICAL OVERVIEW

Before the introduction of the first biochemical methods to purify and (partially) analyze proteins, hemagglutination activity was the only criterion to distinguish lectins from other proteins. Accordingly, lectins were considered a more or less homogeneous group of proteins sharing the ability to clump red blood cells. Once suitable analytical techniques became available, plant lectins were among the first proteins to be studied in some detail. Though it soon turned out that lectins from diVerent plant species exhibited diVerent biochemical and biophysical properties, the limited resolving power of the applied techniques did not allow a subdivision into multiple groups or families. In the mean time, it was established that the agglutination activity of lectins relies on their ability to recognize and bind specific sugars. As a result,

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a great deal of lectin research focused on their sugar binding activity and specificity. Thereby, it became evident that lectins from diVerent sources exhibit diVerent specificities, which eventually led to the concept that plant lectins can be classified into a number of ‘‘specificity groups.’’ Unfortunately, diVerences in specificity could not always be reconciled with diVerences in biochemical/biophysical properties and vice versa indicating that specificity was not an adequate criterion for classification of plant lectins in what at the time were called ‘‘natural groups.’’ The occurrence of such groups was not only inferred from biochemical/biophysical analyses of a steadily increasing number of purified lectins but seemed logical in the context of the discovery of multiple families of plant storage proteins (which were a very important research object in the first phase of plant molecular biology). Immunological analyses provided the first firm indications for the occurrence of plant lectin families with distinct serological properties. Definitive proof followed the first (partial) sequencing of lectin polypeptides. Though these early sequence data left no doubt about the occurrence of multiple lectin families, the available information was very limited and insuYcient for a general classification of all plant lectins. Fortunately, the rapid progress during the 1990s in plant biochemistry and molecular biology yielded a wealth of useful information about numerous lectins and their genes. On the basis of a comprehensive analysis of plant lectin sequences combined with relevant data from plant genome/transcriptome analyses a system was elaborated in 1998, whereby the majority of all plant lectins known at the time were classified into seven families of structurally and evolutionarily related proteins (Van Damme et al., 1998a). Though most ‘‘novel’’ plant lectins identified during the last decade still fit this classification, several recent developments urge a thorough update. First, at least five novel lectin domains have been identified in plants since 1998. Second, the results of a steadily increasing number of genome/transcriptome analyses revealed that most lectin families are far more widespread than can be inferred from the work with the purified proteins. Third, the whole of transcriptome/genome sequencing data indicated that chimerolectins are more widespread in plants than hololectins and provided evidence that most carbohydrate binding domains are found in one or more types of chimeric proteins. At present, 12 diVerent carbohydrate binding domains have been identified in plants. According to biochemical and transcriptome data, all these domains are found in expressed proteins. This section describes—in an alphabetic order—each of the carbohydrate binding domains and corresponding lectin families whereby ample attention is given to the chimerolectins and their domain architecture. Since the purified and characterized lectins represent only a minor fraction of all plant lectins, no adequate survey

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of the taxonomic distribution of the diVerent carbohydrate binding domains can be elaborated on the basis of the ‘‘lectin literature.’’ Therefore, a comprehensive screening was made of the publicly accessible databases (listed in Tables I and II) and the (unpublished) results used to generate a more realistic overview of the occurrence of the diVerent lectin families throughout the terrestrial plants (and if relevant other Viridiplantae). Since there are many errors and incorrect annotations in these databases, all retrieved sequences were carefully checked. Though the results are certainly not final, the analysis of the occurrence of the diVerent carbohydrate binding domains in the (near) completed plant genomes is fairly accurate. However, there remains some uncertainty about the number of genes and the sequences of the proteins (especially those encoded by intron‐containing genes). Screening of the transcriptome databases yielded a wealth of information with respect to the taxonomic distribution of some lectin families but the results should be interpreted with care for two reasons. First, transcriptome data are available for only a (very) limited number of species. Second, apart from a few species, the transcriptome covers only a (small) part of the whole genome. Hence, the results of the transcriptome databases are indicative rather than conclusive. B. DESCRIPTION OF THE DIFFERENT CARBOHYDRATE BINDING DOMAINS/FAMILIES

1. Agaricus bisporus agglutinin homologs Recent work with Marchantia polymorpha revealed that this liverwort expresses at least four functional homologs of lectins that until then were considered typical fungal lectins (Peumans et al., 2007). The prototype of this lectin family is the so‐called Agaricus bisporus agglutinin (ABA) which has been studied in great detail with respect to its biochemical properties, biological activities, and structure. Analysis of the purified lectins indicated that the Marchantia polymorpha ABA homologs (MarpoABA) are dimeric proteins built up of subunits comprising 140–142 amino acid residues (Fig. 1). Specificity studies revealed that MarpoABA (like ABA itself) preferentially interacts with the T antigen (Gal
1,3GalNAc) and that the aYnity of the lectin is strongly enhanced by the presence of high‐density polyvalent glycotopes (Nakamura‐Tsuruta et al., 2006; Wu et al., 2003). The structure of MarpoABA has not been resolved but modeling studies confirmed that this plant lectin has the same overall fold and three‐dimensional structure as the fungal homologs from A. bisporus and Xerocomus chrysenteron. X‐ray diVraction analysis demonstrated that the monomers of these fungal lectins consist of a
‐sandwich built from two bundles of
‐sheets interconnected

TABLE I Occurrence of the DiVerent Carbohydrate Binding Domains and Domain Architectures in (Near) Completed Embryophyta Genomesa Domain/ architecturesb

Physcomitrella patens

Selaginella moellendorYi

Populus trichocarpa

Arabidopsis thaliana

Ricinus communis

Oryza sativa

Sorghum bicolor

Zea mays

Medicago truncatula

Vitis vinifera


c




































þþþ

þþþþ
















þ



















þþ







þ



þþ




EUL 1‐Domain 2‐Domain Vacuolar

þþ þ


þþþ þþþ þþ

þþ

þ

þ

þ þþ

þþ þþ þþ

þþ þþ þ

þ

þ

GNA family GNA Cytoplasm GNA vacuole RLK S‐locGP X‐GNA



þ þ


þþ þþ þþ þ þþþþþ



þþ þþ




þþþ þþþ




þþ þþ


?
þþ þþ




þþ þþ


?
þþ þþ


?
þþ þþ




þþ þþ


ABA Amaranthins Hololectin Am‐aerolysin Am‐kinase CRA Cyanovirin

Hevein Hololectin Class I Chi Class IV Chi PR‐4 He‐RlpA


þ þ
þ

þþ þþ þ
þ


þþ þþ þ



þþ þþ þ



þþ þþ þ


þ þþ þþ þ



þþ þþ þ



þþ þþ þ



þþ þþ þ



þþ þþ þ


Jacalins GJRL MJRL VacJRL Dir‐Jac Jac‐Kelch Kinase‐Jac F‐Box‐Jac NBARC‐Jac


þ









þ







þ








þ

þ
þ



þ








þ
þ
þ
þ


þ
þ ? ? ? ?


þ
þ ? ? ? ?


þ








þ







Leg lectins LegLu LegL LegCyt RK

þþ

þþ

þþ

þþ

þþ

þþþ

þþ

þþþ

þþ

þþþ

þ

þþþ

þ

þþþ

þ

þþþ

þþ þþþ þþ þþþ

þþ

þþþ

LysM Hololec LysM‐Chi CEBiP M RLK

þ ? ? þ

? þ ? þþ



þþ þþ



þþ þþ



þþ þþ



þþ þþ



þþ þþ



þþ þþ



þþ þþ

þ
þþ þþ (continues)

TABLE I Domain/ architecturesb Nictaba N NsN NlN CPL N‐Fbox N‐TIR N‐AIG1 Ricin‐B Hololectin Type 2 RIP a

(continued)

Physcomitrella patens

Selaginella moellendorYi

Populus trichocarpa

Arabidopsis thaliana

Ricinus communis

Oryza sativa

Sorghum bicolor

Zea mays

Medicago truncatula

Vitis vinifera


þ þþ







þþþ




þþ þ
þþþ



þ þ þ
þþþ þ þþ

þ þþ þ
þþþ



? ? þ
þþ ? ?

? ? þ
þþ ? ?

? ? þþ
þþþ ? ?

þ þ þ
þþþ




þ þþ
þþþ











þþ







þþ






þ


þ









Based on the data deposited in the following databases: General database: NCBI: http://www.ncbi.nlm.nih.gov/; proteins, nucleotide sequences, ESTs, trace archives), last update: May 15, 2008 Specific databases: Arabidopsis thaliana: http://www.arabidopsis.org/ (release TAIR8) Physcomitrella patens: http://www.cosmoss.org/ and http://genome.jgi-psf.org/Phypa1_1/ (v1.1 repeat masked main genome sequence) Medicago truncatula: http://www.medicago.org/genome/ (Mt2.O release); http://www.tigr.org/tdb/e2k1/mta1/ (Medicago truncatula Gbrowse Mtr 1.0 pseudomolecule release) Oryza sativa: http://www.tigr.org/tdb/e2k1/osa1/ (Release 5) Populus trichocarpa: http://genome.jgi-psf.org/Poptr1_1/ (Assembly v1.1) Ricinus communis: http://castorbean.tigr.org/ (4X draft assembly) Selaginella moellendorYi: http://selaginella.genomics.purdue.edu/ [best Selaginella contig 12X (11/10/06)]; http://genome.jgi-psf.org/Selmo1/ (v1.0 assembly scaVolds) Sorghum bicolor: http://www.phytozome.net/sorghum; http://genome.jgi-psf.org/Sorbi1/ (v1.0 assembly scaVolds) Vitis vinifera: http://www.genoscope.cns.fr/cgi-bin/ and http://www.vitisgenome.it/ (Vitis vinifera 8X (08/30/2007) Zea mays: http://maize.tigr.org/ (AZM5 assembly) b Domain architectures are explained in Fig. 1 and Figs. 3–12. c Symbols:
, gene absent; þ, at least one gene present; þþ, multiple genes present; þþþ, complex gene families present; ?, remains to be determined.

TABLE II Overall Taxonomical Distribution of the Carbohydrate Binding Domains Found in Embryophytaa Viridiplantae Embryophyta

Carbohydrate binding domain

Bacteria

Fungi

Animals

Chlorophyta

Liverworts

Lycophytes

Mosses

Ferns

Gymnosperms

Angiosperms

ABA Amaranthin CRA Cyanovirin EUL GNA Hevein Jacalins Legume lectin LysM Nictaba Ricin‐B


b

þ
þ
þ þ þ
þ

þ

þ
þ þ þ þ þ
þ






þ
þ þ þ
þ







þ
þ þ



þ

þ þ þ þ
þ þ þ þ


þ
þ þ þ þ þ þ þ þ


(þ)


þ þ þ þ þ þ þ





þ þ þ þ þ þ þ þ



þ

þ þ þ þ þ þ þ



þ þ
þ þ þ þ þ þ þ þ

a

Based on the data deposited in the following databases: Same databases as in the legend to Table I; and some additional databases: Chlamydomonas reinhardtii: http://genome.jgi-psf.org/Chlre3/ (v3.0 assembly) Ostreococcus lucimarinus: http://genome.jgi-psf.org/Ost9901_3/ (Assembly v.2.0) Ostreococcus tauri: http://genome.jgi-psf.org/Ostta4/ (v.2.0 assembly) Volvox carteri: http://genome.jgi-psf.org/Volca1/ (Assembly v1.0) b Symbols:
, absent; þ, present; (þ), possibly present.

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ABA

Agaricus bisporus agglutinin homologs Domain architecture

ABA

Marchantia polymorpha

Example of purified protein ABA

MarpoABA

ABA

Fig. 1. Schematic overview of the identified domain architecture and purified protein with a domain equivalent to the Agaricus bisporus agglutinin (ABA). Only one domain architecture was identified. A representative lectin was isolated from Marchantia polymorpha (the structure of which is indicated in the box).

by a helix–loop–helix motif consisting of two short ‐helices. In addition, the structural analysis revealed that the ABA monomer comprises two distinct binding sites which are located on the opposite side of the helix–loop–helix motif and recognize the two diVerent configurations of a single epimeric hydroxyl (N‐acetylgalactosamine and N‐acetylglucosamine, respectively) (Carrizo et al., 2005). The combined carbohydrate binding sites apparently comprise the entire subunit of the ABA homologs. Hitherto no chimeric proteins comprising (an) ABA domain(s) or corresponding gene(s) have been identified. All identified plant homologs of ABA lack—like their fungal counterparts—a signal peptide and hence are synthesized on free ribosomes in the cytoplasm. MarpoABA undergoes, apart from the removal of the N‐terminal methionine and acetylation of the second amino acid (Ser or Thr), no posttranslational processing. Expression of an EGFP‐tagged MarpoABA isoform in tobacco BY‐2 cells indicated that the lectin resides in the nucleus and cytoplasm. A similar subcellular location has been demonstrated previously for the homolog from the nematophagous fungus Arthrobotrys oligospora (Rose´n et al., 1997). Immunolocalization studies confirmed that MarpoABA is also in the liverwort cells associated with the cytoplasm and, what is of paramount importance, ruled out the possibility that the lectin resides in a fungal endosymbiont (Peumans et al., 2007). Plant homologs of the A. bisporus agglutinin have been isolated exclusively from the liverwort Marchantia polymorpha. In addition, two (identical) expressed sequence tags were identified in the transcriptome of the moss Syntrichia ruralis (syn: Tortula ruralis). However, there is no proof that these two sequences are encoded by the moss cells and not by a possible fungal contaminant or endosymbiont. This caveat is certainly relevant because the genome of another moss (in casu Physcomitrella patens) does not comprise an ABA homolog. There are also no indications for the presence of

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homologous genes in the genome of other land plants (Tables I and II). Accordingly, it can be concluded that A. bisporus agglutinin homologs are confined to a (very) small part of the Embryophyta. Comprehensive screenings of the databases indicated that the A. bisporus agglutinin family is—apart from M. polymorpha and possibly T. ruralis— confined to fungi (Tables I and II). Two ESTs from the insect Aedes aegypti also encode an ABA homolog but no corresponding sequences are found in the genome, indicating that the ESTs are derived most likely from a contaminating fungus. Within fungi, ABA homologs and/or corresponding genes are documented in 20 species belonging to both Ascomycota and Basidiomycota. However, it should be emphasized that most (sequenced) fungal genomes lack ABA genes, which implies that the ABA family is certainly not ubiquitous among fungi. On the basis of the documented occurrence, it seems likely that the ABA domain has been developed by an early fungal ancestor (Fig. 2). The origin of the plant homologs is still

Unikonts CV-N

Nic Fungi

CRA

Ja

ABA

Animals

Am

Angiosperms

Lycophytes

EUL

Slime molds

Gymnosperms Ferns Mosses Liverworts

Excavates Chromoalveolates Glaucophytes Chromists

Chlorophyta (green algae)

Embryophyta

H

Alveolates

Archaea Bacteria

Last universal common ancestor

GNA

Le

LysM

Ri

Fig. 2. Schematic overview of the taxonomic origin of the carbohydrate binding domains found in modern plants. Filled block arrows indicate in which taxonomic division the direct ancestor of the domain most probably evolved. When the origin cannot be assigned to a well‐defined group but only to a lineage, the most probable point of origin is indicated by an open block arrow.

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very puzzling. Phylogenetic analyses revealed a clear anomaly in the dendrogram of the plant and fungal sequences. This phylogenetic discrepancy as well as the very limited occurrence in plants can be regarded as arguments (but certainly not as definitive proofs) in favor of a possible lateral transfer of an ABA gene from a fungus to a plant. 2. Amaranthins The term amaranthin was used for the first time as a trivial name for a lectin isolated from the seeds of A. caudatus. Later on, the term was used in a broader sense for a few lectins found in other Amaranthus species. The lectin from A. caudatus has been studied in detail at the biochemical and structural level. Amaranthin is a homodimer built of 33 kDa protomers comprising 303 amino acid residues. Comprehensive specificity studies demonstrated that the amaranthin preferentially recognizes the T‐antigen disaccharide Gal
(1,3) GalNAc (Rinderle et al., 1989). Structural analysis revealed that the amaranthin subunits comprise two tandem arrayed homologous domains of 150 amino acids. Each domain has a
‐trefoil structure formed by six strands of antiparallel
‐sheets capped by three
‐hairpins into a short
‐barrel. The two domains are linked by a short ‐helical 310 segment (Transue et al., 1997). Native amaranthin is built up of two protomers that are associated head‐to‐tail by noncovalent bonds so that the N‐terminal domain of one protomer faces the C‐terminal domain of the other protomer (Fig. 3). The two T‐antigen disaccharide binding sites are formed by two shallow depressions located at the interfaces of the facing N‐terminal and C‐terminal domains. This peculiar structural feature implies that the amaranthin domain itself possesses no sugar binding site(s) and that a specific spatial arrangement of two domains located on two separate subunits is required to establish a carbohydrate binding site. Hitherto, lectins from Amaranthus species are the only members of the amaranthin family that have been isolated and characterized. In addition, a ‘‘hessian fly responsive’’ gene has been identified in wheat that encodes the so‐called Hfr‐2 protein. This Hfr‐2 protein was described as a (chimeric) ‘‘cytolytic toxin’’ (Hfr‐2) built up of an N‐terminal domain homologous to the amaranthin protomer and a C‐terminal domain sharing a reasonably high sequence identity/similarity with part of the so‐called aerolysin domain (PuthoV et al., 2005). This domain corresponds to the lobe‐forming domain of the channel‐forming toxin aerolysin produced by Aeromonas sp. and some other bacteria. A thorough analysis of the publicly accessible databases of plant genomes and transcriptomes revealed that the amaranthin family is far more complex and fairly widespread among plants (Tables I and II). Besides the genuine

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Amaranthins

Am

Domain architectures

Am

Example of purified protein

Amaranthin domain (Prunus sp.)

Am

Am Amaranthin

Am Am

Am

Am

Amaranthus sp.

Zea mays

Am

Am

Am

Am

Am

Triticum aestivum

Aquilegia formosa ⫻ Aquilegia pubescens

Aerolysin domain Kinase domain

Fig. 3. Schematic overview of the identified domain architectures and purified proteins with amarantin domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

amaranthins and Hfr‐2, three additional types of proteins could be identified (Fig. 3). First, some species of the family Rosaceae express a complex mixture of proteins corresponding to a single amaranthin domain. Second, maize expresses a chimeric protein consisting of an N‐terminal domain comprising a single amaranthin domain and a C‐terminal domain equivalent to that of Hfr‐2. Third, a ‘‘receptor‐like kinase protein’’ was identified in the transcriptome of Aquilegia formosaAquilegia pubescens that consists of an N‐terminal amaranthin domain linked to a C‐terminal kinase domain. This means that the amaranthin family comprises at least two types of hololectins and three types of chimerolectins. However, it is still uncertain whether the amaranthin domains in the proteins other than the genuine amaranthins possess lectin activity.

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None of the identified proteins with an amaranthin domain possesses a signal peptide indicating that they are synthesized on free ribosomes in the cytoplasm. Amaranthin itself undergoes, apart from the removal of the N‐terminal methionine residue, no posttranslational processing (Raina and Datta, 1992). At present, no member of the amaranthin family has been localized at the subcellular level. However, preliminary experiments with an EGFP‐tagged single amaranthin domain protein from Prunus persica demonstrated that the protein resides in the nucleocytoplasmic compartment of tobacco BY‐2 cells (Van Damme et al., 2009a). It is interesting to note that the chimeric amaranthin/(protein) kinase from Aquilegia is also apparently synthesized without a signal peptide. This taken together with the fact that the protein lacks a transmembrane domain suggests that the Aquilegia chimer resides within the cells and hence cannot be considered a receptor kinase. Genuine amaranthins are apparently confined to the family Amaranthaceae. In contrast, homologs of Hfr‐2 are fairly widespread. cDNAs/genes could be retrieved, indeed, in species from distant taxa including Lycophyta (Selaginella), Coniferophyta (Picea, Pinus), Ginkgophyta (Ginkgo biloba), and numerous Magnoliophyta. Within the latter taxon, Hfr‐2 homologs are found in both Liliopsida (wheat and several other grasses) and Eudicotyledons. No homologs of the Aquilegia amaranthin kinase could be retrieved from any other plant genome/transcriptome indicating that this chimeric form might be confined to a single or a few taxonomic groups. Though the amaranthin domain is apparently fairly widespread, it should be emphasized that it is certainly not ubiquitous in plants because no corresponding genes are found in the genomes of P. patens, A. thaliana, Medicago truncatula, Populus trichocarpa, Oryza sativa, and Sorghum bicolor. The amaranthin domain of the genuine amaranthins shares no detectable sequence identity/similarity with any other known lectin or protein. However, BLASTp searches using the sequences of some amaranthin domains from Selaginella moellendorYi as a query indicated a residual similarity to the so‐ called Fascin‐like domain. This domain (119 residues) is the basic building block of the fascin proteins (also called singed proteins) which are a family of eukaryotic actin‐bundling/crosslinking proteins. Fascins were identified in sea urchin, Drosophila, Xenopus, rodents, and humans. They contain four copies of the fascin domain, which itself adopts a
‐trefoil topology and contains an internal threefold repeat. Since the two domains of amaranthin exhibit the same overall fold and topology, one can reasonably conclude that the amaranthin domain is structurally related to the fascin domain. Besides in fascins, the fascin or fascin‐like domain is also found in some bacterial, fungal, and plant proteins. For example, a glycosyl hydrolase family 5
‐ glucosidase was cloned from O. sativa that comprises an N‐terminal

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127

fascin‐like domain fused to a C‐terminal
‐glucosidase domain (Opassiri et al., 2007). It should be mentioned, however, that the fascin‐like domain found in this
‐glucosidase is not recognized in BLASTp searches using the amaranthin domain as a query. Taking into consideration the obvious residual sequence similarity and strong structural resemblance, it is tempting to speculate that the amaranthin domain is evolutionarily related to the fascin domain. If so, it seems likely that the ancestor of the modern amaranthin domain predates the origin of the Viridiplantae. As discussed above, most members of the amaranthin family are chimers of a single or double N‐terminal amaranthin domain fused to a domain that shares a striking sequence similarity with domains three and four of the
‐pore‐forming toxin aerolysin from the Gram‐negative bacterium Aeromonas hydrophila. Aerolysins are a family of cytolytic toxins exported by Aeromonas spp. (Buckley, 1992; Rossjohn et al., 1997). The mature toxins attach by their N‐terminal aerolysin/pertussis toxin domain to eukaryotic cells using glycosylphosphatidylinositol (GPI)‐anchored proteins. After binding, the toxin molecules oligomerize into heptamers, which by interaction of the aerolysin domains of the monomers form channels in the membranes and cause cell death. Besides in bacteria, (part of) the aerolysin domain is also found in some insect (e.g., yolk protein 1 from Hyphantria cunea) and vertebrate (e.g., natterins from the fish Thalassophryne nattereri) proteins. In addition, several fungal proteins have been identified that comprise a (partial) aerolysin domain. Interestingly, some of these fungal proteins (e.g., the hemolytic lectins LSLa and LSLb from Laetiporus sulphureus) are built up of an N‐terminal carbohydrate binding domain and a C‐terminal aerolysin domain (Manchen˜o et al., 2005), and in this respect can be considered structural/functional equivalents of the Hfr‐2 type plant proteins. Moreover, even though the respective carbohydrate binding domains share no detectable sequence similarity, the N‐terminal domain of LSLa and LSLb has a
‐trefoil scaVold very similar to that of the amaranthin domain. To reconstruct the overall molecular evolution of the plant amaranthin family, both the documented taxonomic distribution of the proteins with amaranthin domain(s) and the (residual) sequence/structural similarity with other known protein domains must be taken into consideration. Since it is evident that the Hfr‐2 type proteins are very widespread whereas the hololectins are confined to a few small taxonomic groups, it is tempting to speculate that the whole family evolved from an ancestral Hfr‐2 type chimeric protein. During evolution of the Spermatophyta, the overall architecture of the original chimeric protein remained unchanged. The only documented exception is the maize protein that apparently lost one of the amaranthin

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domains (Fig. 3). Despite the very broad taxonomic distribution over primitive as well as higher Embryophyta, Hfr‐2 homologs are definitively absent from at least half of the sequenced plant genomes. Hence, one can conclude that these genes have been purged from many genomes during evolution of the Embryophyta. To explain the origin of the genuine amaranthins, it is important to realize that these hololectins are apparently confined to (part of) the Amaranthaceae family. Taking into consideration that, for example, in Beta vulgaris genuine amaranthins coexist with the Hfr‐2 type protein, it seems likely that the hololectin originated from the chimeric protein through an evolutionary event whereby an Hfr‐2 type gene was duplicated with a concomitant loss of its aerolysin‐like domain. Though all evidence suggests that this evolutionary event took place during evolution of Amaranthaceae, it might have occurred earlier. A similar scenario applies to the origin of the single amaranthin domain hololectins that are found in some species of the Rosaceae family. Within this family, both single domain hololectins and Hfr‐2 type chimeric proteins occur though there is no evidence yet for a simultaneous occurrence in a single species. The only diVerence with the genuine amaranthins is that in the case of the Rosaceae hololectins, the parental Hfr‐2 type gene lost both its aerolysin‐like domain and one of its amaranthin domains (Fig. 3). Most probably, the underlying evolutionary event took place somewhere during the evolution of the Rosaceae family. In contrast to the origin of the hololectins, which apparently relies on gene duplication followed by domain loss, the evolution of the amaranthin kinase gene (found in Aquilegia) from a parental Hfr‐2 type gene requires a more complex mechanism. A possible—but still speculative—explanation is that a Hfr‐2 type gene was duplicated followed by a domain exchange with a protein kinase gene. Summarizing it can be concluded that the plant amaranthin family evolved most likely from an ancestral Hfr‐2 type gene that was already present in the genome of a very early terrestrial plant (Fig. 2). Taking into account the apparent residual sequence and structural similarity to the fascin and aerolysin‐like domain, respectively, the ancestral Hfr‐2 type gene is most probably not developed by plants but inherited from a common ancestral organism. There is no evidence for the occurrence of Hfr‐2 type genes in prokaryotes but it is obvious that the aerolysin domain and perhaps also the amaranthin/fascin domain have a bacterial origin. 3. Class V chitinase homologs with lectin activity In 2007, a novel lectin was described in the bark of the legume tree Robinia pseudoacacia (black locust) that was unrelated to the genuine legume lectins but shares nearly 50% sequence identity with plant class V chitinases

129

PLANT LECTINS

CRA

Class V chitinase homologs with lectin activity Example of purified protein

Domain architecture CRA

Robinia pseudoacacia

CRA RobpsCRP CRA

Signal peptide

Fig. 4. Schematic overview of the identified domain architecture and purified proteins with a domain equivalent to class V chitinases. Only one domain architecture was identified. A representative lectin was isolated from Robinia pseudoacacia (the structure of which is indicated in the box).

(glycosyl hydrolysase family GH18) (Van Damme et al., 2007a). This novel lectin, called R. pseudoacacia chitinase‐related agglutinin or RobpsCRA is a homodimer of 337 amino acid residue subunits (Fig. 4). Though the lectin shares a high sequence identity with class V chitinases, it is essentially devoid of chitinase activity. RobpsCRA is a weak agglutinin but definitely interacts with a fairly high aYnity with high mannose N‐glycans comprising the proximal pentasaccharide core structure. The structure of RobpsCRA has not been resolved yet but molecular modeling confirmed that the lectin has the same overall fold and three‐ dimensional structure as the human chitotriosidase (hMChi) that of all resolved GH18 proteins shares the highest sequence identity/similarity with RobpsCRA (Fusetti et al., 2002). According to the model, RobpsCRA adopts virtually the same TIM‐barrel fold as hMChi. This fold consists of an inner crown of
‐sheet strands surrounded by an outer crown of ‐helices, and an additional hairpin‐shaped loop composed of three antiparallel strands of
‐sheet that protrudes from one edge of the TIM‐barrel structure. Modeling further indicated that the groove, which normally accommodates the chitin, cannot properly stack the substrate because several solvent‐ exposed hydrophobic residues are replaced by hydrophilic residues in RobpsCRA. Unfortunately, molecular modeling did not yield any indications about the possible location and structure of the carbohydrate binding site. RobpsCRA is synthesized with a signal peptide and follows the secretory pathway. After co‐translational removal of the signal peptide, the protein undergoes no further modifications (and accordingly is not glycosylated).

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Until now, only a single class V chitinase homolog with lectin activity has been isolated from the legume tree R. pseudoacacia (Tables I and II). Closely related homologs are expressed in several other legumes (Medicago truncatula, Lotus japonicus, and Glycine max). There are indications for the presence of similar proteins in other legumes suggesting that chitinase‐related lectins are confined to (part of ) the family Fabaceae. Though rather inconspicuous, the identification of the RobpsCRA family is important because it sheds a new light on the evolution of plant lectins. All evidence suggests that at a given time in the evolution of the Fabaceae family, some species developed a novel carbohydrate binding domain starting from an existing structural scaVold with a diVerent activity (Fig. 2). This implies that there is at least one documented example of a plant carbohydrate binding domain acquired through neofunctionalization. Interestingly, the very same class V chitinases domain was also used by mammals to develop a novel sugar binding domain by a similar (but independent) neofunctionalization process (Van Damme et al., 2007a). 4. Cyanovirin family The Cyanovirin family is named after a virucidal protein that was originally isolated from extracts of the cyanobacterium (blue green alga) Nostoc ellipsosporum in the framework of a screening program setup to identify novel anti‐HIV agents (Boyd et al., 1997). This protein, which was called cyanovirin‐N (CV‐N), irreversibly inactivates human immunodeficiency virus (HIV) and simian immunodeficiency virus. Since the very strong anti‐ HIV activity relies on high aYnity interactions of CV‐N with high‐mannose N‐glycans on the viral surface envelope glycoprotein gp120, it is evident that the protein possesses carbohydrate binding activity and hence can be considered a lectin. CV‐N consists of a single 101 amino acid residue polypeptide with an internal duplication (Gustafson et al., 1997; Fig. 5). NMR and crystallographic studies resolved the structure of CV‐N. Both repeats form equivalent elongated structures built up of a triple‐stranded
‐sheet and a
‐hairpin. Though the two internal repeats readily superimpose, they do not form separate domains because the overall ellipsoid fold depends on multiple contacts between them (Bewley et al., 1998). Two symmetrically related monomers form a dimer by domain swapping in such a way that domain A of one monomer interacts with domain B of its crystallographic symmetry mate and vice versa (Yang et al., 1999). Each CV‐N monomer contains two binding sites located in deep clefts at opposite ends of the elongated structure. Besides N. ellipsosporum, only a few other bacteria contain genes encoding proteins similar to CV‐N. Recently, similar proteins (or corresponding genes/

PLANT LECTINS

Cyanovirin family

CV-N

Domain architecture CV-N

131

Ceratopteris richardii

Example of purified protein CV-N Recombinant CrCVNH from Ceratopteris richardii CV-N

Signal peptide

Fig. 5. Schematic overview of the identified domain architecture and purified protein with a cyanovirin‐N domain. Only one domain architecture was identified. A (recombinant form) of a representative lectin from Ceratopteris richardii was isolated and characterized (the structure of which is indicated in the box).

cDNAs) were identified in several fungi and in the fern Ceratopteris richardii (Percudani et al., 2005). In the mean time, the structure of the (recombinant) C. richardii CV‐N homolog (called CrCVNH) has been resolved (Koharudin et al., 2008) and deposited in the PBD database (PDB: 2JZJ). No similar proteins/genes could be identified in any seed plant. However, comprehensive BLAST searches revealed that the genome of the lycopsid S. moellendorYi contains a complex set of at least 30 genes encoding proteins that are definitely related to CV‐N. For some of these genes, corresponding ESTs have been deposited in the database indicating that S. moellendorYi expresses multiple CV‐N homologs. In addition, a single expressed sequence tag encoding a CV‐N homolog could be identified in the transcriptome of the liverwort M. polymorpha. In contrast to the (numerous) fungal CV‐N homologs, which are all synthesized without a signal peptide, all plant homologs are synthesized with a signal peptide and accordingly follow the secretory pathway. In addition to a signal peptide, the plant proteins also contain an extra C‐terminal fragment that is absent from CN‐V and its fungal homologs. It is still unknown whether this C‐terminal sequence is posttranslationally processed and/or plays a role in the targeting of CV‐N in the plant cell. The current documented occurrence indicates that the CV‐N domain is predominantly found in fungi and more precisely in Ascomycota (Tables I and II). Though there are also a few bacteria that produce CV‐N homologs, a prokaryotic origin of the domain is not evident (Fig. 2). On the contrary, the patchy distribution over only a few species might be indicative for multiple lateral transfers of a CV‐N gene from a fungus to a bacterium. In plants, the

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CV‐N domain seems to be confined to the more primitive Embryophyta (liverworts, Lycopsids, and ferns). It was suggested that the plant and fungal CV‐N domains have a common origin but ‘‘amplified separately in fungi and seedless plants following the separation of these two lineages’’ (Percudani et al., 2005). If so, an early ancestor of all Embryophyta acquired an ancestral CV‐N gene from a common ancestor of fungi and plants. During further evolution of plants, the genes were retained in only a few ‘‘primitive’’ taxa but were purged from the genome of some other lower Embryophyta (e.g., mosses) and all Spermatophyta. According to this scenario, the CV‐N genes must also have been purged from the genome of all Metazoa as well as from that of the Basidiomycota subgroup of the fungi. Vertical inheritance of plant CV‐N genes raises an additional problem because all plant CV‐N homologs contain a signal peptide and a C‐terminal peptide that are both absent from their fungal counterparts. This implies that the ancestral plant gene already acquired two extra sequences. Though such events are diYcult to trace the fact that some genes of S. moellendorYi are interrupted by a phase 0 intron just behind the signal peptide might be indicative for the conversion of a cytoplasmic into a vacuolar/extracellular CV‐N. Hitherto, there is no evidence for the occurrence in plants of chimeric proteins with a CV‐N domain. The same applies to bacteria. However, in fungi, several types of chimeric proteins or corresponding genes have been identified. Interestingly, in one of these fungal chimeric proteins, the CV‐N domain is interrupted by a LysM domain at the boundary between the two internal repeats. As already discussed, the CV‐N domain itself consists of two internal repeats. According to phylogenetic analysis of the individual subdomains of a set of bacterial, fungal, and plant CV‐N sequences, the CV‐N domain arose from a unique ancient duplication that took place in a common (but not explicitly named) ancestor (Percudani et al., 2005).

5. EEA family Lectins from Euonymus europaeus and some other spindle tree species have already been isolated and characterized in the mid 1970s (Paca´k and Kocourek, 1975; Petryniak et al., 1977) but could due to the lack of sequence information not be classified yet. A recent reinvestigation of the lectin and cloning of the corresponding gene revealed that the ‘‘old’’ Euonymus europaeus agglutinin (EEA) represents a novel family of plant lectins (Fouquaert et al., 2008). EEA is a homodimeric protein composed of 152 residue subunits (Fig. 6). Glycan array screening demonstrated that the lectin interacts with both blood group B oligosaccharides and high‐mannose N‐glycans. Since no information is available yet about the structure of EEA, it is still

133

PLANT LECTINS

EUL

EEA family Domain architectures

Example of purified protein

Euonymus europaeus

EUL

EUL

EEA

O. sativa

EUL

EUL

EUL

EUL

EUL

O. sativa

O. sativa

O. sativa

S. moellendorffii

EUL

M. polymorpha

EUL

M. polymorpha

EUL

EUL

O. sativa

EUL

EUL

EUL

Zea mays Signal peptide

Fig. 6. Schematic overview of the identified domain architectures and purified proteins with EUL domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

unclear whether the dual specificity of the lectin relies on the presence of two distinct carbohydrate binding sites. At present, EEA is the only purified and characterized member of this newly identified lectin family. However, BLAST searches revealed that most, if not all, Embryophyta express (a) protein(s) with one or two domain equivalent to the EEA polypeptide. Some of these proteins consist exclusively of an Euonymus lectin (EUL) domain but most of them contain an extra N‐terminal sequence (other than a signal peptide) varying in length between

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5 and up to 200 amino acid residues. In the case of the two‐domain proteins, additional heterogeneity arises from diVerences in the length (and sequence) of the interdomain linker. Furthermore, some proteins contain an extra C‐terminal sequence/domain. On top of this, the overall picture is further complicated by the occurrence of putative proteins with a signal peptide. Since none of the extra domains corresponds to any known structural unit or motif, it is diYcult to make a clear distinction between holo‐ and chimerolectins. For example, an extra N‐terminal or a C‐terminal sequence of less than 20 amino acid residues can hardly be regarded as an extra domain, whereas some long (>200 amino acids) N‐terminal sequences definitely represent a unique structural element. The great majority of all proteins with EUL domain(s) lacks a signal peptide and accordingly are believed to be synthesized on free ribosomes in the cytoplasm. In the case of the Euonymus lectin, the primary translation product is posttranslationally processed by the removal of the first six amino acid residues (Fouquaert et al., 2008). At present, no data have been reported on the subcellular location of proteins with EUL domain(s). However, confocal microscopy of EGFP‐tagged EEA confirmed that in tobacco BY‐2 cells, the lectin resides in the cytoplasm and nucleus. Similar results were obtained with an EGFP‐tagged chimeric homolog (with a long N‐terminal domain) from A. thaliana. In some species, cDNAs/genes could be identified that encode proteins comprising a single EUL domain preceded by a signal peptide. Given the obvious presence of a signal peptide, one can reasonably assume that these particular proteins are synthesized on the endoplasmic reticulum (ER) and follow the secretory pathway. At present, the final destination of the latter proteins is still unknown but to distinguish them from their cytoplasmic/ nuclear homologs, they are referred to as ‘‘vacuolar forms.’’ The identification of the EUL domain as a novel carbohydrate binding module is not only important for the classification of plant lectins but also sheds a new light on the possible function of these lectins in the plant. Some previously identified abscisic acid and salt stress‐responsive proteins in rice and wheat possess one or two EUL domains. This taken together with the fact that proteins with EUL domain(s) are apparently ubiquitous in terrestrial plants might be an indication that lectin‐mediated protein– glycoconjugate interactions play an essential role in some cellular processes in Embryophyta. Though EEA is the only lectin with an EUL domain that has been isolated and characterized, the sequences deposited in the genome and transcriptome databases prove that this carbohydrate binding domain is found in Marchantiophyta, Bryophyta, Filicophyta, Gymnosperms and Angiosperms, mosses,

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and hence is most probably ubiquitous among Embryophyta (Tables I and II). However, as is illustrated below by the overview of the (near) completed plant genomes, not all types of proteins with EUL domain(s) are ubiquitous. S. moellendorYi possesses the most complex (identified) complement of EUL genes comprising 12 and 16 genes encoding single and double domain cytoplasmic proteins, respectively, plus an additional set of six genes encoding (putative) vacuolar single domain homologs. All Poaceae also possess (multiple) genes encoding single and double domain cytoplasmic EUL proteins, and in S. bicolor and Zea mays but not in O. sativa vacuolar homologs are present (and expressed). Also in the moss Physcomitrella, single and double domain genes are found. In contrast, only one or two genes encoding a single domain EUL protein are found in dicotyledonous species. Transcriptome data confirmed that dicots express exclusively single EUL domain proteins but also revealed that in some of them (e.g., Lactuca and Helianthus sp.), the expression of these proteins is controlled by a fairly complex set of genes (up to 20). Gymnosperms express multiple forms of both single and double EUL domain proteins. No exact figure can be given but based on the diversity of the expressed proteins, the genomes of Pinus taeda and Picea sitchensis contain at least 20 EUL genes. In the liverwort M. polymorpha, two diVerently expressed single EUL domain proteins could be identified. No sequences encoding a double domain protein could be retrieved but this does not imply that the corresponding gene(s) are absent from the genome. Since the EUL domain is apparently confined to land plants, it was most probably developed within this taxonomic group. Taking into account the current taxonomic distribution, the EUL must already have been present in the common ancestor of all modern Embryophyta (Fig. 2). Since there is no evidence for the presence in any other line of the Viridiplantae, the best guess is that the EUL domain evolved after the Embryophyta lineage separated from the other Viridiplantae. Once acquired, the EUL domain was retained in the genome of all land plants. Apart from a few exceptions, the sequence of EUL domain itself as well as the domain architecture was fairly well conserved. The complex complement of single and double domain proteins found in P. patens and S. moellendorYi indicates that domain duplication as well as gene duplication occurred during early evolution of land plants. A basic set of single and double domain genes was apparently also retained in ferns and gymnosperms, and in monocotyledonous angiosperms. Other angiosperms possess a much simpler complement of EUL genes. Most dicot species contain a single EUL gene encoding a single domain protein, indeed, indicating that the greater part of the EUL gene complement was purged from the genome after the dicots (including basal Magnoliophyta, Magnoliids, and core

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Eudicotyledons) diverged from the monocots. Some dicots possess (and express) multiple genes. However, it seems that in these cases, a single original ‘‘dicot’’ EUL gene was amplified. 6. GNA family Cloning and X‐ray diVraction analysis revealed that a mannose binding lectin that was originally isolated from the bulbs of the snowdrop (Galanthus nivalis) (Van Damme et al., 1987) represented a novel family of carbohydrate binding plant proteins with a unique three‐dimensional structure. In the mean time, homologs of the so‐called Galanthus nivalis agglutinin (GNA) have been identified in numerous plant species. Since in the beginning, these lectins were exclusively isolated from monocot species, the term ‘‘monocot mannose‐binding lectins’’ was introduced in the mid 1990s as a collective noun for a group of closely related lectins that exhibit an exclusive specificity toward mannose and until then seemed to be confined to a few taxa of the monocotyledons (Liliopsida) (Van Damme et al., 1995). However, this term has become too restrictive since very similar lectins have been identified in plants other than Liliopsida as well as in bacteria (Parret et al., 2003, 2005) and animals (Tsutsui et al., 2003). Therefore, this lectin family is renamed after the first identified member, in casu the Galanthus nivalis agglutinin or GNA (Van Damme et al., 2004a). For the same reason, the carbohydrate binding domain equivalent to the snowdrop lectin is referred to as the ‘‘GNA domain.’’ X‐ray diVraction analysis of several GNA‐related lectins demonstrated that the overall fold of the GNA domain is fairly well conserved. Three four‐stranded
‐sheets organized around a pseudosymmetry axis form a typical
‐prism fold (Hester et al., 1995). Each of the three
‐sheets correspond to one of the three in tandem arrayed homologous subdomains of the GNA domain and harbor a mannose binding site. GNA was originally considered a mannose‐specific lectin that according to structural analyses possesses three similar mannose binding sites per subunit. However, more recent work demonstrated that GNA (and other related lectins) interact only weakly with mannose but exhibit a strong aYnity toward oligomannosides and high‐mannose N‐glycans (Van Damme et al., 2009b). In addition, a reinvestigation of the specificity of a set of two‐domain lectins revealed that some GNA domains specifically interact with complex N‐glycans but are irreactive toward oligomannosides/high‐mannose N‐glycans (Van Damme et al., 2007b). By virtue of the simultaneous presence of both specificity types, some two‐domain lectins (e.g., the tulip lectin TxLC‐I, Van Damme et al., 1996b) are capable of interacting with both high‐mannose and complex N‐glycans. A general remark to be made is that

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the fine specificity of most studied GNA‐related lectins is far from understood. High performance analytical techniques (like glycan microarray analysis) yielded surprising results for diVerent members of the GNA family. Most probably, the specificity of each single GNA domain is a complex composite of the individual specificities of each of the three subdomains. Thereby it should be mentioned that in some lectins, one or two sites might be inactive (Barre et al., 1996). In the case of the sweet protein curculin (Curculigo latifolia fruits), all three sites of the GNA domain are inactive and hence the protein is completely devoid of sugar binding activity (Barre et al., 1997). GNA is synthesized on the ER as a preproprotein comprising a signal peptide followed by a carbohydrate binding domain of 108 amino acid residues and a C‐terminal propeptide (Van Damme and Peumans, 1988; Van Damme et al., 1991). Co‐translational removal of the signal peptide and posttranslational cleavage of the C‐terminal propeptide yields the mature lectin polypeptide. The C‐terminal propeptide is required for proper biosynthesis because it prevents the proprotein to interact with N‐glycosylated proteins present in the lumen of the ER. Though the exact subcellular location has not been determined yet, experiments with EGFP‐tagged GNA clearly demonstrated that the lectin is targeted into the ‘‘vacuolar’’ compartment of tobacco BY‐2 cells (Fouquaert et al., 2007). In contrast to GNA, related lectins from fungi and fishes are synthesized without a signal peptide and presumably reside in the cytoplasmic/nuclear cell compartment (Fouquaert et al., 2006, 2007). It appears, therefore, that depending on the (taxonomic) source, the very same GNA domain is embedded in two diVerent ‘‘molecular environments’’ and accordingly targeted into a diVerent subcellular compartment. To distinguish the genes comprising a sole GNA domain from genes in which the domain is inserted between a signal peptide and a C‐terminal propeptide, they will further be referred to as cytoplasmic and vacuolar GNA homologs, respectively. Hitherto, only vacuolar GNA‐like lectins have been purified from plants. Biochemical analyses of numerous lectins combined with molecular cloning of the corresponding genes demonstrated that the family of GNA‐related plant lectins is a composite of proteins with diVerent molecular structures. One‐domain GNA‐related lectins are composed of 12–14 kDa subunits and exist either as monomers, dimers, or tetramers (Fig. 7). In some lectin genes, the GNA domain is duplicated giving rise to so‐called two‐domain lectins. The two‐domain protomers exist as such or associate into dimers or tetramers giving rise to complex lectin molecules. In some lectins, the composing GNA domains are nearly identical, whereas in others they share only a limited sequence identity. In some but not all cases, the two‐domain

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GNA

GNA family

Domain architectures GNA

Example of purified proteins GNA GNA

Galanthus nivalis

GNA GNA

GNA

GNA

GNA GNA

Monocot sp.

GNA

Selaginella moellendorffii

GNA

GNA

ASA

GNA

GNA GNA

GNA

GNA

GNA GNA

GNA

Brassicaceae

GNA

TxLC-I

Brassicaceae

GNA Selaginella moellendorffii

Signal peptide

PAN/APPLE-like domain

C-terminal peptide

Transmembrane domain

S-locus domain

Kinase domain Unknown N-terminal domain

Fig. 7. Schematic overview of the identified domain architectures and purified proteins with GNA domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

protomers are (partly) cleaved into two polypeptides of (nearly) equal size (Van Damme et al., 1995, 2007b). Vacuolar homologs of GNA have been isolated not only from numerous monocots but also from one dicot (e.g., Hernandia moerenhoutiana; family Hernandiaceae), one gymnosperm (Taxus media, family Taxaceae) (Kai et al., 2004), and one liverwort (M. polymorpha, family Marchantiaceae) (Peumans et al., 2002). An updated screening of genome/transcriptome databases indicated that GNA homologs are expressed by species belonging to the basal Magnoliophyta, the Magnoliids, and the stem Eudicotyledons but not by core Eudicotyledons. In addition, genes encoding vacuolar GNA homologs are found in the genome of S. moellendorYi.

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As reported previously, a few ESTs encoding cytoplasmic GNA homologs were retrieved in the transcriptomes of Z. mays, Triticum aestivum, and M. truncatula (Van Damme et al., 2004a). Since only a total of six ESTs were found and all six sequences were virtually identical to one of the GNA‐like lectins expressed by Fusarium oxysporum, there is some uncertainty about a possible fungal origin of the presumed plant ESTs. However, a screening of the S. moellendorYi genome revealed the presence of at least four diVerent genes encoding a cytoplasmic GNA homolog. Since one of these genes is expressed, one can reasonably conclude that cytoplasmic GNA homologs do occur in plants. Interestingly, the cytoplasmic forms found in the Selaginella genome coexist with multiple vacuolar forms. Additional evidence for the occurrence of cytoplasmic GNAs in plants is provided by an analysis of the transcriptome of flower buds from the basal magnoliophyte Illicium parviflorum (family Schisandraceae). Four percent of the total EST population encodes a mixture of three diVerent cytoplasmic GNAs. Taking into account the abundance of the lectin cDNAs and the fact that the sequences share no high similarity with any known fungal lectin gene, there is little doubt that I. parviflorum ESTs are of a plant origin. The great majority of all identified and characterized plant proteins with GNA domain(s) are hololectins. However, plants also express diVerent types of chimeric proteins with a GNA domain. A well‐known family of such chimeric proteins is the S‐locus glycoproteins from Brassica and other Brassicaceae species, which plays a role in self‐incompatibility and is encoded by extended gene families. The corresponding genes comprise a signal peptide followed by a GNA domain, an S‐locus glycoprotein domain, and a C‐terminal plant PAN/ APPLE‐like (PAN‐A) domain (Fig. 7). Homologs of the S‐locus glycoproteins were also identified in species from other families (e.g., the epidermis‐specific secreted glycoprotein EP1 from Daucus carota and SIEP1L protein from Beta vulgaris) and are probably widespread in seed plants. However, it should be mentioned that in many of the putative S‐locus glycoprotein homologs, either the S‐locus or PAN/APPLE‐like domain or both are poorly conserved. Genes with a domain structure similar to that of the S‐locus glycoprotein genes are also present in the genomes of P. patens and S. moellendorYi. A second type of chimeric proteins with a GNA domain are the S‐locus protein kinases found in Brassicaceae and homologous receptor kinases from other species. S‐locus protein kinases comprise the same domains as the S‐locus glycoproteins but possess an additional transmembrane domain (TMD) and a C‐terminal protein kinase domain (Fig. 7). According to genome/transcriptome data, receptor kinases with a GNA domain are ubiquitous among Embryophyta (except possibly the Marchantiophyta). In most if not all species, the receptor kinases are under the control of a complex gene family.

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Though both the S‐locus glycoproteins and S‐locus receptor kinases and their respective homologs comprise a domain that shares a clearly detectable sequence similarity with the GNA domain, it still remains to be demonstrated that these domains possess carbohydrate binding activity and if so whether they interact with the same glycans as the genuine GNA‐related lectins. Proteins with one or more GNA domain(s) are found in nearly all groups of Eukaryota and bacteria but not in Archaea (Tables I and II). Since the GNA domain is fairly widespread in most major bacterial phyla, one can reasonably assume that it was developed within the Prokaryota and subsequently transferred to eukaryotes (Fig. 2). During evolution and divergence of the eukaryotes, the GNA domain was retained in most but certainly not all lineages. Moreover, even within a single lineage, an extensive loss of the domain occurred. For example, within the vertebrates, the GNA domain is confined to a few fish species. Similarly, within the fungi lineage, GNA domains are exclusively found in a single subgroup of the Ascomycota. The same applies to the Viridiplantae because according to the available information, the GNA domain is exclusively retained in the Embryophyta. Because of the apparent absence from all other Viridiplantae lineages, it is diYcult to trace the origin of the modern plant GNA domains. However, based on the documented taxonomic distribution, it seems likely that the GNA domain was already present in a very early embryophyte. Taking into account that all animal and fungal GNA homologs are cytoplasmic forms, it is tempting to speculate that the Viridiplantae lineage also acquired a gene encoding a cytoplasmic GNA homolog from a common eukaryotic ancestor. This assumption is supported by the presence in the genome of S. moellendorYi of multiple genes with a sole GNA domain. The origin of the far more widespread vacuolar forms can be explained by duplication of a gene with a sole GNA domain followed by insertion between a signal peptide and a C‐terminal propeptide (Van Damme et al., 2004a). Since vacuolar forms are, besides in seed plants, also found in M. polymorpha and S. moellendorYi, the evolutionary event leading to the vacuolar homologs took place most probably in an ancestor of all modern Embryophyta lineages. At present, cytoplasmic and vacuolar forms still coexist in S. moellendorYi but apparently not in any other species for which an adequate genomic coverage is available. This implies that during evolution of the Embryophyta, the cytoplasmic forms were lost from most lineages, whereas the vacuolar homologs were retained in liverworts, lycophytes, and part of the gymnosperms and angiosperms. Though this scenario can readily be reconciled with the documented taxonomic distribution of the GNA‐related lectins, it cannot explain the occurrence of a whole family of cytoplasmic forms in I. parviflorum.

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Besides single domain lectins, the GNA family also comprises a whole set of lectins with a duplicated GNA domain. The origin of these lectins relies most probably on an evolutionary event involving a gene duplication accompanied by domain duplication. Hitherto, two‐domain lectins were exclusively found in monocots. According to a detailed phylogenetic analysis, these proteins do not represent a monophylic group but result from multiple independent domain duplication in tandem insertion events (Van Damme et al., 2007b). The domain architecture of the documented chimeric plant proteins with a GNA domain is indicative for gene duplications accompanied by domain fusion(s). However, it is not clear whether a single fusion took place between a GNA domain and an existing multidomain module (S‐locus/PAN‐A or S‐locus/PAN‐A/TMD/kinase) or the complex genes were formed by multiple successive additions of a single domain. A second problem concerns the relationship between the S‐locus glycoprotein homologs and receptor kinase homologs. Though one tends to assume that the receptor kinase results from a fusion between an S‐locus glycoprotein module and a TMD/kinase module (at least for some genes), it may be the other way around. Some S‐locus glycoprotein genes from Arabidopsis correspond, indeed, to truncated receptor kinase genes. Both S‐locus glycoprotein homologs and receptor kinase homologs are, apart from M. polymorpha, found in all Embryophyta suggesting that their origin predates at least the separation of the moss and vascular plant lineages. 7. Proteins with hevein domains Most plant proteins that were considered as typical chitin binding proteins in the past owe their carbohydrate binding activity to a single or multiple so‐called hevein domains. This particular lectin domain was named after hevein, a 43 amino acid polypeptide present in the latex of Hevea brasiliensis (rubber tree) (Waljuno et al., 1975). As has been corroborated in detail in several previous review papers, the family of chitin binding proteins with hevein domains is highly diverse not only in terms of domain architecture of the mature protomers/proteins but also for what concerns the sometimes complex posttranslational processing of the primary translation products (Raikhel et al., 1993; Van Damme et al., 1998a, b, 2007c). Numerous hololectins have been described that are built up of one or more protomers comprising a single or multiple tandem arrayed hevein domains. In the multidomain protomers, two, three, four, or seven individual hevein domains are separated from each other by short (5–10 residues) linker sequences (Fig. 8). Besides hololectins, several types of chimeric proteins with hevein domain(s) are found in plants. Classical examples are the

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Proteins with hevein domain(s)

H

Domain architectures

Examples of purified proteins

H

Solanum tuberosum

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

H

PL-D H

H

Phytolacca americana

H

H

H

H

H

H

H

H

H

Phytolacca americana

H

H

H

H

H

H

H

H

H

H

WGA

PL-C

Triticum sp.

H

H

H

PL-B

Phytolacca americana

Class-I/IV chitinase; ubiquitous

H

H

Bean class-chitinase Euonymus hevein-like protein

H

H

Class-I chitinase; Brassica juncea

H

H

H

H

H

Brassica juncea class-I chitinase UDA

Barley PR4

H PR4; ubiquitous

H

H

H

H

H

H

H Solanum tuberosum

H

Hevein

H

H

H

Potato lectin

Physcomitrella patens

Chitinase domain Barwin domain

Signal peptide Ser/Pro-rich linker

RlpA domain

Fig. 8. Schematic overview of the identified domain architectures and purified proteins with hevein domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

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class I and class IV chitinases comprising an N‐terminal hevein domain linked to a C‐terminal chitinase domain (Beintema, 1994; Linthorst, 1991). In a few chitinases, the N‐terminal hevein domain is duplicated. Another common chimeric protein consists of an N‐terminal hevein domain linked to a C‐terminal domain equivalent to the so‐called Barwin domain [also known as a pathogenesis‐related (PR) 4 protein]. A less common and completely diVerent chimeric architecture is found in the potato lectin, which consists of two modules of double hevein domains separated by a long (30–50 residue) serine‐proline rich linker (Van Damme et al., 2004b). Though the potato lectin is still considered the prototype of a subfamily of presumed closely related Solanaceae lectins, recent work indicates that the homologs from tomato have a diVerent molecular architecture both for what concerns the number of hevein domains as well as the length of the linker sequences between the hevein domains (Peumans and Van Damme, 2008; Peumans et al., 2003). In addition to these well‐known proteins, genome/transcriptome sequencing data revealed the occurrence in a few primitive Embryophyta of a novel type of chimeric protein comprising an N‐terminal hevein domain and a C‐terminal domain equivalent to the ‘‘Rare lipoprotein A (RlpA)‐like double‐ psi beta‐barrel.’’ RlpA contains a conserved region with a double‐psi ß‐barrel (DPBB) fold. This DPBB fold often possesses an enzymatic domain and is also found in the N‐terminus of pollen allergens. Transcriptome data confirmed that hevein‐DPBB genes are expressed in M. polymorpha, P. patens, and S. moellendorYi but not in seed plants. Since no corresponding protein has been isolated, yet it remains unclear whether the expressed hevein‐DPBBs remain intact or are possibly posttranslationally cleaved. All plant proteins with hevein domain(s) are synthesized with a signal peptide and enter the secretory pathway. Many but not all proteins are further processed by one or more posttranslational modifications. Cleavage of a short C‐terminal propeptide, which in some cases acts as a vacuolar targeting sequence (e.g., in the barley lectin and in some class I chitinases), is documented for both hololectins (e.g., barley lectin) and chimerolectins (e.g., class I chitinases). In other cases, large chimeric precursors are converted into small hololectins through the cleavage of a long C‐terminal propeptide (Lee et al., 1991; Raikhel et al., 1993; Soedjanaatmadja et al., 1995; Van Damme et al., 2007c). For example, hevein itself is produced from a barwin‐like precursor through the removal of the barwin domain. Similarly, the Urtica dioica lectin is the eventual remnant of a long precursor with a C‐terminal chitinase‐like domain (Lerner and Raikhel, 1992). Besides proteolytic modifications, several types of glycosylation are documented. A few proteins (e.g., one of the Phytolocca americana lectins and some class I chitinases) are N‐glycosylated. A totally diVerent mode of glycosylation occurs in

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some Solanaceae lectins (e.g., potato and tomato lectins) which are extensively substituted in their serine/proline rich linker by serine‐linked galactosyl and hydroxyproline‐linked (oligo)arabinosides (Kieliszewski et al., 1994). Several proteins with one or more hevein domains have been analyzed at the structural level. Hevein itself consists of a single 43 amino acid polypeptide that forms a typical structural motif consisting of three antiparallel strands of
‐sheet (
1,
2,
3) associated to two short ‐helical segments (1, 2) (Aboitiz et al., 2004; Andersen et al., 1993). The overall fold is stabilized by four disulfide bridges (between eight extremely conserved Cys residues) that tightly maintain the backbone fold of the secondary structures. Structural analysis of the binding site was achieved by X‐ray crystallography with tri‐N‐acetylchitotriose (NAG3) complexes. Basically, the sugar binding site consists of three subsites. Three aromatic amino acid residues and one serine residue participate to the NAG3 binding. Though the results of the structural analyses fully explain the binding of chito‐oligosaccharides, it remains unclear to what extent the conclusions can be extrapolated to the binding of N‐glycans into the hevein domain. Though the hevein domain is classically considered a typical chitin binding unit, early work with, for example, WGA and UDA already indicated that these proteins also interact with diVerent types of animal glycoproteins. A reinvestigation of the specificity of a quite extended set of lectins with hevein domains using novel high performing analysis techniques (e.g., glycan microarray analysis) not only confirmed the interaction with animal glycoproteins but also revealed that hevein domains have a (much) higher aYnity for (high mannose and/or complex) N‐glycans than for chito‐oligosaccharides (Van Damme et al., 2009b). This implies that the hevein domains—or at least those found in plants—should no longer be regarded as typical chitin binding motifs but as structural units with a more or less pronounced specificity toward N‐glycans. Though the great majority of all genes/proteins with hevein domain(s) were identified in (seed) plants and fungi, genome/transcriptome analyses provide evidence for the occurrence of hevein domains in at least a few other taxa (Tables I and II). For example, a large chimeric protein with an internal hevein domain was identified in the diatom Thalassiosira pseudonana. Some proteins from Caenorhabditis elegans also comprise a small domain that shares a residual sequence similarity but is truncated as compared to the plant and fungal hevein domains. Since hitherto no hevein domain was identified in any prokaryote, one can reasonably assume that it was developed by a eukaryote. The present distribution and domain architecture suggests that the hevein domain originated in an early eukaryote (before the separation of the Viridiplantae and fungi/metazoa) and was vertically

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transmitted in plants and fungi where it further evolved by domain duplication and/or fusion to unrelated domains (Fig. 2). Though the domain is certainly not confined to plants and fungi, it seems to be lost from the genomes of most other eukaryotic lineages. Most, if not all, proteins with hevein domains found in plants have no structurally similar (in terms of domain architecture) counterparts in fungi and vice versa, indicating that the hevein family developed independently in plants and fungi. The diversity in domain architecture of the modern plant proteins with hevein domains requires a complex set of evolutionary events. A first question to address is how the hevein domain was passed from an ancestral eukaryote into modern Viridiplantae. Searches in the databases indicated that the hevein domain is present in some green algae (e.g., Chlamydomonas reinhardtii) as part of chimeric proteins with a yet unidentified overall domain architecture (Suzuki et al., 2000). This observation confirms that the hevein domain was transmitted throughout the Viridiplantae lineage but gives little information about the evolution of the hevein family in plants. It should be mentioned here that the retrieval of the hevein domain in the genome of Chlamydomonas and other chlorophyta is hampered by the fact that the corresponding genes often contain introns within the hevein domain. However, it is almost certain that for none of the proteins identified in Embryophyta, a homolog can be found in any other Viridiplantae lineage. Accordingly, it seems likely that in Embryophyta, the hevein family developed after this lineage separated from the other Viridiplantae. Though it is not certain that the ancestral embryophyte lineage acquired a gene with a sole hevein domain from an earlier Viridiplantae lineage, all evidence suggests that such a gene is at the origin of all hevein domains found in modern land plants. Genes/proteins with a single hevein domain are fairly widespread, indeed, in seed plants as well as in S. moellendorYi. Using a single domain gene as a template, plants were able to develop proteins comprising up to seven hevein domains by domain duplication/multiplication events. Since lectins with multi‐hevein domains are taxonomically patchy distributed, domain amplification events took place most probably at multiple independent occasions in diVerent taxa of especially the angiosperms. In the case of the Solanaceae lectins, domain amplification was apparently accompanied by the insertion of SPPP or similar linker sequences. Amplification of these linkers eventually resulted in lectins with relatively long (10 to more than 50 amino acid residues) sequences of (imperfect) SPPP repeats, which in the past were considered ‘‘extensin‐like’’ domains but are neither evolutionarily nor structurally related to the genuine extensins (Van Damme et al., 2004b). This complex evolutionary event apparently occurred in a not so distant past in a subgroup of the Solanaceae family. To develop

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the chimeric proteins, a single domain gene was fused at its C‐terminus to structurally unrelated domains like a chitinase domain, a barwin domain, and a DPBB domain. Since both the class I and class IV chitinases as well as the pathogenesis‐related class 4 proteins (barwins) occur most probably in all terrestrial plants, the underlying gene fusion events must have taken place in a common ancestor of all Embryophyta and the resulting genes were apparently retained in all lineages. The fusion event yielding the hevein‐DPBB chimers must also have taken place before the separation of the liverworts, mosses, and lycophytes. However, in this particular case, the genes were only retained in the primitive land plants and lost from the genome of the angiosperms. 8. Jacalins Molecular cloning of a previously described T‐antigen disaccharide binding lectin from seeds of the jack fruit tree (Artocarpus integrifolia) revealed that this so‐called ‘‘jacalin’’ was unrelated to any other plant lectin and hence represented a novel family of carbohydrate binding proteins. Since nearly identical homologs were found in several other Artocarpus species and in Maclura pomifera but no similar lectins were identified in any other plant, jacalin was considered the prototype of a minor lectin family confined to a small subgroup of the Moraceae family. However, this assumption appeared incorrect after cloning of a mannose‐specific lectin from the rhizomes of the Convolvulaceae species Calystegia sepium (hedge bindweed) revealed that this protein shares over its entire length (150 amino acid residues) a reasonably high sequence identity with jacalin. Moreover, it soon became evident that similar mannose binding lectins and/or corresponding genes occur in many species covering a very wide taxonomic range. Though clearly related to jacalin, the mannose binding homologs diVer in several aspects. First, there is an obvious diVerence in nominal specificity (T‐antigen disaccharide versus mannose). Second, the protomers of the mannose‐specific homologs are intact whereas these of jacalin are cleaved into two dissimilar polypeptides (Van Damme et al., 1998a; Young et al., 1995). Third, jacalin is synthesized on the ER as a preprotein whereas the mannose‐specific homologs are synthesized without signal peptide. Based on these obvious diVerences, the jacalin‐related lectins are now subdivided into ‘‘galactose‐specific’’ and ‘‘mannose‐specific’’ jacalin‐related lectins (GJRL and MJRL, respectively). The subfamily of GJRLs comprises jacalin and its genuine homologs. GJRLs are built up of cleaved protomers, exhibit a clear preference for galactose over mannose, and are located in the vacuolar compartment (Peumans et al., 2000b). Their documented occurrence is confined to a few genera of the family Moraceae (including Artocarpus, Maclura, and Morus). MJRL are a

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far more complex subfamily than the GJRL. The representatives with the most simple structure (e.g., the Calystegia sepium lectin) are built up of uncleaved protomers (equivalent to the jacalin protomer), exhibit an exclusive specificity toward mannose and are located in the cytoplasmic/nuclear compartment of the cell (Peumans et al., 2000b). Besides these single domain proteins, the MJRL subfamily also comprises lectins built up of protomers consisting of multiple (two to six) in tandem arrayed repeats (Fig. 9). Several of these multi‐domain lectins have been purified and characterized in (some) detail. Well‐known examples are the two‐domain Castanea crenata (Nomura et al., 1998) and three‐domain Parkia platycephala (Mann et al., 2001) lectin, and the myrosinase binding proteins from Brassica napus with four to six jacalin domains (Geshi and Brandt, 1998). Genome/transcriptome analyses indicated that single and/or multiple domain MJRLs are very widespread in seed plants (including gymnosperms) and also occur in lycopods, mosses, ferns, and cycads. For historical reasons (linked to the fact that jacalin was the first member of this family to be identified and isolated), the common carbohydrate module of all these lectins is usually referred to as the ‘‘jacalin domain.’’ It should be emphasized however, that jacalin is due to the intradomain proteolytic cleavage not a representative example for the great majority of all currently identified (uncleaved) jacalin domains. Besides in hololectins, the jacalin domain is also found in diVerent types of chimeric proteins. Several Poaceae species (including important crop plants like wheat, rice, and maize) express proteins consisting of an N‐terminal ‘‘dirigent’’ domain fused to a C‐terminal jacalin domain. Depending on the species, this type of chimerolectin was called
‐glucosidase aggregating factor (Z. mays) (Esen and Blanchard, 2000), VER2 protein and Hfr‐1 (T. aestivum) (Williams et al., 2002; Yong et al., 2003), and OsJAC1/ Os12g0247700 (O. sativa) (Jiang et al., 2006, 2007). No homologs could be retrieved outside the Poaceae family. In the genome of A. thaliana, genes encoding putative proteins consisting of a single (At3g16390 and At2g33070) or two (At3g16410) N‐terminal jacalin domain(s) fused to a sequence of five in tandem Kelch motifs were identified. Hitherto, this type of chimerolectins is exclusively documented in A. thaliana. The A. thaliana genome also contains two genes (At3g59590 and At3g59610) encoding a putative F‐box protein comprising an N‐terminal F‐box followed by an FBA (F‐box associated) domain and a C‐terminal jacalin domain. Thus far, no similar F‐box proteins/genes were identified in any other plant. Three additional chimeric proteins/genes with (a) jacalin domain(s) were found in O. sativa: (i) genes encoding putative proteins consisting of an N‐terminal tyrosine protein kinase domain followed by a duplicated (ABA94732) or triplicated jacalin

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Ja

Jacalins

Examples of purified proteins

Domain architectures Ja

GJRL

Ja Ja Jacalin Ja Ja

Ja Ja Ja

Ja Ja

MJRL

Calsepa

Ja Ja

Ja Ja Ja Ja

Ja Ja

Ja Ja Ja Ja

MornigaM

Heltuba

Ja Ja Ja

VacJRL

Ja

S. moellendorffii

Ja Ja Ja

Ja Ja

Ja Ja Ja

Ja Ja

PPA

CRA

Ja Zea mays Ja

K

Ja

Ja

K K

F

K

K

K

A. thaliana

K

K

Ja

A. thaliana

Ja

Ja

O. sativa

Ja

Ja

Ja

LRR

K

Ja

K

A. thaliana

O. sativa

O. sativa

Signal peptide Dirigent domain K

Kelch motif

F

F-box domain

Kinase domain NB-ARC domain LRR

Leucine-rich repeat

Fig. 9. Schematic overview of the identified domain architectures and purified proteins with jacalin domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

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domain (ABA94721 and ABA94728); (ii) a gene encoding a putative LZ‐ NBS‐LRR class RGA protein (consisting of an N‐terminal NB‐ARC domain followed by a leucine‐rich repeat and a C‐terminal jacalin domain) (BAD23344); and (iii) a gene encoding a putative protein comprising a long unknown N‐terminal domain followed by a Peptidase_C48 domain and a C‐terminal jacalin domain. All chimeric proteins discussed in this section are synthesized without a signal peptide and hence presumably reside in the cytoplasmic/nuclear compartment. This also applies to the putative protein kinases with a C‐terminal jacalin domain, which obviously cannot act as receptor kinases for extracellular signal(s). Structural analyses demonstrated that all jacalin domains, irrespective of whether they are cleaved or not, have the same
‐prism fold formed by three Greek key motifs (Van Damme et al., 2007c). In the GJRLs jacalin and the M. pomifera agglutinin, the 20 amino acid residue
‐chain and 133 amino acid residue ‐chain form three bundles of four antiparallel strands of
‐sheet, which are arranged into a
‐prism fold around a pseudo‐symmetry axis. The (single) monosaccharide binding site is formed by three loops located at one extremity of the
‐prism. X‐ray diVraction analyses confirmed that the uncleaved protomers of MJRL (including artocarpin from A. integrifolia (Pratap et al., 2002), MornigaM from Morus nigra (Rabijns et al., 2005), and the lectins from Calystegia sepium (Bourne et al., 2004), Helianthus tuberosus (Bourne et al., 1999), and Musa acuminata (Clavel et al., 2007; Meagher et al., 2005) have the same
‐prism fold as jacalin. However, it appears that the binding site in the GJRLs is widened as compared to that of the MJRL, which might explain the polyspecific character of the former. It is also noteworthy that in the banana lectin, a second carbohydrate binding site was identified that is located in the vicinity of the first site at the same end of the
‐prism. Jacalin and related lectins were for a long time the only documented representatives of a protein family that was believed to be confined to plants. At present plant proteins still represent the majority of all identified proteins with jacalin domain(s) but it becomes increasingly evident that jacalin or jacalin‐like domains also occur in other eukaryotes and even in prokaryotes (Tables I and II). Genes with a jacalin domain were identified in only six bacterial species from diVerent taxonomic groupings. In animals, a single type of protein was found that comprises a domain similar to jacalin. This so‐ called zymogen granule membrane protein 16 is common in vertebrates. Besides in animals, the jacalin domain is also present in some fungal genes/ proteins where it constitutes the C‐terminal domain of large (800 amino acids) chimeric proteins. The ubiquitous occurrence in land plants and fairly widespread distribution in fungi and vertebrates suggests a monophyletic

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origin, which in turn implies that the ancestral eukaryotic domain predates the separation of fungi/metazoa and Viridiplantae (Fig. 2). It is questionable whether the jacalin domain was developed by a prokaryote. The very limited and patchy distribution rather suggests that a few bacterial species/groups acquired the jacalin domain by lateral transfer from an eukaryote. Genome/transcriptome analyses revealed that jacalin‐related lectins and/or corresponding genes proteins are—apart from the liverwort M. polymorpha— present in and expressed by all Embryophyta for which adequate genomic coverage is available. This implies that the jacalin domain was already present in an early Embryophyta lineage. The origin of the ancestral Embryophyta jacalin domain itself is less clear because no similar domain can be retrieved in the genome/transcriptome of any other Viridiplantae lineage and hence there is no direct link to the presumed ancestral eukaryotic jacalin or jacalin‐ like domain. A plausible explanation is that the domain was lost in the Viridiplantae lineages except in the one leading to the Embryophyta. Irrespective of the unsolved origin of the ancestral Embryophyta jacalin domain, the reconstruction of the molecular evolution of the plant jacalin family raises several additional problems. A first question concerns the molecular environment in which this domain was embedded. Taking into account the widespread occurrence in modern seed plants of cytoplasmic/ nuclear single domain MJRL, it seems logical that an ancestral homolog of these genes is at the origin of the jacalin family in plants. However, this assumption is questioned by the overall structure of the jacalin genes from S. moellendorYi. Screening of the genome/transcriptome databases revealed that this lycophyte expresses three distinct types of single domain lectins which are all synthesized with a signal peptide and a short C‐terminal extension. No putative cytoplasmic/nuclear homologs could be retrieved. Interestingly, the molecular environment of the S. moellendorYi jacalin domains resembles that of the homologous domains found in animal proteins. Moreover, the S. moellendorYi and animal genes share the same exon– intron structure whereas the genes from the angiosperms diVer at this point. Therefore, one cannot preclude that the ancestral Embryophyta jacalin gene encoded a vacuolar/secreted protein. If so, this ancestral ‘‘vacuolar’’ jacalin domain must have been converted at a given point in the evolution of land plants into a ‘‘cytoplasmic’’ form through the removal of the signal peptide from the gene. Taking into account that the signal peptide of the S. moellendorYi jacalin is located on the first exon separated by a zero phase intron from the next exon, the presumed conversion can be achieved by a simple exon loss. After a gene encoding a single domain cytoplasmic/nuclear jacalin was developed, it was vertically transmitted during further evolution of the diVerent Embryophyta lineages and used as a building block for both

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domain duplication and domain fusion events. Gene duplication followed by domain amplification explains the origin of the multidomain jacalins. Genome/transcriptome data revealed that multidomain jacalins are widespread among Embryophyta (including P. patens) but also indicated that there exists a great variation in both the number of genes and the number of jacalin domains per individual gene. Three‐domain jacalins seem to be the most common type as they are found in P. patens and in numerous monocots and dicots. Therefore, it is possible that a three‐domain jacalin gene was already present in an early lineage of land plants. However, it cannot be excluded that multiple independent domain amplifications took place in diVerent taxonomic groups. On the contrary, the unique complex family (at least 30) of jacalin genes with two to six domains found in Arabidopsis and Brassica sp. (but in no other plants) strongly argues for an ‘‘excessive’’ gene/domain amplification within (a subgroup of) the family Brassicaceae. Gene duplication accompanied by fusion to unrelated domain(s) is the most likely mechanism for the origin of the chimeric proteins with (a) jacalin domain(s). Though the available sequence data cover only a small part of the Embryophyta, all evidence suggests that most if not all documented chimeric jacalins are confined to a more or less small taxonomic group. For example, some types are exclusively documented in Arabidopsis or rice. In principle, two explanations are possible: (i) the chimeric genes originated in an early embryophyte but were lost in most lineages or (ii) the chimeric genes were developed by independent evolutionary events in diVerent taxonomic groups. Given the patchy distribution of the currently known chimeric jacalins, the latter theory seems more likely. 9. Proteins with legume lectin domains Seed lectins from a handful of legume species like Canavalia ensiformis ( jack bean), Phaseolus vulgaris (common bean), Glycine max (soybean), Arachis hypogaea (peanut), and Pisum sativum (garden pea) made an outermost important contribution to lectinology as a scientific subdiscipline and were one of the first natural groups of plant lectins to be recognized. Since this type of lectin was only found in legumes, they were commonly referred to as ‘‘legume lectins.’’ Unfortunately, this historical term is now misleading for two reasons. First, lectins structurally and evolutionarily closely related to the genuine legume lectins have also been identified in several species of the Lamiaceae family. Second, in the mean time several other unrelated lectins (e.g., type‐2 RIPs, JRLs, and proteins with hevein domains) were isolated from legumes. Though it is for these reasons preferable to speak in terms of proteins with a domain equivalent to the ‘‘classical legume lectins,’’ the term ‘‘legume lectin’’ is so commonly used that it is hard to replace. However, one

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should keep in mind that the term ‘‘legume lectin’’ has to be reserved for those proteins that are structurally and evolutionary related to, for example, ConA and PHA. All members of the legume lectin family consist of protomers of 250 amino acid residues (Fig. 10). However, the molecular structure of the native proteins is highly diverse depending on (i) the number of subunits (one, two, or four), (ii) the absence/presence of a proteolytic cleavage of the protomer into two smaller polypeptides, (iii) the absence/presence of an interchain disulfide bond, and (iv) the absence/presence of N‐glycans (Van Damme et al., 1998a, 2007c). Numerous legume lectins have been characterized in detail at the structural level (Van Damme et al., 1998a). All protomers, irrespective of whether they consist of an intact or cleaved subunit, exhibit the same canonical overall fold and three‐dimensional structure (Loris et al., 1998). The protomers are built up of a flat seven‐stranded
‐sheet (that forms the back face of the protomer) and a curved six‐stranded
‐sheet (that forms the front face). Both
‐sheets are interconnected by turns and loops to form a flattened dome‐shaped

Proteins with legume lectin domains

Examples of purified proteins

Domain architectures Le Le Le

Le

Le

LegL; legumes

Le

Le

Le

Le

Le

Le

LoLI

LegLc; Medicago truncatula ConA

LegLu; A. thaliana

A. thaliana Signal peptide Transmembrane domain Kinase domain

Fig. 10. Schematic overview of the identified domain architectures and purified proteins with legume domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

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‐sandwich structure, which is commonly referred as the jelly‐roll tertiary fold. An additional
‐sheet of five short
‐strands (the so‐called S‐sheet; Banerjee et al., 1996) stabilizes the overall structure by acting as a linker between the two main
‐sheets. The protomers of all legume lectins contain a single carbohydrate binding site formed by four loops protruding at the top of the dome. Legume lectins also harbor at one side of the
‐sandwich a cluster of hydrophobic residues, which delineate a highly conserved so‐called hydrophobic cavity that is believed to accommodate—at least in some lectins—hydrophobic ligands including the plant hormone auxin (Edelman and Wang, 1978). Another noticeable feature is that legume lectins are metalloproteins with two divalent cations incorporated in their structure: (i) a Ca2þ ion linked to an Asp residue of the carbohydrate binding site, and (ii) a Mn2þ ion linked in the vicinity of the carbohydrate binding site. The great majority of all currently known legume lectins have been identified in legume species. Given the long scientific history and large impact in early lectinology, there are virtually no accessible Fabaceae species that have not been checked for the presence of lectins. Accordingly, it is not surprising that several hundreds of true legume lectins and corresponding genes have been isolated and studied in detail, and that they have been the subject of numerous review papers (Etzler, 1985; Sharon and Lis, 1990; Strosberg et al., 1986; Van Damme et al., 1998a, 2007c). Apart from the true legume lectins, genuine homologs are until now documented only in a few Lamiaceae species (including Glechoma hederacea, Salvia sp., Clerodendron sp., Moluccella laevis) (Alperin et al., 1992; Kitagaki et al., 1985; Medeiros et al., 2000; Vega and Perez, 2006; Wang et al., 2003a, b). Genome/transcriptome analyses revealed that hololectins with a legume lectin domain are not confined to Fabaceae and Lamiaceae but are very widespread among land plants (Table I). All sequenced plant genomes including these from P. patens and S. moellendorYi contain at least one gene encoding a protein with a high similarity to the classical legume lectins. However, the deduced sequences of these genes are considerably longer due to a long (60–150 amino acids) extension at the C‐terminal end of the legume lectin domain. Moreover, apart from M. polymorpha virtually all Embryophyta for which adequate genomic coverage is available express one or more of these novel types of legume lectins. To distinguish this apparently ubiquitous type of legume lectin (in land plants) from the classical Fabaceae lectins (LegL), it will further be referred to as ubiquitous legume lectin (LegLu). Since the genome of M. truncatula (and other legumes) also contains LegLu, it is possible to corroborate the relationships between LegLs and LegLus in the very same species. Screening of the M. truncatula genome databases yielded 15 and two genes encoding LegLs and LegLus, respectively.

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Both types of M. truncatula lectins share roughly 25 and 40% sequence identity and similarity, respectively, indicating that they are only distantly related. This low similarity contrasts the relatively high sequence identity/ similarity (>40 and >60%, respectively) between the LegLus from M. truncatula and, for example, V. vinifera. Both the LegLs and LegLus are synthesized with a signal peptide and accordingly enter the secretory pathway. Surprisingly, screening of the M. truncatula genome also yielded a set of three genes encoding a protein that comprises a (slightly truncated) legume lectin domain but lacks a signal peptide, and hence can be considered a cytoplasmic/nuclear homolog of the ‘‘vacuolar’’ LegLs and LegLus. Unfortunately it is still unclear whether these presumed cytoplasmic lectins are expressed because no complementary EST sequences have been deposited. It should also be mentioned that there is no evidence for the occurrence of similar genes/proteins in any other plant species (including legumes like Glycine max and Lotus japonicus). All members of the legume lectin family purified and characterized thus far are hololectins. However, though no chimeric proteins with a functional legume lectin domain have been isolated yet, genome and transcriptome analyses leave no doubt that most if not all land plants possess an extended family of genes encoding (putative) receptor kinases consisting of an N‐terminal (extracellular) domain resembling the legume lectins and a C‐terminal (cytoplasmic) protein kinase domain (Barre et al., 2002; Herve´ et al., 1996). Since many of these genes are expressed, one can reasonably assume that the corresponding receptor kinases fulfill a yet unidentified role in the plant. However, it is still unclear whether the legume lectin domain is involved in the binding of glycans. Besides in land plants, genes with a legume lectin domain are also found in Ostreococcus sp., which belong to the Prasinophyceae, an early diverging class within Viridiplantae. Both O. lucimarinus and O. tauri possess a gene encoding a primary translation product comprising a signal peptide, a legume lectin domain, and a short C‐terminal extension. All evidence suggests that the resulting protein is a homolog of the legume lectins found in land plants. In addition, O. lucimarinus expresses a large (1009 amino acids) protein comprising a signal peptide, a long unidentified N‐terminal domain, and a legume lectin domain located at the C‐terminus. The identification of proteins with a legume lectin domain is important for two reasons. First, it demonstrates that the legume lectin domain also occurs in lower plants. Second, it appears that the hololectins from Ostreococcus sp. share a higher sequence identity with bacterial homologs than with these from Embryophyta (or any other eukaryote). The latter observation sheds a new light on the origin of the plant legume lectin domain because it provides a strong

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argument in favor of a bacterial origin. Until recently, this link was not obvious because the plant legume lectins shared only a very low residual sequence identity with bacterial legume lectin domains (Van Damme et al., 2007c). Besides in plants, proteins with a domain ‘‘equivalent’’ to legume lectins were identified in several other eukaryotic lineages. Examples are the ERGIC‐53 (ER‐golgi‐intermediate compartment 53 kDa protein) and VIP36 (vesicular‐integral membrane protein 36) proteins, which both play an important role in glycoprotein quality control and topogenesis in animals as well as in fungi but are apparently absent from plants. The current taxonomic distribution (Tables I and II) strongly argues for a bacterial origin (Fig. 2) of the legume lectin domain and a vertical transfer into diVerent eukaryotic lineages. Within Viridiplantae, the domain was retained in some but probably not all lineages (no genes are found in the genomes of, e.g., Volvox and Chlamydomonas). It seems that there was a continuous evolution of the hololectins starting from the earliest Viridiplantae lineages. The ubiquitous presence in Embryophyta and the fairly strong conservation indicate that the LegLu type hololectins form a monophyletic group the origin of which predates the separation of the diVerent embryophyta lineages. Both the narrow taxonomic distribution and the low sequence similarity to the LegLus indicate that the LegL‐type hololectin diverged from the main evolutionary line. The underlying evolutionary event apparently involved gene duplication accompanied by a loss of the greater part of the C‐terminal extension. It also seems likely that the duplicated gene acquired a diVerent promoter for an enhanced expression and evolved independently from the coexisting LegLu toward a storage/defense role. Though the LegLs found in Fabaceae and Lamiaceae share a high sequence identity/similarity, it is questionable whether they have a common ancestor. Taking into account that the Fabaceae and Lamiaceae belong to diVerent lineages of the core eudicotyledons (rosids and asterids, respectively), the ancestral LegL must predate the separation of the asterids and rosids. If so it is diYcult to explain why LegLs are absent from all other rosids and asterids. Therefore, it seems more likely that the Fabaceae and Lamiacaeae LegLs evolved independently within each family through a similar mechanism. This hypothesis is supported by the above‐described composition of the M. truncatula legume lectin family, which is indicative for a still ongoing gene amplification and divergence. Because of the very limited amount of information, no conclusions can be drawn with respect to the origin of the chimeric protein found in Ostreococcus sp. In contrast, the wealth of information about receptor kinases with a legume lectin domain (lecRLKs) allows tracing their possible origin. All Embryophyta (including liverworts) contain more or less extended families

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of lecRLKs. Since no homologs are found in any other organism, one can reasonably assume that the lecRLKs have been developed in an early ancestor of the Embryophyta by a fusion of a legume lectin and a protein kinase domain, and a linker comprising a transmembrane sequence. Once the ancestral lecRLK gene was developed, it was vertically transmitted and amplified. 10. LysM domain During the last five years, evidence accumulated for the occurrence of the so‐called Lysin domain (LysM domain) in diVerent types of plant proteins (Fig. 11). The LysM domain was originally identified in a variety of enzymes involved in bacterial cell wall degradation (Joris et al., 1992) and is believed

LysM domain

LysM

Domain architectures LysM

Examples of purified protein

LysM

LysM

LysM

LysM

Pteris ryukyuensis

Selaginella moellendorffii

LysM

LysM

LysM

LysM

LysM

LysM

LysM

LysM

LysM

Nod-factor receptor

LysM

LysM

LysM

LysM receptor-like kinase

CEBiP Oryza sativa

Chlorophyta LysM

LysM

LysM

LysM

LysM

LysM

Volvox carteri Volvox, Chlamydomonas, Ostreococcus, Picea, Vitis, Physcomitrella

Signal peptide

Kinase domain

Transmembrane domain

Glycoside hydrolase 18

Fig. 11. Schematic overview of the identified domain architectures and purified proteins with LysM domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

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to have—at least in bacteria—a general peptidoglycan binding function. In plants, the LysM domain was first identified in LysM domain‐containing receptor‐like kinases (LysM RLKs) from legumes that are involved in the perception of rhizobial signals and were called Nod Factor Receptors (NFR1 and NFR5) because they interact through their LysM domain with Nod factors (Limpens et al., 2003; Madsen et al., 2003; Radutoiu et al., 2003). Later, similar LysM RLK genes were also found in nonleguminous plants. For example, in the genome of A. thaliana five LysM RLK genes were identified (Zhang et al., 2007). Interestingly, some of these genes apparently play a critical role in chitin signaling and fungal resistance in Arabidopsis (Wan et al., 2008b). Besides in LysM RLKs, LysM domains were also found in a chitin elicitor binding protein (CEBiP) isolated from rice cell membranes (Kaku et al., 2006). CEBiP is a 328 amino acid glycoprotein comprising three in tandem arrayed LysM domains and a short C‐terminal membrane spanning (probably GPI‐anchored) sequence. Finally, a very recent paper reports a new type of plant chitinase from the fern Pteris ryukyuensis that comprises two N‐terminal in tandem arrayed LysM domains and a C‐terminal class IIIb chitinase (Glycoside hydrolase family 18) domain (Onaga and Taira, 2008). No similar LysM containing a chitinase domain has been identified and annotated in any other plant species. However, in the transcriptome of S. moellendorYi, three (incomplete) cDNAs could be retrieved that encode a chimeric protein comprising a single N‐terminal LysM domain fused to a C‐terminal class IIIb chitinase domain. All evidence suggests that the LysM domains found in plants interact with either chitin‐oligosaccharides or structural analogs substituted with fatty acids (as in Nod factors). Since both types of glycans are structurally similar to the bacterial cell wall component peptidoglycan, which is the ‘‘normal substrate’’ of bacterial lysins, one can reasonably assume that the bacterial and plant homologs still share a high structural similarity. At present, no plant LysM domain has been studied at the structural level. However, the structure of the LysM domain from some bacterial proteins (e.g., E. coli membrane‐bound lytic murein transglycosylase D) was resolved (Bateman and Bycroft, 2000). The overall fold corresponds to a
‐‐‐
secondary structure in which the two helices pack onto the same side of an anti‐parallel
‐sheet. A shallow groove located on the surface of the protein most probably acts as a binding site. Both the LysM RLKs (including the legume NFRs), the Pteris ryukyuensis and S. moellendorYi chitinases, are typical chimeric proteins. CEBiP contains only a short extra domain but taking into account that the short C‐terminal membrane spanning sequence most probably represents a GPI anchoring domain, CEBiP can also be considered a chimeric protein.

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This implies that all identified plant proteins with LysM domains are chimerolectins. Moreover, all these proteins are synthesized with a signal peptide and presumably follow the secretory pathway to their final destination into the vacuolar/extracellular compartment (chitinase and CEBiP) or the cell membrane (RLKs and NFR). According to a recent review (Buist et al., 2008), over 4000 proteins from both prokaryotes and eukaryotes were found to contain a single or multiple LysM domains. To explain the wide distribution in bacterial species and eukaryotes (Tables I and II) and the apparent absence from Archaea, two hypotheses were put forward. First, LysM was already present in the common ancestor of all living organisms but was purged from the genome of the archaeal lineage. Second, the LysM domain originated after the separation of bacteria, Archaea and eukaryotes, and was subsequently horizontally transmitted from bacteria to eukaryotes or the other way around. Phylogenetic analyses indicated that plant LysM domains (i.e. the domains found in the RLKs/NFRs and CEBiP) are ancient and that the corresponding genes have evolved through local and segmental duplications (Buist et al., 2008). The identification of LysM‐chitinases in ‘‘primitive’’ Embryophyta‐like ferns and lycophytes supports the latter conclusions. To further corroborate the origin of plant LysM domains, a preliminary screening was made for the possible presence in ‘‘lower’’ Viridiplantae. Thereby, it was found that proteins with LysM domains are quite common in green algae. For example, proteins consisting of two consecutive LysM domains occur in C. reinhardtii, Haematococcus pluvialis, O. tauri, O. lucimarinus, Scenedesmus obliquus, and Volvox carteri. Unlike the RLKs/NFRs, CEBiPs, and LysM‐chitinases, these proteins lack a signal peptide and hence are targeted to a diVerent subcellular location. Interestingly, the same type of protein is also expressed by the moss P. patens, the conifer P. sitchensis, and the angiosperm Vitis vinifera. Since perfectly matching sequences are present in the genomes of P. patens and V. vinifera, there is no doubt that at least in these species the proteins are encoded by plant genes (and not by a possible contaminant or endophyte). Evidently, the latter findings put the evolution of plant genes with LysM domains in a new perspective. All evidence suggests that a cytoplasmic/nuclear protein with a double LysM domain or a predecessor with a single LysM domain is at the origin of the modern plant LysM family. The corresponding gene(s) were retained in most but apparently not all Viridiplantae. Insertion of such a gene between a signal peptide and a chitinase domain possibly gave rise to the LysM‐chitinases. Since these chimeric proteins are absent from green algae but are found in lycophytes and ferns, the presumed domain fusion most probably took place after the Embryophyta diverged from the Chlorophyta. Based on the documented

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taxonomic distribution, the LysM‐chitinase gene was retained in a few ‘‘primitive’’ lineages of the Embryophyta but was already lost in the early seed plant lineage. A comparable evolutionary event involving the insertion of a cytoplasmic/nuclear double domain sequence between a signal peptide and a protein kinase domain might have resulted in the formation of an RLK‐type chimeric gene. Since this type of gene is found in all angiosperms and perhaps also in S. moellendorYi, its origin predates the separation of the vascular plant lineages. According to the current taxonomic distribution, the RLK‐type chimeric gene was retained and amplified in all lineages. Within the Fabaceae family, a diversification occurred whereby an RLK gene was converted into a NFR gene that by multiple duplication events eventually gave rise to the NFR families found in modern legumes. 11. Nictaba family (formerly Cucurbitaceae phloem lectins) The isolation, characterization, and cloning of a jasmonate‐inducible lectin from tobacco leaves (called Nicotiana tabacum agglutinin or Nictaba) (Chen et al., 2002) allowed delineating the carbohydrate binding domain present in the previously described so‐called Cucurbitaceae phloem lectins. Phloem exudates from diVerent Cucurbitaceae species contain high levels of a chitin binding lectin (PP2) that upon exposure to the air forms large complexes with the most abundant structural phloem protein (PP1). The formation of these complexes is not mediated by sugar–protein interactions but depends on the formation of intermolecular disulfide bonds. Characterization of the lectins and cloning of the corresponding cDNAs and genes demonstrated that the PP2 subunits (220–230 amino acid residues) are synthesized without a signal peptide on free ribosomes and undergo, apart from a possible modification of the N‐terminal methionine, no posttranslational processing (Bostwick et al., 1992; Wang et al., 1994). The phloem lectins are exclusively produced in the companion cells of the vascular tissue and are translocated into the phloem sap (Golecki et al., 1999). All PP2 proteins comprise a short C‐terminal cysteine‐rich stretch that enables the formation of intermolecular disulfide bonds between the lectins and PP1. Since the size of the PP2 subunits lies in the same range as that of most other plant lectins and, in addition, no information was available, the Cucurbitaceae lectins were considered hololectins. However, when aligned with the Nictaba polypeptide (165 amino acid residues), which possesses full agglutination activity and hence comprises a complete carbohydrate binding domain, it became evident that Cucurbitaceae lectins comprise an extra N‐terminal sequence of 65 residues and a short (five amino acid residues) C‐terminal extension with two Cys residues (Van Damme et al., 2004a,c) (Fig. 12). Accordingly, the term Cucurbitaceae phloem lectins cannot be used to refer to the embedded

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Nic

Nictaba domain

Examples of purified proteins

Domain architectures N. tabacum

Nic

N NsN

Nic

S. tuberosum

NIN

Nic

S. tuberosum

CPL

Nic

Cucurbita sp

Nic

Nictaba

Nic

Nic Cucurbita pepo phloem lectin (PP2)

Nic F

Nic

T

Nic

AIG1

A. thaliana, O. sativa A. thaliana Nic

A. thaliana

N-terminal domains

F

F-box domain

C-terminal domain

T

TIR domain

AIG1

AIG1 domain

Fig. 12. Schematic overview of the identified domain architectures and purified proteins with Nictaba domain. A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

carbohydrate binding domain. Therefore, it is preferable to introduce a novel, unambiguous term. Taking into account that Nictaba is the only genuine hololectin isolated and characterized thus far, the name ‘‘Nictaba domain’’ seems most appropriate. Nowadays, it is clear that the true Cucurbitaceae phloem lectins represent a special subgroup of a very extended family of plant proteins with a Nictaba domain. The members with the most simple structure are the hololectins with a single Nictaba domain. Nictaba itself can be regarded as the prototype of this subfamily. This tobacco lectin is a homodimeric protein of 165 amino acid residue subunits, which are synthesized in the cytoplasm and undergo, apart from acetylation of the N‐terminal methionine, no posttranslational modifications. After synthesis, Nictaba is partly transported into the nucleus

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where it is believed to interact with numerous glycoproteins. Though originally considered a chitin binding lectin, glycan microarray analyses revealed that Nictaba has a higher aYnity for high‐mannose N‐glycans (Lannoo et al., 2006b). Lectins nearly identical to Nictaba are expressed in several other Solanaceae species (e.g., potato and tomato). Besides in Solanaceae, genes encoding Nictaba homologs were identified in Apium graveolens, A. thaliana (Dinant et al., 2003) (At1g33920), Cucumis melo, and C. sativus. Moreover, a preliminary survey of the genome/transcriptome databases indicates that Nictaba homologs are fairly widespread among dicots but are not common if present at all in monocots, gymnosperms, mosses, and lycophytes. Exploration of the databases revealed the occurrence in many plants of proteins in which the Nictaba domain is preceded by an extra N‐terminal sequence. Two types with an N‐terminal extension of 25 (e.g., At4g19850) and 100 (e.g., At4g19840) amino acid residues, respectively, can be distinguished. Though no detailed overview can be given yet of the taxonomical distribution, it seems that Nictaba homologs with a long N‐terminal extension are very widespread among angiosperms. Homologs with a short N‐terminal extension might be less common. A comprehensive analysis of the Arabidopsis genome/transcriptome revealed the occurrence of chimeric proteins in which a C‐terminal Nictaba domain is fused to an N‐terminal F‐box, TIR‐domain, and AIG1‐domain, respectively. Searches for homologs in other species revealed that F‐box‐Nictaba proteins are expressed in all Embryophyta for which an adequate genomic coverage is available. Moreover, in all sequenced Embryophyta genomes, multiple (up to 20) F‐box‐Nictaba genes occur. No homologs of the other two types of chimeric genes/proteins could be identified yet in any other species. The Nictaba domain shares no detectable sequence similarity with any other known protein or domain, and according to the available information is confined most likely to Embryophyta (Tables I and II). Therefore, it is tempting to speculate that this domain was developed by plants. Given the ubiquitous presence in all Embryophyta and apparent absence from other Viridiplantae, the Nictaba domain was developed most probably in an ancestor of all modern Embryophyta lineages (Fig. 2). After the sole domain was acquired, it was passed into the daughter lineages and used as a building block for the development of additional genes with extra sequences or domains. Because of the lack of detailed information about the complement of genes with a Nictaba domain present in the genomes of diVerent species from various taxa, the overall evolution of the Nictaba family is diYcult to reconstruct. However, for some proteins the origin can be traced reasonably well. For example, the origin of the ubiquitous F‐box‐Nictaba chimers must predate the separation of the Embryophyta lineages. Similarly, all evidence

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suggests that the true Cucurbitaceae phloem lectins are derived from a protein consisting of a Nictaba domain preceded by a long N‐terminal extension. The underlying evolutionary event most probably involved a gene duplication followed by the addition of a short cysteine rich C‐terminal peptide. Given the exclusive occurrence in a few genera of the Cucurbitaceae family, the origin of the true Cucurbitaceae phloem lectins is apparently of a very recent date (in evolutionary terms) and can be situated in a lineage of the Cucurbitaceae that gave rise to the genera Cucumis, Cucurbita, and perhaps some other genera. 12. Ricin‐B family The very first lectin that was discovered, namely ricin from the castor bean (Ricinus communis), is still the prototype of the so‐called ricin‐B family, which is one of the most widespread families of carbohydrate binding proteins found in nature. Structural analysis of ricin revealed that the basic carbohydrate unit consists of three tandemly arrayed subdomains of 40 amino acid residues. This carbohydrate binding domain is commonly referred to as the ‘‘ricin‐B domain.’’ For the sake of clarity it should be mentioned here that a clear distinction must be made between the carbohydrate binding domain or ‘‘ricin‐ B domain’’ and the ‘‘ricin B‐chain,’’ which is the lectin subunit of ricin (and comprises two tandem arrayed ricin‐B domains). The most famous plant lectins of the ricin‐B family are the so‐called type 2 RIPs (Stirpe, 2004; Stirpe and Battelli, 2006). Type 2 RIPs are typical chimeric proteins built up of an A‐chain with a polynucleotide:adenosine glycosidase domain (PAG, formerly called RNA N‐glycosidase or RIP domain, A‐chain) and a B‐chain with a duplicated ricin‐B domain (Fig. 13). Both chains are synthesized on a single precursor and remain linked by an interchain disulfide bridge. Type 2 RIPs are bifunctional proteins possessing both carbohydrate binding and enzymatic activity. By virtue of their PAG domain, type 2 RIPs are capable of (catalytically) inactivating ribosomes by removing a highly conserved adenine residue from the sarcin/ ricin loop of the large ribosomal RNA. As a consequence, type 2 RIPs are extremely potent cytotoxins once they enter the cell. It should be emphasized, however, that only some type 2 RIPs like ricin, abrin, and a few others can be considered genuine toxins. Most other type 2 RIPs are only moderately or even weakly toxic (e.g., type 2 RIPs from elderberry fruits) (Barbieri et al., 2004). The dramatic diVerences in toxicity cannot be ascribed to diVerences in the enzymatic activity of the A‐chain but are due to the ability to penetrate the cell, which itself is largely determined by the carbohydrate binding properties of the B‐chain (Peumans et al., 2001; Stirpe, 2004; Van Damme et al., 2001).

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Detailed X‐ray crystallographic studies of ricin revealed that the lectinic B‐chain consists of two tandemly arrayed homologous ricin‐B domains (referred to as domain one and two, respectively) (Robertus and Monzingo, 2004; Rutenber and Robertus, 1991). Both domains comprise three homologous subdomains (,
, ) and an additional subdomain . Each of the two domains is built up of three bundles of four short
‐strands arranged into a
‐trefoil fold around a threefold symmetry axis. Stabilization of the
‐trefoil is ensured by two (intradomain) disulfide bridges. The B‐chain of ricin comprises two distinct carbohydrate binding sites located in subdomains 1 and 2, respectively. Possibly, a third carbohydrate binding site is located in subdomain
(as in some bacterial ricin‐B domains) (Steeves et al., 1999). Most plant lectins with ricin‐B domains are considered galactose or GalNAc specific. Only a few lectins from Sambucus species definitely preferentially interact with sialylated glycans (Shibuya et al., 1987; Van Damme et al., 1998a, b). Hitherto, type 2 RIPs have been found exclusively in flowering plants. Within the angiosperms, they are fairly widespread. Type 2 RIPs were purified and characterized from Ricinus communis (Euphorbiaceae), Viscum sp. (Santalaceae formerly Loranthaceae), Sambucus sp. (Adoxaceae), Abrus sp. (Fabaceae), Ximenia americana (Olacaceae), Adenia sp. (Passifloraceae), Cinnamomum camphora (Lauraceae), Iris sp. (Iridaceae), and Polygonatum sp. (Ruscaceae). In addition, transcriptome data indicate that type 2 RIPs are also expressed in Elaeis guineensis (Arecaceae), Panax ginseng (Araliaceae), Gossypium sp. (Malvaceae), Ipomopsis aggregata (Polemoniaceae), Malus domestica (Rosaceae), Centaurea and Helianthus sp. (Asteraceae), and Sorghum sp. and Zea mays (Poaceae). It should be emphasized, however (i) that type 2 RIPs are not ubiquitous among the angiosperms and (ii) that they have a rather patchy distribution. For example, within the Fabaceae family, type 2 RIPs are documented in only one species. Though the currently documented distribution is certainly still incomplete, the apparent absence from the genomes of, for example, P. trichocarpa, A. thaliana, M. truncatula, V. vinifera, and O. sativa leaves no doubt that type 2 RIP genes have been lost in many species. Since especially the toxic type 2 RIPs received a lot of interest from scientists in diverse disciplines, lots of eVorts were undertaken to resolve the structure of ricin‐B domains. Detailed studies with ricin and a few other type 2 RIPs allowed reconstructing the biosynthesis of these particular chimerolectins. Basically, type 2 RIPs are synthesized on the ER as complex preproproteins comprising an N‐terminal signal peptide followed by a PAG domain, a linker sequence of 10–20 amino acid residues and a C‐terminal domain with two in tandem

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ricin‐B domains (Frigerio et al., 2001; JolliVe et al., 2004, 2006). After co‐ translational removal of the signal peptide, the resulting propeptide undergoes a series of posttranslational modifications including (i) N‐glycosylation, (ii) interchain disulfide bond formation, (iii) proteolytic excision of the linker between the N‐ and C‐terminal domains, as well as the cleavage of a propeptide to yield a mature lectin protomer consisting of an A‐chain disulfide linked to the B‐chain. The eventual structure of the mature type 2 RIPs depends on the degree and type of oligomerization of the protomers. Some type 2 RIPs are monomers (e.g., ricin) whereas others are dimers (e.g., Ricinus communis agglutinin) or tetramers (e.g., Sambucus nigra agglutinin I). The oligomerization is usually achieved by noncovalent interactions but in some type 2 RIPs interprotomer disulfide bonds are formed (e.g., Sambucus nigra agglutinin I) (Van Damme et al., 1996a). Besides this general scheme, two ‘‘aberrant’’ proteolytic modifications are documented in Sambucus sp. that yield a protein with a totally diVerent structure. First, deletion from the precursor of SNA‐V of a fragment starting from the N‐terminus and ending at residue eight of the normal B‐chain results in a (pseudo)hololectin corresponding to a slightly truncated B‐chain (SNA‐II) instead of the ‘‘normal’’ type 2 RIP SNA‐V (Van Damme et al., 1997b). Second, processing or degradation in elderberry fruits of the type 2 RIP SNA‐I yields a small (pseudo)hololectin corresponding to the sole C‐terminal ricin‐B domain of the parent type 2 RIP (Peumans et al., 1998). Apart from type 2 RIPs, plants also express genes encoding hololectins with ricin‐B domains (Fig. 13). At present, there is no evidence that plants express genes encoding proteins consisting of a single ricin‐B domain. A few sequences encoding proteins with a single ricin‐B domain were deposited in transcriptome databases of, for example, T. aestivum and Populus sp. However, it is still uncertain whether these sequences are really of plant origin. Genes encoding hololectins with a duplicated ricin‐B domain were identified in at least three diVerent angiosperms. Cloning and biochemical analysis revealed that several Sambucus lectins are synthesized as preproproteins that undergo a co‐translational removal of the signal peptide, a posttranslational cleavage of an (35 amino acids) N‐terminal propeptide and N‐glycosylation to yield a glycosylated hololectin built up of two ricin‐B domains (Van Damme et al., 1997b). In cucumber (Cucumis sativus), a root‐ specific xylem sap protein (called XSP30) was identified and partly characterized that corresponds to a hololectin with a double ricin‐B domain. This lectin is also synthesized with a signal peptide but there is no evidence for an additional N‐terminal propeptide. The same applies to a set of four putative XSP30 homologs (some of which are expressed) found in the genome of P. trichocarpa.

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PLANT LECTINS

Ricin-B domain

Ri

Domain architectures

Ri

Ri

Ri

PAG

Examples of purified proteins

Cucumis sativus

Ri

Ri

Sambucus sp.

Ri

Ricinus sp.

Ri

Ri

XSP30 from Cucumis sativus

Ri

Ri

SNA-IV from Sambucus nigra

PAG

Ri

Ri

PAG

Ri

Ri

PAG

Ri

Ri

PAG

Ri

Ri

PAG

Ri

Ri

PAG

Ri

Ri

PAG

Ri

Ri

Ricin

RCA

Ri

Ri

PMRIP-4M

SNA-II from Sambucus nigra

Signal peptide PAG Polynucleotide:adenosine glycosidase domain

Fig. 13. Schematic overview of the identified domain architectures and purified proteins with ricin‐B domain(s). A representative plant species is given for each domain architecture. If available, an example of a purified lectin is presented (in a box) for the diVerent domain architectures.

In contrast to type 2 RIPs, which are apparently confined to Angiosperms, hololectins with ricin‐B domains are also expressed in a liverwort (Table I). According to transcriptome data, M. polymorpha expresses a complex set of genes encoding proteins that comprise a double ricin‐B domain. A closer examination of the deduced sequence indicated that these liverwort proteins

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are synthesized without signal peptide. N‐terminal sequencing confirmed that the mature lectin polypeptide lacks only the N‐terminal Met residue, indicating that the M. polymorpha ricin‐B lectins are synthesized on free ribosomes and thus reside most likely in the cytoplasmic/nuclear compartment. The latter conclusion is of paramount importance because it implies that there is a fundamental diVerence between the liverwort and angiosperm members of the ricin‐B family for what concerns their subcellular location. It is worth mentioning in this context that most nonplant eukaryotic ricin‐B lectins (e.g., from fungi and worms) are cytoplasmic/nuclear proteins. The ricin‐B domain is found in numerous prokaryotic and eukaryotic proteins. Most if not all fungi and metazoa, as well as many other eukaryotes possess proteins with one or more ricin‐B domains (Tables I and II). The great majority of all these proteins are chimeric proteins. Some of these chimerolectins are enzymes with a relatively simple overall structure (e.g., bacterial glycosyl hydrolases, animal glycosyl transferases, plant type 2 RIPs) but others have a very complex domain architecture. The taxonomic distribution and overall phylogeny leave no doubt for a prokaryotic origin of the ricin‐B domain (Fig. 2) and subsequent vertical transfer to an ancestral eukaryote. During evolution and divergence of the eukaryotes, the ricin‐B domain was apparently vertically transmitted into many but certainly not all lineages. Because of the high sequence similarity, a straight vertical inheritance of plant and animal/fungal ricin‐B domains from a common ancestor seems obvious. However, it is not clear how plants acquired the ricin‐B domain. A first problem is the fact that no proteins/genes with a ricin‐B domain are found in Viridiplantae other than the Embryophyta (e.g., in Chlorophyta like Volvox and Chlamydomonas). Assuming that the ancestor of all Viridiplantae inherited the ricin‐B domain from an early eukaryote, the domain must have been lost in most lineages. Second, the ricin‐B domain found in the most primitive line of the Viridiplantae (in casu M. polymorpha) is embedded in cytoplasmic/nuclear proteins whereas all other plant proteins with a ricin‐B domain are synthesized with a signal peptide and follow the secretory pathway. Taking into account that most fungal and animal ricin‐B lectins are also synthesized without a signal peptide, the M. polymorpha proteins can be considered the only plant ricin‐B representative with a ‘‘normal’’ subcellular location. As far as can be concluded from the available data, the cytoplasmic/nuclear ricin‐B domain was not retained in any other embryophyte lineage. Third, the ‘‘vacuolar’’ ricin‐B proteins are also confined to the angiosperm lineage of Embryophyta. A survey of the occurrence within this phylum of the most common type of ricin‐B proteins (namely the type 2 RIPs) indicates a patchy distribution over both monocots and dicots.

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Accordingly, it seems likely that type 2 RIP genes originated in a common ancestor of all modern flowering plants. During evolution, the genes were purged from the genomes of most angiosperms. Given the patchy distribution in, for example, the Poaceae family, this process of gene loss occurred in evolutionary recent terms and is possibly still ongoing. In some species, the opposite process of gene amplification took place. For example, the genome of Ricinus communis contains at least eight type 2 RIP genes. The same applies to Sambucus sp. which expresses a very complex set of both genuine and truncated type 2 RIP genes. It appears that in the latter species an evolutionary event took place whereby a typical type 2 RIP was converted in a (holo)lectin gene through the loss of the PAG domain. Most probably a similar event explains the origin of the hololectins found in Cucumis sativus and perhaps also in Populus sp. At present, it is not clear how the angiosperms acquired type 2 RIP genes. Though it seems very likely that these genes result from a fusion of a PAG and ricin‐B domain, the origin of both domains remains enigmatic. It is diYcult to explain, indeed, that both domains are present in angiosperms but in no other Embryophyta. Hereby the remark should be made that the ricin‐B domain found in the M. polymorpha proteins is more closely related to bacterial proteins than to the B chain of type 2 RIPs, which renders an evolutionary link improbable. In summary, the molecular evolution of the ricin‐B family in plants is far from understood. There is no evidence for a straight classical vertical inheritance of the ricin‐B domain(s) from an ancient eukaryotic ancestor into the diVerent Viridiplantae lineages. Though purely speculative, other mechanisms based on lateral gene transfer should be considered.

13. Others Apart from one exception all currently known/characterized plant lectins for which a minimum of sequence information is available can be classified into one of the 12 families described above. A lectin was isolated and cloned from Dioscorea batatas that is apparently unrelated to all other plant lectins but shares 45% sequence identity with carbonic anhydrase from A. thaliana (Gaidamashvili et al., 2004). Hitherto, no other carbonic anhydrase homologs with carbohydrate binding activity have been identified. Because of the lack of structural data, it is still unclear whether the carbohydrate binding domain of the D. batatas lectin comprises the entire carbonic anhydrase fold or just a subdomain.

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IV. SUGAR BINDING ACTIVITY AND SPECIFICITY OF PLANT LECTINS The discovery in 1952 that the agglutination activity of lectins relies on a binding to specific sugars (Watkins and Morgan, 1952) provided for the first time a biochemical explanation for an until then elusive biological activity. Two decades later, structural analyses of ConA allowed pinpointing the interaction with carbohydrates at a well defined ‘‘sugar binding site’’ (Edelman et al., 1972; Hardman and Ainsworth, 1972). At present, the three‐dimensional structure of most but not all known carbohydrate binding domains has been resolved (Table III). Each domain has its own characteristic overall fold with one or more sugar binding site(s) located at specific positions (Loris et al., 1998; Raval et al., 2004; Robertus and Monzingo, 2004; Sinha et al., 2007; Wright and Hester, 1996). Virtually all identified sugar binding sites consist of a shallow depression exposed at the surface of

TABLE III Overview of Known Three‐Dimensional Structures of Plant Lectins Number of three‐ dimensional structuresa Lectin domain Amaranthin Amaranthus Cyanovirin Ceratopteris GNA domain Allium, Galanthus, Gastrodia, Hippeastrum, Narcissus, Scilla Hevein domain Amaranthus, Hevea, Phytolacca, Triticum, Urtica Jacalin domain Artocarpus, Calystegia, Helianthus, Maclura, Morus, Musa, Parkia Legume domain Bowringia, Canavalia, Cratylia, Dioclea, Dolichos, Erythrina, Glycine, GriVonia, Lathyrus, Lens, Maackia, Pisum, Phaseolus, Psophocarpus, Pterocarpus, Robinia, Ulex, Vicia Ricin‐B domain Abrus, Ricinus, Sambucus, Trichosanthes, Viscum a

Numbers taken from: http://www.cermav.cnrs.fr/lectines/

Free

Complexed

Total

1

1

2

1

0

1

5

9

14

13

14

27

8

24

32

68

138

206

11

13

24

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the protein. Since for most lectin families, three‐dimensional structures of the lectin in complex with a mono‐ or small oligosaccharide have been resolved, the structure–function relationships—in terms of specific recognition of simple sugars—are reasonably well understood. Generally speaking the recognition/binding of simple sugars primarily depends on the interaction of a few amino acid residues located in the lectin carbohydrate binding site with some hydroxyls of the sugars. The specific interaction between the lectin and the carbohydrate involves the formation of a network of hydrogen bonds and is often reinforced by a hydrophobic stacking of the pyranose ring of the sugar to the aromatic ring of Tyr or Phe residues located in the close vicinity of the carbohydrate binding site.

A. PLANT LECTINS POSSESS EXTENDED BINDING SITES THAT PREFERENTIALLY INTERACT WITH COMPLEX GLYCANS

Many plant lectins definitely interact with (a) specific monosaccharide(s). However, the aYnity for monosaccharides is usually very low as compared to that for more complex glycans. The reason for this diVerence in aYnity resides in the atomic structure of the binding sites. Structural analyses revealed that the so‐called monosaccharide binding site accommodates a single well‐defined sugar unit of a bulky N‐glycan chain. Additional amino acid residues located in the vicinity of the primary binding site interact with other sugar units so that a more extended carbohydrate binding site is created. Water‐mediated H‐bonds often participate in the binding of these complex glycan chains. By increasing the number of H‐bonds that anchor the glycan chain to the carbohydrate binding site, the aYnity for complex N‐glycans (in the 10
6–10
8 M range) is usually three to four orders of magnitude higher than that for simple sugars (in the 10
3–10
4 M range). Extended binding sites are found in all plant lectin families for which detailed structural data are available supporting the general idea that complex glycans rather than mono‐ or disaccharides are the natural targets of most plant lectins (Van Damme et al., 1998a, 2007c). It should be mentioned, however, that there are a few documented examples of plant lectins that interact exclusively with a monosaccharide. For example, the lectins from Lamiaceae species (e.g., Salvia sp., Mollucella laevis, Clerodendron sp., and Glechoma hederacea) bind with a high aYnity (in the M range) to GalNAc and GalNAc O‐linked to serine residues (Tn‐antigen) in a protein backbone. Interestingly, the aYnity of these lectins is strongly (up to 104‐fold) enhanced by the presence of multiple clustered Tn‐epitopes, which in a sense mimic complex carbohydrates (Singh et al., 2006).

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1. The specificity of most plant lectins is directed against N‐glycans Numerous plant lectins have been studied in great detail to unravel their sugar binding specificity. For a long time, specificity studies were exclusively based on an indirect approach involving assays whereby the ‘‘competitivity’’ of sugars/glycans was compared in inhibition of agglutination, precipitation, or binding reactions. Though these methods were certainly useful, the need for relatively large quantities of both lectins and glycans often limited the resolving power (especially in terms of the range of glycans that could be screened). As a result, the specificity of most lectins was usually discussed in terms of ‘‘accessible’’ glycans rather than ‘‘potential natural targets.’’ This eventually led to the correct but biased general idea that plant lectins preferentially interact with mono‐ or oligosaccharides moieties of complex O‐ and N‐linked glycans. B. HIGH PERFORMING ANALYSIS TECHNIQUES URGE TO REVISE THE OLD PARADIGMS

During the last decade, several high performance methods were developed that are based on direct measurements of protein–carbohydrate interactions and oVered the additional advantages of automatization and scaling down of assay volumes. Surface plasmon resonance (Linman et al., 2008; Murthy et al., 2008), frontal aYnity chromatography (Hirabayashi, 2004, 2008; Tateno et al., 2007), and glycan microarrays (Blixt et al., 2004) allowed a novel approach to specificity studies. Especially the introduction of glycan microarrays (Van Damme et al., 2009b) revolutionized the analysis of the carbohydrate binding specificity of lectins in general and plant lectins in particular. Glycan microarrays carrying a large number of both natural and synthetic glycan structures allow a very fast and comprehensive screening of the binding properties with a minimal amount of fluorescently labeled lectin (Blixt et al., 2004; Paulson et al., 2006). In addition, by varying the lectin concentration, fairly detailed information can be obtained for what concerns the relative aYnity of the lectin for diVerent glycans. The current glycan microarrays (v3.1) available from the Consortium for Functional Glycomics contain a set of 377 glycan targets, which is considered representative for nonreducing termini of N‐ and O‐glycans in glycoproteins as well as glycolipids. A complete list of the glycans in all versions of the glycan microarray of the CFG is available on the website: (http://www. functionalglycomics.org/static/consortium/resources/resourcecoreh8.shtml). Over the last five years, numerous lectins from diVerent sources have been analyzed by the Consortium using the glycan microarray technology. The results obtained with plant lectins confirmed a preferential binding to

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oligosaccharides and glycans rather than to monosaccharides. More importantly, the glycan microarray analyses allowed—by virtue of unprecedented broad range of test glycans—refining the specificity of many plant lectins. An important conclusion to draw from the data generated by the glycan microarray analyses is that the specificity of most plant lectin domains is directed against N‐glycans in general and high mannose‐type N‐glycans in particular. N‐glycan binding is documented, indeed, for members of all plant lectin families except these with an A. bisporus agglutinin, amaranthin, and LysM domain. Depending on the carbohydrate binding domain, the recognition of the N‐glycans requires a well‐defined set of monosaccharides on the outer branches, the core trimannose, or the core pentasaccharide (Man3GlcNAc2). To illustrate these novel insights generated by the high performance analysis techniques, a few specific cases are briefly discussed here. A first example is the snowdrop lectin. On the basis of hapten inhibition, assays of the agglutination activity GNA was originally described as a lectin with an exclusive specificity toward mannose (Van Damme et al., 1987). More detailed studies using hapten inhibition of the precipitation of glycoproteins allowed refining the specificity and indicated that GNA requires the equatorial hydroxyl groups at the C‐3 and C‐4 positions and an axial group at the C‐2 position of the D‐pyranose ring. Furthermore, the lectin requires a nonreducing terminal D‐mannose residue for the interaction with oligosaccharides, and oligosaccharides with terminal Man(‐1–3)Man units showed the highest inhibitory potency among the manno‐oligosaccharides tested (Shibuya et al., 1988). Glycan microarray analysis confirmed these results but also demonstrated that GNA exhibits a very high aYnity for high mannose (Man9, Man8, Man7, Man6, and Man5) glycans. Since in the latter assays both Man3‐chitobiose and chitotriose were not bound, it could be concluded that the oligomannosyl part of the N‐glycans is required for the high reactivity of the N‐glycans. A second example are the lectins with hevein domains. Numerous lectins of the hevein family (e.g., WGA, UDA, potato, and tomato lectins) have been reported to specifically bind to GlcNAc oligomers (Peumans et al., 1984; Shibuya et al., 1986). In the case of UDA, detailed studies of its carbohydrate binding properties were performed using quantitative precipitation assays and hapten inhibition. It was shown that the carbohydrate binding site of UDA is complementary to an N,N0 ,N00 ‐triacetylchitotriose unit. Analysis of the carbohydrate binding properties of UDA on the glycan microarray confirmed the strong binding to the (GlcNAc)3‐oligomer, but also revealed that the lectin has a higher aYnity for high mannose N‐glycans (e.g., Man9, Man8, Man6, and Man5) and that this high‐aYnity interaction requires the core pentasaccharide (Man3GlcNAc2). Similar results were

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obtained for the tomato lectin using a lectin blot analysis (Oguri, 2005). It should be emphasized, however, that this obvious preference for N‐glycans over chito‐oligosaccharides cannot be extrapolated to all hevein domains. For instance, WGA and DSL (Datura stramonium agglutinin) are more reactive toward glycans containing terminal GlcNAc or substituted GlcNAc. The tobacco agglutinin is another example of a lectin that was originally considered a chitin binding protein (Chen et al., 2002) but upon glycan microarray analysis turned out to exhibit a clear preference for high mannose and complex N‐glycans over GlcNAc oligomers. In this case also, the high aYnity for the N‐glycans requires the presence of the core Man3GlcNAc2 pentasaccharide (Lannoo et al., 2006b). Finally, a recent reinvestigation by the glycan microarray technique demonstrated that the Euonymus europaeus agglutinin, which was described as a blood group B substance‐specific lectin, definitely also interacts with high mannose N‐glycans (Fouquaert et al., 2008). 1. The issue of multi‐specificity Early specificity studies already indicated that several lectins interact with a very broad range of oligosaccharides or glycans. Moreover, some extreme cases were reported where plant lectins apparently interact with structurally unrelated glycans. Because of the limitations inherent to methods based on competitive binding assays, it was often diYcult to explain the apparent polyspecific character in terms of either extended binding sites or multiple independently acting binding sites. Fortunately, the introduction of frontal aYnity electrophoresis and glycan microarrays allowed a renewed approach to this problem by a more detailed and extended dissection of the specificity. Two case studies illustrate the progress that could be made in this area. First, a reinvestigation of the specificity of a set of plant lectins with a duplicated GNA domain revealed that most of these lectins exhibit a clear dual specificity directed against high mannose and complex N‐glycans, respectively, and that this dual specificity relies on the simultaneous presence of at least two diVerent independently acting binding sites (Van Damme et al., 2007b). Second, as already mentioned above, glycan microarray analysis revealed that the ‘‘canonical’’ blood group B saccharide‐specific lectin from Euonymus europaeus (Teneberg et al., 2003; Yamamoto and Sakai, 1981) reacts not exclusively with the blood group B substance and related oligosaccharides but also binds with a fairly high aYnity to high mannose N‐glycans (Fouquaert et al., 2008). A careful examination of the glycan microarray data indicates that even lectins with a presumed ‘‘simple’’ specificity (in terms of recognition of a single type of glycan) might exhibit a more complex carbohydrate binding

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pattern. For example, the specificity of GNA is most probably the sum of the specificities of the three individual binding sites. The same applies most likely to all lectins that comprise multiple (unidentical) carbohydrate binding domains and/or domains with multiple binding sites. 2. The issue of carbohydrate binding defective lectin homologs Several cases have been reported of plant proteins that are clearly evolutionarily related to genuine lectins but exhibit no agglutinating or sugar binding activity. Classical examples are the sweet protein curculin (GNA family) (Barre et al., 1997), arcelins (Legume lectin family) (Mirkov et al., 1994), and Sambucus nigra lectin‐related protein (SNLRP, Ricin‐B family) (Van Damme et al., 1997a). The latter protein is devoid of agglutinating activity and fails to bind to any of the commonly used immobilized sugars (e.g., galactose, N‐acetylgalactosamine, N‐acetylneuraminic acid) or glycoproteins (e.g., fetuin, asialofetuin, mucin, asialomucin, ovomucoid, and thyroglobulin). Molecular modeling and docking studies using the B‐chain of ricin as a model indicated that the replacement of some critical amino acids in the potential binding sites of SNLRP is responsible for the apparent loss of carbohydrate binding activity. However, when analyzed by glycan microarrays, SNLRP strongly reacted with GlcNAc‐oligomers (pentamer, hexamer, trimer) as well as with numerous glycan structures containing GlcNAc residues substituted with galactose residues or sialic acid. Accordingly, SNLRP can no longer be considered a carbohydrate binding deficient RIP. Two important lessons should be taken from this particular example. First, it is precocious to consider a protein carbohydrate binding deficient on the basis of assays with a limited set of sugars/glycans. Second, one should be extremely careful in interpreting predictions made by molecular modeling and docking experiments.

V. BIOSYNTHESIS, TOPOGENESIS, AND SUBCELLULAR LOCATION OF PLANT LECTINS A. AN OLD PARADIGM: PLANT LECTINS FOLLOW THE SECRETORY PATHWAY

Until the mid 1990s it was more or less generally accepted that the great majority of all plant lectins are synthesized on the ER and follow the secretory pathway to their final destination into the vacuolar compartment (including specialized organelles derived thereof like storage protein vacuoles) or more rarely the extracellular space. This paradigm was based

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on a quite impressive amount of data from numerous studies of the biosynthesis, topogenesis, and subcellular location of various lectins combined with cDNA cloning of the corresponding genes. Initially there were no good reasons to contest this paradigm because virtually all lectins known at the time followed the secretory pathway, indeed. Classical examples that contributed to our general understanding of the biosynthesis, posttranslational modifications, and topogenesis are a number of legume lectins (PHA, pea lectin, ConA), jacalin, the snowdrop lectin, ricin, and some elderberry type 2 RIPs, and diVerent types of lectins with hevein domains (including WGA, hevein, UDA, and class I and class IV chitinases). Since the results of these studies have been described in detail in a previous review paper, they are not repeated here (except for a brief summary included in the sections describing the diVerent carbohydrate binding domains/lectin families) (Van Damme et al., 1998a, 2007c).

B. EXPERIMENTAL EVIDENCE FOR THE OCCURRENCE OF NONSECRETORY PLANT LECTINS

Molecular cloning of genes encoding the phloem lectins from Curbita sp. provided the first evidence for the synthesis of a plant lectin on free ribosomes in the cytoplasm (Bostwick et al., 1992; Wang et al., 1994). However, this particular aspect received little attention. The concept of ‘‘cytoplasmic’’ plant lectins was eventually introduced after a homolog of jacalin was identified in Calystegia sepium that was synthesized without a signal peptide and—as was demonstrated by immunocytochemical localization—resides in the cytoplasm and nucleus of the rhizome cells (Peumans et al., 2000b). Apart from demonstrating that some plant lectins are located in the cytoplasm and the nucleus, the identification of the Calystegia sepium agglutinin (Calsepa) as a mannose binding homolog of the galactose binding vacuolar jacalin provided for the first time unambiguous evidence for the occurrence of both cytoplasmic and vacuolar homologs within a single plant lectin family. In the mean time, several other cytoplasmic homologs of Calsepa have been isolated and characterized. For example, rice seedlings accumulate a mannose‐specific jacalin‐related lectin upon treatment with jasmonate or NaCl (Zhang et al., 2000). Expression of a fusion construct in which the rice lectin sequence was fused to EGFP revealed fluorescence in the nucleus and the cytoplasm of the tobacco cells whereas the vacuole was completely devoid of signal (Van Damme et al., 2009a). Since these findings confirm the predicted cytoplasmic/nuclear location of the rice lectin, one can reasonably assume the same applies to all other homologs.

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A few years after the discovery of the cytoplasmic jacalin homologs, a new family of cytoplasmic plant lectins was identified. Chen et al. (2002) reported the characterization and molecular cloning of a jasmonic acid methyl ester‐ inducible lectin (called Nictaba) in the leaves of Nicotiana tabacum (var. Samsun NN). Immunocytochemical localization studies revealed that the lectin is located in the nucleus and the cytoplasm of leaf cells, but not in vascular tissues (Chen et al., 2002). This nucleocytoplasmic location of Nictaba was confirmed using confocal microscopy of tobacco BY‐2 cells expressing a Nictaba sequence fused to EGFP (Lannoo et al., 2006b). Since the Nictaba sequence comprises a putative nuclear localization signal (NLS), sequence experiments were designed to check the possible role of this NLS. Mutation of the NLS (102KKKK105) into (102KTAK105) completely abolished the transport of EGFP‐labeled Nictaba into the nucleus of the BY‐2 cells indicating that the NLS is required and is suYcient for transport of the lectin from the cytoplasm into the nucleus. Screening of transcriptome databases revealed that some plants (T. aestivum, Z. mays, and M. truncatula) express proteins that due to the absence of a signal peptide and a C‐terminal propeptide can be considered cytoplasmic homologs of the vacuolar GNA‐related lectins. Transient expression of such a putative cytoplasmic GNA homolog from maize (tagged with EGFP) confirmed that the lectin resides in the nucleus and cytoplasm of BY‐2 cells. The same applies to the cytoplasmic GNA homologs found in fishes and fungi (Fouquaert et al., 2006; Van Damme et al., 2009a). Molecular cloning of the Euonymus europaeus agglutinin, which was recently identified as the prototype of a novel plant lectin family, revealed that this protein is also synthesized without a signal peptide. Preliminary localization experiments of an EUL‐EGFP fusion protein confirmed the predicted nucleocytoplasmic localization (Van Damme et al., 2009a). Biochemical and molecular analyses demonstrated that the ABA‐homologs expressed in the liverwort M. polymorpha are, like their fungal counterparts, synthesized on free ribosomes. Confocal microscopy confirmed that an EGFP‐tagged M. polymorpha lectin is targeted into the nucleus and cytoplasm of tobacco BY‐2 cells. Finally, a comparison of the amino acid sequence of mature amaranthin and the primary translation product encoded by the corresponding cDNA/ gene indicated that this lectin is also synthesized in the cytoplasm. To check the presumed cytoplasmic location, an amaranthin‐like sequence from Prunus persica fused to EGFP was transiently expressed by BY‐2 cells and analyzed by confocal microscopy. Fluorescence was predominantly associated with the nucleus and to a lesser extent with the cytoplasm (Van Damme et al., 2009a).

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Summarizing, one can conclude that there is ample evidence for the occurrence of cytoplasmic/nuclear lectins in plants. Moreover, cytoplasmic lectins are not confined to a single family but are already documented in six diVerent families. It is also noteworthy that most of the recently identified novel plant lectin families are typical cytoplasmic/nuclear proteins. C. GENOME/TRANSCRIPTOME DATA INDICATE THAT THE MAJORITY OF ALL PLANT LECTINS IS SYNTHESIZED ON FREE RIBOSOMES AND RESIDES IN THE NUCLEOCYTOPLASMIC COMPARTMENT

Though the majority of all purified and characterized plant lectins represent vacuolar forms, this does not imply that cytoplasmic/nuclear forms are less common. On the contrary, the opposite is more likely. The present ratio of vacuolar forms over cytoplasmic/nuclear forms is strongly biased by the fact that until a decade ago virtually all plant lectin research was focused on a few families with almost exclusively vacuolar forms (e.g., legume lectins, hevein family, ricin‐B family, GNA family). Once more attention was given to ‘‘novel’’ lectin families, a rapidly increasing number of cytoplasmic/nuclear lectins from diVerent lectin families were isolated and characterized. The total number of purified cytoplasmic/nuclear lectins is still relatively low. However, all evidence suggests that the rare examples documented thus far are just members of large families of cytoplasmic/nuclear plant lectins. To assess this assumption, a comprehensive screening was made of the publicly accessible genome and transcriptome databases. Though still incomplete, this screening clearly indicates that the majority of all lectin genes encode proteins that are synthesized in the cytoplasm and reside in the cytoplasmic/ nuclear compartment. Moreover, it appears that cytoplasmic/nuclear forms occur in all families except the (large) hevein family and the small cyanovirin and class V chitinase families. Another important conclusion to be drawn is that in most—though not all—cases the present vacuolar forms evolved from an original cytoplasmic form (e.g., vacuolar GNAs, jacalin) (Van Damme et al., 2004a). These findings demonstrate that cytoplasmic/nuclear lectins are the rule rather than the exception. Evidently, this final conclusion has important consequences for what concerns the physiological role of plant lectins.

VI. EXPRESSION OF LECTINS IN PLANTS A. GENERAL ASPECTS

For practical reasons research was for a long time concentrated on those plant lectins that are (very) abundant in seeds or vegetative storage tissues like bark, bulbs, and rhizomes (Van Damme et al., 1998a). Since about

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15 years, there is a tendency to pay more attention to plant lectins that are weakly expressed in nonstorage tissues (like leaves, roots, flowers). This change in interest eventually resulted in the discovery of several novel lectins that are not constitutively expressed but are induced by a specific treatment with a biotic/abiotic stress factor. Along with this evolution in the choice of model systems, the general ideas about the temporal and spatial regulation of the expression of plant lectins (dramatically) changed. Basically, plant lectins can be subdivided in two major groups for what concerns their expression and induction pattern, namely a group of developmentally regulated highly expressed lectins and a group of weakly expressed inducible lectins. For the sake of simplicity, these two groups will further be referred to as (i) highly expressed ‘‘classical’’ lectins and (ii) inducible low expressed ‘‘novel’’ lectins, respectively. Both the subdivision and terminology are rather arbitrary. Moreover, it should be emphasized that this arbitrary subdivision applies to most but certainly not all lectins.

B. HIGHLY EXPRESSED ‘‘CLASSICAL’’ LECTINS

The term ‘‘highly expressed’’ refers to the fact that these lectins represent an important part of the total protein content of the tissues in which they occur. Since these abundant lectins were the preferred (and almost sole) research subjects in the early days of lectinology, the term ‘‘classical’’ is added just to refer to this fact. Examples of highly expressed ‘‘classical’’ lectins are the abundant storage protein‐like lectins found in seeds and vegetative storage tissues. Since the plant physiological aspects of these lectins have been reviewed, this topic is not further discussed here (Van Damme et al., 1998a). The basic principle is that plants accumulate at a given developmental stage large amounts of lectins in seeds or vegetative storage organs. These lectins represent a harmless store of nitrogen that can readily be degraded in situ to provide the seedlings or newly formed shoots with amino acids for a rapid growth and development. Storage protein‐like lectins are particularly common in the legume lectin and GNA family. In addition, they are found in the jacalin (e.g., jacalin itself), ricin‐B (e.g., type 2 RIP in the bark of Sambucus sp.), hevein (e.g., Datura stramonium seed lectin), and amaranthin (amaranthin from A. caudatus seeds) families. The great majority of all highly expressed ‘‘classical’’ lectins are vacuolar forms. However, there are also a few documented examples of cytoplasmic forms (e.g., amaranthin and the mannose‐specific jacalin‐related lectin from Morus nigra bark).

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The term ‘‘inducible low expressed novel lectins’’ refers to three distinct aspects that distinguish this group of lectins from the above‐described highly expressed ‘‘classical’’ lectins. First, the lectins are not constitutively expressed but only upon induction; second, even after induction the eventual expression level remains low, and third, the identification of these lectins is of a recent date. Evidence accumulated during the last decade that some plants synthesize lectin(s) in response to stress situations like drought, high salt concentration, wounding, or treatment with some plant hormones. In general, the expression level of the induced lectins remains low. Most strikingly, all inducible lectins identified thus far reside in the nucleus and the cytoplasm of the cells. On the basis of these observations, the concept was developed that lectin‐ mediated protein–carbohydrate interactions in the cytoplasm and the nucleus play an important or possibly even crucial role in the stress physiology of the plant cell (Van Damme et al., 2004a, b). Since this group of lectins was not included in previous review papers, a comprehensive overview is given below. The very first inducible lectin to be purified and characterized was a mannose‐specific jacalin‐related lectin (called Oryza sativa agglutinin or Orysata) from NaCl‐treated rice seedlings (Hirano et al., 2000; Zhang et al., 2000). Sequence analysis revealed that Orysata corresponded to a previously described salt‐inducible protein (as SalT) (Claes et al., 1990). Orysata cannot be detected in untreated plants but is rapidly expressed in roots and sheaths after exposure of whole plants to salt or drought stress, or jasmonic acid and abscisic acid treatment (Claes et al., 1990; De Souza Filho et al., 2003; Moons et al., 1997). Interestingly, the lectin is also expressed in excised leaves after infection with an incompatible Magnaporthe grisea race (Kim et al., 2003; Qin et al., 2003) as well as during senescence (Lee et al., 2001). An in silico search for proteins and genes comprising domain(s) equivalent to Orysata revealed that these sequences are widespread. In the mean time, jasmonate‐inducible homologs of Orysata have been identified not only in several other Gramineae species (Van Damme et al., 2004d) but also in Helianthus tuberosus (Nakagawa et al., 2000) and Ipomoea batatas (Imanishi et al., 1997). Recently, a mannose‐binding jacalin‐related lectin was purified from barley coleoptiles (Grunwald et al., 2007). Horcolin (Hordeum vulgare coleoptile lectin) shares high sequence homology to the highly light‐inducible barley protein HL#2 (Po¨tter et al., 1996). Horcolin mRNA is upregulated under salt‐stress conditions. It appears that barley

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plants possess two closely related lectins, which are regulated and expressed independently of each other. Grunwald et al. (2007) suggested that horcolin may play a role in the perception or transduction of biotic/abiotic stress‐ induced signals or in the handling of altered environmental conditions. Within the Gramineae family, several inducible chimeric stress/defense‐ related proteins containing a C‐terminal jacalin domain fused to an N‐terminal ‘‘dirigent’’ domain were identified. One of these proteins (called VER2) is specifically expressed during vernalization (Yagi et al., 2002; Yong et al., 2003). The VER2 proteins in rice and in wheat were shown to be jasmonate inducible. Expression of wheat VER2 was also induced by gibberellins (Yong et al., 2003). Expression and characterization of the recombinant rice protein has proven that VER2 is a genuine lectin which can interact with mannose (Jiang et al., 2006). Since overexpression of the gene in rice suppresses coleoptile and stem elongation, it was suggested that this lectin plays an important role in rice growth and development (Jiang et al., 2007). A diVerent homolog of VER2 is induced in wheat plants in response to the initiation of first‐instar larval feeding of specific biotypes of Hessian fly (Mayetiola destructor). The corresponding gene is referred to as Hessian fly responsive gene 1 (Hfr‐1) (Williams et al., 2002). Another homolog, which is high expressed during systemic acquired resistance, was previously described as the ‘‘wheat chemically induced gene 1’’ or WCI‐1 (Go¨rlach et al., 1996). A second type of inducible lectin was identified in leaves of tobacco plants exposed to jasmonate methyl ester. This tobacco lectin (called Nictaba) cannot be detected in untreated plants but accumulates at fairly high concentrations in leaves and in no other tissues. Nictaba is synthesized in the cytoplasm and partly translocated into the nucleus. Apart from jasmonates no other inducing agents/factors except insect (caterpillar) feeding could be identified (Lannoo et al., 2007). Homologs of Nictaba are expressed in several other Solanaceae species but it is not clear whether they are also jasmonate inducible. Curiously, homologs of the Nictaba gene(s) are apparently absent from the genome of several Nicotiana species (Lannoo et al., 2006a). A third type of inducible lectin was already described in 1995 as an abscisic acid (ABA) and salt responsive protein but was only recently recognized as a lectin after molecular cloning of the Euonymus europaeus agglutinin (EEA) revealed that this lectin and the salt‐inducible rice protein share a common domain (Fouquaert et al., 2008). EEA shares a marked sequence identity/ similarity with the so‐called family of rice OSR40 proteins which were found to be induced in roots of a salt‐tolerant rice variety by abscisic acid (ABA) and salt stress (Moons et al., 1995, 1997). Moons et al. (1997) suggested a role

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for OSR40c1 in the adaptation of the roots to a hyperosmotic environment and involvement in water loss prevention and cell wall rigidity. The data deposited in the Rice Expression Database confirm that all members of the OSR40 protein family are strongly induced by salt stress and treatment with the phytohormone ABA. Homologs of the rice OSR40 protein family are expressed most probably in all Embryophyta. This obvious ubiquitous occurrence suggests that the EUL domain plays a universal role in stress‐related physiological processes. Evidence for stress‐related induction was also obtained from studies with maize and banana. Riccardi et al. (2004) have reported increased levels of OSR40 proteins when maize plants are subjected to water deficit. In addition, it was shown that the quantitative variations of OSR40 proteins in maize leaves are linked to diVerences in ABA accumulation. However, the proteins of the OSR40 family are also found in relatively large quantities in unstressed maize coleoptiles. Recently, two isoforms of the OSR40 family were also identified in banana (Musa spp.) (Samyn et al., 2007). In shoot meristem cultures of banana, OSR40 was slightly upregulated after high sucrose stress. In a dehydration‐tolerant banana variety, the OSR40 proteins represented a very abundant protein under high sucrose concentrations. In contrast, the dehydration‐tolerant variety contained much lower levels of OSR40. Therefore, a role in dehydration tolerance was also put forward for OSR40 proteins in banana (Carpentier et al., 2007). In silico expression analysis (Fouquaert, 2008) revealed that the EUL‐ homolog from Arabidopsis (At2g39050) is developmentally regulated with a high expression in senescent leaves and in flowers. Microarray expression analyses of leaf mesophyll cells and guard cells (Leonhardt et al., 2004) revealed that At2g39050 is weakly expressed in the mesophyll cells of 5‐week‐old leaves, but is specifically expressed in the guard cells that form the stomata. Moreover, when the leaves were floated on a 100‐M ABA solution, an extremely high expression of At2g39050 was observed in these guard cells. Analyses of the expression of the EUL‐protein in Arabidopsis upon diVerent environmental stresses revealed that expression of this gene is strongly upregulated in the shoots by treatment with 300 mM mannitol (mimicking osmotic stress) and 150 mM NaCl (salt stress). These results leave no doubt that in Arabidopsis plants, the EUL domain is associated with stress adaptation. The strong induction in the guard cells of stomata after exposure of leaves to ABA suggests that the EUL homolog from Arabidopsis plays a role in the regulation of the closure of the stomata to protect the plant against adverse environmental conditions like drought stress and salt stress.

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VII. WHY DO PLANTS EXPRESS LECTINS? A. CLASSICAL CONCEPTS: MANY HYPOTHESES BUT POOR EXPERIMENTAL EVIDENCE

Since the early discovery of lectins, scientists tried to answer the question why (some) plants accumulate blood clumping proteins. Because of the obvious toxicity of the first lectins isolated (e.g., ricin, abrin, Robinia lectin), it was believed that plant lectins are toxins. Though there is no doubt that some lectins are potent toxins, indeed, toxicity is the exception rather than the rule. Only a few type 2 RIPs (e.g., ricin, abrin, Adenia toxins) can be considered lethal poisons and in this particular case the toxicity requires the presence of an enzymatic domain (for reviews see Barbieri et al., 2004; Stirpe and Battelli, 2006). There are also a few poisonous lectins that owe their toxicity to their carbohydrate binding domain. A classical example is the lectin from beans (Phaseolus vulgaris, PHA) that was incriminated of several documented cases of fatal food poisoning. Experiments with animals revealed that the strong toxicity of PHA is not based on a direct cytotoxic activity but follows a degeneration of intestinal epithelium through disruption of the microvilli, alteration of the cytoskeleton, and a change in the activity of brush border enzymes (Kik et al., 1989). Many other lectins (diVerent legume lectins, jacalin, and WGA) provoke similar but far less dramatic eVects as PHA (Jordinson et al., 1999; Nakata and Kimura, 1985; Pusztai et al., 1993) and therefore are now considered antinutrients rather than toxins. For those lectins that exhibit no toxic eVects, alternative functions had to be searched. Many hypotheses have been put forward but in general very little experimental support could be obtained (for reviews see Van Damme et al., 1998a). A notorious example is the presumed role of legume lectins in the establishment of the symbiosis with nitrogen fixing bacteria of the genus Rhizobium (reviewed by Hirsch, 1999). Despite a tremendous amount of eVorts, including the use of transgenic plants, no unambiguous evidence could be presented that legume lectins play an essential role in this process. On the contrary, the discovery in legumes of LysM domain‐containing receptor‐like kinases (LysM RLKs) that bind with a high specificity and aYnity Nod factors, proved a far more logic and convincing mechanism. Yet, the symbiosis hypothesis still persists. One of the major diYculties in the search for the physiological role of plant lectins is the fact that most of them are present in fairly high to extremely high concentrations and accordingly can hardly be involved in any process based on the recognition of specific signal molecules. For example, how can one explain that some legumes accumulate up to 10% lectin (on a total

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protein basis) in their seeds just to attract a few bacteria to the root hairs of the seedlings? Another problem concerns the subcellular location of these lectins. Taking into account that they are almost invariably translocated into the vacuolar compartment, it is diYcult to envisage how they can be involved in essential cellular processes that are merely confined to the cytoplasmic/ nuclear compartment. Finally, the striking diVerences in the ‘‘overall biology’’ of the classical lectins combined with their patchy taxonomic distribution exclude any general role. Therefore, the idea was gradually developed that these lectins are accessory proteins used for diVerent purposes unrelated to vital processes. Eventually the concept was proposed that most classical lectins can be considered storage/defense‐related proteins (Peumans and Van Damme, 1995; Van Damme et al., 1998a). 1. Most classical lectins can be considered storage/defense‐related proteins Abundant or readily detectable lectins are fairly widespread in seeds and vegetative tissues. In general, these lectins represent 0.1–10% of the total protein. In view of the high lectin concentrations, it is hard to believe that these lectins will interact with specific receptors in the plant. In addition, analyses of the biosynthesis/topogenesis as well as spatio‐temporal expression patterns also provided indirect indications against a specific endogenous role. As already mentioned above, the majority of lectins in seeds and storage tissues is synthesized through the secretory pathway, resulting in a localization of the mature proteins in vacuoles or related organelles, or in the extracellular compartment. In addition, most of these plant lectins are synthesized as inactive precursors that become activated only after sequestration in the specialized organelles, and therefore are inactive at the time they pass through, for example, the ER compartment. Furthermore, most of these plant lectins, irrespective of the family to which they belong, bind exclusively to complex animal N‐ and O‐glycans, and therefore are said to have a clear preference for foreign over plant‐specific glycans (Peumans et al., 2000a). The lack of complementarity between the sugar binding specificity of most plant lectins and the structure of the carbohydrates present on conspecific glycoconjugates is a major argument against a specific endogenous role. Taking into account that many of these abundant lectins are developmentally regulated in a similar way as storage proteins, the concept was developed that these plant lectins are not involved in specific recognition phenomena in the plant cell itself but are a special class of aspecific constitutively expressed defense proteins that help the plant to cope with attacks by predators, such as phytophagous invertebrates and/or herbivorous animals. It is suggested that these lectins combine a defense‐related role with a function as a storage protein and whenever appropriate can be recruited by the plant for defense

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purposes (Peumans and Van Damme, 1995). Evidently, the concept of lectins as defense‐ and/or storage‐related proteins is only applicable for plant lectins that are expressed in high concentration and have a carbohydrate binding specificity that would allow them to recognize and bind, for example, glycoproteins exposed along the intestinal tract of predators. The interaction of plant lectins with several carbohydrate structures in, for example, fungi and insects, and the detrimental eVect thereof on the predator was shown in several studies involving, for example, application of the lectin to the diet or growth medium of the predator and/or studies on growth and development of predators on lectin expressing plants. For instance, the lectins with hevein domains, WGA and UDA, aVect the growth and development of fungi (Broekaert et al., 1989; Van Parijs et al., 1991). It should be mentioned, however, that this antifungal activity of lectins is relatively weak when compared to other antifungal plant proteins (e.g., defensins, thionins). In contrast to fungi and bacteria, many plant lectins are moderately to highly toxic for insects and higher animals (Murdock and Shade, 2002; Pusztai and Bardocz, 1996; Pusztai et al., 1990; Van Damme, 2008). Furthermore, several research groups have provided evidence that there exists a link between toxicity of particular lectins and their capability to recognize and bind animal or insect glycans. B. NOVEL CONCEPTS

Two novel developments urged to update the above‐described concepts regarding the physiological role of plant lectins. First, it becomes increasingly evident that plants accumulate lectins in response to specific external factors. Second, it now appears that cytoplasmic/nuclear lectins are the rule rather than the exception, and that accordingly most plant lectins are at least in principle capable of interacting with specific glycoconjugates in the ‘‘vital’’ compartments of the plant cell. 1. Jasmonate‐ and stress‐induced lectins As already discussed above, the identification of three diVerent types of jasmonate‐ and/or stress‐inducible lectins with an experimentally confirmed cytoplasmic/nuclear location put the search for the physiological role of plant lectins in a totally diVerent perspective because it implied that lectins might well be involved in specific cellular processes within the plant cell. If so, one can reasonably expect that these lectins interact with glycoproteins or other glycoconjugates present in either the cytoplasm or the nucleus, or both. Specificity studies revealed that at least the tobacco lectin has the right specificity to interact with endogenous glycoproteins and more precisely

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with N‐glycosylated glycoproteins carrying high mannose‐type N‐glycans. Conclusive evidence for both in vitro and in situ interactions between Nictaba and nuclear/cytoplasmic tobacco proteins was obtained using Far Western blots. These blots clearly demonstrated that Nictaba reacts with many proteins present in a crude extract from purified nuclei (Lannoo et al., 2006b). PNGase treatment of the proteins almost completely abolished the interaction with Nictaba, suggesting that Nictaba reacts with N‐glycans. In addition, the interaction of the lectin with these nuclear proteins was inhibited by GlcNAc oligomers. Therefore, one can reasonably assume that Nictaba interacts through its carbohydrate binding activity with endogenous glycoprotein receptors in the cytoplasmic/nuclear compartment. This taken together with the nucleocytoplasmic location and the induction by jasmonate strongly argues for a specific role of Nictaba in jasmonate‐inducible or jasmonate‐dependent physiological processes (Chen et al., 2002; Van Damme et al., 2004a). Hitherto, no similar results were reported for any other inducible cytoplasmic/nuclear plant lectin but it seems likely that the conclusions drawn from the studies with Nictaba also apply to at least some other lectins. 2. Insect‐induced lectins Besides jasmonate and stress, insects can also act as an inducing factor for the expression of lectins in plants. In the case of the tobacco lectin, insect herbivory by larvae of the Egyptian cotton leaf worms (Spodoptera littoralis) can partly replace the eVect of jasmonate, most probably by triggering the synthesis of endogenous jasmonates in the leaves (Lannoo et al., 2007). Interestingly, wounding did not induce lectin expression, presumably because the rise in jasmonate levels in the leaf due to diVerent types of wounding is only transient. Previously, Zhu‐Salzman et al. (1998) reported that the expression of the GlcNAc binding GriVonia simplicifolia lectin II (GS‐II) in leaves is systemically (but not locally at the site of treatment) upregulated after wounding as well as by jasmonic acid treatment, whereas insect attack did only upregulate GS‐II expression in systemic leaves but not in local leaves. It should be mentioned that in contrast to the tobacco lectin, which can only be detected after induction, the GriVonia lectin is constitutively expressed at low but detectable levels in leaf tissue. More extensive research was done on wheat plants infested by Hessian fly (Mayetiola destructor Say). Wheat plants respond to an infestation with first‐ instar Hessian fly larvae by the expression of several unique genes including at least three diVerent lectin genes: (i) Hfr‐1 comprising an N‐terminal disease response or dirigent domain and a C‐terminal jacalin domain (Subramanyam et al., 2006, 2008; Williams et al., 2002), (ii) Hfr‐2 consisting of a duplicated

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N‐terminal amaranthin domain fused to a C‐terminal domain similar to channel‐forming lytic toxins (PuthoV et al., 2005) and (iii) Hfr‐3, a homolog of the WGA (Giovanini et al., 2007). Though not all induced genes have been identified yet, the current state of the art clearly argues for the involvement of a set of lectins in the resistance of wheat to Hessian fly. At present, it is not entirely clear how the induced lectins contribute to the plant’s resistance. Some evidence points toward a direct toxic eVect on the insects but other mechanisms cannot be excluded. C. DEVELOPING RESEARCH AREAS

The impact of the recently obtained novel insights in the taxonomic distribution, subcellular location, molecular evolution, expression, and carbohydrate binding specificity on the unraveling and fine tuning of the physiological role of plant lectins is for the time being still limited. However, it can be expected that pioneering work will soon lead to breakthroughs in our understanding of what lectins can do in plants. To illustrate the legitimate expectations, three topics are discussed here in some detail. 1. Involvement of plant lectins in protein–carbohydrate interactions within the nucleocytoplasmic compartment As discussed above in detail, plant lectins are primarily cytoplasmic/nuclear proteins and hence are located in those cell compartments where the great majority of all essential cellular processes take place. In principle, the cytoplasmic/nuclear lectins can interact with any glycoconjugate carrying exposed complementary carbohydrate(s) present in the same compartments, and by doing so aVect processes in which these glycoconjugates are involved. Unfortunately, at present little information is available about the presence of endogenous carbohydrate ligands and glycosylated receptors for the nucleocytoplasmic lectins. Therefore, this issue can be discussed only in general terms. a. Glycosylation in nucleus and cytoplasm. Both cytoplasmic and nuclear proteins from plants and animals are known to undergo a highly dynamic posttranslational modification involving the addition/removal of O‐GlcNAc at serine and threonine residues (Hart et al., 2007). O‐GlcNAc modification is essential for normal functioning and survival in mammalian cells, which illustrates the importance of this simple modification in basic cellular processes. Evidence has been presented that this glycosylation of nucleocytoplasmic proteins modulates signaling but also influences protein expression, degradation, and traYcking in animal cells. In addition, recent experiments

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have indicated that O‐GlcNAc serves as a nutrient and stress sensor (Hart et al., 2007; Zachara and Hart, 2006). In contrast to animal cells, virtually no data were reported on O‐GlcNAc modification in plant cells. However, all evidence suggests that also plant proteins are modified by O‐GlcNAc. b. N‐glycosylation. N‐glycosylation is a very common modification of countless proteins in eukaryotic cells. The process of the attachment of the N‐glycans and subsequent modifications in the ER is for a great deal conserved in all eukaryotes. However, further modifications of the N‐glycans downstream the ER are quite diVerent in, for example, plants, animals, and yeast. N‐glycosylation starts in the ER by the transfer of a precursor oligosaccharide to the Asn residue of an N‐glycosylation consensus sequence (Asn‐X‐Thr/Ser) and is further modified by the coordinated action of several glycosyltransferases and glycosidases. Proper folding and targeting of the newly synthesized glycoproteins is ensured by an eYcient quality control system within the ER. This system involves two lectins, called calnexin and calreticulin, that interact with monoglucosylated oligosaccharide chains present on newly formed N‐linked glycoproteins and play an important role in folding of glycoproteins and targeting of misfolded proteins for degradation (Spiro, 2004). Interplay between the transmembrane region of calnexin and ER degradation‐enhancing ‐mannosidase‐like protein (EDEM), a mannose‐ specific lectin, is involved in selection of misfolded glycoproteins for dislocation into the cytosol where they can be degraded by proteasomes. Another group of L‐type lectins (ERGIC‐53, VIP36, VIPL) is involved in transport of folded glycoproteins to the Golgi (Banerjee et al., 2007). It was recently shown that these lectins have diVerent intracellular distributions and dynamics in the ER‐Golgi compartment as well as diVerent sugar binding properties (Kamiya et al., 2008). Glycoproteins emerging from the ER are transported to the Golgi complex where they are further modified by the removal and attachment of a variety of diVerent monosaccharides. Proteins carrying N‐glycans that leave the Golgi complex are generally destined for secretion and reside in the extracellular compartment. However, during the past few years, evidence accumulated that proteins carrying N‐glycans are also present in the cytosol and nucleus of animal cells (Belanger et al., 2005; Kane et al., 2002; Wozniak et al., 1989) and preliminary data indicate that the same applies to plant cells. Besides glycoproteins, the cytoplasm also contains free oligosaccharides that are generated in the ER from lipid‐linked oligosaccharides as well as from misfolded proteins (Chantret and Moore, 2008; Spiro, 2004) and via the cytoplasm eventually end up in the lysosomes where they are degraded.

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In mammalian cells, the transport of these free oligosaccharides from the ER into the cytosol is an eYcient process but clearance from the cytosol to the lysosomes is much less eYcient, resulting in an accumulation in the low micromolar concentrations of the free oligosaccharides in the cytoplasm. Free N‐glycans are also found in diVerent plant cells (Priem et al., 1993). In Ginkgo biloba, two diVerent types of free N‐glycans were identified: (i) a high mannose type structure with one GlcNAc residue at the reducing end (Man9‐5GlcNAc1) and (ii) a complex type structure with an N‐acetylchitobiose unit at the reducing end (Man3Xyl1Fuc1GlcNAc2). Both types basically diVer from the free glycans in animal cells (Kimura and Matsuo, 2000). Free N‐glycans are ubiquitous in developing and growing plant cells at micromolar concentrations (Maeda and Kimura, 2006). They are found primarily in the cytosol fraction (Kimura et al., 2002). At present, the physiological relevance of these free oligosaccharides in the cytosol is still unclear. It was suggested that they are involved in signaling (Moore, 1999; Priem et al., 1993), protein assembly, and oligomerization (Kimura et al., 2002). In summary, all evidences strongly indicate that the cytoplasmic/nuclear compartment of the plant cell contains both free and protein‐linked glycans. Since most of these glycans are high aYnity ligands for the few nucleocytoplasmic lectins that hitherto were studied in detail for what concerns their specificity, one can reasonably assume that the coexisting nucleocytoplasmic lectins and glycans can at least in principle interact with each other. As already mentioned above, experiments with the jasmonate‐induced tobacco leaf lectin confirmed an interaction with nuclear glycoproteins (Lannoo et al., 2006b). Since it is well established now that some nucleocytoplasmic lectins are only expressed in response to well‐defined biotic and abiotic stimuli, the hypothesis was put forward that these plant lectins act as regulators of intracellular signaling through binding to N‐glycosylated receptors in the nucleus or the cytoplasm. At present, this hypothesis is supported by only a few documented cases. However, transcriptome data indicate that most nucleocytoplasmic lectins are upregulated by stress factors. Moreover, the fact that several of the inducible nucleocytoplasmic lectins are apparently ubiquitous in Embryophyta or seed plants also argues for a general role in the stress physiology of plants. 2. F‐box proteins with a lectin domain: Involvement in glycoprotein degradation Lectins do not only function in proper folding of glycoproteins (see the examples of calreticulin and calnexin above), recent evidence also suggests that they are also involved in glycoprotein degradation (Anelli and Sitia, 2008; Wang and Ng, 2008). Protein quality control occurs primarily in the ER and is based on N‐linked glycosylation. The attachment of N‐glycans to

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nascent proteins in the ER lumen facilitates proper protein folding, yet the retention of high mannose N‐glycans on misfolded or unassembled glycoproteins serves as a marker for ubiquitin‐related degradation (Helenius and Aebi, 2004; Meusser et al., 2005). This ER‐associated degradation (ERAD) pathway involves a retrograde transfer of the glycoproteins from the ER into the cytosol for degradation by the 26S proteasome system. Recognition of target proteins and subsequent ubiquitylation occurs through the SCF complex, a multiprotein complex named after three of its constituents (S‐phase kinase‐associated protein 1 or Skp1, Cullin‐1/CDC53 and an F‐box protein) which is located in the nucleus and cytoplasm and binds target proteins and ubiquitylates them prior to recognition by the nucleocytoplasmic 26S proteasome (Smalle and Vierstra, 2004). The F‐box protein determines the substrate specificity of the SCF complex since it directs target proteins to the SCF complex. F‐box proteins exhibit a typical bipartite structure (Kipreos and Pagano, 2000): the N‐terminal conserved F‐box domain of 40 amino acid residues is required for direct interaction with the SCF complex (Bai et al., 1996) whereas the C‐terminal domain is used to recruit the target protein into the SCF complex. DiVerent substrate‐specific motifs involved in protein–protein interaction have been identified in the C‐terminal domains of many F‐box proteins. Examples are Trp‐Asp (WD) 40 domains, leucine‐rich repeats (LRRs), Kelch domains, Armadillo repeats, tetratricopeptide repeats, zinc fingers, and proline‐rich domains (Petroski and Deshaies, 2005). Recently, experiments with animal cells revealed that a small family of cytosolic mammalian F‐box proteins with a C‐terminal sugar binding domain (SBD) targets mis‐ or unfolded glycoproteins for degradation (Yoshida et al., 2002, 2003). Biochemical and structural analyses demonstrated that two of these sugar binding F‐box proteins (or Fbs proteins), FBXO2 and FBXO6 (also called Fbs1 and Fbs2, respectively), preferentially bind to free N‐glycans with a high‐mannose oligosaccharide (Man3–9) attached to a core chitobiose (GlcNAc2) and to high‐mannose N‐linked glycoproteins (Glenn et al., 2008). Fbs1 and Fbs2 do not bind to Man3, Man4, or Man5 structures lacking GlcNAc2, demonstrating the importance of the chitobiose core. In general, the core GlcNAc2 of N‐glycans present on correctly folded (native) proteins is not accessible for lectins. However, in unfolded (denatured) N‐linked glycoproteins, the chitobiose core of the glycan is more accessible for the SBD of Fbs1 and Fbs2. Yoshida et al. confirmed that SCF complexes with either Fbs1 and Fbs2 recognize denatured ERAD substrates in the cytosol and act in glycoprotein degradation/homeostasis (Yoshida, 2007; Yoshida et al., 2005). In plants, no homologs of the Fbs proteins have been described yet, although 700 (putative) F‐box proteins have been identified in the

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annotated genomes of Arabidopsis thaliana and rice (Gagne et al., 2002; Jain et al., 2007; Kuroda et al., 2002; Lechner et al., 2006). However, plants express a whole battery of F‐box proteins with a C‐terminal Nictaba domain that can be considered functional analogs of the mammalian Fbs proteins. Genome/transcriptome data leave no doubt that F‐box‐Nictaba proteins are ubiquitous in land plants and are evolutionarily fairly well conserved (Lannoo et al., 2008). No F‐box‐Nictaba proteins were isolated and characterized yet. However, the high sequence similarity between the Nictaba domain of the F‐box proteins and the corresponding domain of Nictaba itself and the Cucurbitaceae phloem lectins strongly indicates that it has a comparable carbohydrate binding activity and specificity directed against the core pentasaccharide of N‐glycans. Therefore, it seems justified to consider at least some of the plant F‐box proteins as functional analogs of the mammalian Fbs1 and Fbs2 proteins. 3. Receptor kinases with a lectin domain: Involvement in cell signaling A particular class of chimeric lectins that deserve special attention are the receptor‐like kinases (RLKs) with a carbohydrate binding domain (or at least a domain related to one of the documented plant carbohydrate binding domains). RLKs are one of the largest gene families in the Arabidopsis genome with more than 600 members representing nearly 2.5% of Arabidopsis protein coding genes (Shiu and Bleecker, 2001, 2003). Hitherto, RLKs have been found exclusively in plants (Krupa et al., 2006). They consist of an N‐terminal extracellular ligand binding domain, a hydrophobic single transmembrane‐spanning domain, and a C‐terminal intracellular catalytic kinase domain (Walker, 1994). As such, their structure resembles that of the animal receptor tyrosine kinases and the receptor serine/threonine‐specific kinases of the TGF
R (transforming growth factor
receptor) family (Cock et al., 2002; Shiu and Bleecker, 2001). However, most plant kinase domains do not phosphorylate tyrosine residues but are serine/threonine specific (Nishiguchi et al., 2002) or act on histidine residues (Hwang et al., 2002). The extracellular domains of the plant receptor kinases can be highly diverse which enables the RLK to selectively respond to diverse extracellular signals. One third of the RLKs found in Arabidopsis contain leucine‐rich repeat sequences, implicated in protein–protein interactions, in their extracellular ligand binding domain (Shiu and Bleecker, 2001, 2003). Little is known about their functional role but it has been suggested that plant RLKs may have a role in a diverse range of signaling processes such as self‐incompatibility reactions in Brassica species via S receptor kinase signaling (Takasaki et al., 2000), brassinosteroid signaling via the BRI1 kinase (Belkhadir and Chory, 2006), meristem development controlled by CLAVATA1 (DeYoung and Clark,

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2001), leaf development controlled by Crinkly4 (GiVord et al., 2005) and bacterial/disease resistance mediated by Xa21 in rice (Song et al., 1995), FLS2 in Arabidopsis (Chinchilla et al., 2006), and Eix2 in tomato (Mariano et al., 2004). Disruption in the plant RLK activities leads to abnormal plant development. Until now, only limited information is available about the ligands that are recognized by plant RLKs (see Cock et al., 2002 for an overview). Herve´ and coworkers were the first to identify putative extracellular lectin‐ like domains in plant receptor kinases (Herve´ et al., 1996, 1999). Meanwhile, it has become clear that lectin‐like RLKs (lecRK) are a large superfamily of plasma membrane‐located RLKs exclusively expressed in plants containing diVerent types of extracellular lectin domains such as legume lectin‐like domains, LysM domains, and GNA domains, and conserved intracellular (functional) serine/threonine kinase domains (Fig. 14) (Barre et al., 2002; Krupa et al., 2006). The first Arabidopsis lecRLK genes identified, Ath.lecRK‐a1‐4, were reported by Herve´ et al. (1996, 1999). Meanwhile, 42 lecRKs and nine related soluble legume lectin sequences could be identified in Arabidopsis (Barre et al., 2002). The Ath.lecRKs consist of an N‐terminal signal peptide sequence, an extracellular legume lectin‐like domain which is heavily N‐glycosylated, a short single‐pass transmembrane helical domain, a juxtamembrane domain, and an intracellular serine/threonine kinase domain, capable (in most cases) to autophosphorylate a Ser residue. It is unlikely that most of these Ath. lecRKs can bind monosaccharides because some of the crucial amino acid residues in the binding site are substituted. However, the hydrophobic cavity seems to be conserved in the lectin domains of the Ath.lecRKs and is assumed to bind complex glycans and/or hydrophobic ligands such as auxins and/or cytokinins. Andre´ et al. (2005) demonstrated the lectin domain of PnLPK, a poplar lecRK highly homologous to the Ath.lecRKs interacts with carbohydrates such as ‐L‐rhamnose, (neo)glycoproteins, and asialofetuin. Gouget et al. (2006) reported that the lectin domain of an Ath.lecRK (At5g60300) was devoid of any sugar binding activity but instead could mediate plasma membrane–cell wall adhesions through protein–protein interactions based on RGD (Arg‐Gly‐Asp) sequence recognition. Several papers report the involvement of lecRKs in developmental processes. For example, Riou et al. (2002) observed that the expression of Ath. lecRK‐a1 is induced in natural senescence of cotyledons and leaves. SGC lecRK, another Arabidopsis lecRK, was shown to be involved in pollen development, probably by oligoglucan and oligogalacturonic acid signaling (Wan et al., 2008a). An RLK from Gossypium is believed to be involved in fiber development in cotton bolls (Zuo et al., 2004).

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A Transmembrane lecRLKs A. thaliana, P. nigra, M. truncatula, O. sativa

Le

LysM

LysM

A. thaliana, L. japonicus, O. sativa

LysM

A. thaliana, L. japonicus, O. sativa

GNA B

Soluble cytoplasmic/nuclear lectin-kinases

Am

Am

Aquilegia formosa x Aguilegia pubescens

Ja

Ja

O. sativa

Ja

Ja

Ja

O. sativa

Legume lectin domain

Transmembrane domain

LysM

LysM domain

S-locus domain

GNA

GNA domain

PAN/APPLE-like domain

Am

Amaranthin domain

Kinase domain

Ja

Jacalin domain

Le

Fig. 14. Schematic representation of the overall structure of protein kinases with a lectin domain. (A) Membrane‐integrated lectin receptor‐like kinases and (B) soluble cytoplasmic/nuclear kinases with one or more lectin domains.

LecRKs might also be involved in wound signaling and defense‐related pathways. Riou et al. (2002) reported an induced expression of Ath.lecRK‐a1 in leaves after mechanical wounding and application of oligogalacturonic acids, which suggested that Ath.lecRK‐a1 participates in the jasmonate‐ independent wound‐inducible pathway. Ath.lecRK2 might be involved in salt stress and ethylene signaling (He et al., 2004). The poplar lecRK PnLPK gene is upregulated by wounding but is, unlike Ath.lecRK‐a1, predominantly expressed in roots rather than in other organs (Nishiguchi et al., 2002). Two Medicago lecRKs genes with a root‐specific expression were highly induced

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upon nitrogen starvation and repressed after rhizobial inoculation or addition of lipochitooligosaccharide Nod factors. These MtlecRKs were shown to interact through the hydrophobic cavity present in their legume lectin‐like domain with a Nod factor secreted by the rhizobium Sinorhizobium meliloti and could increase nodulation on Medicago roots, implying a role for MtlecRKs in legume–rhizobia symbiosis (Navarro‐Gochicoa et al., 2003). Also the Lotus japonicus RLKs, having extracellular LysM lectin domains, were shown to be involved in Nod factor recognition from a bacterial microsymbiont (Madsen et al., 2003; Radutoiu et al., 2003). Tobacco BY‐2 cells express legume lectin‐like RLKs that are responsive toward bacterial and yeast elicitors (Sasabe et al., 2007). Lastly, LecRKs might also be involved in fungal recognition events, as is the case for a plasma membrane‐located B‐lectin domain containing lecRK from rice (Chen et al., 2006). As summarized above, the lectin(‐like) domains of lecRKs might recognize extracellular signals by diVerent mechanisms and transmit these to the intracellular plant compartment where the kinase domain can activate by protein phosphorylation a multitude of signaling pathways involved in plant development and defense.

VIII. THE PLANT LECTINOME The information generated by transcriptome and genome analyses gives a fairly comprehensive and accurate overview of the carbohydrate binding domains and annex protein families found in plants. Evidently, the overall picture is still incomplete because less than a dozen plant genomes are (nearly) completed and transcriptome data are also available for only a limited number of species. Moreover, apart from a few exceptions, the transcriptome data cover only part of the genome and are often biased by the nature of the libraries. However, two important conclusions can already be drawn. First, plants possess a very complex set of genes encoding a multitude of proteins with one or more carbohydrate binding domains. Second, there are apparently huge diVerences between the ‘‘set of lectin genes’’ present in the genomes of diVerent plant species. These diVerences concern both the type of lectins and the number of genes. At present, there is no simple explanation for the observed diVerences. In principle, one can reasonably assume that ubiquitous lectins (like class I chitinases, F‐box proteins with a Nictaba domain, proteins with an EUL domain, receptor kinases with a GNA, or legume lectin domain) fulfill a universal role in all Embryophyta whereas those with a (very) narrow taxonomic distribution (e.g., jacalin, most legume lectins) are accessory proteins with a well‐defined

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but unspecific function (e.g., as defense or storage protein). However, the situation is less clear for lectins that are present in most but not all species or taxa. A possible explanation might be that some carbohydrate binding domains are components of a redundant system and that accordingly one or more lectin families can be lost in a lineage without loss of an essential function. To address this issue, one needs a detailed overview of the full complement of lectin genes (which can be defined as ‘‘the lectinome’’) present in the genomes of diVerent species spanning a broad taxonomic range. Once a minimal number of plant lectinomes have been corroborated, comparative analyses will allow tracing how basic genome evolution processes (like genome duplication, gene duplication, gene elimination, transposon insertion, and gene evolution) steered the evolution of the plant lectinome (Gaut and Ross‐Ibarra, 2008; Leitch and Leitch, 2008; Tang et al., 2008). Comprehensive plant lectinome analyses will also be extremely useful to compare the arsenal of carbohydrate binding proteins found in plants and other organisms. There are undoubtedly fundamental diVerences between the lectinomes of plants, fungi, and animals. Moreover, it seems that plants possess diVerent unique lectins that are involved most probably in cellular processes that are confined to Embryophyta. Accordingly there are fundamental diVerences between the ‘‘glycobiology’’ of plant, fungal, and animal cells.

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Clock Control Over Plant Gene Expression

ANTOINE BAUDRY AND STEVE KAY

Section of Cell and Developmental Biology, Division of Biological Sciences, University of California, San Diego, La Jolla, California 92093

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Biological Pathways Under the Influence of the Clock in Plants . . . . . . . . . . . A. Leaf Movements ................................................................ B. Ion Fluxes and Stomatal Aperture .......................................... C. Photosynthesis, Carbon Fixation, and Plant Fitness ..................... D. Other Metabolic Processes .................................................... E. Developmental Responses .................................................... F. Plant Hormone Production and Responses to Stress ..................... III. Interplay Between the Clock and Gene Expression. . . . . . . . . . . . . . . . . . . . . . . . . A. Characteristics of the Circadian Transcriptome ........................... B. mRNA Decay and Oscillations in Gene Expression ...................... C. Clock Impact on Transcription .............................................. IV. Architecture of the Plant Clock . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Timing of CAB2 Expression1 (TOC1) and the Pseudo‐Response Regulators (PRRs) ............................................................. B. Gigantea ......................................................................... C. Factors Mediating Light Input Into the Clock ............................ D. Factors Independent of Light ................................................ V. Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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0065-2296/08 $35.00 DOI: 10.1016/S0065-2296(08)00402-3

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ABSTRACT Endogenous time‐keepers known as circadian clocks are setting the pace of multiple metabolic and physiological processes. These complex biological oscillators have arisen in eukaryotic organisms and have been shown to enhance fitness and to confer a competitive advantage by several mechanisms. In general, the plant circadian clock participates in the resonance of the internal biology with optimal time‐of‐day, in coupling the diVerent steps common to one pathway, in gating or partitioning incompatible physiological activities to distinct phases, and in sensing the changes in the seasons and when to flower. Original transcriptional as well as post‐transcriptional mechanisms are involved in the diVerent steps of clock function, from the integration of environmental clues (light quality, temperature) to the interlocked feedback loops composing the core of the oscillator, and the resulting genome scale changes in gene expression. Our current understanding of the intimate mechanisms influencing gene expression at these diVerent steps of clock progression is the focus of this chapter.

I. INTRODUCTION Early evidence of the existence of endogenous rhythms in living organisms was provided by the geophysicist Jean‐Jacques d’Ortous de Mairan (1729). He described the leaf movements of the Sensitive plant (Mimosa pudica) as folding at dusk and opening at dawn. While initially attributed to phototropism, they were observed to still occur when the plant is transferred to darkness. His understanding was that the Sensitive is able to ‘‘sense the sun without seeing it.’’ In this brief publication, he suggested to test the eVect of temperature or of artificially inverted day–night cycles and even extended his observation to patients suVering from sleep disorders. Since this pioneer experiment, biological rhythms with an
24‐h period have been shown to control a wide array of metabolic activities and behaviors in plants (Fig. 1) as well as in other organisms, from unicellular algae (Synechococcus elongatus, Chlamydomonas reinhardtii), to fungi (Neurospora crassa), insects (Drosophila), and animals (rodents, humans). Recent breakthroughs in genomic research have revealed that at the molecular level, underlying these cyclic activities in plants, massive changes occur in gene expression (Fig. 2) in anticipation and in response to predictable changes in the environment. This chapter will first present the pervasive influence of the clock on the biology of the plant and then the focus will shift to the current understanding of the underlying mechanisms broadly aVecting gene expression. Finally, the nature of the known elements composing the plant circadian clock will be described.

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Input Pathway Light Temperature Circadian clock

Gene expression Leaf movements Photosynthesis Secondary metabolism Stomatal aperture Calcium fluxes Photoperiodic flowering Hormone production Hypocotyl elongation Stress responses Output Pathway

Fig. 1. Illustration of the relations between the circadian clock, the input, and the output pathways.

II. BIOLOGICAL PATHWAYS UNDER THE INFLUENCE OF THE CLOCK IN PLANTS A. LEAF MOVEMENTS

Probably the most obvious output of the plant clock, as highlighted by Ortous de Mairan’s experiment, is the control over leaf and petal movements. Similar to the Sensitive plant, folding of the leaves of the rain tree (Samanea saman) is under both clock and light regulation (Moshelion et al., 2002). These movements are believed to arise from periodic volume changes of the cells of the leaf motor organs (pulvini) and variations in osmotic forces due to Kþ‐ion fluxes across the plasma membrane. In this case, the mechanism of clock action would be through the local induction, in the pulvini, of a cycling expression of several genes encoding putative Kþ‐ion channels. However, in other plant species, rhythmic leaf movements appear to be the consequence of the general action of the clock on cell growth and expansion. For instance, hypocotyl elongation in Arabidopsis (Arabidopsis thaliana) exhibits an oscillatory pattern with peaks of growth occurring at subjective dusk (Nozue et al., 2007). Growing leaves also show circadian changes in position: they rise during the day and fall at night, presumably because of anti‐phased oscillations in the elongation of the abaxial and adaxial cells of the petiole. Imaging of leaf or cotyledon movements is routinely used as a direct read‐out of clock function in Arabidopsis (Edwards and Millar, 2007).

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ZT

0

4

8 12 16 20 24 28 32 36 40 44

Peak at ZT0 370 genes

Peak at ZT4 300 genes

Peak at ZT8 422 genes

Peak at ZT12 205 genes Peak at ZT16 163 genes Peak at ZT20 249 genes

146 cycling TF

Fig. 2. All phases are represented in the circadian transcriptome. A heatmap of the expression profiles of the 1708 Arabidopsis genes displaying a high‐amplitude (pMMC
< 0.05) cosine‐wave‐shaped cycling pattern is represented. The bottom panel displays the expression pattern of 146 Arabidopsis cycling TFs. Genes are ordered depending on their phase of expression during a 2‐day time‐course in LL. Peak expression levels are in black and troughs are in white. B. ION FLUXES AND STOMATAL APERTURE

The cytosolic concentration of free Ca2þ ions fluctuates over the course of the day and is under the control of both light and the circadian clock. Using transgenic tobacco (Nicotiana tabacum) and Arabidopsis plants expressing a Ca2þ‐sensitive luminescent aequorin has revealed that these robust oscillations are synchronized and not restricted to specific cell types (Johnson et al., 1995). High intracellular Ca2þ concentrations occur at midday and lower concentrations at night. Fluctuations in free Ca2þ ions are particularly important for the regulation of the aperture of the stomata. The stomatal conductance, resulting from these circadian fluxes, is also at its peak at

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midday, inducing the turgescence of the guard cells and the opening of the stomata (Hennessey and Field, 1991; Somers et al., 1998). Through the modulation of these ion fluxes, in addition to influencing the diverse biological and developmental events mediated by Ca2þ signaling, the clock participates in coupling enhancement of gas exchange through the stomata with a critical peak in photosynthesis activity. C. PHOTOSYNTHESIS, CARBON FIXATION, AND PLANT FITNESS

A great number of genes associated with chlorophyll synthesis, heme production, chlorophyll accumulation, and synthesis of chlorophyll a/b‐binding proteins (CAB) show a cycling pattern of expression with a peak at midday (Harmer et al., 2000). This corresponds to a peak in the activity of the light‐ harvesting complex and it is believed to balance for a rapid light‐induced turnover of proteins of this complex during the day (Dodd et al., 2005). Accordingly, carbon assimilation has been shown to oscillate under constant light (LL), and with constant CO2 levels in several plant species (Hennessey and Field, 1991). Enhancement of photosynthesis during the day is one mechanism by which the clock might increase plant fitness. Dodd and coworkers demonstrated that the accumulation of chlorophyll, carbon fixation, and consequently overall biomass production is increased when the internal period conferred by the clock matches up with that of the external environment (Dodd et al., 2005). For instance, the fitness of wild‐type Arabidopsis plants is enhanced when they are grown in an environment with a 24‐h period, whereas the short‐ or long‐period mutants timing of CAB2 expression1–1 ‘‘(toc11;  = 20.7 h) and zeitlupe (ztl;  = 27.132.5 h)’’ is more adapted to 20‐ or 28‐h period lengths, respectively. In addition, plants with no rhythm (arrhythmia) display a greater disadvantage than do plants with a non‐ matching period. These results highlight two important clock functions: the coupling (resonance) of biological pathways with the optimal time of day and the resulting advantage conferred by such an optimization. D. OTHER METABOLIC PROCESSES

In addition to its implication in carbon homeostasis, the clock is involved in nitrogen and sulfur metabolism (Harmer et al., 2000) and genes involved in the assimilation of these mineral nutrients are phased from the end of subjective night to the beginning of the day. In addition, secondary metabolism is also clock controlled. For instance, enzymes of the phenylpropanoid pathway are co‐regulated by transcription factors (TF) of the R2R3‐MYB family such as PRODUCTION OF ANTHOCYANIN PIGMENTS1 (PAP1,

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Borevitz et al., 2000) in Arabidopsis. Clock control of PAP1 expression coordinates the activation of these enzymes before dawn, activating the production of protective pigments in anticipation of the onset of harmful UV radiations (Harmer et al., 2000). Similarly, the timing of production of volatile compounds of floral scents is tightly controlled by both diurnal and circadian mechanisms (Dudareva and Pichersky, 2000; Dudareva et al., 2003). Production of the volatile terpenes of the scent of snapdragon (Antirrhinum majus) peaks at midday and is coincident with the activity of potential pollinators (Dudareva et al., 2003). Likewise, flowers pollinated by nocturnal insects tend to have maximal scent production in the early evening (Dudareva and Pichersky, 2000). E. DEVELOPMENTAL RESPONSES

Clock‐mediated eVects on development have been particularly well documented for photoperiodic control of flowering and hypocotyl growth (Nozue et al., 2007; Suarez‐Lopez et al., 2001; Yanovsky and Kay, 2002). In these pathways, the circadian clock participates in the induction of switches in developmental programs under specific environmental cues and especially permissive light conditions. The main action of the clock is in fine‐tuning of the expression pattern of key regulators of flowering (CONSTANS, CO) or hypocotyl growth (PHYTOCHROME INTERACTING FACTOR4/5, PIF4/5), while light controls the activity of the product of these genes. In the case of photoperiodic flowering, the clock is responsible for the evening peak of CO expression (Suarez‐Lopez et al., 2001; Yanovsky and Kay, 2002). Upon seasonal extension of the photoperiod, CO expression progressively coincides with inducible light conditions that stabilize CO protein (Jang et al., 2008; Liu et al., 2008; Valverde et al., 2004) and result in a switch from vegetative to reproductive growth. By contrast, light has a repressive eVect on hypocotyl growth by destabilizing PIF4 and PIF5 proteins (Nozue et al., 2007), but the clock participates in restricting hypocotyl growth to a specific phase by delaying the expression of these genes until the end of the night. Interestingly, clock control of cytosolic Ca2þ concentrations could also participate in establishing permissive conditions for flowering induction and hypocotyl growth (Imaizumi et al., 2007; Love et al., 2004), thus highlighting the pervasive action of the clock in regulating specific biological pathways. F. PLANT HORMONE PRODUCTION AND RESPONSES TO STRESS

Production of the plant hormones indole acetic acid (IAA, auxin) and ethylene is under circadian control, with a peak in the middle of the subjective day (Jouve et al., 1999; Thain et al., 2004). For auxin, the clock acts at

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multiple steps, from production to response, with one or more genes involved in each step being under clock regulation (Covington and Harmer, 2007). As a consequence, half of the genes known to be responsive to auxin signaling are also rhythmically expressed, and growth responses to this hormone are gated by the clock (Covington and Harmer, 2007; Jouve et al., 1999). Again, the clock induces oscillations in the expression of ACC SYNTHASE8 gene (ACS8), a key step in the production of an ethylene precursor (Thain et al., 2004). Furthermore, the clock might also have a broader impact on the early events of stress response since the expression of several genes known to be common to multiple abiotic and biotic stresses is also influenced by the clock (Fowler et al., 2005; Walley et al., 2007). The reasons for such regulation are still poorly understood; however, one possibility is that the clock acts in desensitizing these pathways at a time when they are not required or could be potentially deleterious for plant growth and development if active (Fowler et al., 2005).

III. INTERPLAY BETWEEN THE CLOCK AND GENE EXPRESSION A. CHARACTERISTICS OF THE CIRCADIAN TRANSCRIPTOME

With the recent development of microarray technology, high‐throughput monitoring of variations in gene expression has proved to be an invaluable tool for the analysis of the circadian transcriptome. Although the real proportion of genes under circadian regulation is still a matter of debate, independent research groups have all provided compelling evidences for massive oscillations of the transcriptome happening in constant conditions (Blasing et al., 2005; Edwards et al., 2006; Harmer et al., 2000; Michael et al., 2008; SchaVer et al., 2001; Fig. 2). Depending on the stringency of the criteria used for the analysis, 6–16% of the expressed genes display a robust cosine‐ wave‐shaped cycling pattern in constant light (Edwards et al., 2006; Harmer et al., 2000; SchaVer et al., 2001). However, introduction of additional expression patterns and consideration of spike changes of expression in the analysis has allowed Michael et al. (2008) to greatly extend this proportion to almost one third (31%) of the transcriptome. Still, this number is likely to be an underestimate because computational assessment of low amplitude oscillations remains challenging. Also, specific transcript turnover could mask some oscillations occurring at the transcriptional level, as suggested by enhancer trapping experiments (Michael and McClung, 2003). All phases of expression are represented but with a majority of transcripts peaking few

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hours before dawn or dusk (Covington and Harmer, 2007; Michael et al., 2008; Fig. 2). This is consistent with the implication of the clock in the anticipation of these two major transitions in environmental conditions. Circadian oscillations are a subset of the oscillating transcriptome seen in driven conditions. Comparison of microarray time‐courses obtained under diVerent constant (light, LL, or darkness, DD) or driven conditions (photocycles, LD, or thermocycles, HC) has revealed complex interactions of the clock with the other main sources of fluctuations in gene expression: light, temperature, and sucrose (Blasing et al., 2005; Michael et al., 2008). An interaction has been shown in driven conditions, in which light and sucrose induce changes in the amplitude or phase of circadian‐driven oscillations. For instance, diurnal variations in sugar metabolism (starch, sucrose, and trehalose) are a source of fluctuations in gene expression with sugar‐induced genes peaking in the light, whereas sugar‐repressed genes peak in the dark. A significant subset of circadian‐regulated genes (25%) shows a reinforcement of their expression by the super‐imposition of these diurnal changes of sugars (Blasing et al., 2005). By contrast, comparison of short day (StD) and long day (LgD) time‐courses suggest that light and especially the duration of the photoperiod can influence the phase of circadian‐regulated genes (Michael et al., 2008). In these two conditions, morning and evening co‐regulated gene clusters maintain a 12‐h phase diVerence, but they display a 4‐h phase‐lag in LgD compared to StD. As a consequence, the peak of the cluster of evening genes precedes dusk by
6 h in LgD and by
2 h in StD. Similarly, a subset of transcripts, including CAB and all genes peaking in the middle of the day, is shifted to a later phase under LgD. Another intriguing influence of extended photoperiods has been described for a subset of
500 circadian‐regulated genes, which show a second peak of expression under these conditions (Michael et al., 2008). For most genes, thermocycles do not influence the phase of expression. However, for specific sets of genes involved in the cell cycle, DNA, and protein synthesis, thermocycles can antagonize the eVect of light and will dictate the phase over photocycles. Finally, the light stimulus might also be important for sustaining circadian oscillations under constant conditions, since the proportion of circadian transcripts decreases from 31% to 6% under DD compared to LL (Michael et al., 2008). B. mRNA DECAY AND OSCILLATIONS IN GENE EXPRESSION

As reported for mouse ALBUMIN or the Arabidopsis CAB1 genes (Millar and Kay, 1991; Wuarin and Schibler, 1990), long mRNA half‐lives might mask rhythmic expression generated by transcriptional rhythms, suggesting that coordination between transcriptional and post‐transcriptional mechanisms is

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a prerequisite to generating oscillations in mRNA levels. Accordingly, studies on mRNA stability have identified circadian transcripts as being a substantial part of relatively unstable and rapidly degraded transcripts (Gutierrez et al., 2002). In addition to a coordinated action with transcriptional rhythms, specific regulation occurring at the mRNA level could also play an active role in shaping an oscillatory pattern or balancing transcriptional fluctuations. Indeed, loss of a light‐activated negative regulator of CATALASE3 (CAT3) transcript accumulation was suggested to explain the diVerences between the oscillatory pattern observed in LL and a clamped high expression seen in DD (Zhong et al., 1997). Similarly, nuclear run‐on experiments have shown that transcriptional mechanisms are not suYcient to explain the oscillatory expression pattern of the NITRATE REDUCTASE gene (NIA2; Pilgrim et al., 1993). Finally, the transcript of the core clock component CIRCADIAN CLOCK ASSOCIATED1 (CCA1; Wang and Tobin, 1998) displays changes in stability under LD, suggesting that regulation of mRNA decay pathways might be one way for external cues to impact clock‐regulated gene expression (Yakir et al., 2007). In general, these post‐transcriptional mechanisms remain poorly understood. However, recent analyses of CCR‐LIKE (CCL) and SENESCENCE ASSOCIATED GENE1 (SEN1) mRNA decay suggest that one mechanism of clock action might be through the downstream instability degradation pathway (DST; Lidder et al., 2005). Depending of the time‐of‐day, CCL displays variation in its mRNA stability persisting under free‐running conditions. Disruption at this level of regulation in the dst1 mutant leads to a phase‐lag of CCL oscillations correlated with alterations of the half‐life of its mRNA. A similar phase‐lag is also observed for dst1 leaf movements, suggesting that this mRNA decay pathway could play a broader role in the generation of oscillations. Circadian regulation of gene expression could be also mediated by the clock control of RNA‐binding proteins such as COLD AND CIRCADIAN REGULATED2 (CCR2), also known as GLYCINE RICH PROTEIN7 (GRP7; Hassidim et al., 2007; Staiger, 2001). Oscillations in CCR2 transcripts are generated by rhythmic transcriptional activation and remain particularly robust under DD, making it an invaluable tool to monitor the activity of the clock in this condition. Early identification of a genuine RNA‐ binding motif (an N‐terminal RNA Recognition Motif—RRM) in CCR2 suggests that it might confer rhythmic expression through direct interactions with target transcripts (Staiger, 2001). Mutagenesis in the RRM impairs interactions of CCR2 with its targets, including its own mRNA, a transcript for another RNA‐binding protein, GRP8, and other rhythmically and non‐ rhythmically‐regulated transcripts (Schoning et al., 2007). Negative

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regulation of mRNA stability by CCR2 has been linked with the generation of anti‐sense transcripts and would involve the non‐sense‐mediated mRNA decay pathway (NMD; Schoning et al., 2007).

C. CLOCK IMPACT ON TRANSCRIPTION

1. A transcriptional feedback loop is at the core of the clock Although the molecular nature of the components involved might diVer, the general architecture of the eukaryotic clocks rely on interlocking feedback loops, based on transcription and regulated protein turnover (Alabadi et al., 2001; Farre et al., 2005; Harmer et al., 2001; Fig. 3). In plants, light induces the expression of CCA1 (Wang and Tobin, 1998) and LATE ELONGATED HYPOCOTYL (LHY; SchaVer et al., 1998), which encode single MYB domain TFs involved in the induction of clock‐regulated genes activated early in the day, among them, PSEUDO REPONSE REGULATOR7 (PRR7) and PRR9 (Farre et al., 2005). At the same time, CCA1/LHY compose a negative arm of a transcriptional loop directly repressing genes with peak expression in the evening (Alabadi et al., 2001), including TOC1, another member of the PRR family (Strayer et al., 2000). As the levels of CCA1/LHY fall during the course of the day, in part due to the repressive action of PRR7/9 on their expression, TOC1 transcription is alleviated and TOC1 protein level rises (Alabadi et al., 2001; Farre et al., 2005). In an opposing action, TOC1 and additional clock components then feed back to activate (directly or indirectly) the expression of CCA1/LHY, therefore allowing the re‐initiation of the oscillator (Alabadi et al., 2001). The transcriptional mechanisms leading to the repression of TOC1 expression by CCA1/LHY are well documented. These homologous TFs bind directly to the promoter of TOC1 to a specific DNA sequence defined as the evening element (EE; Alabadi et al., 2001; Harmer and Kay, 2005; Perales

Light

Light

CCA1 LHY

Fig. 3.

PRRs

Y

TOC1 LUX

ELF4

Interlocked feedback loops at the core of the Arabidopsis oscillator.

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and Mas, 2007). The binding of CCA1 to TOC1 promoter oscillates during the day and correlates with a reduction in histone H3 acetylation at this locus, consistent with rhythmic changes in chromatin structure (Perales and Mas, 2007). Potentially, along with CCA1/LHY binding, an unknown histone deacetylase would be recruited, thus locking down TOC1 expression till the end of the day. However, the simultaneous ability of CCA1/LHY to activate directly the transcription of morning‐phased genes (PRR7/9 and CAB2) remains intriguing and suggests that additional co‐factors could influence CCA1/LHY activity. The antagonistic mechanisms by which TOC1 and PRR7/9 influence CCA1/LHY expression are unknown and remain an important area of investigation. However, LUX ARRHYTHMO (LUX, also known as PHYTOCLOCK1), a MYB‐related TF of the GARP sub‐family, was recently shown to be important for the progression of the oscillator (Hazen et al., 2005; Onai and Ishiura, 2005). LUX expression peaks in the evening and is co‐regulated with TOC1. In the lux mutant, the oscillations of several clock outputs are arrhythmic in LL including CCA1 and LHY expressions that are clamped low. It is tempting to speculate that LUX might participate, along with TOC1, in the activation of CCA1/LHY (Fig. 3). 2. Identification of phase‐associated CIS‐elements It is likely that co‐regulated genes peaking at a specific time of day are responsive to the same pathway and might share a common promoter motif. Consequently, computational analyses have been successful in identifying several CIS‐elements mediating morning, evening, and other phases of expression (Harmer and Kay, 2005; Harmer et al., 2000; Hudson and Quail, 2003; Michael and McClung, 2002; Michael et al., 2008). Although most of the work on the characterization of these modules has been done in Arabidopsis, some evidence suggests that they are conserved in rice (Oryza sativa ssp. japonica) and poplar (Populus trichocarpa), despite the large evolutionary distance separating these species (Jiao et al., 2005; Michael et al., 2008). a. The evening module. In addition to the core oscillator interlocking feedback loop components, the EE has been shown to control the evening phase of expression of a large number of genes. Initially identified as a motif common to the promoters of a sub‐group of 31 cycling genes with an evening phase of expression, the EE was defined as the nine nucleotide sequence AAAATATCT (Harmer et al., 2000). Its ability to confer an evening phase of expression was confirmed by targeted mutagenesis of the EE sequences found in CCR2 (Harmer et al., 2000), TOC1 (Alabadi et al., 2001), and CAT3 (Michael and McClung, 2002) promoters. It is also presumed to be involved in the regulation of LUX, GIGANTEA (GI ), and EARLY FLOWERING4

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(ELF4) promoters (Hazen et al., 2005; Khanna et al., 2003; Mizoguchi et al., 2002). Furthermore, multimerization of four copies of an EE of the CCR2 promoter was suYcient to confer robust evening‐phased circadian oscillations to a luciferase reporter (Harmer and Kay, 2005). Although the necessity of associating several copies of this motif to generate robust oscillations might seem artificial, the EEs can be found more than once in several evening promoters and even four times in the CCR2 promoter. The potential additive eVects of the multimerization might then reflect a true additive eVect between several similar elements. However, this is not a general trend, suggesting that in native promoters, other motifs might cooperate with the EE to confer the observed high‐amplitude oscillations. Functional EEs can also be found in cycling promoters that peak at a diVerent phase. For instance, PRR9 expression peaks in the morning despite the presence of a genuine EE in its promoter. When isolated and multimerized, the PRR9 EE shows the same temporal activity as the CCR2 EE (Harmer and Kay, 2005). Thus, depending on the context of the promoter, the phase dictated by the presence of an EE is antagonized by other CIS‐ elements. Another attractive hypothesis is that the EE of the PRR9 promoter still contributes to the oscillations, but a new phase is conferred through cooperation with other CIS‐elements. Similar complex interactions are likely responsible for the contribution of the CCA1‐binding site (CBS, AAMAATCT, where M sets for A or C) to the midday activity of the CAB2 promoter (Carre and Kay, 1995; Wang et al., 1997). Although the EE and the CBS are almost identical in sequence, computational analysis with the CBS does not show the same strong link with circadian regulation and with a specific phase of the day (Harmer and Kay, 2005). Still, multimers of some CBSs isolated from the PRR9 and CCR2 promoters confer a robust evening‐ phased rhythm of expression similar to that of the EE (Harmer and Kay, 2005), suggesting an intrinsic activity that confers an evening phase of expression. However, the CBS appears to be less eYcient than the EE in dictating the phase of expression within native promoters and might rely more on interactions with other motifs. One possibility is that the nucleotide changes at the center of the motif weaken the interaction with CCA1/LHY (Harmer and Kay, 2005). Sequences close to the EE might also be involved in conferring an evening phase of expression. Computational analyses have identified another motif of seven nucleotides in length: CTTATCC, called the GATA motif (Hudson and Quail, 2003; Michael et al., 2008). The EE and GATA motifs share the same core sequence TATC (or GATA in reverse complement) that was identified in the regulatory regions of many light and circadian‐responsive genes of monocots and dicots plants, including genes related to photosynthesis,

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such as RBCS and CAB (Terzaghi and Cashmore, 1995). The core of the circadian GATA motif is also related to the I‐box of sequence GATAA. However, the EE and the GATA motif have diVerent flanking sequences, which are known to be involved in the specificity of the binding of TFs (Terzaghi and Cashmore, 1995). Therefore, the proteins binding to the GATA motif might be distinct and/or respond to diVerent signals than the ones interacting with the EE. Consistently, computational analysis predicts a 3–6 h earlier phase of expression for the GATA motif (Michael et al., 2008; Fig. 4). In other organisms, GATA‐binding proteins have been identified that contain a type IV zinc finger DNA‐binding domain. WHITE COLLAR‐1 (WC‐1) and WC‐2, two members of this sub‐class of TFs, are at the core of the circadian clock and blue‐light signaling in Neurospora crassa (Ballario et al., 1996; Linden and Macino, 1997). In Arabidopsis, this DNA‐binding domain is the signature of the GATA TF family which contains 30 members (Bi et al., 2005; Manfield et al., 2007; Reyes et al., 2004; Teakle et al., 2002). Although some of these GATA factors have been shown to bind in vitro to the GATA elements, their involvement in light or circadian regulation remains unclear. Instead, as mentioned above, the MYB‐related factors CCA1/LHY have been shown to interact with the EE in vitro and in planta, and this interaction is mainly responsible for transcriptional repression (Alabadi et al., 2001; Perales and Mas, 2007). However, the EE interacts with multiple TFs, including activators of transcription that would be required to explain the oscillations observed in the multimerization experiments. Consistently, the EE‐binding activity fluctuates during the day with a peak preceding the evening phase and is still detected in nuclear extracts of cca1 lhy double mutants (Harmer and Kay, 2005). CCA1/LHY are members

ME - CCACAC G-box - GCCACGTG GATA - CTTATCC EE - AAAATATCT TBX - AAACCCT G-box + GATA

0

12

24

Fig. 4. Schematic representation of the diVerent phases of expression conferred by the phase‐associated CIS‐elements.

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of a sub‐family of MYB‐related factors which contains 13 other members that are candidates for EE‐binding proteins (Carre and Kim, 2002; Riechmann et al., 2000; Yanhui et al., 2006). In addition, the recent use of a protein microarray containing 119 MYB‐ and 47 MYB‐related factors revealed that several members of these families can interact with the EE in vitro. Four of these EE‐binding proteins display a clock‐regulated expression in microarray time‐courses with an afternoon to evening phase and therefore might contribute to the EE activity (Gong et al., 2008). b. The morning module. As noticed for the EE and the GATA evening module, the identified CIS‐elements that mediate a circadian expression phased in the morning belong to a category of elements also tightly linked to light regulation (Hudson and Quail, 2003; Jiao et al., 2007; Michael et al., 2008). Two main elements have been identified, the morning element (ME, CCACAC) and the circadian G‐box (GCCACGTG). Again, a conserved sequence is at the core of these two elements, here CCAC, but diVerent flanking sequences are likely responsible for variations in phase and mediation of the occurrence of a peak before dawn for the ME and after dawn for the G‐box (Michael et al., 2008; Fig. 4). The wide distribution of these elements indicates that they might be involved in the phase determination of many output genes but their presence in several core clock gene promoters also suggests involvement in the core of the oscillator (Michael and McClung, 2002). However, even if sequences related to these motifs are suYcient to confer a morning phase of expression when multimerized, an in‐depth functional characterization of their circadian properties still remains to be performed. Indeed, the close proximity of an EE and a CCACG sequence in the PRR9 promoter led to the fortuitous discovery that a morning phase is associated with this second motif in the context of a mutated EE multimer (Harmer and Kay, 2005). Implication of sequences similar to the ME in light transduction was recently suggested by the comparison of the promoter sequences of a set of genes responsive to far‐red light signaling (Hudson and Quail, 2003; Jiao et al., 2005). In this analysis, a motif of consensus sequence GCCAC was found to be enriched in the promoters of genes activated by the phytochrome A pathway (PHYA) and was named SORLIP1 for ‘‘sequence over‐represented in light‐induced promoters.’’ Nothing is known about the function of this sequence either for the clock or light signaling. By contrast, the G‐box is one of the most well‐studied CIS‐element in plants, although many aspects of its functionality remain poorly understood. Promoter elements with the core hexameric motif CACGTG are known to be crucial for the activity of many light‐regulated promoters (CAB, RBCS, CHS, RCA) as well as promoters induced by stress (responsive to ABA,

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ethylene and jasmonic acid) and defence responses (Menkens et al., 1995). Approximately one‐tenth of all genes in the Arabidopsis genome possess a G‐box in the 500 bp immediately preceding the translation start site, which makes this relatively frequent motif twice as common in promoter regions compared to coding regions or the genome as a whole (Hudson and Quail, 2003). In order to reconcile this wide distribution of the G‐box with the specificity of its responses to particular stimuli, it was first hypothesized that it could respond to general regulators of transcription (Menkens et al., 1995). The specificity of the response would then be mediated by the promoter context and interactions with an adjacent factor. On this line, extensive work on light‐activated promoters has revealed that rather than relying on a single element, the combination between several diVerent motifs is important (Jiao et al., 2007). Consistently, recent computational analysis confirmed that the G‐box and the I‐box form a composite light‐responsive unit and are associated in a subset of promoters involved in photosynthesis including CAB proteins, diVerent photosystem I reaction center subunits, photosystem II associated proteins, and ferredoxin (Vandepoele et al., 2006). Whether these interactions are the result of additive or synergistic regulations remains unknown, but this would not be a general trend since no systematic association of the G‐box with a specific motif has been found apart from a possible tendency for duplication and the association with another G‐box (Freeling et al., 2007). Interestingly, the related E‐box core element (CANNTG) frequently in full G‐box form is also known to be an important motif of insect and mammalian circadian promoters. However, generation of high‐amplitude oscillations by this motif would require a combination of a canonical E‐box (CACGTG) and an E‐box‐like sequence (CANNTG), in tandem, with a short interval between them (Munoz et al., 2002; Nakahata et al., 2008; Paquet et al., 2008). Variations in the sequences directly flanking the G‐box is another mechanism thought to mediate the specificity of the response of this motif to certain stimuli (Hudson and Quail, 2003; Jiao et al., 2005; Menkens et al., 1995). Particular flanking sequences responsible for PHYA‐mediated induction of gene expression have been identified, whereas other sequences would mediate PHYA repression (Hudson and Quail, 2003). In addition, the flanking sequences might be involved in the tissue specificity of the light response and distinct sequences would be associated with cotyledon and/or root expression (Jiao et al., 2005). Consistently, extensive characterization of G‐box‐binding proteins (GBP) by gel‐retardation assays has shown great variations in the aYnity for the diVerent elements containing a CACGTG core depending on the nature of the flanking sequences (Foster et al., 1994). As in other organisms, the plant GBPs belong to the bHLH and bZIP

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families of TFs (Heim et al., 2003; Jakoby et al., 2002). Members of these families can form homo‐ as well as hetero‐dimers and can acquire unique DNA‐binding activities through these interactions, thus enhancing the variety of potential G‐box‐binding complexes and conferring another level of complexity (Fairchild et al., 2000; Schindler et al., 1992, Toledo‐Ortiz et al., 2003). Light‐regulated interactions of the bZIP proteins HY5, GBF, and the bHLH factors homologous to PIF3 with the G‐box is particularly well‐ documented (Chattopadhyay et al., 1998; Mallappa et al., 2006; Martinez‐ Garcia et al., 2000; Menkens et al., 1995). However, their potential implication in the activity of the circadian G‐box remains unclear (Oda et al., 2004; Viczian et al., 2005). In addition, similar to what has been observed for the EE, the circadian G‐box likely interacts with multiple GBPs, including both activators and inhibitors of transcription.

3. CIS‐elements associated with other phases of the day In addition to the morning and the evening module, it is likely that several additional motifs may participate in phasing gene expression to other phases of the day. A group of motifs, all containing a stretch of three C, was recently shown to confer a midnight phase of expression and it includes the telobox (TBX, AAACCCT), the protein box (PBX, GGCCCAT), and the sucrose box (SBX, AAGCCC) (Michael et al., 2008; Fig. 4). These motifs are largely over‐represented among promoters of genes associated with protein synthesis. Intriguingly, under thermocycles only, the phase conferred by this motif is the opposite (morning) of the phase conferred in light cycles (midnight). The TBX was shown to interact in vitro with an Arabidopsis protein containing a purine ‐like DNA‐binding domain, a signature of proteins interacting with purine‐rich motifs (Manevski et al., 2000; Tremousaygue et al., 2003). TBX was initially implicated in cell‐cycle regulation and sugar signaling, in combination with other motifs, the specific night phase conferred by the TBX is thought to limit the coincidence between DNA replication and harmful UV irradiation (Michael et al., 2008). As mentioned above, complex interactions between diVerent sets of CIS‐ elements conferring morning and evening phases of expression might also result in new phases of expression. For example, in the promoters of genes involved in photosynthesis, motifs conferring a morning phase and similar to the G‐box are associated with evening‐phased motifs of the GATA sub‐type (Vandepoele et al., 2006). The resulting phase of expression is intermediate to the phase conferred by these motifs when they are isolated, and genes have a midday peak of expression (Dodd et al., 2005; Harmer et al., 2000; Fig. 4).

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IV. ARCHITECTURE OF THE PLANT CLOCK Despite its pervasive action in generating oscillations in gene expression, only a few genuine TFs have been identified as central clock components, the MYB‐related factors CCA1/LHY (SchaVer et al., 1998; Wang and Tobin, 1998) and LUX (Hazen et al., 2005; Onai and Ishiura, 2005). In contrast, genetic screens for altered clock or flowering phenotypes have led to the identification of a collection of proteins of unknown function and, for the most part, unique to plants. These factors might be involved in regulating the activity of known or yet to be discovered central clock components. A. TIMING OF CAB2 EXPRESSION1 (TOC1) AND THE PSEUDO‐RESPONSE REGULATORS (PRRs)

Mapping of the toc1–1 mutation implicated a sub‐family of proteins containing an atypical phospho‐accepting receiver‐like domain (pseudo‐receiver domain, PR) and a basic motif also found in the CO family of TFs (CCT domain for CO, CO‐like, and TOC1) in the plant clock (Strayer et al., 2000). The short‐period phenotype exhibited by this mutant in a variety of rhythms (CAB2:LUC, CCR2:LUC, and the stomatal conductance) as well as the dose‐dependent eVect of TOC1 protein on the period determination (Mas et al., 2003a; Somers et al., 1998) led to the placement of TOC1 at the core of the oscillator. Consistent with its nuclear localization (Makino et al., 2000; Strayer et al., 2000), TOC1 expression in the evening is presumed to activate, directly or indirectly, the transcription of CCA1/LHY (Alabadi et al., 2001). TOC1, also called PRR1, is the founding member of a small gene family of five members (TOC1 ¼ PRR1, PRR3, PRR5, PRR7, PRR9) all containing the PR and CCT domains in their sequence (Matsushika et al., 2000; Strayer et al., 2000). The PRR genes display a robust circadian expression with successive waves of expression spanning from the morning to the evening in the order PRR9, PRR7, PRR5, and PRR3/TOC1, these two latter evening PRRs peaking at the same phase of the day. Although toc1 mutations have the most dramatic eVect on the clock, mutant and over‐expression analyses have demonstrated the importance of the other PRRs in clock progression. First, the morning PRRs, PRR7 and PRR9, participate in CCA1/LHY expression as components of a feedback loop. The single mutants display a long‐period phenotype that is enhanced in the double mutant prr7 prr9 (Farre et al., 2005; Salome and McClung, 2005). Accordingly, PRR9 over‐ expressing plants have a short‐period phenotype (Matsushika et al., 2002), whereas highly increased levels of PRR7 severely compromise the clock (Farre and Kay, 2007). In an opposing action, PRR3 over‐expression,

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similarly to a moderate increase in TOC1 levels, lengthens the period (Mas et al., 2003a; Murakami et al., 2004). PRR5 over‐expression does not have a strong eVect on period determination. Instead, the amplitude of expression of the morning genes CCA1 and especially LHY was shown to be decreased (Sato et al., 2002). Such low amplitude of CCA1/LHY expression has also been recently described for PRR7 over‐expressing plants (Farre and Kay, 2007). Together with the additive eVects on clock progression observed in prr9 prr7, prr7 prr5, and prr9 prr7 prr5 combinations of mutations (Farre et al., 2005; Nakamichi et al., 2005a,b; Niinuma et al., 2008; Salome and McClung, 2005), the results of the over‐expression experiments suggest that the PRRs fulfill partially overlapping functions at the core of the oscillator. Consistent with a dose‐dependent eVect of several of the PRRs on the period determination of the oscillator, the accumulation of these proteins is tightly regulated and turns out to be a key checkpoint of clock progression. Blue‐light activation of the ZTL F‐box protein was shown to target TOC1 and PRR5 proteins for degradation by the proteasome and is thought to refine the circadian wave‐forms of expression of these two proteins (Kiba et al., 2007, Mas et al., 2003b). Similarly, PRR7 protein turnover was shown to be under both circadian and light regulation (Farre and Kay, 2007). In addition, recent insights on PRR3 function suggest that tissue‐specific regulated turnover of TOC1 protein might have an important role in particular clock outputs such as photoperiodic flowering (Para et al., 2007). In contrast with the quasi ubiquitous activity of the TOC1 promoter, PRR3 promoter activity is restrained to the vascular tissue. The temporal activity of vascular‐ specific reporters is more aVected by the prr3 mutation than by reporters widely expressed such as CAB2:LUC and CCR2:LUC, in accordance with a tissue‐specific activity of the PRR3 protein. In these cells, through a direct interaction with TOC1, PRR3 appears to specifically antagonize ZTL action, and be responsible for a local enrichment in TOC1 protein. Although the general mode of action of the PRRs on clock progression remains unknown, several lines of evidence point toward an implication in transcriptional regulation. First, TOC1 was implicated in several direct interactions with TFs, including the seed developmental regulator ABSCISIC ACID INSENSITIVE3 (ABI3; Kurup et al., 2000) as well as several members of the PIF sub‐family (Yamashino et al., 2003). However, these interactions were characterized only in yeast and their real significance in planta remains poorly understood. More recently, the functional analysis of the CCT domain of CO has revealed a structural similarity with a sub‐ domain of proteins of the Heme Activator Protein complex (HAP; Wenkel et al., 2006). Through direct protein interactions, mediated by the CCT domain, and involving AtHAP3 and AtHAP5, CO would take part in a

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heterotrimeric HAP complex promoting flowering. HAP complexes are CCAAT‐box‐binding complexes, a CIS‐element present in
25% of eukaryotic promoters. The gene family encoding the HAP proteins has greatly expanded in plants with 26 potential homologs of AtHAP3 and AtHAP5. Complex interaction matrices between these AtHAPs and the 40 CCT‐ containing proteins found in Arabidopsis is believed to specify the activation of multiple transcriptional programs (Wenkel et al., 2006) and would link the CCT domain with DNA‐binding complexes. B. GIGANTEA

Identified in several independent genetic screens, the gi mutant displays a pleiotropic phenotype, consequence of GI implication in multiple pathways, including a role in clock regulation (Eimert et al., 1995; Fowler et al., 1999; Huq et al., 2000; Park et al., 1999). gi was first identified as a late flowering mutant insensitive to the photoperiod (Eimert et al., 1995; Fowler et al., 1999; Park et al., 1999). This mutant was also shown to accumulate increased amounts of starch in photosynthetic tissues, independently of the late flowering characteristics (Eimert et al., 1995). In addition, gi was isolated in a screen for PHYB signaling deficient mutants and it has a long hypocotyl phenotype under red light (Huq et al., 2000). However, more recently, gi was also shown to be impaired in the inhibition of hypocotyl elongation under blue light and very low fluences of far‐red light (VLFR), suggesting that its role in hypocotyl elongation is not restricted to PHYB signaling (Martin‐Tryon et al., 2007; Oliverio et al., 2007). On this line, several other aspects of PHYA signaling under VLFR are aVected in gi such as cotyledon unfolding and seed germination (Oliverio et al., 2007). Clock defects in gi include a short‐period phenotype for most of the outputs tested, such as leaf movements and CAB2:LUC rhythms (Fowler et al., 1999; Park et al., 1999). In accordance with a role of GI in light signaling, the fluence‐dependent period lengthening is less pronounced and the clock is less aVected under DD than under LL in gi (Park et al., 1999). At the core of the oscillator, the expression of CCA1 and LHY was shown to be altered with reduced amplitude (Fowler et al., 1999; Park et al., 1999). As observed for cca1 lhy double mutant, an early flowering phenotype would be expected from this short‐period mutant (Mizoguchi et al., 2005). In contrast, gi is late flowering and genetic analyses have demonstrated that this apparent inconsistency is the consequence of an intimate action of GI in flowering regulation and an activation of CO expression (Mizoguchi et al., 2005; Sawa et al., 2007). GI would then function in several pathways in addition to the clock and some of the pleiotropic phenotypes of gi mutants represent separable

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roles for the protein (Martin‐Tryon et al., 2007; Mizoguchi et al., 2005). On this line, an allele of gi (gi‐200) has been identified that is aVected in the clock but not in CO expression, suggesting that GI function in flowering can be biochemically separated from its function in the clock (Martin‐Tryon et al., 2007). The cloning of GI revealed that it encodes a large plant‐specific protein (1173 AA; Fowler et al., 1999; Huq et al., 2000; Park et al. 1999). Despite the presence of several putative trans‐membrane domains in its sequence (Fowler et al., 1999; Park et al., 1999), GI was shown to be localized throughout the nucleoplasm (Huq et al., 2000; Mizoguchi et al., 2005). The levels of GI protein are cycling with an accumulation during the day and a decline at night (David et al., 2006), but the nuclear localization is not aVected by diVerent light and dark treatments (Huq et al., 2000). GI protein was shown to interact directly with several key proteins and to directly modulate their activity. First, an interaction with SPINDLY (SPY) was demonstrated. Encoding an O‐linked
‐N‐acetyl glucosamine transferase, SPY is working as a negative regulator of gibberellin signaling (Tseng et al., 2004). Genetic analyses suggest that GI would act in antagonizing SPY action, potentially by inhibiting SPY‐mediated post‐translational modifications of specific clock components. Another mechanism of GI function is its direct interaction with the F‐box proteins ZTL and FKF1 (Kim et al., 2007; Sawa et al., 2007). Activated by blue light, GI–ZTL interaction results in stabilizing both GI and ZTL proteins, thus refining their respective circadian wave‐forms of expression. As a consequence, the amplitude of TOC1 protein rhythms is enhanced (Kim et al., 2007). Similarly, GI–FKF1 interaction depends on blue‐light activation (Sawa et al., 2007). Chromatin immuno‐precipitation experiments have shown that this complex is assembled on CO chromatin and promotes flowering by targeting for degradation an inhibitor of CO transcription, CDF1. C. FACTORS MEDIATING LIGHT INPUT INTO THE CLOCK

1. Early flowering3 ELF3 was first identified through a mutation (elf3) causing early flowering under both StD and LgD conditions (Zagotta et al., 1996). However, elf3 is also impaired in several aspects of seedling photomorphogenesis and has elongated hypocotyl and petioles as well as pale leaves, phenotypes that are usually associated with defects in light perception. Later work suggested that most of the aspects of elf3 phenotype could also be linked to clock dysfunctions. Indeed, all the clock outputs analyzed in this mutant (CAB2 and CCR2 expression, leaf movements) are totally arrhythmic after one day in LL

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(Covington et al., 2001; Hicks et al., 1996; McWatters et al., 2000), which makes it a remarkably severe phenotype. In addition, rhythms under LD conditions are aVected. For instance, CAB2:LUC displays an increased response to dawn activation and a box‐shaped pattern enhanced under extended daylength (as opposed to the wild‐type wave‐forms; Hicks et al., 1996). In contrast, no such dramatic impairment of the clock can be seen under DD, and CCR2:LUC displays robust oscillations in these conditions with a period similar to the wild type, although with a reduced amplitude (Covington et al., 2001). Temperature entrainment can also bypass the requirement for ELF3 function and partially restore rhythmic CAB2 expression (McWatters et al., 2000). The light‐dependence of this mutant phenotype suggests that ELF3 is not a core clock component but instead it is thought to regulate the light input. The strong acute responses of CAB2:LUC in the mutant after light pulses during subjective night are consistent with the role of ELF3 in the gating of light signaling (McWatters et al., 2000). During the arrest of the oscillator in the mutant in LL, TOC1 expression gets clamped at a high level and CCA1/LHY expression collapses (Alabadi et al., 2001; SchaVer et al., 1998). This corresponds to the peak of ELF3 transcript and protein expression (Covington et al., 2001; Liu et al., 2001). Thus, it was hypothesized that ELF3 functions by allowing progression of the clock through a light‐sensitive phase around dusk by repressing the light input into the oscillator. Cloning of ELF3 revealed that it encodes a protein of 695 amino acids without any previously characterized functional domains with the exception of the presence of a proline‐rich region, an acidic region, and a threonine/ glutamine‐rich region that could potentially play a role in transcriptional activation (Hicks et al., 2001). These regions are conserved in several plant proteins, including a very similar protein in Arabidopsis named ESSENCE OF ELF3 CONSENSUS (EEC, Liu et al., 2001); however, a great variation has been noted in the number of glutamines composing the glutamine‐rich region in diVerent Arabidopsis wild‐type accessions (7–29 elements; Tajima et al., 2007). The absence of a genuine DNA‐binding domain suggests that ELF3 would act as a co‐factor. In accordance with this hypothesis, antibodies raised against ELF3 and immunoblots performed after sub‐cellular fractionation revealed the presence of the protein in the nuclear fraction (Liu et al., 2001). The accumulation of the protein in this compartment oscillates with a peak occurring at dusk in both LD and LL, and mainly follows the activity of the promoter (Covington et al., 2001; Liu et al., 2001). However, a stronger accumulation of the protein in LL, and the fact that it is hardly detected in DD, suggests a potential implication for light in ELF3 stabilization (Liu et al., 2001). In addition to the formation of homo‐dimers,

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a yeast two‐hybrid screen has revealed the ability of the N‐terminal part of the protein to interact with the C‐terminal portion of PHYB (Liu et al., 2001). Interestingly, genetic analyses indicate that ELF3 is involved in PHYB signaling in early morphogenesis (hypocotyl elongation) and this interaction could account for some aspects of ELF3 function (Liu et al., 2001). 2. Early flowering4 Similarly to elf3, elf4 displays a pleiotropic phenotype: early flowering independent of the photoperiod, elongated hypocotyl and petioles, and a severe impairment of the clock in LL (Doyle et al., 2002). However, some discrete diVerences exist between these two mutants. For instance, the long hypocotyl phenotype in elf4 is sensitive to the photoperiod and is more pronounced under StD, whereas elf3 is elongated under both StD and LgD (Doyle et al., 2002). Moreover, the total leaf number at the time of flowering is significantly higher in elf4 than in elf3, indicating that elf3 flowers earlier than elf4 (Doyle et al., 2002). Concerning the clock, the oscillations of several outputs (CAB2, CCR2, and leaf movements) are strongly impaired and after the first day in LL, the amplitude of the oscillations damps and a great variability is observed in the period of these weak rhythms (Doyle et al., 2002). An enhanced response to dawn of several morning reporters as well as a greater sensitivity of CAB2: LUC to light pulses in the mutant indicate that ELF4 is also involved in the gating of light input into the clock (McWatters et al., 2007). Interestingly, light signaling pathways downstream of the PHYs have been shown to converge at the regulation of ELF4 expression. A rapid induction of ELF4 is seen within 1 h of a far‐red light pulse and is characteristic of a response to PHYA (Tepperman et al., 2001). Long‐term response of ELF4 to a red‐light pulse is aVected in phyB; and elf4 seedlings are impaired in several aspects of PHYB‐mediated de‐etiolation, consistent with an action of ELF4 downstream of PHYB (Khanna et al., 2003). Finally, ELF4 expression in etiolated seedlings exposed to red light was changed in an elf3 background. Along with the strong similarities between the corresponding mutants, this result is consistent with an action of ELF3 and ELF4 in the same pathway, ELF4 acting downstream of ELF3 (Kikis et al., 2005). The expression of several core clock genes is severely impaired in elf4. After 1 day of release in LL, CCA1 and LHY expression collapses, whereas TOC1 is clamped at a high level (Doyle et al., 2002; Kikis et al., 2005; McWatters et al., 2007). ELF4 requirement for the expression of CCA1/ LHY was also observed in etiolated seedlings exposed to red light (Kikis et al., 2005). By contrast, TOC1 expression was unchanged in these experiments, suggesting that elf4 would act on the clock mainly by inducing the

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expression of the morning genes. ELF4 function appears to be similar to that of ELF3 and TOC1 (Kikis et al., 2005). CCA1 and LHY can feedback negatively to regulate ELF4 expression, thus composing another key regulatory feedback loop proposed to act at the core of the oscillator (Kikis et al., 2005; McWatters et al., 2007; Fig. 3). ELF4 encodes a 111 AA protein with no significant homology to proteins of known function (Doyle et al., 2002; Khanna et al., 2003). It is the founding member of a small family containing five other genes in Arabidopsis. Interestingly, a stretch of 26 AA is strictly conserved in all the members, indicating importance for this motif. Consistent with a role in gene regulation, a fusion between ELF4 and GFP is targeted to the nucleus of onion epidermal cells (Khanna et al., 2003). 3. Sensitivity to red‐light reduced1 (SRR1) Identified in a screen for mutants with an elongated hypocotyl at low intensities of white light (Staiger et al., 2003), srr1 has reduced sensitivity to red light and is impaired in several PHYB‐mediated processes such as chlorophyll biosynthesis, petiole elongation, and flowering. However, the phyB srr1 double mutant displays stronger phenotypes than the individual single mutants in white light, suggesting that SRR1 might also act independently of PHYB. In constant conditions, clock outputs (CAB2, CCR2, CAT3, and leaf movements) as well as clock components (CCA1, TOC1, FKF1, and GI ) have a shorter period, lower amplitude, and faster damping in srr1. The acute response of CAB2:LUC to light is enhanced, consistent with a function of SRR1 in the gating of light input. By contrast with elf3 and elf4, the clock is also aVected in DD (short period and fast damping), indicating that its clock action is not restricted to light signaling. Despite the absence of any recognizable domain in the sequence of SRR1 protein, homologs can be found in other organisms and suggest a conservation of its function in eukaryotes. A functional SRR1–GFP fusion revealed the presence of the protein in both the cytoplasm and the nucleus. SRR1 expression is not under the control of the clock but is induced by red light. 4. Time for coVee (TIC) The tic mutation was identified in an EMS‐mutagenized population of CAB2:LUC plants and displays a reduced amplitude and a short‐period phenotype (Hall et al., 2003). CCR2:LUC in tic has also a similar low amplitude in LL with a large variability in period length, although neither of these two reporters becomes arrhythmic in the mutant. A partial defect in circadian gating was revealed in tic which would be consistent with a function of TIC in the mid to late subjective night. In a tic elf3 double mutant

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background, impairment of the clock is more dramatic than in elf3 or tic alone, suggesting that TIC is involved in a light input pathway distinct from ELF3. Accordingly, tic plants display diVerent morphological phenotypes than elf3 and are small with a short hypocotyl. They also flower early under both StD and LgD. The impairment of the expression of several core clock genes in the mutant indicates that TIC is working close to the oscillator (Ding et al., 2007; Hall et al., 2003). The amplitude of the morning genes CCA1/ LHY is reduced but LHY is more severely aVected than CCA1, suggesting that it is a dominant genetic target of TIC (Ding et al., 2007). Interestingly, this would also suggest that CCA1 and LHY expressions are influenced by distinct pathways. However, diVerences between the phenotype of tic and cca1 lhy also indicate that TIC might have additional target genes besides LHY. TIC encodes a nuclear protein of 1550 amino acids and one homolog can be found in the Arabidopsis genome (Ding et al., 2007). TIC mRNA and protein levels do not cycle, suggesting that the clock does not feedback on TIC expression. The presence of an ATP/GTP‐binding site A motif (P‐loop) in its sequence might relate TIC activity to elements of a kinase cascade. 5. Light‐insensitive period1 (LIP1) lip1–1 was identified as a short‐period mutant for the CAB2:LUC reporter in LL (Kevei et al., 2007). This short‐period phenotype was also described for CCR2:LUC (both in LL and DD) and leaf movement rhythms, as well as for CCA1, LHY, and TOC1 expression; no eVect was seen on the amplitude of these rhythms. In the wild type, the period of the oscillator is known to increase with lower fluence rates of light, but no increase in period length is seen in lip1–1, suggesting a reduced sensitivity of the mutant to light quality. By contrast, the phase response curves of CCR2:LUC indicate hypersensitivity to light resetting in the first half of the subjective night. The cloning of LIP1 revealed that it encodes an atypical GTPase localized in both nuclear and cytosolic compartments. Its ability to hydrolyze GTP was verified in vitro. LIP1 is believed to function as a negative factor controlling the light‐dependent shortening of the period. 6. Far‐red elongated hypocotyl3 (FHY3) Specifically impaired in PHYA signaling, fhy3 shows decreased responsiveness to constant monochromatic far‐red light inhibition of hypocotyl elongation, cotyledon opening, and production of anthocyanins (Whitelam et al., 1993). A light‐dependent impairment of the clock has also been described in this mutant with arrhythmicity of CAB2:LUC in LL (Allen et al., 2006). However, unlike in elf3 and tic mutants, which display arrhythmicity under all light conditions, the clock is still running under blue light in fhy3,

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although the period is shorter. Thorough analysis of light input defects, and more precisely release from light experiments and phase response curves, has shown that fhy3 is defective in the gating of light input during the subjective day. Along with its action in other PHYA signaling pathways, FHY3 is another element in the complex machinery of the gating of light input into the clock. It is a member of the family of Mutator‐like transposing elements and new insights into FHY3 mode of action have recently been provided (Lin et al., 2007). It has been shown that FHY3 functions as a TF and directly activates the expression of FHY1 and FHL, two proteins required for PHYA translocation into the nucleus. Sequences homologous to the FHY3 DNA‐ binding site (CACGCGC) are found in the CCA1 and ELF4 promoters, and FHY3 can interact with these promoters in yeast one‐hybrid experiments (Lin et al., 2007). A direct activation of CCA1 by FHY3 would be consistent with the decrease in CCA1 expression observed in fhy3 (Allen et al., 2006). D. FACTORS INDEPENDENT OF LIGHT

1. TEJ tej was identified as a long‐period mutant in a genetic screen for altered CAB2:LUC expression (Panda et al., 2002). An
2 h lengthening of the period was described for all tested clock outputs without any change in amplitude. This eVect was independent of light quality, suggesting that TEJ does not function in light input into the oscillator. TEJ encodes a poly ADP‐ribose glycohydrolase (PARG), an enzyme degrading poly ADP‐ribose polymers (pADPr) resulting from the action of polymerases (PARP). Steady‐ state mRNA of TEJ and PARP do not cycle in Arabidopsis. An over‐ accumulation of pADPr was found in the seedlings of tej and this phenotype can be antagonized by the use of inhibitors of poly ADP ribosylation, which also rescues the slowing of the oscillator. This direct correlation between poly ADP ribosylation and period length was proposed to result from the modification of a target TF that may either enhance expression of a repressor or suppress a positive acting element of the clock thus leading to slowing the pace of the oscillator. 2. FIONA1 (FIO1) A recent genetic screen has identified fio1 as a novel early flowering mutant insensitive to photoperiod with an elongated hypocotyl (Kim et al., 2008). Similarly to tej, fio1 displays a 2–3 h lengthening of the period of all clock outputs, independently of light quality and temperature. FIO1 encodes a nuclear protein conserved in prokaryotes and eukaryotes with a DUF890 domain, a signature of the methyltransferase superfamily. It is believed to

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function as an S‐adenosyl‐L‐methionine‐dependent methyltransferase. Like TEJ and SRR1, genes homologous to FIO1 can be found in other organisms. Similarly, it does not fulfill all the criteria of a classical clock component as the clock does not control FIO1 expression and FIO1 over‐expression does not aVect the clock. However, like TEJ, FIO1 function may be required to modulate the activity of a core clock component controlling the pace of the oscillator.

V. CONCLUSION AND PERSPECTIVES In the past years, tremendous progress has been made in our understanding of the plant clock. Genetic approaches conducted in the model species Arabidopsis has led to the identification of several genes involved in the light input pathways or directly at the core of the oscillator (this chapter). Extensive functional analysis of these clock components has allowed proposing an interlocked feedback loop model for the plant clock relying on transcriptional mechanisms and regulated protein turnover (Alabadi et al., 2001; Farre et al., 2005; Harmer et al., 2001; Kikis et al., 2005; McWatters et al., 2007; Fig. 3). Recent modeling approaches have confirmed the eYciency of these feedback loops to confer robust oscillations at the core of the oscillator (Locke et al., 2005, 2006; Zeilinger et al., 2006). However, these approaches have also highlighted that some components still need to be identified in order to complete our model. Among those challenges facing the clock field is the identification of the mechanisms of CCA1 and LHY expression that link the evening components of the oscillator to the morning components. Although TOC1 repression by CCA1/LHY is well documented (Alabadi et al., 2001; Harmer and Kay, 2005; Perales and Mas, 2007), the mechanisms leading to its activation remain also to be characterized. A great number of factors have been implicated in the light input into the oscillator and this pathway appears to be complex. It is likely that resolving the mechanisms of light input is one key to deciphering the mechanisms of CCA1, LHY, and TOC1 activation. Use of the microarray technology coupled with computational biology has proved to be an invaluable tool for assessing the extent of clock influence on gene expression and identifying the pathways targeted by the clock (Blasing et al., 2005; Edwards et al., 2006; Harmer et al., 2000; Michael et al., 2008; SchaVer et al., 2001). One‐third of all plant genes oscillate in LL, indicating that the clock is a major factor driving rhythms in gene expression. However, interactions between the clock and other important factors of fluctuations in gene expression (light, temperature, nutrient availability) are responsible for the oscillatory pattern observed in diurnal conditions (Blasing et al., 2005;

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Michael et al., 2008). While the molecular bases of these interactions remain to be investigated, a comprehensive view of the mechanisms responsible for clock‐driven oscillations in gene expression is emerging. The clock is acting at multiple levels by controlling the phase of expression and/or activity of key components of transcription and mRNA decay machineries, such as for instance TFs (Fig. 2) and RNA‐binding proteins. Several CIS‐elements have been identified that are implicated in specific phases of expression: the ME and the circadian G‐box confer a morning‐phased expression, whereas the GATA, EE and TBX are associated with an evening to midnight expression (Harmer and Kay, 2005; Harmer et al., 2000; Hudson and Quail, 2003; Michael and McClung, 2002; Michael et al., 2008). More work need to be done on the characterization of these motifs and their respective binding proteins. In addition, these elements might be suYcient to confer rhythmic expression in certain conditions; however, some evidences also suggest that they might cooperate with other elements that remain to be identified (Harmer and Kay, 2005). Another attracting field of investigations is the nature of the interactions (cooperation or antagonism) between morning‐ and evening‐phased CIS‐elements when they are present in the same promoter (Vandepoele et al., 2006). Interestingly, some of these transcriptional mechanisms broadly aVecting clock outputs are also present at the core of the oscillator and their study should bring new insights in the central mechanisms of the interlocked feedback loop. Finally, although major progresses in the field have been performed, thanks to the wealth of genetic resources available for the model species Arabidopsis, new models are emerging for the study of the plant clock. The recent completion of the genome sequences of the unicellular algae Chlamydomonas reinhardtii (Matsuo et al., 2008; Merchant et al., 2007) as well as of several higher plants (for instance rice and poplar; GoV et al., 2002; Tuskan et al., 2006; Yu et al., 2002) is opening new areas of investigation. These new resources raise many interesting questions for our field as for instance: are the same genes present in other plants, are they in identical or diVerent copy numbers, and is their function conserved or slightly diVerent? Answers to these questions are likely to give interesting insights into the evolution of the biological clock in plants.

ACKNOWLEDGEMENTS We thank the members of the Kay Laboratory for critical reading of the manuscript and helpful comments during the preparation of this chapter. Work on the plant clock in the Kay Laboratory is supported by National Institute of Health Grants GM‐56006 and GM‐67837.

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Late Embryogenesis Abundant Proteins

MING‐DER SHIH,* FOLKERT A. HOEKSTRA,{ AND YUE‐IE C. HSING*

*Institute of Plant and Microbial Biology, Academia Sinica, Taipei 11529, Taiwan { Laboratory of Plant Physiology, School of Experimental Plant Sciences, Wageningen University, Wageningen NL‐6703 BD, The Netherlands

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Water Status in Living Organisms ........................................... B. Seed Development and Lea Proteins ........................................ II. Classification and Occurrence of Lea Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Classical Classification......................................................... B. Popp Classification ............................................................. C. The Annotation of Arabidopsis Lea Genes ................................ III. The Structure of Lea Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Lea Proteins in Solution Status .............................................. B. Lea Proteins in Dry Status .................................................... C. Hydrophobic Lea Proteins .................................................... D. Hydrophilic Lea Proteins Are Members of Natively Unfolded Proteins IV. The Physiological Roles of Lea Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Accumulation of Lea Proteins During Dehydration ................ B. Ectopic Expression of Lea Proteins.......................................... C. In Vitro Studies ................................................................. D. Ion Scavenger ................................................................... E. Recalcitrant Seeds .............................................................. F. Other Possible Mode of Action .............................................. V. Conclusion and Future Perspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Advances in Botanical Research, Vol. 48 Incorporating Advances in Plant Pathology Copyright 2008, Elsevier Ltd. All rights reserved.

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0065-2296/08 $35.00 DOI: 10.1016/S0065-2296(08)00404-7

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ABSTRACT During the late maturation stage of seed development, water content decreases greatly. One of the most striking characteristics of mature orthodox seeds is their ability to withstand severe desiccation. Mechanisms of plant drought/desiccation tolerance have been studied by numerous groups, and a broad range of molecules have been identified to play some roles. Examples are proline, oligosaccharide, and late embryogenesis abundant (LEA) proteins, and so on. LEA proteins were first described from mature cotton seeds decades ago. Since then, many LEA proteins were identified from vascular and nonvascular plants, fungi, algae, and microbes, as well as anhydrobiotic animals such as protozoa, nematodes, insects, and crustaceans, and so on. The extensive distribution of LEA genes among diverse taxa implies that these genes might be primitive yet important and therefore maintained by these species. As a result of evolution, they may have a certain universal function—osmoprotection. Hydrophilic LEA proteins are members of natively unfolded proteins in solution. After the removal of bulk cytoplasmic water, the structures of LEA proteins undergo desiccation‐ induced folding. These biophysical features suggest that LEA proteins may carry out a bipartite function under diVerent water states. During drought, LEA proteins may establish a water shell and decrease ion strength. After desiccation, they may enhance the bioglass strength and act as a water replacement to stabilize cellular components.

I. INTRODUCTION A. WATER STATUS IN LIVING ORGANISMS

Water is the most important element for any living organism. Dehydration of a living cell has great physical and chemical eVects on cellular and macromolecular structures and usually causes irreparable changes, which ultimately lead to cell death. Hence, in terms of critical water level, two types of tolerance for living organisms are distinguished. ‘‘Drought tolerance’’ can be described as tolerance to moderate dehydration, with no bulk cytoplasmic water, which is present with
23% water on a fresh weight basis or
0.3 g H2O/g dry weight. ‘‘Desiccation tolerance’’ usually refers to tolerance to further dehydration, when the hydration shell is gradually lost (Hoekstra et al., 2001). Some organisms can survive under extreme aridity, even with removal of 99% of their body water. The dry organisms may continue in this unique living state and resume vital metabolism upon rehydration (Crowe et al., 1992; Keilin, 1959). Tissues with low water content can be found in both animal and plant kingdoms, such as in seeds or pollen of plants, yeast cells, fungal spores, nematodes, rotifers, and embryos of some crustaceans such as brine shrimp (Artemia franciscana) (reviewed in Alpert, 2006; Crowe et al., 1992). In higher plants, orthodox seeds or pollen grains that contain

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low water content can maintain germination ability for years, decades, or even centuries under favorable conditions (reviewed in Hoekstra et al., 2001). The leaves and roots of resurrection plants can be dehydrated to a water content of less than 10% and lose life phenomena; within a few hours after rehydration, dried tissues resume vital metabolism and life cycle (Bartels et al., 1990; Bianchi et al., 1991). Desiccation tolerance in plants is a complex and multifaceted response (Alpert, 2006; Buitink and Leprince, 2004; Clegg, 1986). The response may include the structural or component alteration of the cell wall, organelles, or organs (Farrant, 2000; Farrant et al., 2003; Jones and McQueen‐Mason, 2004; Peeva and Maslenkova, 2004; Tuba et al., 1996; Vicre et al., 2004a,b); induction of the repair system (Potts, 1996; Smith‐Espinoza et al., 2003; Wilson et al., 2004); removal of free radicals (Franc¸a et al., 2007; Leprince et al., 1996; Oliver et al., 2001; Sunkar et al., 2003); and accumulation of macromolecules (Buitink and Leprince, 2004; Collins and Clegg, 2004; Crowe et al., 1992, 1998, 2005; Dure et al., 1989; Hoekstra, 2005; Hoekstra and Golovina, 2002; Murphy et al., 2001; Oliver et al., 2001; Quartacci et al., 2002; Xu et al., 1996). During drying, the accumulation of macromolecules such as oligosaccharides or proteins greatly increases cytoplasmic viscosity and usually causes the formation of bioglasses. Thus, bioglasses have been suggested to provide intracellular protection against the denaturation of large molecules to stabilize plasma membranes (Burke, 1986). Oligosaccharides are usually considered the main component in bioglasses. In nematodes, trehalose increases to 20% dry weight during slow dehydration (Madin and Crowe, 1975). Trehalose also accumulates in yeast and bacterial spores during dehydration and appears to be required for survival in the absence of water (Coutinho et al., 1988; Crowe et al., 1984). In plant systems, sucrose and certain oligosaccharides such as raYnose or stachyose as well as cyclitols instead of trehalose accumulate in large quantities during seed maturation (Hoekstra and Van Roekel 1988; Horbowicz and Obendorf, 1994; Koster and Leopold, 1988). Pollen grains may contain as much as 25% dry weight in the form of sucrose (Hoekstra et al., 1992). Seeds may contain large quantities of nonreducing oligosaccharides depending on the species (e.g.,
17–20% for soybean) (Amuti and Pollard, 1977). The biophysical features of bioglasses are quite diVerent from that of sugar glasses (Wolkers et al., 1998a,b, 1999). That is, other cellular components may play important roles in the formation or functions of bioglasses. The major seed proteins involved in bioglasses formation should be late embryogenesis abundant (LEA) proteins (Blackman et al., 1991, 1992, 1995; Buitink and Leprince, 2004; Oliver et al., 2001; Wolkers et al., 1998a, 1999, 2001).

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Embryogenesis in flowering plants represents a series of stages to develop a miniature plant within the seed. The initial stage is fertilization, and the development of the zygote leads to the formation of a viable embryo, which will undergo subsequent development. The cellular expansion stage is characterized by increased dry mass, and thus provides energy for the processes of germination. In most plant species, the final stage of seed development, maturation, is initiated by a natural reduction in seed water content, which will eventually drop to
10%. During this stage, the gene expression and protein profiles change greatly and are usually associated with achieving desiccation tolerance, increasing abscisic acid (ABA) content, or developing the capacity for seed germination (Goldberg et al., 1989; Hughes and Galau, 1989; Rosenberg and Rinne, 1986; Skriver and Mundy, 1990). These proteins are distinct from storage proteins because of their relatively late accumulation during embryonic development. Moreover, their mRNAs are maintained at a high level in the dehydrated mature embryos, while the transcripts of storage protein genes completely degrade during the last embryogenesis stage (Goldberg et al., 1989). LEA proteins have been suggested to be associated with desiccation tolerance. They are called ‘‘late embryogenesis abundant’’ because they are more abundant during late embryogenesis than during mid‐embryogenesis (Galau et al., 1986). LEA proteins were first described from embryos of mature wheat (Triticum aestivum) and cotton (Gossypium hirsutum) (Cuming and Lane, 1979; Dure et al., 1981). From mature wheat embryos, a dominant protein was found in a cell‐free translation system (Cuming and Lane, 1979). Because this protein was the most conspicuous protein labeled by [35S]methionine during early germination (0–3 h), it was designated the early‐methionine‐labeled (Em) protein. Thereafter, Em mRNAs and corresponding proteins declined rapidly in level (Cuming, 1984; Thompson and Lane, 1980). Transcript analysis of all stages of cotton embryogenesis indicated that LEA mRNAs appeared during late embryogenesis and were maintained at a high level in the dehydrated mature embryos. Some could be induced in dissected immature embryos with exogenous ABA treatment. After germination, these mRNAs and their corresponding proteins declined quickly in level (Dure et al., 1981; Galau and Dure, 1981; Galau et al., 1986). The sequence of Em cDNA indicated high homology with the cotton D‐11 LEA gene (Baker et al., 1988; Litts et al., 1987). Subsequently, the rice (Oryza sativa) Rab21 (Mundy and Chua, 1988), barley (Hordeum vulgare) and maize (Zea mays) dehydrin (Close et al., 1989), carrot (Daucus carota) Dc3 and Dc8 (Choi et al., 1987; Wilde et al., 1988), barley HVA1 (Hong et al., 1988), and

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rape (Brassica napus) LEA76 (Harada et al., 1989) were cloned and sequenced from mature embryos, aleurones, or somatic embryos. These cDNA sequences shared similar sequence motifs and biochemical properties and thus were suggested to share similar functions (Dure et al., 1989). Since then, hundreds of LEA proteins from vascular to nonvascular plants have been isolated (see reviews in Close, 1996, 1997; Cuming, 1999; Dure et al., 1989; Joh et al., 1995; Knight et al., 1995). Interestingly, although LEA genes were first identified from developing seeds, many LEA‐like genes were induced by only ABA or other environmental stresses (see reviews in Bray, 1994; Bray et al., 1993; Cuming, 1999; Ingram and Bartels, 1996; Wang et al., 2003). In addition, various LEA‐like genes have been identified from microbes (Mueller et al., 1992; Vo¨lker et al., 1994), fungi (Abba’ et al., 2006; Eichinger et al. 2005), protozoa (Katinka et al., 2001), bdelloid rotifers (TunnacliVe et al., 2005), nematodes (Browne et al., 2002; Gal et al., 2004), insects (Kikawada et al. 2006), and crustaceans (Hand et al., 2006; Wang et al., 2007). All these animal LEA‐like genes are expressed in dehydrated or anhydrobiosis spores, embryos/fetuses/nymphs, or somatic tissues. These animal LEA‐like transcripts occur only under stress conditions and thus are suggested to be associated with desiccation tolerance. Since the early era of identification of LEA proteins, hypotheses have focused on their protective roles in the acquisition of desiccation tolerance. The physical characteristics of LEA proteins may provide some clues to their mode of action. Typically, the primary structures of many LEA proteins include predominantly charged and uncharged polar amino acid residues but not hydrophobic ones, indicating the LEA proteins are overwhelmingly hydrophilic. Thus, LEA proteins were suggested to provide water‐ or salt‐ binding capacity and association of the subcellular matrix, salt bridges, or ‘‘surface‐solvation’’ moieties (reviewed in Close et al., 1993; Dure, 1993b; Dure et al., 1989). Transgenic/transformation studies also provided direct evidence of a desiccation protection function (e.g., Imai et al., 1996; Kazuoka and Oeda, 1994; Nylander et al., 2001; Xu et al., 1996). The functions of LEA proteins have been gradually revealed after more than two decades, but their detailed functions still remain to be resolved.

II. CLASSIFICATION AND OCCURRENCE OF LEA PROTEINS Early analysis on transcript abundance showed that LEA genes contained two distinct classes, LEA or LEA‐A subgroups, by their temporal pattern of expression during embryogenesis. LEA‐A genes were expressed at a slightly

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earlier stage in seed development than LEA genes (Galau et al., 1986; Hughes and Galau, 1989, 1991). A more common classification of LEA genes was then projected by deduced protein structural domains or chemical characteristics. Currently, two criteria are used. The traditional criterion was initially described by Dure et al. (1989). On the basis of amino acid sequence and conserved motifs, LEA proteins were classified into five to nine subclasses (Bies‐Etheve et al., 2008; Bray, 1994; Cuming, 1999; Dure et al., 1989). The second criterion was provided by a computer analysis, ‘‘Protein or Oligonucleotide Probability Profile’’ (POPP), which showed over‐ or under‐ representation of particular amino acids in protein sequences (Wise, 2002, 2003). After clustering by consensus POPP, LEA proteins were reclassified into at least four groups (Wise, 2003; Wise and TunnacliVe, 2004). Despite the diVerent criteria of classification, the primary structures of most LEA proteins shared similar biophysical features. The Kyte–Doolittle hydropathy algorithm (Kyte and Doolittle, 1982) revealed that most LEA proteins were hydrophilic (Baker et al., 1988; Dure et al., 1989). A. CLASSICAL CLASSIFICATION

1. LEA I proteins This group includes the wheat Em protein (Cuming, 1984; Cuming and Lane, 1979; Litts et al., 1987) and its homolog, the cotton D‐19 protein (Baker et al., 1988). Both proteins contain high sequence homology and similar protein characteristics (Baker et al., 1988; Dure et al., 1989). LEA I proteins isolated from various plant and bacteria species all were rich in Gly (
20%) and charged amino acids (
40%). Early work divided LEA I proteins into two subgroups according to the repeat number of hydrophilic 20‐mer conserved sequences [(R/G)S(R/K)GGQTRKEQLGXEGTXEM] near the carboxyl terminus (Espelund et al., 1995; Gaubier et al., 1993). For example, Arabidopsis had two Em genes: AtEm1, which contained four repeats of 20‐mer motifs, and AtEm6, with one 20‐mer motif (Gaubier et al. 1993). A similar 20‐mer motif (GRKGGEATSKNHDKEFYQEI) was also found in a Bacillus subtilis gsiB gene, which encoded a general stress protein and was induced during sporulation or by glucose or phosphate starvation, limited oxygen, heat, oxidation, and salinity (Mueller et al. 1992; Stacy and Aalen, 1998; Vo¨lker et al., 1994). In a recent proteomic study, LEA I protein was recognized in dehydrated cysts of brine shrimp (Wang et al., 2007). With typical LEA I protein sequences as a query, we identified more than 100 LEA I entries from public databases, including GenBank (http://www.ncbi.nlm.nih. gov) and TIGR Plant Transcript Assemblies database (http://plantta.tigr. org). The list of sequences and basic characteristics are published on our Web

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site: "http://ipmb.sinica.edu.tw/soja/LEA/review_list/. The average molecular weight of these LEA I proteins is 11.5 kDa (maximum 20.3, minimum 6.8). For the net electric charge, 58, 26, and 16% of these proteins have a pI value <6, between 6 and 9, and >9, respectively. For these groups, most proteins are small and acidic to neutral.

2. LEA II proteins The LEA II proteins, also termed D‐11 (Baker et al., 1988), RAB (responsive to ABA) (Mundy and Chua, 1988), or dehydrin proteins (Close et al., 1989), were originally identified from cotton developing embryos. Several cold‐regulated (COR) proteins were also demonstrated to be similar to LEA II proteins (e.g., Gilmour et al., 1992). To date, more than 100 LEA II proteins have been identified in a range of plants of widely divergent taxa, including angiosperms, gymnosperms, pteridophytes, bryophytes, fungi, algae, and cyanobacteria (Abba’et al., 2006; Campbell and Close 1997; Close, 1997; Dure et al., 1989; Eichinger et al., 2005; Rorat, 2006; Saavedra et al., 2006; Velten and Oliver, 2001; Wisniewski et al., 1996). All LEA II entries are listed at the Web site: http://ipmb.sinica.edu.tw/soja/LEA/review_list/. LEA II proteins had three specific, characteristic sequence ‘‘modules’’ that defined this class of protein (Close, 1997; Close et al., 1993). First, all the LEA II proteins contained one to multiple 15‐mer lysine‐rich conserved sequences (EKKGIMDKIKEKLPG) (Galau and Close, 1992), designated as the ‘‘K‐domain,’’ near the C‐terminus. The second motif, located at the N‐terminus, was designated the ‘‘Y‐domain’’ with the consensus sequence of (V/T)DEYGNP, which had similar amino acid sequence to the nucleotide binding domain found in chaperones of plants and bacteria (Martin et al., 1993). The third feature, termed the ‘‘S‐segment,’’ was a stretch of contiguous Ser or Thr residues acting as a site of protein phosphorylation (Vilardell et al., 1990). The S‐segment was suggested to be phosphorylated by CK2 or other Ser/Thr protein kinases (Plana et al., 1991; Riera et al., 2004) and to be associated with a signal for nucleus and nucleolus localization (Goday et al., 1994; Riera et al., 2004). Moreover, these motifs were embedded in amorphous regions with Gly/Ser‐, Gly/Thr‐, or Glu/Lys‐rich residues designated as ‘‘‐domains’’ (Close, 1997; Close et al., 1993; Dure, 1993b). According to the presence, order and number of the Y‐, S‐, and K‐domains, LEA II proteins had been classified into five subgroups. These subgroups included the proteins ‘‘YnSK2’’ (where n is the number of repeat), ‘‘Kn,’’ ‘‘KnS,’’ ‘‘SKn,’’ and ‘‘YnKn,’’ respectively (Close, 1996, 1997). The molecular weight of the LEA II proteins is from 5.3 to 66.3 kDa (average 18.3 kDa). The wide range of molecular weight in LEA II proteins is caused by the repeat number of K‐segments. For the net electric charge, 7, 58, and 35% of

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these proteins have a pI value <6, between 6 and 9, and >9, respectively, which suggests that most LEA II proteins are neutral to basic. 3. LEA III proteins Like LEA I proteins, LEA III proteins, also called D‐7 proteins, are found across plant, fungi, microbial, and animal kingdoms (see review in Cuming, 1999; Dure et al., 1989; TunnacliVe and Wise, 2007). To date, more than 100 entries can be identified in public databases (see list at http://ipmb.sinica. edu.tw/soja/LEA/review_list/). For net electric charge, 20, 37, and 43% of these proteins have a pI value <6, between 6 and 9, and >9, respectively, which suggests that most LEA III proteins are neutral to basic. The particular feature found in these LEA III proteins is the tandem‐repeating 11‐mer sequence. Some of them contain the subsequent duplication and divergence of the 22‐mer sequence, which gives rise to additional copies of the 11‐mer sequence. A large variation in molecular weight among LEA III proteins is caused by the number of tandem repeats, which range from 5 to more than 30. The average molecular weight of the LEA III proteins is 25.5 kDa (maximum 67.2, minimum 7.2). These 11‐mer motifs had been proposed to be linked by ionic bridges and played important physiological roles (Dure et al., 1989). The LEA III proteins, comprising subgroups D‐7, Dc8, GmPM8, and D‐29, are characterized by the diVerent consensus sequences of 11‐mer repeats, such as TAGAAKEKAXE, KAKETKDAAAE, TKDYAGDAAQK, and (E/Q)XK(E/Q)KX(E/D/Q) (where  presents a hydrophobic residue) for D‐7, Dc8, GmPM8, and D‐29 subgroups, respectively (Baker et al., 1988; Curry and Walker‐Simmons, 1993; Dure et al., 1989; Hsing et al., 1995). Although 11‐mer repeats of microbial or animal LEA III proteins may not be in tandem, their consensus sequences still fit with the above subgroups. For example, PvLEA1, PvLEA2, and PvLEA3 from Polypedilum vanderplanki (Kikawada et al., 2006) belonged to the GmPM8, Dc8, and D‐29 subgroups, respectively, whereas AfrLEA1 and AfrLEA2 from Artemia franciscana (Hand et al., 2006; Wang et al., 2007) belong to the Dc8 and D‐29 subgroups, respectively. 4. LEA IV proteins This group, also called D‐113 proteins, was isolated from several species, including cotton D‐113 (Baker et al., 1988), tomato and its wild relative Le25 (Cohen et al., 1991; Kahn et al., 1993), sunflower (Helianthus annuus) Ha ds11 (Almoguera and Jordano, 1992), soybean GmPM1 and GmPM16 (Chen et al., 1992; Shih et al., 2004), Arabidopsis AtLEA4-1 (PAP260) and AtLEA4-5 (PAP51) (Parcy et al., 1994), and Phaseolus vulgaris PvLEA‐18

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(Colmenero‐Flores et al., 1999). To date, more than 100 entries have been identified from widely divergent taxa in the plant kingdom, including angiosperms, gymnosperms, pteridophytes, and bryophytes (see list at http://ipmb. sinica.edu.tw/soja/LEA/review_list/). The average molecular weight of the LEA IV proteins is 12.6 kDa, with the maximum of 18.8 and minimum of 8.4. LEA IV proteins lack consensus motifs or sequences like LEA I, LEA II, or LEA III proteins. Generally, the N‐terminal domains contain several conserved small and charged amino acid residues, whereas the variable C‐terminal regions include a large amount of nonpolar amino acid residues. LEA IV proteins contain high proportions of small amino acid residues such as Gly or Ala and charged ones such as Asp, Gly, Lys, or Arg and are thus highly hydrophilic (Dure, 1993b). For the net electric charge, 22, 22, and 56% of these proteins have a pI value <6, between 6 and 9, and >9, respectively, which suggests that most LEA IV proteins are basic. Like LEA I, II, III proteins, LEA IV proteins are heat soluble, that is, they do not precipitate after boiling for 10 min. On the basis of sequence alignments, this group contains at least four diVerent subgroups: D113, GmPM16, PvLEA‐18, and GmPM28.

5. LEA V Cotton LEA D‐34 and D‐95 proteins (Baker et al., 1988; Galau et al., 1993) are representatives of the group V LEA proteins. Several homologues from cotton or other plant species have been recognized, including maize RAB28 (Pla et al., 1991), carrot ECP31 (Kiyosue et al., 1992), cotton D‐95 and D‐73 (Galau et al., 1993), Arabidopsis AtECP31 and AtRAB28 (Arenas‐Mena et al., 1999; Yang et al., 1996), Craterostigma plantagineum pcC27‐45 (Piatkowski et al., 1990), tomato ER5 (Zegzouti et al., 1997), hot pepper (Capsicum frutescens) CaLEA6 (Kim et al., 2005), Medicago truncatula MtPM25 (Boudet et al., 2006), and soybean GmPM22, GmPM24, GmPM25, and GmPM26. Currently, about 100 entries have been identified (see list at http://ipmb.sinica.edu.tw/soja/LEA/review_list/). The average molecular weight of the LEA V proteins is 18.1 kDa (maximum 38.5, minimum 5.3). For the net electric charge, 87, 6, and 7% of these proteins have a pI value <6, between 6 and 9, and >9, respectively, which suggests that most LEA V proteins are acidic. The members of this group are distinctively diVerent from the other four groups of LEA proteins by their containing a high proportion of hydrophophic residues and lacking the feature of boiling solubility. Hence, LEA I, II, III, and IV proteins are so‐called hydrophilic LEA proteins, whereas LEA V proteins are hydrophobic.

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6. Atypical LEA proteins Atypical LEA proteins, as with other groups of LEA proteins, accumulate in seeds during late embryogenesis and disappear after the early hours of germination. They may also accumulate in other organs during various environmental stresses. This group does not contain any motif or protein characteristics defined in other LEA proteins; examples are barley HVA22 (Shen et al., 1993); rice WSI76 (Takahashi et al., 1994); Arabidopsis AtDI19, AtLEA9-1 (M10) and AtLEA9-2 (M17) (Gosti et al., 1995; Raynal et al., 1999); wheat AWPM‐19 and WCOR413 (Breton et al., 2003; Koike et al., 1997); and soybean GmPM4 (Hsing et al., 1998). The expression pattern suggests that deduced products of these genes are associated with the development of late embryogenesis. However, unlike typical hydrophilic LEA proteins, they may be carrier proteins, galactinol synthases, fiber proteins, or plasma membrane proteins. Thus, the members of this group fit with the primary definition of LEA proteins but have diverse protein characteristics and functions. B. POPP CLASSIFICATION

POPP is a suite of interrelated software tools that allow for discovering what is statistically ‘‘unusual’’ in the composition of an unknown protein or for clustering proteins into families based on peptide composition (Wise, 2002). The basic principle of POPP is to amend the defect of local‐alignment sequence‐comparison tools. For example, proteins or protein domains with similar function are likely to contain the same mono‐, di‐, or tripeptides over‐ or under‐represented. However, several algorithms such as BLAST typically score matches over longer oligopeptides but introduce the disadvantage of noncontiguous matches; thus, they might miss similarities (Wise and TunnacliVe, 2004). By POPP, LEA proteins are reclassified into at least four groups (Wise, 2003; Wise and TunnacliVe, 2004) as follows: Group 1: This group includes the traditional LEA I proteins and is also split into two subgroups. Although the criterion of subgroups is based on diVerent consensus POPPs, it is the same as the traditional criterion for LEA I proteins. However, the two subgroups belong to two diVerent superfamilies (SFs). Group 1a proteins (SF4) share POPP vectors with histone H4 and with other chromosomal and nuclear proteins. The predicted function of group 1a proteins is DNA binding nuclear protein. Group 1b proteins (SF6), by contrast, match a group of proteins with broader functions, such as RNA or ATP binding and gyrase or chaperone activities. Group 2: In addition to the traditional LEA II proteins, some LEA IV proteins are included in this group. The consensus POPPs suggest that this

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group comprises two subgroups, 2a and 2b: Group 2a proteins include all the group 2 proteins not associated with cold tolerance and all group 2 proteins that are accumulated in late embryogenesis; group 2b proteins include those associated with cold tolerance, together with a few that are not produced during late embryogenesis. Group 2a proteins belong to two diVerent superfamilies. SF1 is the cluster of DNA unwinding‐ or repair‐related proteins, whereas SF10 proteins share POPP vectors with DNA binding proteins and cytoskeleton proteins. Group 2b proteins (SF3) share the cluster of DNA unwinding‐ or repair‐related proteins, as well as cytoskeleton, Ca2þ binding, and chaperone proteins. Group 3: The new group 3 includes the traditional LEA III and the remainder of the LEA IV proteins. Subgroups, corresponding to groups 3a (SF2) and 3b (SF5), can be identified by their POPP consensus peptides. The key words and phrases associated with SF2 proteins are chaperone, coiled coil, tropomyosin, phosphorylation, stress, filament, cytoskeleton, neurofilament, actin binding, and rotamase. By contrast, SF5 proteins contain nuclear and DNA binding proteins, as well as histones, mainly H1 histones. Group 4: This group includes LEA V proteins and corresponds to SF7. Group 4 proteins share characteristics with a cluster of chaperone and chromatin‐associated nuclear proteins. To compare the traditional and POPP classifications, LEA I and LEA V proteins remain distinct, as unique groups. The classical LEA IV proteins are reclassified into group 2 or group 3 by their individual motifs. If the number of mismatches allowing for the 11‐mer motif in group 3 proteins is increased by 1 to 5, the set of matching LEA proteins includes several group 2 proteins, such as barley DHN1 (LEA II) and DHN2 (LEA II), rice Rab16D (LEA II), soybean GmPM1 (LEA IV), and tomato Le25 (LEA IV). In addition, the group 3 protein C. plantagineum pcC30‐6 (LEA III) has a poly‐lysine shutter, whereas another two group 3 proteins, wheat WCS19 (LEA III) and Wcor14a (LEA III), have poly‐serine shutters (i.e., the 2S motif). Thus, some group 2 and group 3 proteins may be related. C. THE ANNOTATION OF ARABIDOPSIS LEA GENES

Before the era of genome sequencing, LEA gene cloning usually depended on the transcript pools from tissues of interest. In general, this strategy allows for identifying one to several genes of interest but cannot be used to establish an overall inventory. The genome sequencing project provides a powerful tool to undertake an exhaustive analysis of all LEA gene families in one species. By using a custom bioinformatic pipeline, one can search for genes similar to those previously described as LEA. Hence, Bies‐Etheve et al. (2008)

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analyzed the A. thaliana genome sequence by using several known LEA genes as reference sequences and revealed 50 LEA genes in its genome. Combining sequence‐similarity and conserved‐domains analysis, these 50 LEA genes were divided into nine diVerent groups: Groups 1–4: correspond to the classical LEA classification. The Arabidopsis genome contains two genes in group 1, nine in group 2, six in group 3, and five in group 4. Group 5: contains six genes encoding D‐34 type (LEA V) proteins. Group 6: contains four genes related to cotton D‐73 (Galau et al., 1993) and Arabidopsis AtDi21 (Gosti et al., 1995). Group 7: contains three genes for the D‐95 type (LEA V) proteins. Group 8: contains three genes similar to P. vulgaris PvLEA‐18 (Colmenero‐ Flores et al., 1999). This gene belongs to the LEA IV group in the classical classification. Group 9: contains two genes, that is, the Arabidopsis M10 and M17. These two genes are assigned to the atypical LEA group in the classical classification.

III. THE STRUCTURE OF LEA PROTEINS Because of the importance of LEA proteins in desiccation tolerance, many studies endeavor to determine the structures of LEA proteins. The secondary structures of several LEA proteins have been characterized by circular dichroism (CD), nuclear magnetic resonance (NMR), or Fourier transform IR (FTIR) spectroscopy. In addition, the three‐dimensional structures of D‐95 (LEA V) proteins have been determined. A. LEA PROTEINS IN SOLUTION STATUS

Identification of the fine structures of LEA proteins is a main goal for functional studies. Most of the algorithms (e.g., Chou and Fasman, 1978; Rost and Liu, 2003; Rost and Sander, 1993) predicted that the LEA I to LEA IV proteins should adopt large amounts of defined secondary structures, such as ‐helix or ‐sheet. For example, the 20‐mer motif for LEA I, the K‐segment for LEA II, 11‐mer repeats for LEA III, or the hydrophilic domain for LEA IV was predicted to form amphiphilic ‐helical structures (Dure, 1993a; Imai et al., 1996). After spectroscopy analysis, however, more and more hydrophilic LEA proteins were found to contain disordered structures in solution state.

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1. LEA I proteins Even before the first LEA gene was cloned and sequenced, the purified Em proteins from wheat embryos were used for studying the biophysical characteristics. The far‐UV CD spectrum revealed that Em proteins in KCl‐ HEPES buVer adopts mainly, about 70%, random coiled and little ‐helical conformations. Gel filtration and hydrodynamic experiments showed that Em proteins were globular proteins. The addition of Ca2þ, Mg2þ, or Mg2þ‐ ATP failed to change the shape of CD spectra. Em proteins still contain over 50% random coiled conformations in the presence of trifluoroethanol (TFE), a common ‐helical‐promoting cosolvent. In addition, the 20‐mer motif also adopted a random‐coil conformation. These results suggested that Em proteins lack defined secondary structures (Gilles et al., 2007; McCubbin et al., 1985). The pea (Pisum sativum) P11 proteins were purified from pea seed axis and characterized by CD and NMR spectroscopy. Spectra from both techniques indicated that P11 proteins lacked ordered conformations and had high thermostability (Russouw et al., 1995, 1997). The carrot EMB‐ 1 proteins were characterized by 1H NMR spectroscopy under both aqueous conditions and conditions simulating desiccation, suggesting that EMB‐1 protein had no defined secondary or tertiary structure in solution and the protein backbones were highly flexible on a sub‐nanosecond time scale (Eom et al., 1996). Far‐UV CD spectra of various temperatures indicated that the soybean GmD‐19 proteins were highly disordered in solution state and underwent the temperature‐induced unfolding process. CD diVerence spectra indicated that the proteins contained a few extended helicals known as poly (L‐proline)‐type II (PPII) conformations, which usually appeared in disordered polypeptides at low temperature (Arnott and Dover, 1968; Gokce et al., 2005; Sreerama and Woody, 1994, 1999; Woody, 1992). In the presence of TFE or sodium dodecylsulfate (SDS), both being ‐helix promoting agents, the spectra indicated that the proteins adopted restricted ‐helical conformations. Although SDS micelles might induce the conformational change, incubation of GmD‐19 proteins with phospholipids had no eVect on protein structures (Soulages et al., 2002). Finally, FTIR analysis showed that M. truncatula MtEm6 proteins adopted 47% defined secondary structures in the fully hydrated condition (Boudet et al., 2006).

2. LEA II proteins CD analysis of the purified G50 dehydrins from maize kernels showed that the protein comprised
75% disordered structures (Ceccardi et al., 1994). The far‐UV spectrum of recombinant C. plantagineum DSP16 proteins revealed a large amount of disordered conformations with partial helical

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structures. 1H NMR spectra results also suggested that the proteins were essentially unstructured. In the presence of high concentration of TFE, the far‐UV CD spectra of DSP16 proteins gave the typical features of ‐helix. Moreover, in the presence of 7 M guanidinium chloride, the proteins underwent denaturant‐dependent structural transitions, while also showed that the proteins contained typical PPII helical features in natural condition (Lisse et al., 1996). Far‐UV CD spectroscopy revealed the protein conformations of the purified DHN from cowpea with or without the addition of SDS. In the absence of SDS, the proteins lacked well‐defined secondary structures, but in the presence of SDS, the proteins adopted ‐helical conformations (Ismail et al., 1999b). A similar strategy was used for studies of CuCOR19, a citrus cold‐responsive dehydrin. The CD spectra of CuCOR19 proteins showed that the major secondary structures were random coiled in the solution state but ‐helical in the presence of SDS (Hara et al., 2001). The CD spectra of purified maize DHN1 proteins revealed the characteristics of random coils. Incubation of DHN1 with phosphatidic acid‐derived small unilamellar vesicles shifted the structures to ‐helical conformations, as compared to findings with SDS (Koag et al., 2003). The structural property of soybean GmDHN1 proteins also indicated a large amount of disordered structures in solution state. Far‐UV spectra and their diVerence spectrum on temperature scans revealed that GmDHN1 proteins underwent the temperature‐induced unfolding processes and contained a few PPII helices at low temperature. The addition of TFE or SDS induced the proteins to increase ‐helical structures. CD spectra of GmDHN1 proteins incubated with multilamellar liposomes of dimyristoyl phosphatidylcholine or dimyristoyl phosphatidylglycerol revealed no diVerence in structures as compared to treatment with GmDHN1 proteins alone (Soulages et al., 2003). Finally, the full‐length sequences or conserved K‐segments of four typical Arabidopsis dehydrins, COR47, LTI29, LTI30, and RAB18, were analyzed by CD spectroscopy. The spectra of these proteins suggested that full‐length dehydrins adopted overall disordered conformations, whereas the K‐segments adopted significant PPII helices at low temperature. In the presence of 90% TFE, the extent of the ‐helical structures of these four dehydrins was not the same. After estimation, the maximal helix content ranged from 2 to 50%. Although K‐segments were predicted to adopt ‐helical structures by predicting algorithms, the helical propensities for the K‐segments of these dehydrins were relatively weak and substantially lower than that of the full‐length dehydrins. These dehydrins also showed no conformational change in the presence of salts or increase of protein concentration (Mouillon et al., 2006).

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3. LEA III proteins The results of FTIR analysis for D‐7 LEA proteins from Typha latifolia pollen revealed completely disordered structures (i.e.,
25% turn structures and 75% random‐coiled conformations) in the hydrated state. Aphelenchus avenae LEA proteins, AavLEA1 (Browne et al., 2002), were analyzed by CD and FTIR spectroscopy. Far‐UV CD spectra revealed that AavLEA1 contained disordered structures and a temperature‐induced unfolded process in solution. The spectra of temperature scans suggested the existence of PPII helices at low temperature (Goyal et al., 2003). CD and FTIR spectroscopy showed that the pea seed mitochondrial LEA proteins, LEAM (also known as PsLEAm), were an intrinsically disordered protein. In the presence of TFE or SDS, the spectra of LEAM showed typical ‐helices (Grelet et al., 2005; Tolleter et al., 2007). 4. LEA IV proteins Both far‐UV CD and FTIR spectroscopy indicated that soybean GmPM16 proteins adopted highly disordered conformations in solution. In addition, GmPM16 showed no conformational change in the presence of oligosaccharides such as sucrose, raYnose, stachyose, or trehalose, or in solutions of pH 4.8, 7.0, or 9.4. The addition of SDS or TFE promoted the formation of ‐helical structures in the GmPM16 proteins (Shih et al., 2004). Furthermore, two other soybean proteins, the basic GmPM1 and the acidic GmPM28, also adopted huge amounts of disordered conformation and had no interaction with oligosaccharides in the fully hydrated state. CD diVerence spectra indicated that the proteins contained a few extended helical, or PPII, conformations at low temperatures. Both proteins had less intrinsic ability to adopt ‐helical structures than GmPM16 proteins (Shih et al., unpublished data). B. LEA PROTEINS IN DRY STATUS

Because of the accumulation of LEA proteins during late embryogenesis with its extremely low water content, the structure of these proteins in the dry state should be useful information for protein functions. T. latifolia D‐7 LEA protein was the first protein used to measure the protein secondary structures in the dehydrated state. The results of FTIR spectra suggested that the proteins adopted defined secondary structures, such as ‐helix and ‐sheet. The proportions of defined secondary structures depended on the rate of drying, fast or slow (Wolkers et al., 2001). The temperature‐scanning FTIR spectra of GmPM1 proteins illustrate the real‐time conformational changes during water loss, and are shown in Fig. 1. The decreasing density in amide II

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Fig. 1. Temperature‐scanning FTIR spectra of GmPM1 proteins. Each shade represents a diVerent temperature, from 10 to 104 8C. (Panel A) The shift of maximal absorption peaks in the amide I (1700–1600 cm1), amide II (1580– 1520 cm1), and amide III (1470–1430 cm1) regions. (Panels B and C) Details of maximal absorption peaks in amide I and II regions.

and the increasing one in amide III suggest the reduction of hydrogen bonds between water molecules and protein side chains. The peak shifts that appear at amide I and amide II suggest an increase in ‐helix/random coil ratios during temperature rise. The FTIR spectra of other LEA proteins, such as A. avenae AavLEA1, soybean GmPM16, M. truncatula MtEm6, and pea LEAM, also revealed similar conformational changes in dehydrated status (Boudet et al., 2006; Goyal et al., 2003; Shih et al., 2004; Tolleter et al., 2007). In addition, in the presence of oligosaccharides, abundant in dry pollen

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grains or seeds, T. latifolia D‐7 or soybean GmPM16 adopted mainly ‐ helical structures and might prevent the formation of intermolecular extended ‐sheets. The dehydrated mixture of sucrose and the LEA protein showed a higher glass transition temperature (Tg) and lower wave number‐ temperature coeYcient (WTC) than the dehydrated oligosaccharide alone (Shih et al., 2004; Wolkers et al., 2001). Recently, two novel LEA III proteins with unusual 11‐mer repeats from bdelloid rotifers (Adineta ricciae), ArLEA1 and ArLEA2, were reported. Both protein sequences were very similar; ArLEA1A was longer by 44 amino acid residues which corresponded to 10th–13th 11‐mer sequences. However, the far‐UV CD spectroscopy of ArLEA1A and ArLEA1B proteins in solution and dry state suggested diverse structural features. ArLEA1A proteins adopted disordered structures and performed dehydrated conformational changes, while the shorter ArLEA1B proteins contained large amounts of defined secondary structures under both conditions. Moreover, ArLEA1B had a stronger propensity to interact with dry phospholipid membranes than ArLEA1A and AavLEA1 (Pouchkina‐Stantcheva et al., 2007). TunnacliVe and colleagues (Goyal et al., 2003; TunnacliVe and Wise, 2007; Wise and TunnacliVe, 2004) pointed out that interaction between LEA proteins and oligosaccharides was not unique because other non‐LEA proteins, such as poly‐L‐lysine or BSA protein, were able to interact with oligosaccharides (Crowe et al., 1997; Wolkers et al., 1998c). LEA proteins and other similar proteins that carried out protection functions might share similar mechanisms. Besides, all FTIR spectra of studied LEA proteins showed typical ‐helical and ‐sheet conformations (Goyal et al., 2003; Tolleter et al., 2007; Wolkers et al., 2001). The maximum bands of these spectra appeared around 1654 cm1, whereas the maximum bands of typical coiled‐coil or filament proteins appeared around 1643 cm1 in the amide I region (Heimburg et al., 1996, 1999). The band shift in the amide I region is a critical signature to distinguish the typical ‐helical structure and typical coiled‐coil conformation, suggesting that LEA III proteins might not form coiled coil conformation. The studied LEA III proteins adopted diverse proportions of secondary structures, although all contained 11‐mer repeats. Thus, LEA proteins may follow individual but unknown processes to fold up three‐dimensional structures in the dry state. C. HYDROPHOBIC LEA PROTEINS

In typical LEA proteins, the LEA V group is unique among other hydrophilic LEA groups because of its hydrophobicity and heat instability. Thus, LEA V proteins may have the advantage to determine fine structures. In fact, the

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first, and currently the only three‐dimensional structures of LEA proteins, At1g01470.1, an Arabidopsis D‐95 type LEA protein, had been reported (Singh et al., 2005). The structures of At1g01470.1 in aqueous phase, which were determined by NMR spectroscopy, revealed an ‐fold consisting of one ‐helix and seven ‐strands that formed two antiparallel ‐sheets. This ‐fold occupied
50% of the protein conformations. The homologous structures of two antiparallel ‐sheets were similar to a fibronectin type III (Fn3) domain (Brummendorf and Rathjen, 1996). The closest structure homologs to At1g01470.1 were the barrel 1 domain of human blood coagulation factor XIII zymogen (Kraulis, 1991; Yee et al., 1994) and bacteria apaG protein (Cicero et al., 2007). However, the structures of these homologs lacked an ‐helix in the C‐terminus. In addition, both proteins might form a hetero‐tetramer or homo‐dimer, but no evidence suggests that At1g01470.1 could form a polymer. On the other hand, FTIR analysis showed that the M. truncatula D‐34 type LEA protein, MtPM25, adopted 51% defined secondary structures in solution status. Interestingly, MtPM25 proteins also adopted dehydration‐induced conformation changes. There were 65 and 80% defined secondary structures after fast and slow drying, respectively (Boudet et al., 2006). D. HYDROPHILIC LEA PROTEINS ARE MEMBERS OF NATIVELY UNFOLDED PROTEINS

In biophysical strategy, the structure–function paradigm is a widely used formula, that is, Primary amino‐acid sequence ! Three dimensional structure ! Function Three‐dimensional structure is taken to be a prerequisite for protein function, and native protein structure equates with ordered three‐dimensional structure. In the late 1980s, several cases suggested that proteins lacking a fixed structure do not necessarily lack function. Afterwards, proteins with largely disordered conformations, now called ‘‘natively unfolded proteins’’ or ‘‘intrinsically disordered proteins,’’ became a new frontier of protein function study (Dunker et al., 2001; Schweers et al., 1994; Uversky et al., 2000). Natively unfolded proteins behaved almost like random coils and should be extremely flexible, essentially noncompact (extended), and had little or no ordered secondary structure under physiological conditions (Uversky, 2002a,b). Proteins with large disordered conformations were not a rare phenomenon. A genome‐wide prediction indicated that the disordered proteins were common in eukaryote genomes. In the Arabidopsis or Drosphila geneome, for example, 29 or 41% of proteins were predicted to be

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partially, and 8 or 17% to be wholly, disordered, respectively (Dunker et al., 2001). After comparing the disordered regions of a large number of proteins, Dunker and colleagues pointed out that the disordered regions shared some common sequence features. They consisted of low sequence complexity, with amino‐acid compositional bias, essentially noncompact (extended), and high flexibility. Furthermore, most of the intrinsically disordered proteins, being mainly depleted in Ile, Leu, Val, Trp, Phe, Try, Cys, and Asn residues, were enriched in Glu, Lys, Arg, Gly, Gln, Ser, Pro, and Ala residues (reviewed in Dunker et al., 2001; Uversky, 2002a,b). Although natively unfolded proteins lacked defined secondary structures, they possessed various important functions such as regulation of transcription and cell cycle control (Dunker et al., 2001; Wright and Dyson, 1999). It had been suggested that some natively unfolded proteins were correlated with several neurodegenerative disorders, such as Alzheimer’s disease, Down’s syndrome, and Parkinson’s disease (reviewed in Uversky, 2002a). For example, the possible pathogenic factor of Parkinson’s disease, ‐synuclein, was a natively unfolded protein and could interact with elevated levels of several metals, such as iron, mercury, zinc, or aluminum, which were risk factors for Parkinson’s disease, to adopt a large amount of ordered conformations (Uversky et al., 2001). Thus, the natively unfolded proteins lacking rigid structures under physiological condition might represent a functional advantage for their large plasticity allowing them to interact eYciently with several diVerent targets (Wright and Dyson, 1999). Because the primary protein sequences associated tendency of protein folding, analysis of biochemical and biophysical features for 20 amino acid residues is able to predict the protein folding status. Uversky et al. (2000) suggested that the mean hydropathicity (hHim) calculated by Kyte–Doolittle hydropathy for unfolded proteins should be smaller than their boundary hydropathicity (hHib) calculated by the equation hHib ¼ (hRiþ1.151)/2.785, where hRi is the mean net charge. Garbuzynskiy et al. (2004) provided the average number of contacts and an optimal set of artificial parameters for 20 amino acid residues. After calculation of 90 natively unfolded proteins and 80 ideally folded proteins, the mean average number of contacts per amino acid residue was 20.05  0.58 for unfolded proteins and 20.91  0.25 for folded proteins, while the mean artificial parameter per amino acid residue was 3.0  1.2 for unfolded proteins and 0.6  0.5 for folded proteins. Table I shows the amino acid composition and three protein folding parameters of several typical LEA proteins. The amino acid composition of LEA proteins reveals that hydrophilic LEA proteins (LEA I–IV) are rich in small and hydrophilic amino acid residues and poor in nonpolar ones. By contrast, LEA V proteins and a small heat shock protein Hsp22, which contain defined

TABLE I Comparison of Amino Acid Content (mol%) and Calculated Structural Parameters for Several LEA Proteins Small1 Group I

II

III

IV

V Hsp22

Species

Accession

Arabidopsis Arabidopsis Soybean Rice Rice Soybean Soybean Rice Cotton Cotton Soybean Carrot Cotton Soybean Soybean Soybean Cotton Cotton Soybean

Z11157 Z11158 AF004805 AK287564 X57327 AF004807 M94012 D26538 X13203 X13201 Z22872 X16131 M19406 M80666 AF004810 AF117724 M19389 M88322 X63198

Hydroxyl2

Nonpolar3

Hydrophilic4

mol% 23.9 25.7 21.0 21.5 22.4 23.5 27.0 16.3 14.5 27.9 25.3 26.7 29.1 30.6 20.3 18.0 28.0 11.3 7.8

13.0 7.9 11.4 9.7 14.0 18.7 17.7 7.6 19.3 14.0 15.7 9.4 13.9 19.1 9.8 22.5 11.0 14.6 6.8

10.9 12.5 11.4 16.1 10.1 13.3 7.1 10.9 13.8 8.1 7.0 11.5 8.5 9.8 14.3 13.5 21.6 29.8 21.9

A small heat shock protein Hsp22 is used as an out‐group control. Boundary line for folded and unfolded proteins is i 20.91  0.25/20.05  0.58, ii 0.6  0.5/3.0  1.2 (Garbuzynskiy et al., 2004). 1 Gly þ Ala, 2 Ser þ Thr, 3 Ile þ Leu þ Met þ Val, 4 amide þ acid þ basic þ hydroxyl, 5 ideal protein‐folding parameters.

pI 59.8 57.9 64.8 59.1 56.1 55.4 55.8 72.8 62.8 61.0 60.0 56.4 53.3 53.2 54.1 59.6 44.7 44.4 45.3

7.7 6.0 5.3 9.8 11.1 9.8 6.6 7.3 4.9 7.8 7.7 5.6 5.7 10.2 9.8 4.4 4.6 4.6 7.7

Mean/boundary hydropathicity

Number of i contacts

Artificialii,5 parameters

0.3131/0.4133 0.3170/0.4298 0.3015/0.4201 0.3311/0.4210 0.3338/0.4432 0.3730/0.4198 0.3365/0.4244 0.1860/0.4211 0.3620/0.4306 0.3361/0.4133 0.3555/0.4133 0.3701/0.4172 0.3434/0.4220 0.3723/0.4257 0.3555/0.4241 0.3519/0.4375 0.4620/0.4255 0.4731/0.4276 0.4426/0.4095

19.3725 19.4684 19.3255 19.5777 19.6050 19.6895 19.6055 19.0348 20.0537 19.4250 19.6130 19.5957 19.6168 19.5421 19.9875 19.7509 20.1508 20.8626 20.7919

4.2748 4.3897 4.0448 4.3844 4.7411 4.0009 4.5101 5.6832 2.7349 4.4222 3.7786 3.5772 4.1625 4.3949 3.2315 3.5679 1.9736 0.5756 0.9308

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secondary structures, have more nonpolar residues. The protein‐folding parameters also suggest that all listed hydrophilic LEA proteins fit with unfolded ones. For LEA V proteins, the protein folding parameters of M88322, a cotton D‐95, fit with folded proteins. However, the mean average number of contacts of M19389, a cotton D‐34, locates in the region of an unfolded boundary, which may be caused by extremely high contents of small amino acid residues. Other parameters still suggest that D‐34 protein should be folded. Thus, these parameters may be used to find out most LEA genes in a genome‐wide prediction analysis. The consensus POPP analysis of various LEA proteins also indicated similar profiles of over‐ and under‐represented peptides (Wise, 2002, 2003). And, spectroscopy analyses provide direct evidence to define hydrophilic LEA proteins as natively unfolded proteins. Most folded proteins contain a hydrophobic core with the side chain stabilizing the folded conformation and charged or polar side chains on the surface where they interact with surrounding water molecules. By contrast, natively unfolded proteins perform mainly the water molecule–protein interaction in lieu of intramolecularly hydrophobic interaction. Hence, they fail to obtain defined structures. Moreover, unfolded LEA proteins may carry additional water molecules. Hydrodynamic data suggested that the molecular volumes of wheat Em (LEA I) and nematode AavLEA1 (LEA III) were significantly larger than those of a typical globular protein of equivalent size, which indicated a high level of associated water. For example, the nematode AavLEA1 had
20‐fold more associated water than a typical globular protein (Goyal et al., 2003; McCubbin et al., 1985). Natively unfolded proteins may change their conformation by interacting with other molecules such as proteins, nuclear acids, or metal ions. Although hydrophilic LEA proteins interact with SDS micelles and perform the protein‐folding process, no evidence exists to show that hydrophilic LEA proteins could interact with sugars, metal ions, or even phospholipids in solution. Gel electrophoresis results suggested that nematode AavLEA1 and soybean GmPM8 might oligomerize in vitro (Goyal et al., 2003; Hsing, unpublished data). Again, no further evidence showed that AavLEA1, GmPM8, or other related LEA III proteins would adopt quaternary structures in vivo. However, LEA proteins did perform conformational changes during dehydration, because superficial groups of proteins no longer bound hydrogen with water and instead formed inter‐ and intramolecular hydrogen bond with one another (Prestrelski et al., 1993). With dehydration, the biochemical features of LEA proteins changed. They started to interact with various oligosaccharides, although no interaction occur in solution. By comparison with pure oligosaccharide glasses, the presence of LEA

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proteins increased both the Tg and the average strength of hydrogen bonding. This suggested that the proteins acted synergistically with the oligosaccharide in the formation of the glassy matrix (Shih et al., 2004; Wolkers et al., 2001). Although both T. latifolia D‐7 and nematode AavLEA1 proteins belonged to the D‐7‐type subgroup and contained five 11‐mer repeats, they formed a diverse proportion of secondary structures after dehydration (Goyal et al., 2003; Wolkers et al., 2001), perhaps because they lacked an identical folding mechanism. In addition, the folding kinetics of disordered LEA proteins should be very diVerent to that of other folded proteins. Thus, the folding mechanism of LEA proteins or other natively unfolded proteins in specific conditions might be a new area for protein kinetics.

IV. THE PHYSIOLOGICAL ROLES OF LEA PROTEINS In general, orthodox seeds undergo continuous dehydration during maturation and maintain low water content. All orthodox seeds can withstand dehydration to
5% [0.053 g H2O g1 dry weight (gg1)] of water content, even when maturation drying is not completed prior to shedding (Berjak and Pammenter, 2002). Thus, considering their redundancy and high amounts, LEA proteins have been suggested as one of the key factors in the acquisition of desiccation tolerance. Because LEA proteins disappear quickly during seed germination, their constituent amino acid residues may be utilized while the seed storage proteins are at an early stage of degradation. Thus, LEA proteins may also play an atypical storage role during germination. Nevertheless, this role should be viewed as a natural consequence of the recycling system. A. THE ACCUMULATION OF LEA PROTEINS DURING DEHYDRATION

For orthodox seeds, desiccation‐intolerant developing seeds or dissected immature embryos acquired drought tolerance after de novo synthesis of LEA proteins by precocious maturation (artificial dehydration), treatment with exogenous ABA or osmoticum such as polyethylene glycol or manitol (Bartels et al., 1988; Blackman et al., 1991, 1995; Hsing and Wu, 1992; Hsing et al., 1990; Rosenberg and Rinne, 1989). These results support the association between desiccation tolerance and LEA proteins. Using soybean early‐ germinating seedlings as a model, Blackman et al. (1995) demonstrated that the increased LEA protein level might reduce the electrolyte leakage after desiccation and subsequent rehydration. Exogenous ABA or osmoticum treatment arrested the germinating process in dissected embryos or germinating seeds; the dormant seeds failed to germinate after imbibition and the level

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of LEA proteins in dormant seeds remained constant. Using nondormant seeds, it was demonstrated that LEA protein level declined rapidly after imbibition (Hong et al., 1992; Morris et al., 1990; Ried and Walker‐ Simmons, 1993; Shen et al., 1993). In two closely related cowpea lines (F6 siblines), the chilling‐tolerant line containing a 35‐kDa dehydrin showed maximal percentage emergence and slower leakage of electrolytes during seed imbibition than the genetically similar line without the dehydrin. The relation between physiological and molecular data suggested that this gene might be a useful marker in breeding programs (Ismail et al., 1997, 1999a). Seven winter wheat cultivars were used for the study on dehydrin accumulation during the exposure of drought stress. Three cultivars expressed a 24‐kDa dehydrin after 4‐day stress treatment and at the subsequent sampling dates while no dehydrins were detected in nonstress control plants. Dehydrin expression was significantly delayed in the remaining cultivars. The presence of dehydrins was related to the acquisition of drought tolerance characterized by a greater maintenance of shoot dry matter production (Lopez et al., 2003). LEA proteins accumulate in not only seeds but also vegetative tissues after exogenous ABA treatment or environmental stresses such as chilling, freezing, drought, and salinity stimulation. Barley HVA1 protein accumulated in whole 3‐day‐old seedlings under exogenous ABA treatment. By contrast, in 7‐day‐old seedlings, HVA1 proteins were detected only in roots, but scarcely (Hong et al., 1992). In young wheat seedlings, several LEA III proteins accumulated in shoots and scutellar tissues under severe dehydration (more than 90% water loss). After rehydration, shoots and scutellar tissues resumed growth. The roots showing accumulation of mRNA but not protein did not resume growth and were replaced by new developing roots from scutellar tissue. Hence, the accumulation of LEA III proteins in severely dehydrated seedlings was associated with tissue dehydration tolerance (Ried and Walker‐ Simmons, 1993). Stress‐tolerant varieties of some crops also revealed a high accumulation of LEA proteins. A comparison of protein profiles after ABA treatment revealed the levels of some LEA II and LEA III proteins significantly higher in roots from salt‐tolerant rice varieties than those from salt‐ sensitive ones (Moons et al., 1995). Therefore, increasing endogenous LEA proteins or the addition of extra LEA proteins may increase plant adaptability under extreme environments. In anhydrobiotic plants, such as resurrection plants, mosses, and green alga, many LEA proteins appeared during drought or even during an anhydrobiosis state (Bewley et al., 1993; Hellwege et al., 1996; Honjoh et al., 1995; Joh et al., 1995; Knight et al., 1995). Freezing‐ induced WCS120, a dehydrin‐like protein, accumulated near the plasma membrane but did not integrate into the lipid bilayer (Danyluk et al., 1998).

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M.‐D. SHIH ET AL. B. ECTOPIC EXPRESSION OF LEA PROTEINS

A single LEA gene introduced into model systems may provide systematic tool to measure the degree of stress tolerance. Tobacco (Nicotiana tabacum) plants expressing three diVerent groups of LEA genes from C. plantagineum were unable to reveal any significant improvement over control plants under various osmotic stresses (Iturriaga et al., 1992). Overexpression of Arabidopsis RAB18 (LEA II) in Arabidopsis plants failed to improve freezing or drought tolerance (Lang and Palva, 1992). Transgenic rice harboring barley HAV1 gene (LEA III) was the first successful attempt to confer resistance to drought or salinity stresses (Xu et al., 1996). Transgenic wheat and oat (Avena sativa) harboring HVA1 showed increased desiccation tolerance, biomass productivity, and water use eYciency under high salt, osmotic, or drought conditions via membrane protection (Babu et al., 2004; Maqbool et al., 2002; Sivamani et al., 2000). The accumulation of wheat WCS19, a cold‐regulated chloroplast LEA III protein, in transgenic Arabidopsis after cold acclimation allowed for decreased LT50 (the temperature at which 50% of the total ion leakage occurred) and increased resistance to photoinhibition in leaves under freezing stress, which suggested that WCS19 proteins enhanced freezing tolerance (NDong et al., 2002). In transgenic rice expressing wheat PMA1595 (LEA I) or PMA80 (LEA II), the accumulation of PMA1595 or PMA80 proteins was associated with increased salt‐ and drought‐stress tolerance (Cheng et al., 2002). Accumulation of Arabidopsis AtRAB28 (LEA V) protein improved the germination rate under standard conditions or salt and osmotic stress (Borrell et al., 2002). The accumulation of the citrus dehydrin CuCOR19 in tobacco conferred less electrolyte leakage than control plants under freezing temperature. Transgenic plants also showed earlier germination and better seedling growth than control plants with chilling treatment (Hara et al., 2003). Wheat WCor410 (LEA II) gene introduced into strawberry improved chilling tolerance at 5 8C after previous acclimation treatment (Houde et al., 2004). Rapeseed (Brassica napus) LEA III gene ME‐leaN4 introduced into Chinese cabbage (Brassica campestris) or lettuce (Lactuca sativa L.) resulted in improved drought tolerance (Park et al., 2005a,b). The constitutive expression of maize dehydrin rab17 gene in transgenic Arabidopsis gave high sugar and proline content; in addition, these plants showed more tolerance to high salinity conditions and recovered faster from mannitol treatment than nontransformed control plants (Figueras et al., 2004). However, the transgenic plants showed germination and growth inhibition under high salt conditions, a condition mediated by RAB17 proteins phosphorylation status under desiccation (Riera et al., 2004). The accumulation of a hot pepper hydrophobic LEA V protein, CaLEA6, in transgenic tobacco enhanced tolerance to dehydration and salt

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stresses but not chilling (Kim et al., 2005). ABA‐ and oxidant‐induced Arabidopsis LEA V gene, AtLEA6‐1 (previously called Sag21 or AtLEA5), was introduced into Arabidopsis and increased the root growth and shoot biomass, both in optimal condition and under H2O2 stress. However, the photosynthesis of transgenic plants was more susceptible to drought (Miller et al., 1999; Mowla et al., 2006). A potato (Solanum sogarandinum) dehydrin gene, DHN24, was introduced into cucumber (Cucumis sativus) and increased chilling and freezing tolerance (Rorat et al., 2006; Yin et al., 2006). Additionally, a series of experiments with recombinant yeasts (Saccharomyces cerevisiae) illustrated that increased stress tolerance is directly attributable to the accumulation of LEA proteins. For instance, yeasts harboring tomato Le4 (LEA II), barley HVA1 (LEA III), or tomato Le25 (LEA II) genes were studied under various stresses. Overexpression of these proteins increased tolerance to high concentrations of KCl and to freezing stress but not to sorbitol treatment or heat‐shock stress. However, LE25 and HVA1 proteins but not LE4 proteins also improved the growth rate of yeasts in the media containing a high concentration of NaCl (Imai et al., 1996; Zhang et al., 2000). In addition, the accumulation of wheat Em proteins increased tolerance to a high concentration of NaCl, KCl, or even sorbitol but not to freezing or heat‐shock stress (Swire‐Clark and Marcotte, 1999). Yu et al. (2005) introduced two wheat LEA III genes, TaLEA2 and TaLEA3, into yeast to examine the eVects of these genes on osmotic stress conditions. The results suggested that overexpression of each gene improved sorbitol, salt, or freezing tolerance in transgenic yeast cells. Moreover, accumulation of Arabidopsis AtLEA6‐1 proteins in the oxidant‐sensitive yeast mutant,
yap1, increased the oxidative stress tolerance of
yap1 (Mowla et al., 2006). Nematode AavLEA1 (LEA III) gene transferred into the human cell line T‐Rex293 prevented the aggregation of a wide range of other proteins both in vivo and in vitro (Chakrabortee et al., 2007). In the prokaryote system, soybean GmPM11 (LEA I), GmPM2 (LEA III), and GmPM30 (LEA III) proteins oVered protection against salt or cold stresses, but soybean ZLDE‐ 2 (LEA II) protein was ineVective in protection test (Lan et al., 2005; Liu and Zheng, 2005). In contrast, Arabidopsis LEA II and LEA IV proteins prevented E. coli growth under normal conditions (Campos et al., 2006). These results suggest that diVerent LEA proteins may have unique contributions to cellular protection against stresses. C. IN VITRO STUDIES

To date, because of the limitations of technology, the use of in vitro analyses, such as cryoprotective or desiccation‐protective assay, for LEA proteins is the common strategy to study the protein functions. Without the addition of

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cryoprotectant, the freeze/thaw cycle results in loss of lactate dehydrogenase (LDH, EC 1.1.1.27) activities (Carpenter and Crowe, 1988; Tamiya et al., 1985). Several LEA II proteins, including spinach COR85 (Kazuoka and Oeda, 1994), wheat WSC120 (Houde et al., 1995), peach (Prunus persica) PCA60 (Wisniewski et al., 1999), citrus (Citrus unshiu) CuCOR19 (Hara et al., 2001), barley P‐80 (Bravo et al., 2003), soybean 26/27‐kDa dehydrin (Momma et al., 2003), and citrus fruit (Citrus clementinaCitrus reticulate) dehydrin CrCOR15 (Sanchez‐Ballesta et al., 2004), had been used in cryoprotective assays. Compared with other cryoprotectants such as sucrose and BSA, as well as other proteins, these LEA proteins produced the lowest PD50 value (i.e., the concentration of added molecule required for 50% residual LDH activity after a freeze/thaw cycle). Thus, these dehydrins provided eVective cryoprotection by preventing the freezing denaturation of LDH. Two alga (Chlorella vulgaris C‐27) LEA III proteins, HIC6 and HIC12, and nematode AavLEA1 were also more eVective than sucrose or BSA in protecting LDH level after a freeze/thaw cycle (Goyal et al., 2005; Honjoh et al., 2000). In addition, the results of a cryoprotection assay of artificial shortened HIC6 11‐mer peptides suggested that the cryoprotective activity decreased with a decrease in the repeating unit of the 11‐mer motifs (Honjoh et al., 2000). In addition to their cryoprotection role, some LEA proteins had been demonstrated to have radical scavenging or dehydration protection activity. Citrus CuCOR19 proteins, used to measure the reducing activity of free radicals, were able to remove hydroxyl and peroxyl radicals, suggesting that CuCOR19 proteins directly scavenged radicals (Hara et al., 2004). The pea mitochondria LEA III protein, LEAM, and nematode AavLEA1 also protected LDH, fumarase, and rhodanese in a drying treatment (Goyal et al., 2005; Grelet et al., 2005). In addition, LEAM or lysozyme dried with 1‐steroyl‐2‐oleoyl‐phosphatidylcholine 18:0/18:1 (SOPC) liposome was measured by diVerential scanning calorimetry to monitor the gel–liquid crystalline phase transition temperature after dehydration. The results revealed a major interaction of LEAM but not lysozyme with SOPC liposomes in the dry state (Tolleter et al., 2007). Bdelloid rotifers ArLEA1 and ArLEA2 were reported to prevent the aggregation of desiccation‐sensitive enzymes. However, ArLEA1 only contained partial activity as ArLEA2 had (Pouchkina‐Stantcheva et al., 2007). D. ION SCAVENGER

Several LEA II proteins had been demonstrated to contain Ca2þ‐binding activity. For example, several acidic dehydrins containing the S‐segments showed phosphorylation‐dependent Ca2þ binding (Alsheikh et al., 2003;

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Heyen et al., 2002). Calcium was involved with various plants developmental and signal transduction processes (e.g., Bush, 1995; Hetherington and Brownlee, 2004). Several studies showed that the ability of plants to maintain calcium level was essential to survive through abiotic stresses (Knight et al., 1996; Minorsky and Spanswick, 1989). However, not every S‐segment‐ containing dehydrin had the function of phosphorylation‐dependent Ca2þ binding. The neutral LEA II protein DHR18 contained the S‐segment but had no Ca2þ‐binding activity (Alsheikh et al. 2005). LEA II proteins also had high aYnity to several heavy metal ions such as Fe3þ, Cu2þ, Zn2þ, Mn2þ, and Fe2þ (Herzer et al., 2003; Kruger et al., 2002; Svensson et al., 2000). The aYnity might be caused by a high proportion of His residues in dehydrins (Grossoehme et al., 2006; Svensson et al., 2000).

E. RECALCITRANT SEEDS

Compared with orthodox seeds, recalcitrant seeds underwent little or no maturation dehydration and remain desiccation sensitive during development. Such seeds were shad hydrated, with the minimum water content appearing to be
23% on a wet mass basis. Generally, the water content could range from 30 to 80% (Berjak and Pammenter, 2002; Hoekstra et al., 2001). Unlike orthodox seeds, most recalcitrant seeds could not survive during dehydration or chilling (less than 10 8C) and thus were not able to store for long periods. Some species, such as Avicennia marina, which were extremely dehydration sensitive, did not express LEA genes (Farrant et al., 1992, 1996). However, most recalcitrant seeds still accumulated LEA proteins (the current LEA proteins identified from recalcitrant seeds all belonged to LEA II proteins) and responded to ABA, drought, or temperature stimulation (Farrant et al., 1996; Finch‐Savage et al., 1994; Gee et al., 1994; Han et al., 1997; Kovach and Bradford, 1992a,b). In addition, oligosaccharides, which existed in orthodox seeds, accumulated in some recalcitrant seeds (Farrant et al., 1993; Finch‐Savage and Blake, 1994). The glass formation occurred only at water content
10–15%, whereas such content was well below the lethal limit for recalcitrant seeds (Berjak and Pammenter, 2002; Sun and Leopold, 1994). In summary, LEA proteins and oligosaccharides accumulated in some recalcitrant seeds, and bioglasses might form during dehydration. However, before these protection mechanisms operate, slowly dried recalcitrant seeds already lost viability at water contents far above the essential condition these mechanisms need (Berjak and Pammenter, 2008).

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Recently, two reports suggested that LEA proteins might involve in embryo development or genetic diversity (Manfre et al., 2006; Pouchkina‐Stantcheva et al., 2007). A T‐DNA insertion allele of the Arabidopsis LEA I AtEm6 gene showed that mutant seeds displayed premature seed dehydration and maturation at the distal end of siliques. Complementary test was suYcient to rescue the phenotype of the atem6‐1 mutant allele, suggesting ATEM6 protein was required for normal seed development. However, severe abi3‐4 mutant that no mRNA signal was detectable for both Arabidopsis Em genes did not show premature seed dehydration (Nambara et al., 1992; Parcy et al., 1994). The sequence diVerence between ArLEA1A and ArLEA1B mainly appeared at four 11‐mer repeats indel, while ArLEA1B might have more eYciency than ArLEA1A had. The authors concluded that the functional divergence of former alleles was essential to adopt low genetic diversity, which was the consequence of asexual reproduction for bdelloid rotifers (Pouchkina‐Stantcheva et al., 2007). In plants, several LEA proteins contained similar sequence diversity within the same genome. For example, two soybean LEA III proteins, GmPM8 and GmPM10, shared high homology with an amino acid identity of
90%, with 31 11‐mer repeats for GmPM8 and 32 repeats for GmPM10 (Hsing et al., 1995). Using two Arabidopsis Em genes, AtEm1 and AtEm6, as model, the temporal and spatial studies suggested that these two genes had diverse regulation pathway (Bies et al., 1998; Vicient et al., 2000). Hence, the biological features and evolutionary mechanism for LEA proteins sequence diversity still need further investigation.

V. CONCLUSION AND FUTURE PERSPECTS The term ‘‘late embryogenesis abundant’’ has been used for more than 20 years to describe genes/proteins. However, numerous LEA genes were found not expressed during seed development and expressed only in vegetative tissues. In addition, considering the animal LEA proteins, the term ‘‘late embryogenesis abundant’’ cannot be applied to all situations. Dure et al. (1989) suggested a substitute term, water stress protein, which is not acknowledged widely, possibly because it refers to proteins induced only during water stress. We suggest retaining the label LEA, without emphasizing the feature ‘‘late embryogenesis abundant.’’ LEA proteins were identified almost 30 years ago. From their presence in mature seeds to anhydrobiotic animals, LEA proteins have come to be seen

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as an amazing component with roles in universal dehydration tolerance. Recent studies suggest that LEA proteins may have functions in (1) the molecular coat of the water shell, (2) ion scavenging, and (3) molecular chaperoning. The biophysical features of LEA proteins suggest that they may perform a bipartite function under diVerent cellular water states. Thus, the diVerent protective mechanisms may act at diVerent stages of water loss. Moreover, because a number of nonreducing oligosaccharides could be involved in drought and desiccation (Amuti and Pollard, 1977), the relation between LEA proteins and nonreducing oligosaccharides and the role of oligosaccharides alone should not be ignored. Upon water loss, from the normal range (
80%) to the drought condition (above 23%), the decrease in number of water molecules and thus cellular volume causes crowding of cytoplasmic components, and the cellular contents become increasingly viscous. With the removal of bulk cytoplasmic water, the hydrophilic surface area of proteins and membranes is thermodynamically unfavorable and thus causes abnormal protein aggregation/denaturation and membrane fusion. LEA proteins containing large amounts of water molecules may establish the water shell of intermolecules. That is, the water shell is connected by the hydrogen bonds among LEA–water–oligosaccharide–water–cellular components. Hence, LEA proteins may interrupt the abnormal protein aggregation and membrane fusion because they provide new hydrogen bonds from the water molecules they absorb to the surface of proteins and membranes. In this condition, LEA proteins also act as ion scavengers, because their primary structures contain a high proportion of charged and polar amino acid residues. Studies of model plants or microbes harboring heterogenic LEA genes suggest that the accumulation of LEA proteins may reduce salt damage. In orthodox seeds or other anhydrobiotic tissues, numerous transcription factors, enzymes, or other structure proteins that are desiccation sensitive, are required for protection during seed maturation or anhydrobiosis. During desiccation (from
23 to
10% or lower water content on a dry weight basis), loss of intra‐water molecules causes the dehydrated‐induced LEA proteins to refold and interact with oligosaccharides to perform bioglass. Intracellular bioglasses have been suggested to play a role both in desiccation tolerance and in storage stability. However, simple sugar mixtures or glasses do not tolerate desiccation unless other bioglass components, such as LEA proteins, are involved in the glasses (Koster, 1991; Wolkers et al., 1998a, 1999). Hence, considering the interaction between LEA proteins and oligosaccharides and the role of bioglasses in desiccation, LEA proteins should perform their functions in bioglasses in anhydrobiotic organisms (Walters, 1998; Wolkers et al., 1998a, 1999, 2001; Shih et al., 2004). Under desiccation,

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LEA proteins should directly interact with proteins, oligosaccharides, and the plasma membrane and may enhance bioglass strength, as well as act as a water replacement to stabilize cellular components.

ACKNOWLEDGMENTS This work was partially supported by grants from the Taiwan National Science Council (NSC) and Academia Sinica (AS), and the Netherlands Organization for Scientific Research (NWO) to YICH and FAH. The authors also thank to all the young scholars who worked on LEA gene/ protein research in their labs.

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AUTHOR INDEX

Numbers in bold refer to pages on which full references are listed.

A Aalen, R. B., 216, 253 Abba, S., 215, 217, 240 Aboitiz, N., 144, 193 Adenot, X., 32, 52 Aebi, M., 188, 197 Ainsworth, C. F., 77, 79, 81, 85, 89, 94, 168, 197 Allen, E., 31–32, 52 Allen, T., 92–93, 96 Aloni, R., 3–4, 6, 18, 24, 34, 42, 52 Alperin, D. M., 153, 193 Alpert, P., 212–213, 240 Alsheikh, M. K., 236–237, 240 Ambudkar, S., 21, 52 Amuti, K. S., 213, 239, 240 Anders, N., 23, 52 Andersen, N. H., 144, 193 Andre´, S., 190, 193 Anelli, T., 187, 193 Aravind, L., 27, 63 Arenas-Mena, C., 219, 240 Ariel, F. D., 29, 52 Arnott, S., 223, 240 B Babu, R. C., 234, 240 Bai, C., 188, 193 Baima, S., 5, 32–33, 52 Baker, J., 214, 216–219, 240 Ballario, P., 81, 96 Balusˇka, F., 22, 52 Bao, N., 33, 52 Barbieri, L., 162, 181, 194 Bardocz, S., 183, 203 Barre, A., 137, 154, 173, 190, 194 Bartels, D., 213, 215, 232, 240, 248 Barton, M. K., 27, 61 Baskin, T. I., 44, 52 Bateman, A., 157, 194 Battelli, M. G., 162, 181, 205 Baucher, M., 5–6, 16, 32, 52 Beintema, J. J., 143, 194 Belanger, K. D., 186, 194 Benezra, R., 29, 52

Benjamins, R., 22, 53 Benkova´, E., 21, 53 Bennett, M. J., 18, 22, 53, 60 Berjak, P., 237, 241 Berjak, P., 232, 237, 241 Bewley, C. A., 130, 194 Bewley, J. D., 233, 241 Bi, Y. M., 81, 96 Bianchi, G., 213, 241 Bies, N., 238, 241 Bies-Etheve, N., 216, 221, 241 Blackman, S. A., 213, 232, 241 Blake, P. S., 237, 244 Blakeslee, J. J., 22, 53 Blanchard, D. J., 147, 196 Blasing, O. E., 74, 76, 94, 96 Bleecker, A. B., 41, 65, 189, 205 Blilou, I., 20–21, 53 Blixt, O., 170, 194 Boerjan, W., 4, 46, 53 Bohlenius, H., 16, 53 Bohn-Courseau, I., 40, 66 Bonifacino, J. S., 23, 53 Bonke, M., 6, 15, 39–40, 53 Borevitz, J. O., 46, 53, 74, 98 Borrell, A., 234, 241 Bostwick, D. E., 159, 174, 194 Boudet, J., 219, 223, 226, 228, 241 Bourne, Y., 149, 194 Boutte´, Y., 20, 53 Bowman, J. L., 50, 56 Boyd, M. R., 130, 195 Boyd, W. C., 110, 194 Bradford, K. J., 237, 249 Brandt, A., 147, 197 Bravo, L. A., 236, 241 Bray, E. A., 215, 241 Breton, G., 220, 241 Broekaert, W. F., 183, 195 Brown, D. M., 45, 53 Browne, J., 215, 225, 242 Brownlee, C., 237, 246 Brummendorf, T., 228, 242 Buckley, J. T., 127, 195 Buist, G., 158, 195

258

AUTHOR INDEX

Buitink, J., 213, 242 Burk, D. H., 44, 53 Burke, M. J. 213, 242 Busch, M., 23, 53 Bush, D., 237, 242 Busse, J. S., 10, 17, 54 Bycroft, M., 157, 194 C Campbell, S. A., 217, 242 Campos, F., 235, 246 Can˜o-Delgado, A. I., 5, 15, 34, 37, 54 Candela, H., 24, 54 Carland, F. M., 15, 25, 54 Carlsbecker, A., 6, 54 Carpenter, J. F., 236, 242 Carpentier, S. C., 180, 195 Carre, I. A., 80, 82, 96 Carrizo, M. E., 122, 195 Cashmore, A. R., 81, 103 Ceccardi, T. L., 223, 242 Chakrabortee, S., 235, 242 Chantret, I., 186, 195 Chapple, C. C. S., 5, 54 Chattopadhyay, S., 84, 96 ChaVey, N., 16, 54 Chen, R., 20, 54 Chen, X., 192, 195 Chen, Y., 159, 172, 175, 184, 195, Chen, Z. Y., 218, 242 Cheng, Z. Q., 234, 242 Chinchilla, D., 190, 195 Choe, S., 5, 37, 54 Choi, J. H., 214, 242 Chou, J. H., 222, 242 Christensen, S. K., 22, 54 Chua, N. -H., 217, 250 Cicero, D. O., 228, 242 Cilia, M. L., 8, 54 Claes, B., 178, 195 Clark, S. E., 41, 54, 189 Clavel, C., 149, 195 Clegg, J. S., 213, 242–243 Close, T. J., 215, 217, 242–243 Cock, J. M., 41, 54, 189–190, 195 Cohen, A., 218, 243 Collins, C. H., 213, 243 Colmenero-Flores, J. M., 218, 222, 243 Coutinho, C., 213, 243 Coutinho, P. M., 45, 54 Covington, M. F., 74, 76, 89, 96 Crowe, J. H., 212–213, 227, 243 Cuming, A. C., 214–216, 218, 243 Curry, J., 218, 243 D Danon, A., 49, 54 Danyluk, J., 233, 243

Datta, A., 126, 204 David, K. M., 88, 96 de Dorlodot, S., 12, 14, 55 de Leon, B. G., 35, 55 del Pozo, O., 48, 55 Delwiche, C. F., 4, 55 Demura, T., 17, 55 Dengler, N., 24–25, 55, 59, 62 Deshaies, R. J., 188, 202 De Souza Filho, G., 178, 195 Deyholos, M. K., 6, 65 DeYoung, B. J., 41, 55, 189, 195 Di Laurenzio, L., 27, 55 Dinant, S., 161, 196 Ding, Z., 92, 96 Dixon, H. B. F., 111, 196 Doblin, M. S., 45, 55 Dodd, A. N., 73, 84, 96 Donaldson, J. G., 23, 55 Donaldson, L. A., 46, 55 Doolittle, R. F., 216, 249 d’Ortous de Mairan, J. J., 70, 102 Douglas, C., 14–16, 58 Dover, S. D., 223, 240 Doyle, M. R., 90–91, 96 Drickamer, K., 114, 206 Dudareva, N., 74, 96–97 Dunker, A. K., 228–229, 243 Dure, L., 213–218, 222, 238, 244 Dyson, H. J., 229, 255 E Ebringerova, A., 45, 55 Edelman, G. M., 153, 196 Edwards, K. D., 71, 74, 94, 97 Eichinger, L., 215, 217, 244 Eimert, K., 87, 97 Elfstrand, M., 110, 196 Elge, S., 5, 55 Emery, J. F., 5, 15, 28–29, 32, 55 Eom, J. W., 223, 242 Esen, A., 147, 196 Eshed. Y., 15, 29, 31, 55, 56 Espelund, M., 216, 244 Etzler, M. E., 153, 196 Evert, R. F., 10, 17, 54 F Fahlgren, N., 32, 56 Fairchild, C. D., 84, 97 Farrant, J. M., 213, 237, 244 Farre, E. M., 77, 85–86, 94, 97 Fasman, G. D., 222, 242 Field, C. B., 73, 98 Figueras, M., 234, 244 Finch-Savage, W. E., 237, 244 Fisher, K., 40–41, 56

259

AUTHOR INDEX Fletcher, J. C., 40–41, 56, 67 Floyd, S. K., 50, 56 Foster, R., 83, 97 Fouquaert, E., 132, 134, 137, 172, 175, 179–180, 196 Fowler, S., 74, 87–88, 97 Frana, M. B., 213, 245 Freeling, M., 83, 97 Frigerio, L., 164, 196 Friml, J., 20–23, 56 Fukuda, H., 6, 10, 16–17, 27, 33, 37–38, 43, 47, 50, 56, 58, 62 Funk, V., 47, 56 Fusetti, F., 129, 196 G Ga¨lweiler, L., 20, 25, 56, 63 Gabaldon, C., 50, 56 Gagne, J. M., 189, 196 Gaidamashvili, M., 167, 196 Gal, T. Z., 215, 245 Galau, G. A., 214, 216–217, 219, 245, 247 Gallagher, K., 27, 56 Garbers, C., 22, 56 Garbuzynskiy, S. O., 229, 245 Garcia, D., 30–31, 57 Gardiner, J. C., 45, 57 Gaubier, P., 216, 245 Gaut, B. S., 193, 197 Gee, O. H., 237, 245 Geisler, M., 21–22, 57 Geldner, N., 23, 57 Geshi, N., 147, 197 Ghouse, A. K. M., 11, 58 Gilles, G. J., 223, 245 Gilmour, S. J., 217, 245 Giovanini, M. P., 185, 197 GiVord, M. L., 190, 197 Glenn, K. A., 188, 197 Goday, A., 217, 245 Goff, S. A., 95, 97 Goicoechea, M. 47, 57 Gokce, I., 223, 245 Goldberg, R. B., 214, 245 Goldstein, I. J., 110, 197 Golecki, B., 159, 197 Golovina, E. A., 213, 246 Gong, W., 82, 97 Goodman, H. M., 14, 57 Go¨rlach, J., 179, 197 Gosti, F., 220, 222, 245 Gouget, A, 190, 197 Goyal, K., 226–227, 231–232, 236, 245 Grebe, M., 23, 57 Green, K. A., 15, 57 Grelet, J., 225, 236, 246

Groover, A., 50, 57 Grossoehme, N. E., 237, 246 Grunwald, I., 178–179, 197 Gustafson, K. R., 130, 197 Gutierrez, R. A., 77, 97 H Hall, A., 91–92, 98 Han, B., 237, 246 Hand, S. C., 215, 218, 246 Hansen, G., 48, 57 Hara, M., 224, 234, 236, 246 Harada, J. J., 215, 246 Hardman, K. D., 168, 197 Harmer, S. L., 73–74, 76–77, 79–80, 82, 84, 94–98 Hart, G. W., 186 Hassidim, M., 77, 98 Hatsugai, N., 48, 57 Hazen, S. P., 79–80, 85, 98 He, X. -J., 191, 197 Heidstra, R., 27, 66 Heim, M. A., 84, 98 Heimburg, T., 227, 246 Heinze, T., 45, 55 Helariutta, Y., 6, 17, 27, 57 Helenius, A., 188, 197 Hellwege, E. M., 233, 246 Hennessey, T. L., 73, 98 Herve´, C., 154, 190, 197–198 Herzer, S., 237, 246 Hester, G., 136, 168, 198 Hetherington, A. M., 237, 246 Heyen, B. J., 237, 246 Hicks, K. A., 89, 98 Higuchi, M., 34–35, 58 Hirabayashi, J., 170, 198 Hirano, K., 178, 198 Hirsch, A. M., 81, 198 Hoekstra, F. A., 212–213, 237, 246–247 Hong, B., 214, 233, 247 Honjoh, K., 233, 247 Honjoh, K. I., 236, 247 Horbowicz, M., 213, 247 Houde, M., 234, 236 Hsing, Y. I. C., 218, 220, 238, 247 Hudson, M. E., 31, 79–80, 82–83, 98 Hughes, D. W., 214, 247 Hunter, C., 32, 58 Huq, E., 87–88, 98 Hutchison, C. E., 35–36, 58 Hwang, I., 35–36, 58, 189, 198 Hyun, Y., 30, 58

Igarashi, M., 5, 58 Iliev, I., 16, 58

I

260 Imai, R., 215, 222, 235, 247 Imaizumi, T., 74, 98 Imanishi, S., 178, 198 Ingram, J., 215, 248 Inoue, T., 34–35, 58 Iqbal, M., 11, 58 Ishiura, M., 79, 85, 100, 102 Ismail, A. M., 224, 233, 248 Ito, Y., 17, 41, 47, 208 Iturriaga, G., 234, 248 Iwakawa, H., 31, 58 Iwasaki, T., 37, 58 J Jackson, C. L., 23, 53, 65 Jackson, D., 8, 54 Jackson, R. B., 12, 14, 23, 64 Jacobs, P. J., 18, 66 Jacobs, W. P., 18, 58 Jain, M., 189, 198 Jakoby, M., 84, 98 Jang, S., 74, 98 Jansson, S., 14–16, 58 Jiang, J. F., 147, 179, 198 Jiao, Y., 79, 82–83, 98, 99 Joh, T., 215, 233, 248 Johnson, C. H., 72, 99 Jolliffe, N. A., 164, 198 Jones, A. M., 50, 57 Jones, L., 213, 248 Jordinson, M, 181, 198 Joris, B., 156, 198 Jouve, L., 74, 99 Jung, J. H., 33, 58 Jurgens, G., 40, 66 K Kader, J. C., 39, 58 Kahn, T. L., 218, 248 Kai, G., 138, 199 Kakimoto, T., 35, 59 Kaku, H., 157, 199 Kamiya, Y., 186, 199 Kane, R., 186, 199 Kang, J., 24, 33, 59 Katinka, M. D., 215, 248 Katsuhara, K., 50, 59 Kawaoka, A., 47, 59 Kawasaki, T., 50, 59 Kay, S. A., 80, 82, 85–86, 94–95, 98, 101–102, 104 Kazuoka, T., 215, 236, 248 Keilin, D., 212, 248 Kerstetter, R. A., 28, 59 Kevei, E., 92, 99 Khanna, R., 80, 90–91, 99 Kiba, T., 36, 59, 86, 99

AUTHOR INDEX Kidner, C. A., 15, 26, 27, 31, 59 Kieliszewski, M. J., 144, 199 Kik, M. J., 181, 199 Kikawada, T., 215, 218, 248 Kikis, E. A., 90–91, 94, 99 Kim, H. S., 219, 235, 248 Kim, J., 15, 33, 59 Kim, J. Y., 82, 96 Kim, Y. S., 29–30, 51, 59, 93 Kipreos, E. T., 188, 199 Kitagaki, H., 153, 199 Kiyosue, T., 219, 248 Kleine-Vehn, J., 23, 59 Knight, C. D., 215, 233, 248 Knight, H., 237, 249 Koag, M. C., 224, 249 Koch, A. J., 25, 59 Kocourek, J., 132, 202 Koharudin, L. M., 131, 199 Koike, M., 220, 249 Koizumi, K., 23, 25, 59 Komamine, A., 16, 37, 38, 56 Kondo, T., 41, 60 Koster, K. L., 213, 239, 249 Kovach, D. A., 237, 249 Kramer, E. M., 18, 60 Kraulis, P. J., 228, 249 Kruger, C., 237, 249 Krupa, A., 189–190, 199 Kubo, M., 6, 15, 41–42, 45, 60 Kuroda, H., 189, 199 Kurup, S., 86, 99 Kyte, J., 216, 249 L Lam, E., 48–49, 55, 67 Lan, Y., 235, 249 Landsteiner, K. 110, 199 Lane, B. G., 214, 216, 243, 250, 253 Lang, V., 234, 249 Lannoo, N., 161, 172, 175, 179, 184, 187, 189, 200 Lechner, E., 189, 200 Lee, C., 45, 60 Lee, H. I., 143, 200 Lee, I., 30, 58 Lee, R. H., 178, 200 Lehmann, K., 47–48, 60 Leitch, A. R., 193, 200 Leonhardt, N., 180, 200 Leopold, A. C., 213, 237, 249, 253 Leprince, O., 213, 242, 249 Lerner,D. R., 143, 200 Li, H., 15, 31, 60 Liang, H., 49, 60 Lidder, P., 77, 99 Lim, C., 46, 64 Limpens, E., 157, 200

AUTHOR INDEX Lin, R., 22, 60, 93, 99 Lin, W. C., 31, 60 Linden, H., 81, 99 Linman, M. J., 170, 200 Linthorst, H. J. M., 143, 200 Lis, H., 153, 204 Lisse, T., 224, 249 Litts, J. C., 214, 216, 249 Liu, J., 222, 252 Liu, X. L., 74, 89–90, 99 Liu, Y., 235, 249 Locke, J. C., 94, 99, 100 Lopez, C. G., 233, 249 Loris, R., 152, 168, 200 Love, J., 74, 100 Luschnig, C., 20–21, 60 M Ma¨ho¨nen, A. P., 15, 34–36, 60, 61 Macino, G., 81, 99 Madin, K. A. C., 213, 249 Madsen, E. B., 157, 192, 200 Maeda, M., 187, 200 Maher, E. P., 20, 60 Makino, S., 85, 100 Mallappa, C., 84, 100 Manchen˜o, J. M., 127, 200 Mancuso, S., 22, 61 Manevski, A., 84, 100 Manfield, I. W., 81, 100 Manfre, A. J., 238, 250 Mann, K., 147, 201 Maqbool, B., 234, 250 Mariano, A. C., 190, 201 Martienssen, R. A., 15, 59 Martin, J., 217, 250 Martindale, S. J. B., 20, 60 Martinez-Garcia, J. F., 84, 100 Martin-Tryone, E. L., 87–88, 100 Mas, P., 79, 81, 85–86, 94, 100, 102 Maslenkova, L., 213, 251 Mathur, J., 5, 37, 61 Matsubayashi, Y., 41, 61 Matsuo, S., 187, 199 Matsuo, T., 95, 100 Matsushika, A., 85, 100 Mattsson, J., 18, 24–25, 61 Mayer, U., 23, 61 McClung, C. R., 74, 79, 82, 85–86, 95, 99, 101–102, 104–105 McConnell, J. R., 15, 27, 33, 61 McCormick, S., 41, 54 McCubbin, A. G., 223, 231, 250 McCully, M. E., 5, 62 McQueen-Mason, S., 213, 248 McWatters, H. G., 89–91, 94, 100 Meagher, J. L., 149, 201

261

Medeiros, A., 153, 201 Meijer, A. H., 4, 6, 11–12, 18, 64 Meinhardt, H., 25, 61 Menkens, A. E., 83–84, 101 Merchant, S. S., 95, 101 Meusser, B., 188, 201 Michael, T. P., 74, 76, 79–80, 82, 84, 94–95, 98, 101 Michniewicz, M., 23, 61 Milioni, D., 17, 41, 61 Millar, A. J., 71, 76–97, 101 Miller, J. D., 235, 250 Minorsky, P. V., 237, 250 Mirkov, T. E., 173, 201 Mitchison, G. J., 24, 61 Mitsuda, N., 46, 61 Mizoguchi, T., 80, 87–88, 101 Momma, M., 236, 250 Monzingo, A. F., 163, 168, 204 Moons, A., 178–179, 201, 233, 250 Moore, S. E., 186, 187, 201 Morgan, W. T. J., 110, 168, 208 Morris, D. A., 20, 61 Morris, P. C., 233, 250 Moshelion, M., 71, 101 Motose, H., 5, 17, 38–39, 62 Mouillon, J. M., 224, 250 Mowla, S. B., 235, 250 Mu¨ller, A., 20, 62 Mueller, J. P., 215–216, 250 Mundy, J., 214, 217, 250 Munoz, E., 83, 101 Murakami, M., 86, 101 Murdock, L. L., 183, 201 Murphy, A. S., 22, 57 Murphy, D. J., 213, 250 Murthy, B. N., 170, 201 N Nagata, T., 20, 37, 62 Nakagawa, R., 178, 201 Nakahata, Y., 83, 101 Nakamichi, N., 86, 101 Nakamura-Tsuruta, S., 117, 201 Nakashima, T., 49, 62 Nakata, S., 181, 201 Nambara, E., 238, 250 Navarro-Gochicoa, M. -T., 192, 201 NDong, C., 234, 250 Nelson, T., 25, 62 Ng, D. T. W., 187, 208 Niinuma, K., 86, 101 Nishiguchi, M., 189, 191, 202 Nomura, K., 147, 202 Nozue, K., 71, 74, 102 Nylander, M., 215, 251

262

AUTHOR INDEX

O Obara, K., 47, 62 Obendorf, R. L., 213, 247 O’Brien, T. P., 5, 62 Oda, A., 84, 102 Oda, Y., 15, 62 Oeda, K., 215, 236, 248 Oguri, S., 172, 202 Ohashi-Ito, K., 38, 62 Okada, K., 20, 62 Oliver, A. E., 213, 251 Oliverio, K. A., 87, 102 Onaga, S., 157, 202 Onai, K., 79, 85, 102 ¨ nnerud, H., 46, 62 O Opassiri, R., 127, 202 Otsuga, D., 28, 32, 62 P Paca´k, F., 132, 202 Pagano, M., 188, 199 Palme, K., 20, 63 Palva, E. T., 234, 249 Pammenter, N. W., 232, 237, 241, 244, 246 Panda, S., 93, 102 Paquet, E. R., 83, 102 Para, A., 86, 102 Parcy, F., 218, 238, 251 Park, B. J., 234, 251 Park, C. M., 33, 58 Park, D. H., 87–88, 102 Parret, A. H., 136, 202 Patzlaff, A., 47, 63 Paulson, J. C., 170, 202 Peeva, V., 213, 251 Pekker, I., 30, 63 Pen˜a, M. J., 45, 63 Peragine, A., 32, 63 Perales, M., 79, 81, 94, 102 Percudani, R., 131–132, 202 Persson, S., 45, 63 Petra´sˇek, J., 20–21, 63 Petroski, M. D., 188, 202 Petryniak, J., 132, 202 Peumans, W. J., 111, 117, 122, 137–138, 143, 146–147, 164, 171, 174, 182–183, 202, 203, 206 Piatkowski, D., 219, 251 Pichersky, E., 74, 96 Pilgrim, M. L., 77, 102 Pla, M., 219, 251 Plana, M., 217, 251 Pollard, C. J., 213, 239, 240 Ponting, C. P., 27, 63 Po¨tter, E., 178, 203 Potts, M., 213, 251

Pouchkina-Stantcheva, N. N., 227, 236, 238, 251 Pratap, J. V., 149, 203 Prestrelski, S. J., 231, 251 Priem, B., 187, 203 Prigge, M. J., 15, 33, 63 Prusinkiewicz, P., 25, 64 Pusztai, A., 181, 183, 203 Puthoff, D. P., 122, 185, 203 Pysh, L. D., 27, 63 Q Qin, Q. -M., 178, 203 Quail, P. H., 79, 80, 82–83, 97–99, 102, 103 Quartacci, M. F., 213, 251 R Rabijns, A., 149, 203 Radutoiu, S., 157, 192, 203 Raikhel, N. V., 141, 143, 204 Raina, A., 126, 204 Rashotte, A. M., 20, 63 Rathjen, F. G., 228, 242 Raubitschek, H., 110, 199 Raval, S. 168, 204 Raynal, M., 220, 251 Reed, J. C., 49, 67 Reguera, R. M., 110, 194 Renkonen, K. O., 110, 204 Reyes, J. C., 81, 102 Rhoades, M. W., 28, 33, 63 Riccardi, F., 180, 204 Riechmann, J. L., 82, 102 Ried, J. L., 233, 251 Riera, M., 217, 234, 251 Rinderle, S. J., 124, 204 Rinne, R. W., 214, 232, 252 Riou, C., 190–191, 204 Roberts, A. W., 44, 63 Robertus, J. D., 163, 168, 204 Rolland-Lagan, A. G., 25, 64 Rorat, T., 235, 252 Rose´n, S., 122, 204 Rosenberg, L. A., 214, 232, 252 Ross-Ibarra, J., 193, 197 Rossjohn, J., 127, 204 Rost, B., 222, 252 Rotari, V. I., 48, 64 Roussel, M. R., 46, 64 Russouw, P. S., 223, 252 Rutenber, E., 163, 204 S Saavedra, L., 217, 252 Sachs, T., 4, 12, 18, 24–25, 64 Sakagami, Y., 41, 60, 61

AUTHOR INDEX Sakai, H., 36, 64 Sakai, I., 172, 208 Salome, P. A., 85, 104 Samyn, B., 180, 204 Sanchez-Ballesta, M. T., 236, 252 Sander, C., 222, 252 Santelia, D., 22, 64 Sasabe, M., 192, 204 Sato, E., 86, 102 Savidge, R., 16, 58 Sawa, M., 87–88, 102 Scarpella, E., 4, 6, 11–12, 18, 25, 64 Schaffer, R., 74, 77, 85, 89, 94, 103 Schenk, H. J., 12, 14, 64 Scheres, B., 15, 34, 36, 64 Schibler, U., 76, 84, 103, 104 Schoning, J. C., 77, 103 Schrader, J., 50, 64 Schweers, O., 228, 252 Scorrano, L., 49, 64 Shade, R. E., 183, 201 Sharma, V. K., 41, 64 Sharon, N., 153, 204 Sheen, J., 35–36, 58, 198 Shen, Q., 233, 252 Shevell, D. E., 23, 64 Shibaoka, H., 37, 58 Shibuya, N., 163, 171, 204 Shih, M. D., 22, 218, 225, 239, 252 Shiu, S. H., 41, 65, 189 Sidler, M., 22, 65 Sieburth, L., 5, 9, 12, 18, 66 Sieburth, L. E., 6, 24–25, 41, 65 Siegfried, K. R., 31, 65 Singh, S., 228, 252 Singh, T., 168, 205 Sinha, S., 168, 205 Sitia, R., 187, 193 Sivamani, E., 234, 252 Skriver, K., 214, 252 Smalle, J., 188, 205 Smith, A. K., 124, 206 Smith, L. G., 27, 56 Smith-Espinoza, C. J., 213, 253 Soedjanaatmadja, U. M., 143, 205 Somers, D. E., 73, 85, 103 Soulages, J. L., 223–224, 253 Spanswick, R. M., 237, 250 Spiro, R. G., 186, 205 Sreerama, N., 223, 253 Stacy, R. A., 216, 253 Staiger, D., 77, 91, 103 Steeves, R. M., 163, 205 Steeves, T. A., 42, 65 Steiner-Lange, S., 46, 65 Steinmann, T., 5, 20, 23, 25, 65 Stirpe, F., 162, 181, 205

263

Strayer, C., 77, 85, 103 Strosberg, A. D., 153, 205 Suarez-Lopeze, P., 74, 103 Subramanyam, S., 184, 205 Sun, W. Q., 237, 253 Sun, Y., 48, 65 Sunkar, R., 213, 253 Sussex, I. M., 42, 65 Suzuki, L., 145, 205 Suzuki, S., 35, 55 Suzuki, T., 34, 65 Svensson, J., 237, 253 Swarup, R., 20, 22, 65 Szekeres, M., 37, 65 Szyjanowicz, P. M., 45, 65 T Taira, T., 157, 202 Taiz, L., 8, 11–12, 65 Tajima, T., 89, 103 Takahashi, R., 220, 253 Takasaki, T., 189, 205 Tamagnone, L., 47, 66 Tamiya, T., 236, 253 Tang, G., 33, 66 Tang, H., 193, 206 Tateno, H., 170, 206 Taylor, G., 14–15, 66 Taylor, M. E., 114, 206 Taylor, N. G., 45, 66 Teakle, G. R., 81, 103 Ten Hove, C. A., 27, 66 Teneberg, S. 172, 206 Tepperman, J. M., 90, 103 Terasaka, K., 22, 66 Terzaghi, W. B., 81, 103 Thain, S. C., 74, 103 Thompson, E. W., 214, 253 Thompson, M. V., 8, 66 Thompson, N. P., 18, 66 Timmermans, M. C., 26–27, 31, 59 Tobin, E. M., 77, 85, 104 Toledo-Ortize, G., 84, 103 Tolleter, D., 225–227, 236, 253 Torres-Ruiz, R. A., 40, 66 Traas, J., 40, 66 Transue, T. R., 124, 206 Treml, B. S., 20, 66 Tremousaygue, D., 84, 103 Tseng, T. S., 88, 104 Tsutsui, S., 136, 206 Tuba, Z., 213, 253 Tunnacliffe, A., 227, 253, 254 TunnacliVe, A., 216, 218, 253 Turner, S., 5–10, 12, 14–15, 17–18, 40–41, 43–44, 47–50, 66 Tuskan, G. A., 15–16, 95 Tyree, M. T., 7–8, 66

264

AUTHOR INDEX

U Uversky, V. N., 228–229, 254 V Valverde, F., 74, 104 van Bel A. J., 8, 66 Van Damme, E. J. M., 111, 116, 126, 129–131, 136–140, 141, 143, 145–146, 149, 152–153, 155, 159, 162–164, 170–172, 174–178, 181–184, 202, 206, 207 Van Parijs, J., 183, 207 van Raemdonck, D., 16, 66 Van Roekel, T., 213, 247 Vandepoele, K., 83, 95, 104 Vega, N., 153, 207 Velten, J., 217, 254 Vicient, C. M., 238, 254 Vicre, M., 213, 254 Viczian, A., 84, 104 Vierstra, R. D., 188, 196, 295 Vieten, A., 20–21, 56, 66 Vilardell, J., 217, 254 Vo¨lker, U., 215–216, 254 W Waites, R., 31, 67 Waljuno, K., 141, 207 Walker, J. C., 189, 207 Walker-Simmons, M. K., 218, 233, 243, 251 Walley, J. W., 74, 104 Walters, C., 239, 254 Wan, J., 157, 190, 207–208 Wang, H., 22, 37, 50, 60, 67 Wang, J. L., 153, 196 Wang, X., 187, 206 Wang, M. -B., 174, 208 Wang, W., 215, 254 Wang, Z. Y., 77, 80, 85, 104 Watanabe, N., 49, 67 Watkins, W. M., 110, 168, 208 Wenkel, S., 29–30, 51, 67, 86–87, 98, 104 Whitelam, G. C., 92, 96, 104 Wilde, H. D., 214, 254 Williams, C. E., 15, 31–33, 40, 147, 179, 184, 208 Wilson, C., 213, 254 Wise, M. J., 216, 218, 220, 227, 231, 253, 254

Wisniewska, J., 21, 67, 217, 236 Wolkers, W. F., 213, 225, 227, 232, 239, 255 Woody, R. W., 223, 255 Wozniak, R. W., 186, 208 Wright, P. E., 168, 229, 255 Wu, A. M., 117, 208 Wu, S., 232, 247 Wuarin, J., 76, 104 X Xu, D., 20, 31, 49, 213, 215, 234, 255 Y Yagi, F., 179, 208 Yakir, E., 77, 104 Yamamoto, R., 17, 37–38, 67 Yamamoto, S., 172, 208 Yamashino, T., 86, 104 Yang, H. P., 219, 255 Yang, Y., 22, 67 Yang, F., 22, 130, 208 Yanhui, C., 82, 104 Yanovsky, M. J., 74, 104 Ye, Z. H., 4–7, 10–11, 14–15, 17, 28, 43–44, 53, 67, 68 Yee, V. C., 228, 255 Yin, Z. M., 235, 255 Yokoyama, A., 36, 67 Yong, W. D., 147, 179, 208 Yoshida, Y., 188, 208–209 Young, N. M., 146, 209 Yu, J. N., 49, 95, 235, 255 Z Zachara, N. E., 186, 209 Zagotta, M. T., 88, 105 Zaı´malova´, E., 20, 68 Zegzouti, H., 219, 255 Zeiger, E., 8, 11–12, 65 Zeilinger, M. N., 94, 105 Zhang, L., 157, 235, 255 Zhang, W., 174, 178, 209 Zhao, C., 50, 68 Zhong, H. H., 77, 105 Zhong, R, 15, 28, 46, 63, 68 Zhong, R. Q., 44, 68 Zhou, A., 5, 68 Zhu-Salzman, K., 184, 209 Zimmermann, M. H., 7–8, 66 Zuo, K., 190, 209

SUBJECT INDEX

A ACC SYNTHASE8 gene (ACS8), 75 Actin, vascular development, 45 Agaricus bisporus agglutinin homologs, 117, 122–124 embryophyta genomes, 118–121 Amaranthins aerolysin domains, 127 fascin-like domain, 126–127 Hfr–2 protein homology, 126 morphology, 124 protein morphology, 125 taxonomic distribution, 127–128 Antirrhinum Majus, 74 Arabidopsis genetic analysis of, 14 weak points, 15 Arabidopsis thaliana, 71 Artocarpus integrifolia -prism fold, 149 chimerolectins, 147, 151 Embryophyta lineage, 150 jacalin-related lectins, 146–147 purified proteins, 148 taxonomic groupings, 149, 151 Atypical LEA proteins, 220 AUX1, AUX family, 22

B BAX inhibitor-1, 49 Brassinosteroid critical roles of, 37 ZeHB12 expression of, 38 Brefeldin A, PAT studies, 23

C Canalization hypothesis, 24 Carbohydrate binding domains Agaricus bisporus agglutinin homologs, 117, 122–124 embryophyta genomes, 118–121 Amaranthins aerolysin domains, 127 fascin-like domain, 126–127 Hfr–2 protein homology, 126 morphology, 124 protein distribution, 125 taxonomic distribution, 127–128

Artocarpus integrifolia -prism fold, 149 chimerolectins, 147, 151 Embryophyta lineage, 150 jacalin-related lectins, 146–147 purified proteins, 148 taxonomic groupings, 149, 151 Cucurbitaceae phloem lectins. See Nictaba family cyanovirin-N (CV-N) protein, 130–132 Dioscorea batatas, 167 Euonymus europaeus agglutinin (EEA) Euonymus lectin (EUL) domain architecture, 133–134 gene characters, 134–135 structural composition, 132–133 taxonomic distribution, 135–136 Galanthus nivalis agglutinin (GNA) cloning and X-ray diVraction analysis, 136–137 cytoplasmic homologs, 139 domain architecture, 140–141 S-locus protein kinases, 139–140 synthesis, 137 vacuolar lectins, 137–138 Hevea brasiliensis domains post-translational processing, 141–143 tri-N-acetylchitotriose (NAG3) complexes, 144–145 viridiplantae lineage, 145–146 legume lectin domains, 151 bacterial origins, 154–155 chimeric protein origin, 155–156 hololectins’ prevalence, 153, 154 LegL and LegLu relationship, 153, 155 molecular structure, 152 promoter’s structure, 152–153 taxonomic distribution, 118–121, 155 Lysin (LysM) domain domain architecture, 156–157 structure, 157 taxonomic distribution, 158–159 Nictaba family domain architecture, 160 Embryophyta lineage, 161 lectin formation, 159 taxonomic distribution, 161 plant lectins, 114–115 Ricin-B family domain architecture, 163–165

266

SUBJECT INDEX

Carbohydrate binding domains (cont. )

molecular evolution, 167 structure analysis, 162–163 taxonomic distribution, 163, 166 Robinia pseudoacacia (RobpsCRA) morphology, 129 secretory pathway synthesis, 129–130 Chlorophyll a/b-binding proteins (CAB), 73 Circadian clock free Ca2þ ions, cytosolic concentration of, 72 pathways, 71 switch induction, 74 Cytokinin AHP6 locus, 36 His-Asp phosphorelay mechanism, 35 procambial cells, 34 receptor genes, 34 E Early flowering3 (ELF3) cloning, 89 essence of elf3 consensus, 89 seedling photomorphogenesis, aspects of, 88 Early flowering4 (ELF4), 90–91 Euonymus europaeus agglutinin (EEA) Euonymus lectin (EUL) domain architecture, 133–134 gene characters, 134–135 structural composition, 132–133 taxonomic distribution, 135–136 F Far-red elongated hypocotyl3 (FHY3), 92–93 FIONA1 (FIO1), 93–94 G Galanthus nivalis agglutinin (GNA) cloning and X-ray diVraction analysis, 136–137 cytoplasmic homologs, 139 domain architecture, 140–141 S-locus protein kinases, 139–140 synthesis, 137 vacuolar lectins, 137–138 GIGANTEA (GI) clock defects, 87 cloning of, 88 Gnetales, 8 GNOM proteins, 23 H Helix-loop-helix domain, 29 Hemagglutinin, 110

Hevea brasiliensis domains post-translational processing, 141–143 tri-N-acetylchitotriose (NAG3) complexes, 144–145 viridiplantae lineage, 145–146 His-Asp phosphorelay mechanism, 36 L Late embryogenesis abundant proteins arabidopsis LEA genes genome sequencing project, 221 groups, 222 lea proteins, classification and occurrences Atypical LEA proteins, 220 average molecular weight, 219 cotton developing embryos, 217 cotton LEA D-34 and D-95 proteins, 219 criteria used, 216 D-11 proteins, 217 D-113 proteins, 218 D-7 proteins, 218 distinct classes, 215 domain and segment, 217 Kyte–Doolittle hydropathy algorithm, 216 wheat Em and cotton D-19 proteins, 216 living organisms, water status desiccation tolerance, 212–213 drought tolerance, 212 oligosaccharides, 213 tolerance, types, 212 POPP classification basic principles of, 220 subgroups, 221 seed development and lea proteins early-methionine-labeled protein, 214 embryogenesis, 214 late embryogenesis abundant, 214 surface-solvation moieties, 215 LEA proteins hydrophilic LEA proteins amino acid content, comparison, 230 gel electrophoresis, 231 hydrodynamic data, 231 protein folding, 229 three-dimensional structure, 228 hydrophobic LEA proteins hydrophobicity and heat instability, 227 MtPM25 proteins, 228

SUBJECT INDEX in dry status GmPM1 proteins, temperature-scanning FTIR spectra, 226 secondary structures, 225 sucrose, dehydrated mixture, 227 in solution status CD analysis, 223 CuCOR19 proteins, 224 EMB-1 proteins, 223 fine structures, identification of, 222 gel filtration and hydrodynamic experiments, 223 soybean GmPM16 proteins, 225 physiological roles dehydration, accumulation, 232–233 ectopic expression, 234–235 in vitro studies, 235–236 ion scavenger, 236–237 mode of action, 238 orthodox seeds, 232 recalcitrant seeds, 237 storage role, 232 structure of hydrophilic lea proteins, 228–232 hydrophobic LEA proteins, 227–228 in dry status, 225–227 in solution status, 222–225 LEA I proteins, 216–217, 223 LEA II proteins, 217–218, 223–224 LEA III proteins, 218, 225 LEA IV proteins, 218–219, 225 LEA V, 219 Lectin, 110–111 Lectinomes, 193 Legume lectin domains, 151 bacterial origins, 154–155 chimeric protein origin, 155–156 hololectins’ prevalence, 153, 154 LegL and LegLu relationship, 153, 155 molecular structure, 152 promoter’s structure, 152–153 taxonomic distribution, 118–121, 155 Lens culinaris, 110 Light input, plant clock factors mediating early flowering3, 88–90 early flowering4, 90–91 far-red elongated hypocotyl3, 92–93 light-insensitive period1, 92 sensitivity to red-light reduced1, 91 time for coVee, 91–92 independent factors fiona1, 93–94 tej, 93 Light-insensitive period1 (LIP1), 92 Lignin terrestrial biopolymer, 46 Lysin (LysM) domain domain architecture, 156–157

267

structure, 157 taxonomic distribution, 158–159 M Meristems, vascular plants, 9 N Nicotiana tabacum, 72 Nictaba family domain architecture, 160 Embryophyta lineage, 161 lectin formation, 159 taxonomic distribution, 161 P Phaseolus vulgaris, 110 Phloem, 8–9 Phloem fibers, 11 Phytochrome interacting factor4/5 (PIF4/5), 74 Pisum sativum, 110 Plant gene expression, clock control ACC SYNTHASE8 gene, 75 biological pathways carbon fixation, 73 developmental responses, 74 ion fluxes and stomatal aperture, 72–73 leaf movements, 71 metabolic processes, 73–74 photosynthesis and plant fitness, 73 plant hormone production and stress responses, 74–75 circadian transcriptome characteristics analysis, 75 circadian-regulated genes, 76 evening module cycling promoters, 80 GATA motif, 80 nine nucleotide sequence, 79 WC-1 and WC-2, 81 LUX ARRHYTHMO, 79 morning module elements identified, 82 G-box-binding proteins, characterization, 83 phytochrome A pathway, 82 mRNA decay and oscillations, 76–78 plant clock, architecture gigantea, 87–88 light independent factors, 93–94 light inputs, 88–93 TOC1 and PRRS, 85–87 PRR7 and PRR9, 78 transcription, impacts CIS-elements association, 84 phase-associated CIS-elements, identification, 79–84 transcriptional feedback loop, 78–79

268

SUBJECT INDEX

Plant lectins biochemical analyses, 115–116 and carbohydrate binding Agaricus bisporus, 117–124 Amaranthins, 124–128 cyanovirin-N (CV-N) protein, 130–132 defective lectin homologs, 173 Dioscorea batatas, 167 Euonymus europaeus agglutinin (EEA), 132–136 Galanthus nivalis agglutinin (GNA), 136–141 Hevea brasiliensis domains, 141–146 legume lectin domains, 151–156 Lysin (LysM) domain, 156–159 Nictaba family, 159–161 Ricin-B family, 162–167 Robinia pseudoacacia (RobpsCRA), 129–130 cell signaling, receptor kinases, 189–192 classical lectins insect-induced lectins, 184 jasmonate/stress-induced proteins, 183–184 storage/defense-related proteins, 182 cytoplasmic lectins, discovery, 175 expression and induction patterns classical concepts, 181–182 ‘‘classical lectins, 177, 182 inducible ‘‘novel lectins, 178–179 novel concepts, 183 glycoprotein degradation research, 188–189 hevein domain lectins, 171–172 multi-specificity lectins, 172–173 nonsecretory lectins, evidence for, 174 paradigms carbohydrate binding specificity, 113–114 definition and nomenclature, 111 general biology and physiological role, 113 occurrence and taxonomical distribution, 112 ribosome inactivating proteins (RIPs), 112–113 protein–carbohydrate interactions research glycosylation, 185 N-glycosylation, 186–187 snowdrop lectin, hapten inhibition, 171 specificity studies, new approaches, 170–171 stress-related induction, 180 sugar binding activity

aYnity, complex glycans, 169 specificity, N glycans, 170, 171 three-dimensional structures of domains 168 synthesis on free ribosomes, 176 tobacco agglutinin, 172 transcriptome databases, 116–117 Polar auxin transport ABC transporters, 21–22 AtPGP19 function, 22 AUX1, 22 basipetal transport, 18 brefeldin A, 23 GNOM protein, 23 influx and eZux proteins, 20 molecular mechanism of, 19 PIN localization, 21 POPP classification basic principles of, 220 interrelated software tools, 220 LEA protein groups, 220–221 Populus powerful model system, 16 tree-specific traits, 15 Populus trichocarpa, 79 Production of anthocyanin pigments1 (PAP1), 73 Pseudo-response regulators (PRRs), 85–87. See also TOC1 R Reaction-diVusion model, 25 Ricin, 109–110 Ricin-B family domain architecture, 163–165 molecular evolution, 167 structure analysis, 162–163 taxonomic distribution, 163, 166 Robinia pseudoacacia (RobpsCRA) morphology, 129 secretory pathway synthesis, 129–130 Root growth cell growth stages, 13 developmental stages in angiosperms, 13 major function, 12 stele (vascular cylinder), 13 S Samanea saman, 71 Sensitivity to red-light reduced1 (SRR1), 91 Shoot growth procambial cells, 10 protoxylem and metaxylem, 11 secondary growth, 11 Sieve tube, phloem, 8 Stele, root growth, 13

SUBJECT INDEX T Taxol MT-stabilizing drug, 43 TEJ, 93 Time for coVee (TIC), 91–92 Timing of CAB2 expression1 (TOC1) CCA1/LHY expression, 86 heme activator protein complex, 86 mapping, 85 photoperiodic flowering, 86 pseudo-response regulators, 85 robust circadian expression, 85 Tracheids, water-conducting cells, 42 V Vascular development system adaxial-abaxial axis determinants AS1 and AS2, 31 ETT and ARF4, 30 apl mutant, 40 brassinosteroid critical roles, 37 ZeHB12 expression of, 38 cell death atbil-1 and atbil-2, 49–50 BAX inhibitor-1, 49 PCD, 48–49 vacuolar processing enzyme, 48 xylem, 47 conductive cells, 3 continuity and regulatory signals auxin, 18 stem veins functions, 17 cytokinin AHP6 locus, 36 distanct types, 36 His-Asp phosphorelay mechanism, 35 procambial cells, 34 receptor genes, 34 diVerentiation process, 5 genetic and molecular biological approaches, 5–6 HD-ZIP IIIs and kanadis gain-of-function, 28 loss-of-function, 28–29 PHB and PHV, 27–28 putative transcription factor, 28 HD-ZIP IIIs and miRNA ATHB8, 32 START domain, 33 inhibtors, HD-ZIP IIIs regulation helix-loop-helix, 29 little zippers, 29 ZPR3 protein, 30 major classes, 4 model systems arabidopsis thaliana, 14–15 populus, 15–16 zinnia, 16–17

269

MYB coiled-coil-type transcription factor family, 39 phloem intercalated with xylem mutant, 40–41 physiological, biochemical, and molecular approaches, 4–5 phytohormone functioning, 51 plant growth dicots and monocots, comparison, 11–12 meristems, 9 phloem fibers, 11 procambial cell formation, 10 protoxylem and metaxylem, 11 secondary growth, 11 shoot growth, 10–11 polar auxin transport ABC transporters, 21–22 AUX1, 22 basipetal transport, 18 brefeldin A, 23 GNOM protein, 23 influx and eZux proteins, 20 molecular mechanism, 19 PIN proteins, 20–21 radial patterning, regulation of adaxial and abaxial identities, 26 adaxial–abaxial patterning, 27 asymmetric cell divisions, 27 root growth cell growth stages, 13 concentric cell layers, 12–13 development stages in angiosperms, 13 major function of, 12 stele, 13 SAM identity genes, 50 secondary wall formation actin, 45 cell wall types, 43 cortical microtubules, 43–44 fragile fiber mutants, 44 hemicellulose, 45 in wood, 44–45 lignin, 46 taxol, MT-stabilizing drug, 43 stem development stages, 3 strand formation models canalization hypothesis, 24–25 pattern of veins, leaves, 24 reaction–diVusion model, 25 tracheary element (TE) diVerentiation diVerentiation processes, 43 water-conducting cells, 42 vascular tissues bundles, 6 essential roles, 6 phloem, 8–9 xylem, 7–8

270 Vascular development system (cont. )

vascular-related NAC-domain proteins, 42 xylogen BLAST search, 39 in vitro Zinnia culture, 38–39 TE diVerentiation, 39 Vascular-related NAC-domain proteins, 42 Vicia sativa, 110

SUBJECT INDEX X Xylem, 7–8 Xylogen BLAST search, 39 in vitro Zinnia culture system, 38 TE differentiation, 39 Z Zinnia stages in, 17 xylem diVerentiation, 16