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
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
VOLUME 54
AMSTERDAM • BOSTON • HEIDELBERG • LONDON
NEW YORK • OXFORD • PARIS • SAN DIEGO
SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO
Academic Press is an imprint of Elsevier
Academic Press is an imprint of Elsevier 32 Jamestown Road, LondonNW17BY,UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands First edition 2010 Copyright � 2010 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:
[email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material 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: 978-0-12-380870-7 ISSN: 0065-2296 For information on all Academic Press publications visit our website at www.elsevierdirect.com Printed and bound in the United States of America 10 11 12 13 10 9 8 7 6 5 4 3 2 1
Working together to grow libraries in developing countries www.elsevier.com | www.bookaid.org | www.sabre.org
CONTRIBUTORS TO VOLUME 54
´ LE ` NE CITERNE HE UMR de Ge´ne´tique Ve´ge´tale, CNRS—Univ Paris-Sud—INRA—AgroParisTech, Ferme du Moulon, 91190 Gif-sur-Yvette, France CATHERINE DAMERVAL UMR de Ge´ne´tique Ve´ge´tale, CNRS—Univ Paris-Sud—INRA—AgroParisTech, Ferme du Moulon, 91190 Gif sur-Yvette, France JONG-CHIN HUANG Department of Chemical Biology, National Pingtung University of Education, Pingtung, 90003, Taiwan, ROC LOUIS JOHN IRVING Graduate School of Life and Environmental Science, University of Tsukuba, 1-1-1 Tennodai, Tsukuba 305-8572, Japan HIROYUKI ISHIDA Graduate School of Agricultural Science, Tohoku University, 1-1 Tsutsumidori Amamiyamachi, Aoba-ku, Sendai 560-0043, Japan FLORIAN JABBOUR Universite´ Paris-Sud, Laboratoire Ecologie, Syste´matique, Evolution, CNRS UMR 8079, AgroParisTech, Orsay, F-91405, France; and Institute for Systematic Botany and Mycology, University of Munich, Menzinger Strasse 67, 80638 Munich, Germany GUANG-YUH JAUH Institute of Plant and Microbial Biology, Academia Sinica, NanKang, Taipei 11529, Taiwan, ROC; and Biotech nology Center, National Chung-Hsing University, Taichung, 402, Taiwan, ROC AMANE MAKINO Graduate School of Agricultural Science, Tohoku University, 1-1 Tsutsumidori Amamiyamachi, Aoba-ku, Sendai 560-0043, Japan SOPHIE NADOT Universite´ Paris-Sud, Laboratoire Ecologie, Syste´ma tique, Evolution, CNRS UMR 8079, AgroParisTech, Orsay, F-91405, France YUJI SUZUKI Graduate School of Agricultural Science, Tohoku Uni versity, 1-1 Tsutsumidori Amamiyamachi, Aoba-ku, Sendai 560-0043, Japan HUEI-JING WANG Institute of Plant and Microbial Biology, Academia Sinica, NanKang, Taipei 11529, Taiwan, ROC GEORGE CHUCK Plant Gene Expression Center, United States Department, Agriculture-Agriculture Research Service, Albany, CA 94710, United States
CONTENTS OF VOLUMES 35–53 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
Contents of Volume 36 PLANT VIRUS VECTOR INTERACTIONS
Edited by R. Plumb
x
CONTENTS OF VOLUMES 35–53
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 Volume 37 ANTHOCYANINS IN LEAVES
Edited by K. S. Gould and D. W. Lee
CONTENTS OF VOLUMES 35–53
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
xi
xii
CONTENTS OF VOLUMES 35–53
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 35–53
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
xiii
xiv
CONTENTS OF VOLUMES 35–53
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
CONTENTS OF VOLUMES 35–53
Nutrient Sensing and Signalling in Plants: Potassium and Phosphorus ANNA AMTMANN, JOHN P. HAMMOND,
PATRICK ARMENGAUD AND PHILIP J. WHITE
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
xv
xvi
CONTENTS OF VOLUMES 35–53
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 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
CONTENTS OF VOLUMES 35–53
xvii
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 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
xviii
CONTENTS OF VOLUMES 35–53
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 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 VOLUMES 35–53
xix
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 MARIANA JOVANOVIC, VALE´ RIE LEFEBVRE, PHILIPPE
LAPORTE, SILVINA GONZALEZ-RIZZO, CHRISTINE
LELANDAIS-BRIE´ RE, FLORIAN FRUGIER, CAROLINE
HARTMANN AND MARTIN CRESPI
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
xx
CONTENTS OF VOLUMES 35–53
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
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
Contents of Volume 48 Molecular Mechanisms Underlying Vascular Development JAE-HOON JUNG, SANG-GYU KIM, PIL JOON SEO
AND CHUNG-MO PARK
Clock Control Over Plant Gene Expression ANTOINE BAUDRY AND STEVE KAY
CONTENTS OF VOLUMES 35–53
Plant Lectins ELS J. M. VAN DAMME, NAUSICAA LANNOO
AND WILLY J. PEUMANS
Late Embryogenesis Abundant Proteins MING-DER SHIH, FOLKERT A. HOEKSTRA
AND YUE-IE C. HSING
Contents of Volume 49 Phototropism and Gravitropism in Plants MARIA LIA MOLAS AND JOHN Z. KISS Cold Signalling and Cold Acclimation in Plants ERIC RUELLAND, MARIE-NOELLE VAULTIER,
ALAIN ZACHOWSKI AND VAUGHAN HURRY
Genome Evolution in Plant Pathogenic and Symbiotic Fungi GABRIELA AGUILETA, MICHAEL E. HOOD,
GUISLAINE REFRE´ GIER AND TATIANA GIRAUD
Contents of Volume 50 Aroma Volatiles: Biosynthesis and Mechanisms of Modulation During Fruit Ripening BRUNO G. DEFILIPPI, DANIEL MANRI´QUEZ,
KIETSUDA LUENGWILAI AND MAURICIO
´ LEZ-AGU ¨ ERO
GONZA Jatropha curcas: A Review NICOLAS CARELS You are What You Eat: Interactions Between Root Parasitic
Plants and Their Hosts
LOUIS J. IRVING AND DUNCAN D. CAMERON
xxi
xxii
CONTENTS OF VOLUMES 35–53
Low Oxygen Signaling and Tolerance in Plants FRANCESCO LICAUSI AND PIERDOMENICO PERATA Roles of Circadian Clock and Histone Methylation in the Control of Floral Repressors RYM FEKIH, RIM NEFISSI, KANA MIYATA,
HIROSHI EZURA AND TSUYOSHI MIZOGUCHI
Contents of Volume 51 PAMP-Triggered Basal Immunity in Plants ¨ RNBERGER AND BIRGIT KEMMERLING THORSTEN NU Plant Pathogens as Suppressors of Host Defense JEAN-PIERRE ME´ TRAUX, ROBERT WILSON JACKSON,
ESTHER SCHNETTLER AND ROB W. GOLDBACH
From Nonhost Resistance to Lesion-Mimic Mutants:
Useful for Studies of Defense Signaling
ANDREA LENK AND HANS THORDAL-CHRISTENSEN Action at a Distance: Long-Distance Signals in Induced Resistance MARC J. CHAMPIGNY AND ROBIN K. CAMERON Systemic Acquired Resistance R. HAMMERSCHMIDT Rhizobacteria-Induced Systemic Resistance ¨ FTE DAVID DE VLEESSCHAUWER AND MONICA HO Plant Growth-Promoting Actions of Rhizobacteria STIJN SPAEPEN, JOS VANDERLEYDEN AND YAACOV OKON
CONTENTS OF VOLUMES 35–53
xxiii
Interactions Between Nonpathogenic Fungi and Plants M. I. TRILLAS AND G. SEGARRA Priming of Induced Plant Defense Responses UWE CONRATH Transcriptional Regulation of Plant Defense Responses MARCEL C. VAN VERK, CHRISTIANE GATZ
AND HUUB J. M. LINTHORST
Unexpected Turns and Twists in Structure/Function of PR-Proteins that Connect Energy Metabolism and Immunity MEENA L. NARASIMHAN, RAY A. BRESSAN,
MATILDE PAINO D’URZO, MATTHEW A. JENKS
AND TESFAYE MENGISTE
Role of Iron in Plant–Microbe Interactions P. LEMANCEAU, D. EXPERT, F. GAYMARD, P. A. H. M. BAKKER AND J.-F. BRIAT Adaptive Defense Responses to Pathogens and Insects LINDA L. WALLING Plant Volatiles in Defence MERIJN R. KANT, PETRA M. BLEEKER, MICHIEL VAN WIJK,
ROBERT C. SCHUURINK AND MICHEL A. HARING
Ecological Consequences of Plant Defence Signalling MARTIN HEIL AND DALE R. WALTERS
Contents of Volume 52 Oxidation of Proteins in Plants—Mechanisms and Consequences LEE J. SWEETLOVE AND IAN M. MØLLER
xxiv
CONTENTS OF VOLUMES 35–53
Reactive Oxygen Species: Regulation of Plant Growth and Development HYUN-SOON KIM, YOON-SIK KIM, KYU-WOONG HAHN,
HYOUK JOUNG AND JAE-HEUNG JEON
Ultraviolet-B Induced Changes in Gene Expression and Antioxidants in Plants S. B. AGRAWAL, SURUCHI SINGH AND
MADHOOLIKA AGRAWAL
Roles of �-Glutamyl Transpeptidase and �-Glutamyl Cyclotransferase in
Glutathione and Glutathione-Conjugate Metabolism in Plants
NAOKO OHKAMA-OHTSU, KEIICHI FUKUYAMA AND
DAVID J. OLIVER
The Redox State, a Referee of the Legume–Rhizobia Symbiotic Game DANIEL MARINO, CHIARA PUCCIARIELLO, ALAIN PUPPO
AND PIERRE FRENDO
Contents of Volume 53 Arabidopsis Histone Lysine Methyltransferases FRE´DE´RIC PONTVIANNE, TODD BLEVINS,
AND CRAIG S. PIKAARD
Advances in Coffea Genomics ALEXANDRE DE KOCHKO, SE´ LASTIQUE AKAFFOU, ALAN
ANDRADE, CLAUDINE CAMPA, DOMINIQUE CROUZILLAT,
ROMAIN GUYOT, PERLA HAMON, RAY MING, LUKAS
A. MUELLER, VALE´ RIE PONCET, CHRISTINE TRANCHANT DUBREUIL, AND SERGE HAMON
Regulatory Components of Shade Avoidance Syndrome ` JAIME F. MARTI´NEZ-GARCI´A, ANAHIT GALSTYAN, MERCE
´ SALLA-MARTRET, NICOLAS CIFUENTES-ESQUIVEL, MARC ¸ AL
GALLEMI´, AND JORDI BOU-TORRENT
CONTENTS OF VOLUMES 35–53
xxv
Responses of Halophytes to Environmental Stresses with Special
Emphasis to Salinity
KSOURI RIADH, MEGDICHE WIDED, KOYRO HANS-WERNER, AND ABDELLY CHEDLY Plant Nematode Interaction: A Sophisticated Dialogue PIERRE ABAD AND VALERIE M. WILLIAMSON Optimization of Nutrition in Soilless Systems: A Review ´ NGELES CALATAYUD ELISA GORBE AND A
Pollen Germination and Tube Growth
HUEI-JING WANG,* JONG-CHIN HUANG† AND
GUANG-YUH JAUH*,þ,1
*
Institute of Plant and Microbial Biology, Academia Sinica, NanKang, Taipei 11529, Taiwan, ROC † Department of Chemical Biology, National Pingtung University of Education, Pingtung, 90003, Taiwan, ROC þ Biotechnology Center, National Chung-Hsing University,
Taichung, 402, Taiwan, ROC
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Pollen Adhesion and Hydration on the Stigma . . . . . . . . . . . . . . . . . A. Initial Adhesion between Pollen and Stigma . . . . . . . . . . . . . . . B. Pollen–Stigma Interaction During Cross-Linking Adhesion . . . . C. Specific Protein Pairs in Pollen–Stigma Cross-Linking Adhesion D. Pollen Hydration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Pollen Tube Growth in the Style . . . . . . . . . . . . . . . . . . . . . . . . III. Endomembrane Trafficking in the Tip Region Contributes to Rapid
and Polar Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Increased Cellular Modification and Differentiation in
Germinating Pollen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Exo/Endocytotic Vesicles Maintain Polar and Rapid
Tube Elongation in Pollen Tubes . . . . . . . . . . . . . . . . . . . . . . . C. Small GTPASE and its Regulatory Components Contribute
to Membrane Trafficking in Pollen Tube Polar Growth . . . . . . . D. Roles of Phospholipids, their Derivatives and Phospholipase
C in Membrane Trafficking of Elongating Pollen Tubes . . . . . .
2
4
4
5
7
8
10
14
15
18
21
23
Huei-Jing Wang and Jong-Chin Huang have been contributed equally to this work 1
Corresponding author: E-mail:
[email protected] Advances in Botanical Research, Vol. 54 Copyright 2010, Elsevier Ltd. All rights reserved.
0065-2296/10 $35.00
DOI: 10.1016/S0065-2296(10)54001-1
2
H.-J. WANG ET AL.
IV. Roles of Actin Cytoskeleton, Ionic and Regulatory Proteins in Regulation of Pollen Tube Growth. . . . . . . . . . . . . . . . . . . . . . . . . . A. Actin-Binding Proteins Control Actin Remodelling and Dynamics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Actin-Associated Motor Proteins Mediate Vesicles or Organelle Movement. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Ionic and Protein Regulators Contribute to Oscillatory Pollen Tube Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Global Analysis of Gene Expression in Pollen Tubes . . . . . . . . . . . . VII. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
26 27 29 31 36 37 39 39
ABSTRACT Seeds and grains are staple food supply around the world and their production is the result of successful double fertilization of female gametophytes by male gametophytes. In flowering plants, the highly reduced haploid male gametophyte (pollen grain) plays a critical role in fertilization and crop production through the generation of a pollen tube that delivers the male gametes (two sperm cells) to the egg and central cells within the female gametophyte (embryo sac) for double fertilization. Double fertilization and the functional specialization of the male gametophyte are two key innovations behind the evolutionary success of angiosperms. Pollen tube elongation involves typical polar cell growth and provides a marvelous system to study the fundamental events of morpho genesis, cellular compartmentation and signalling networks in establishing and main taining cellular identities. Furthermore, pollen germination on the stigmatic papilla cell surface and pollen tube elongation in the style offer an exceptional system to study the molecular and cellular mechanisms involved in cell-to-cell communication and signal ling. Here, we summarize current knowledge and important discoveries of genes/proteins and mechanisms essential for pollen germination, pollen tube elongation and guidance in the style that highlight new insights into the elements regulating the complex interaction between these two partners during fertilization. In addition, during the past decade, study of plant biology has been enhanced by the development of advanced instruments and valuable data: highly annotated genomes, comprehensive genetic and transcriptomic resources, readily available T-DNA insertional mutants and revolutionary microscope and imaging systems. The simultaneous availability of these tremendous resources and tools opens a new era to conduct genome-wide and systematic investigation of the molecular and cellular mechanisms governing pollen germination and tube elongation.
I. INTRODUCTION In most animals, germ-line cells differentiate from somatic cells during early embryogenesis and remain as a distinct stem cell population throughout their lives. Nevertheless, plants have a complex life cycle
POLLEN GERMINATION AND TUBE GROWTH
3
with the alteration of diploid sporophytic and haploid gametophytic phases, and the germ lines formed inside flowers are derived from a few somatic-origin cells (Ko¨hler and Grossniklaus, 2005). In flowering plants, after landing on the stigmatic surface of the pistil, pollen grains hydrate and germinate, and each one extrudes a polarized outgrowth to form a pollen tube. Pollen tubes elongate rapidly within pistil tissues, targeting ovules, until the sperm cells are discharged within the cytoplasm of the degenerating synergids (Sandaklie-Nikolova et al., 2007). This highly reduced haploid male gametophyte plays a critical role in fertilization and crop production by delivering the resident male gametes (two sperm cells) to the egg and central cells within the female gametophyte (embryo sac) for double fertilization (Cheung and Wu, 2008). Seed development is initiated after the double fertilization of the egg and central cells by the two sperm cells, giving rise to the diploid embryo and the triploid endo sperm (Ko¨hler and Grossniklaus, 2005). Double fertilization and the functional specialization of the male gametophyte are two key innovations behind the evolutionary success of angiosperms (Borg et al., 2009). Under standing the molecular and cellular events in pollen germination and tube elongation will provide new insights into the regulatory elements that control male germ-line identity and male molecules involved in the inter action with the female partner during fertilization. The journey of pollination starts from adhesion of pollen on stigmatic papillae, and complex cell-to-cell interactions determine the reception or rejection of pollen. After hydration, the pollen tube forms and penetrates into female tissue, elongates along the stylar transmitting track and finally reaches the micropyle and enters into the embryos sac to release two sperm cells for double fertilization in the female gametophyte. Both the male and female partners play important roles during all these processes. The system is unique for study of cell-to-cell interactions and signalling and has been used to reveal fundamental knowledge of many important cellular events. In addition, the pollen tube in angiosperms is a single and extremely reduced male gametophytic cell, and its elongation involves a specialized polar cell growth. This unique apical growth offers an excellent cell model to study the molecular and cellular mechanisms in morphogenesis, cellular compartmen tation and signalling networks essential for establishing and maintaining cellular identities. In this chapter, we summarize the cellular and molecular events regulating pollen germination and pollen tube growth. In addressing the rapid and marked modification of cellular compartmentation during pollen germination, we do not cover other important aspects during pollen formation, such as anther development, microsporogenesis, microgameto genesis, self-incompatibility and fertilization.
4
H.-J. WANG ET AL.
II. POLLEN ADHESION AND HYDRATION
ON THE STIGMA
A. INITIAL ADHESION BETWEEN POLLEN AND STIGMA
Pollen grains transfer from a quiescent state to a germination program when they land on the stigma. Successful landing and adhesion is the essential first step to initiate an intimate mating dialogue between both male and female gametophytes. Before tightly adhering on the stigma, pollen grains must be captured by stigma. In plants with wet stigmas, such as Lilium, the initial adhesion factors are present in the stigmasecreted exudates. Liquid surface tension on the wet stigma is suggested to be sufficient to support pollen capture (Heslop-Harrison, 1981; Heslop-Harrison and Shivanna, 1977). In plants with dry stigmas, the compositions in the pollen cell walls determine the fate of the reorganiza tion step. The pollen wall is complex, with high divergence of compo nents among species (Blackmore et al., 2007). In general, most pollen walls are composed of two major distinct layers: the exine, accumulated primarily by sporopollenin, and the intine, mainly composed of pectin and cellulose. In addition, different protein- and lipid-rich materials form a pollen coat (tryphine) anchor on the exine. Exine has a key role in the initial pollen–stigma adhesion in dry-stigma plants. Binding assay results clearly revealed that a defective pollen coat in Arabidopsis eceriferum (cer6-2) mutated pollen does not affect initial binding significantly (Zinkl et al., 1999), but removal of the pollen coat via chemical extrac tion showed a similar result. For example, several treatments, including salts, chaotropes, divalent cation chelators, reducing or oxidizing reagents, protease and lipase, did not disrupt the initial pollen–stigma adhesion, but certain detergents did. These results indicate that the initial pollen–stigma adhesion is likely through hydrophobic interaction caused by a primary substance in the exine and hydrophobic moiety-rich sporopollenin (Zinkl et al., 1999). Exine purified from Arabidopsis pollen binds to stigma with high affinity, which indicates that the initial adhesion is exine dependent (Zinkl et al., 1999). Pollen grains from mutant less adherent pollen (lap1) revealed markedly aberrant exine and were defective in adhesion (Zinkl and Preuss, 2000). Exine-involved pollen capture was also shown to be a species-specific property (Zinkl et al., 1999). Several other genes or proteins participating in exine formation have been studied in Arabidopsis; examples are MALE STERILITY 2 (Aarts et al., 1997), DEX1 (Paxson-Sowders et al., 2001), faceless pollen-1 (Ariizumi et al., 2003), NEF1(Ariizumi et al., 2004), b-1,3
POLLEN GERMINATION AND TUBE GROWTH
5
glucan synthases (Dong et al., 2005; Nishikawa et al., 2005), MALE STERI LITY1 (Ito et al., 2007; Yang et al., 2007), cytochrome P450 (Morant et al., 2007), CER3 (Rowland et al., 2007), small GTPase Rop (Zhang and McCormick, 2007), exine 1 (Ariizumi et al., 2008), RUPTURED POLLEN GRAIN1 (Guan et al., 2008), KNS2 (Suzuki et al., 2008), fatty acyl-CoA synthetase (de Azevedo Souza et al., 2009), LAP3 (Dobritsa et al., 2009b) and long-chain fatty acid omega-hydroxylase (Dobritsa et al., 2009a). Mutations in these genes showed various defects in exine formation. Although the initial adhesion was found to be exine dependent, further dissection of the mole cular mechanism of the first binding step is difficult because the biosynthetic process and the exact structure of sporopollenin are unclear (Piffanelli et al., 1998; Scott, 1994). New findings in molecular genetics will promote an understanding of sporopollenin synthesis and exine development. The mechanism of pollen capture in the stigmatic aspect is also difficult to define. The waxy cuticle on the surface of a dry stigma might participate in this initial pollen capture. Acetone treatment of stigmatic papillae from Brassica oleracea causes disabled pollen capture (Heizman et al., 2000) because of lack of wax on the surface of cuticle papilla. The hydrophobic interaction might account for pollen capture between lipophilic molecules of the exine wall and lipid-rich stigma cuticle, but the molecular mechanism of the stigmatic cuticle layer in the partner selection task remains unclear. After initial adhesion of pollen on the stigma, pollen germination is determined by the pollen population effect: the number of pollen grains affecting pollen germination. Phytosulphokine-a (PSK-a), a sulphated pen tapeptide, could stimulate pollen germination in vitro and might be the determinant of the pollen population effect (Chen et al., 2000). PSK-a is a signalling molecule with multiple functions (Matsubayashi et al., 2001), is released from cultured pollen with less than 2-hrs incubation and can significantly enhance germination of low-density cultured pollen (Chen et al., 2000). A leucine-rich repeat receptor kinase (LRR-RK) might function as a PSK-a receptor (Matsubayashi and Sakagami, 2000; Matsu bayashi et al., 2002, 2006a, b; Shinohara et al., 2007), but the detailed mechanism remains to be deciphered. B. POLLEN–STIGMA INTERACTION DURING CROSS-LINKING ADHESION
After pollen grains land on the stigma, the germination program moves forward to the next stage, pollen–stigma cross-linking (Swanson et al., 2004), in which the interactions between pollen and stigma result in a tighter connection than with pollen capture. The interaction of the pollen coat (tryphine) and stigma forming the “foot” structure has a key role during
6
H.-J. WANG ET AL.
this adhesion in plants with a dry stigma (Elleman et al., 1992). This pollen foot, a phenomenon called “coat conversion”, still exists when the pollen coat in other regions is removed by treating pollinated stigma with nonpolar solvent cyclohexane (Elleman and Dickinson, 1986, 1990). Removal of the pollen coat in the mutant cer6-2 or by chemical treatment adversely affects adhesion but has no effect on pollen capture (Heizman et al., 2000; Luu et al., 1997; Zinkl et al., 1999). The pellicle, a membrane-like proteinaceous layer on the stigmatic papillae cuticle (Gaude and Dumas, 1986), and the membrane-like exinic outer layer (EOL) of pollen (Gaude and Dumas, 1984) are involved in early pollen–stigma cross-linking. EOL is involved in pollen adhesion in Brassica (Heizman et al., 2000; Luu et al., 1997; Zinkl et al., 1999), but its composition has not been characterized. In B. oleracea, the pellicle of protease-treated stigma collapses, and pollen adhesion is adversely affected (Luu et al., 1997; Stead et al., 1980). This adverse effect recovered within 2 hr after protease treatment, but recovery of adhesion is inhibited after immersion of the protease-treated stigma in cycloheximide, a protein translation inhibitor. These results reveal that the pellicle can be repaired by depositing renewable proteins (Roberts et al., 1984; Stead et al., 1980). Although the definite composition of the pellicle remains uncertain, cyto chemical analyses indicated the presence of glycoproteins and nonspecific esterases (Heslop-Harrison et al., 1975; Knox et al., 1976; Mattsson et al., 1974), as well as ATPases, adenylate cyclases and peroxidases (Gaude and Dumas, 1986; Pandey, 1967), on the stigma surface of several dry-stigma plants. It will be interesting to know the roles of these proteins or enzymes in pollen–stigma adhesion. Recently, a stigma-specific peroxidase (SSP) from Senecio squalidus was identified (McInnis et al., 2005) and appeared in the cytosol of stigmatic papillae and in the pellicle, as revealed by immunolocalization studies (McInnis et al., 2006b). Peroxidases catalyse the breakdown of H2O2, which along with other reactive oxygen species (ROS) are crucial molecules in cell signalling in various aspects (Foreman et al., 2003; Hancock et al., 2006; Laloi et al., 2004; Meinhard et al., 2002; Neill et al., 2002a, b; Rentel and Knight, 2004). In S. squalidus and Arabidopsis, high amounts of ROS or H2O2 are localized to the stigmatic papillae, and their amounts were reduced by applying pollen grains or nitric oxide (NO) on stigmas (Hiscock et al., 2007; McInnis et al., 2006a, b). Recent study suggested that pollen grains could generate NO (Hiscock et al., 2007) and NO might function as a signalling molecule in pollen tube guidance (Prado et al., 2004; 2008). Although it is intriguing that high amounts of stigmatic ROS or H2O2 and high levels of peroxidase activity coexist in receptive, unpollinated stigmas, the regulation process might be involved in harmonizing this situation.
POLLEN GERMINATION AND TUBE GROWTH
7
Pollen-synthesized NO may be a triggering signal molecule to signal the stigma for pollen grain landing, to reduce the amount of ROS or H2O2, perhaps through the function of an SSP or other unidentified factors and unknown mechanisms, and eventually to promote pollen adhesion and germination. C. SPECIFIC PROTEIN PAIRS IN POLLEN–STIGMA CROSS-LINKING ADHESION
Two stigmatic S-locus-related proteins, S-locus-related 1 (SLR1) and S-locus glycoprotein (SLG), have been implicated in pollen–stigma cross-linking adhesion in Brassica. SLR1 protein accumulates in the stigma papillae cell wall (Umbach et al., 1990) and shows a highly conserved property across plant taxa (Lalonde et al., 1989). Both antisense suppression of SLR1 expression and pretreatment of wild-type stigmas with anti-SLR1 antibodies significantly weakened pollen–stigma adhesion but not during initial pollen capture (Luu et al., 1999). SLR1 interacts with its pollen counterpart by binding to S-locus-related binding proteins, SLR1-BP1 and SLR1-BP2, which are members of class A pollen coat proteins (PCP-A) (Doughty et al., 1993; Hiscock et al., 1995; Takayama et al., 2000). However, not all of the Brassica group use SLR1-BP as the interactive partner of SLR1, because the expression of SLR1-BP was detected only in the A genometype chromosomes, such as Brassica napus (AACC), Brassica juncea (AABB) and Brassica campestris (AA), by northern blot and PCR-based analysis (Takayama et al., 2000). Like SLR1, SLG expresses in stigmatic papillae, and its proteins are secreted and accumulated in cell walls (Kandasamy et al., 1989; Umbach et al., 1990). The role of SLG in pollen–stigma adhesion was revealed by masking SLG by treatment with monoclonal anti-SLG antibo dies on the stigma to block this adhesion event (Luu et al., 1999). SLG binds PCP-A1 in vitro, which suggests another “potential” pollen–stigma adhesion mechanism with this interaction (Doughty et al., 1993, 1998; Hiscock et al., 1995). SLG–PCP-A1 interaction is not the comprehensive phenomenon in Brassica because SLG is absent in some S-haplotypes (Suzuki et al., 2000, 2003). However, PCP-A1 is not the only partner of SLG: at least 10 proteins in the pollen coat of B. campestris were identified to interact with SLG (Takayama et al., 2000). The above data imply that SLG may play multiple roles in the pollen–stigma interaction and one of them mediates pollen– stigma adhesion. Although evidence shows SLR1 and SLG included in the process of pollen–stigma adhesion, the detailed mechanism of their work is unclear. Their interaction may be associated with the “coat conversion” phenomenon described previously, but what causes this extensive alteration of biochemical property is also unknown.
8
H.-J. WANG ET AL. D. POLLEN HYDRATION
Mature pollen grains are metabolically quiescent and highly desiccated. Hydra tion occurs when pollen grains land on the stigma and/or are incubated in culture medium to trigger a further metabolic program for the next germina tion stage. In plants with a wet stigma, pollen grain hydration is thought to be passive and unregulated, because pollen grains locate in the environment with plentiful liquid—the lipid-rich stigma exudates. Stigma exudates of Nicotiana are essential for pollen hydration and germination. Ablation of the stigma secretory zone and its secretion by cytotoxic transgenic STIG1-barnase plants resulted in stigmaless tobacco, failed to support pollen hydration and resulted in sterility, but application of exogenous stigma exudates, even from Petunia, restored female fertility (Goldman et al., 1994; Wolters-Arts et al., 2002). Certain lipids in stigma exudates of Nicotiana tabacum, especially triacylglycer ides, were able to restore pollen hydration and female fertility of transgenic STIG1-barnase plants (Wolters-Arts et al., 1998, 2002). In contrast to wet-stigma plants, in dry-stigma species, hydration of pollen is more intricate, but accumulating evidence suggests the presence of pollen– stigma discrimination during hydration (Hiscock and Dickinson, 1993; Hulskamp et al., 1995). The stigmatic cuticle and pollen coat are crucial in pollen hydration. The stigma cuticle is a waxy layer with multiple functions. Unlike impermeability in other plant epidermal cells, that of the stigma cuticle, supposed to be modulated by pollination signals, makes pollen hydration accessible. The mentor effect, whereby compatible pollen can trigger hydration and germination of incompatible pollen in mixed popula tions on the stigma, modulates permeability of the stigma cuticle affected by pollination, which may induce a localized change in the cuticle (Hulskamp et al., 1995; Knox et al., 1987; Preuss et al., 1993). In the fiddlehead mutant, the cuticle of other plant epidermal cells changes their water-insulating property, which results in ectopic organ fusion involving interactions between epidermal cells (Lolle et al., 1992, 1997). The FIDDLEHEAD gene encodes a b-ketoacyl CoA synthase necessary for synthesis of longchain lipids (Pruitt et al., 2000; Yephremov et al., 1999). Pollen grains could hydrate and germinate on the surface of non-stigmatic tissues because of the collapse of cuticle integrity in the mutant (Lolle and Cheung, 1993; Lolle et al., 1997). Therefore, mutations disturbing cuticle development and affect ing its permeability might promote pollen hydration on other non-stigmatic tissue. This stigma cuticle is unique and critical in hydrating pollen grains by receiving pollination signals. In the male aspect of pollination, Arabidopsis ECERIFERUM (CER) mutants revealed the importance of lipid metabolism of the pollen coat
POLLEN GERMINATION AND TUBE GROWTH
9
during pollen hydration. These mutants were male sterile and showed an altered waxy cuticle of stem and leaf (Hannoufa et al., 1993; Hulskamp et al., 1995; Jenks et al., 1995; Koornneef et al., 1989; McNevin et al., 1993; Preuss et al., 1993; Rashotte et al., 2004). Among these mutants, cer1, cer3 and cer6 were identified by screening male-sterile genes defective in pollen hydration on the stigma (Hulskamp et al., 1995; Preuss et al., 1993). Pollen grains from these mutants cannot hydrate and germinate on the stigma surface but hydrate and germinate in vitro or in a high-humidity environment. The combination of the different cer mutants did not restore male fertility in mixed pollination experiments, but cer-mutant pollen hydration could be rescued by co-incubation with wild-type pollen (Hulskamp et al., 1995). These data suggest that some pollen–stigma recognition factors may be lost in those mutated pollen grains. All three mutants showed a defective metabolic step in lipid biosynthesis. CER1 encodes a protein that shares high similarity to maize GLOSSY1, which acts as an aldehyde decarbonylase (the mutant accumulates C30 aldehydes and lacks C29 alkanes) (Aarts et al., 1995; Hannoufa et al., 1993; Hansen et al., 1997; Kunst and Samuels, 2003; McNevin et al., 1993). CER3 was previously known as nuclear E3 ubiquitin ligases (At5g02310), but recent evidence showed that it was actually encoded by WAX2/YRE/FLP1 (At5g57800), a gene with unclear function in wax biosynthesis (the mutant is deficient in C29 alkanes, alcohols and ketones) (Garzon et al., 2007; Hannoufa et al., 1996; Jenks et al., 1995; Kunst and Samuels, 2003; Rowland et al., 2007). CER6 encodes a long-chain fatty acid condensing enzyme (the mutant accumulates C24–C26 lipids) (Fiebig et al., 2000; Kunst and Samuels, 2003). Exogenous application of lipids, such as triacylglycerides and mineral oil, can also rescue cer mutants, but the detailed mechanism of how these lipids are transformed and act is still unknown because supply of non-deficient lipids could also rescue the mutated phenotype (Preuss et al., 1993; Wolters-Arts et al., 1998; Zinkl and Preuss, 2000). These findings suggest that the presence of abundant lipids during the pollen–stigma interaction may overcome regulatory bar riers in pollen hydration and the lipids of the pollen coat may form the hydraulic channel to conduct fluid from the stigma to desiccated pollen (Swanson et al., 2004). In addition to lipids, certain proteins in the stigma and pollen coat have key roles in pollen hydration. Cycloheximide-treated stigma elevated the rate of incompatible pollen hydration in Brassica, which suggests that continued protein synthesis is required to regulate pollen hydration (Sarker et al., 1988). Proteomic analysis of the Arabidopsis pollen coat showed that most of the corresponding genes locate in two genomic clusters: one encodes six lipases and the other contains six lipid-binding oleosin genes.
10
H.-J. WANG ET AL.
All six glycine-rich proteins (GRPs) contain a consensus oleosin domain and a glycine-rich unique repetitive motif and are indispensable in pollen hydration; one example is GRP17 (Mayfield and Preuss, 2000; Mayfield et al., 2001). Variation in individual repeat sequences of GRP between species and even ecotypes may evolve to form species barriers. Rapid evolution of the GRP cluster also suggests that members of this protein family are potential candidates for species-specific recognition in the pol len–stigma interaction (Fiebig et al., 2004; Mayfield et al., 2001; Schein et al., 2004). Similar to GRP17, the pollen extracellular matrix (ECM) lipase 4 (EXL4) also participates in pollen hydration on the Arabidopsis stigma: exl4-1 showed a reduced pollen hydration rate, but pollen morphol ogy and fertility were normal. EXL4 functioning in combination with GRP17 to promote the initiation of hydration implies that the oleosin domain of GRP17 may solubilize lipids, making them accessible by EXL4 and other pollen coat lipases. Therefore, changes in lipid composition at the pollen–stigma interface, possibly mediated by EXLs, are essential for efficient hydration (Updegraff et al., 2009). E. POLLEN TUBE GROWTH IN THE STYLE
Pollen tubes growing in the style perceive complex and essential signals sent from the gynoecium for nutrients and guidance cues to complete their journey of double fertilization. In vivo pollen tube growth during plant sexual reproduction can be divided into six stages: (1) stigma penetration, (2) growth in the style and transmitting tissue, (3) emergence from the transmitting tissue, (4) funicular guidance, (5) micropylar gui dance and (6) pollen reception (Johnson and Lord, 2006). Many interac tions occur between the pollen and pistil in this journey, but how the pollen tube receives and interprets the diverse signals within the pistil to reach the embryo sac, the female gametophyte, is still unclear. However, growing evidence suggests that pollen tube guidance is regulated by collaboration between sporophytic (stigma, style and transmitting tissue) and gametophytic (embryo sac) cells of the female tissue. Two major hypotheses have been proposed to explain the guidance of pollen tube growth during pollination: ECM-derived adhesive molecule-based archi tectural constraints and chemotropic guidance cues (Lord, 2003). Genetic (in Arabidopsis) and in vitro tube guidance (in Torenia fournieri) studies suggest that most flower plants probably use both mechanisms to guide pollen tube towards the ovule and that multiple signals sent from the female tissues are required for tube guidance in each stage during pollina tion (Johnson and Lord, 2006).
POLLEN GERMINATION AND TUBE GROWTH
11
In the past few years, several potential chemotropic signalling molecules have been identified from different plant species. Glycosylated tobacco transmitting tissue-specific (TTS) proteins were found effective as attrac tants in promoting pollen tube growth in vitro and attracting pollen tubes in semi-in vivo pollen tube growth medium. Within the style, TTS proteins displayed a gradient of increasing glycosylation from the stigmatic end to the ovarian end of the transmitting tissue, the same direction as pollen tube growth (Wu et al., 1995). This finding indicates that pollen tubes might be able to perceive changes in sugar concentration associated with TTS proteins. The TTS glycoprotein NaTTS from Nicotiana alata also stimulated pollen tube elongation in vitro and attracted pollen tubes in a semi-in vivo pollen tube culture system (Wu et al., 2000). NaTTS and 120K, another pistil arabinogalactan protein in N. alata, provide their C-terminal domain for binding of pollen proteins (NaPCCP and NaSBP1) and might contribute to signalling and trafficking inside pollen tubes growing in planta (Lee et al., 2008, 2009). In an in vitro adhesion bioassay of lily pollination, a stylar pectin was necessary for pollen tube adhesion (Mollet et al., 2000). Besides pectin, polypeptides such as stigma/ style cysteine-rich adhesin (SCA) and chemocyanin in lily have shown promise as chemotropic attractants produced and secreted by the pistil and are involved in adhesion or guidance of the pollen tube (Chae et al., 2007; Kim et al., 2003; Mollet et al., 2000; Park and Lord, 2003; Park et al., 2000). In addition to the ECM components, the navigation of pollen tubes towards ovules and reception of the pollen tube by the embryo sac needs other known signals hidden in the ovule. Higashiyama et al. (Higa shiyama et al., 2001; Okuda et al., 2009) suggested synergid cells of T. fournieri as the source of chemotropic attractants, and recently they found that synergid cells secreted defensin-like LUREs that could attract cultured pollen tubes (Higashiyama et al., 2001; Okuda et al., 2009). Maize ZmEA1 is exclusively expressed in the egg apparatus, and downregulation of ZmEA1 led to ovule sterility caused by loss of close-range pollen tube guidance to the micropyle (Marton et al., 2005). The POP2 gene encodes a transaminase that degrades GABA, whose concentration gradient increases along the pollen path in the pistil and affects pollen tube gui dance when the pop2 mutant appears in both the female and male game tophytes (Ma, 2003; Palanivelu et al., 2003). In addition, GABA localizes in the pistil along the pollen tube growth path, with higher concentration in cells surrounding the micropyle. The pop2 mutant with mutation in a transaminase for normal GABA metabolism exhibited more than 100-fold accumulation of GABA in the style. The high concentration of GABA in the style of the pop2 mutant may disturb the guidance of the pollen tube.
12
H.-J. WANG ET AL.
These findings led to the hypothesis that GABA forms a chemotropic gradient that directs the pollen tube to the micropyle. Several instances of well-characterized genes involved in pollen tube guidance have been reported in Arabidopsis. Pollen tube guidance needs the normal function of the Central Cell Guidance (CCG) gene that encodes a potential transcrip tion factor with an N-terminal conserved zinc b-ribbon domain expressed in the central cell (Chen et al., 2007). Magatama mutants (maa1 and maa3) show a disturbed later step of pollen guidance, as well as wandering and polyspermy phenotypes of pollen tubes (Shimizu and Okada, 2000; Shi mizu et al., 2008). Reception of the pollen tube fails in the feronia (fer)/ sire´ne (sir) mutant, which disrupts pollen tube growth arrest, and the tube continues to grow inside the female gametophyte without releasing sperm cells (Huck et al., 2003; Rotman et al., 2003). Further study revealed FER encoding a kinase-active receptor-like kinase (RLK) and accumulating asymmetrically in the synergid membrane at the filiform apparatus. FER protein might play a key role in the signalling pathway required for successful fertilization (Escobar-Restrepo et al., 2007; McCormick, 2007). The LORELEI gene is expressed in synergid cells before fertilization and encodes a small plant-specific putative GPI-anchored protein that might act in the signalling pathway informing the embryo sac of the arrival of a pollen tube (Capron et al., 2008). Recent study of Arabidopsis abstinence by mutual consent (amc) showed overgrowth of the pollen tube and pre vention of sperm cell discharge only when an amc pollen tube encountered an amc female gametophyte (Boisson-Dernier et al., 2008). Although much progress in understanding the chemotropic molecules has been achieved, how pollen tubes recognize and respond to these cues is still unclear. One hypothesis is that some putative transmembrane receptors localized on the pollen tube tip specifically interact with ECM cues and convert the signals into intracellular responses to change the direction of the tip. Several approaches have been used to identify the promising male determinants involved in this cell–cell recognition during pollination. The first is the genetic approach. Ideally, the pollen tube guidance mutants should have wild-type germination and growth rates but fail to target ovules. Actually, genetic study is a valuable but time-consuming approach to address this question. For example, Johnson et al. (2004), by use of tagging with a pollen-specific marker gene, identified 30 hapless (hap) mutants with abnormal pollen tube growth phenotypes within the ovary. Among the pollen mutants analysed, 10 could extend tubes and grow the length of the pistil; however, these mutants showed reduced ability to target ovules. Future work will be necessary to identify more hap genes to associate gene functions with specific defects in individual guidance steps.
POLLEN GERMINATION AND TUBE GROWTH
13
Another approach to identifying receptors for guidance cues is to study genes expressed by the tube that are homologous to any known and well-characterized receptors. For example, more than 600 receptor-like kinase (RLK) genes are present in the Arabidopsis genome; 10% are pollen expressed and about 90% of these are pollen specific (Honys and Twell, 2003). Members of the RLK gene family have been functionally character ized in tomato. LePRK1 and LePRK2 are pollen-specific RLKs localized to the surface of growing tube tips and interact with each other at this region (Muschietti et al., 1998). Interestingly, this interaction, as well as the phosphorylation of LePRK2, can be disrupted by incubation with a style extract in vitro (Wengier et al., 2003). This finding suggests that LePRKs may be involved in perception of ECM cues expressed by cells along the tube growth path. Three putative ECM ligands interacting with the extra cellular domain of LePRKs are LAT52 (Tang et al., 2002), LeSTIG1 and LeSHY (Tang et al., 2004). Loss of LAT52 function leads to pollen tube growth defects, but LeSTIG1 has been shown to stimulate tube growth in vitro. Tang et al. (2004) hypothesized that the LePRK1/2 dimer switches ligands from LAT52 to LeSTIG1 on contact with the stigma, and this switch is critical for initiation of tube growth. To identify their cytoplasmic partners, Kaothien et al. (2005) revealed a pollen-specific and membraneassociated phosphorylated protein, KPP, whose overexpression leads to defects in tip-localized actin dynamics and results in loss of apical polarity. In addition, KPP interacts with the cytoplasmic domains of LePRK1 and LePRK2 in vitro. All of these features imply that KPP is involved in LePRK-mediated signal transduction events that regulate tube growth. However, it is not yet known whether LePRK1 and/or LePRK2 mediate phosphorylation of KPP in pollen and whether incubation with stigma extracts or LeSTIG1 alters the LePRK–KPP interaction or the phosphor ylation status of KPP. In the past few decades, many histological, cytological and genetic studies have provided plentiful and important information about events and genes involved in pollen adhesion and hydration on the stigmatic papilla cell surface. However, many unanswered questions, such as those related to the molecular metabolism of sporopollenin and exine during pollen formation, regulatory factors participating in signal communication between pollen and stigma during adhesion, and lipid composition and conversion on the pollen surface during hydration, remain to be elucidated. Recent genome-wide studies have revealed many gene expression profiles, and candidate genes involved in pollen germination and tube growth have been accumulated in recent genome-wide studies. Furthermore, the functional characterization of diverse T-DNA insertional mutants has explained the gene networks
14
H.-J. WANG ET AL.
participating in these events. Transcriptomic and proteomic approaches combined with systematic characterization of available mutants defective in pollen adhesion and hydration will be powerful tools to greatly advance our understanding of pollen germination and tube growth in the pistil.
III. ENDOMEMBRANE TRAFFICKING IN THE TIP
REGION CONTRIBUTES TO RAPID AND POLAR
GROWTH
Pollen tubes have eminent growth ability among all plant tissues. For instance, the growth rates of lily and tobacco pollen tubes are approximately 200–300 and 25–100 nm/sec, respectively, in medium-based in vitro germina tion (Chen et al., 2002; Hepler et al., 2001; Hwang et al., 2005; Kost et al., 1998). The fast features are attributed to polarized intracellular organization in the tip region. In the tip of the growing pollen tube, four regions in the pollen tube structure are apex, apical flank, subapical region and shank region. These four regions are discriminated by dynamic and unapparent boundaries according to the distribution of vesicles and organelles. Abun dant membrane exchange, including endocytosis and exocytosis, occurs in the apex of the growing tube. Massive exocytotic vesicles fuse to the extreme apical domain to provide the membrane and cell wall materials, which contribute to tube elongation. Otherwise, the endocytotic vesicles recycle excess membrane resources from the membrane and engulf ECM compo nents, which are secreted from female tissues. Behind the apical region, the subapical region is a passage that contains exo/endocytotic vesicles targeting to the apical growth domain or recycling to organelles. The subapical region is adjacent to the dome of the tip region. In the subapical region, many organelles, including plastid, mitochondria, endoplasmic reticulum (ER) and the Golgi, flow along the tube cortex from the shank, turn around and flow forward to the shank region along the axial of the tube. The bidirectional cytoplasmic streaming maintains the dynamic distribution of the ER and Golgi, which are pivotal organelles to maintain secreted pathways and function as a membrane exchange centre to accept endocytotic vesicles and split exocytotic vesicles, to reach the subapical region of the pollen tube (Cai and Cresti, 2009; Cheung and Wu, 2008). In growing tubes, the apical dome includes the apex and apical flank, where visualized organelles are absent and small vesicles are present, called the clear zone. With this elabo rate structure and integrated organization, the tip of the pollen tube provides the efficient formation of a new membrane and cell wall to the growth domain to achieve rapid tube growth.
POLLEN GERMINATION AND TUBE GROWTH
15
A. INCREASED CELLULAR MODIFICATION AND DIFFERENTIATION IN
GERMINATING POLLEN
Pollen developed within the anther provides a good system for investigating the regulation of several fundamental cellular events such as cell division, cell fate determination, gene regulation, cell–cell communication and cellular differen tiation during pollination (Lord and Russell, 2002). In addition to the mRNAs, proteins, rRNAs and diverse stored bioactive small molecules (Taylor and Hepler, 1997a), carbohydrates and lipids are the major reserves in the mature pollen grain that provide most of the energy required for pollen germination and tube growth (Miki-Hirosige and Nakamura, 1983). Various kinds of sac charides have been identified in different cell compartmentations of mature pollens, including polysaccharides (starch in amyloplasts, fructans in vesicle or cytosol and callose in vesicles), oligosaccharides (sucrose in the cytosol) and monosaccharides (glucose and fructose in the cytosol) (Pacini, 1996). These cytoplasmic saccharides may adjust osmotic pressure (Pacini, 1996) or prevent the plasma membrane from denaturing (Hoekstra et al., 1991) to keep pollen vital for longer periods during pollen dehydration, exposure, dispersal and even pollen germination. Plastids function as a storage compartment of carbohy drate reserves during pollen development and participate in the promotion of pollen tube growth. Cle´ment and Pacini (2001) indicated that in most angios perm species, only proplastids and amyloplasts are found in the pollen, and one or two cycles of amylogenesis (starch synthesis) or amylolysis (starch degrada tion) occur with unknown mechanisms during pollen development. In addition, the occurrence of amylogenesis in the developing microspore or pollen is always preceded by vacuolation, which suggests that starch synthesis or degradation may be involved in the vacuolation process, probably by regulating osmotic pressure in the vacuoles providing or storing osmoticum as glucose monomers. During lily pollen maturation, amyloplast membranes start to disappear gradually, and starch granules are scattered in the cytoplasm, but redifferentia tion of proplastids to amyloplasts is observed in the cytoplasm of lily pollen after 30-min germination and is continuously present in the cytoplasm of elongating pollen tubes (Jiang et al., 2007; Southworth and Dickinson, 1981; Fig. 1). In germinating lily pollen grains, organelles such as amyloplasts, mitochondria and the Golgi apparatus are poorly differentiated in desiccated mature pollen grain, but thereafter they start to differentiate after pollen germination (Fig. 1A, C and E). The Golgi apparatus and round and elongated mitochondria show distinguishable secreted vesicles and cisternae and cristae, respectively. How ever, one of the most obvious changes is the redifferentiation of amyloplasts from proplastids and the accumulation of many starch granules inside. Because a similar phenomenon was found in pollen incubated in medium with pentaer ythritol replacing sucrose, this amylogenesis seems the autonomous biogenesis
16
H.-J. WANG ET AL.
A
B
C
D
E
F
Fig. 1. Organellar differentiation and organization in germinating pollen. Lily (A, C, E) and Arabidopsis (B, D, F) pollen grains germinated in culture medium for 60 min were fixed and embedded in Spurr’s resin for ultrastructural studies. The development of amyloplasts during pollen germination was found in both pollen grains (C, D). Arrows in B–F indicate the tubular endoplasmic reticulum (ER)-like double-membrane structures surrounding the differentiating amyloplasts (C, D) and lipid bodies (E, F). a, amyloplast; g, Golgi apparatus; l, lipid body; m, mitochondria; v, vacuole.
POLLEN GERMINATION AND TUBE GROWTH
17
of amyloplasts during pollen germination (Wang and Jauh, unpublished data). In addition, an interesting phenomenon was found in germinating lily pollen. First, some unique tubular ER-like membrane structures (arrows in Fig. 1C and E; Jiang et al., 2007) were closely associated with and/or enveloped developing amyloplasts and lipid bodies. A similar event was found in germinating Arabi dopsis pollen (Fig. 1B, D and F), and sometimes even more than one layer of tubular ER-like membranes were found outside the amyloplasts and lipid bodies (arrows in Fig. 1D and F) as well as in those in vivo-grown lily pollen tubes (Jiang et al., 2007). These phenomena are similar to autophagosomes found in nutrient-depleted yeast (Levine and Klionsky, 2004; Noda et al., 2002). Autophagy involves the formation of cytosolic vesicles, which contain some compartments of the cytoplasm. These vesicles will further target and fuse with the vacuole, where the compartments of the cytoplasm will be degraded and recycled. During starvation, yeast cells will initiate the autophagy pathway to degrade and recycle cytosolic compartments for survival. The process begins with the formation of the autophagosome, which is a double-membrane vesicle composed of a portion of the cytosol. This vesicle continues to fuse to the vacuole, which contains proteases to break down the autophagosome, thus completing the autophagy process. In addition to its role in nutrient recycling during starvation (Bassham et al., 2006; Moriyasu and Hillmer, 2000), autop hagy may also have important roles in a more general response to various abiotic stresses to assist in cell survival, such as regulation of programmed cell death in pathogen-infected plant cells (Liu et al., 2005; Seay and Dinesh-Kumar, 2005), and oxidative stress (Xiong et al., 2007), and programmed cell death during senescence (Bassham, 2007; Filonova et al., 2000; Gaffal et al., 2007). The source of membranes used for autophagosome formation in different eukaryotic organisms is controversial; examples are those originating from the trans-Golgi network (TGN), provacuolar compartment and ER, as well as de novo synthesized membranes. Although in germinating lily pollen, morphological evidence revealed that the membrane source involved in autophagosome-like compartment formation is tubular ER, provacuoles or TGN-derived vesicles may also be involved in autophagosome formation. To further verify the membrane source, the potential inhibitor of the default secretory pathway, brefeldin A (BFA), was used (Wang and Jauh, unpub lished data). The differentiation of most organelles in lily pollen was not affected by germination in medium containing 1 mg/ml BFA for 1 hr. How ever, the autophagosome-like formation was enhanced by BFA. The differ entiated amyloplasts contained large amounts of starch granules or lipid bodies, and the amyloplast-enclosed lipid body, as well as a portion of cytoplasm, was completely enclosed by the double membrane. The lipid body encircled by tubular ER was not affected by BFA. The ER nature of
18
H.-J. WANG ET AL.
the membrane was confirmed by immunogold labelling with various anti bodies to an ER-localized HSP70 molecular chaperone (BIP), a-amylase and a papain-type proteinase (SH-EP) directly transported to vacuoles from the ER via ER-derived vesicles (Toyooka et al., 2001) in germinating lily pollen. These results indicated that the tubular ER would be the origin of membrane used for autophagosome-like compartment formation. In most eukaryotic cells, autophagy occurs under nutrient stress, but in germinating pollen, it takes place under a normal developmental process. Considering the char acteristics of the survival strategy and rapid events of pollen germination and tube growth, the autophagic machinery is an efficient and fast strategy to degrade reserves for the production of energy and structural components and to ensure successful pollination. However, further investigation of other components involved in autophagic and vacuolar biogenesis such as starch degradation enzymes, other proteases, the source of the tubular membrane structure and the components involved in the formation of the autophagosome-like special organelles are needed to elucidate the mechan ism and function of this autophagic machinery in pollen germination. Interestingly, recent genetic studies of autophagy-related genes in Arabidopsis suggested that autophagy plays a pivotal role in male gametophyte develop ment. On the basis of sequence similarity to proteins involved in yeast autop hagy, a number of Arabidopsis orthologues, AtATGs, with predicted function in various steps in autophagy were functionally and genetically characterized (Bassham, 2007; Inoue et al., 2006). Among all AtATGs, only AtATG6 was essential for pollen tube germination, because other AtATG homozygous mutants were fertile (Fujiki et al., 2007; Qin et al., 2007). The pollen viability of atatg6 showed normal development but male sterility. Harrison-Lowe and Olsen (2008) suggested that the potential defect of atatg6 occurs after micro sporogenesis. Investigating the ultracellular modification of atatg6 to reveal the possible autophagic machinery in regulating pollen germination is of interest. B. EXO/ENDOCYTOTIC VESICLES MAINTAIN POLAR AND RAPID
TUBE ELONGATION IN POLLEN TUBES
Technical advances such as high-resolution microscopy, living cell staining with fluorescent dye and expressing vesicle-targeted fluorescent proteins have allowed for tracing the dynamic events of vesicle trafficking in living tubes. Exocytosis, which involves delivering cytoplasmic membrane-localized pro tein, secreted protein and ECM components synthesized from the ER and Golgi apparatus to cytoplasmic membrane, is a fundamental cellular process for cell expansion, especially in rapid and polar growing pollen tubes. In the past decade, two studies used transformed pollen tubes expressing green fluorescent
POLLEN GERMINATION AND TUBE GROWTH
19
protein (GFP)-labelled secretive proteins, membrane protein and secreted GFP to observe cytoplasmic streaming, a reverse fountain and the inverted coneshaped structure by fluorescence microscopy. In the subapical and apical regions, inverted cone-shaped vesicles were monitored by GFP-tagged cyto plasmic membrane proteins and secreted proteins to show that massive exocy totic vesicles are delivered to the apical membrane of the pollen tube (Cheung et al., 2002; de Graaf et al., 2005). However, the surprising discovery was that the subapical region is the targeting domain of exocytosis instead of apicaltargeted exocytosis. Observing the localization of two cytoplasmic membranetargeting proteins, pectin methylesterase (PME) and Hþ-ATPase, provides evidence of subapical-targeted exocytosis. In tobacco pollen tubes expressing PME–GFP fusion proteins, GFP localization revealed PME localized in the apical region but not in the subapical region (Bosch et al., 2005). Interestingly, no GFP signals were present in the inverted cone, which is filled with massive exocytotic vesicles, so the apical targeting of PME might be through the sub apical region targeting to the membrane domain and undergoing anterograde flow towards the apex. Protein targeting to the cytoplasmic membrane through an inverted cone-mediated and apex-targeting route seems not to explain the occupation of proteins in the apical membrane. This model is also supported by the expression pattern of Hþ-ATPase. Apical region-excluded localization of Hþ-ATPase was observed by immunostaining and GFP-tagged protein (Certal et al., 2008; Lefebvre et al., 2005). About 80% of secreted membrane is recycled by endocytosis (Picton and Steer, 1983). On in-depth assay of endocytosis in the pollen tube, vital styryl dyes FM4-64 and FM1-43, which form a fluorescent structure after they are incorporated into the cytoplasm membrane and sequentially engulfed into endocytotic vesicles, applied to living pollen tubes revealed inverted coneshaped staining in the apical region (Camacho and Malho, 2003; Parton et al., 2001). The inverted cone-shaped staining by FM dye revealed that the endocytotic vesicles are abundant in the apical domain. However, combining FM4-64 staining with ultrastructure observation revealed another distinct endocytosis pathway in the subapical region of the pollen tube (Moscatelli et al., 2007). In this research, the most prevalent endocytosis, with a clathrin dependent manner, was revealed in the subapical region. Sustained staining revealed that the FM-labelled vesicles derived from the subapical membrane are recycled to the secretory pathway through the Golgi apparatus, are distributed in an inverted cone and flow through cytoplasm streaming by a reverse fountain. The dynamic endocytotic vesicle recycling suggests that maintaining the equilibrium of membrane distribution between the cytoplas mic membrane and intracellular vesicles or organelles is efficient for rapid growth of pollen tubes (Fig. 2A).
20
H.-J. WANG ET AL. Subapical regin
Shank
A
Apical Apex flank Retro gra d e
por ns tra t
Ptdlns-PLC
Ptdlns(4,5)P2
Clathrin-coated endocytic vesicle
PtdlnsPK
DAG
Exocytic vesicle
Rab11
Endocytic vesicle
Rab2
Golgi
B
GAP
GDP-Rac-Rho
GDI
PtdlnsPK
GTP-Rac-Rho
Ptdlns(4,5)P2
GEF
C
Villin/gelsolin ADF
Fig. 2.
AIP GTP-Rac-Rho
(Continued)
Actin
Proton flux
Formin
Calcium flux
Ptdlns(4,5)P2
POLLEN GERMINATION AND TUBE GROWTH
21
C. SMALL GTPASE AND ITS REGULATORY COMPONENTS CONTRIBUTE
TO MEMBRANE TRAFFICKING IN POLLEN TUBE POLAR GROWTH
Small GTPases are identified in all eukaryotic cells and are important in various cellular processes. Two small GTPase families, including Rac-Rop and Rab, are identified as regulators of polar growth in pollen tubes. The activity of small GTPases, which is required for downstream signalling, is
Fig. 2. Endomembrane trafficking, small GTPases and actin remodelling regulate pollen tube growth. Three biological events, endomembrane trafficking, regulation of Rac-Rho GTPases and actin remodelling, involved in tube growth are shown. Dynamic vesicle transport through endocytosis and exocytosis in the dome of pollen tubes targets the cell membrane and cell wall materials to the apex and recycles excess membrane, extracellular matrix and the membrane-bound signalling molecule, diacyl glycerol (DAG), to the intracellular compartment (A). The dynamic distribution of phosphatidylinositol (4,5)-bisphosphate (PtdIns[4,5]P2) and its derivative lipid DAG are also shown. PtdIns phospholipase C (PI-PLC) localized at the subapical region and shank of tubes catalyses apex-localized PtdIns[4,5]P2 into DAG and inositol 1,4,5-trisphosphate (Ins[1,4,5]P3). The membrane-bound DAG is generated at the subapical region and then retargeted to the apex by an endomembrane trafficking pathway. Two Rab small GTPases, small vesicle-localized Rab11 and Golgi-localized Rab2, participating in endomembrane transport are also shown. The activity regulation of small Rac-Rho small GTPases, which is mediated by GTPase activating factor (GAP), guanine nucleotide-exchange factor (GEF) and guanine nucleotide dissociation inhibitor (GDI), shows specific distribution (B). The apical membrane-localized GEF promotes exchange of GDP to GTP to activate Rac-Rho GTPase. Activated GTPbound Rac-Rho GTPase associating with the membrane flows to the subapical membrane by retrograde transport and meets GAP. Subapical region-resident GAP enhances intrinsic GTPase activity of Rac-Rho GTPase to change to an inactive GDP-bound form by GTP hydrolysis. Following inactivation of Rac-Rho GTPase in the subapical membrane, the cytoplasmic GDI promotes GDP-bound Rac-Rho GTPase dissociation from the GAP and recycles it to the apical membrane. GDP-bound Rac-Rho GTPase is changed to the GTPbound form by the apical membrane-localized GEF. As mentioned here, GTP-bound RacRho GTPase associates with PtdlnsPK to enhance its activity. Several actin-binding proteins (ABPs) and ionic factors whose activity is to control actin dynamics are shown in (C). The thick and stable actin cables are only in the shank of the pollen tube. In the subapical and apical regions, abundant, fine actin filaments exist and show high dynamic organization. Three ABPs, ADF, villin and gelsolin, which have actin filament severing activity, show different activity regulated by spatial distribution of calcium and proton in the tip of pollen tubes. The proton influx in the apical domain and efflux in the subapical region temporarily creates a slight alkaline banding in the apical region of pollen tubes. The ADF activity to sever actin filaments is increased by association with AIP in a slightly alkaline condition as shown in the visible form. By contrast, low-activity ADF, which is phosphorylated by the Rac-Rho GTPase pathway, associates with apical membrane-localized PtdIns[4,5]P2 and localizes in other regions, whose slightly acidic condition in pollen tubes is shown in the transparent form. The tip-focused calcium gradient is made by calcium influx in the apical domain. The activities of villin and gelsolin, which promote actin severing, are upregulated by a high calcium condition in the tip apex, the activities of the actin-severing factor, ADF, villin and gelsolin, are considered to provide more ends of actin filaments by actin polymerization for fast tube growth. Actin-polymerizing protein, formin, is associated with the membrane domain to polymerize thin actin filaments.
22
H.-J. WANG ET AL.
determined by cycling between the active GTP-bound and inactive GDPbound forms. The GTP binding of GTPases promoting conformational change allows the interaction with effectors to trigger downstream signalling. Small GTPase regulatory proteins, which control small GTPase activity, are of three classes: (1) the guanine nucleotide exchange factors (GEFs) promote exchange of GDP to GTP to activate small GTPases; (2) the GTPase-activating factors (GAPs) inactivate small GTPases by stimulating low intrinsic GTPase activity to convert the GTP-bound to the GDP-bound form; and (3) the guanine nucleotide dissociation inhibitors (GDIs) prevent small GTPases from changing to the active GTP-bound form and keep GEFs away. Plant Rac-Rop GTPases belonging to the eukaryotic Rho GTPase super family of the Rho and Rac family in animals and the Cdc42 family in yeast are especially restricted to the membrane domain to function as a coordina tor in actin cytoskeleton organization and membrane trafficking. Apical membrane-associated Rac-Rop GTPases participating in polar pollen tube growth have been described (Kost et al., 1999; Li et al., 1999). In gain-of-function assays, overexpressing Rac-Rop GTPases or expressing constitutively active (CA) counterparts, which are maintained predomi nantly in an actively GTP-bound state, leads to depolarized pollen tube growth, which suggests that excess activated Rac-Rop GTases cause secre tive vesicles to ectopically fuse to the tip membrane. By contrast, the expres sion of dominant-negative (DN) Rac-Rop GTPases, which preferentially bind GDP and are believed to compete with their wild-type counterparts for endogenous Rho-Rop GTPase interation factors, leads to inhibited tube growth, which suggests that attenuation of DN Rac-Rob GTPases in the pollen tube blocks secretion in the tip region. In addition to Rac-Rob proteins, the regulatory components of Rac-Rop GTPases, such as RhoGAPs, RhoGDIs and RhoGEFs, also critically control polar growth through local restriction and activity regulation of Rac-Rop. Shank regionlocalized tobacco RhoGAP1 (NtRhoGAP1) is a negative regulator of NtRac5 to restrict activated GTP-bound NtRac5 in the apical region to ensure polar growth (Klahre and Kost, 2006). NtRhoGDI2, a negative regulator of Rac-Rho signalling, disassociates NtRac5 from the cytoplasmic membrane to the cytoplasm and forms an NtRhoGDI2–NtRac5 heterodi mer (Klahre et al., 2006). Sequel studies provide evidence suggesting that NtRhoGDI2 internalizing NtRac5 seems to be required for redirecting to the apical domain (Klahre et al., 2006). Moreover, studies of the expression of newly identified Arabidopsis RhoGEFs in tobacco pollen tubes showed RhoGEFs involved in polar tube growth and targeted to the tip region, where Rac-Robs localize (Gu et al., 2006; Zhang and McCormick, 2007). Results of functional studies of RhoGEF, RhoGAP and RhoGDI revealed
POLLEN GERMINATION AND TUBE GROWTH
23
that the spatiotemporal regulation of Rac-Rho GTPase activity and recy cling provide a dynamic Rac-Rho GTPase-mediated polar growth model. The targeting of exocytotic vesicles to the apical domain of the pollen tube relies on active and apical membrane-bound Rac-Rho GTPases. With con tinuous tube growth, retrograde transported Rac-Rho GTPases from the tube apex to the shank region are inactive by shank region-resident RhoGAPs and are internalized into the cytoplasm by RhoGDIs. Internalization of inactive Rac-Rho GTPases allows them to retarget to the apical region, where apical-resident RhoGEFs activate Rac-Rho GTPases (Fig. 2B). Another group of small GTPases involved in membrane trafficking in pollen tubes are Ras-related small GTPases, Rabs. Because Rab GTPases function in vesicle fission or fusion and determine targeting specificity, they are critical for membrane trafficking (Stenmark, 2009). Similar to Rac-Rob GTPase studies, expressing the CA and DN form of Rab GTPases in pollen tubes is effective to address the function of Rab GTPase in pollen tube growth. Two plant Rab GTPases were identified to participate in pollen tube growth by controlling membrane trafficking in two membrane compartments (Fig. 2A). In trans formed tobacco pollen tubes, GFP-fused NtRab2 localized to the Golgi appa ratus. Downregulating NtRab2 activity by overexpressing its DN form inhibited pollen tube growth and significantly accumulated GFP-labelled cargo protein in the secretory system between the ER and Golgi apparatus but did not greatly impair cell surface targeting, which suggests that NtRab2 determines the membrane trafficking route between the ER and Golgi appa ratus (Cheung et al., 2002). Moreover, inverted cone-targeted NtRab11 func tions as a regulator of apex-focused membrane trafficking. Altering activity by overexpressing the CA and DN forms of NtRab11 inhibited exocytic and recycled vesicles in the inverted cone region and compromised the delivery of secretory proteins to the ECM, which underscores that the apex-focused membrane secretion mediated by NtRab11 is important for tube growth (de Graaf et al., 2005). Although only a few plant Rab GTPases have been described, the conserved mechanism of Rab GTPase, which controls the tar geting specificity of membrane trafficking, seems similar. Further functional characterization of plant Rab GTPase will bring more insight into Rab determined vesicle identity for intermembrane targeting. D. ROLES OF PHOSPHOLIPIDS, THEIR DERIVATIVES AND PHOSPHOLIPASE C IN MEMBRANE TRAFFICKING OF ELONGATING POLLEN TUBES
Phosphatidylinositol (4,5)-bisphosphate (PtdIns[4,5]P2) is a secondary mes senger generated by phosphatidylinositol monophosphate kinase (PtdlnsPK) to promote the fusion of secretory vesicles with the plasma membrane in
24
H.-J. WANG ET AL.
animal cells (Cremona and De Camilli, 2001; Hay et al., 1995). However, a similar role of PtdIns[4,5]P2 in exocytosis has not been shown in plants. Early work showing Rac-Rop GTPase regulating PtdlnsPK activity to synthesize PtdIns[4,5]P2 in the apex of pollen tubes seems to correspond to where active Roc-Rop GTPases localize (Kost et al., 1999). Interestingly, PtdIns[4,5]P2 functions as a Rho GDI displacement factor (RhoGDF) to promote disassociation between GDIs and Rho GTPases in the animal system (DerMardirossian and Bokoch, 2005; Faure et al., 1999). Previous study suggesting that NtGDI2 promotes NtRac5 recycling (see previous discussion) implies that PtdIns[4,5]P2 and Rac-Rop GTPases are activated by a positive feedback loop, whereby Rac-Rop GTPases promote an increased level of PtdIns[4,5]P2 by activating PtdlnsPK, and sequential dis association of RhoGDI-bound Rac-Rop GTPases in the cytoplasm allows for freeing Rac-Rop GTPases to retarget to the apical membrane (Fig. 2B). Despite the model presented here, the function of PtdIns[4,5]P2 needs further elucidation. Focusing on the downstream metabolism of PtdIns[4,5]P2, the PtdIns [4,5]P2-specific phospholipase C (PtdIns-PLC) isoforms NtPLC3 and PetPLC were identified in tobacco and Petunia and found to localize in the shank region adjacent to the apical region where PtdIns[4,5]P2 loca lizes in elongating pollen tubes, which suggests that PLCs maintain the spatial restriction of PtdIns[4,5]P2 (Dowd et al., 2006; Helling et al., 2006). Downregulating in vivo NtPLC3 activity in pollen tubes by a specific chemical inhibitor-depolarized pollen tube growth in pollen ger mination caused PtdIns[4,5]P2 to spread to a much larger area of the cytoplasmic membrane at the tip region. This finding suggests that PtdInsPLCs mediating the spatial restriction of PtdIns[4,5]P2 is essential for polar tube growth by a PtdIns[4,5]P2-mediated positive feedback loop of Rac-Rop GTPase activity. Two signalling molecules, diacyl glycerol (DAG) and inositol 1,4,5-trisphosphate (Ins[1,4,5]P3), are generated from PtdIns[4,5]P2 by PtdIns-PLC catalysis. In animal cells, DAG is an important signalling molecule activating the downstream effector, protein kinase C (PKC), to trigger divergent biological processes, such as cell proliferation (Yang and Kazanietz, 2003). However, the signalling activity of DAG and the identification of plant PKC homologues have not been reported in plant cells or in pollen tubes. Use of a specific GFP-labelled marker bound to DAG in vivo revealed that the membrane-integrated DAG specifically localized at the tip region of growing pollen tubes (Helling et al., 2006). With the activity of PtdIns-PLCs decreased by treatment with a chemical inhibitor, the DAG-specific GFP marker was prevented from association with the plasma membrane of the tip region,
POLLEN GERMINATION AND TUBE GROWTH
25
which indicates that DAG accumulation in the tip region requires the activity of PtdIns-PLCs. Presumably, DAG is generated by shank region-targeted PtdIns-PLCs and is expected to be transported away from the tip apex along the retrograde flow of the plasma membrane. Further work supports that endocytosis in the shank region, and vesicles recycling to the apex of the tip are required for DAG targeting to the apical region in growing pollen tubes (Fig. 2A). Thus, DAG abundance precisely restricted at the tip region of the pollen tube seems to be functionally required for polar tube growth. Moreover, overlapping the localization of important signalling molecules, such as Rac-Rop GTPases and their regulatory components, implies that DAG plays an important role in signalling transduction in tube growth. The signalling activity of the second product of PtdIns-PLCs, Ins[1,4,5]P3, was reported in animal cells as a key regulator of Ca2þ influx into the cytoplasm but not in plant cells (Clapham, 1995). Further study in plants will provide more insights into the connection of Ca2þ and these secondary signalling molecules, DAG and Ins[1,4,5]P3. Despite the many identified factors involved in controlling membrane trafficking in polar pollen tube growth, the intercellular signal transduc tion between the female transmitting tract and the pollen tube determin ing tube growth polarity in vivo is still unclear. Remaining questions include how the guiding signal communicates to sustain tube growth polarity and the nature of regulatory factors elicited from female ECM ligands and perceived by ligand-corresponding receptor-like proteins on the pollen tube plasma membrane. As mentioned previously, several RLKs and extracellular guiding factors have been discovered, but the mechanisms underlining these specific interactions remain to be explored. Moreover, uncharacterized receptors or ligands participating in pollen tube guidance may interact with identified partners to trigger down stream responses for determining the direction of tube growth. Clarifying the relationships between receptors and ligands is critical to illuminate polar tube growth in vivo. In addition to the early signal perception, how the perceived signals transduced from RLKs such as Rac-Rop GTPases and PtdIns-PLCs to downstream machineries to regulate tube growth direction by controlling exocytosis is still unclear. Many characterized protein kinases expressed in the pollen tube, including calcium-dependent protein kinases (CDPKs), may relay the signals in the signalling cas cades. Further understanding the mechanism regulating the network among protein kinases and upstream receptors or downstream effectors will help elucidate the integrity of signalling transduction presiding over polar tube growth.
26
H.-J. WANG ET AL.
IV. ROLES OF ACTIN CYTOSKELETON, IONIC
AND REGULATORY PROTEINS IN REGULATION
OF POLLEN TUBE GROWTH
Observations of the subcellular structure in pollen tubes have revealed that an elaborate and dynamic actin cytoskeleton distributes in the cytoplasm, and different actin structures preferentially partition in pollen tubes. To reveal the actin structures in pollen tubes, phalloidin-based fluorescent dyes are commonly used to visualize actin distribution in fixed pollen tubes (Foissner et al., 2002). Because advanced observation of dynamic actin filaments can efficiently and directly provide more information on how actin regulates tube growth, the technique of living cell observation with GFP-labelled actin-binding protein (ABP) to visualize actin has been broadly used. One practical actin indicator in living cells is GFP-fused mouse Talin actin-binding domain (GFP-mTalin). Time-lapsed images obtained from GFP-mTalin-expressed pollen tubes by confocal florescent microscopy revealed the subtle and dynamical actin filaments in the apical region of pollen tubes (Fu et al., 2001; Kost et al., 1998). Similar observa tions occurred with expression of other GFP-labelled ABPs, tobaco actin depolymerized factor 1 (NtADF1) and lily LIM1 (LlLIM1) (Allwood et al., 2002; Wang et al., 2008a). Notably, high expression of the actin indicators in pollen tubes inhibits pollen tube growth and leads to artificial patterns of actin structures resulting from the natural actin binding and remodelling property of these indicators, so the expression dosage should be considerable for observation. However, combining observations of actin structures from fixed and living pollen tubes can provide a more reliable blueprinting of actin structures. In the shank region of growing pollen tubes, long, thick, lowdynamic actin cables longitudinally extend along the shank to reach the base of the reverse fountain, where the organelles move along bidirectional cyto plasm streaming. By contrast, the fine, short, flexible and highly dynamic actin filaments predominantly occupy the apical and subapical region of growing pollen tubes containing massive small vesicles, shown as an inverted cone-shaped region by FM dye labelling (Fig. 2C). The roughly compart mentalized distribution of diverse actin structures in the tip of pollen tubes seems to be tracks for vesicles or organelle delivery, and the diversity of actin structures is due to direct and indirect control by numerous regulatory factors, such as ABPs, Hþ and Ca2þ (see following discussion). In addition to actin filaments, microtubules have been reported to parti cipate in the movement of vegetative nuclei and generative cells in tobacco pollen tubes (Astrom et al., 1995). In vitro glide microtubule assay with newly identified kinesin motor proteins from pollen tubes revealed the
POLLEN GERMINATION AND TUBE GROWTH
27
microtubules also sufficient for short-range and slow movement of the organelles and vesicles (Romagnoli et al., 2003, 2007). However, pollen tube treatment with the microtubule-depolymerized drug oryzalin showed that the microtubule does not significantly affect pollen tube growth but, rather, affects the direction of tube growth, which suggests that microtu bules, at least, are involved in polar growth in an actin filament-coordination manner (Gossot and Geitmann, 2007). Polar pollen tube growth may rely on the interaction of actin filaments and microtubules to form an integral cytoskeleton structure for intracellular membrane trafficking and adjust the direction of growth. A. ACTIN-BINDING PROTEINS CONTROL ACTIN REMODELLING
AND DYNAMICS
To date, numerous articles have comprehensively described that several ABPs control plant development and pollen tube growth by their specific functions in actin structure regulation, such as capping, severing, bundling, branching, nucleation, polymerization and depolymerization (Higaki et al., 2007; Hussey et al., 2006; Ren and Xiang, 2007; Staiger, 2000). The control of actin dynamics by ABPs in pollen tubes is usually coordinated with other regulatory factors, such as Hþ, Ca2þ and PtdIns[4,5]P2, reported as having critical roles for pollen tube growth, to rapidly and precisely respond to extrinsic signals (Foissner et al., 2002; Lovy-Wheeler et al., 2006; Monteiro et al., 2005). Here, we briefly introduce how the interaction of ABPs and signalling factors triggers actin remodelling to promote pollen tube growth. ADF possesses actin filament binding, severing and depolymerizing activ ity. Overexpressing NtADF1 in tobacco pollen tubes severely inhibits pollen tube growth and interrupts actin organization (Allwood et al., 2002). This study also indicated that the binding activity of NtADF1 is decreased when it is phosphorylated through a Rac-Rob GFPase-dependent signalling path way. Biochemical assay showed that a slight alkaline condition enhances the actin-binding activity of NtADF1, which suggests that the intracellular Hþ gradient, which gives rise to an acidic tip and an alkaline subapical region, thus forming a banding region in oscillation (see following discussion), is correlated with the activity of NtADF1. In tobacco pollen tubes coexpres sing NtADF1 and AIP1, an actin-interacting protein that can enhance the actin-depolymerizing activity of NtADF1, predominantly colocalize with fine and short actin filaments in the alkaline subapical region (Allwood et al., 2002; Ketelaar et al., 2004). However, apical membrane-localized PtdIns[4,5]P2 directly binds to NtADF1 and inhibits its activity. These findings suggest that the spatial activity restriction of NtADF1 in the
28
H.-J. WANG ET AL.
alkaline and AIP-presented subapical region increases the dynamics of actin filaments within the region of abundant vesicle transport and prevents the destruction of newly formed actin filaments near the apical membrane (Fig. 2C). Profilin, a monomeric ABP, possesses modulating actin nucleation activ ity and enhances actin polymerization activity. Excess profilins leading to disturbed actin organization and inhibited pollen tube growth suggests that the proper amount of profiling is critical for pollen tube growth (Vidali et al., 2001). Some profiling is sensitive to Ca2þ, which displays a concentration gradient distributed in the tip region in an oscillatory fluctuation manner; thus, the spatiotemporal regulation of profilin activity seems to persist in tube growth (Kovar et al., 2000). The versatile ABPs, villins/gelsolins, are actin severing, bundling, capping and nucleating proteins. Two villins/gelsolins were identified in lily and reported as actin regulators in pollen tube growth. Overexpression of ABP29 in lily pollen abolishes the actin structure. Biochemical assay with recombinant ABP29 protein revealed that it accelerates actin nucleation, blocks barbed ends and severs actin filaments in a Ca2þ- and PtdIns[4,5] P2-regulated manner (Xiang et al., 2007). In addition, ABP41 binding to actin filaments is Ca2þ dependent, and use of a specific antibody to block ABP41 activity in the pollen tube leads to inhibited pollen tube growth (Fan et al., 2004). Capping proteins (CPs) bind to the growing end of actin filaments and modulate polymerization. Interestingly, Arabidopsis AtCP1 possesses a reg ulating capping activity that is inhibited by phosphatidic acid (PA), a cata lytic product of phospholipase D (PLD), to result in promoted actin polymerization (Huang et al., 2006; Meijer and Munnik, 2003; Wang, 2005). Moreover, changing PLD activity or PA level inhibits tube growth and destruction of actin filaments (Monteiro et al., 2005; Potocky et al., 2003). This finding suggests that CPs may be an effecter involved in PA- or PLD-mediated actin polymerization. Recently, a novel plant ABP, LIM, identified from tobacco functioned as actin-bundling proteins to stabilize actin filaments against depolymerization (Thomas et al., 2006). A lily pollen-enriched LIM, LlLIM1, revealed that LIMs participate in pollen tube growth by stabilizing actin filaments simul taneously regulated by Ca2þ and Hþ (Wang et al., 2008a). Overexpressing LlLIM1 in the pollen tube severely inhibits tube growth, and mild expression of GFP-labelled LlLIM1 in pollen tubes has a moderate effect on tube growth and leads to formation of an asterisk-shaped, thick actin structure and abnormal vesicle aggregation in the subapical region of the tip. This finding suggests that overstable actin filaments without plasticity and
POLLEN GERMINATION AND TUBE GROWTH
29
flexibility in the tip region cause blocked tip-focused exocytosis and inhibit tube growth. The actin-nucleating protein, formin, stimulates de novo actin assembly to produce actin filaments from the cell membrane. Arabidopsis formin homo logue 1 (AFH1) ectopically expressed in tobacco pollen tubes promotes the formation of short actin cables adjacent to the cell membrane, which suggests that it stimulates actin assembly along the cell periphery (Cheung and Wu, 2004). Moderately expressing AFH1 in pollen tubes slightly enhances tube growth, but overexpressing AFH1 leads to depolarized tube growth. Although ectopic expression studies have revealed the possible role of formins in tube growth, further study with pollen-expressed formin homologues is necessary to elucidate its native function during pollen tube growth. B. ACTIN-ASSOCIATED MOTOR PROTEINS MEDIATE VESICLES OR
ORGANELLE MOVEMENT
The myosin superfamily, actin-based molecular motor proteins, exists in all eukaryotic cells and functions as mobile components with mechanical mov ing properties along actin cables by consuming ATP. The Arabidopsis gen ome contains 17 myosin-like genes classified into two classes: 4 myosin VIII genes and 13 myosin XI genes (Reddy and Day, 2001). Class VIII myosins predictably associate with the cell membrane and plasmadesmata. More over, class XI myosins, which are similar to animal and fungal myosin V, are likely involved in movement of organelles, including chloroplasts, mitochon dria, ER, peroxisome and the Golgi apparatus (Liebe and Menzel, 1995; Nebenfuhr et al., 1999; Van Gestel et al., 2002). Because C-terminal tails of myosin are considered to interact with organelles and determine the specifi city of cargo, the association of the yellow fluorescent protein (YFP)-fused C terminus of myosins and organelle markers has been examined to reveal the cargo specificity of class XI myosins (Reisen and Hanson, 2007). The mitochondria-, Golgi apparatus- and small unidentified organelle-localized YFPs indicate the presence of the cargo specificity of class XI myosins. In addition, the Arabidopsis mutants mya2-1 and mya2-1, which disrupt the gene encoding one member of the class XI myosins, show abolished vesicle movement and polar auxin transport; thus, class XI myosins may also be involved in small vesicle trafficking (Holweg and Nick, 2004). Despite numerous myosin-related functional studies in plants, the direct functional characterization of myosins in pollen tubes is absent. Limited studies invol ving antibodies against animal myosin V to reveal the localization of plant class XI myosins have indicated that class XI myosins associate with
30
H.-J. WANG ET AL.
organelles and vesicles in pollen tubes (Miller et al., 1995). To learn more about the function of myosin in pollen tube growth, identifying pollenexpressed myosins and further examining their function with pollen tubes used as material are needed. Since the diverse activities of most ABPs present in the tip region can be controlled by protein modification and chemical compounds such as Hþ, Ca2þ and PtdIns[4,5]P2, the spatiotemporal distribution of these ABPs and regulatory chemicals are critical to sustain tube growth. For example, the tip-localized ADFs possessing actin-depolymerized activity are antagonistic to actin-stabilized proteins, LIMs, which are present in all tubes, and both participate in tube growth because disturbing the activity of each leads to growth arrest. Therefore, the activity of co-presented ABPs should be con trolled intermittently and distributively by regulatory molecules. PtdIns[4,5] P2 is localized in the tip membrane, and the gradient distribution of Hþ and Ca2þ occurs periodically in the tip region. Thus, the precise layouts of regulatory molecules can explain how the diverse ABPs perform their func tions in the tip region without conflict. A question is how the pollen tube establishes the periodically tip-focused Hþ and Ca2þ gradients (see following discussion). Another important cytoskeletal component not fully discussed here is the function of microtubules and microtubule-associated proteins (MAPs) in pollen tube elongation. In plant cells, dynamic microtubules form four distinct assemblies in the plant cell cycle to guide Golgi-derived vesicles to a site where they will fuse to form the new cell plate that separates the daughter cells. The four assemblies are the interphase cortical array involved in cell expansion, the preprophase band of microtubules determining the plane of cell division, the spindle involved in daughter chromosome separa tion and the formation of phragmoplast in late anaphase (Cai, 2010). The organization and functions of actin microfilaments in regulating pollen tube elongation have been described in great detail; however, the precise functions and mechanisms of microtubule/MAPs in pollen tube elongation remain unknown. From studies of several microtubule-disturbing agents, such as colchicine and oryzalin, microtubules were suggested to have a lesser role during in vitro pollen tube growth (Cai and Cresti, 2009 and cited refer ences). Nevertheless, accumulating evidence suggests that microtubules have a role in organelle movement and, interestingly, may functionally cooperate with actin microfilaments through uncharacterized ABPs or MAPs to con trol the trafficking of organelles and secreted vesicles during pollen tube elongation (Cai and Cresti, 2009). This putative functional cooperation of two important cytoskeletal components may play more important role in vivo, where pollen tube elongation is much faster than in vitro. Functional
POLLEN GERMINATION AND TUBE GROWTH
31
identification and investigation of the essential components involved in this hypothesis, such as the putative proteins mediating the interaction between actin microfilaments and microtubules and the posttranslational modifica tion of microtubules, will be important to reveal the mechanisms of the cytoskeleton network during pollen tube elongation.
V. IONIC AND PROTEIN REGULATORS CONTRIBUTE TO OSCILLATORY POLLEN TUBE GROWTH Pollen tubes possessing a non-linear and quasi-sinusoidal growth rate repre sented as a universal biological event in different plant species reveal a general and oscillatory growth pattern, with periods of 20–90 sec and ampli tudes of three- to fourfold difference between two top peaks of growth rate (Feijo et al., 2001; Parton et al., 2003). Behind the oscillatory growth rate referred as the major output, numerous cellular events within the apical or subapical region also have phase-shifted oscillation patterns with same period of growth rate as individual circuits. Example activities include dynamically changed structure of actin filaments, ionic level and ion influx; metabolic activities; endomembrane trafficking; and cell wall formation. However, because disruption of these individual circuits can perturb oscillatory growth and other circuitous events, these circuits are shown to have hierarchic regulatory relationships in a feedback loop persisting in oscillatory tube growth. Three biological events, cell wall formation, apical membrane secretion and metabolic activity, which intuitively participate in pollen tube growth, show oscillatory patterns that mirror growth rate in pollen tube growth. Cell wall formation directly occurring in the tube apex contributing to tube elongation was discovered to be oscillatory by measuring the change in cell wall thickness in the growing tip of lily (McKenna et al., 2009). During oscillatory growth, the thickness of the apical cell wall preceding the peak growth rate suggests that fast tube growth after abundant vesicle secretion via an apex-targeted exocytic pathway occurs in each period of the growth rhythm. Similar observations with use of FM dye to measure the abundance of vesicles in inverted cone regions reflects oscillatory secretion (Parton et al., 2001). However, the congregate of mitochondria at the subapical region accompanying the peak metabolic activity of NAD(P)H dehydrogen ase, which corresponds to an ATP synthetic reaction, anticipates growth maximum by 5–10 sec (Cardenas et al., 2006). Both circuits, generated by energy and incorporating the cell wall/membrane to the tip region, are referred to as fundamental processes of tube growth and show frame-shifted
32
H.-J. WANG ET AL.
patterns to that of growth oscillation, which implies that a predictable and complicated signalling network mediates the regulation of different biologi cal events. Two ionic factors, Ca2þ and Hþ, exhibit greatly changing and oscillatory gradients in the tip of growing tubes and are considered to have pivotal roles in regulation of biological events such as actin remodelling. The tip-focused [Ca2þ] gradient, observed only in growing pollen tubes, exhibits a steep decrease in level from the apex to the subapical region along the axis of pollen tubes. During tube oscillatory growth, the change in [Ca2þ] level in the extreme apex can be as much as fourfold, with a range between 700 and more than 3000 nM (Holdaway-Clarke et al., 1997; Pierson et al., 1996). The [Ca2þ] gradient seems to be maintained by Ca2þ influx at the pollen tube tip membrane through putative apical strength-activated calcium channels (Dutta and Robinson, 2004). In oscillatory pollen growth, the maximal subcellular [Ca2þ] level at the tip is the same as the phase of growth rate. However, the maximal influx of [Ca2þ] at the pollen tube apex follows the growth peak and lags by about 11 seconds, which suggests that fast tube growth might change the membrane strength to increase the influx of [Ca2þ] by activating putative apical strength-activated calcium channels. However, Arabidopsis mutants defective in a cell membrane-associated Ca2þ-ATPase are male deficient and show decreased seed setting and reduced pollen growth rates in vitro, which suggests that Ca2þ-ATPase might be required for forming a tip-focused Ca2þ gradient (Schiott et al., 2004). The down stream signalling proteins of Ca2þ involved in pollen tube growth, such as calmodulin-like protein (CML) and CDPKs, have been identified, which reflects the importance of Ca2þ (Myers et al., 2009; Yoon et al., 2006; Zhou et al., 2009). In Arabidopsis, a functional survey of pollen-specific calcium regulatory proteins reveals that one CML, three CDPKs and three calci neurin B-like (CBL) proteins participate in pollen tube growth. The tobacco pollen tubes overexpressing cytoplasm-localized Arabidopsis CDPK32 (AtCDPK32) show reduced length and significantly swollen tips of growing tubes. However, perturbation of a Petunia plasma membrane-localized CDPK also causes depolarization of pollen tubes (Yoon et al., 2006). These studies imply tip-focused Ca2þ regulation of polar tube growth mediated by cytoplasmic CDPK. Recently, two plasma membrane-localized Arabidopsis CDPKs, AtCDPK34 and AtCDPK17, were reported to be critical regulators of tube growth. AtCDPK34 and AtCDPK17 double mutants show reduced growth rate and failed fertilization. Interestingly, the double mutant harboring a calcium-insensitive CDPK34 transgene can not recover fertility, which supports a mechanistic model that AtCDPK17 and AtCDPK34 induce Ca2þ signals to increase the rate of pollen tube tip
POLLEN GERMINATION AND TUBE GROWTH
33
growth (Myers et al., 2009). Otherwise, except for the calcium-related signal ling molecules mentioned above, several ABPs can be directly regulated by calcium to modulate actin-binding activity (see previous discussion). Calcium-sensitive profilin and villins/gelsolins regulating actin dynamics might be critical for oscillatory tube elongation by rhythmic [Ca2þ] in the tip of pollen tubes. The tip-focused Hþ gradient in the pollen tube also exhibits oscillatory change. During tube growth, a slight alkaline region emerges around the base of the clear zone and anticipates the fast growth peak of oscillatory tube growth (Feijo et al., 1999; Lovy-Wheeler et al., 2006). The polarized cytoplasmic Hþ gradient is perhaps maintained by Hþ influx at the apex and efflux along the subapical membrane by membrane-localized Hþ-ATPase. Modifying tip-focused Hþ gradient by chemical processes, such as acidification and alkalization, leads to reorganization of the actin cytoskeleton, especially in the apical domain, which suggests that maintain ing oscillatory Hþ gradient is critical for tube growth. In pollen tubes with application of sodium acetate, chemical-induced acidification leading to destruction of actin fringes in alkaline region seems to be due to the reduction of ADF/AIP activity (see previous discussion). Thus, a proposed model supposes that the increasing pH in the subapical alkaline region stimulates the fragmenting activity of ADF/AIP, which in turn generates more ends for actin polymerization to support faster growth. In addition to ADF, another ABP, LlLIM1, preferably bound and stabilized to actin filaments in acidic and low Ca2þ conditions, might participate in the region with oscillatory and marked change in Ca2þ and Hþ levels. With the converging activity of ion-regulated ABPs and ionic oscillations in the tip region, activated ADF/AIP, located in the alkaline band, generates fringe actin filaments for increasing actin polymerization, which in turn supports faster growth rates and the beginning of proton and calcium influx. This situation inactivates ADF/AIP, activates LIM, decreases actin polymeriza tion and retards growth. With maximal accumulation of [Ca2þ] after tube growth retardation and concomitant increase and stabilization of pH, the activity of LIM decreases and ADF/AIP increases to repeat the cycle of events (Fig. 3). The activity of Rac-Rop signalling proteins to regulate apical actin assembly shows the same frequency as growth oscillation, leads to maximal growth rate and exhibits a similar phase with apical actin filaments, but apparently ahead of tip-localized Ca2þ, as revealed by measuring the inten sity of expressing GFP-tagged mTalin and Arabidopsis RIC4 (AtRIC4) in pollen tubes (Hwang et al., 2005). RICs, a small family of Rac-Rop targets, are localized in the tip membrane or cytoplasm and participate in polarity
34
H.-J. WANG ET AL.
A
Affinity of L|LIM1 binding to F-actin High Medium Low
pH
7.5
6.5
F-actin filaments/cables
B
2+ [Ca ]
3 µM
100 nM
C Efficiency of vesicle transport High Medium Low
Golgi apparatuses/secreted vesicles
Growth rate 2+ [Ca ]c in the apical zone pH in the apical zone Golgi apparatuses/secreted vesicles in apical zone L|LIM1 strong binding to F-actin 25–70s
Golgi apparatuses/ pH [Ca2+] secreted vesicles 3 µM 7.5 High
(µm/s)
Growth rate 0.15
6.5
0.05
Fig. 3.
(Continued)
100 nM
Low
POLLEN GERMINATION AND TUBE GROWTH
35
of tube growth mediated by positive- and negative-regulated actin polymer ization (Gu et al., 2005; Wu et al., 2001). AtRIC4, which promotes apical actin polymerization, and another member of the RIC family, AtRIC3, have adverse activity in enhancing actin filament disassembly in the apex by enhancing intracellular [Ca2þ] level. Because versatile Ca2þ regulates the activity of some ABPs, CAMs and CDPKs, the circuit of Rac-Rop activity seems to be a pivotal role to control polar growth and actin dynamics by modulating RIC4-dependent calcium accumulation in the tip region of pollen tubes. Oscillatory Rac-Rop activity seems to be the central circuit that governs and integrates other individual circuits by regulating the tip-focused Ca2þ level. However, the tip-focused oscillatory Hþ level is not involved in this Rac-Rop-dominant regulatory network. Hþ influx or efflux mediated by tip-localized ATPases may be controlled by the strength of the tip mem brane, so the control of the Hþ level should be regulated by other circuits that contribute to tube growth and are dominantly regulated by Rac-Rop activity. However, the activity of ATPases might be regulated by a mechanism such as ATPase-binding proteins, independent of Rac-Rop activity, and play coordinate roles with Rac-Rop activity to control tube growth.
Fig. 3. Oscillatory calcium and proton gradients coupled with actin dynamics regulate oscillatory tube growth. The special and temporal changes of oscillatory events in growing pollen tubes, such as actin dynamics, calcium and proton gradients and vesicle transport, potentially linked with each other, are shown. In (A) and (B), the oscillatory proton gradient and the calcium gradient couple with actin dynamics regulated by LlLIM1, an actin-binding protein regulated simultaneously by proton and calcium in three time points, reflecting slow growth, moderate growth preceding fast growth and fast growth. The vesicle transport level also has an oscillatory pattern shown in (C). (D) shows the curves of growth rate, apical pH, apical calcium level and the apical level of Golgi apparatuses or secreted vesicles corresponding to (A), (B) and (C). At the time of slow growth, the Golgi apparatuses or secreted vesicle accumulation mediated by LlLIM1 strongly binding to actin to promote actin bundling is controlled by low apical pH and low apical calcium level. After slow growth, accumulating vesicles start to be targeted to the apical membrane domain to promote tube elongation and trigger calcium import by activating strength-gating channels. At the same time, the increase in apical calcium level and subapical pH downregulates LlLIM1-binding activity and promotes actinsevering activity of ADF1 to generate fine actin filaments for further fast growth. Abundant vesicles targeted to the apex drive the fast tube growth rate, which persists by the formation of many long, fine actin filaments by formin polymerization. By fast growing, large proton and calcium imports are activated by strength-gating channels to form the acidic and high calcium condition at the tip region. After abundant exocytosis, decreased rate of vesicle transport leads to slow growth. Low pH value and high calcium level following growth slowing enhances LlLIM1-binding activity to stabilize actin filaments and enter another period of oscillatory growth.
36
H.-J. WANG ET AL.
VI. GLOBAL ANALYSIS OF GENE EXPRESSION
IN POLLEN TUBES
The activity of preexisting and newly synthesized transcripts and proteins during pollen germination and tube growth has been demonstrated; how ever, recent proteomic and transcriptomic studies highlight the global mole cular information of important proteins and genes involved in pollen germination and tube growth. For example, 322 unique proteins were identified from mature pollen of Oryza sativa via the proteomic approach, and most were first identified in pollen (Dai et al., 2006). Further compar ison of the protein profiles of mature and germinating rice pollen revealed 186 differentially expressed proteins, 66 with development-stage specificity and 120 with expression level changes (Dai et al., 2007). Proteome mapping of Arabidopsis mature pollen showed more than 100 proteins differentially expressed in pollen (Holmes-Davis et al., 2005; Noir et al., 2005). Similar results were obtained by comparing the protein profiles of pollen and germinating pollen tubes of gymnosperms, such as Pinus strobus (Fernando, 2005), Picea meyeri (Chen et al., 2006) and (Wu et al., 2008) Pinus bungeana. Large-scale transcriptome analysis of Arabidopsis pollen showed reduced complexity and a unique composition with greater proportions of selectively expressed (11%) and enriched (26%) genes than those in vegetative tissues examined (Pina et al., 2005). Interestingly, genome-wide comparison of developing Arabidopsis microspores in bicellular and tricellular stages revealed a decline in the proportion of diverse mRNA species but an increase in the proportion of male gametophyte-specific transcripts (Honys and Twell, 2004). In addition, transcriptomic analysis revealing the composition of transcripts with levels changed from desiccated mature pollen to hydrated pollen and to growing pollen tubes strongly suggested that de novo initiation of the transcriptional program occurs at this stage (Wang et al., 2008b). Recently, Borges et al. (2008) found the transcriptome in sperm cells with distinct and diverse transcriptional profiles as compared with sporophytic tissues and even pollen. Gene expression patterns affected by cold stress were also characterized by serial analysis of gene expression (Lee and Lee, 2003). Transcriptome analysis results are also available for pollen of soybean (Glycine max) and maize (Haerizadeh et al., 2009; Ma et al., 2008). Results from global expression analysis of pollen are consistent with the message that every factor necessary for pollen germination might be ready and the activity is undergoing when this program is turned on. After germi nation, another program is executed to promote tube growth. Although all of this knowledge improves our understanding of differentially expressed
POLLEN GERMINATION AND TUBE GROWTH
37
genes and proteins associated with pollen and provides insights into the molecular programs that separate the germination and growth of pollen tubes from mature pollen grains, the targets of those researches focus on pollen incubated in medium (in vitro-grown pollen tube). The limitations of in vitro media are well known: pollen tubes never grow as fast or as long in vitro as they do in vivo (Taylor and Hepler, 1997b). Molecular information about in vivo-grown pollen tubes is not covered by the global analyses we describe. As well, all of the known molecules generated by pistils, which can adhere to and guide the pollen tube, do not exist in common germination medium; therefore some communication between pollen and pistil will not be reflected in the profiles. Even some existing unknown interactions occur ring in vivo might complicate gene expression profiles. Some genes might not turn on or off or even increase or decrease their quantities in the inappropri ate environment without intimate mating dialogue. In other words, the true story must be uncovered under the real in vivo condition. Almost all pollen-expressed genes described in the literature were revealed by in vitro or semi-in vivo conditions, and little information is available for in vivo-grown pollen tubes. Taking the advantage of the hollow style of Lilium longiflorum (Thunb.), we collected in vivo-grown pollen tubes from cut-opened styles and established a transcriptomic database of transcripts enriched and/or specifically expressed in the in vivo-grown pollen tube after suppression subtractive hybridization screening (Huang and Jauh, unpub lished data). We found transcripts specifically or highly expressed in the in vivo-grown pollen tubes. Functional assay revealed that some transcripts could promote pollen tube growth rate in vivo, and further exploration of in vivo pollen tube-enriched or pollen tube-specific genes are in progress. By combining semi-in vivo system and microarray analysis approaches, Qin et al. (2009) provided convenience data that with the assistance from pistil, pollen tubes receive competent of guidance cues by altering the expression profiles of genes involved in signal transduction, transcription and pollen tube growth. Functional characterization of these promising candidates will elucidate the elegant molecular regulatory network of how pollen tube navigates through the pistil and optimizes its elongation for successful double fertilization.
VII. CONCLUSIONS For a long time, pollen germination, tube elongation and double fertiliza tion have been fascinating research topics for plant biologists. Reception or rejection of pollen by the female stigma is an evolutionary strategy that
38
H.-J. WANG ET AL.
plants have adapted to ensure successful sexual reproduction, species pre servation and biodiversity generation (Cheung et al., 2010). Important molecular and cellular mechanisms involved in these events include diverse signalling, cellular differentiation and endomembrane trafficking of pollen germination and tube growth, as well as the regulatory elements that control male germ-line identity and synthesis of male molecules involved in the interaction with the female partner during fertilization. Early histological and cytological studies of pollen germination and pollination events, com bined with functional characterization of mutants defective in these events, have established the foundation of the current biology of this area. How ever, in this post-genomic era, we now have an even better opportunity to explore these important research topics. Arabidopsis has been widely used as a model plant organism because of its advantages and the availability of valuable resources for basic research into plant genetics and molecular biology. The resources include a highly annotated genome, comprehensive genetic and transcriptomic data, readily available T-DNA insertional mutants and newly developed tools for cell biology study. In addition, recent advances in the development of technologies in molecular and imaging tools have provided a great opportunity to explore the temporal and spatial aspects of pollen germination and pollination. By taking advan tage of the improved imaging power of intravital two-photon excitation microscopy in deeply penetrating plant tissues, plant biologists can precisely track the pollination event in real time without rupturing tissue (Cheung et al., 2010). This innovatory technology, combined with stable (e.g. transgenic plants) or transient (e.g. particle bombardment) transforma tion approaches as well as other cell biology tools (e.g. GFP-tagged pro teins), is extremely important to functionally investigate the genes involved in diverse cellular aspects such as cytoskeleton-mediated cytoplasmic streaming and endomembrane trafficking, growth guidance and the hypothetical functional cooperation between actin microfilaments and microtubules in in vivo-grown pollen tubes. Despite immense efforts during the past few decades to understand the genes involved in these processes, considerable information is still needed to elucidate the full molecular mechanisms of fertilization, aspects that are crucial for plant breeding. However, the availability of the advanced tools and resources opens a new era for genome-wide and systematic investigation of the genes and mechanisms regulating pollen germination, tube growth and double fertilization. A good understanding of these processes provides critical information that will contribute greatly to plant breeding and will catalyse further translational innovations within the life sciences.
POLLEN GERMINATION AND TUBE GROWTH
39
ACKNOWLEDGEMENTS
This work was supported by research grants from Academia Sinica (Taiwan), the National Science Council (96-2311-B-001-023-MY3 and 98-2313-B-001-001-MY3, Taiwan) and the Li Foundation (USA) to G.Y. Jauh.
REFERENCES Aarts, M.G., Hodge, R., Kalantidis, K., Florack, D., Wilson, Z.A., Mulligan, B.J., et al., 1997. The Arabidopsis MALE STERILITY 2 protein shares similarity with reductases in elongation/condensation complexes. Plant J. 12, 615–623. Aarts, M.G., Keijzer, C.J., Stiekema, W.J., Pereira, A., 1995. Molecular character ization of the CER1 gene of Arabidopsis involved in epicuticular wax biosynthesis and pollen fertility. Plant Cell 7, 2115–2127. Allwood, E.G., Anthony, R.G., Smertenko, A.P., Reichelt, S., Drobak, B.K., Doonan, J.H., et al., 2002. Regulation of the pollen-specific actindepolymerizing factor LlADF1. Plant Cell 14, 2915–2927. Ariizumi, T., Hatakeyama, K., Hinata, K., Inatsugi, R., Nishida, I., Sato, S., et al., 2004. Disruption of the novel plant protein NEF1 affects lipid accumula tion in the plastids of the tapetum and exine formation of pollen, resulting in male sterility in Arabidopsis thaliana. Plant J. 39, 170–181. Ariizumi, T., Hatakeyama, K., Hinata, K., Sato, S., Kato, T., Tabata, S., et al., 2003. A novel male-sterile mutant of Arabidopsis thaliana, faceless pollen-1, pro duces pollen with a smooth surface and an acetolysis-sensitive exine. Plant Mol. Biol. 53, 107–116. Ariizumi, T., Kawanabe, T., Hatakeyama, K., Sato, S., Kato, T., Tabata, S., et al., 2008. Ultrastructural characterization of exine development of the transient defective exine 1 mutant suggests the existence of a factor involved in constructing reticulate exine architecture from sporopollenin aggregates. Plant Cell Physiol. 49, 58–67. Astrom, H., Sorri, O., Raudaskoski, M., 1995. Role of microtubules in the movement of the vegetative nucleus and generative cell in tobacco pollen tubes. Sex. Plant Reprod. 8, 61–69. Bassham, D.C., 2007. Plant autophagy—more than a starvation response. Curr. Opin. Plant Biol. 10, 587–593. Bassham, D.C., Laporte, M., Marty, F., Moriyasu, Y., Ohsumi, Y., Olsen, L.J., et al., 2006. Autophagy in development and stress responses of plants. Autophagy 2, 2–11. Blackmore, S., Wortley, A.H., Skvarla, J.J., Rowley, J.R., 2007. Pollen wall devel opment in flowering plants. New Phytol. 174, 483–498. Boisson-Dernier, A., Frietsch, S., Kim, T.H., Dizon, M.B., Schroeder, J.I., 2008. The peroxin loss-of-function mutation abstinence by mutual consent disrupts male-female gametophyte recognition. Curr. Biol. 18, 63–68. Borg, M., Brownfield, L., Twell, D., 2009. Male gametophyte development: a mole cular perspective. J. Exp. Bot. 60, 1465–1478. Borges, F., Gomes, G., Gardner, R., Moreno, N., McCormick, S., Feijo, J.A., et al., 2008. Comparative transcriptomics of Arabidopsis sperm cells. Plant Phy siol. 148, 1168–1181.
40
H.-J. WANG ET AL.
Bosch, M., Cheung, A.Y., Hepler, P.K., 2005. Pectin methylesterase, a regulator of pollen tube growth. Plant Physiol. 138, 1334–1346. Cai, G., 2010. Assembly and disassembly of plant microtubules: tubulin modifica tions and binding to MAPs. J. Exp. Bot. 61, 623–626. Cai, G., Cresti, M., 2009. Organelle motility in the pollen tube: a tale of 20 year. J. Exp. Bot. 60, 495–508 Camacho, L., Malho, R., 2003. Endo/exocytosis in the pollen tube apex is differen tially regulated by Ca2þ and GTPases. J. Exp. Bot. 54, 83–92. Capron, A., Gourgues, M., Neiva, L.S., Faure, J.E., Berger, F., Pagnussat, G., et al., 2008. Maternal control of male-gamete delivery in Arabidopsis involves a putative GPI-anchored protein encoded by the LORELEI gene. Plant Cell 20, 3038–3049. Cardenas, L., McKenna, S.T., Kunkel, J.G., Hepler, P.K., 2006. NAD(P)H oscil lates in pollen tubes and is correlated with tip growth. Plant Physiol. 142, 1460–1468. Certal, A.C., Almeida, R.B., Carvalho, L.M., Wong, E., Moreno, N., Michard, E., et al., 2008. Exclusion of a proton ATPase from the apical membrane is associated with cell polarity and tip growth in Nicotiana tabacum pollen tubes. Plant Cell 20, 614–634. Chae, K., Zhang, K., Zhang, L., Morikis, D., Kim, S.T., Mollet, J.C., et al., 2007. Two SCA (stigma/style cysteine-rich adhesin) isoforms show structural differences that correlate with their levels of in vitro pollen tube adhesion activity. J. Biol. Chem. 282, 33845–33858. Chen, Y., Chen, T., Shen, S., Zheng, M., Guo, Y., Lin, J., et al., 2006. Differential display proteomic analysis of Picea meyeri pollen germination and pollentube growth after inhibition of actin polymerization by latrunculin B. Plant J. 47, 174–195. Chen, Y.H., Li, H.J., Shi, D.Q., Yuan, L., Liu, J., Sreenivasan, R., et al., 2007. The central cell plays a critical role in pollen tube guidance in Arabidopsis. Plant Cell 19, 3563–3577. Chen, Y.F., Matsubayashi, Y., Sakagami, Y., 2000. Peptide growth factor phytosul fokine-a contributes to the pollen population effect. Planta 211, 752–755. Chen, C.Y., Wong, E.I., Vidali, L., Estavillo, A., Hepler, P.K., Wu, H.M., et al., 2002. The regulation of actin organization by actin-depolymerizing factor in elongating pollen tubes. Plant Cell 14, 2175–2190. Cheung, A.Y., Boavida, L.C., Aggarwal, M., Wu, H.M., Feijo´, J.A., 2010. The pollen tube journey in the pistil and imaging the in vivo process by two-photon microscopy. J. Exp. Bot. 61, 1907–1915. Cheung, A.Y., Chen, C.Y., Glaven, R.H., de Graaf, B.H., Vidali, L., Hepler, P.K., et al., 2002. Rab2 GTPase regulates vesicle trafficking between the endo plasmic reticulum and the Golgi bodies and is important to pollen tube growth. Plant Cell 14, 945–962. Cheung, A.Y., Wu, H.M., 2004. Overexpression of an Arabidopsis formin stimulates supernumerary actin cable formation from pollen tube cell membrane. Plant Cell 16, 257–269. Cheung, A.Y., Wu, H.M., 2008. Structural and signaling networks for the polar cell growth machinery in pollen tubes. Annu. Rev. Plant Biol. 59, 547–572. Clapham, D.E., 1995. Calcium signaling. Cell 80, 259–268. Cle´ment, C., Pacini, E., 2001. Anther plastids in angiosperms. Bot. Rev. 67, 54–73. Cremona, O., De Camilli, P., 2001. Phosphoinositides in membrane traffic at the synapse. J. Cell Sci. 114, 1041–1052. Dai, S., Chen, T., Chong, K., Xue, Y., Liu, S., Wang, T., 2007. Proteomics identi fication of differentially expressed proteins associated with pollen
POLLEN GERMINATION AND TUBE GROWTH
41
germination and tube growth reveals characteristics of germinated Oryza sativa pollen. Mol. Cell Proteomics 6, 207–230. Dai, S., Li, L., Chen, T., Chong, K., Xue, Y., Wang, T., 2006. Proteomic analyses of Oryza sativa mature pollen reveal novel proteins associated with pollen germination and tube growth. Proteomics 6, 2504–2529. de Azevedo Souza, C., Kim, S.S., Koch, S., Kienow, L., Schneider, K., McKim, S.M., et al., 2009. A novel fatty acyl-CoA synthetase is required for pollen develop ment and sporopollenin biosynthesis in Arabidopsis. Plant Cell 21, 507–525. de Graaf, B.H., Cheung, A.Y., Andreyeva, T., Levasseur, K., Kieliszewski, M., Wu, H.M., 2005. Rab11 GTPase-regulated membrane trafficking is crucial for tip-focused pollen tube growth in tobacco. Plant Cell 17, 2564–2579. DerMardirossian, C., Bokoch, G.M., 2005. GDIs: central regulatory molecules in Rho GTPase activation. Trends Cell Biol. 15, 356–363. Dobritsa, A.A., Nishikawa, S., Preuss, D., Urbanczyk-Wochniak, E., Sumner, L.W., Hammond, A., et al., 2009b. LAP3, a novel plant protein required for pollen development, is essential for proper exine formation. Sex. Plant Reprod. 22, 167–177. Dobritsa, A.A., Shrestha, J., Morant, M., Pinot, F., Matsuno, M., Swanson, R., et al., 2009a. CYP704B1 is a long-chain fatty acid omega-hydroxylase essential for sporopollenin synthesis in pollen of Arabidopsis. Plant Physiol. 151, 574–589. Dong, X., Hong, Z., Sivaramakrishnan, M., Mahfouz, M., Verma, D.P., 2005. Callose synthase (CalS5) is required for exine formation during microga metogenesis and for pollen viability in Arabidopsis. Plant J. 42, 315–328. Doughty, J., Dixon, S., Hiscock, S.J., Willis, A.C., Parkin, I.A., Dickinson, H.G., 1998. PCP-A1, a defensin-like Brassica pollen coat protein that binds the S locus glycoprotein, is the product of gametophytic gene expression. Plant Cell 10, 1333–1347. Doughty, J., Hedderson, F., McCubbin, A., Dickinson, H., 1993. Interaction between a coating-borne peptide of the Brassica pollen grain and stigmatic S (self-incompatibility)-locus-specific glycoproteins. Proc. Natl. Acad. Sci. U.S.A. 90, 467–471. Dowd, P.E., Coursol, S., Skirpan, A.L., Kao, T.H., Gilroy, S., 2006. Petunia phos pholipase C1 is involved in pollen tube growth. Plant Cell 18, 1438–1453. Dutta, R., Robinson, K.R., 2004. Identification and characterization of stretchactivated ion channels in pollen protoplasts. Plant Physiol. 135, 1398–1406. Elleman, C.J., Dickinson, H.G., 1986. Pollen-stigma interactions in Brassica. IV. Structural reorganization in the pollen grains during hydration. J. Cell Sci. 80, 141–157. Elleman, C.J., Dickinson, H.G., 1990. The role of the exine coating in pollen–stigma interactions in Brassica oleracea L. New Phytol. 114, 511–518. Elleman, C.J., Franklin-Tong, V., Dickinson, H.G., 1992. Pollination in species with dry stigmas: the nature of the early stigmatic response and the pathway taken by pollen tubes. New Phytol. 121, 413–424. Escobar-Restrepo, J.M., Huck, N., Kessler, S., Gagliardini, V., Gheyselinck, J., Yang, W.C., et al., 2007. The FERONIA receptor-like kinase mediates male-female interactions during pollen tube reception. Science 317, 656–660. Fan, X., Hou, J., Chen, X., Chaudhry, F., Staiger, C.J., Ren, H., 2004. Identification and characterization of a Ca2þ-dependent actin filament-severing protein from lily pollen. Plant Physiol. 136, 3979–3989. Faure, J., Vignais, P.V., Dagher, M.C., 1999. Phosphoinositide-dependent activation of Rho A involves partial opening of the RhoA/Rho-GDI complex. Eur. J. Biochem. 262, 879–889.
42
H.-J. WANG ET AL.
Feijo, J.A., Sainhas, J., Hackett, G.R., Kunkel, J.G., Hepler, P.K., 1999. Growing pollen tubes possess a constitutive alkaline band in the clear zone and a growth-dependent acidic tip. J. Cell Biol. 144, 483–496. Feijo, J.A., Sainhas, J., Holdaway-Clarke, T., Cordeiro, M.S., Kunkel, J.G., Hepler, P.K., 2001. Cellular oscillations and the regulation of growth: the pollen tube paradigm. Bioessays 23, 86–94. Fernando, D.D., 2005. Characterization of pollen tube development in Pinus strobus (Eastern white pine) through proteomic analysis of differentially expressed proteins. Proteomics 5, 4917–4926. Fiebig, A., Kimport, R., Preuss, D., 2004. Comparisons of pollen coat genes across Brassicaceae species reveal rapid evolution by repeat expansion and diver sification. Proc. Natl. Acad. Sci. U.S.A. 101, 3286–3291. Fiebig, A., Mayfield, J.A., Miley, N.L., Chau, S., Fischer, R.L., Preuss, D., 2000. Alterations in CER6, a gene identical to CUT1, differentially affect longchain lipid content on the surface of pollen and stems. Plant Cell 12, 2001–2008. Filonova, L.H., Bozhkov, P.V., Brukhin, V.B., Daniel, G., Zhivotovsky, B., von Arnold, S., 2000. Two waves of programmed cell death occur during formation and development of somatic embryos in the gymnosperm, Nor way spruce. J. Cell Sci. 113, 4399–4411. Foissner, I., Grolig, F., Obermeyer, G., 2002. Reversible protein phosphorylation regulates the dynamic organization of the pollen tube cytoskeleton: effects of calyculin A and okadaic acid. Protoplasma 220, 1–15. Foreman, J., Demidchik, V., Bothwell, J.H., Mylona, P., Miedema, H., Torres, M. A., et al., 2003. Reactive oxygen species produced by NADPH oxidase regulate plant cell growth. Nature 422, 442–446. Fu, Y., Wu, G., Yang, Z., 2001. Rop GTPase-dependent dynamics of tip-localized F-actin controls tip growth in pollen tubes. J. Cell Biol. 152, 1019–1032. Fujiki, Y., Yoshimoto, K., Ohsumi, Y., 2007. An Arabidopsis homolog of yeast ATG6/VPS30 is essential for pollen germination. Plant Physiol. 143, 1132–1139. Gaffal, K.P., Friedrichs, G.J., El-Gammal, S., 2007. Ultrastructural evidence for a dual function of the phloem and programmed cell death in the floral nectary of Digitalis purpurea. Ann. Bot. 99, 593–607. Garzon, M., Eifler, K., Faust, A., Scheel, H., Hofmann, K., Koncz, C., et al., 2007. PRT6/At5g02310 encodes an Arabidopsis ubiquitin ligase of the N-end rule pathway with arginine specificity and is not the CER3 locus. FEBS Lett. 581, 3189–3196. Gaude, T., Dumas, C., 1984. A membrane-like structure on the pollen wall surface in Brassica. Ann. Bot. 54, 821–825. Gaude, T., Dumas, C., 1986. Organization of stigma surface components in Brassica: a cytochemical study. J. Cell Sci. 82, 203–216. Goldman, M.H., Goldberg, R.B., Mariani, C., 1994. Female sterile tobacco plants are produced by stigma-specific cell ablation. EMBO J. 13, 2976–2984. Gossot, O., Geitmann, A., 2007. Pollen tube growth: coping with mechanical obsta cles involves the cytoskeleton. Planta 226, 405–416. Gu, Y., Fu, Y., Dowd, P., Li, S., Vernoud, V., Gilroy, S., et al., 2005. A Rho family GTPase controls actin dynamics and tip growth via two counteracting downstream pathways in pollen tubes. J. Cell Biol. 169, 127–138. Gu, Y., Li, S., Lord, E.M., Yang, Z., 2006. Members of a novel class of Arabidopsis Rho guanine nucleotide exchange factors control Rho GTPase-dependent polar growth. Plant Cell 18, 366–381. Guan, Y.F., Huang, X.Y., Zhu, J., Gao, J.F., Zhang, H.X., Yang, Z.N., 2008. RUPTURED POLLEN GRAIN1, a member of the MtN3/saliva gene
POLLEN GERMINATION AND TUBE GROWTH
43
family, is crucial for exine pattern formation and cell integrity of micro spores in Arabidopsis. Plant Physiol. 147, 852–863. Haerizadeh, F., Wong, C.E., Bhalla, P.L., Gresshoff, P.M., Singh, M.B., 2009. Genomic expression profiling of mature soybean (Glycine max) pollen. BMC Plant Biol. 9, 25. Hancock, J., Desikan, R., Harrison, J., Bright, J., Hooley, R., Neill, S., 2006. Doing the unexpected: proteins involved in hydrogen peroxide perception. J. Exp. Bot. 57, 1711–1718. Hannoufa, A., McNevin, J., Lemieux, B., 1993. Epicuticular waxes of eceriferum mutants of Arabidopsis thaliana. Phytochemistry 33, 851–855. Hannoufa, A., Negruk, V., Eisner, G., Lemieux, B., 1996. The CER3 gene of Arabidopsis thaliana is expressed in leaves, stems, roots, flowers and apical meristems. Plant J. 10, 459–467. Hansen, J.D., Pyee, J., Xia, Y., Wen, T.J., Robertson, D.S., Kolattukudy, P.E., et al., 1997. The glossy1 locus of maize and an epidermis-specific cDNA from Kleinia odora define a class of receptor-like proteins required for the normal accumulation of cuticular waxes. Plant Physiol. 113, 1091–1100. Harrison-Lowe, N.J., Olsen, L.J., 2008. Autophagy protein 6 (ATG6) is required for pollen germination in Arabidopsis thaliana. Autophagy 4, 339–348. Hay, J.C., Fisette, P.L., Jenkins, G.H., Fukami, K., Takenawa, T., Anderson, R.A., et al., 1995. ATP-dependent inositide phosphorylation required for Ca(2þ) activated secretion. Nature 374, 173–177. Heizman, P., Luu, D.T., Dumas, C., 2000. Pollen-stigma adhesion in the Brassica ceae. Ann. Bot. 85 (Supplement A), 23–27. Helling, D., Possart, A., Cottier, S., Klahre, U., Kost, B., 2006. Pollen tube tip growth depends on plasma membrane polarization mediated by tobacco PLC3 activity and endocytic membrane recycling. Plant Cell 18, 3519–3534. Hepler, P.K., Vidali, L., Cheung, A.Y., 2001. Polarized cell growth in higher plants. Annu. Rev. Cell Dev. Biol. 17, 159–187. Heslop-Harrison, Y., 1981. Stigma characteristics and angiosperm taxonomy. Nord. J. Bot. 1, 401–420. Heslop-Harrison, J., Heslop-Harrison, Y., Barber, J., 1975. The stigma surface in incompatibility responses. Proc. R. Soc. B Biol. Sci. 188, 287–297. Heslop-Harrison, Y., Shivanna, K.R., 1977. The receptive surface of the angiosperm stigma. Ann. Bot. 41, 1233–1258. Higaki, T., Sano, T., Hasezawa, S., 2007. Actin microfilament dynamics and actin side-binding proteins in plants. Curr. Opin. Plant Biol. 10, 549–556. Higashiyama, T., Yabe, S., Sasaki, N., Nishimura, Y., Miyagishima, S., Kuroiwa, H., et al., 2001. Pollen tube attraction by the synergid cell. Science 293, 1480–1483. Hiscock, S.J., Bright, J., McInnis, S.M., Desikan, R., Hancock, J.T., 2007. Signaling on the stigma: potential new roles for ROS and NO in plant cell signaling. Plant Signal. Behav. 2, 23–24. Hiscock, S.J., Dickinson, H.G., 1993. Unilateral incompatibility within the Brassi caceae: further evidence for the involvement of the self-incompatibility (S)-locus. Theor. Appl. Genet. 86, 744–753. Hiscock, S.J., Doughty, J., Willis, A.C., Dickinson, H.G., 1995. A 7-kDa pollen coating-borne peptide from Brassica napus interacts with S-locus glycopro tein and S-locus-related glycoprotein. Planta 196, 367–374. Hoekstra, F.A., Crowe, J.H., Crowe, L.M., 1991. Desiccation tolerance of Papaver dubium L. pollen during its development in the anther. Plant Physiol. 88, 626–632.
44
H.-J. WANG ET AL.
Holdaway-Clarke, T.L., Feijo, J.A., Hackett, G.R., Kunkel, J.G., Hepler, P.K., 1997. Pollen tube growth and the intracellular cytosolic calcium gradient oscillate in phase while extracellular calcium influx is delayed. Plant Cell 9, 1999–2010. Holmes-Davis, R., Tanaka, C.K., Vensel, W.H., Hurkman, W.J., McCormick, S., 2005. Proteome mapping of mature pollen of Arabidopsis thaliana. Proteo mics 5, 4864–4884. Holweg, C., Nick, P., 2004. Arabidopsis myosin XI mutant is defective in organelle movement and polar auxin transport. Proc. Natl. Acad. Sci. U.S.A. 101, 10488–10493. Honys, D., Twell, D., 2003. Comparative analysis of the Arabidopsis pollen tran scriptome. Plant Physiol. 132, 640–652. Honys, D., Twell, D., 2004. Transcriptome analysis of haploid male gametophyte development in Arabidopsis. Genome Biol. 5, R85. Huang, S., Gao, L., Blanchoin, L., Staiger, C.J., 2006. Heterodimeric capping protein from Arabidopsis is regulated by phosphatidic acid. Mol. Biol. Cell 17, 1946–1958. Huck, N., Moore, J.M., Federer, M., Grossniklaus, U., 2003. The Arabidopsis mutant feronia disrupts the female gametophytic control of pollen tube reception. Development 130, 2149–2159. Hulskamp, M., Kopczak, S.D., Horejsi, T.F., Kihl, B.K., Pruitt, R.E., 1995. Identi fication of genes required for pollen-stigma recognition in Arabidopsis thaliana. Plant J. 8, 703–714. Hussey, P.J., Ketelaar, T., Deeks, M.J., 2006. Control of the actin cytoskeleton in plant cell growth. Annu. Rev. Plant Biol. 57, 109–125. Hwang, J.U., Gu, Y., Lee, Y.J., Yang, Z., 2005. Oscillatory ROP GTPase activation leads the oscillatory polarized growth of pollen tubes. Mol. Biol. Cell 16, 5385–5399. Inoue, Y., Suzuki, T., Hattori, M., Yoshimoto, K., Ohsumi, Y., Moriyasu, Y., 2006. AtATG genes, homologs of yeast autophagy genes, are involved in constitu tive autophagy in Arabidopsis root tip cells. Plant Cell Physiol. 47, 1641–1652. Ito, T., Nagata, N., Yoshiba, Y., Ohme-Takagi, M., Ma, H., Shinozaki, K., 2007. Arabidopsis MALE STERILITY1 encodes a PHD-type transcription factor and regulates pollen and tapetum development. Plant Cell 19, 3549–3562. Jenks, M.A., Tuttle, H.A., Eigenbrode, S.D., Feldmann, K.A., 1995. Leaf epicuti cular waxes of the eceriferum mutants in Arabidopsis. Plant Physiol. 108, 369–377. Jiang, P.-L., Wang, C.-S., Hsu, C.-M., Jauh, G.-Y., Tzen, J.T.C., 2007. Stable oil bodies sheltered by a unique oleosin in lily pollen. Plant Cell Physiol. 48, 812–821. Johnson, M.A., Lord, E.M., 2006. Extracellular guidance cues and intracellular signaling pathways that direct pollen tube growth. In: The Pollen Tube (Malho´, R. (Ed.)), Plant Cell Monographs Volume 3/2006, Springer Berlin Heidelberg. pp 223–242. Johnson, M.A., von Besser, K., Zhou, Q., Smith, E., Aux, G., Patton, D., et al., 2004. Arabidopsis hapless mutations define essential gametophytic functions. Genetics 168, 971–982. Kandasamy, M.K., Paolillo, D.J., Faraday, C.D., Nasrallah, J.B., Nasrallah, M.E., 1989. The S-locus specific glycoproteins of Brassica accumulate in the cell wall of developing stigma papillae. Dev. Biol. 134, 462–472. Kaothien, P., Ok, S.H., Shuai, B., Wengier, D., Cotter, R., Kelley, D., et al., 2005. Kinase partner protein interacts with the LePRK1 and LePRK2 receptor kinases and plays a role in polarized pollen tube growth. Plant J. 42, 492–503.
POLLEN GERMINATION AND TUBE GROWTH
45
Ketelaar, T., Allwood, E.G., Anthony, R., Voigt, B., Menzel, D., Hussey, P.J., 2004. The actin-interacting protein AIP1 is essential for actin organization and plant development. Curr. Biol. 14, 145–149. Kim, S., Mollet, J.C., Dong, J., Zhang, K., Park, S.Y., Lord, E.M., 2003. Chemo cyanin, a small basic protein from the lily stigma, induces pollen tube chemotropism. Proc. Natl. Acad. Sci. U.S.A. 100, 16125–16130. Klahre, U., Becker, C., Schmitt, A.C., Kost, B., 2006. Nt-RhoGDI2 regulates Rac/ Rop signaling and polar cell growth in tobacco pollen tubes. Plant J. 46, 1018–1031. Klahre, U., Kost, B., 2006. Tobacco RhoGTPase ACTIVATING PROTEIN1 spa tially restricts signaling of RAC/Rop to the apex of pollen tubes. Plant Cell 18, 3033–3046. Knox, R.B., Clarke, A., Harrison, S., Smith, P., Marchalonis, J.J., 1976. Cell recog nition in plants: determinants of the stigma surface and their pollen inter actions. Proc. Natl. Acad. Sci. U.S.A. 73, 2788–2792. Knox, R.B., Gaget, M., Dumas, C., 1987. Mentor pollen techniques. Int. Rev. Cytol. 107, 315–332. Ko¨hler, C., Grossniklaus, U., 2005. Seed development and genomic imprinting in plants. In: Progress in Molecular and Subcellular Biology: Epigenetics and Chromatin (Jeanteur, P. (Ed.)), Springer-Verlag, Berlin-Heidelberg, pp. 237–262. Koornneef, M., Hanhart, C.J., Thiel, F., 1989. A genetic and phenotypic description of eceriferum (cer) mutants in Arabidopsis thaliana. J. Hered. 80, 118–122. Kost, B., Lemichez, E., Spielhofer, P., Hong, Y., Tolias, K., Carpenter, C., et al., 1999. Rac homologues and compartmentalized phosphatidylinositol 4, 5-bisphosphate act in a common pathway to regulate polar pollen tube growth. J. Cell Biol. 145, 317–330. Kost, B., Spielhofer, P., Chua, N.H., 1998. A GFP-mouse talin fusion protein labels plant actin filaments in vivo and visualizes the actin cytoskeleton in growing pollen tubes. Plant J. 16, 393–401. Kovar, D.R., Drobak, B.K., Staiger, C.J., 2000. Maize profilin isoforms are func tionally distinct. Plant Cell 12, 583–598. Kunst, L., Samuels, A., 2003. Biosynthesis and secretion of plant cuticular wax. Prog. Lipid Res. 42, 51–80. Laloi, C., Apel, K., Danon, A., 2004. Reactive oxygen signalling: the latest news. Curr. Opin. Plant Biol. 7, 323–328. Lalonde, B.A., Nasrallah, M.E., Dwyer, K.G., Chen, C.H., Barlow, B., Nasrallah, J.B., 1989. A highly conserved Brassica gene with homology to the S-locus-specific glycoprotein structural gene. Plant Cell 1, 249–258. Lee, C.B., Kim, S., McClure, B., 2009. A pollen protein, NaPCCP, that binds pistil arabinogalactan proteins also binds phosphatidylinositol 3-phosphate and associates with the pollen tube endomembrane system. Plant Physiol. 149, 791–802. Lee, J.Y., Lee, D.H., 2003. Use of serial analysis of gene expression technology to reveal changes in gene expression in Arabidopsis pollen undergoing cold stress. Plant Physiol. 132, 517–529. Lee, C.B., Swatek, K.N., McClure, B., 2008. Pollen proteins bind to the C-terminal domain of Nicotiana alata pistil arabinogalactan proteins. J. Biol. Chem. 283, 26965–26973. Lefebvre, B., Arango, M., Oufattole, M., Crouzet, J., Purnelle, B., Boutry, M., 2005. Identification of a Nicotiana plumbaginifolia plasma membrane H(þ)-ATPase gene expressed in the pollen tube. Plant Mol. Biol. 58, 775–787. Levine, B., Klionsky, D.J., 2004. Development by self-digestion; molecular mechan isms and biological functions of autophagy. Dev. Cell 6, 463–477.
46
H.-J. WANG ET AL.
Li, H., Lin, Y., Heath, R.M., Zhu, M.X., Yang, Z., 1999. Control of pollen tube tip growth by a Rop GTPase-dependent pathway that leads to tip-localized calcium influx. Plant Cell 11, 1731–1742. Liebe, S., Menzel, D., 1995. Actomyosin-based motility of endoplasmic reticulum and chloroplasts in Vallisneria mesophyll cells. Biol. Cell 85, 207–222. Liu, Y., Schiff, M., Czymmek, K., Talloczy, Z., Levine, B., Dinesh-Kumar, S.P., 2005. Autophagy regulates programmed cell death during the plant innate immune response. Cell 121, 567–577. Lolle, S.J., Berlyn, G.P., Engstrom, E.M., Krolikowski, K.A., Reiter, W.D., Pruitt, R.E., 1997. Developmental regulation of cell interactions in the Arabidopsis fiddlehead-1 mutant: a role for the epidermal cell wall and cuticle. Dev. Biol. 189, 311–321. Lolle, S.J., Cheung, A.Y., 1993. Promiscuous germination and growth of wildtype pollen from Arabidopsis and related species on the shoot of the Arabidopsis mutant, fiddlehead. Dev. Biol. 155, 250–258. Lolle, S.J., Cheung, A.Y., Sussex, I.M., 1992. Fiddlehead: an Arabidopsis mutant constitutively expressing an organ fusion program that involves interactions between epidermal cells. Dev. Biol. 152, 383–392. Lord, E.M., 2003. Adhesion and guidance in compatible pollination. J. Exp. Bot. 54, 47–54. Lord, E.M., Russell, S.D., 2002. The mechanisms of pollination and fertilization in plants. Annu. Rev. Cell Dev. Biol. 18, 81–105. Lovy-Wheeler, A., Kunkel, J.G., Allwood, E.G., Hussey, P.J., Hepler, P.K., 2006. Oscillatory increases in alkalinity anticipate growth and may regulate actin dynamics in pollen tubes of lily. Plant Cell 18, 2182–2193. Luu, D.T., Heizmann, P., Dumas, C., 1997. Pollen-stigma adhesion in Kale is not dependent on the self-(in)compatibility genotype. Plant Physiol. 115, 1221–1230. Luu, D.T., Marty-Mazars, D., Trick, M., Dumas, C., Heizmann, P., 1999. Pollen-stigma adhesion in Brassica spp involves SLG and SLR1 glycoproteins. Plant Cell 11, 251–262. Ma, H., 2003. Plant reproduction: GABA gradient, guidance and growth. Curr. Biol. 13, R834–836. Ma, J., Skibbe, D.S., Fernandes, J., Walbot, V., 2008. Male reproductive develop ment: gene expression profiling of maize anther and pollen ontogeny. Genome Biol. 9, R181. Marton, M.L., Cordts, S., Broadhvest, J., Dresselhaus, T., 2005. Micropylar pollen tube guidance by egg apparatus 1 of maize. Science 307, 573–576. Matsubayashi, Y., Ogawa, M., Kihara, H., Niwa, M., Sakagami, Y. (2006b). Disruption and overexpression of Arabidopsis phytosulfokine receptor gene affects cellular longevity and potential for growth. Plant Physiol. 142, 45–53. Matsubayashi, Y., Ogawa, M., Morita, A., Sakagami, Y., 2002. An LRR receptor kinase involved in perception of a peptide plant hormone, phytosulfokine. Science 296, 1470–1472. Matsubayashi, Y., Sakagami, Y., 2000. 120- and 160-kDa receptors for endogenous mitogenic peptide, phytosulfokine-a, in rice plasma membranes. J. Biol. Chem. 275, 15520–15525. Matsubayashi, Y., Shinohara, H., Ogawa, M. (2006a). Identification and functional characterization of phytosulfokine receptor using a ligand-based approach. Chem. Rec. 6, 356–364. Matsubayashi, Y., Yang, H., Sakagami, Y., 2001. Peptide signals and their receptors in higher plants. Trends Plant Sci. 6, 573–577.
POLLEN GERMINATION AND TUBE GROWTH
47
Mattsson, O., Knox, R.B., Heslop-Harrison, J., Heslop-Harrison, Y., 1974. Protein pellicle of stigmatic papillae as a probable recognition site in incompatibility reactions. Nature 247, 298–300. Mayfield, J.A., Fiebig, A., Johnstone, S.E., Preuss, D., 2001. Gene families from the Arabidopsis thaliana pollen coat proteome. Science 292, 2482–2485. Mayfield, J.A., Preuss, D., 2000. Rapid initiation of Arabidopsis pollination requires the oleosin-domain protein GRP17. Nat. Cell Biol. 2, 128–130. McCormick, S., 2007. Plant science. Reproductive dialog. Science 317, 606–607. McInnis, S.M., Costa, L.M., Gutierrez-Marcos, J.F., Henderson, C.A., Hiscock, S.J., 2005. Isolation and characterization of a polymorphic stigma-specific class III peroxidase gene from Senecio squalidus L. (Asteraceae). Plant Mol. Biol. 57, 659–677. McInnis, S.M., Desikan, R., Hancock, J.T., Hiscock, S.J. (2006a). Production of reactive oxygen species and reactive nitrogen species by angiosperm stigmas and pollen: potential signalling crosstalk? New Phytol. 172, 221–228. McInnis, S.M., Emery, D.C., Porter, R., Desikan, R., Hancock, J.T., Hiscock, S.J. (2006b). The role of stigma peroxidases in flowering plants: insights from further characterization of a stigma-specific peroxidase (SSP) from Senecio squalidus (Asteraceae). J. Exp. Bot. 57, 1835–1846. McKenna, S.T., Kunkel, J.G., Bosch, M., Rounds, C.M., Vidali, L., Winship, L.J., et al., 2009. Exocytosis precedes and predicts the increase in growth in oscillating pollen tubes. Plant Cell 21, 3026–3040. McNevin, J.P., Woodward, W., Hannoufa, A., Feldmann, K.A., Lemieux, B., 1993. Isolation and characterization of eceriferum (cer) mutants induced by T-DNA insertions in Arabidopsis thaliana. Genome 36, 610–618. Meijer, H.J., Munnik, T., 2003. Phospholipid-based signaling in plants. Annu. Rev. Plant Biol. 54, 265–306. Meinhard, M., Rodriguez, P.L., Grill, E., 2002. The sensitivity of ABI2 to hydrogen peroxide links the abscisic acid-response regulator to redox signalling. Planta 214, 775–782. Miki-Hirosige, H., Nakamura, S., 1983. Growth and differentiation of amyloplasts during male gamete development in Lilium longiflorum. In Pollen: Biology and Implications for Plant Breeding (Mulcahy, D.L., Ottaviano, E., (Eds.)), Elsevier Science Publisher, Biomedical, New York, pp. 141–147. Miller, D.D., Scordilis, S.P., Hepler, P.K., 1995. Identification and localization of three classes of myosins in pollen tubes of Lilium longiflorum and Nicotiana alata. J. Cell Sci. 108 (Pt 7), 2549–2563. Mollet, J.C., Park, S.Y., Nothnagel, E.A., Lord, E.M., 2000. A lily stylar pectin is necessary for pollen tube adhesion to an in vitro stylar matrix. Plant Cell 12, 1737–1750. Monteiro, D., Castanho Coelho, P., Rodrigues, C., Camacho, L., Quader, H., Malho, R., 2005. Modulation of endocytosis in pollen tube growth by phosphoinositides and phospholipids. Protoplasma 226, 31–38. Morant, M., Jorgensen, K., Schaller, H., Pinot, F., Moller, B.L., WerckReichhart, D., et al., 2007. CYP703 is an ancient cytochrome P450 in land plants catalyzing in-chain hydroxylation of lauric acid to provide building blocks for sporopollenin synthesis in pollen. Plant Cell 19, 1473–1487. Moriyasu, Y., Hillmer, S., 2000. Autophagy and Vacuole Formation. Sheffield Academic Press, Sheffield, England, pp 71–89. Moscatelli, A., Ciampolini, F., Rodighiero, S., Onelli, E., Cresti, M., Santo, N., et al., 2007. Distinct endocytic pathways identified in tobacco pollen tubes using charged nanogold. J. Cell Sci. 120, 3804–3819.
48
H.-J. WANG ET AL.
Muschietti, J., Eyal, Y., McCormick, S., 1998. Pollen tube localization implies a role in pollen-pistil interactions for the tomato receptor-like protein kinases LePRK1 and LePRK2. Plant Cell 10, 319–330. Myers, C., Romanowsky, S.M., Barron, Y.D., Garg, S., Azuse, C.L., Curran, A., et al., 2009. Calcium-dependent protein kinases regulate polarized tip growth in pollen tubes. Plant J. 59, 528–539. Nebenfuhr, A., Gallagher, L.A., Dunahay, T.G., Frohlick, J.A., Mazurkiewicz, A. M., Meehl, J.B., et al., 1999. Stop-and-go movements of plant Golgi stacks are mediated by the acto-myosin system. Plant Physiol. 121, 1127–1142. Neill, S.J., Desikan, R., Clarke, A., Hurst, R.D., Hancock, J.T., 2002b. Hydrogen peroxide and nitric oxide as signalling molecules in plants. J. Exp. Bot. 53, 1237–1247. Neill, S., Desikan, R., Hancock, J., 2002a. Hydrogen peroxide signalling. Curr. Opin. Plant Biol. 5, 388–395. Nishikawa, S., Zinkl, G.M., Swanson, R.J., Maruyama, D., Preuss, D., 2005. Callose (b-1, 3 glucan) is essential for Arabidopsis pollen wall patterning, but not tube growth. BMC Plant Biol. 5, 22. Noda, T., Suzuki, K., Ohsumi, Y., 2002. Yeast autophagosomes: de novo formation of a membrane structure. Trends Cell Biol. 12, 231–235. Noir, S., Brautigam, A., Colby, T., Schmidt, J., Panstruga, R., 2005. A reference map of the Arabidopsis thaliana mature pollen proteome. Biochem. Biophys. Res. Commun. 337, 1257–1266. Okuda, S., Tsutsui, H., Shiina, K., Sprunck, S., Takeuchi, H., Yui, R., et al., 2009. Defensin-like polypeptide LUREs are pollen tube attractants secreted from synergid cells. Nature 458, 357–361. Pacini, E., 1996. Types and meaning of pollen carbohydrate reserves. Sex. Plant Reprod. 9, 362–366. Palanivelu, R., Brass, L., Edlund, A.F., Preuss, D., 2003. Pollen tube growth and guidance is regulated by POP2, an Arabidopsis gene that controls GABA levels. Cell 114, 47–59. Pandey, K.K., 1967. Origin of genetic variability: combinations of peroxidase iso zymes determine multiple allelism of the S gene. Nature 213, 669–672. Park, S.Y., Jauh, G.Y., Mollet, J.C., Eckard, K.J., Nothnagel, E.A., Walling, L.L., et al., 2000. A lipid transfer-like protein is necessary for lily pollen tube adhesion to an in vitro stylar matrix. Plant Cell 12, 151–164. Park, S.Y., Lord, E.M., 2003. Expression studies of SCA in lily and confirmation of its role in pollen tube adhesion. Plant Mol. Biol. 51, 183–189. Parton, R.M., Fischer-Parton, S., Trewavas, A.J., Watahiki, M.K., 2003. Pollen tubes exhibit regular periodic membrane trafficking events in the absence of apical extension. J. Cell Sci. 116, 2707–2719. Parton, R.M., Fischer-Parton, S., Watahiki, M.K., Trewavas, A.J., 2001. Dynamics of the apical vesicle accumulation and the rate of growth are related in individual pollen tubes. J. Cell Sci. 114, 2685–2695. Paxson-Sowders, D.M., Dodrill, C.H., Owen, H.A., Makaroff, C.A., 2001. DEX1, a novel plant protein, is required for exine pattern formation during pollen development in Arabidopsis. Plant Physiol. 127, 1739–1749. Picton, J.M., Steer, M.W., 1983. Membrane recycling and the control of secretory activity in pollen tubes. J. Cell Sci. 63, 303–310. Pierson, E.S., Miller, D.D., Callaham, D.A., van Aken, J., Hackett, G., Hepler, P.K., 1996. Tip-localized calcium entry fluctuates during pollen tube growth. Dev. Biol. 174, 160–173. Piffanelli, P., Ross, J.H.E., Murphy, D.J., 1998. Biogenesis and function of the lipidic structures of pollen grains. Sex. Plant Reprod. 11, 65–80.
POLLEN GERMINATION AND TUBE GROWTH
49
Pina, C., Pinto, F., Feijo, J.A., Becker, J.D., 2005. Gene family analysis of the Arabidopsis pollen transcriptome reveals biological implications for cell growth, division control, and gene expression regulation. Plant Physiol. 138, 744–756. Potocky, M., Elias, M., Profotova, B., Novotna, Z., Valentova, O., Zarsky, V., 2003. Phosphatidic acid produced by phospholipase D is required for tobacco pollen tube growth. Planta 217, 122–130. Prado, A.M., Colaco, R., Moreno, N., Silva, A.C., Feijo, J.A., 2008. Targeting of pollen tubes to ovules is dependent on nitric oxide (NO) signaling. Mol. Plant 1, 703–714. Prado, A.M., Porterfield, D.M., Feijo, J.A., 2004. Nitric oxide is involved in growth regulation and re-orientation of pollen tubes. Development 131, 2707–2714. Preuss, D., Lemieux, B., Yen, G., Davis, R.W., 1993. A conditional sterile mutation eliminates surface components from Arabidopsis pollen and disrupts cell signaling during fertilization. Genes Dev. 7, 974–985. Pruitt, R.E., Vielle-Calzada, J.P., Ploense, S.E., Grossniklaus, U., Lolle, S.J., 2000. FIDDLEHEAD, a gene required to suppress epidermal cell interactions in Arabidopsis, encodes a putative lipid biosynthetic enzyme. Proc. Natl. Acad. Sci. U.S.A. 97, 1311–1316. Qin, Y., Leydon, A.R., Manziello, A., Pandey, R., Mount, D., Denic, S., et al., 2009. Penetration of the stigma and style elicits a novel transcriptome in pollen tubes, pointing to genes critical for growth in a pistil. PLoS Genet. 5, e1000621. Qin, G., Ma, Z., Zhang, L., Xing, S., Hou, X., Deng, J., et al., 2007. Arabidopsis AtBECLIN 1/AtAtg6/AtVps30 is essential for pollen germination and plant development. Cell Res. 17, 249–263. Rashotte, A.M., Jenks, M.A., Ross, A.S., Feldmann, K.A., 2004. Novel eceriferum mutants in Arabidopsis thaliana. Planta 219, 5–13. Reddy, A.S., Day, I.S., 2001. Analysis of the myosins encoded in the recently completed Arabidopsis thaliana genome sequence. Genome Biol. 2, RESEARCH0024. Reisen, D., Hanson, M.R., 2007. Association of six YFP-myosin XI-tail fusions with mobile plant cell organelles. BMC Plant Biol. 7, 6. Ren, H., Xiang, Y., 2007. The function of actin-binding proteins in pollen tube growth. Protoplasma 230, 171–182. Rentel, M.C., Knight, M.R., 2004. Oxidative stress-induced calcium signaling in Arabidopsis. Plant Physiol. 135, 1471–1479. Roberts, I.N., Harrod, G., Dickinson, H.G., 1984. Pollen-stigma interactions in Bras sica oleracea. II. The fate of stigma surface proteins following pollination and their role in the self-incompatibility response. J. Cell Sci. 66, 255–264. Romagnoli, S., Cai, G., Cresti, M., 2003. In vitro assays demonstrate that pollen tube organelles use kinesin-related motor proteins to move along microtubules. Plant Cell 15, 251–269. Romagnoli, S., Cai, G., Faleri, C., Yokota, E., Shimmen, T., Cresti, M., 2007. Microtubule- and actin filament-dependent motors are distributed on pol len tube mitochondria and contribute differently to their movement. Plant Cell Physiol. 48, 345–361. Rotman, N., Rozier, F., Boavida, L., Dumas, C., Berger, F., Faure, J.E., 2003. Female control of male gamete delivery during fertilization in Arabidopsis thaliana. Curr. Biol. 13, 432–436. Rowland, O., Lee, R., Franke, R., Schreiber, L., Kunst, L., 2007. The CER3 wax biosynthetic gene from Arabidopsis thaliana is allelic to WAX2/YRE/FLP1. FEBS Lett. 581, 3538–3544.
50
H.-J. WANG ET AL.
Sandaklie-Nikolova, L., Palanivelu, R., King, E.J., Copenhaver, G.P., Drews, G.N., 2007. Synergid cell death in Arabidopsis is triggered following direct inter action with the pollen tube. Plant Physiol. 144, 1753–1762. Sarker, R.H., Elleman, C.J., Dickinson, H.G., 1988. Control of pollen hydration in Brassica requires continued protein synthesis, and glycosylation in neces sary for intraspecific incompatibility. Proc. Natl. Acad. Sci. U.S.A. 85, 4340–4344. Schein, M., Yang, Z., Mitchell-Olds, T., Schmid, K.J., 2004. Rapid evolution of a pollen-specific oleosin-like gene family from Arabidopsis thaliana and clo sely related species. Mol. Biol. Evol. 21, 659–669. Schiott, M., Romanowsky, S.M., Baekgaard, L., Jakobsen, M.K., Palmgren, M.G., Harper, J.F., 2004. A plant plasma membrane Ca2þ pump is required for normal pollen tube growth and fertilization. Proc. Natl. Acad. Sci. U.S.A. 101, 9502–9507. Scott, R.J., 1994. Pollen exine-the sporopollenin enigma and the physics of pattern. In: Scott, R.J., Stead, M.A. (Eds.), Molecular and Cellular Aspects of Plant Reproduction. Cambridge University Press, Cambridge, pp. 49–81. Seay, M.D., Dinesh-Kumar, S.P., 2005. Life after death: are autophagy genes involved in cell death and survival during plant innate immune responses? Autophagy 1, 185–186. Shimizu, K.K., Ito, T., Ishiguro, S., Okada, K., 2008. MAA3 (MAGATAMA3) helicase gene is required for female gametophyte development and pollen tube guidance in Arabidopsis thaliana. Plant Cell Physiol. 49, 1478–1483. Shimizu, K.K., Okada, K., 2000. Attractive and repulsive interactions between female and male gametophytes in Arabidopsis pollen tube guidance. Devel opment 127, 4511–4518. Shinohara, H., Ogawa, M., Sakagami, Y., Matsubayashi, Y., 2007. Identification of ligand binding site of phytosulfokine receptor by on-column photoaffinity labeling. J. Biol. Chem. 282, 124–131. Southworth, D., Dickinson, D.B., 1981. Ultrastructural changes in germinating lily pollen. Grana 20, 29–35. Staiger, C.J., 2000. Signaling to the actin cytoskeleton in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 257–288. Stead, A.D., Roberts, I.N., Dickinson, H.G., 1980. Pollen-stigma interaction in Brassica oleracea: the role of stigmatic proteins in pollen grain adhesion. J. Cell Sci. 42, 417–423. Stenmark, H., 2009. Rab GTPases as coordinators of vesicle traffic. Nat. Rev. Mol. Cell Biol. 10, 513–525. Suzuki, G., Kakizaki, T., Takada, Y., Shiba, H., Takayama, S., Isogai, A., et al., 2003. The S haplotypes lacking SLG in the genome of Brassica rapa. Plant Cell Rep. 21, 911–915. Suzuki, T., Kusaba, M., Matsushita, M., Okazaki, K., Nishio, T., 2000. Character ization of Brassica S-haplotypes lacking S-locus glycoprotein. FEBS Lett. 482, 102–108. Suzuki, T., Masaoka, K., Nishi, M., Nakamura, K., Ishiguro, S., 2008. Identification of kaonashi mutants showing abnormal pollen exine structure in Arabidop sis thaliana. Plant Cell Physiol. 49, 1465–1477. Swanson, R., Edlund, A.F., Preuss, D., 2004. Species specificity in pollen-pistil interactions. Annu. Rev. Genet. 38, 793–818. Takayama, S., Shiba, H., Iwano, M., Asano, K., Hara, M., Che, F.S., et al., 2000. Isolation and characterization of pollen coat proteins of Brassica campestris that interact with S locus-related glycoprotein 1 involved in pollen-stigma adhesion. Proc. Natl. Acad. Sci. U.S.A. 97, 3765–3770.
POLLEN GERMINATION AND TUBE GROWTH
51
Tang, W., Ezcurra, I., Muschietti, J., McCormick, S., 2002. A cysteine-rich extra cellular protein, LAT52, interacts with the extracellular domain of the pollen receptor kinase LePRK2. Plant Cell 14, 2277–2287. Tang, W., Kelley, D., Ezcurra, I., Cotter, R., McCormick, S., 2004. LeSTIG1, an extracellular binding partner for the pollen receptor kinases LePRK1 and LePRK2, promotes pollen tube growth in vitro. Plant J. 39, 343–353. Taylor, L.P., Hepler, P.K. (1997a). Pollen germination and tube growth. Annu. Rev. Plant Physiol. Plant Mol. Biol. 48, 461–491. Taylor, L.P., Hepler, P.K. (1997b). Pollen germination and tube growth. Annu. Rev. Plant Physiol. Plant Mol. Biol. 48, 461–491. Thomas, C., Hoffmann, C., Dieterle, M., Van Troys, M., Ampe, C., Steinmetz, A., 2006. Tobacco WLIM1 is a novel F-actin binding protein involved in actin cytoskeleton remodeling. Plant Cell 18, 2194–2206. Toyooka, K., Okamoto, T., Minamikawa, T., 2001. Cotyledon cells of Vigna mungo seedlings use at least two distinct autophagic machineries for degradation of starch granules and cellular components. J. Cell Biol. 154, 973–982. Umbach, A.L., Lalonde, B.A., Kandasamy, M.K., Nasrallah, J.B., Nasrallah, M.E., 1990. Immunodetection of protein glycoforms encoded by two independent genes of the self-incompatibility multigene family of Brassica. Plant Physiol. 93, 739–747. Updegraff, E.P., Zhao, F., Preuss, D., 2009. The extracellular lipase EXL4 is required for efficient hydration of Arabidopsis pollen. Sex. Plant Reprod. 22, 197–204. Van Gestel, K., Kohler, R.H., Verbelen, J.P., 2002. Plant mitochondria move on F-actin, but their positioning in the cortical cytoplasm depends on both F-actin and microtubules. J. Exp. Bot. 53, 659–667. Vidali, L., McKenna, S.T., Hepler, P.K., 2001. Actin polymerization is essential for pollen tube growth. Mol. Biol. Cell 12, 2534–2545. Wang, X., 2005. Regulatory functions of phospholipase D and phosphatidic acid in plant growth, development, and stress responses. Plant Physiol. 139, 566–573. Wang, H.J., Wan, A.R., Jauh, G.Y. (2008a). An actin-binding protein, LlLIM1, mediates calcium and hydrogen regulation of actin dynamics in pollen tubes. Plant Physiol. 147, 1619–1636. Wang, Y., Zhang, W.Z., Song, L.F., Zou, J.J., Su, Z., Wu, W.H. (2008b). Transcriptome analyses show changes in gene expression to accompany pollen germination and tube growth in Arabidopsis. Plant Physiol. 148, 1201–1211. Wengier, D., Valsecchi, I., Cabanas, M.L., Tang, W.H., McCormick, S., Muschietti, J., 2003. The receptor kinases LePRK1 and LePRK2 associate in pollen and when expressed in yeast, but dissociate in the presence of style extract. Proc. Natl. Acad. Sci. U.S.A. 100, 6860–6865. Wolters-Arts, M., Lush, W.M., Mariani, C., 1998. Lipids are required for directional pollen-tube growth. Nature 392, 818–821. Wolters-Arts, M., Van Der Weerd, L., Van Aelst, A.C., Van Der Weerd, J., Van As, H., Mariani, C., 2002. Water-conducting properties of lipids during pollen hydra tion. Plant Cell Environ. 25, 513–519. Wu, X., Chen, T., Zheng, M., Chen, Y., Teng, N., Samaj, J., et al., 2008. Integrative proteomic and cytological analysis of the effects of extracellular Ca(2þ) influx on Pinus bungeana pollen tube development. J. Proteome Res. 7, 4299–4312. Wu, G., Gu, Y., Li, S., Yang, Z., 2001. A genome-wide analysis of Arabidopsis Rop interactive CRIB motif-containing proteins that act as Rop GTPase targets. Plant Cell 13, 2841–2856.
52
H.-J. WANG ET AL.
Wu, H.M., Wang, H., Cheung, A.Y., 1995. A pollen tube growth stimulatory glycoprotein is deglycosylated by pollen tubes and displays a glycosylation gradient in the flower. Cell 82, 395–403. Wu, H.M., Wong, E., Ogdahl, J., Cheung, A.Y., 2000. A pollen tube growthpromoting arabinogalactan protein from Nicotiana alata is similar to the tobacco TTS protein. Plant J. 22, 165–176. Xiang, Y., Huang, X., Wang, T., Zhang, Y., Liu, Q., Hussey, P.J., Ren, H., 2007. ACTIN BINDING PROTEIN 29 from Lilium pollen plays an important role in dynamic actin remodeling. Plant Cell 19, 1930–1946. Xiong, Y., Contento, A.L., Bassham, D.C., 2007. Disruption of autophagy results in constitutive oxidative stress in Arabidopsis. Autophagy 3, 257–258. Yang, C., Kazanietz, M.G., 2003. Divergence and complexities in DAG signaling: looking beyond PKC. Trends Pharmacol. Sci. 24, 602–608. Yang, C., Vizcay-Barrena, G., Conner, K., Wilson, Z.A., 2007. MALE STERI LITY1 is required for tapetal development and pollen wall biosynthesis. Plant Cell 19, 3530–3548. Yephremov, A., Wisman, E., Huijser, P., Huijser, C., Wellesen, K., Saedler, H., 1999. Characterization of the FIDDLEHEAD gene of Arabidopsis reveals a link between adhesion response and cell differentiation in the epidermis. Plant Cell 11, 2187–2201. Yoon, G.M., Dowd, P.E., Gilroy, S., McCubbin, A.G., 2006. Calcium-dependent protein kinase isoforms in Petunia have distinct functions in pollen tube growth, including regulating polarity. Plant Cell 18, 867–878. Zhang, Y., McCormick, S., 2007. A distinct mechanism regulating a pollen-specific guanine nucleotide exchange factor for the small GTPase Rop in Arabidop sis thaliana. Proc. Natl. Acad. Sci. U.S.A. 104, 18830–18835. Zhou, L., Fu, Y., Yang, Z., 2009. A genome-wide functional characterization of Arabidopsis regulatory calcium sensors in pollen tubes. J. Integr. Plant Biol. 51, 751–761. Zinkl, G.M., Preuss, D., 2000. Dissecting Arabidopsis pollen-stigma interactions reveals novel mechanisms that confer mating specificity. Ann. Bot. 85, 15–21. Zinkl, G.M., Zwiebel, B.I., Grier, D.G., Preuss, D., 1999. Pollen-stigma adhesion in Arabidopsis: a species-specific interaction mediated by lipophilic molecules in the pollen exine. Development 126, 5431–5440.
Molecular Mechanisms of Sex Determination in Monoecious and Dioecious Plants
GEORGE CHUCK
Plant Gene Expression Center, United States Department, AgricultureAgriculture Research Service, Albany, CA 94710, United States
I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII. XIII. XIV. XV. XVI.
1
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sex Determination in Cucurbitaceae . . . . . . . . . . . . . . . . . . . . . . . Sex Determination in Melon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Positional Specificity of the Sex Determination Signal . . . . . . . . . . Sex Determination in Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genes Involved in Feminization . . . . . . . . . . . . . . . . . . . . . . . . . . Genes Involved in Masculinization . . . . . . . . . . . . . . . . . . . . . . . . silkless1—A Gene Involved in Perception and Protection from
the Sex Determination Signal . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genetic Interaction Between Sex-Determining Genes . . . . . . . . . . . Molecular Identity of the Ts2 Gene . . . . . . . . . . . . . . . . . . . . . . . . Molecular Identity of Ts1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Are Cytokinins Involved with Ts1 and Ts2 in Lower Floret
Abortion? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Molecular Identity of Class II Tasselseeds-Ts4. . . . . . . . . . . . . . . . Molecular Identity of Ts6, A Dominant Gain-of-Function
Mutant of ids1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Maize Pistil Abortion Pathway may be Under Small RNA
Control. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Relationship Between Maize AP2 Genes and Sex
Determination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
54
56
57
59
60
63
64
66
66
67
69
70
71
73
74
75
Corresponding author: E-mail:
[email protected] Advances in Botanical Research, Vol. 54 Copyright 2010, Elsevier Ltd. All rights reserved.
0065-2296/10 $35.00
DOI: 10.1016/S0065-2296(10)54002-3
54
G. CHUCK
XVII. Hormones, AP2 Genes and MADS Box Genes—What is the Connection To Sex Determination? Evidence and Speculation . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
76 78 78
ABSTRACT In order to enhance outcrossing, plants have evolved a variety of different mechanisms to promote sex determination, a process by which flowers become male or female. Sex determination has evolved multiple times independently, and thus there is no single sex determination mechanism or gene. There is, however, a common set of pathways involving plant hormones that has been uncovered through the cloning of sex determi nation genes from both monocots and dicots. Why these pathways were selected for presents an interesting puzzle that links hormone signalling to multiple developmental pathways including those involving floral homeotic genes as well as small RNAs.
I. INTRODUCTION Sex determination is a process that leads to distinction and separation of structures responsible for production of male and female gametes. The ultimate goal of sex determination is to encourage genetic fitness by promoting out crossing, thus enhancing heterozygosity of alleles and decreasing inbreeding. While plants are generally hermaphroditic, i.e. produce stamens and pistils on a single flower, they have evolved a range of different systems and mechanisms to facilitate outcrossing. These include several mechanisms to prevent selffertilization such as genetic self-incompatibility and heterostyly, in which the stamens and styles of a hermaphroditic flower are of different lengths. A variety of different model systems have been used to uncover the diverse pathways that control sex determination. For example, the genetics of sex determination has been well studied in homosporous ferns such as Ceratopteris richardii, in which loci responsible for generation of male, female and hermaphroditic sex types have been described (Tanurdzic and Banks, 2004). In addition, other basal plants, including bryophytes and lycophytes, also undergo sex determination although they possess distinctly different reproductive structures compared to angiosperms. Within the flowering plants, sex determination has been well studied in Silene latifolia, which has an X and Y sex chromosomal system as found in many animals (Charlesworth, 2008). The great diversity of sexual expression in angiosperms argues against a single mechanism underlying the sex determination process that has likely evolved multiple times independently
SEX DETERMINATION IN MONOCOTS AND DICOTS
55
(Ainsworth et al., 1998). This has been confirmed through the cloning of genes involved in sex determination in monoecious and dioecious plants and finding that they operate in a variety of different pathways. A different sex determination system involves sexual dimorphism, i.e. morphological distinction between male and female flowers through differ ential growth, repression or abortion of sex organs. This process can lead to monoecious plants, i.e. plants that produce unisexual flowers on the same individual, or dioecious plants that produce unisexual flowers on different individuals. In addition, plants may display some combination of both at different times. The common ancestor of all angiosperms was most likely a hermaphroditic flower (Cronquist, 1988), indicating that sex determination is a derived condition imposed on a basic developmental floral plan. Recent studies of sexual dimorphism in two economically important angiosperm species have confirmed this and helped elucidate the molecular mechanism behind sex deter mination in monoecious and dioecious plants. These include members of the dioecious Cucurbitaceae, such as melon and cucumber, in which formation of bisexual and unisexual flowers is dependent on the inheritance of three different loci. By contrast, monoecious maize plants undergo sex determination based on the activity of genes involving at least in two separate pathways for the devel opment of male versus the female flowers (Dellaporta and Calderon-Urrea, 1994; Irish and Nelson, 1989). In both species, sex determination occurs by selective arrest of either the male stamens or the female carpels during various stages of development. Several sex determination loci have now been cloned from both groups of plants and have been shown to involve highly conserved hormone biosynthesis pathways, including those for ethylene, gibberellic acid (GA) and jasmonic acid (JA). While there are no functional parallels between these two pathways in cucurbits and maize, these results point to hormones as being necessary for growth and development of sex-specific floral organs. The goal of this chapter is to place these newly cloned sex determination genes within the context of the overall scheme for plant development. The molecular identity of these sex determination genes should help explain the mechanism underlying early classical experiments on hormonal control of sex determina tion, and may help clarify why hormone pathways were selected for this process in the first place. In addition, this new data should allow connections to be made between the genes responsible for sex-specific floral organ initiation, such as including the well-studied floral homeotic genes, with hormone signalling. The fact that many hormone pathways have intersecting signalling pathways may also explain why different classes of hormones can cause the same sex determi nation effects in different plants. Thus, it is clear that plants have not created unique sex determination genes, but have co-opted existing hormone pathways that influence a variety of other pathways in order to promote outcrossing.
56
G. CHUCK
II. SEX DETERMINATION IN CUCURBITACEAE The cucurbits have been a model system for the analysis of sexual dimorph ism for decades. Cucumber (Cucumis sativus L.) is a monoecious plant in which sex determination is genetically controlled by three loci, F, A and M. The semi-dominant F (female) locus controls the degree of femaleness, while the A (androecious) locus is epistatic to F and is also required for expression of femaleness (Pierce and Wehner, 1990). Combining of the f and a alleles intensifies the androecious (primarily male) phenotype of f (Robinson et al., 1976). The M (andromonoecious) locus is required for selective abortion of the male reproductive organs (Perl-Teves, 1999). The combination of the M-ff alleles makes monoecious flowers, the most common form. M-F- results in gynoecious female flowers, mmF- makes hermaphroditic flowers, while mmff flowers are andromonoecious. A wealth of research implicated cucurbit sex determination genes as functioning in hormone metabolism pathways, specifically, gibberellins (GA) and ethylene. In cucumber, application of GA acts as a masculinizing agent, while ethylene acts as a feminizing agent (Perl-Teves, 1999). Treat ment of monoecious lines with ethylene causes increased female flower development (Iwahori et al., 1970; MacMurray and Miller, 1968; Shannon and De La Guardia, 1969). These data, however, do not simply indicate that each hormone controls each different sex, or that the balance of these two hormones controls sex determination. For example, by treating various dimorphic forms with both hormones and hormone inhibitors, it was shown that ethylene may be the main determinate of sex-specific expression, and may in fact be downstream of the GA signal (Yin and Quinn, 1995). A model explaining the action of the F and M genes posits that F controls the range and gradient of ethylene synthesis along the shoot to promote femaleness, while M encodes a product that perceives the ethylene signal at a local level. Thus, presence of the F allele will allow all flower primordia to produce enough ethylene to produce female floral organs, while the presence of the M allele mediates the ethylene signal to repress male floral organs (Yamasaki et al., 2001). The cloning of the F and M genes from cucumber confirmed this model for sex determination, but raised several questions as well. The clue that ethylene was involved in F gene function led researchers to clone and map genes from cucumber controlling ethylene biosynthesis such as the 1-aminocyclopropane-1 carboxylic acid synthase genes (ACS) (Kamachi et al., 1997) that catalyze one of the first committed steps in the ethylene biosynthetic pathway (Oeller et al., 1991). One member of this gene family was found to genetically map to the F locus and be tightly linked to it (Mibus
SEX DETERMINATION IN MONOCOTS AND DICOTS
57
and Tatlioglu, 2004; Trebitsh et al., 1997). Interestingly, monoecious lines have a single copy of this gene, while gynoecious lines have an additional copy called CS-ACS1G. This duplicated copy was found to likely corre spond to the F locus (Mibus and Tatlioglu, 2004). The expression of both CS-ACS genes were found to be upregulated by ethylene in monoecious and gynoecious plants, but not andromonoecious ones (Kamachi et al., 1997; Yamasaki et al., 2001), thus clearly linking ethylene to female cucumber development. In situ localization of CS-ACS transcript showed high levels of expression in female flower buds in pistil and ovule primordia (Saito et al., 2007). The other player in this pathway, the M locus, was also cloned through a candidate gene and map-based cloning approach and shown to correspond to CS-ACS2. The m genotype was found to be caused by poly morphisms of conserved amino acids within the active site of the enzyme (Boualem et al., 2009; Li et al., 2009). These polymorphisms cause reduced enzyme activity when tested in Escherichia coli (Li et al., 2009) or in vitro (Boualem et al., 2009). Thus, in contrast to earlier models it appears that the product of the M locus does not perceive the ethylene signal or mediate the ethylene response, but may in fact be responsible for production of ethylene itself. Surprisingly, application of exogenous ethylene to mm plants does not appear to rescue the phenotype (Yamasaki et al., 2001), indicating that the M locus may also somehow indirectly affect perception of ethylene.
III. SEX DETERMINATION IN MELON These same ethylene biosynthetic genes appear to also control sex determina tion in melon. In melon, sex determination is dependent on the inheritance of two alleles, a (andromonoecious) and g (gynoecious) (Poole and Grimball, 1939). Wild-type plants are monoecious, having the genotype A-G-, while hermaphrodites are aagg. In addition, andromonoecious plants are aaG while gynoecious plants are AAgg (Kenigsbuch and Cohen, 1989). A chromo some walk to the A locus identified the melon homologue of the CmACS-7 gene as a candidate, which was confirmed by the isolation of ethyl methane sulfonate (EMS)-induced missense alleles that recapitulated the a phenotype (Boualem et al., 2008). Transcripts from the A locus were found in carpel primordia of female and hermaphroditic flowers, much like M, and phyloge netic analysis indicated that the melon A locus is the likely orthologue of the cucumber M gene (Boualem et al., 2009), consistent with their similar func tions. Andromonoecious aa lines contain a single missense mutation in CmACS-7 that causes a decrease in enzyme activity. These data, together with the fact that a mutants are still capable of initiating female floral organs,
58
G. CHUCK
indicate that the A locus is necessary for female development in monoecious lines, but not in hermaphroditic lines. Thus, the A gene itself is not required for carpel initiation and may function to suppress stamen development instead. Finally, the a mutation is monophyletic, and arose once from the wild-type A allele, and was likely the target of selection (Boualem et al., 2008). The recessive g allele causes either unisexual female development in combination with the A gene or hermaphroditic flowers when A is mutated. A chromosome walk to the g locus showed that the mutant is caused by insertion of an hAT transposon near a C2H2 zinc finger transcription factor called CmWIP1 (Martin et al., 2009), several of which are known to be involved in carpel development in Arabidopsis (Crawford et al., 2007). Targeted EMS-induced lesions in this transcription factor were isolated, several of which converted monoecious lines to gynoecy, thus confirming that g does in fact correspond to the CmWIP1 gene. In hermaphroditic and gynoecious plants, the transposon insertion causes spreading of DNA methylation to nearby genes, including g, thereby inactivating them. In young male flowers, G is expressed in carpel primordia that will later abort, but is absent in gynoecious or hermaphroditic lines where carpels do not abort, indicating that G expression in carpels prevents their devel opment (Martin et al., 2009). The expression of G relative to CmACS-7, a.k.a. the A gene, indicated that both were expressed in mutually exclusive patterns, where A is only expressed when G is missing. This indicated some regulatory interaction between the two, where G may be a repressor of A (Martin et al., 2009). Taken together, a model for how sex determination is mediated by the action of the G gene, and its target, the A gene (Fig. 1) posits that G has dual function in repressing carpel development and
Allele G A
G
Carpel
Gene CmWIP1 CmACS-7
A
Stamens
Male flowers
G
Carpel
Function Transcription regulation Ethylene biosynthesis
A
Stamens
Female flowers
G
Carpel
A*
Stamens
Hermaphrodite flowers
Fig. 1. Sex determination in Cucumis melo. Model for sex determination in melon based on the expression of the G and A genes (Martin et al., 2009), where G is a repressor of carpels and A is a repressor of stamens. Grey letters represent inactivation. The A* allele has reduced activity due to an amino acid substitution and thus cannot repress stamens (Boualem et al., 2008).
SEX DETERMINATION IN MONOCOTS AND DICOTS
59
promoting stamen development through repression of A in male monoe cious flowers. In female monoecious flowers, G is not expressed, allowing both carpel initiation and expression of A, which blocks stamen develop ment. In hermaphrodites, lack of G allows carpel development and expres sion of a, but since this variant has an amino acid substitution within the active site (Boualem et al., 2009), it has reduced function and thus cannot repress stamen development.
IV. POSITIONAL SPECIFICITY OF THE SEX
DETERMINATION SIGNAL
It is interesting that genes that control biosynthesis of a diffusible hormone such as ethylene are capable of repressing floral organ development in such a precise manner. The stamens and carpels are usually in adjacent whorls of the flower in most angiosperms, and yet ethylene affects floral development in just one whorl. Such precise activation of ethylene biosynthesis requires either temporally and spatially specific promoters driving the ethylene signal, or limited competency of cells to respond to it in a whorl-specific manner. As floral patterning is known to involve expression of the floral homeotic MADS box genes (Coen and Meyerowitz, 1991), several groups have analysed how this positional information is transduced to actuate sex determi nation. For example, using cucumber, Kater et al. asked whether abortion of floral organs occurs in a position-dependent manner, or whether abortion is a long-range signal that affects reproductive organs regardless of position. This question was explored using the class B homeotic mutant green petals (gp), which is caused by loss of the CUM26 MADS box gene (Kater et al., 2001). In male gp flowers, the third whorl stamens are replaced by carpels that ectopically express the AGAMOUS-related MADS box gene CUM1, thus demonstrating that female floral organs can still develop in flowers that possess the male sex determination program. In transgenic cucumber plants overexpressing CUM1, carpelloid sepals were found in the first whorls and staminoid petals were found in the second whorls in both male and female flowers (Kater et al., 2001). These plants have effectively been converted from monoecious to bisexual hermaphrodites, despite the presence of an active sex determination program. In summary, the sex determination pro cess appears to only operate in the inner reproductive whorls based on their position and not on their organ identity, i.e. in female flowers it is specific to the third whorl and in male flowers it is specific to the fourth whorl. These studies raise the question of how the A and G genes are regulated by positional information to carry out stamen or carpel abortion in melon.
60
G. CHUCK
The above results seem to imply that these genes do not seem to be activated by floral organ identity factors, and since sex determination occurs after initiation of floral organs, it is possible that the involvement of floral homeotic transcription factors is minimal. Nevertheless, recent research on how MADS box genes form higher-order complexes to pro mote identities specific to each whorl (Liu and Mara, 2009) continues to present an attractive mechanism to explain how A and G are activated in such specific locations. In addition, the fact that MADS box genes play a role in fruit ripening points to a direct link with ethylene metabolism, and thus sex determination. Indeed, there is evidence that ethylene may be capable of inducing expression of class C MADS box genes in banana (Liu et al., 2009), indicating that MADS genes may in fact be downstream of the ethylene signal. Other MADS box genes, however, may be upstream of the signal. For example, the tomato AGAMOUS-like gene, TAGL1, is capable of activating transcription of the ACS2 promoter in vitro (Itkin et al., 2009). In summary, there is good evidence that there is a connection between ethylene metabolism and MADS gene function, though the nature of their interaction and how it relates to sex determination remains to be established.
V. SEX DETERMINATION IN MAIZE Maize is a classic system to understand the genetics of monoecy in angiosperms. Maize mutants affecting sex determination have been col lected and analyzed for nearly a century (Neuffer et al., 1997) and have uncovered the hormone-related pathways that control sex determination. The flowers of maize, called florets, have different sexual identities based on their position in the plant. Maize produces two types of inflorescences, the tassel and the ear (Fig. 2A). Upon floral transition, the vegetative meristem is transformed into an inflorescence meristem (IM) that differ entiates into a tassel primordium. Tassels are located in a terminal posi tion and contain male florets, while the ears are located in axillary positions several nodes below and contain female florets. Ears and tassels also differ by the presence of branch meristems (BM), which form side branches in the tassel but are absent in the ear. The IM initiates a series of unique lateral meristems in a sequential pattern (Fig. 2B). First, it initiates ordered rows of spikelet pair meristems (SPM) in a spiral phyllotaxy. Then, each SPM initiates a pair of spikelet meristems (SM) in a distichous phyllotaxy. The SM produces a structure called the spikelet, a highly contracted floral branch system that is the fundamental floral unit of all
61
SEX DETERMINATION IN MONOCOTS AND DICOTS
A
IM
B
C
SPM
SM BM
D
WT E
F
G
ts4 H
SM
I G
SM OG
G
UF
G
LF
Wild type
G
ts4 SM
ids1 G ts4 Ts6
G
F G
F
F F F
G
ts4 in situ
Fig. 2. Development of maize florets and analysis of ts4. (A) Wild-type male tassel inflorescence. (B) SEM of wild-type tassel showing development of lateral meristems. (C) SEM of wild-type tassel spikelet showing development of upper and lower floral meristems. Upper floral meristem is more advanced and is beginning to initiate floral organs. (D) SEM of wild-type ear spikelet showing further development of floral organs in upper floret. Florets are hermaphrodites at this stage. (E) Older ear spikelet showing upper and lower florets with well-developed floral organs. The silk is formed from fused carpels. The gynoecium of the lower floret (arrowhead) has begun to undergo cell death. (F) ts4 tassel showing silks. (G) SEM of ts4 tassel spikelets displaying indeterminate SM with several extra florets. Florets lack gynoecial cell death. (H) In situ hybridization of the ts4 microRNA precursor transcript. Expression is seen as a ring at the base of the SM. (I) Model for floret initiation based on negative regulation of ids1 by the ts4 microRNA. ts4 (red) limits ids1 (blue) activity in order to keep protein levels below a threshold to allow floret initiation in only two locations. When the microRNA is missing in ts4 mutants, or, in microRNA-resistant versions of ids1 such as in Ts6, ectopic IDS1 protein results in extra florets that no longer responds to sex determination signals. IM, inflorescence meristem; SPM, spikelet pair meristem; SM, spikelet meristem; BM, branch meristem; UF, upper floret; LF, lower floret; G, gynoecium; S, stamen; OG, outer glume; si, silk. (See Color Insert.)
62
G. CHUCK
grass species (Clifford, 1987). Each SM initiates two sterile leaves, called glumes, followed by two lemmas, each of which contains a floral meristem (FM) in its axil (Fig. 2C). The FM forms several distinct lateral organs, including a bract leaf called the palea, a pair of lodicules, three stamens, and finally central fused carpels collectively called a silk (Fig. 2D and E). In the ear, the lower floret aborts to produce a single floreted spikelet (Fig. 2E), while in the tassel both florets develop (Cheng et al., 1983). Until the sex determination program initiates, floret development between the male and female inflorescence is nearly identical. Moreover, the ordered rows of lateral primordia found in the inflorescence allow an accurate assessment of developmental time, as the youngest primordia are found at the tip of the inflorescence and gradually become older the farther away they grow. This stereotypical developmental pattern of FM development makes maize an attractive model to study sex determination. Indeed, several classical maize mutations that alter floral development have provided insight into how sexual identity is acquired and maintained in a timely fashion (Dellaporta and Calderon-Urrea, 1994). Sex determination in maize is characterized by gross morphological and histological differences in the stamens and carpel primordia. In the tassel, the early fused carpels, i.e. the gynoecium, begins to disintegrate soon after initiation. This process is evident by histological differences in the gynoecial initial cells that become highly vacuolated, and eventually disintegrate (Cheng et al., 1983). An understanding of why these gynoecial cells abort came from analysing their DNA integrity using DAPI staining (CalderonUrrea and Dellaporta, 1999). Diminished DAPI staining, indicating nuclear fragmentation, was observed in the subepidermal cells of the tassel gynoecial cells in a stereotypical pattern, demonstrating that this tissue-specific cell death is a developmentally programmed process (Calderon-Urrea and Dellaporta, 1999). This pattern of gynoecial-specific nuclear degeneration is conserved in distantly related members of the Andropogoneae grass tribe (Le Roux and Kellogg, 1999) and perhaps may be a defining characteristic of the panicoid family of grasses that include maize. In the ear, the stamen initials of the upper floret abort, while both the stamens and gynoecial initials of the lower floret abort (Fig. 2E). One of the earliest signs of sex determination in the ear is a distinct lack of stamen primordia expression of CYCLIN B (Kim et al., 2007), a gene needed for the G to M phase cell cycle progression. Interestingly, these aborting stamens specifically express at high levels a putative negative regulator of mitosis called WEE1 (Kim et al., 2007). WEE1 encodes a Thr/Tyr protein kinase, but whether this is responsible for the lack of CYCLIN B expression in aborting stamens is unclear. In contrast to the gynoecium of the tassel, the
SEX DETERMINATION IN MONOCOTS AND DICOTS
63
aborting stamen cells of the ear appear to lack signs of DNA fragmentation that would normally reflect a cell death-specific process such as apoptosis. Together, these results indicate that one of the initial steps in specifying female florets in maize is a simple cell cycle block within stamen cells, but whether this is a cause or an effect of the sex determination program remains unclear. After the floret sexes have been specified, secondary sexual characters unrelated to stamen or carpel growth further differentiate the male tassel and female ear inflorescences. This can be seen in the glumes that become green and elongated in the tassel, but reduced and white in the ear. In addition, the inflorescence stem becomes thicker and wider in the ear, but remains thinner in the tassel. These secondary phenotypes appear be under control of downstream targets of the sex determination process, and may be factors selected for in the domestication of maize from teosinte.
VI. GENES INVOLVED IN FEMINIZATION Plant hormones such as GA have long been implicated in sex determination in maize. For example, sustained treatment of maize tassels with varying concentrations of GA has been shown to cause a lack of pistil abortion in maize tassels (Nickerson, 1959). This lack of pistil abortion can also be seen in the tassels of plants grown at low ambient light intensity, and such plants appear to also have higher levels of endogenous GAs (Rood et al., 1980). Comparison of GA-like substances of male and female maize inflorescences showed higher levels present in ears, consistent with GA being a feminizing agent (Rood et al., 1980). Genetic proof supporting this assertion comes from the identification of a distinct class of masculinizing dwarf mutants that are known to have defects in GA biosynthesis. These mutants have low levels of GA and can be rescued by GA spraying (Phinney and Spray, 1982) and include the an1, d1, d2, d3 and d5 mutants (Neuffer et al., 1997), each of which were shown to affect a different step in the synthesis of the gibberellin GA1 (Phinney, 1984). Collectively, these mutants, also known as the andromonoecious dwarves, all produce perfect florets in the ear, i.e. the florets do not undergo stamen abortion in the female inflorescence. The d3 gene was cloned by transposon tagging and shown to encode a cytochrome P450 enzyme (Winkler and Helentjaris, 1995). The an1 gene was also cloned by transposon tagging and found to be chloroplastlocalized plant cyclase involved in ent-kaurene biosynthesis, one of the early steps in GA biosynthesis (Bensen et al., 1995). Thus, levels of
64
G. CHUCK
endogenous GAs clearly play a role in promoting stamen abortion in the ears, but may also play a role in suppressing carpel abortion in the tassel. A second class of andromonoecious dwarves includes the dominant dwarf mutants such as D8 and D9. D8 has normal levels of GA1, and yet cannot be rescued with exogenous GA (Fujioka et al., 1988). Clonal analysis of the Miniplant allele of D8 showed that the mutation is cell-autonomous, con sistent with it being involved in GA perception rather than a diffusible signal (Harberd and Freeling, 1989). The cloning of the D8 gene showed that it encodes the orthologue of the Arabidopsis GIBBERELLIN INSEN SITIVE gene, a member of the DELLA class of nuclear signalling mole cules known to negatively regulate the GA response (Peng et al., 1999). The D8 allele is a dominant negative form of the gene that can no longer respond to the GA and constitutively represses the GA response. Thus, biosynthesis of GA as well as signalling work together as feminizing factors during normal maize floral development.
VII. GENES INVOLVED IN MASCULINIZATION A second group of maize mutants, the tasselseeds, has the opposite effect of the dwarves, causing a lack of pistil abortion within male tassel florets. These mutants fall into two classes based on their phenotypes, called class I and class II (Irish, 1997b). The class I tasselseeds, including tassel seed1 (ts1) and tassel seed2 (ts2) (Nickerson and Dale, 1955), cause a complete transforma tion of the male tassel florets to female. This transformation includes fem inization of secondary sexual characteristics, such as reduction of the glumes and enlargement of the inflorescence stem. The ears of these mutants also display irregular rows of kernels in the ear due to lack of lower floret abortion, indicating that the ts1 and ts2 genes function in both inflorescences regardless of sex. Mosaic analysis showed that wild-type functional tassel sectors could be derived late in ts2 development, after the SM branches from the SPM, indicating that ts2 gene function is only required late in develop ment (DeLong et al., 1993). Transposon excision events of mutable alleles that restore ts2 gene function only occur in subepidermal cell layers, which makes sense in the light of the fact that these cell lineages are the ones that give rise to the gametes (Dawe and Freeling, 1990). Interestingly, subepider mal wild-type sectors in ts2 mutants are capable of conferring a wild-type phenotype on mutant epidermal tissue, a fact that points to the TS2 gene product, or a downstream effector, functioning non-cell autonomously (DeLong et al., 1993). The remaining class I tasselseeds include the dominant mutants Ts3 and Ts5, which display phenotypes similar to ts1 and ts2,
SEX DETERMINATION IN MONOCOTS AND DICOTS
65
though not as severe, and with variable penetrance (Nickerson and Dale, 1955; Neuffer et al., 1997). These latter mutants await further phenotypic and molecular characterization. The class II tasselseeds, including ts4 and the dominant mutant Ts6 mutants, also cause a lack of pistil abortion in the tassel, but display addi tional phenotypes. The recessive ts4 mutant was first described in 1928 as causing irregular branching within the inflorescence as well as feminization of the tassel due to a lack of pistil abortion (Phipps, 1928) (Fig. 2F). The ts4 mutant alters timing and determinacy of the SM by extending the period of FM initiation (Irish, 1997b). Scanning electron microscopy (SEM) of ts4 mutant tassels revealed that the SPM are indeterminate and branch-like (Fig. 2G). In addition, the ts4 SM is indeterminate and produces multiple florets in a random pattern (Fig. 2G). Thus, the ts4 gene has a wide range of functions, specifying male FM sexuality as well as SPM and SM determinacy. The Ts6 mutant phenotype is very similar to ts4, and also causes a feminized tassel with indeterminate branching (Nickerson and Dale, 1955). The dominant Ts6 mutant displays SM indeterminacy and initiates extra FMs before initiating floral organs (Irish, 1997a). In support of this, Ts6 mutant SMs overexpress the meristem-specific homeobox gene knotted1 (Jackson et al., 1994) for longer periods of time, indicating that SM differ entiation is delayed in the mutant. Thus, one can argue that Ts6 is a heterochronic mutant, in which the SM does not terminate in production of a floret, and instead delays differentiation in order to initiate extra florets. Apart from the lack of pistil abortion, these extra florets appear to undergo development in a manner similar to wild type (Irish, 1997b). Clonal analysis showed that Ts6 is likely to function cell autonomously (Johri and Coe, 1983), i.e. encodes a non-diffusible gene product, and thus is unlikely to be directly involved in hormone pathways. Since Ts6 is dominant, one explana tion for its function is that it overproduces a factor that causes meristem indeterminacy. Together, the class II ts4 and Ts6 mutants have been interpreted as a novel class of tasselseeds based on their unique branching phenotypes as well as their genetic interactions with the ts1 and ts2 mutants (Irish et al., 1994). In certain genetic backgrounds, both ts4 and Ts6 mutant ears continue to undergo stamen suppression as in wild type, indicating that the lack of sex determination in these mutants involves a pathway differ ent from the dwarf mutants. The existence of a class of tassel seed mutants that affects both meristem branching as well as sex determination suggests that these seemingly disparate processes share a unique pathway in maize.
66
G. CHUCK
VIII. SILKLESS1—A GENE INVOLVED
IN PERCEPTION AND PROTECTION FROM
THE SEX DETERMINATION SIGNAL
The recessive silkless1 (sk1) mutation affects female floral organ develop ment in the ear. Mutant plants lack silks in the ear due to pistil abortion, rendering the plants female sterile (Jones, 1925). Secondary sexual characters such as glume morphology and stem thickening remain female, and the male florets of the tassel are unaffected, suggesting that the SK1 gene product is required only for pistil development in the ear. Analysis of pistil develop ment in sk1 ears showed nuclear fragmentation of subepidermal cells in a pattern reminiscent of pistil abortion in male florets (Calderon-Urrea and Dellaporta, 1999). Interestingly, the timing of cell death in sk1 female florets correlates with the expression of ts2 mRNA by in situ hybridization (Calderon-Urrea and Dellaporta, 1999), indicating a potential regulatory relationship between the two genes.
IX. GENETIC INTERACTION BETWEEN
SEX-DETERMINING GENES
Combinations between the various sex determination mutants described above have been made to determine how many different pathways are involved, and to place the genes in a genetic hierarchy. Double mutants between d1 and ts1, ts2, ts4 or Ts5 were additive, showing that sex determi nation in the male and female florets operate by separate pathways (Irish et al., 1994). By contrast, ts1/ts2 double mutants resembled the ts1 or ts2 single mutants, which may indicate that the genes function in the same pathway (Irish et al., 1994). Double mutants between the class II tasselseeds with the class I tasselseeds resulted in a synergistic phenotype, where the tassel was enlarged with increased branching compared to the single mutants (Irish et al., 1994). This observation may point to the class I and II tasselseed genes acting in different sex determination pathways that function in parallel. The most enlightening double mutant combinations were found between sk1 and both classes of tasselseeds. The sk1/ts2 double mutant was first reported by Jones (Jones, 1932), and repeated more recently by Irish (Irish et al., 1994). Both researchers found that the presence of sk1 greatly reduced the severity of the ts2 feminization phenotype of the tassel, making it appear nearly normal. In the double mutant ear, pistil development is also normal, except for the fact that the second floret does not abort, much like the ts2
SEX DETERMINATION IN MONOCOTS AND DICOTS
67
single mutant. Thus, ts2 appears to be epistatic to sk1, and is likely to act in the same pathway. This interaction can be explained by models where the wild-type sk1 gene product functions to suppress the function of ts2 within the ear florets (Dellaporta and Calderon-Urrea, 1994). Thus, if one considers the ts2 gene product as a promoter of pistil-specific cell death in wild type, as seen in the tassel and lower ear florets, then sk1 would function to promote pistil development in the upper ear florets by blocking the ts2 death signal. This is consistent with data showing that timing of sk1 nuclear fragmenta tion in mutant pistils coincides with the timing of ts2 gene expression (Calderon-Urrea and Dellaporta, 1999). Double mutant combinations between sk1 and the class II tasselseeds such as ts4 and Ts6 also seem to normalize the feminized tassel (Irish et al., 1994; Mustyatsa and Miku, 1975), but do not seem to affect the branching defects found in the Tasselseed single mutants (G. Chuck unpublished observations). Since Ts6 is dominant, its functional relationship to sk1 is still difficult to discern. ts4, however, may also be epistatic to sk1 much like ts2, but only with respect to the pistil abortion phenotype (Irish et al., 1994; G. Chuck unpublished observations). Since ts4 is a negative regulator of gene expression (discussed below), and not simply a cell death promoter like ts2, this genetic relationship could be explained by the fact that targets of ts4 may indirectly function in promoting ts2 gene expression (G. Chuck unpublished observations).
X. MOLECULAR IDENTITY OF THE TS2 GENE The ts2 gene was cloned by transposon tagging by isolating the flanking sequences from an Ac transposable element inserted into the ts2 gene (DeLong et al., 1993). The ts2 sequence showed similarity to short-chain dehydrogenases, a gene family with a broad range of substrates. ts2 was most similar, however, to a particular subclass known to function as hydroxysteroid dehydrogenases. This fact would seem to indicate that ts2 is involved in metabolism of a steroid-like molecule, or perhaps a GA-like molecule (DeLong et al., 1993). Since the ts2 gene product was predicted to be non-cell autonomous, this molecule would be able to diffuse to adjacent cell layers. While this seemed like an attractive hypothesis, direct biochemical evidence for this model was lacking. However, in vitro studies of the activity of bacterially produced TS2 protein provided further clarification on what its actual enzymatic activity and true substrate might be. TS2 was found to oligomerize as a tetramer, and CD spectroscopy showed that TS2 binds to a nucleotide cofactor such as NAD or NADP as predicted (Wu et al., 2007). Ligand-binding screens were used to test the ability of TS2 to bind to a variety of different substrates, and significant binding was
68
G. CHUCK
found with several different classes of steroid molecules. Surprisingly, no binding was found with plant hormones, including abscisic acid, JA and, more impor tantly, GA (Wu et al., 2007). While this is consistent with the genetic data showing that the dwarf and class I tasselseed mutants operate in separate path ways, it raises the question of how application of GA is able to mimic the ts2 mutant phenotype in the tassel. ts2 RNA expression was found in a very specific tissue in the subepidermal layers of the gynoecium of the male florets late in development (DeLong et al., 1993), at the time when pistil abortion normally occurs. While this expression makes sense in light of the ts2 tassel phenotype, in ears, ts2 RNA was found in the pistil primordia of both the upper and lower florets (Calderon-Urrea and Dellaporta, 1999). This was rather unexpected since ts2 is a putative cell death signal, and the pistil of the upper floret of the ear is functional. A possible explanation for this observation may be that the pistils of the upper floret may be protected from the cell death effects of ts2 by activity of the sk1 gene. In a ts1 mutant background, ts2 in situ expression was undetectable (Calderon-Urrea and Dellaporta, 1999), con sistent with the earlier genetic results showing an epistatic relationship between the two genes. This result would seem to indicate that ts1 is required for ts2 RNA accumulation, implying some kind of regulatory function for ts1, though the cloning of ts1 later proved otherwise. The ts2 gene has a similar function in related grasses such as Eastern gammagrass (Tripsacum dactyloides), demonstrating an ancestral function for ts2 in Zea and Tripsacum. The apical spikelets of Eastern gammagrass are male due to pistil abortion, while the basal spikelets are female. These basal spikelets also show abortion of the lower floret, much like the maize ear. In the gynomonoecious sex form1 (gsf1) mutant, pistil abortion of the apical spikelets and lower floret abortion of the basal spikelets do not occur, mimicking the ts2 mutant phenotype in maize. The TS2 gene from gammagrass was cloned and found to map to the GSF1 locus (Li et al., 1997), and the gsf1 phenotype was found to be associated with an internal deletion of the gene. In support of the conserved genetic function of gsf1, wide crosses between gammagrass and maize were done to create gsf1/ts2 hybrids that failed to complement the mutant phenotype. Taken together, the evidence indicates that the gsf1 mutant phenotype is most likely caused by loss of function of the gammagrass ortho logue of ts2. Moreover, the expression pattern of gsf1 is also highly similar to ts2 and is found in the subepidermal layers of the pistil primordia prior to abortion. Thus, these studies would seem to indicate that the ts2 gene is a conserved target of evolution to separate the sexes in grasses. If the above assertion is true, phylogenetic analysis of bisexual and uni sexual grasses should show that ts2 is under purifying selection in the latter
SEX DETERMINATION IN MONOCOTS AND DICOTS
69
group. Phylogenetic analysis of ts2-related genes from 74 different species showed that true ts2 orthologues only exist in moncots, but is under purify ing selection in both groups (Malcomber and Kellogg, 2006). ts2 is clearly serving an important biological function to be under such selective pressure, but if it is not playing a role in sex determination in some of the unisexual grasses, what is it doing? A survey of ts2 transcription from maize, sorghum and rice showed that ts2 expression is not simply specific to floret primordia, but is universally expressed in a variety of tissues, including roots and leaves (Malcomber and Kellogg, 2006). Perhaps ts2 is not simply a sex-determining gene, but has been co-opted in a specific clade of grasses to function as such, making it likely that ts2 has a more general function during plant develop ment (Malcomber and Kellogg, 2006). Analysis of ts2 homologues in dicots seems to support this view. White campion (Silene latifolia) is a dioecious model for sex determination in which plants produce either male or female flowers on each plant. A ts2 homologue cloned from white campion was analysed to determine whether it was involved in pistil abortion in male flowers. While this gene was expressed only in male flowers, it was not expressed in the aborting gynoecium, but in the tapetum of the anthers instead (Lebel-Hardenack et al., 1997). This expression pattern was con served in the dicot Arabidopsis as well. Since the tapetal cells of dicots undergo a type of cell death since they break down during pollen formation, it may be more useful to view the ts2 gene family as general factors control ling cell death rather than strictly as sex-determining genes.
XI. MOLECULAR IDENTITY OF TS1 Since ts1 appears be necessary for ts2 expression, it was important to determine if ts1 showed homology to regulators of gene expression. The ts1 gene was cloned by chromosome walking, and its molecular identity was confirmed by the analysis of eight mutant alleles. ts1 does not in fact regulate gene expression, but shows extensive homology to class 2 plastid-localized 13-lipoxygenases (Acosta et al., 2009). These genes are known to function in the biosynthesis of jasmonates (Wasternack, 2007). ts1 is widely expressed in a diversity of different tissues, much like ts2. Fluorescently tagging of the protein in onion epidermal cells confirmed the plastid localization of ts1. Surprisingly, ts1 RNA localization was not specific to pistil primordia like ts2, but was found at the base of spikelets far away from the florets (Acosta et al., 2009). Since the genetic results with ts1/ts2 double mutants put both genes in the same pathway, it was expected that their expression patterns should overlap. This unusual expression pattern would seem to indicate that
70
G. CHUCK
the jasmonates produced by ts1 would have to function non-cell autono mously, moving from the base of the spikelet, across numerous cell layers, all the way to the floret. Nevertheless, it is clear that the ts1 gene is involved in synthesis of JA, since direct measurements of JA levels in ts1 mutants was nearly 10-fold lower than in wild type (Acosta et al., 2009). In addition, application of exogenous JA rescued the ts1 phenotype, demonstrating that lack of JA is responsible for the mutant phenotype. In addition, exogenous JA also rescued the ts2 mutant phenotype, a fact that points to the true substrate of ts2 being an intermediate in the JA pathway. Why the earlier in vitro substrate binding studies with TS2 failed to find JA as a candidate is still unclear, but perhaps TS2 recognizes a JA intermediate with a different structure rather than JA itself. An explanation of why ts2 expression is dependent on ts1 may lie in the fact that many JA biosynthetic genes are upregulated by JA, and may positively autoregulate (Acosta et al., 2009). Previous studies in Arabidopsis showed that JAs are needed for proper pollen development and maturation (Ishiguro et al., 2001). The observation that JAs may be important for pistil abortion in maize presents a novel function for this class of hormones, and shows that monocots have co-opted the JA biosynthetic pathway to serve a unique sex determination function. While these findings are clearly important for understanding the precise pathway that initiates the sex determination process, several outstanding questions remain. Both ts1 and ts2 must function in the ear since the mutants fail to abort the lower florets. What is JA doing in the ear, and is it performing a novel function compared to the tassel? What is the ts1 expression pattern in the ear, and does it differ from the tassel where expression was found at the base of spikelets rather than in pistils? Finally, both ts1 and ts2 are widely expressed in a variety of tissues, raising the questions of how tissue specificity of the sex determination is acquired and perceived. A crucial next step is the isolation of the receptor for this sex determination signal, whether it be JA itself or some other intermediate in the JA pathway. Finally, cloning the sk1 gene will be important to under stand how some tissues are protected from the ts1 and ts2 cell death signals.
XII. ARE CYTOKININS INVOLVED WITH TS1 AND TS2 IN LOWER FLORET ABORTION? A different hormone may play a role with ts1 and ts2, specifically in lower floret abortion. An Arabidopsis senescence-induced promoter for the cysteine protease gene SAG12 (Gan and Amasino, 1995) was used to drive
SEX DETERMINATION IN MONOCOTS AND DICOTS
71
the isopentenyl transferase gene (IPT), which is involved in cytokinin bio synthesis. In transgenic plants, ectopic cytokinin produced in senescing tissues might be expected to keep cells alive for longer periods of time as found in dicots (Gan and Amasino, 1995). When put into transgenic maize, the transgene behaved similarly to Arabidopsis, and was induced in senescing leaf tissue (Young et al., 2004). Interestingly, the maize transformants had normal tassels, but lacked lower floret abortion in the ear. Indeed, the lower florets were completely functional and were capable of being fertilized and producing viable embryos (Young et al., 2004). Why cytokinin suppresses lower floret cell death in the ear but not pistil abortion in the tassel or stamen abortion in the ear is still unknown. Perhaps it is possible that cytokinin induces expression of cell death protection signals such as the sk1 gene in the lower floret. It will be interesting to see how cytokinins affect sk1 expression to test this hypothesis.
XIII. MOLECULAR IDENTITY OF CLASS II
TASSELSEEDS-TS4
Cloning of the ts4 locus revealed a second pathway that may have indirect effects on the sex determination pathway. Through chromosome walking, ts4 was identified as a miR172 gene known to regulate the APETALA2 (AP2) family of transcription factors (Chuck et al., 2007b). Five miR172 genes exist in maize, but ts4, otherwise known as zma-MIR172e, is unique, differing by one base within the microRNA-binding site from the other four. Four transposon-induced alleles of ts4 were isolated, three of which con tained Helitron transposon insertions in the promoter, and one derived from targeted mutagenesis using Mutator transposons (Bensen et al., 1995). Taken together, these alleles confirmed that ts4 corresponds to this unique miR172 gene. In Arabidopsis, miR172 is known to negatively regulate its AP2 gene targets at the level of translation (Aukerman and Sakai, 2003; Chen, 2004). The Arabidopsis AP2 gene is a regulator of floral organ identity that causes lack of petals and homeotic transformations of the outermost whorls to carpels in the mutant (Bowman et al., 1989). In normal flowers, AP2 func tions to restrict stem cell formation through negative regulation of WUSCHEL, and defines the expression boundary of the floral organ iden tity genes APETALA3 (AP3) and PISTILLATA (PI) (Zhao et al., 2007). In maize, the equivalent targets of AP2 and their roles in sex determination are unknown. No loss-of-function phenotype has been described for any miR172 gene in any plant species. Thus, the ts4 mutation represents a unique
72
G. CHUCK
opportunity to functionally dissect the negative regulatory relationship between mir172 and its AP2 targets in monocots. zma-MIR172e is expressed in young shoot apices starting at 14 days after planting, and in tassel and ear tissue at high levels. In situ hybridiza tion using the ts4 pri-microRNA transcript as a probe showed expression at the base of the SM above the outer glume near the region where FMs are predicted to form (Chuck et al., 2007b) (Fig. 2H). Expression persists within this domain until SM and FMs are initiated, after which expression can also be seen as a ring at the base of the upper FM prior to anther and carpel initiation as well as the lower FM (Fig. 2I). In summary, ts4 is expressed in a circular domain at the base of the meristems that it affects, indicating that it negatively regulates AP2 transcription factor(s) in this area. A minimum of five AP2 target genes exists for ts4, several of which have been functionally characterized. For example, one potential target is glossy 15 (gl15) that has been shown to be targeted by zma-MIR172 (Lauter et al., 2005). The gl15 gene is involved in controlling the specifica tion of juvenile versus adult leaf fates (Moose and Sisco, 1996). Double mutants between ts4 and gl15, however, were additive, indicating that the two genes act in different pathways. Another potential target of ts4 is the floral patterning gene indeterminate spikelet1 (ids1). The ids1 gene plays a major role in the specification of SM fates by terminating growth of the SM by transforming it into the upper floret. In ids1 mutants, this termination step either does not occur or is delayed, and the SM continues to initiate extra florets (Chuck et al., 1998). If ids1 is in fact negatively regulated by ts4, then loss of ids1 should suppress the ts4 mutant phenotype since ids1 would no longer be inappropriately expressed. The ts4;ids1 double mutant incompletely suppressed both the indeterminate SM branching and sex determination defects of ts4 mutants (Chuck et al., 2007b), indicating that it is a target of ts4. Furthermore, the SM of the rice heterochronic supernumerary bract (snb) mutant also under goes a delayed transition to FM production in a manner very similar to ids1, and is caused by loss of a closely related AP2 transcription factor (Lee et al., 2006). In maize, the orthologue of the snb gene is called sister of indetermi nate spikelet1 (sid1) (Chuck et al., 2008). A phylogenetic analysis suggests that sid1 and ids1 share a common ancestor and might even possess a similar function. This gene duplication could potentially explain the incomplete suppression of ts4 mutants by ids1 mutants (Chuck et al., 2008). Proof that ids1 and sid1 have similar functions comes from the fact that a dramatic enhancement of the ids1 mutant phenotype was seen in double mutants. In the tassel spikelets of the double mutant, no florets were found compared to the ids1 single mutant, which initiates several fertile florets. SEM images of
SEX DETERMINATION IN MONOCOTS AND DICOTS
73
the ids1/sid1 double mutant confirmed that FM were not initiated, and that the mutant SM continuously initiate bracts compared to ids1 alone. This phenotype indicates that both ids1 and sid1 are necessary for the transition of the SM to produce FM. While this phenotype is very different from AP2 mutants in Arabidopsis that continue to initiate FMs ids1/sid1 double mutants also display floral organ identity defects similar to AP2 (Chuck et al., 2008), and ectopically express reproductive organ markers such as ZAG1 and ZMM2 (Schmidt et al., 1993) in lateral organs. This is reminis cent of AP2 mutants in Arabidopsis that ectopically express AGAMOUS LIKE MADS box transcription factors in the outer whorls of the flowers (Drews et al., 1991). Thus, while ids1/sid1 appear to share a subset of common functions with AP2 genes in dicots, in monocots these genes have taken on novel functions during the floral transition.
XIV. MOLECULAR IDENTITY OF TS6, A DOMINANT GAIN-OF-FUNCTION MUTANT OF IDS1 Further proof that ids1 is a target of ts4 came from cloning the Ts6 gene by chromosome walking. Ts6 was mapped to the same chromosomal location as ids1, which raised the possibility that both mutations were in fact alleles of the same gene. This possibility was confirmed by sequencing ids1 from two different Ts6 alleles, both of which contained nucleotide substitutions in the same base pair of the microRNA-binding site relative to the progenitor allele (Chuck et al., 2007b). This microRNA-binding site mutation releases ids1 from negative regulation by ts4, thus explaining the dominant nature of the Ts6 mutant. In summary, the suppression of the ts4 mutant phenotype by ids1 alleles, and the similar phenotypes of Ts6 and ts4 demonstrate that the ids1 (a.k.a. Ts6) gene is a true target gene of ts4. This explains why the mutant phenotypes of ts4 and Ts6 are so similar—the same gene, ids1, is being affected by each mutation. The roles of ids1 and sid1 during floral development are confusing in the light of their gain-of-function and loss-of-function phenotypes. On one hand, the loss-of-function mutant phenotype makes more florets, as seen in ids1. On the other hand, the gain-of-function mutant phenotype for ids1 also makes extra florets, as shown by the ts4 and Ts6 mutant phenotypes. How can the gain-of-function and loss-of-function phenotypes be similar? One explanation is that a negative regulatory relationship exists between ids1 and sid1, similar to how other AP2 genes regulate their homologues in Arabidopsis (Mathieu et al., 2009). This is supported by analysis of sid1 expression in ids1 mutants that showed much higher levels of transcript
74
G. CHUCK
compared to wild type (Chuck et al., 2008). Thus, sid1 appears to be under negative regulation by ids1, and in ids1 mutants, sid1 levels increase to abnormally high levels, which then cause the SM to initiate extra FMs. Therefore, the loss-of-function and gain-of-function phenotypes for ids1 are in fact the opposite of one another—the gain of function making extra florets and the complete loss of function making no florets (Fig. 2I). As will be discussed later, the lack of pistil abortion in the ts4 and Ts6 mutants may be a reflection of the ability of ectopic IDS1 to initiate extra florets outside of the domain where the sex determination signal operates.
XV. THE MAIZE PISTIL ABORTION PATHWAY MAY
BE UNDER SMALL RNA CONTROL
While microRNAs such as miR172 play a role in pistil development by negatively regulating ids1, other small RNAs may do the same. Interestingly, lack of pistil abortion in tassels is often seen in mediator of paramutation1 (mop1) mutants of maize. mop1 mutants are caused by loss of an RNAdependent RNA polymerase gene similar to RDR2 in Arabidopsis (Alleman et al., 2006), resulting in decreased production of siRNAs targeting direct repeats. A recent analysis of microRNA expression in mop1 lines showed that levels of miR156 are reduced, while expression of SPL targets are increased in feminized mop1 inflorescences (Hultquist and Dorweiler, 2008). The miR156 microRNA that targets the SQUAMOSA PROMOTER BINDING LIKE (SPL) class of transcription factors that functions to specify the juvenile phase of plant development in Arabidopsis and maize (Chuck et al., 2007a; Wu and Poethig, 2006). Paradoxically, deep sequencing of small RNAs from mop1 inflorescences demonstrated that the overall levels of most microRNAs, including miR156 and miR172, were increased an average of 5.3-fold (Nobuta et al., 2008). Nevertheless, given that miR156 levels appear to be altered in mop1, how can this lead to a tasselseed phenotype? One answer is that there appears to be a converse negatively regulatory relationship between miR156 and miR172 levels (Chuck et al., 2009), as demonstrated by the fact that high levels of miR156 suppress miR172 (Chuck et al., 2007a). The molecular basis for this relationship may be explained by experiments showing that miR156 targets SPL genes that are necessary for transcription of miR172 (Wu et al., 2009). Thus, if miR156 levels are in fact high in mop1 mutants, they would cause miR172 levels to be low since the levels of SPL factors would also be low. Lowered miR172 levels would thereby mimic a ts4 loss-of-function mutant, though this hypothesis has yet to be tested.
SEX DETERMINATION IN MONOCOTS AND DICOTS
75
Other small RNA mutants also cause tasselseed-like mutant phenotypes. The required to maintain repression 6 (rmr6) mutant of maize was originally isolated as a suppressor of the purple plant paramutant state in maize (Hollick et al., 2005). rmr6 homozygotes display a variety of mutant phenotypes, one of which is a lack of pistil abortion in the tassel (Parkinson et al., 2007). In addition, double mutant analysis between rmr6 and sk1 revealed sk1 to be epistatic. One model explaining this relationship is that rmr6 is a negative regulator of sk1 activity, and that in rmr6 mutants, sk1 is ectopically expressed, thus protecting all pistils in the tassel from the cell death signals derived from the ts1 and ts2 genes (Parkinson et al., 2007). Cloning of the rmr6 gene showed it to be defective in the NRPD1a gene that encodes the largest subunit of the Pol IV polymerase known to be important for production of �24 nucleotide small RNAs (Erhard et al., 2009). The siRNAs produced from Pol IV have been shown to be important for siRNA dependent DNA methylation of heterochromatin (Onodera et al., 2005). In the light of this, one possible scenario for the tasselseed phenotype of rmr6 is that lack of methylation of the sk1 gene causes ectopic expression in the tassel, resulting in a lack of pistil abortion.
XVI. THE RELATIONSHIP BETWEEN MAIZE AP2
GENES AND SEX DETERMINATION
How can AP2 genes such as ids1 affect sex determination in the maize tassel? IDS1 protein is normally found in pistil primordia in ears, and is ectopically expressed in pistils of ts4 tassels (Chuck et al., 2007b). Loss-of function mutants of ids1 have reduced fertility and unfused pistils, while full-loss-of-function mutants in double mutant combinations with sid1 do not make florets or pistils at all. Taken together, these data indicate that ids1 may be important for pistil growth and development. In addition, ids1 may have an important function in negatively regulating AGAMOUS-like MADS box genes, similar to how AP2 functions in Arabidopsis. This is supported by the observation that AGAMOUS-like MADS box genes are ectopically expressed in the lateral organs of the spikelet of ids1/sid1 double mutants (Chuck et al., 2008). Gain-of-function studies of Arabidopsis expressing miR172-resistant forms of AP2 caused meristems to have more stem cells due to prolonged expression of the WUSCHEL and AGAMOUS genes (Zhao et al., 2007). In the light of this, perhaps in ts4 and Ts6 mutants, the lack of downregulation of ids1 causes spikelets to have more stem cells, and thus greater capacity to initiate FMs. Alternatively, the ectopic IDS1 in these tasselseeds may cause downregulation of ZAG1,
76
G. CHUCK
which causes meristem indeterminacy in maize when mutated (Mena et al., 1996). Such indeterminate meristems can then produce extra florets as seen in ts4 and Ts6. Since the extra FMs form in random positions as opposed to a distichous phyllotaxy as in wild type, they may not be in the proper position to receive the sex determination signal, and thus display a tassel seed phenotype. Other floral homeotic mutants in maize provide support for this view. The maize silky (si) mutant also displays silks in the tassel due to homeotic transformation of stamens to carpels, and is caused by loss of function of a B class MADS box gene similar to APETALA3 in Arabidopsis (Ambrose et al., 2000). These ectopic pistils within the male florets of si tassels demonstrate that pistils in an abnormal position are resistant to the sex determination signal. Such models are in line with studies in cucumber showing that the sex determination process acts not on identity of the floral organs, but on their positions within the whorls of the flower (Kater et al., 2001).
XVII. HORMONES, AP2 GENES AND MADS BOX
GENES—WHAT IS THE CONNECTION TO SEX
DETERMINATION? EVIDENCE AND SPECULATION
Hormones, including GA and JA, clearly play a role in sex determination in maize; thus it seems obvious that there must be a connection between their synthesis and homeotic gene function. There is evidence in Arabidopsis that GA may act downstream of AP2, since application of exogenous GA to ap2 1 mutants rescues the mutant phenotype under certain conditions (Okamuro et al., 1997). Other AP2 genes such as LEAFY PETIOLE function as positive regulators of GA-induced germination (Ward et al., 2006). More over, genes that may be repressed by AP2, including the AGAMOUS-like genes, may function as direct regulators of GA metabolic genes (Wang et al., 2004). AGAMOUS itself has been proposed to be a positive regulator of GA biosynthesis by binding to and regulating the GA4 gene, which catalyses one of the last steps in GA biosynthesis (Go´mez-Mena et al., 2005). Feedback regulation may also happen, since GA signalling has also been proposed to regulate AG (Yu et al., 2004), indicating that the levels of hormone and floral homeotic regulators may be in mutual homeostasis. Finally, AG is also a direct regulator of JA biosynthesis, since it binds directly to the promoter of the DAD1 gene, a phospholipase that catalyses the first step in JA biosynth esis (Ito et al., 2007), which may be important for stamen development. In the light of these recent advances, it is clear that there are direct connec tions between sex determination genes and floral homeotic regulators, with
SEX DETERMINATION IN MONOCOTS AND DICOTS
77
hormones being the common link. Untangling these correlations to firmly establish a causal relationship with sex determination presents a major challenge for the future. The results described above can now help elevate the discussion of hor mones, sex determination and floral homeotic regulators over earlier models describing simple hormone phenomenology. Numerous experiments over the years showed that spraying of hormones caused a host of developmen tal abnormalities, including a lack of sex determination. Such results always had to be carefully interpreted since it was never clear which specific pathways were being affected. Now that hormone signalling is well understood, and clear regulatory relationships between hormone path ways and transcription factors appear to exist, hormone phenomenology can be viewed in a new light. For example, one can imagine GAs and AP2 genes converging on AG-like genes in maize to ultimately influence the synthesis of JAs. These JAs may directly feed into the pathway controlled by the ts1 and ts2 genes to produce the pistil abortion signal in male florets. While purely speculative, such models now have the potential to be tested directly. Furthermore, in the light of all the new associations described between hormone pathways, it may even be possible to connect all the various hormones involved in sex determination in cucumber, melon and maize directly with the GA pathway. For example, the late flowering phenotype of the constitutive triple response 1 mutant defective in ethylene signalling can be corrected by spraying exogenous GA (Achard et al., 2008). While this connection between ethylene and GA may seem odd at first glance, it has been shown that ethylene indirectly regulates GA signalling by con trolling levels and stability of the DELLA proteins that modulate the GA response (Achard et al., 2003). In fact, multiple hormone pathways appear to converge on the DELLA signalling pathway (Achard et al., 2006), including JA (Navarro et al., 2008). It is tempting to speculate that all the sex determination hormones, including ethylene, GA and JA, mediate their effects via DELLA signalling, although direct evidence for this has yet to be generated. The groundwork for how sex determination can be mediated through hormones has now been laid, and further systems biology approaches to make these connections stronger will have to be done. System-wide approaches to identify DELLA targets and how they are expressed in male or female flowers will help make putative associations between hormones and sex determination stronger. The next challenge is to deter mine how the sex determination signals are perceived, and how that information is translated to cause organ-specific cell death. Though not
78
G. CHUCK
discussed at length in this review, this field has the promise to have a profound effect on agriculture, especially in the production of hybrid seed. This fact almost guarantees that further research in the area will be done, not only in the model plant systems mentioned here but in diverse crop plants as well.
ACKNOWLEDGEMENTS Thanks to China Lunde for reviewing this manuscript. G.C. is supported by DOE grant DE-AI02-08ER15962 and AFRI grant 2009-03484.
REFERENCES Achard, P., Cheng, H., De Grauwe, L., Decat, J., Schoutteten, H., Moritz, T., et al., 2006. Integration of plant responses to environmentally activated phyto hormonal signals. Science 311, 91–94. Achard, P., Renou, J.P., Berthome´, R., Harberd, N.P., Genschik, P., 2008. Plant DELLAs restrain growth and promote survival of adversity by reducing the levels of reactive oxygen species. Curr. Biol. 18, 656–660. Achard, P., Vriezen, W.H., Van Der Straeten, D., Harberd, N.P., 2003. Ethylene regulates Arabidopsis development via the modulation of DELLA protein growth repressor function. Plant Cell 15, 2816–2825. Acosta, I.F., Laparra, H., Romero, S.P., Schmelz, E., Hamberg, M., Mottinger, J.P., et al., 2009. Tasselseed1 is a lipoxygenase affecting jasmonic acid signaling in sex determination of maize. Science 323, 262–265. Ainsworth, C., Parker, J., Buchanan-Wollaston, V., 1998. Sex determination in plants. Curr. Top. Dev. Biol. 38, 167–223. Alleman, M., Sidorenko, L., McGinnis, K., Seshadri, V., Dorweiler, J., White, J., et al., 2006. An RNA-dependent RNA polymerase is required for paramu tation in maize. Nature 442, 295–298. Ambrose, B.A., Lerner, D.R., Ciceri, P., Padilla, C.M., Yanofsky, M.F., Schmidt, R.J., 2000. Molecular and genetic analyses of the silky1 gene reveal conservation in floral organ specification between eudicots and monocots. Mol. Cell 5, 569– 579. Aukerman, M.J., Sakai, H., 2003. Regulation of flowering time and floral organ identity by a microRNA and its APETALA2-like target genes. Plant Cell 15, 2730–2741. Bensen, R.J., Johal, G.S., Crane, V.C., Tossberg, J.T., Schnable, P.S., Meeley, R.B., et al., 1995. Cloning and characterizatin of the maize An1 gene. Plant Cell 7, 75–84. Boualem, A., Fergany, M., Fernandez, R., Troadec, C., Martin, A., Morin, H., et al., 2008. A conserved mutation in an ethylene biosynthesis enzyme leads to andromonoecy in melons. Science 8, 836–838. Boualem, A., Troadec, C., Kovalski, I., Sari, M., Perl-Treves, R., Bendahmane, A., 2009. A conserved ethylene biosynthesis enzyme leads to andromonoecy in two Cucumis species. PLoS ONE 4, e6144.
SEX DETERMINATION IN MONOCOTS AND DICOTS
79
Bowman, J.L., Smyth, D.R., Meyerowitz, E.M., 1989. Genes directing flower devel opment in Arabidopsis. Plant Cell 1, 37–52. Calderon-Urrea, A., Dellaporta, S.L., 1999. Cell death and cell protection genes determine the fate of pistils in maize. Development 126, 435–441. Charlesworth, D., 2008. Plant sex chromosomes. Genome Dyn. 4, 83–94. Chen, X., 2004. A microRNA as a translational repressor of APETALA2 in Arabi dopsis flower development. Science 303, 2022–2025. Cheng, P.C., Greyson, R.I., Walden, D.B., 1983. Organ initiation and the develop ment of unisexual flowers in the tassel and ear of Zea mays. Am. J. Bot. 70, 450–462. Chuck, G., Candela, H., Hake, S., 2009. Big impacts by small RNAs in plant development. Curr. Opin. Plant Biol. 12, 81–86. Chuck, G., Cigan, M., Saeteurn, K., Hake, S., 2007a. The heterochronic maize mutant Corngrass1 results from overexpression of a tandem microRNA. Nat. Genet. 39, 544–549. Chuck, G., Meeley, R., Hake, S., 1998. The control of maize spikelet meristem fate by the APETALA2-like gene indeterminate spikelet1. Genes Dev. 12, 1145–1154. Chuck, G., Meeley, R., Hake, S., 2008. Floral meristem initiation and meristem cell fate are regulated by the maize AP2 genes ids1 and sid1. Development 135, 3013–3019. Chuck, G., Meeley, R.B., Irish, E., Sakai, H., Hake, S., 2007b. The maize tasselseed4 microRNA controls sex determination and meristem cell fate by targeting Tasselseed6/indeterminate spikelet1. Nat. Genet. 39, 1517–1521. Clifford, H.T., 1987. Spikelet and floral morphology. In: Grass Systematics and Evolution, (Soderstrom, T.R., Hilu, K.W., Campbell, C.S., Barkworth, M.E. (Eds.)), Smithsonian Institution Press, Washington, DC, pp. 21–30. Coen, E.S., Meyerowitz, E.M., 1991. The war of the whorls: genetic interactions controlling flower development. Nature 353, 31–37. Crawford, B., Ditta, G., Yanofsky, M., 2007. The NTT gene is required for transmit ting-tract development in carpels of Arabidopsis thaliana. Curr. Biol. 17, 1101–1108. Cronquist, A., 1988. The Evolution and Classification of Flowering Plants. New York Botanical Gardens, Bronx, New York. Dawe, R.K., Freeling, M., 1990. Clonal analysis of the cell lineage in the male flower of maize. Dev. Biol. 142, 233–245. DeLong, A., Calderon-Urrea, A., Dellaporta, S.L., 1993. Sex determination gene TASSELSEED2 of maize encodes a short-chain alcohol dehydrogenase required for stage-specific floral organ abortion. Cell 74, 757–768. Dellaporta, S.L., Calderon-Urrea, A., 1994. The sex determination process in maize. Science 266, 1501–1505. Drews, G.N., Bowman, J., Meyerowitz, E.M., 1991. Negative regulation of the Arabidopsis homeotic gene AGAMOUS by the APETALA2 product. Cell 65, 991–1002. Erhard, K.F., Stonaker, J.L., Parkinson, S.E., Lim, J.P., Hale, C.J., Hollick, J.B., 2009. RNA polymerase IV functions in paramutation in Zea mays. Science 323, 1201–1205. Fujioka, S., Yamane, H., Spray, C.R., Katsumi, M., Phinney, B.O., Gaskin, P., et al., 1988. The dominant non-gibberellin-responding dwarf mutant (D8) of maize accumulates native gibberellins. Proc. Natl. Acad. Sci. U.S.A. 85, 9031–9035. Gan, S., Amasino, R.M., 1995. Inhibition of leaf senescence by autoregulated production of cytokinin. Science 270, 1986–1988.
80
G. CHUCK
Go´mez-Mena, C., de Folter, S., Costa, M.M., Angenent, G.C., Sablowski, R., 2005. Transcriptional program controlled by the floral homeotic gene AGAMOUS during early organogenesis. Development 132, 429–438. Harberd, N.P., Freeling, M., 1989. Genetics of dominant gibberellin-insensitive dwarfism in maize. Genetics 121, 827–838. Hollick, J.B., Kermicle, J.L., Parkinson, S.E., 2005. Rmr6 maintains meiotic inheri tance of paramutant states in Zea mays. Genetics 171, 725–740. Hultquist, J., Dorweiler, J., 2008. Feminized tassels of maize mop1 and ts1 mutants exhibit altered levels of miR156 and specific SBP-box genes. Planta 229, 99–113. Irish, E., 1997a. Experimental analysis of tassel development in the maize mutant tassel seed 6. Plant Physiol. 114, 817–825. Irish, E.E., 1997b. Class II tassel seed mutations provide evidence for multiple types of inflorescence meristems in maize (Poaceae). Am. J. Bot. 84, 1502–1515. Irish, E.E., Langdale, J.A., Nelson, T.M., 1994. Interactions between tasselseed genes and other sex determining genes in maize. Dev. Genet. 15, 155–171. Irish, E.E., Nelson, T., 1989. Sex determination in monoecious and dioecious plants. Plant Cell 1, 737–744. Ishiguro, S., Kawai-Oda, A., Ueda, J., Nishida, I., Okada, K., 2001. The DEFECTIVE IN ANTHER DEHISCIENCE gene encodes a novel phospholipase A1 catalyzing the initial step of jasmonic acid biosynthesis, which synchronizes pollen maturation, anther dehiscence, and flower opening in Arabidopsis. Plant Cell 13, 2191–2209. Itkin, M., Seybold, H., Breitel, D., Rogachev, I., Meir, S., Aharoni, A., 2009. TOMATO AGAMOUS-LIKE 1 is a component of the fruit ripening regulatory network. Plant J. 60, 1081–1095. Ito, T., Ng, K.H., Lim, T.S., Yu, H., Meyerowitz, E.M., 2007. The homeotic protein AGAMOUS controls late stamen development by regulating a jasmonate biosynthetic gene in Arabidopsis. Plant Cell 19, 3516–3529. Iwahori, S., Lyons, J.M., Smith, O.E., 1970. Sex expression in cucumber plants as affected by 2-chloroethylphosphonic acid, ethylene and growth regulators. Plant Physiol. 46, 412–415. Jackson, D., Veit, B., Hake, S., 1994. Expression of maize KNOTTED1 related homeobox genes in the shoot apical meristem predicts patterns of morpho genesis in the vegetative shoot. Development 120, 405–413. Johri, M.M., Coe, E.H., 1983. Clonal analysis of corn plant development I. The development of the tassel and the ear shoot. Dev. Biol. 97, 154–172. Jones, D.F., 1925. Heritable characters in maize XXIII. Silkless. Heredity 16, 339–341. Jones, D.F., 1932. The interaction of specific genes determining sex in dioecious maize. Proceedings of the Sixth International Congress of Genetics, 2, 104–107. Kamachi, S., Sekimoto, H., Kondo, H., Sakai, S., 1997. Cloning of a cDNA for a 1-aminocyclopropane-1-carboxylate synthase that is expressed during development of female flowers at the apices of Cucumis sativus L. Plant Cell Physiol. 38, 1197–1206. Kater, M., Franken, J., Carney, K., Colombo, L., Angenent, G., 2001. Sex determi nation in the monoecious species cucumber is confined to specific floral whorls. Plant Cell 13, 481–493. Kenigsbuch, D., Cohen, Y., 1989. The inheritance of gynoecy in muskmelon. Genome 33, 317–320. Kim, J.C., Laparra, H., Caldero´n-Urrea, A., Mottinger, J.P., Moreno, M.A., Dellaporta, S.L., 2007. Cell cycle arrest of stamen initials in maize sex determina tion. Genetics 177, 2547–2551.
SEX DETERMINATION IN MONOCOTS AND DICOTS
81
Lauter, N., Kampani, A., Carlson, S., Goebel, M., Moose, S.P., 2005. microRNA172 down-regulates glossy15 to promote vegetative phase change in maize. Proc. Natl. Acad. Sci. U.S.A. 102, 9412–9417. Lebel-Hardenack, S., Ye, D., Koutnikova, H., Saedler, H., Grant, S.R., 1997. Conserved expression of a TASSELSEED2 homolog in the tapetum of the dioecious Silene latifolia and Arabidopsis thaliana. Plant J. 12, 515–526. Le Roux, L., Kellogg, E., 1999. Floral development and the formation of unisexual spikelets in the Andropogoneae (Poaceae). Am. J. Bot. 86, 354–366. Lee, D.Y., Lee, J., Moon, S., Park, S.Y., An, G., 2006. The rice heterochronic gene SUPERNUMERARY BRACT regulates the transition from spikelet mer istem to floral meristem. Plant J. 49, 64–78. Li, D., Blakey, C.A., Dewald, C., Dellaporta, S.L., 1997. Evidence for a common sex determination mechanism for pistil abortion in maize and in its wild relative Tripsacum. Proc. Natl. Acad. Sci. U.S.A. 94, 4217–4222. Li, Z., Huang, S., Liu, S., Pan, J., Zhang, Z., Tao, Q., et al., 2009. Molecular isolation of the M gene suggests that a conserved-residue conversion induces the formation of bisexual flowers in cucumber plants. Genetics 182, 1381–1385. Liu, Z., Mara, C., 2009. Regulatory mechanisms for floral homeotic gene expression. Semin. Cell Dev. Biol., e pub. Liu, J., Xu, B., Hu, L., Li, M., Su, W., Wu, J., et al., 2009. Involvement of a banana MADS-box transcription factor gene in ethylene-induced fruit ripening. Plant Cell Rep. 28, 103–111. MacMurray, A.L., Miller, C.M., 1968. Cucumber sex expression modified by 2-chloroethanephosphonic acid. Science 162, 1397–1398. Malcomber, S.T., Kellogg, E.A., 2006. Evolution of unisexual flowers in grasses (Poaceae) and the putative sex-determination gene, TASSELSEED2 (TS2). New Phytol. 170, 885–899. Martin, A., Troadec, C., Boualem, A., Rajab, M., Fernandez, R., Morin, H., et al., 2009. A transposon-induced epigenetic change leads to sex determination in melon. Nature 461, 1135–1138. Mathieu, J., Yant, L., Mu¨rdter, F., Ku¨ttner, F., Schmid, M., 2009. Repression of flowering by the miR172 target SMZ. PLoS Biol. 7, e1000148. Mena, M., Ambrose, B., Meeley, R.B., Briggs, S.P., Yanofsky, M.F., Schmidt, R.J., 1996. Diversification of C-function activity in maize flower development. Science 274, 1537–1540. Mibus, H., Tatlioglu, T., 2004. Molecular characterization and isolation of the F/f gene for femaleness in cucumber (Cucumis sativus L.). Theor. Appl. Genet. 109, 1669–1676. Moose, S.P., Sisco, P.H., 1996. glossy15, an APETELA2-like gene from maize that regulates leaf epidermal cell identity. Genes Dev. 10, 3018–3027. Mustyatsa, S.I., Miku, V.E., 1975. Phenotypic descriptions of double homozygotes in terms of certain sex genes in maize. Genetika 11, 10–14. Navarro, L., Bari, R., Achard, P., Liso´n, P., Nemri, A., Harberd, N.P., et al., 2008. DELLAs control plant immune responses by modulating the balance of jasmonic acid and salicylic acid signaling. Curr. Biol. 18, 650–655. Neuffer, M.G., Coe, E.H., Wessler, S.R., 1997. Mutants of Maize. Cold Spring Harbor Laboratory Press, Plainview, New York. Nickerson, N.H., 1959. Sustained treatment with gibberellin acid of five different kinds of maize. Ann. Mo. Bot. Gard. 47, 19–37. Nickerson, N.H., Dale, E.E., 1955. Tassel modifications in Zea mays. Ann. MO Bot. Gard. 42, 195–211. Nobuta, K., Lu, C., Shrivastava, R., Pillay, M., De Paoli, E., Accerbi, M., et al., 2008. Distinct size distribution of endogeneous siRNAs in maize: evidence
82
G. CHUCK
from deep sequencing in the mop1-1 mutant. Proc. Natl. Acad. Sci. U.S.A. 105, 14958–14963. Oeller, P., Lu, M., Taylor, L., Pike, D., Theologis, A., 1991. Reversible inhibition of tomato fruit senescence by antisense RNA. Science 254, 437–439. Okamuro, J.K., Szeto, W., Lotys-Prass, C., Jofuku, K.D., 1997. Photo and hormonal control of meristem identity in the Arabidopsis flower mutants apetala2 and apetala1. Plant Cell 9, 37–47. Onodera, Y., Haag, J.R., Ream, T., Nunes, P.C., Pontes, O., Pikaard, C.S., 2005. Plant nuclear RNA polymerase IV mediates siRNA and DNA methylationdependent heterochromatin formation. Cell 120, 613–622. Parkinson, S.E., Gross, S.M., Hollick, J.B., 2007. Maize sex determination and abaxial leaf fates are canalized by a factor that maintains repressed epige netic states. Dev. Biol. 308, 462–473. Peng, J., Richards, D.E., Hartley, N.M., Murphy, G.P., Devos, K.M., Flintham, J.E., et al., 1999. Green revolution’ genes encode mutant gibberellin response modulators. Nature 400, 256–261. Perl-Teves, R., 1999. Male to Female Conversion along the Cucumber Shoot: Approaches to Studying Sex Genes and Floral Development in Cucumis sativus. Bios Scientific Publishers, Oxford, UK. Phinney, B.O., 1984. Gibberellin A1, Dwarfism and the Control of Shoot Elongation in Higher Plants. Cambridge University Press, Cambridge. Phinney, B.O., Spray, C., 1982. Chemical Genetics and the Gibberellin Pathway in Zea Mays L. Plant Growth Substances. Academic Press, New York, pp. 101–110. Phipps, I.F., 1928. Heritable characters in maize. XXXI. Tassel-seed4. J. Hered. 19, 399–404. Pierce, L.K., Wehner, T.C., 1990. Review of genes and linkage groups in cucumber. Hort. Sci. 25, 605–615. Poole, C.F., Grimball, P.C., 1939. Inheritance of new sex forms in Cucumis melo L. J. Hered. 30, 21–25. Robinson, R.W., Munger, H.M., Whitaker, T.W., Bohn, G.M., 1976. Genes of Cucurbitaceae. Hort. Sci. 11, 554–568. Rood, S., Pharis, R., Major, D., 1980. Changes of endogenous gibberellin-like sub stances with sex reversal of the apical inflorescence of corn. Plant Physiol. 66, 793–796. Saito, S., Fujii, N., Miyazawa, Y., Yamasaki, S., Matsuura, S., Mizusawa, H., et al., 2007. Correlation between development of female flower buds and expression of the CS-ACS2 gene in cucumber plants. J. Exp. Bot. 58, 2897–2907. Schmidt, R.J., Veit, B., Mandel, M.A., Mena, M., Hake, S., Yanofsky, M.F., 1993. Identification and molecular characterization of ZAG1 the maize homolog of the Arabidopsis floral homeotic gene AGAMOUS. Plant Cell 5, 729–737. Shannon, S., De La Guardia, M.D., 1969. Sex expression and the production of ethylene induced by auxin in cucumber (C. sativus L.). Nature 223, 186. Tanurdzic, M., Banks, J.A., 2004. Sex-determining mechanisms in land plants. Plant Cell 16, S61–S71. Trebitsh, T., Staub, J.E., O’Neill, S., 1997. Identification of a 1-aminocyclopropane 1-carboxylic acid synthase gene linked to the female (F) locus that enhances female sex expression in cucumber. Plant Physiol. 113, 978–995. Wang, H., Caruso, L.V., Downie, A.B., Perry, S.E., 2004. The Embryo MADS domain protein AGAMOUS-like 15 directly regulates expression of a gene encoding an enzyme involved in gibberellin metabolism. Plant Cell 16, 1206–1219.
SEX DETERMINATION IN MONOCOTS AND DICOTS
83
Ward, J.M., Smith, A.M., Shah, P.K., Galanti, S.E., Yi, H., Demianski, A.J., et al., 2006. A new role for the Arabidopsis AP2 transcription factor, LEAFY PETIOLE, in gibberellin-induced germination is revealed by the misexpres sion of a homologous gene, SOB2/DRN-LIKE. Plant Cell 18, 29–39. Wasternack, C., 2007. Jasmonates: an update on biosynthesis, signal transduction and action in plant stress response, growth and development. Ann. Bot. 100, 681–697. Winkler, R., Helentjaris, T., 1995. The maize Dwarf3 gene encodes a cytochrome P450 mediated early step in gibberellin biosynthesis. Plant Cell 7, 1307–1317. Wu, X., Knapp, S., Stamp, A., Stammers, D.K., Jo¨rnvall, H., Dellaporta, S.L., et al., 2007. Biochemical characterization of TASSELSEED 2, an essential plant short-chain dehydrogenase/reductase with broad spectrum activities. FEBS J. 274, 1172–1182. Wu, G., Park, M.Y., Conway, S.R., Wang, J.W., Weigel, D., Poethig, R.S., 2009. The sequential action of miR156 and miR172 regulates developmental timing in Arabidopsis. Cell 138, 750–759. Wu, G., Poethig, R.S., 2006. Temporal regulation of shoot development in Arabidop sis thaliana by miR156 and its target SPL3. Development 133, 3539–3547. Yamasaki, S., Fujii, N., Matsuura, S., Mizusawa, H., Takahashi, H., 2001. The M locus and ethylene-controlled sex determination in andromonoecious cucumber plants. Plant Cell Physiol. 42, 608–619. Yin, T., Quinn, J.A., 1995. Tests of a mechanistic model of one hormone regulating both sexes in Cucumis sativus (Cucurbitaceae). Am. J. Bot. 82, 1537–1546. Young, T.E., Giesler-Lee, J., Gallie, D.R., 2004. Senescence-induced expression of cytokinin reverses pistil abortion during maize flower development. Plant J. 38, 910–922. Yu, H., Ito, T., Zhao, Y., Peng, J., Kumar, P., Meyerowitz, E.M., 2004. Floral homeotic genes are targets of gibberellin signaling in flower development. Proc. Natl. Acad. Sci. U.S.A. 101, 7827–7832. Zhao, L., Kim, Y., Dinh, T.T., Chen, X., 2007. mir172 regulates stem cell fate and defines the inner boundary of APETALA3 and PISTILLATA expression domain in Arabidopsis floral meristems. Plant J. 51, 840–849.
The Evolution of Floral Symmetry
HE´LE`NE CITERNE,* FLORIAN JABBOUR,†,‡ SOPHIE NADOT†
AND CATHERINE DAMERVAL*,1
*
UMR de Ge´ne´tique Ve´ge´tale, CNRS—Univ Paris-Sud—INRA— AgroParisTech, Ferme du Moulon, 91190 Gif-sur-Yvette, France † Universite´ Paris-Sud, Laboratoire Ecologie, Syste´matique, Evolution, CNRS UMR 8079-AgroParisTech, Orsay, F-91405, France ‡ Institute for Systematic Botany and Mycology, University of Munich, Menzinger Strasse 67, 80638 Munich, Germany
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Definitions of Symmetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Symmetry and Flower Development . . . . . . . . . . . . . . . . . . . . . . . . . A. Establishment of Symmetry at Various Stages During
Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Impact of Growth and Organ Elaboration on Floral symmetry . C. Developmental Trajectories and Flower Symmetry . . . . . . . . . . IV. Evolution of Flower Symmetry. . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Distribution of Symmetry among Extant Angiosperms . . . . . . . B. Emergence of Zygomorphy during Angiosperm Evolution in
Relation to Insect Diversification . . . . . . . . . . . . . . . . . . . . . . . C. Architecture of Flowers and Inflorescences—What is Their
Impact on Floral Symmetry . . . . . . . . . . . . . . . . . . . . . . . . . . . V. The Significance of Symmetry in Plant–Pollinator Interactions . . . . . A. Zygomorphy and Outcrossing Strategies . . . . . . . . . . . . . . . . . .
86
88
93
93
94
95
97
97
98
100
108
109
All authors contributed equally to this review 1
Corresponding author: E-mail:
[email protected] Advances in Botanical Research, Vol. 54 Copyright 2010, Elsevier Ltd. All rights reserved.
0065-2296/10 $35.00
DOI: 10.1016/S0065-2296(10)54003-5
86
H. CITERNE ET AL.
B. Pollinator Preferences and their Perception of Symmetry . . . . . . C. Floral Symmetry and Pollination Syndromes . . . . . . . . . . . . . . . D. Variability of Floral Traits in Zygomorphic and Actinomorphic Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Molecular Bases of Flower Symmetry. . . . . . . . . . . . . . . . . . . . . . . . A. The Floral Symmetry Gene Regulatory Network in Antirrhinum Majus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. CYC-like Genes are Implicated in the Control of Zygomorphy in Diverse Lineages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Genetic Mechanisms Underlying Changes in Floral Symmetry. . D. Evolution of CYC-like Genes: Functional Implications . . . . . . . E. Beyond CYC: Conservation and Divergence of Other Components of the Floral Symmetry Network . . . . . . . . . . . . . VII. Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
112 112 113 115
115 118 121 122 124 126
128
128
ABSTRACT Symmetry is a defining feature of floral diversity. Here we review the evolutionary and ecological context of floral symmetry (adding new data regarding its distribution), as well as the underlying developmental and molecular bases. Two main types of symmetry are recognized: radial symmetry or actinomorphy and bilateral symmetry or zygomor phy. The fossil record suggests that zygomorphy evolved in various lineages 50 MY (million years) after the emergence of angiosperms, coinciding with the diversification of specialized insect pollinators. Among extant angiosperms, zygomorphy is a highly homoplastic trait, and is associated with species radiation thereby satisfying the defini tion of key innovation. The evolution of symmetry may be influenced by clade-specific floral and inflorescence characteristics, possibly indicating different underlying con straints. Ecological studies suggest that zygomorphy may promote cross-fertilization through increased precision in pollen placement on the pollinator’s body. The molecular bases of flower symmetry are beginning to be unravelled in core eudicots, and available evidence underlines the repeated recruitment of CYC2 genes, associated with frequent gene duplications. Future prospects are discussed, emphasizing symmetry as a model character for understanding the evolutionary bases of homoplastic floral traits.
I. INTRODUCTION With more than 260,000 extant species, angiosperms represent about 90% of terrestrial plant biodiversity. The flower, which is a synapomorphy of the group, is a fascinating structure in many respects, having a well-conserved ground plan but tremendous diversity in the size, colour, shape and number of its parts. As a component of human environment it participates in shaping our feeling of beauty.
THE EVOLUTION OF FLORAL SYMMETRY
87
Symmetry is one of the major features taking part in this perception. There are two principal types of floral symmetry, radial and bilateral (Section II), the latter having evolved several times independently in angios perms (Section IV). Bilateral symmetry is therefore a homoplastic trait, which poses fascinating questions concerning the homology of underlying developmental and genetic processes, and the evolutionary forces at work in the different occurrences. Indeed, such recurrent innovations provide researchers with ideal models to address the question of the relative impor tance of historical contingency, physical and developmental constraints and selection, in the course of organismal evolution. As with many other architectural traits, the type of symmetry is often an integral part of a species’ definition, even though more or less important deviations from the characteristic type can be observed in natural popula tions (Section V). The first floral symmetry mutant was described by Linnaeus, based on an atypical sample of Linaria vulgaris harvested by Magnus Zio¨berg in 1742 in Roslagen (Sweden). The flower was radially symmetric with five nectar spurs, contrasting with the normal bilaterally symmetric Linaria flower with just a single spur. Linnaeus called it “Peloria,” after the Greek word for monster. He proposed that it arose through fertilization of a normal Linaria by pollen from an alien species (Linnaeus (1744), discussed in Gustafsson, 1979). Darwin was aware of peloric forms in a number of species, and he remarked that many Labiateae and Scrophulariaceae species are prone to such abnormal shapes. He supposed that pelorism was due to an arrest of development or to reversion. He made reciprocal crosses between peloric and normal snap dragon, and observed that none of the offsprings exhibited peloria; he reported that “the crossed plants, which perfectly resembled the common snapdragon, were allowed to sow themselves, and out of a hundred and twenty-seven seedlings, eighty-eight proved to be common snapdragons, two were in an intermediate condition between the peloric and normal state, and thirty-seven were perfectly peloric, having reverted to the structure of their one grand-parent” (1868). Darwin failed to interpret this segregation (not significantly different from a Mendelian 3:1 segregation for one domi nant gene) and explained the results in the context of his pangenesis hypothesis, which has now been totally dismissed. Hugo De Vries investi gated extensively peloric Linaria; he observed that the typical peloria repro duces five times the ventral part of the normal flower, and suspected that repetitions of other parts could also occur. Indeed, he reported a rare regular variant with a tubular corolla lacking spurs (cited in Gustafsson, 1979). Nowadays, several peloric mutants are commercialized as horticul tural varieties (e.g. in Antirrhinum, Sinningia and orchids).
88
H. CITERNE ET AL.
The elucidation of the genetic bases of peloria in Anthirrhinum and Linaria came more than two centuries after the discovery of peloria, through the work of E. Coen and R. Carpenter’s group (Section VI). Since these groundbreaking results, several research groups have been investigating the genetic origin of symmetry in a growing number of plant families, relying mostly on a candidate gene approach. In recent years, results have been obtained that point to a key role of the TCP gene family in independent occurrences of bilateral symmetry. These advances have prompted several excellent reviews in the last year (Busch and Zachgo, 2009; Hileman and Cubas, 2009; Jab bour et al., 2009a; Preston and Hileman, 2009; Rosin and Kramer, 2009). At the dawn of a new era in evolutionary biology opened up by highthroughput DNA sequencing technologies and functional genomics, it may be of interest to examine what we know about floral symmetry, not only from a genetic but also from evolutionary, developmental and ecological points of view. This review explores these various fields in an attempt to summarize existing knowledge and open new prospects for future research.
II. DEFINITIONS OF SYMMETRY Symmetry is a geometrical concept that can be applied to either living organisms or non-living objects. In biology, rotational and reflection sym metries are generally sufficient to describe the range of forms (Almeida and Galego, 2005; Manuel, 2009). Rotational symmetry is defined as the rotation of an object by an angle of 360˚/n (n>1) that does not change the object. In reflection (flip or mirror) symmetry, an axis can be defined such that two points on a perpendicular line to this axis are at equal distance from it; in other words, this axis defines two mirror images. These two types of sym metry have formed the basis for the discrete categories used to describe flower symmetry. Floral symmetry is generally defined from an “en face” view at anthesis, taking into consideration a planar projection of the flower, which justifies the use of the term “axis” of symmetry. Very few studies of floral symmetry integrate the three-dimensional structure of the flower (Leppik, 1972), where it is more appropriate to talk about “planes” of symmetry. The classification reflecting the three dimensions is complex and unwieldy, and simple defini tions of flower types are generally preferred. Nevertheless, taking into account the three-dimensional structure may be important for understand ing the adaptive value of particular shapes in relation to interactions with pollinators. Although in name floral symmetry refers to the entire structure with all its constitutive parts (sepals, petals, androecium and gynoecium), the
THE EVOLUTION OF FLORAL SYMMETRY
89
descriptions apply primarily to the perianth (particularly the corolla) and sometimes to the androecium. The symmetry of the gynoecium is often described independently from other floral organs. It is generally defined on the basis of ovule placentation, that is, according to internal compartmenta lization since the carpels are often partially or totally fused (with fused ovary walls, styles and stigma). Moreover, the gynoecium is often affected by a reduction in carpel number compared to the merism of the perianth. It is therefore frequently left out when characterizing floral symmetry. Flowers appear predominantly symmetrical and rarely asymmetrical. Among symmetrical flowers, two major types are classically recognized: radial symmetry—also called polysymmetry or actinomorphy (from the Greek word aktis a&: sunray), and bilateral symmetry—also called monosymmetry or zygomorphy (from the Greek word zugon on: a device joining two objects together). Actinomorphy is characterized by both rotational and reflection symmetry. In actinomorphic flowers, all organs of a same type (i.e. sepals, petals or stamens) are identical in shape and size, and evenly distributed around the floral receptacle (Fig. 1B–E). Zygomorphy has only reflection symmetry along a single axis (Fig. 1G–J). The term zygomorphy was first introduced by the German botanist Alexander Braun (1835). Zygomorphic flowers have also been referred to as irregular (Sprengel, 1793), which is misleading in suggesting an absence of symmetry, or as symmetrical (Mohl, 1837; Wydler, 1844), which does not strictly differentiate between radial and bilateral symmetry. Although inappropriate, these terms are still being used in the literature (see, for instance, Coen et al., 1995; Luo et al., 1996). A rarer type of symmetry is disymmetry, where two different orthogonal symmetry axes can be distinguished (Fig. 1N). It occurs in a few clades of magnoliids (Winteraceae (Ronse De Craene et al., 2003)), basal eudicots (e.g. Fumarioideae (pers. obs.) and Eupteleaceae (Ren et al., 2007)), and core eudicots (Oleaceae (Sehr and Weber, 2009), Brassicaceae (Ronse De Craene et al., 2002; Rudall and Bateman, 2002), Begoniaceae (Rudall and Bateman, 2002) and Balanophoraceae (Eberwein et al., 2009)). For most zygomorphic flowers, the single symmetry axis is vertically oriented, passing through the inflorescence apex (adaxial or dorsal side) and the subtending bract (abaxial or ventral side). Consistently, bilaterally symmetrical flowers are also described as dorsoventrally asymmetrical (Carpenter and Coen, 1990; Coen, 1991). Cases of oblique zygomorphy (where the symmetry axis deviates from the dorsoventral position) and transverse zygomorphy (where the symmetry axis is horizontal) occur in some families. Oblique zygomorphy is found in Sapindaceae and Vochysiaceae (Eichler, 1878), Musaceae (Lane, 1955; Schumann, 1900;
90
H. CITERNE ET AL.
(A)
(B)
(C)
(F)
(G)
(H)
(D)
(E)
(I)
(J)
(K)
(L)
(M)
Flag flower (N)
(O)
(P)
Lip flower (Q)
(R)
Fig. 1. Different types of floral symmetry illustrated by examples from monocots and eudicots. Symmetry types are represented in A (actinomorphy), F (zygomorphy), M (disymmetry) and O (asymmetry). Red dotted lines: symmetry axes. B: Hibiscus sp. (Malvaceae, eudicot), C: Aquilegia vulgaris (Ranunculaceae, eudicot), D: Nigella damascena (Ranunculaceae, eudicot), E: Iris pseudacorus (Iridaceae, monocot), G: Corydalis sp. (Papaveraceae s.l., eudicot), H: Orchis militaris (Orchidaceae, monocot), I: Lobelia tupa (Campanulaceae, eudicot), J: Alstroemeria sp. (Alstroemeriaceae, monocot), K: flag flower: Lathyrus sp. (Fabaceae, eudicot), L: lip flower: Lamium galeobdolon (Lamiaceae, eudicot), N: Lamprocapnos spectabilis (Papaveraceae s.l., eudicot), P: Vinca minor (Apocynaceae, eudicot), Q: Tibouchina urvilleana (Melastomataceae, eudicot), R: Strelitzia reginae (Strelitziaceae, monocot). Photographs: F. Jabbour, except 1N: C. Damerval. (See Color Insert.)
Winkler, 1930), Marantaceae (Kunze, 1985), Solanaceae (Tucker, 1999), Moringaceae, Bretschneideraceae (now included in Akaniaceae) (Ronse De Craene et al., 1998, 2000, 2002) and Heliconiaceae (Kirchoff et al., 2009). Transverse zygomorphy is found in Sabiaceae (Wanntorp and Ronse De Craene, 2007) and Papaveraceae (Corydalis and Fumaria). In the latter, however, rotation of the flower pedicel results in vertically oriented flowers at maturity (Fig. 1G).
THE EVOLUTION OF FLORAL SYMMETRY
91
The single symmetry axis of zygomorphic flowers can originate from an unequal distribution of organs at maturity and/or the superimposition of secondary identities on the basic sepal, petal or stamen identity. Examples of unequal distribution of organs of a same identity can be found in Asteridae, where the corolla is organized following a few conserved pat terns, the most common being 2|3 (two petals in the dorsal position|three in the ventral position), 4|1 and 0|5 (Donoghue et al., 1998). The concept of secondary identity translates morphological differentiation (including micromorphological specificities) within a given organ type. For example, in the species Antirrhinum majus (Veronicaceae, Asteridae) where the corolla has an upper lip formed by two fused petals and a lower lip formed by the three other petals (2|3 type), three petal identities (dorsal - single petal—two petals, lateral—two petals and ventral—one petal) are recorded (Corley et al., 2005; Luo et al., 1996). Similarly, in Fabaceae, the standard (dorsal—single petal), wings (lateral—two petals) and keel (ventral—two petals more or less fused) can be considered as having three different petal identities. The combination of unequal distribu tion and secondary identities of petals makes the corolla of many zygo morphic flowers appear bilabiate, leading to the definition of two main types of flowers, namely, lip (or gullet—Faegri and van der Pijl, 1966) and flag (Endress, 1994). The distinction comes essentially from the placement of sexual organs in the upper (lip type) or lower (flag type) part of the flower (see Section V). Lip flowers are essentially found in Lamiales (Fig. 1L), Campa nulales, Zingiberales and Orchidales. Flag flowers are encountered in Faba ceae (Fig. 1K), in Polygalaceae and in Papaveraceae (Proctor et al., 1996). In rare cases such as in tribe Delphinieae (Ranunculaceae), secondary identities develop on spirally inserted organs (Jabbour et al., 2009b). Symmetry is not constant within natural populations, and small devia tions can occur around a main type (see Section V). In addition, within the discrete categories defined above, a quantitative element can be added to classify flowers according to the degree of differentiation or deviation from radial or bilateral symmetry they exhibit. For instance, flowers can be described as almost actinomorphic, slightly zygomorphic or almost zygomorphic (Endress, 1999). There are three main developmental causes for such deviations. First, spiral phyllotaxis implies that organs sharing the same identity are not inserted on a same plane, resulting in flowers that are, strictly speaking asymmetric, even though they can appear actinomorphic or zygomorphic. This is the case in most members of Ranunculaceae, for instance, in tribe Delphinieae with “zygomorphic” flowers, and in Adonis and Nigella that have “actinomorphic” flowers. Second, the curvature of organs or groups of organs can result in a heterogeneous spatial
92
H. CITERNE ET AL.
distribution of organs and can also create imperfectly symmetrical flowers (e.g. both the androecium and gynoecium are curved in Geranium (zygomorphic; Geraniaceae), Solanum (actinomorphic; Solanaceae) and Gladiolus (zygomorphic; Iridaceae). Finally, the degree of zygomorphy can depend on the position of the flower along the inflorescence. In several groups with actinomorphic flowers, the flower can become slightly zygo morphic due to the bending of floral organs when compressed laterally by neighbouring flowers (Endress, 1999). Very few species have asymmetric flowers (Fig. 1P–R). Asymmetric flowers with chaotic organization occur in a few basal angiosperms, where the innermost perianth organs and the stamens are irregularly arranged from inception (e.g. in some Zygogynum species (Winteraceae), Endress, 1999). Asymmetry can also be the result of precise developmental processes that are reproducible among members of the same species (e.g. in Fabaceae, Lamiales, Orchidaceae and Zingiberales). In this case, asymmetry can be found in all floral parts (e.g. in Vochysiaceae (Tucker, 1999)) or in just a single organ type (e.g. in Senna (Caesalpinioideae), where asymmetry affects only the gynoecium). It can be due to a reduction in organ number, such as in Cannaceae and Valerianaceae (e.g. flowers in Canna and Centranthus have a single lateral stamen). Another form of asymmetry is enantiomor phy, an asymmetry polymorphism resulting in flowers of two types that are mirror images. It can be due to the formation of both left- and rightcontorted (sinistrorse or dextrorse) corollas (e.g. Wachendorfia (Haemodor aceae), Endress, 1999, 2001a; Helme and Linder, 1992; Senna (Fabaceae), Marazzi and Endress, 2008; Banksia (Proteaceae), Renshaw and Burgin, 2008), or the deflection of the style to the left or to the right (enantiostyly; see Graham and Barrett, 1995) such as in Wachendorfia paniculata (Hae modoraceae) (Endress, 2001a; Jesson and Barrett, 2002; Jesson et al., 2003; Ornduff and Dulberger, 1978; Tucker, 1996, 1999) and Paraboea rufescens (Gesneriaceae) (Gao et al., 2006). In most enantiostylous species, style deflection is associated with a single pollinating anther opposite the style. Monomorphic enantiostyly, in which individuals exhibit both flower morphs (e.g. Solanum rostratum (Endress, 2006)) has been described in at least 10 monocot and eudicot families, whereas dimorphic enantiostyly, where the two morphs occur on separate plants, has been recorded only in seven species belonging to three monocot families (reviewed in Jesson and Barrett, 2002, 2003). Rarely, only one morph occurs within a species (Endress, 1999) (e.g. Strobilanthinae (Acanthaceae); Moylan et al., 2004). Examination of the developmental process leading to enantiostyly has shown that it resulted from unequal cell division rates at the base of the style (Douglas, 1997; Jesson et al., 2003).
THE EVOLUTION OF FLORAL SYMMETRY
93
III. SYMMETRY AND FLOWER DEVELOPMENT Studies of flower development have benefited from the development of scanning electronic microscopy in the mid 20th century, and various authors have made remarkable contributions to our understanding of flower devel opment and symmetry (e.g. Endress, 1999; Ronse De Craene, 2003; Tucker, 2003a). The developmental processes underlying the different types of floral symmetry at anthesis appear highly diverse and provide information regard ing the evolution of floral diversity.
A. ESTABLISHMENT OF SYMMETRY AT VARIOUS STAGES DURING
DEVELOPMENT
In the first stages of development, phyllotaxis and direction of organ initiation are crucial parameters influencing meristem symmetry (Dong et al., 2005; Tucker, 2002, 2003b). There are two types of phyllotaxis, spiral and whorled. In some taxa there is a combination of both spiral and whorled phyllotaxis, with some organs inserted on a spiral and others on whorls (e.g. Aquilegia (Ranunculaceae) where all organs are inserted on whorls, except sepals (Tucker and Hodges, 2005)). In spiral phyllotaxis, organs are initiated one at a time, with an equal time interval (plastochron) between organs of a same type. In whorled phyllotaxis, initiation of organs of a same type can be synchronous or unidirectional. Usually, and provided that growth is homo geneous, the first organs initiated are the largest at maturity (Goebel, 1905) but there are numerous exceptions (e.g. papilionoid corollas and androecia). An exhaustive list of taxa spanning all major angiosperm clades in which organo genesis follows a unidirectional order is given by Tucker (1999). Zygomorphy can be observed before organ initiation, and persist through out development, or can appear later at various stages of development. For instance, the floral meristem of A. majus has initially the form of a loaf (oval, thus disymmetric), then becomes pentagonal and lastly zygomorphic (Vincent and Coen, 2004). According to the authors, zygomorphy is estab lished in this species at the 15th plastochron among the 58 identified, that is, after 9% of the floral developmental sequence, with the acquisition of dorsal and ventral identities. Another instance of early establishment of zygomorphy during development is Lotus japonicus (Fabaceae) (Feng et al., 2006), where the initiation of floral organs is unidirectional (Dong et al., 2005). Zygomorphy can also be established late in development. The developmental processes underlying late-onset zygomorphy can include heterogeneous growth, heterochrony (a temporal shift from the ancestral condition in a developmental process (Douglas and Tucker, 1996; Rudall and Bateman, 2004; Tucker, 1999))
94
H. CITERNE ET AL.
and/or late elaboration of structures such as glands or spurs (Tucker, 1999). Late zygomorphy appears to be frequent in taxa embedded in groups with predomi nantly actinomorphic flowers (Endress, 1999), such as Ranunculaceae (Jabbour et al., 2009b). In the tribe Delphinieae (Ranunculaceae), it originates from heterogeneous growth of petals and sepals after ontogenesis is completed, and the elaboration of spurs on the two petals and single sepal on the adaxial side (Jabbour et al., 2009b). In Iberis amara, which belongs to Brassicaceae, a family with predominantly actinomorphic flowers, dorsal and ventral petals begin to grow in a heterogeneous way only after the onset of stamen initiation, leading to a zygomorphic mature flower (Busch and Zachgo, 2007). Both early and late zygomorphy occur in the Asteridae. For instance, zygo morphy is apparent from the onset of organ initiation in the subfamily Oroban chaceae, but it is preceded by an actinomorphic stage during development in the Plantaginaceae, Bignoniaceae and Lecythidaceae (Tucker, 1999). B. IMPACT OF GROWTH AND ORGAN ELABORATION ON FLORAL SYMMETRY
Reduction, suppression and differential elaboration of organs determine structural symmetry sensu Rudall and Bateman (2003), as opposed to zygo morphy caused or reinforced by differential petal colouration (Fig. 1J), displacement or unequal organ expansion during development. Organ abortion, which can result from totally suppressed or early arrested growth, is a major determinant of zygomorphy. As a result of heterochrony, an organ can become progressively aborted at an earlier stage until its total suppression (e.g. Li and Johnston, 2000; Mitchell and Diggle, 2005). One or several organ types can be affected. A large monocot group with mostly zygomorphic flowers by organ reduction is Poales sensu lato (Kellogg, 2000; Rudall and Bateman, 2004). The three grass lodicules are hypothesized to be homologous to a single perianth whorl, based on morphological, develop mental and genetic evidence (see, for instance, Schmidt and Ambrose, 1998). Since the dorsal lodicule is absent from most derived grasses (e.g. Hordeum, Pooideae), the presence of only two ventral lodicules renders the grass flowers structurally zygomorphic (Rudall and Bateman, 2004). The female flowers of Stephania dielsiana (Menispermaceae) have a single sepal, two petals and a single carpel, which makes them zygomorphic due to organ reduction, compared to the trimerous actinomorphic male flowers (Wang et al., 2006). In Sinningia cardinalis, A. majus and Rehmannia angulata (all belonging to different families within Lamiales), the dorsal stamen is reduced to a staminode and the degree of reduction increases from the former to the latter, reinforcing the zygomorphic shape of the flower (Endress, 1998). Strong morphological differentiation at the perianth level is often associated
THE EVOLUTION OF FLORAL SYMMETRY
95
with alterations in the androecium, including stamen reduction (staminodes) or even abortion (Rudall and Bateman, 2004). In Gesneriaceae, for instance, the strength of corolla zygomorphy was found to be associated with alteration in stamen number (Endress, 1997). In zygomorphic Proteaceae, a bilabiate perianth is associated with ventral (e.g. Placospermum) or dorsal (e.g. Synaphea) staminodes (Douglas, 1997; Douglas and Tucker, 1996). Differential organ elaboration contributes to bilateral symmetry at matur ity. It includes fusion, curvature (see Section II) and the formation of glands or spurs. A well-known example of differential fusion of organs of a same identity is found in the bilabiate corolla of A. majus, but also in the ligulate flowers of Asteraceae (which can have two reduced and three large fused petals (2|3), three fused petals only (Asteroideae), five fused petals (Cichorioideae) or one reduced and four large fused petals (e.g. Barnadesia) (Ronse De Craene, 2010)). Another example is found in Proteaceae where tepals are fused postgenitally and their partitioning is either equal, resulting in an actinomorphic flower, or unequal, rendering the flower zygomorphic (e.g. Lomatia). Spurs are floral appendages that appear late during development. Their origin is highly diverse, developing on sepals (e.g. Impatiens (Balsaminaceae)), petals (e.g. Viola (Violaceae)), receptacular hypanthia (e.g. Tropaeolum (Tro paeolaceae)), stamen–petal tubes (e.g. Diascia (Scrophulariaceae)) or at the base of the ovary (e.g. Pelargonium). The formation of spurs can affect the symmetry of a flower. When the number of spurs is equal to the merism of the flower (e.g. Epimedium (Berberidaceae), Aquilegia (Ranunculaceae) and Halenia (Gentianaceae)), the flower is actinomorphic. Flowers with a single spur (e.g. Corydalis), or a pair of spurs (e.g. Diascia, Delphinium, Dicentra), are zygomorphic or disymmetric. The presence of a single spur can also determine the orientation of the symmetry axis. The development of a spur in species of Tropaeolum changes the symmetry from oblique to median zygomorphy (Ronse De Craene and Smets, 2001). It has been shown that in Asteridae the evolution of floral symmetry is tightly correlated with that of spurs, and that zygomorphy is a prerequisite for the evolution of single or paired spurs (Jabbour et al., 2008). C. DEVELOPMENTAL TRAJECTORIES AND FLOWER SYMMETRY
Following organ initiation, the major determinants of floral symmetry are organ growth, differentiation and distribution of mature organs. The symmetry of mature flowers can be largely independent of phyllotaxis and organ initiation, and flowers with either whorled (with or without unidirectional initiation) or spiral phyllotaxis can appear actinomorphic or zygomorphic. Figure 2 proposes theoretical examples combining three developmental processes taking part in the establishment of flower symmetry at anthesis,
96
H. CITERNE ET AL. Developmental processes Initiation I
Growth G
Homogeneous Gα
Homogeneous Iα
Heterogeneous Gβ
Homogeneous Gα
Heterogeneous Iβ
Heterogeneous Gβ*
Symmetry of adult flower Differential elaboration D Absent Dα
Iα | Gα | Dα Actinomorphy without change of symmetry during development
Present Dβ
Iα | Gα | Dβ Late zygomorphy
Absent Dα
Iα | Gβ | Dα Zygomorphy with a change of symmetry during development
Present Dβ
Iα | Gβ | Dβ Zygomorphy with a change of symmetry during development
Absent Dα
Iβ | Gα | Dα Early zygomorphy
Present Dβ
Iβ | Gα | Dβ Early zygomorphy
Absent Dα
Iβ | Gβ* | Dα Actinomorphy with a change of symmetry during development
Present Dβ
Iβ | Gβ | Dβ Zygomorphy with changes of symmetry during development
Fig. 2. Theoretical developmental trajectories combining different states for three processes (organ initiation, growth and differential elaboration) resulting in different types of floral symmetry. Two states are considered for each process: synchronous (Ia) or asynchronous (Ib) initiation, homogeneous (Ga) or heterogeneous (Gb) growth, and absence (Da) or presence (Db) of differential elaboration. Combining the two states for the three developmental processes results in eight theoretical outcomes. For example, the Ib | Gb | Da trajectory has an actinomorphic outcome because the heterogeneous growth compensates for the unidirectional initiation of organs (indicated by Gb*). Black circle: floral meristem. Black/gray disk: organ primordium. Black star: elaborated organ. The colour of disks is lighter for organs initiated later. The size of disks is proportional to the primordium growth rate. Stars of different shapes represent differentiated organs. Red line: the single axis of symmetry in zygomorphic stages. (See Color Insert.)
THE EVOLUTION OF FLORAL SYMMETRY
97
namely, initiation, growth and differential elaboration of organs of a same type. Two states are considered here for each process: synchronous (Ia) or asynchronous (Ib) initiation, homogeneous (Ga) or heterogeneous (Gb) growth and absence (Da) or presence (Db) of differential elaboration. The combination of these three processes results in eight developmental trajec tories, producing either zygomorphic or actinomorphic flowers at maturity (Fig. 2). Although this representation oversimplifies complex developmental processes, it serves to illustrate how similar states at maturity may result from different developmental pathways, suggesting that the underlying molecular agents controlling floral symmetry may also be different. Actino morphic flowers can originate from disymmetric or zygomorphic develop mental stages (Endress, 1994; Ronse De Craene and Smets, 1994). Tsou and Mori (2007) report cases where symmetry changes more than once during flower development, such as in Cariniana micrantha (Lecythidaceae) where flowers are successively zygomorphic (sepals initiate asynchronously, Ib), then almost actinomorphic (when sepals are initiated, petals initiate and grow synchronously, Gb), and finally zygomorphic (a hood is derived from the abaxial rim of the ring meristem, Db) (Endress, 1994; Tsou and Mori, 2007). A similar situation is found in the genus Couroupita (Lecythidaceae) in which the upper half of the developing flower is initially retarded at first, resulting in an early zygomorphic stage. The flower becomes actinomorphic when stamens and carpels initiate and then zygomorphic again when the androecium proliferates and forms a tongue-like structure with sterile sta mens (Endress, 1999, Tsou and Mori, 2007). Floral zygomorphy thus relies on complex and potentially numerous developmental trajectories, and this relates to the highly homoplastic nature of this trait in adult flowers. A detailed knowledge of symmetry changes during development is important for (1) understanding symmetry transitions among related species, (2) understanding the repeated establishment of bilateral symmetry across angiosperms and (3) interpreting genetic data underlying these morphological changes.
IV. EVOLUTION OF FLOWER SYMMETRY A. DISTRIBUTION OF SYMMETRY AMONG EXTANT ANGIOSPERMS
Zygomorphy has always been considered a derived trait in angiosperms compared to actinomorphy. Studies that have attempted to infer the ancestral features of the first angiosperms (e.g. Doyle and Endress, 2000; Endress and Doyle, 2009) conclude that the first angiosperms had actinomorphic flowers.
98
H. CITERNE ET AL.
The exact number of transitions toward zygomorphy throughout all angios perms is unknown. The estimated numbers given in papers that deal with the evolution of zygomorphy vary according to the paper (e.g. more than 25 in Cubas, 2004, at least 38 in Zhang et al., 2010). However, it generally reflects the number of families in which zygomorphy is found, but not the actual number of transitions from actinomorphy to zygomorphy. Indeed, such transitions can potentially happen several times within a family. The exact number of transitions can only be obtained through the detailed reconstruc tion of the evolution of the character “floral symmetry” (i.e. character optimization) on a robust and well-resolved phylogenetic tree of angios perms. Variation in the number of families displaying zygomorphy may be due to changes in the classification of angiosperms. We conducted a detailed phylogenetic study of the evolution of zygomorphy in angiosperms using updated phylogenies based on the latest classification (APG3: The Angiosperm Phylogeny Group, 2009 and http://www.mobot.org/MOBOT/ research/APweb). Our results indicate that zygomorphy evolved only once in “basal angiosperms” (a paraphyletic assemblage consisting of all angiosperm taxa that have diverged before the divergence of monocots and eudicots), at least 23 times independently in monocots (see Section IV.C for more detail) and at least 46 times independently in eudicots (see Figs. 4 and 5). The number of independent transitions from actinomorphy to zygomorphy is therefore much higher (at least 70, almost twice the highest number given in the literature) than all previously estimated numbers. Many speciose taxa present strongly zygomorphic flowers (like, for instance, Faboideae, Orchidaceae, Poaceae or the order Lamiales), which is consistent with the hypothesis that zygomorphy could play a positive role in speciation rates. This was rigorously tested using a phylogenetic frame work comparing species richness in sister clades differing in their floral symmetry (Sargent, 2004). In 15 out of 19 sister pairs identified, the lineage with zygomorphic flowers is significantly more diverse than its sister group with actinomorphic flowers, which gives strong support to the hypothesis that zygomorphy is a key innovation. B. EMERGENCE OF ZYGOMORPHY DURING ANGIOSPERM EVOLUTION IN RELATION TO INSECT DIVERSIFICATION
The first known angiosperm remains are pollen grains dated to the Hauter ivian (130–136 Ma, million years ago) (Fig. 3; Feild and Arens, 2007; Friis et al., 2006; Frohlich, 2006). The first fossil of a whorled pentamerous flower with both petals and sepals, considered as a eudicot representative, is recorded in the Cennomanian (Basinger and Dilcher, 1984), while fossil
THE EVOLUTION OF FLORAL SYMMETRY
99
Myr 23 PALEOGENE
Oligocene 34 Eocene 56
Late
65 Maastrichtian 71 Campanian 83 Santonian
Fossils of flower; black: first remains related to Nympheales; white: first cyclic eudicot flower; light gray: fossils ancestral to zygomorphic flowers; dark gray: zygomorphic flower First bee fossil: Melittosphex burmensis
Coniacian
89 Turonian 93 Cennomanian 100 Albian 112 Aptian 125 130
Barremian Hauterivian Valanginian 140 Berriasian 145
JURASSIC
Early
CRETACEOUS
Paleocene
Fossils of monoaperturate (black) and triaperturate (white) pollen grains
Fig. 3. Timescale showing the first appearance of important floral features during angiosperm evolution, based on the fossil record. The vertical black bars indicate two major diversification periods, which coincide with the appearance of new floral traits (from Crepet, 2008; Crepet and Niklas, 2009; Dilcher, 2000; Friis et al., 2001, 2006, 2010; Poinar and Danforth, 2006).
flowers with spirally inserted floral parts are dated to the Barremian–Aptian (Crepet, 2008). Transition to a whorled organization of the flower possibly opened the way for further floral innovations, which appear especially numerous during the Turonian geological stage, coinciding with a period of radiation leading to angiosperm dominance in some floras of the midCretaceous (Crepet, 2008; Crepet and Niklas, 2009; Friis et al., 2010). Bilateral symmetry is thought to have first evolved during this first angios perm radiation, based on Turonian fossils with asymmetric flowers with staminodal nectaries that could be considered “precursors” of zygomorphic flowers, as suggested by their resemblance to the flowers of extant taxa adapted to specialist pollinators (Crepet, 1996, 2008). Remains of clearly zygomorphic flowers, as well as brush flowers (with numerous long stamens), are recorded in Paleocene–Eocene deposits (Fig. 3; Crepet and Niklas, 2009; Dilcher, 2000).
100
H. CITERNE ET AL.
The coevolution of plants and insects has been considered for a long time to be the primary cause of radiation of plants and of some insect groups (correspondence between Saporta and Darwin (1877) cited in Friedman, 2009; Grant, 1949; Grimaldi, 1999; but see Waser, 1998; Gorelick, 2001). Reconstructing the evolution of pollination modes on the phylogeny of extant basal angiosperms clearly indicates that insect pollination is the ancestral state (Hu et al., 2008). Early flowering plants may have been pollinated by a wide diversity of insects such as beetles, primitive moths, various flies and possibly sphecid wasps ancestral to bees (Bernhardt, 2000; Grimaldi, 1999; Hu et al., 2008). Extant bees, that comprise many extant pollinators of zygomorphic flowers, constitute a derived natural group of spheciform wasps (vegetarian wasps) that almost certainly originated in the Mid to Late Cretaceous (Grimaldi, 1999; Poinar and Danforth, 2006). Corbiculate bees (honeybees, bumblebees, orchid bees and stingless bees) extensively diversified in the Early Tertiary (Grimaldi and Engel, 2005). Interestingly, the emergence of floral innovations and derived pollinators co-occurs with the angiosperm radiations of the Turonian (89–93.5 Ma) and the Upper Paleocene Lower Eocene periods (Crepet and Niklas, 2009). In addition, a significant correlation was observed between angiosperm species number and insect family number during Cretaceous–Tertiary geological stages. Even though correlations cannot be considered to necessarily reflect causative influence of one group on the other, it may indicate reciprocal driving mechanisms for diversification (Crepet, 1996). The fossil records thus indicate that zygomorphy evolved in several plant lineages during the same period as the rise of some bee families, supporting the hypotheses of coevolution with these insects as the triggering mechanism for floral symmetry evolution (e.g. Neal et al., 1998). C. ARCHITECTURE OF FLOWERS AND INFLORESCENCES—WHAT IS THEIR IMPACT ON FLORAL SYMMETRY
Perianth symmetry is only one of the numerous floral features that can present variation. Because bilateral symmetry affects the shape of the meristem sometimes from the earliest stages of floral development, the issue of how changes in floral symmetry may have been constrained or canalysed by other features of the flower or the inflorescence architecture during the course of evolution can be raised. When flowers are grouped in inflorescences, they become necessarily constrained by neighbouring flow ers during their development, which may potentially affect flower shape at adult stage. Intrinsic features of flowers such as the number of organ primordia could also be prone to have such an effect, by exerting sterical
THE EVOLUTION OF FLORAL SYMMETRY
101
constraints on flower shape. In the following paragraphs, we examine the relationship between floral symmetry and selected features of flowers and inflorescences.
1. Flower Symmetry and Inflorescences It has been suggested that the symmetry of flowers is somewhat linked to the way they are organized in inflorescences (Coen and Nugent, 1994; Rudall and Bateman, 2010). Inflorescence architecture among angiosperms is very diverse, which has led to a complex and sometimes ambiguous typology (Prenner et al., 2009 and references therein). Two basic types can be distin guished based on the fate of the terminal meristem. In cymose inflorescences, the terminal meristem forms a flower, and inflorescence growth results from the development of one or more lateral axes, which in turn reiterate this pattern (sympodial growth). All axes terminate in a flower. In racemose inflorescences, the terminal meristem promotes inflorescence growth by producing lateral meristems that will produce either flowers or secondary axes reiterating the main axis pattern (monopodial growth). Terminal meristems do not produce flowers but eventually become exhausted. Inflor escences can be simple or compound, associating diversely cymose and/or racemose modules (Prenner et al., 2009). Inflorescence axes are observed in fossil records as early as flowers, but their interpretation is very difficult. The particular architecture of the repro ductive unit of Archaefructus (Barremian–Aptian), now considered to be related to Nympheales, has been interpreted either as a multipartite naked flower with an elongated axis (Sun et al., 2002) or as an ebracteate racemose inflorescence bearing simple unisexual and naked flowers (Friis et al., 2003). Several spike-like or even compound inflorescences from the midCretaceous, densely covered with small flowers, have been found (Friis et al., 2006). Parkin (1914) suggested that the primitive inflorescence type is determinate, meaning in its simplest expression a solitary flower (discussed in Rudall and Bateman, 2010). Morphological analyses of extant “basal” taxa and fossil records in a phylogenetic framework suggest grouping of flowers in inflorescence rather than solitary as the ancestral state, but the ancestral state for inflorescence remains equivocal (Endress and Doyle, 2009). This result apparently comes from the authors’ interpretation of Archaefructus and the inflorescence of Nympheales as racemose, which is a matter of debate (Rudall and Bateman, 2010). Classically, it is stated in the literature that radially symmetric flowers are found in both racemose and cymose inflorescences whereas zygomorphic flowers preferentially occur in racemose inflorescences (Coen and Nugent,
102
H. CITERNE ET AL.
1994; Dahlgren et al., 1985). Indeed, the meristems of grouped flowers are embedded in an asymmetric morphogenetic field defined by the flower subtending bract toward the ventral side and the terminal inflorescence meristem toward the dorsal side (Coen and Nugent, 1994). The existence of different cellular or physiological contexts for terminal and lateral mer istems can be illustrated by terminal peloria that occurs in species that normally produce zygomorphic flowers grouped into racemose inflores cences. In the centroradialis mutant of A. majus, the inflorescence meristem shifts to a floral identity, and the resulting terminal flower is radially sym metric, very similar to lateral ones in the cycloidea mutant (Clark and Coen, 2002). Terminal peloria in eudicots have also been reported in species belonging to the Lamiales, Ranunculaceae (Rudall and Bateman, 2004) and Fumarioideae (Cysticapnos vesicarius, pers. obs.). Morphogenetic gra dients may also account for the different symmetry of central and marginal flowers in derived “flower-like” inflorescences, such as the radiate capitula in Asteraceae, the corymb of I. amara or the umbels in some Apiaceae (e.g. in Daucus carota, pers. obs.). It can be speculated that a prerequisite for the evolution of zygomorphy is the emergence of asymmetric morphogenetic fields in an inflorescence. We examined the relationship between floral symmetry and inflorescence growth pattern (monopodial versus sympodial) by conducting a detailed comparative study of the evolution of both characters in monocots, taking into account the most recent phylogenetic advances in this large clade. Figure 4 presents two mirror phylogenetic trees of the monocots on which flower symmetry (left-hand tree) and inflorescence type (right-hand tree) have been optimized using Maximum Parsimony. It shows that zygomorphy evolved at least 23 times independently from actinomorphy throughout monocots, and not only in the context of a racemose (indeterminate) inflor escence. Zygomorphy evolved together with single flowers in various families, for example, in Arachnites uniflora (Corsiaceae), in Thismia americana (Thismiaceae), in Tecophilaea cyanocrocus (Tecophilaeaceae), in Paphiopedilum appletonianum (Orchidaceae) and in Gethyllis atropurpureum (Amaryllidaceae) and it is found in association with cymose (at least the terminal units) inflorescences in several families of Zingiberales (in Musaceae, Heliconiaceae and Strelitziaceae), in Commelinales (in Haemodoraceae and Commelinaceae), in Liliales (in Alstroemeriaceae) and in Asparagales (in Doryanthaceae and Amaryllidaceae). Flowers in some of these taxa may be quite strongly zygomorphic, like in Zingiberales or Gilliesia (Amaryllidaceae) for example (with organ reduction and synor ganization), indicating that zygomorphy is not necessarily precluded by the sympodial growth of cymose inflorescences. In other words, in monocots,
103
THE EVOLUTION OF FLORAL SYMMETRY Type of symmetry
Type of inflorescence Racemose
Actinomorphy
At least terminal units cymose
Zygomorphy
Panicle Single flowers
Asymmetry No perianth *
*
* ** *
*
*
*
** *
** *
** * ** * ** * * ** * ** *
A
Fig. 4. (A) Mirror trees of monocots showing the evolution of perianth symmetry and inflorescence type, optimized using Maximum Parsimony. Left tree: optimization of perianth symmetry. Several colours on the same branch denote ambiguity in the ancestral state. Right tree: optimization of inflorescence type. Several colours on the same branch denote ambiguity in the ancestral state. Coloured lines refer to the orders of monocots. Asterisks indicate taxa that produce flowers possessing more than six stamens. The topology of the tree was established using information from the Angiosperm Phylogeny website (http://www.mobot.org/MOBOT/research/APweb/) and detailed phylogenies obtained from the literature when necessary. Species represented in this tree were selected according the following criteria: (1) all monocot families are represented by at least one species, and (2) families in which there is variation for at least one of the characters examined are represented by two or more species. Botanical descriptions were mostly obtained from Dahlgren et al. (1985). (B) From left to right and top to bottom: names of the terminal taxa (species) included in the tree. (See Color Insert.)
104
H. CITERNE ET AL.
Costus speciosus (Costaceae)
Hedychium coronarium (Zingiberaceae)
Canna glauca (Cannaceae)
Maranthochloa cuspidata (Maranthaceae)
Heliconia magnifica (Heliconiaceae)
Orchidenta maxillarioides (Lowiaceae)
Ravenala madagascariensis (Strelitziaceae)
Phenakospermum guinanense (Strelitziaceae)
Strelitzia reginae (Strelitziaceae)
Musa acuminata (Musaceae)
Haemodorum corymbosum (Haemodoraceae)
Anigozanthos flavidus (Haemodoraceae)
Wachendorfia paniculata (Haemodoraceae)
Heteranthera callifolia (Pontederiaceae)
Pontederia lanceolata (Pontederiaceae)
Philydrum lanuginosum (Phylidraceae)
Hanguana malayana (Hanguanaceae)
Tradescantia sillamontana (Commelinaceae)
Commelina forskalaei (Commelinaceae)
Dasypogon bromeliifolius (Dasypogonaceae)
Poa trivialis (Poaceae)
Oryza sativa (Poaceae)
Ochlandra stridula (Poaceae)
Arundinaria gigantea (Poaceae)
Imperata cylindrica (Poaceae)
Joinvillea plicata (Joinvilleaceae)
Ecdeicolea monostachya (Ecdeiocoleaceae)
Flagellaria guineensis (Flagellariaceae)
Dapsilanthus disjunctus (Restionaceae)
Centrolepis fascicularis (Centrolepidaceae)
Anarthria prolifera (Anarthriaceae)
Lipocarpha occidentalis
Evandra aristata (Cyperaceae)
Scirpus californicus (Cyperaceae)
Carex praeclara (Cyperaceae)
Distichia sp. (Juncaceae)
Juncus castaneus (Juncaceae)
Thurnia sphaerocephala (Thurniaceae)
Mayaca fluviatilis (Mayacaceae)
Eriocaulon taishanense (Eriocaulaceae)
Eriocaulon decangulare (Eriocaulaceae)
Abolboda linearifolia (Eriocaulaceae)
Orectanthe sceptrum (Xyridaceae)
Xyris lacerata (Xyridaceae)
Rapatea paludosa (Rapateaceae)
Pitcairnia xanthocalyx (Bromeliaceae)
Dyckia remotifolia (Bromeliaceae)
Billbergia nutans (Bromeliaceae)
Typha latifolia (Typhaceae)
Sparganium erectum (Sparganiaceae)
Retispatha dumetosa (Arecaceae)
Nypa fruticans (Arecaceae)
Caryota mitis (Arecaceae)
Phoenix dactylifera (Arecaceae)
Phytelephas macrocarpa (Arecaceae)
Phytelephas aequatorialis (Arecaceae)
Synechantus warscewiczianus (Arecaceae)
Cocos nucifera (Arecaceae)
Howea balmoreana (Arecaceae)
Dypsis lutescens (Arecaceae)
Dypsis lantzeana (Arecaceae)
Dypsis mirabilis (Arecaceae)
Neuwiedia inae (Orchidaceae)
Apostasia odorata (Orchidaceae)
Paphiopedilum appletonianum (Orchidaceae)
Vanilla planifolia (Orchidaceae)
Ophrys insectifera (Orchidaceae)
Eulophia andamanensis (Orchidaceae)
Aspidistra dodecandra (Asparagaceae)
Asparagus officinalis (Asparagaceae)
Yucca baccata (Asparagaceae)
Hosta japonica (Asparagaceae)
Aphyllanthes monspeliensis (Asparagaceae)
Sowerbaea juncea (Asparagaceae)
Lomandra insularis (Asparagaceae)
Trichlora lactea (Amaryllidaceae)
Miersia chilensis (Amaryllidaceae)
Leucocoryne purpurea (Amaryllidaceae)
Gillesia graminea (Amaryllidaceae)
Solaria miersiodes (Amaryllidaceae)
Allium vineale (Amaryllidaceae)
Gethyllis atropurpureum (Amaryllidaceae)
Gethyllis ciliaris (Amaryllidaceae)
Sprekelia formosissima (Amaryllidaceae)
Habranthus robustus (Amaryllidaceae)
Lycoris aurea (Amaryllidaceae)
Galanthus nivalis (Amaryllidaceae)
Asphodelus aestivus (Xanthorrhoeaceae)
Haworthia integra (Xanthorrhoeaceae)
Hemerocallis fulva (Xanthorrhoeaceae)
Simethis planifolia (Xanthorrhoeaceae)
Phormium cookianum (Xanthorrhoeaceae)
Arnocrinum gracillimum (Xanthorrhoeaceae)
Xanthorrhoea preissii (Xanthorrhoeaceae)
Xeronema callistemon (Xeronemataceae)
Moraea aristata (Iridaceae)
Isophysis tasmanica (Iridaceae)
Iris germanica (Iridaceae)
Geosiris aphylla (Iridaceae)
Aristea biflora (Iridaceae)
Fig. 4.
(Continued)
Gladiolus segetum (Iridaceae)
Crocosmia masoniorum (Iridaceae)
Crocosmia paniculata (Iridaceae)
Freesia laxa (Iridaceae)
Crocus angustifolius (Iridaceae)
Romulea citrina (Iridaceae)
Doryanthes palmeri (Doryanthaceae)
Doryanthes ensifolia (Doryanthaceae)
Tecophilaea cyanocrocus (Tecophilaeaceae)
Zephyra elegans (Tecophilaeaceae)
Conanthera bifolia (Tecophilaeaceae)
Cyanella lutea (Tecophilaeaceae)
Cyanella hyacinthoides (Tecophilaeaceae)
Cyanastrum johnstonii (Tecophilaeaceae)
Ixiolirion montanum (Ixioliriaceae)
Borya spetentrionalis (Boryaceae)
Astelia pumila (Asteliaceae)
Lanaria plumosa (Lanariaceae)
Pauridia longituba (Hypoxidaceae)
Curculigo latifolia (Hypoxidaceae)
Curculigo racemosa (Hypoxidaceae)
Hypoxis decumbens (Hypoxidaceae)
Blandfordia grandiflora (Blandfordiaceae)
Calochortus nuttallii (Calochortaceae)
Corsia unguiculata (Corsiaceae)
Arachnites uniflora (Corsiaceae)
Smilax aspera (Smilacaceae)
Heterosmilax japonica (Smilacaceae)
Heterosmilax longiflora (Smilacaceae)
Heterosmilax seisuiensis (Smilacaceae)
Gagea lutea (Liliaceae)
Ripogonum scandens (Ripogonaceae)
Philesia magellanica (Philesiaceae)
Paris quadrifolia (Melanthiaceae)
Chamaelirium luteum (Melanthiaceae)
Chionographis chinensis (Melanthiaceae)
Veratrum album (Melanthiaceae)
Campynema lineare (Campynemataceae)
Colchicum automnale (Colchicaceae)
Petermannia cirrosa (Petermanniaceae)
Luzuriagaria radicans (Luzuriagaceae)
Bomarea pardina (Alstroemeriaceae)
Alstroemeria aurantiaca (Alstroemeriaceae)
Asplundia multistaminata (Cyclanthaceae)
Pandanus candelabrum (Pandanaceae)
Croomia pauciflora (Stemonaceae)
Pentastemona egregia (Stemonaceae)
Triuris hyalina (Triuridaceae)
Barbacenia purpurea (Velloziaceae)
Vellozia prolifera (Velloziaceae)
Dioscorea communis (Dioscoreaceae)
Dioscorea melanophyma (Dioscoreaceae)
Dioscorea convolvulacea (Dioscoreaceae)
Trichopus zeylanicus (Dioscoreaceae)
Stenomeris cumingiana (Dioscoreaceae)
Burmannia madagascariensis (Burmanniaceae)
Thismia americana (Thismiaceae)
Afrothismia pachyantha (Thismiaceae)
Oxygyne triandra (Thismiaceae)
Narthecium ossifragum (Nartheciaceae)
Japonolirion osense (Petrosaviaceae)
Petrosavia stellaris (Petrosaviaceae)
Potamogeton pectinatus (Potamogetonaceae)
Althenia filiformis (Potamogetonaceae)
Zannichellia palustris (Potamogetonaceae)
Zostera marina (Zosteraceae)
Posidonia oceanica (Posidoniaceae)
Cymodocea nodosa (Cymodoceaceae)
Ruppia spiralis (Ruppiaceae)
Maundia triglochinoides (Juncaginaceae)
Triglochin maritimum (Juncaginaceae)
Lilaea scilloides (Juncaginaceae)
Aponogeton hexatepalus (Apotonogetonaceae)
Aponogeton proliferus (Apotonogetonaceae)
Aponogeton madagascariensus (Apotonogetonaceae)
Aponogeton distachyos (Apotonogetonaceae)
Scheuchzeria palustris (Scheuchzeriaceae)
Sagittaria platyphylla (Alismataceae)
Wiesneria triandra (Alismataceae)
Hydrocleys nymphoides (Limnocharitaceae)
Limnocharis flava (Limnocharitaceae)
Butomopsis latifolia (Limnocharitaceae)
Halophila ovalis (Hydrocharitaceae)
Thalassia testudinum (Hydrocharitaceae)
Vallisneria americana (Hydrocharitaceae)
Hydrilla verticillata (Hydrocharitaceae)
Najas marina (Hydrocharitaceae)
Egeria densa (Hydrocharitaceae)
Elodea nuttallii (Hydrocharitaceae)
Stratiotes aloides (Hydrocharitaceae)
Hydrocharis morsus-ranae (Hydrocharitaceae)
Limnobium spongia (Hydrocharitaceae)
Butomus umbellatus (Butomaceae)
Pleea tenuifolia (Tofieldiaceae)
Tofieldia pusilla (Tofieldiaceae)
Lemna minor (Araceae)
Cryptocoryne crispatulata (Araceae)
Pistia stratiotes (Araceae)
Pothos chinensis (Araceae)
Anthurium ramoncaracasii (Araceae)
Acorus calamus (Acoraceae)
Zingiberales
Commelinales Dasypogonaceae
Poaceae
Arecaceae
Hosta japonica (Asparagaceae)
Liliales
Pandanales
Dioscoreaceae Petrosaviales
Alismatales
Acorales
B
THE EVOLUTION OF FLORAL SYMMETRY
105
bilateral symmetry does not occur exclusively in flowers produced by lateral meristems. In eudicots, zygomorphic flowers are generally assembled in racemose inflorescences. A rare exception is the cymose inflorescence of the zygomorphic Capnoides sempervirens (Fumarioideae), even though zygomorphy is more fluctuant in the terminal flower than in lateral flowers (pers. obs.).
2. Floral Constraints on the Evolution of Symmetry A previous study exploring the relationships between floral symmetry, merism, number of stamens and presence of spurs in Ranunculales, the earliest-diverging order in the eudicots, showed that zygomorphy evolved three times independently and in very different architectural contexts in this group (Damerval and Nadot, 2007). Another study conducted in the large Asterid clade, where numerous transitions toward zygomorphy have occurred (sometimes followed by reversals to actinomorphy) has shown that zygomorphy is almost never associated with polyandry (i.e. a number of stamens higher than twice the merism) in this derived eudicot clade (Jabbour et al., 2008). This study highlights the fact that floral symmetry may not evolve completely independently from other floral features. In particular, it suggests that an increase in stamen number could impede the dorsoventralization of the flower. A similar situation was found in monocots (Fig. 4) in which only one co-occurrence of polyandry (defined here as more than six stamens, six being twice the most widespread type of merism in monocots) and zygomorphy is observed, within the genus Aponogeton from the basal order Alismatales. In Rosids, however, several co-occurrences of zygomorphy and polyan dry are recorded (Fig. 5). Among the 11 (at least) transitions toward zygomorphy and the more than 25 transitions toward polyandry (defined as over twice the merism) recorded in the phylogenetic tree of rosid families, co-occurrences of both character states are observed five times. They are found in Emblingiaceae (which produce dimerous flow ers with eight or nine stamens), in Begoniaceae, which have dimerous disymmetric rather than truly zygomorphic flowers, in Resedaceae, Cleomaceae (in which however, most zygomorphic genera have flowers with few stamens) and in Chrysobalanaceae. Truly zygomorphic flowers with numerous stamens are found only in Resedaceae and Chrysobala naceae, suggesting that the establishment of zygomorphy might be con strained in a polyandrous context, like in the Asterids. Furthermore, like in the Asterids the presence of spurs (here a single spur) is conditioned to zygomorphy (Fig. 5). The main difference lies in the fact that in
106
H. CITERNE ET AL.
Type of perianth symmetry
Number of stamens
Polysymmetry
Twice merism or less More than twice merism (polyandry)
Monosymmetry No perianth
Variable
* * *
*
**
*
*
*
A
Fig. 5. (A) Mirror trees of Rosids showing the evolution of perianth symmetry and the state of the androecium (number of stamens) relatively to the merism, optimized using Maximum Parsimony. Left tree: optimization of perianth symmetry. Several colours on the same branch denote ambiguity in the ancestral state. Right tree: optimization of the state of the androecium (number of stamens). Several colours on the same branch denote ambiguity in the ancestral state. Coloured lines refer to the orders of Rosids. Asterisks indicate taxa that produce spurred flowers. The topology of the tree was established using information from the Angiosperm Phylogeny website (http://www.mobot.org/MOBOT/research/APweb/). All families are included and represent the terminal taxa of the tree. When zygomorphy is present in addition to actinomorphy within a family, it concerns closely related taxa, therefore the number of transitions at the family level is a good proxy for the actual number of transitions. Botanical descriptions were obtained from the AP website, from eFloras (http://www.efloras. org/), and from Delta (http://delta-intkey.com/angio/www/index.htm). (B) From left to right and top to bottom: names of the terminal taxa (families) included in the tree. (See Color Insert.)
107
THE EVOLUTION OF FLORAL SYMMETRY Celastraceae Lepidobotryaceae Huaceae Oxalidaceae Connaraceae Brunelliaceae Cephalotaxaceae Elaeocarpaceae Cunoniaceae Linaceae Irvingiaceae Ixonanthaceae Humiriaceae Pandaceae Ochnaceae Hypericaceae Podostemaceae Calophyllaceae Bonnetiaceae Clusiaceae Centroplacaceae Malpighiaceae Elatinaceae Peraceae Rafflesiaceae Euphorbiaceae Picrodendraceae Phyllanthaceae Balanopaceae Chrysobalanaceae Euphronaceae Dichapetalaceae Trigoniaceae Caryocaraceae Achariaceae Goupiaceae Lacistemataceae Salicaceae Violaceae Passifloraceae Putranjivaceae Lophopyxidaceae Ctenolophonceae Erythroxylaceae Rhizophoraceae Fabaceae-Faboideae Fabaceae-Mimosoideae Fabaceae-Caesalpinioideae Suraniaceae Polygalaceae Quillajaceae Rosaceae Rhamnaceae Eleagnaceae Dirachnaceae Barbeyaceae Ulmaceae Cannabaceae Moraceae Urticaceae Corynocarpaceae Coriariaceae Cucurbitaceae Tetramelaceae Begoniaceae Datiscaceae Anisophylleaceae Nothofagaceae Fagaceae Myricaceae Rhoipteleaceae Juglandaceae Ticodendraceae Betulaceae Casuarinaceae Geraniaceae Melianthaceae Francoaceae Ledocarpaceae Vivianaceae
Fig. 5.
(Continued)
Combretaceae Onagraceae Lythraceae Penaeaceae Alzateaceae Crypteroniaceae Melastomataceae Vochysiaceae Myrtaceae Stachyceraceae Crossosomataceae Guatemalaceae Staphyleaceae Geissolomataceae Ixerbaceae Strasburgeriaceae Aphloiaceae Picramniaceae Nitrariaceae Kirkiaceae Burseraceae Anacardiaceae Simaroubaceae Meliaceae Rutaceae Sapindaceae Biebersteiniaceae Gerrardinaceae Tapisciaceae Dipentodontaceae Neuradaceae Thymeleaceae Sphaerosepalaceae Bixaceae Dipterocarpaceae Sarcolaenaceae Cistaceae Cytinaceae Muntingiaceae Malvaceae Akaniaceae Tropaeolaceae Moringaceae Caricaceae Setchellanthaceae Limnanthaceae Koeberliniaceae Bataceae Salvadoraceae Emblingiaceae Pentadiplandraceae Gyrostemonaceae Resedaceae Tovariaceae Capparaceae Brassicaceae Cleomaceae Krameriaceae Zygophyllaceae Vitaceae Peridiscaceae Cercidiphyllaceae Daphniphyllaceae Hamamelidaceae Altingiaceae Paeoniaceae Crassulaceae Aphanopetalaceae Tetracarpaceae Penthoraceae Haloragaceae Iteaceae Grossulariaceae Saxifragaceae Dilleniaceae Gunneraceae Myrothamnaceae
Celastrales Oxalidales
Malpighiales
Fabales
Rosales
Cucurbitales
Fagales
Melianthales Myrtales
Crossosomatales Picramniales Sapindales Huerteales Malvales
Brassicales
Zygophyllales Vitales
Saxifragales
Dillenialese Gunnerales
B
108
H. CITERNE ET AL.
Rosids, unlike in Asterids, zygomorphy has evolved in a limited number of families and does not characterize large clades (with the exception of Faboideae (Fabaceae)). Polyandry has evolved frequently throughout the group and represents a synapomorphy of the speciose subfamily Mimo soideae (Fabaceae) as well as of Rosaceae and of the genus Begonia (Begoniaceae). One striking feature is that many families display varia tion in the number of stamens among genera. Rosids are mostly char acterized by corollas with free petals whereas Asterids have corollas with fused petals. Could it be that the former allows more flexibility in floral organ number than the latter? Constraints in the evolution of morphological traits may stem from three different sources that are not necessarily independent: physical, selective and genetic. Our results suggest that inflorescence and floral architecture do not influence the evolution of floral symmetry in the same way in all clades, which invalidates a general role of physical constraints on the evolution of zygomorphy per se. We focused on the possible evolutionary antagonism between polyandry and zygomorphy in flowers. From a physical point of view, it is possible to conceive that the spatial constraints exerted by numer ous stamen primordia on the floral meristem are strong at the beginning of development, but can become relaxed as development proceeds, allowing for late-onset zygomorphy. From an adaptive point of view, polyandry and zygomorphy may be viewed as redundant for pollination efficiency. We argue that polyandry emerging in a zygomorphic context (or the reverse) may not be positively selected. Indeed, there are few examples of taxa associating both traits. The unequal distribution of this association between plant groups (near absent in Asterids but present in Rosids and Ranuncula ceae) could suggest variation in the genetic networks underlying both traits. For instance, in Asterids, the antagonism of polyandry and zygomorphy could be linked to the role of symmetry genes in inhibiting stamen develop ment (see Section VI). It would be of major interest to decipher the genetic mechanisms involved in taxa where zygomorphy and polyandry co-occur, such as in Resedaceae (Rosids) or in the Delphinieae (Ranunculales).
V. THE SIGNIFICANCE OF SYMMETRY IN
PLANT–POLLINATOR INTERACTIONS
In this section, we explore the ecological aspects of symmetry and its possible adaptive value. For ease of comparison, we consider only the flower as the study object, not lower- (bilabiate structures within flowers such as the meranthia defined by Westerkamp and Classen-Bockhoff (2007)) or higher
THE EVOLUTION OF FLORAL SYMMETRY
109
order (flower-like inflorescences such as capitula of Asteraceae) structures. We examine to what extent bilateral symmetry can be considered as one among many mating strategies increasing gene flow and thus potentially genetic diversity within species. For insects (as for other animal flower visitors), flowers are potential energetic food sources. In this context, sym metry can be perceived as an indicator of the quality and/or quantity of reward/food, even though floral mimicry may alter this potential relation ship (pollination deceit). The capacity of pollinators to perceive symmetry and discriminate between different types lays the foundation for pollinatormediated selection of flower shape, which gives an insight into the potential role of symmetry in plant population dynamics and species diversification.
A. ZYGOMORPHY AND OUTCROSSING STRATEGIES
Zygomorphy results in a polarized visual signal emitted by the flower, to which participates the orientation of the symmetry axis. This axis is generally vertically oriented, thus matching the symmetry plane of flying visitors in approach. Some studies showed that flower orientation plays a role per se in orienting the approach and landing behaviour of pollinators (Fenster et al., 2009; Ushimaru and Hyodo, 2005; Ushimaru et al., 2009), and vertical orientation has been suggested as being the first evolutionary step toward the evolution of zygomorphy (Fenster et al., 2009). Morphological differ entiation further restricts pollinator access and movement within flowers, often resulting in improved precision in pollen placement and subsequent increase in cross-fertilization. Zygomorphy thus appeared as one of numer ous contrivances for decreasing selfing and its detrimental effects on offsprings. In addition, precise pollen placement could form the basis for reproductive isolation, and thus may promote species diversification.
1. Attributes of Zygomorphic Flowers Promoting Cross-Pollination The visual signal emitted by zygomorphic flowers is generally borne by the corolla, with its brilliant colours and polarized morphology. Consistently, among 38 insect-pollinated Mediterranean species, zygomorphic ones allo cate significantly more biomass to the corolla than actinomorphic ones (Herrera, 2009). In order to ensure reproductive success, a balance must be achieved between the amount of pollen deposited on visitors and especially pollina tors, and the amount actually transferred to the stigma of another flower. This is all the more crucial when pollinators are pollen feeders. This is achieved by adaptations aiming to limit pollen wastage (e.g. poricidal
110
H. CITERNE ET AL.
Fig. 6. Interaction between a solitary bee and a flower of Agapanthus africanus (Amaryllidaceae). Due to the ventral position and curvature of the stamens, pollen deposition is sternotribic. The style is longer than the filaments, so that the pollinator comes into contact with the stigma before reaching the anthers. This arrangement favors cross-pollination. Photograph: S. Nadot.
anthers in buzz-pollinated flowers, which is encountered in some beepollinated species) and to increase precision in pollen placement on the pollinator’s body (Fig. 6). In this context, zygomorphy of the perianth is very often supplemented by various devices. For example, rewards—nectar or oil, less often resins—may be more or less concealed or not easily accessible, in nectar spur or flower throat. In Antirrhinum and Linaria, for example, the lower lip is inflated and pressed against the upper lip (“personate” flower), creating a physical obstacle in front of the nectaries. Such flowers select for strong bees able to insert their head between the two lips and open the corolla. Nectar guides are especially elaborate in zygo morphic flowers, participating in the internal symmetry, are often yellow— possibly mimicking anther colour—and attractive to bees (Endress, 1994). Bilabiate flowers of the lip and flag types (see Section II) are characterized by contrasted placement of sexual organs. In both types, the lower part of the flower serves as a landing platform for non-hovering pollinators. Stamens and stigma are protected by the upper lip in lip-type flowers, and pollen deposition on pollinators is usually nototribic (on the back). In flag flowers,
THE EVOLUTION OF FLORAL SYMMETRY
111
stamens and style are enclosed in the lower part of the bilabiate flower, and pollen deposition is sternotribic (on the ventral part of pollinators). In addition, special mechanical devices can ensure pollen application, powered by pollinators as they land on the flower (e.g. trigger system in Medicago sativa) or as they move in during visitation (e.g. the motile stamens of Salvia) to reach the reward.
2. Comparison of Zygomorphy with other Mating Strategies Promoting Outcrossing Mating strategies promoting cross-pollination include herkogamy (spatial separation of sexual organs, including various types of stylar polymorph isms), dichogamy (temporal separation of male and female maturity, i.e. protandry or protogyny), self-incompatibility systems, unisexual flowers, borne on the same (monoecy) or different (dioecy) individuals, and various combinations of both uni- and bisexual flowers (e.g. gynomonoecy, gynodioecy). These systems coexist with zygomorphy to a variable extent. Darwin (1877, cited in Barrett, 2010) considered heterostyly somewhat functionally redundant with zygomorphy as morphological adaptations promoting cross-pollination, which is consistent with the rare occurrence of both characters simultaneously. Barrett et al. (2000) found distyly in a rare species of the zygomorphic genus Salvia, possibly as a response to a new environment where protandry was not sufficient to limit intrafloral mating. In some zygomorphic species, differential spatial arrangements of reproduc tive parts have been observed, such as flexistyly (a reciprocal combination of herko- and dichogamy) in Alpinia species (Zingiberaceae), inversostyly (reciprocal vertical positioning of sexual organs) in Hemimeris species (Scro phulariaceae) or enantiostyly (Section II, and reviewed in Barrett, 2010). In most enantiostylous species, style deflection is associated with a single polli nating anther in opposite direction to the style. This particular configuration results in pollen deposited on the pollinator’s flank by one type of flower coming into contact with the stigma of its mirror-image flower (Jesson and Barrett, 2003). In addition, in some enantiostylous species, anther dimorph ism evolved, with the non-pollinating anthers specialized in pollinator feed ing. An association between zygomorphy and enantiostyly has been observed in monocots (Jesson and Barrett, 2003). Among the two forms of dichogamy, protogyny is common in wind-, beeand fly-pollinated flowers, while protandry is predominant in flowers polli nated by bees and butterflies (e.g. Endress, 2010). Consistently, an associa tion between protandry and zygomorphy has been observed in Asteridae (Kalisz et al., 2006).
112
H. CITERNE ET AL.
B. POLLINATOR PREFERENCES AND THEIR PERCEPTION OF SYMMETRY
Insects constitute the most speciose group of extant plant pollinators, even though pollination by specific groups of birds, bats and non-flying mammals also occurs (Cronk and Ojeda, 2008; Endress, 1994; Fleming et al., 2009). Symmetry as a visual cue may be recognized because of innate preferences or learning abilities. Insect were already diverse by the Permian (Whitfield and Kjer, 2008), which means that their vision began to evolve well before the emergence of angiosperms, and innate preferences or visual bias may have been recruited to improve plant–insect relationship up to flower pollination. For instance, Biesmeijer et al. (2005) established a parallel between floral guides (high frequency of a dark centre, with radial stripes or dots), insectivorous pitchers (dark centres, stripes and peripheral dots) and the appearance of the entrance of the nest of stingless bees. They proposed that plants exploit the perceptual bias of insects to attract them to specific displays such as flowers. In tests with artificial flowers, many insect species belonging to Lepidop tera, Coleoptera, Hymenoptera and Diptera have been found to prefer the largest and most symmetric flowers (Mo¨ller, 2000; Mo¨ller and Sorci, 1998; Wignall et al., 2006). Preference for larger flowers is most probably related to the low resolution of the composite insect eye (Chittka and Raine, 2006). Bees as a whole constitute the most important group of pollinators with about 20,000 species (Grimaldi and Engel, 2005), including insects with different social behaviour (solitary or social), size and various adaptations for nectar and pollen collection (e.g. Krenn et al., 2005; Thorp, 2000). Bees have high learning abilities. They are able to discriminate bilateral and radial symmetry from asymmetry. At a short distance, internal flower symmetry marked, for instance, by nectar guides may reinforce symmetry perception (Lehrer, 1999). Among bilaterally symmetrical patterns, bees prefer the patterns with vertically oriented symmetry plane, and among radially sym metric patterns, the ones with radiating bars rather than concentric circles (Giurfa et al., 1999). Preference for bilaterally symmetric shapes was demon strated to be innate in flower-naive bumblebees (Rodriguez et al., 2004). In many other studies, it is not always clear whether discrimination is based on innate preference or experience from natural conditions where particular shapes may be linked to the availability of different rewards (Lehrer, 1999).
C. FLORAL SYMMETRY AND POLLINATION SYNDROMES
The concept of pollination syndrome has been widely debated since its definition in the 19th century by Federico Delpino (Fenster et al., 2004;
THE EVOLUTION OF FLORAL SYMMETRY
113
Ollerton et al., 2009; Tripp and Manos, 2008 and references therein). It translates the observation that similar suites of flower traits can be found in evolutionarily unrelated taxa as a result of convergent selection by the same pollinating agent (Faegri and van der Pijl, 1966; Fenster et al., 2004; Proctor et al., 1996). Functional groups of pollinators have been defined to account for the observation that many species have flowers visited by large arrays of pollinator species, and conversely some pollinators visit a large array of species with different flower shapes. Analysing the Carlinville (Illinois) flora, Fenster et al. (2004) found that 61% of 86 zygomorphic species were pollinated by one functional group, significantly more than the 52% observed among 192 actinomorphic species. The traditional bee pollination syndrome includes a well-marked tridimensional form—more or less tubular flowers and most commonly zygomorphic—yellow, blue or purple colour, and nectar and pollen rewards (Faegri and van der Pijl, 1966; Proctor et al., 1996). This is not to say that all zygomorphic flowers are bee-pollinated. Indeed, it is believed the shape associated with bee pollination may have paved the way for further diversification, for example, bird pollination consistently evolved from bee pollination, and some bird-pollinated species have strongly zygomorphic flowers (e.g. Lotus maculatus—Cronk and Ojeda, 2008).
D. VARIABILITY OF FLORAL TRAITS IN ZYGOMORPHIC AND
ACTINOMORPHIC SPECIES
Because of their specific interaction with a limited number of different pollinators, it has been proposed that species with zygomorphic flowers should experience stronger pollinator-mediated stabilizing selection for flower shape and size than species with actinomorphic flowers (Berg, 1959; Gong and Huang, 2009; Wolfe and Krstolic, 1999). Consistently, various studies demonstrate lower variability in flower size in zygomorphic species than in actinomorphic ones (Herrera et al., 2008; Ushimaru and Hyodo, 2005; van Kleunen et al., 2008; Wolfe and Krstolic, 1999). While the type of symmetry is generally consubstantial with species defini tion, within-species variability around a main type exists, and has been reported to be partly genetically controlled (Mo¨ller and Shykoff, 1999). Departure from perfect symmetry is generally measured as the difference between the longest and the shortest petal in actinomorphic flowers, and between the “right” and the “left” petal in zygomorphic ones (e.g. Mo¨ller and Eriksson, 1994). More integrative approaches have been attempted, relying on geometric modelling of shape (Frey et al., 2007; Go´mez et al., 2006), which capture more spatial information.
114
H. CITERNE ET AL.
Small randomly directed deviations from perfect symmetry in natural populations are defined as fluctuating asymmetry (Endress, 1999; Mo¨ller, 2000; Rudall et al., 2002). Factors limiting such asymmetry are synorganiza tion and bilateral symmetry. Highly synorganized flowers such as those encountered in orchids (zygomorphic—Rudall and Bateman, 2002) or Apocynaceae (actinomorphic) exhibit low fluctuating asymmetry (Endress, 1999). Low fluctuating asymmetry is also observed in zygomorphic species compared to actinomorphic ones (Mo¨ller, 2000), while leaf asymmetry does not differ between the two categories of plants, suggesting that repro ductive traits are subject to differential selective pressures in the two groups, in contrast to vegetative traits. However, zygomorphic flowers tend to be larger than actinomorphic ones, and larger flowers generally exhibit less fluctuating asymmetry than smaller ones; thus, it is difficult to separate the actual effect of size and symmetry on the level of fluctuating asymmetry (Mo¨ller, 2000). The capacity to better control random variation may be an indication of “genotype quality,” and the most symmetrical flowers of some species have been shown to be the richest in nectar (Mo¨ller, 1995, 2000). In some species, a low degree of asymmetry was associated with a better seed set (Mo¨ller, 2000 for review), but in other ones this association does not hold (Botto-Mahan et al., 2004; Frey et al., 2005; Weeks and Frey, 2007). Flower visitation and reproductive success can be affected by a large number of uncontrolled causes, from environmental factors to biological ones, which may explain the lack of consistency of these results. In addition to the variability of individual traits, the level of floral inte gration measured by the correlations between the size of floral parts, is also expected to be higher in zygomorphic than in actinomorphic species because of the fit with pollinator morphology. Harder and Johnson (2009) found such a trend in their compilation of 56 studies on 43 animal-pollinated species. An integrative view of corolla shape and symmetry has been obtained by means of geometric morphometrics in the Brassicaceae species Erysimum mediohispanicum (Go´mez et al., 2006, 2008b). Shape variations are mainly found in the width of the petals and their relative distribution, generating symmetry ranging from actinomorphy to disymmetry and zygomorphy. The first study, conducted over 2 years in a single population, demonstrates that the main beetle pollinator preferentially visits disymmetric and zygo morphic corollas. In addition, the zygomorphic shape exhibits a higher fitness, measured by seedling survival (Go´mez et al., 2006). In a more extensive study involving three different populations visited by a larger diversity of pollinator assemblages, it was found that different pollinators
THE EVOLUTION OF FLORAL SYMMETRY
115
preferred different flower shapes, and that between-population variability in shape can be accounted for by preference of the major local pollinator. Pollen and nectar production also varied significantly with corolla shape (Go´mez et al., 2008a). The most rewarding flowers matched the artificial flower shape preferentially visited by bees, suggesting that bees use the visual cue as an indicator of reward amount. Significant phenotypic selection on flower shape was observed in all populations of this species (Go´mez et al., 2006, 2008b), thus giving an insight in the mechanisms of flower shape evolution mediated by reward and driven by pollinator preference. To summarize, zygomorphy results in tighter flower–pollinator interac tion than actinomorphy, and probably contributes to increased outcrossing rates. Several lines of arguments thus support the hypothesis that zygomor phy is an adaptive trait that may have brought about species divergence and species radiation in the past. However, extant populations generally exhibit low diversity in floral symmetry, making it difficult to compare the selective values of different types of symmetry.
VI. MOLECULAR BASES OF FLOWER SYMMETRY A. THE FLORAL SYMMETRY GENE REGULATORY NETWORK IN ANTIRRHINUM MAJUS
The molecular signals controlling floral symmetry were first described, and are best understood, in A. majus (Veronicaceae, Lamiales). Wild-type A. majus flowers have strongly differentiated organs along the dorsoventral axis parti cularly in the second and third whorls (petals and stamens). The two dorsal, two lateral and single ventral petals differ in size, shape, epidermal cell type and internal symmetry; in particular, the dorsal petal lobes are large and asymmetric whereas the ventral petal lobe is smaller and bilaterally symme trical. The dorsal stamen is arrested to form a staminode, whereas the lateral and ventral stamen pairs differ in filament length and pilosity. Unequal devel opment along the dorsoventral axis is apparent at the start of organogenesis, with dorsal organs delayed in their initiation (Luo et al., 1996). Two closely related genes CYCLOIDEA (CYC) and DICHOTOMA (DICH) have been identified as master control genes for bilateral symmetry by forward genetic screens (Luo et al., 1996, 1999). Cyc:dich double mutants have completely radially symmetric flowers with all organs resembling the ventral phenotype. Single cyc mutants have ventralized lateral organs and dorsal organs with lateralized features, while dich mutants display altera tions of the internal symmetry of the dorsal petals. CYC and DICH are two
116
H. CITERNE ET AL.
closely related DNA-binding transcription factors belonging to the TCP gene family (Cubas et al., 1999b; Luo et al., 1996, 1999). Both genes are expressed in the dorsal region of the floral meristem throughout its devel opment (Luo et al., 1996, 1999). CYC and DICH expression is detectable prior to organogenesis at the junction of the inflorescence and floral mer istem. After all organs are initiated, their expression is limited to the two dorsal petals and staminode, with DICH having a more restricted expression in the dorsal half of the dorsal petals (Luo et al., 1996, 1999). CYC, like other members of the TCP gene family, is believed to affect development by regulating patterns of cell growth and proliferation (reviewed in Cubas et al., 2001; Martı´n-Trillo and Cubas, 2009). In the dorsal staminode, CYC expression correlates with the downregulation of cell cycle genes such as HISTONE H4 and CYCLIN D3B (Gaudin et al., 2000). Although the initial effect of CYC expression on the floral meristem is growth retardation, at later stages of development its effect as a growth suppressor or promoter is dependent on organ identity rather than positional cues (Clark and Coen, 2002; Coen and Meyerowitz, 1991; Luo et al., 1996). CYC and DICH promote dorsal identity in A. majus flowers. By contrast, ventral identity is controlled by DIVARICATA (DIV), a gene encoding an MYB transcription factor with two imperfect repeats (R2R3) of the DNA-binding MYB domain (Almeida et al., 1997; Galego and Almeida, 2002). In loss-of-function div mutants, the ventral region of the corolla acquires lateral identity (Almeida et al., 1997). DIV is transcribed in all floral organs early in development and is inhibited post-transcriptionally in the dorsal and lateral regions through the expression of CYC and DICH (Galego and Almeida, 2002). At later stages of development when ventral petals become differentiated from lateral petals, DIV is strongly induced in the inner layer of epidermal cells of the ventral and adjacent parts of the lateral corolla lobes (Galego and Almeida, 2002). DIV promotes the expres sion of a MIXTA-like MYB gene AmMYBML1 required for the develop ment of ventral-specific petal epidermal cell types, in conjunction with B-class MADS box genes (Perez-Rodriguez et al., 2005). A gene regulatory network has been proposed for the control of floral symmetry in A. majus (Costa et al., 2005). CYC is activated upon floral induction; the molecular trigger is unknown but appears to be independent of floral meristem identity genes, as CYC is also expressed in the adaxial region of young axillary shoots adjacent to the inflorescence (Clark and Coen, 2002). Asymmetric expression in axillary meristems suggests that CYC responds to a positional cue or gradient within these meristems (Clark and Coen, 2002). The persistent expression of CYC during floral development is thought to be maintained by B- and C-function MADS
THE EVOLUTION OF FLORAL SYMMETRY
117
proteins such as DEFICIENS and PLENA (Clark and Coen, 2002) as well as self-positive feedback (Costa et al., 2005). One direct target of CYC and DICH is RADIALIS (RAD), a single-repeat MYB transcription factor that has TCP-binding sites in its promoter region and intron (Corley et al., 2005; Costa et al., 2005). RAD is required to mediate most of the effects of CYC and DICH; however, residual asymmetry is found in rad mutants suggesting some effects of CYC are independent of RAD (Corley et al., 2005; Costa et al., 2005). RAD is closely related to DIV, but has lost the C-terminal MYB II domain (Corley et al., 2005). Although direct antagonism of RAD and DIV remains to be demonstrated, this could operate by direct competition for molecular targets (Corley et al., 2005). RAD is believed to act nonautonomously on lateral organ development by inhibiting DIV (Corley et al., 2005). This may occur by cell-to-cell movement of RAD proteins, or alternatively by the activation of a downstream signalling molecule that affects lateral development (Corley et al., 2005). The gene interactions described above are summarized in Fig. 7.
Floral induction
B-and C-function MADS box genes RAD – independent pathway CYCLIN D3B
CYC RAD
DIV
DICH
DIV
DIV
DIV
AmMYBML1 B -function MADS box genes
Fig. 7. Major gene interactions regulating floral symmetry in Antirrhinum majus. Gene transcription and proposed interactions are shown in the different regions (dorsal (blue), lateral (green) and ventral (orange)) of the floral meristem. Arrows indicate upregulation, lines terminated by a perpendicular line indicate repression, and dashed lines for the repression of DIV by RAD in lateral regions represent putative RAD protein movement or indirect interaction. (See Color Insert.)
118
H. CITERNE ET AL.
B. CYC-LIKE GENES ARE IMPLICATED IN THE CONTROL OF ZYGOMORPHY IN DIVERSE LINEAGES
The extent to which these genes are implicated, and their interactions con served, in the elaboration of bilaterally symmetrical flowers has been exam ined in diverse groups of angiosperms (Fig. 8). Most studies have focused on
Asterales Apiales a,b,c–CYC2 (1-2)
Dipsacales Aquifoliales ASTERID Lamiales Solanales
d-CYC/DICH
e-VmCYC1/VmCYC2
Gontianales Garryales Fabales Rosales
f,g,h-LegCYC1/LegCYC2 (LST1) f,g-LegCYC3 (KEW1)
Malpighiales ROSID
Myrtales
i,j-CYC2B (1-2)
i-CYC2A, j-CYC2A/CYC2B-3
Brassicales Malvales
k-IaTCP1
Santalales Caryophyllales Saxifragales Gunnerales
Fig. 8. Summary of expression patterns of CYC-like genes (CYC2 clade) during late developmental stages in the corolla of representative zygomorphic core eudicot species (phylogeny derived from the Angiosperm Phylogeny website). Asterales: a. Gerbera hybrida (Broholm et al., 2008), b. Senecio squalidus (Kim et al., 2008), c. Helianthus annuus (Chapman et al., 2008); Lamiales: d. Antirrhinum majus (Luo et al., 1996, 1999), e. Veronica montana (Preston et al., 2009); Fabales: f. Lotus japonicus (Feng et al., 2006), g. Pisum sativum (Wang et al., 2008), h. Lupinus nanus (Citerne et al., 2006); Malpighiales: i. Byrsonima crassifolia, j. Janusia guaranitica (Zhang et al., 2010); Brassicales: k. Iberis amara (Busch and Zachgo, 2007). Although the predominant expression domain is dorsal (and lateral), ventral expression is found in Asterales. Expression is also detected on the abaxial side in I. amara but is weaker (in yellow) than on the dorsal side (orange). The effect on petal growth and development (acting as growth promoter or suppressor) varies across species. (See Color Insert.)
THE EVOLUTION OF FLORAL SYMMETRY
119
homologues of CYC/DICH. The Lamiales have evolved zygomorphic flow ers from an ancestor with actinomorphic flowers (Coen and Nugent, 1994; Donoghue et al., 1998; Endress, 2001b), and it is therefore unsurprising that CYC-like genes are implicated in the control of bilateral symmetry in other members of this clade. In particular, the persistent expression on the dorsal side of the developing flower of CYC homologues has been described in other zygomorphic species of Veronicaceae (Cubas et al., 1999a; Hileman et al., 2003; Preston et al., 2009) and Gesneriaceae (Du and Wang, 2008; Gao et al., 2008; Song et al., 2009; Zhou et al., 2008). Notably, variations in the pattern of stamen development and the degree of petal differentiation along the dorsoventral axis have frequently been associated with modifications of CYC-like gene expression. For example, in Mohavea confertiflora (Veroni caceae), the abortion of both dorsal and lateral stamens coincides with an expansion of the expression domain of CYC and DICH homologues from the dorsal region to the lateral stamen primordia (Hileman et al., 2003). Similarly, in Chirita heterotricha (Gesneriaceae), an expanded expression domain (i.e. in both dorsal and lateral regions of the flower) of one CYC homologue coincides with the abortion of dorsal and lateral stamens (Gao et al., 2008). In the Lamiales, however, stamen abortion per se is not necessarily associated with CYC expression, particularly on the ventral side (Preston et al., 2009; but see Song et al., 2009). CYC-like genes have been recruited for the control of floral symmetry in families that have evolved zygomorphy independently of the Lamiales. Within Rosids, these have been implicated in the control of dorsal (and sometimes lateral) petal identity in Fabaceae, Brassicaceae and Malpighia ceae (Busch and Zachgo, 2007; Feng et al., 2006; Wang et al., 2008; Zhang et al., 2010). In Papilionoideae (Fabaceae), two closely related CYC-like genes are expressed in the dorsal region of developing flowers (Citerne et al., 2006; Feng et al., 2006; Wang et al., 2008); one of these, LOBED STANDARD 1 (LST1), is an important determinant of dorsal petal identity, promoting cellular proliferation and epidermal cell differentiation (Feng et al., 2006; Wang et al., 2008). The other copy appears to have less effect on phenotype, but may act redundantly to control dorsal petal development (Wang et al., 2008). A third CYC homologue expressed in the dorsal and lateral regions of the developing flower, KEELED WINGS 1 (KEW1), is also a regulator of dorsoventral asymmetry, and determines lateral petal identity (Feng et al., 2006; Wang et al., 2008). The petals of lst1:kew1 double mutants have ventral identity in both L. japonicus and Pisum sativum (Feng et al., 2006; Wang et al., 2008). Similar expression is found in duplicate CYC-like genes in zygomorphic Malpighiaceae (Zhang et al., 2010). As in Fabaceae, paralogues exhibit
120
H. CITERNE ET AL.
either dorsal or dorsolateral expression late in floral development, a pattern that is not found in their closest relatives with actinomorphic flowers. Gene duplication, and consequently functional divergence, has occurred indepen dently in Fabaceae and Malpighiaceae (Citerne et al., 2003; Zhang et al., 2010), and the extent of functional redundancy and specificity remains to be demonstrated in Malpighiaceae. Within Brassicaceae, I. amara has been a case study for the control of late-onset zygomorphy (Busch and Zachgo, 2007). I. amara flowers are tetramerous with two reduced dorsal petals and two enlarged ventral petals. A shift occurs during flower development: petals are initiated simultaneously and grow equally until relatively late in development when, at the onset of stamen differentiation, unequal adaxial–abaxial petal growth becomes apparent. A shift is also observed in the expression of the homologue of CYC in I. amara IaTCP1, which is expressed equally early in development but becomes strongly expressed in the two dorsal petals relative to the ventral petals at later developmental stages (Busch and Zachgo, 2007). The effect of IaTCP1 decreases petal growth and is the opposite of what is observed in Antirrhinum and Fabaceae (i.e. promoter of petal growth during late developmental stages), indicative of functional divergence. Constitutive expression of IaTCP1 in Arabidopsis produces a similar phenotype to when the endogenous gene TCP1 is constitutively expressed, that is, repressed cell division reducing vegetative and petal growth, suggesting that DNA targets and interacting proteins are conserved in Brassicaceae (Busch and Zachgo, 2007). By contrast, the effect on petal growth of heterologous expression of Antirrhinum CYC in Arabidopsis is enlargement by cell expansion suggesting that targets and interacting proteins are not conserved between Antirrhinum and Brassicaceae (Busch and Zachgo, 2007; Costa et al., 2005). In Asteraceae (Asterid clade like Lamiales), CYC-like genes also regulate dorsoventral asymmetry but in a novel manner, as a ventralizing factor (Broholm et al., 2008; Kim et al., 2008). In radiate inflorescences, both actinomorphic (disc) and zygomorphic (ray) flowers are present: the outer most flowers develop enlarged fused petal lobes on the ventral side (the ligule), and have aborted stamens. Expression of a subset of CYC-like genes was found predominantly in ray flowers (Broholm et al., 2008; Chap man et al., 2008; Kim et al., 2008), in particular on the ventral side promoting ligule development (Broholm et al., 2008). In Gerbera hybrida, the effects of constitutive expression of GhCYC2 differ not only with organ type (increas ing growth of petals and reducing growth of stamens) but also according to flower type and position along the capitulum radius (Broholm et al., 2008). There is less evidence for the involvement of CYC-like genes in the control of zygomorphy outside the core eudicots. In rice, RETARDED PALEA1
THE EVOLUTION OF FLORAL SYMMETRY
121
(REP1) promotes the differentiation of the palea and the lemma (which together function as a calyx surrounding the stamens and carpel) by regulat ing cellular expansion and differentiation (Yuan et al., 2009). In Fumarioi deae (Papaveraceae), bilaterally symmetric flowers are characterized by the development of a nectar spur in one of the two outer petals. The asymmetric expression of one CYC-like gene in the spurred petal of C. sempervirens could indicate a role in floral zygomorphy but remains to be demonstrated functionally (Damerval et al., 2007). C. GENETIC MECHANISMS UNDERLYING CHANGES IN FLORAL SYMMETRY
Modification of key development regulators appears to underlie morpholo gical evolution (e.g. Doebley and Lukens, 1998; Wilson et al., 1977; Rosin and Kramer, 2009). Changes in the timing, duration and localization of CYC-like gene expression have repeatedly been implicated in changes in floral symmetry. Case studies have provided examples of different muta tional mechanisms. In L. vulgaris, naturally occurring radially symmetrical mutants have lost CYC expression through extensive methylation of pro moter and ORF (Cubas et al., 1999a). Surveys of epigenetic alteration of gene expression in plants suggest this mode of regulation may play a role in morphological evolution (Kalisz and Purrugannan, 2004; Rapp and Wendel, 2005); however, no other example has been described so far in the context of floral symmetry. In Senecio, interspecific hybridization has been shown to have played a part in the evolution of a floral symmetry polymorphism (Kim et al., 2008). In Senecio vulgaris, a species with typically non-radiate inflores cences bearing only disc florets, a radiate form has evolved by introgres sion of an allele at the RAY locus from Senecio squalidus with radiate inflorescences. The RAY locus consists of two CYC2 paralogues, and as in Gerbera, one of these genes appears to promote ventral identity in ray florets. These genes are specifically expressed in the outer florets, and are differentially expressed in the two forms. It is believed that changes in cisregulatory regions, rather than the ORF, may underlie the differences between the two morphs. There are numerous cases of species derived from zygomorphic lineages that have evolved actinomorphic flowers secondarily. Diverse types of changes in CYC-like gene expression have been described. In Plantago lanceolata (Veronicaceae), a wind-pollinated genus with radial tetramerous flowers, expression of the CYC-homologue PlCYC is detected in flowers only at later stages of development in all four stamens (in the anther con nective and stamen filament) and transiently in the ovaries (Reardon et al.,
122
H. CITERNE ET AL.
2009). The actinomorphy in P. lanceolata is therefore correlated with a lack of both early expression and asymmetric expression in petals. The function of PlCYC is unknown, but has been proposed to delay stamen development and therefore promote dichogamy. Unlike other members of Veronicaceae (Preston et al., 2009), P. lanceolata has only one CYC-like gene, which could suggest a functionally significant gene-loss event (Reardon et al., 2009). In Bournea leiophylla (Gesneriaceae), the transition from a zygomorphic pattern in the early stages of floral development to actinomorphy at anthesis correlates with the downregulation of the dorsal expression of a CYC-like and a RAD-like gene (Zhou et al., 2008). By contrast, in Cadia purpurea (Fabaceae), the derived radial symmetry of the corolla coincides with an expansion of the expression domain of one CYC-like gene to all petals (Citerne et al., 2006). It remains to be determined whether these heterochro nic and heterotopic changes in gene expression are caused by modifications in their cis-regulatory regions or in the function or nature of their trans acting regulators. D. EVOLUTION OF CYC-LIKE GENES: FUNCTIONAL IMPLICATIONS
It is believed that morphological evolution proceeds by tinkering of existing genetic pathways (Jacob, 1977). What is the context of CYC-like gene evolution that makes them a common player in the repeated evolution of floral zygomorphy in many lineages? Members of the TCP gene family are transcription factors that bind to DNA through their characteristic basic helix–loop–helix domain (bHLH) (Martı´n-Trillo and Cubas, 2009). CYC together with its homologue in maize TEOSINTE BRANCHED 1 (TB1) belong to a clade of class II TCP genes (the ECE clade), whose members are generally characterized by a second short conserved hydrophilic domain (R domain) and a conserved motif of amino acids termed “ECE”. Character ized genes in this clade appear to have a predominant role in growth repres sion (Martı´n-Trillo and Cubas, 2009). TB1 is a suppressor of axillary meristem growth (Doebley et al., 1997), but also affects floral development by suppressing stamen growth in female flowers (Hubbard et al., 2002). Two major duplication events have occurred in the ECE clade, prior to the divergence of the core eudicots (Howarth and Donoghue, 2006). All genes implicated so far in dorsoventral asymmetry of flowers belong to the same CYC2 clade (Howarth and Donoghue, 2006), whereas genes from the CYC1 and CYC3 clade in Arabidopsis appear to have a role like TB1 in the development of axillary buds (Aguilar-Martı´nez et al., 2007; Finlayson, 2007). This could reflect sub/neofunctionalization of major ECE-CYC lineages in the core eudicots, where the effects on floral development such
THE EVOLUTION OF FLORAL SYMMETRY
123
as stamen suppression of the CYC/TB1 ancestor were retained and subse quently modified in the CYC2 clade. The dorsal expression of many CYC2 genes is believed to be shared by the common ancestor of Rosids and Asterids (Cubas et al., 2001). In Arabidopsis thaliana (Brassicaceae), which has radially symmetrical flowers, the homo logue of CYC TCP1, is transiently expressed in the dorsal region of the floral meristem prior to organogenesis (Cubas et al., 2001). Modification of this incipient asymmetry through its persistent expression during organ primor dia development could account for the repeated evolution of zygomorphy (Cubas et al., 2001). However, evidence of ventral and radial expression of CYC2 genes in different lineages suggests lability in the response to local signals along the dorsoventral axis in the floral meristem. For example, early expression of IaTCP1 in I. amara is very weak and ubiquitous (Busch and Zachgo, 2007), and differs from that of Arabidopsis TCP1, which is transi ently expressed on the dorsal side of the floral meristem. Without expression data from other Brassicaceae, it is not clear whether the early asymmetric pattern is ancestral or derived. Similarly in Malpighiales, the actinomorphic relatives of the zygomorphic members of family Malpighiaceae (which have “typical” CYC dorsal expression) differ in their expression of CYC-like genes; in the closest relative these are expressed uniformly in late-stage flowers, whereas in the next closest relative no CYC expression is detected at this stage (Zhang et al., 2010). The role of CYC2 genes in petal develop ment also appears to be labile, probably reflecting differences in their inter action with other proteins. In different lineages, these genes can either promote or repress growth through cell proliferation and/or expansion, and are often associated with cellular differentiation. Independent duplication of CYC2 genes appears to be a common phe nomenon in core eudicots, for example, in Veronicaceae (Preston et al., 2009), Gesneriaceae (Citerne et al., 2000; Smith et al., 2004, 2006), Asteraceae (Broholm et al., 2008; Chapman et al., 2008), Dipsacales (Howarth and Donoghue, 2005), Fabaceae (Citerne et al., 2003; Fukuda et al., 2003) and Malpighiales (Zhang et al., 2010). Correlation between floral form and copy number has been postulated in Dipsacales but the significance of duplications specific to zygomorphic lineages remains to be demonstrated (Howarth and Donoghue, 2005). There appears to be flexibility in the fate of duplicate CYC-like genes from the CYC2 clade, providing scope for morphological evolution and the elaboration of complex flowers. The duplication, and consequent subfunc tionalization, of CYC and DICH is specific to the Antirrhineae (Gu¨bitz et al., 2003; Hileman and Baum, 2003). However, in L. vulgaris, although both CYC and DICH orthologues have been identified (Gu¨bitz et al., 2003;
124
H. CITERNE ET AL.
Hileman and Baum, 2003), loss of CYC activity appears to be sufficient to generate a fully radial flower (Cubas et al., 1999a) and the function of Linaria DICH is unknown. The extent of redundancy and/or functional divergence between paralogues varies among lineages. Nevertheless, there has been little evidence of shifting patterns of selection acting on CYC-like genes, either between actinomorphic and zygomorphic lineages or between duplicate copies (Hileman and Baum, 2003 (Veronicaceae); Smith et al., 2006 (Gesneriaceae); Reardon et al., 2009 (Veronicaceae); but see Ree et al., 2004 (Fabaceae)), suggesting that gene biochemical function, beyond the establishment of zygomorphy, may be conserved at least between closely related taxa. In Helianthus annuus (Asteraceae), however, positive selection was detected at multiple sites in the TCP and R domains in CYC2 paralogues with divergent expression patterns (Chapman et al., 2008). E. BEYOND CYC: CONSERVATION AND DIVERGENCE OF OTHER
COMPONENTS OF THE FLORAL SYMMETRY NETWORK
MYB genes form one of the largest families of transcription factors in plants and many play a key role in plant development (Du et al., 2009). Recent reports show that R2R3-MYB genes can also have a role in cell cycle regulation (reviewed in Cominelli and Tonelli, 2009). In addition, functional relationships have been described between MYB and bHLH proteins (e.g. in the production of specialized epidermal cells (Du et al., 2009; Ramsay and Glover, 2005)). Therefore, MYB genes may be involved in different floral symmetry gene networks. The involvement of RAD-like genes in the floral symmetry pathway appears to be conserved in the Lamiales. Expression of the homologue of Antirrhinum RAD has been described in other species of Veronicaceae (Preston and Hileman, 2009) as well as Gesneriaceae (Zhou et al., 2008), where all show strong expression in the dorsal region of the developing flower coinciding with CYC-like gene expression. However, in Arabidopsis, RAD-like genes (AtRLs) do not appear to be activated either by the endo genous CYC homologue TCP1 (Baxter et al., 2007) or by constitutively expressed Antirrhinum CYC (Costa et al., 2005). However, constitutive expression of Antirrhinum RAD does have developmental effects in Arabi dopsis, repressing vegetative growth and development (Baxter et al., 2007). Therefore, although the ancestor of RAD probably had developmental functions, both cis- and trans-acting regulators have diverged since the separation of Antirrhinum and Arabidopsis lineages, and the co-option of RAD in the regulation of floral symmetry may be specific to the Lamiales (or Asterids) (Baxter et al., 2007; Costa et al., 2005).
THE EVOLUTION OF FLORAL SYMMETRY
125
Little is known about the function of DIV-like genes outside of Antirrhi num. In Bournea (Gesneriaceae), a genus with flowers showing bilateral symmetry only during the early stages of development, expression of two DIV homologues was detected in floral organ primordia irrespective of position along the dorsoventral axis (Zhou et al., 2008). Outside the Lamiales, expression of five DIV-like genes in Heptacodium (Caprifoliaceae, Dipsacales) suggests these are for the most part widely transcribed in floral organs (Howarth and Donoghue, 2009). However, the interpretation of expression patterns of DIV-like genes is complicated by the fact that they may be, as DIV in Antirrhinum, regulated posttranscriptionally (Galego and Almeida, 2002). Downstream targets and interacting proteins have not yet been identified in other lineages. In Pisum (Fabaceae), the SYMMETRIC PETALS 1 locus affects the internal asymmetry of petals and appears to be antagonized by KEW and LST1 (Wang et al., 2008). Micromorphological differences between dorsal, lateral and ventral petal epidermis in Papilionoideae (Fabaceae) may also implicate MIXTA-like genes, as in Antirrhinum (Ojeda et al., 2009). A different regulatory pathway involving MADS-box transcription fac tors has been invoked for the elaboration of zygomorphic flowers in Orch idaceae (Mondrago´n-Palomino and Theissen, 2008, 2009; Tsai et al., 2004). In Phalaenopsis equestris, DEF-like paralogues were found to be differen tially expressed in floral organs; in particular PeMADS4 is specifically expressed in the ventral lip (Tsai et al., 2004). In radially symmetric forms of P. equestris with three lip-like internal tepals, ectopic expression of PeMADS4 was detected in each internal tepal suggesting this change in gene transcription may be associated with the loss of zygomorphy (Tsai et al., 2004). A model has been proposed where morphological differentia tion within the perianth of orchids (i.e. inner versus outer tepals and lateral versus ventral inner tepals) is associated with different combinations of four functionally divergent duplicate DEF-like genes (Mondrago´n-Palomino and Theissen, 2008, 2009). According to this model, outer tepal identity is established by one duplicate gene pair (clades 1þ2), whereas inner tepal identity is established by the combination of clade 1þ2þ3 genes and inner ventral identity by the combination of clade 1þ2þ3þ4 genes, with variations in floral morphology attributed to changes in expression of clade 3 and 4 genes (Mondrago´n-Palomino and Theissen, 2008, 2009). The applicability of this framework remains to be demonstrated in other Orchidaceae. Never theless, it suggests that novel pathways directly involving genes other than CYC may control floral symmetry in certain lineages, although the involve ment of TCP genes providing positional cues is not ruled out.
126
H. CITERNE ET AL.
VII. PERSPECTIVES
Floral symmetry is an ideal system for investigating enduring questions in evolutionary biology such as (1) what is the genetic basis of convergent evolution and (2) to what extent can natural morphological novelties result from gradual or saltational evolution? It has been proposed that repeated evolution of traits controlled by few regulatory loci of major effect is likely to have a common genetic basis (Gompel and Prud’homme, 2009; Wood et al., 2005). In all cases examined so far, the genetic control of floral symmetry involves major regulatory genes where changes in activity can dramatically alter phenotype. In the core eudicots, the repeated involvement of CYC2 genes in the elaboration of zygomorphic flowers suggests that a preexisting genetic pathway is preferentially modified in this group. Never theless, many questions remain regarding the establishment and ancestral function of this pathway. CYC2 genes are present and appear functional in actinomorphic core eudicot species; however, their function is unknown. A wider survey of CYC2 expression and function needs to be carried out, particularly in actinomorphic lineages, to establish whether early dorsal expression is indeed the ancestral state. In addition, zygomorphy in the core eudicots appears to be associated with the duplication and functional divergence of the CYC-like gene lineage. This begs the question whether zygomorphy in lineages outside of the core eudicots could also be controlled by TCP genes, and if not, what alternative pathways could underlie this convergence. The example of Orchidaceae suggests alternative mechanisms involving subfunctionalization of organ identity genes, but this scenario is linked with gene duplications that are specific to this family, and cannot be generalized to other zygomorphic monocot lineages such as Poaceae or Zingiberales. Reconstructing character evolution in a phylogenetic frame work has shown that the flower and inflorescence contexts for the evolution of symmetry may differ from one clade to another, suggesting different constraints that may reveal different genetic networks underlying symmetry. It is unclear whether changes in floral symmetry over evolutionary time have taken place through gradual or saltational events, for example, through the appearance of hopeful monsters (Goldschmidt, 1940) possibly involving homeotic mutants. A new symmetry phenotype emerging in a population may be maintained because it is able to reproduce vegetatively or self-fertilize, and because of diverse evolutionary forces, possibly including pollinatormediated selection. In the case where self-fertilization is possible, it may nevertheless lead to inbreeding depression and low selective value of the novel phenotype. As a rare mutant, it may have more chance to survive in small populations escaping drift than in large populations, where it will be
THE EVOLUTION OF FLORAL SYMMETRY
127
easily counter-selected. Very few studies have been able to precisely quantify the variability of symmetry in natural populations. This probably comes from the difficulty to formally assess deviations from a dominant type in a manner that encompasses the complexity of flower morphology, but also from an apparent lack of within- species variability. Different fitness associated with different types, as shown in Erysimum mediohispanicum (Go´mez et al., 2006, 2008b), could constitute a basis for further evolution through gradual changes. On another hand, the absence (or extreme rarity) of coexistence of two markedly different types of symmetry in a same species may be the result of rapid loss of new types, or reproductive isolation promoted by plant– pollinator interaction leading to successful speciation. In the Fumarioideae (Papaveraceae), where zygomorphy is due to the loss of one of two nectar spurs, it is possible to find degrees in spur reduction in a same inflorescence in Capnoides, but also between species of Corydalis (Lide´n, 1986). This could represent an example of gradual changes associated with species divergence. It has been suggested that homeotic mutants could have played a rare but important role in establishing new plant lineages (Theissen, 2006), although this remains to be demonstrated. As far as floral symmetry is concerned, most homeotic mutants that have been described have radially symmetric flowers derived from zygomorphic types. In the case of L. vulgaris, the actinomorphic epimutant may only reproduce vegetatively so that its fitness may not be linked to its floral phenotype, and its evolutionary significance is thus difficult to assess (Theissen, 2000, 2006). Species that have evolved radially symmetric flowers secondarily frequently show changes in CYC-like genes (such as loss-of-function or heterotopic expression), which could be sufficient to account for their phenotype. By contrast, no zygomorphic mutants are known in actinomorphic species, suggesting that complex genetic pathways are established over time, consistent with the hypothesis of evolution of zygomorphy through gradual genetic changes. The hypothesis of a major ancestral event possibly relaxing genetic con straints and opening the way for gradual transformations reinforcing the initial change cannot, however, be easily dismissed. Phylogenetic analyses pointing to clades exhibiting different degrees in the “severity” of zygomor phy derived from an actinomorphic ancestor could help evaluate this hypothesis. In particular, phenotypes such as “nearly actinomorphic” or “nearly zygomorphic” could constitute intermediate evolutionary steps toward the evolution of structural zygomorphy. Analysis of symmetry gene networks will benefit in the near future from large-scale genomic studies made easier by high-throughput sequencing. Such extensive comparative analyses in well-chosen species may help unravel the evolutionary steps in the making of floral symmetry.
128
H. CITERNE ET AL.
ACKNOWLEDGEMENTS
We thank our colleagues for fruitful discussions, and an anonymous reviewer for constructive comments. HC was supported by a fellowship from the Agence Nationale de la Recherche program ANR-07-BLAN 0112-02, and FJ by a fellowship from the Ministe`re de l’Enseignement Supe´rieur et de la Recherche, France.
REFERENCES Aguilar-Martı´nez, J.A., Poza-Carrio´n, C., Cubas, P., 2007. Arabidopsis BRANCHED1 acts as an integrator of branching signals within axillary buds. Plant Cell 19, 458–472. Almeida, J., Galego, L., 2005. Flower symmetry and shape in Antirrhinum. Int. J. Dev. Biol. 49, 527–537. Almeida, J., Rocheta, M., Galego, L., 1997. Genetic control of flower shape in Antirrhinum majus. Development 124, 1387–1392. Barrett, S.C.H., 2010. Darwin’s legacy: the forms, function and sexual diversity of flowers. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 365, 351–368. Barrett, S.C.H., Wilken, D.H., Cole, W.W., 2000. Heterostyly in the Lamiaceae: the case of Salvia brandegeei. Plant Syst. Evol. 223, 211–219. Basinger, J.F., Dilcher, D.L., 1984. Ancient bisexual flowers. Science 224, 511–513. Baxter, C.E., Costa, M.M., Coen, E.S., 2007. Diversification and co-option of RADlike genes in the evolution of floral asymmetry. Plant J. 52, 105–113. Berg, R.L., 1959. A general evolutionary principle underlying the origin of develop mental homeostasis. Am. Nat. 93, 103–105. Bernhardt, P., 2000. Convergent evolution and adaptive radiation of beetlepollinated angiosperms. Plant Syst. Evol. 222, 293–320. Biesmeijer, J.C., Giurfa, M., Koedam, D., Potts, S.G., Joel, D.M., et al., 2005. Convergent evolution: floral guides, stingless bee nest entrances, and insec tivorous pitchers. Naturwissenschaften 92, 444–450. Botto-Mahan, C., Pohl, N., Medel, R., 2004. Nectar guide fluctuating asymmetry does not relate to female fitness in Mimulus luteus. Plant Ecol. 174, 347–352. Braun, A., 1835. Dr. Carl Schimper’s Vortra¨ge u¨ber die Mo¨glichkeit eines wissenschaftlichen Versta¨ndnisses der Blattstellung, nebst Andeutung der hauptsa¨chlichen Blattstellungsgesetze und insbesondere der neuentdeckten Gesetze der Aneinanderreihung von Cyclen verschiedener Maasse. Flora 18, 145–192. Broholm, S.K., Ta¨htiharju, S., Laitinen, R.A., Albert, V.A., Teeri, T.H., Elomaa, P., 2008. A TCP domain transcription factor controls flower type specification along the radial axis of the Gerbera (Asteraceae) inflorescence. Proc. Natl. Acad. Sci. U.S.A. 105, 9117–9122. Busch, A., Zachgo, S., 2007. Control of corolla monosymmetry in the Brassicaceae Iberis amara. Proc. Natl. Acad. Sci. U.S.A. 104, 16714–16719. Busch, A., Zachgo, S., 2009. Flower symmetry evolution: towards understanding the abominable mystery of angiosperm radiation. BioEssays 31, 1181–1190. Carpenter, R., Coen, E., 1990. Floral homeotic mutations produced by transposon mutagenesis in Antirrhinum majus. Genes Dev. 4, 1483–1493.
THE EVOLUTION OF FLORAL SYMMETRY
129
Chapman, M.A., Leebens-Mack, J.H., Burke, J.M., 2008. Positive selection and expression divergence following gene duplication in the sunflower CYCLOIDEA gene family. Mol. Biol. Evol. 25, 1260–1273. Chittka, L., Raine, N.E., 2006. Recognition of flowers by pollinators. Curr. Opin. Plant Biol. 9, 428–435. Citerne, H.L., Luo, D., Pennington, R.T., Coen, E., Cronk, Q.C., 2003. A phyloge nomic investigation of CYCLOIDEA-like TCP genes in the Leguminosae. Plant Physiol. 131, 1042–1053. Citerne, H.L., Mo¨ller, M., Cronk, Q.C.B., 2000. Diversity of cycloidea-like genes in Gesneriaceae in relation to floral symmetry. Ann. Bot. 86, 167–176. Citerne, H.L., Pennington, R.T., Cronk, Q.C., 2006. An apparent reversal in floral symmetry in the legume Cadia is a homeotic transformation. Proc. Natl. Acad. Sci. U.S.A. 103, 12017–12020. Clark, J.I., Coen, E.S., 2002. The cycloidea gene can respond to a common dorso ventral prepattern in Antirrhinum. Plant J. 30, 639–648. Coen, E., 1991. The role of homeotic genes in flower development and evolution. Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 241–279. Coen, E., Nugent, J.M., Luo, D.A., Bradley, D., Cubas, P., Chadwick, M., et al., 1995. Evolution of floral symmetry. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 350, 35–38. Coen, E.S., Meyerowitz, E.M., 1991. War of the whorls: genetic interactions control ling floral development. Nature 353, 31–37. Coen, E.S., Nugent, J.M., 1994. Evolution of flowers and inflorescences. Develop ment 120, 107–116. Cominelli, E., Tonelli, C., 2009. A new role for plant R2R3-MYB transcription factors in cell cycle regulation. Cell Res. 19, 1231–1232. Corley, S.B., Carpenter, R., Copsey, L., Coen, E., 2005. Floral asymmetry involves an interplay between TCP and MYB transcription factors in Antirrhinum. Proc. Natl. Acad. Sci. U.S.A. 102, 5068–5073. Costa, M.M., Fox, S., Hanna, A.I., Baxter, C., Coen, E., 2005. Evolution of regulatory interactions controlling floral asymmetry. Development 132, 5093–5101. Crepet, W.L., 1996. Timing in the evolution of derived floral characters: Upper Cretaceous (Turonian) taxa with tricolpate and tricolpate-derived pollen. Rev. Palaeobot. Palynol. 90, 339–359. Crepet, W.L., 2008. The fossil record of angiosperms: requiem or renaissance? Ann. Mo. Bot. Gard. 95, 3–33. Crepet, W.L., Niklas, K.J., 2009. Darwin’s second “abominable mystery”: why are there so many angiosperm species? Am. J. Bot. 96, 366–381. Cronk, Q., Ojeda, I., 2008. Bird-pollinated flowers in an evolutionary and molecular context. J. Exp. Bot. 59, 715–727. Cubas, P., 2004. Floral zygomorphy, the recurring evolution of a successful trait. Bioessays 26, 1175–1184. Cubas, P., Coen, E., Zapater, J.M., 2001. Ancient asymmetries in the evolution of flowers. Curr. Biol. 11, 1050–1052. Cubas, P., Lauter, N., Doebley, J., Coen, E., 1999b. The TCP domain: a motif found in proteins regulating plant growth and development. Plant J. 18, 215–222. Cubas, P., Vincent, C., Coen, E., 1999a. An epigenetic mutation responsible for natural variation in floral symmetry. Nature 401, 157–161. Dahlgren, R.M.T., Clifford, H.T., Yeo, P.F., 1985. The Families of the Monocoty ledons (Structure, Evolution, and Taxonomy), Springer, Berlin. Damerval, C., Le Guilloux, M., Jager, M., Charon, C., 2007. Diversity and evolution of CYCLOIDEA-like TCP genes in relation to flower development in Papaveraceae. Plant Physiol. 143, 759–772.
130
H. CITERNE ET AL.
Damerval, C., Nadot, S., 2007. Evolution of perianth and stamen characteristics with respect to floral symmetry in Ranunculales. Annals of Botany 100, 631–640. Darwin, C., 1868. The Variation of Animals and Plants Under Domestication, J. Murray, London. Darwin, C., 1877. The different forms of flowers on plants of the same species, John Murray, London. Dilcher, D., 2000. Toward a new synthesis: major evolutionary trends in the angios perm fossil record. Proc. Natl. Acad. Sci. U.S.A. 97, 7030–7036. Doebley, J., Lukens, L., 1998. Transcriptional regulators and the evolution of plant form. Plant Cell 10, 1075–1082. Doebley, J., Stec, A., Hubbard, L., 1997. The evolution of apical dominance in maize. Nature 386, 485–488. Dong, Z.C., Zhao, Z., Liu, C.W., Luo, J.H., Yang, J., Huang, W.H., et al., 2005. Floral patterning in Lotus japonicus. Plant Physiol. 137, 1272–1282. Donoghue, M.J., Ree, R.H., Baum, D.A., 1998. Phylogeny and the evolution of flower symmetry in the Asteridae. Trends Plant Sci. 3, 311–317. Douglas, A.W., 1997. The developmental basis of morphological diversification and synorganization in flowers of Conospermeae (Stirlingia and Conosper minae, Proteaceae). Int. J. Plant Sci. 158, S13–S48. Douglas, A.W., Tucker, S.C., 1996. Comparative floral ontogenies among Persoo nioideae including Bellendena (Proeaceae). Am. J. Bot. 83, 1528–1555. Doyle, J.A., Endress, P.K., 2000. Morphological phylogenetic analysis of basal angiosperms: comparison and combination with morphological data. Int. J. Plant Sci. 161, S121–S153. Du, H., Zhang, L., Liu, L., Tang, X.F., Yang, W.J., Wu, Y.M., et al., 2009. Biochemical and molecular characterization of plant MYB transcription factor family. Biochemistry (Moscow) 74, 1–11. Du, Z.Y., Wang, Z.Y., 2008. Significance of RT-PCR expression patterns of CYC-like genes in Oreocharis benthamii (Gesneriaceae). J. Syst. Evol. 46, 23–31. Eberwein, R., Nickrent, D.L., Weber, A., 2009. Development and morphology of flowers and inflorescences in Balanophora papuana and B. elongata (Bala nophoraceae). Am. J. Bot. 96, 1055–1067. Eichler, A.W., 1878. Blu¨thendiagramme Constructert und Erla¨utert, Engelmann, Leipzig. Endress, P.K., 1994. Diversity and Evolutionary Biology of Tropical Flowers, Cambridge University Press, Cambridge, UK. Endress, P.K., 1997. Antirrhinum and the Asteridae—evolutionary changes of floral symmetry. Symp. Soc. Exp. Biol. 51, 133–140. Endress, P.K., 1998. Antirrhinum and Asteridae—evolutionary changes of floral symmetry. Symp. Soc. Exp. Biol. 51, 133–140. Endress, P.K., 1999. Symmetry in flowers: diversity and evolution. Int. J. Plant Sci. 160, S3–S23. Endress, P.K., 2001a. Evolution of floral symmetry. Curr. Opin. Plant Biol. 4, 86–91. Endress, P.K., 2001b. The flower in extant basal angiosperms and inferences on ancestral flowers. Int. J. Plant Sci. 162, 1111–1140. Endress, P.K., 2006. Angiosperm floral evolution, morphological developmental framework. Adv. Bot. Res. 44, 1–61. Endress, P.K., 2010. The evolution of floral biology in basal angiosperms. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 365, 411–421. Endress, P.K., Doyle, J.A., 2009. Reconstructing the ancestral angiosperm flower and its initial specializations. Am. J. Bot. 96, 22–66. Faegri, K., van der Pijl, L., 1966. The Principle of Pollination Ecology, Pergamon Press, Oxford.
THE EVOLUTION OF FLORAL SYMMETRY
131
Feild, T.S., Arens, N.C., 2007. The ecophysiology of early angiosperms. Plant Cell Environ. 30, 291–309. Feng, X., Zhao, Z., Tian, Z., Xu, S., Luo, Y., Cai, Z., et al., 2006. Control of petal shape and floral zygomorphy in Lotus japonicus. Proc. Natl. Acad. Sci. U.S.A. 103, 4970–4975. Fenster, C.B., Armbruster, W.S., Dudash, M.R., 2009. Specialization of flowers: is floral orientation an overlooked first step? New Phytol. 183, 502–506. Fenster, C.B., Armbruster, W.S., Wilson, P., Dudash, M.R., Thomson, J.D., 2004. Pollination syndromes and floral specialization. Annu. Rev. Ecol. Evol. Syst. 35, 375–403. Finlayson, S.A., 2007. Arabidopsis Teosinte Branched 1-like 1 regulates axillary bud outgrowth and is homologous to monocot Teosinte Branched1. Plant Cell Physiol. 48, 667–677. Fleming, T.H., Geiselman, C., Kress, W.J., 2009. The evolution of bat pollination: a phylogenetic perspective. Ann. Bot. 104, 1017–1043. Frey, F.M., Davis, R., Delph, L.F., 2005. Manipulation of floral symmetry does not affect seed production in Impatiens pallida. Int. J. Plant Sci. 166, 659–662. Frey, F.M., Robertson, A., Bukoski, M., 2007. A method for quantifying rotational symmetry. New Phytol. 175, 785–791. Friedman, W.E., 2009. The meaning of Darwin’s “abominable mystery”. Am. J. Bot. 96, 5–21. Friis, E.M., Doyle, J.A., Endress, P.K., Leng, Q., 2003. Archaefructus—angiosperm precursor or specialized early angiosperm? Trends Plant Sci. 8, 369–373. Friis, E.M., Pedersen, K.R., Crane, P.R., 2001. Fossil evidence of water lilies (Nympheales) in the Early Cretaceous. Nature 410, 357–360. Friis, E.M., Pedersen, K.R., Crane, P.R., 2006. Cretaceous angiosperm flowers: inno vation and evolution in plant reproduction. Palaeogeogr. Palaeoclimatol. Palaeoecol. 232, 251–293. Friis, E.M., Pedersen, K.R., Crane, P.R., 2010. Diversity in obscurity: fossil flowers and the early history of angiosperms. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 365, 369–382. Frohlich, M.W., 2006. Recent developments regarding the evolutionary origin of flowers. Adv. Bot. Res. 44, 63–127. Fukuda, T., Yokoyama, J., Maki, M., 2003. Molecular evolution of cycloidea-like genes in Fabaceae. J. Mol. Evol. 57, 588–597. Galego, L., Almeida, J., 2002. Role of DIVARICATA in the control of dorsoventral asymmetry in Antirrhinum flowers. Genes Dev. 16, 880–891. Gao, J.-Y., Ren, P.-R., Yang, Z.-H., Li, Q.-J., 2006. The pollination ecology of Paraboea rufescens (Gesneriaceae): a buzz-pollinated tropical herb with mirror-image flowers. Ann. Bot. 97, 371–376. Gao, Q., Tao, J.H., Yan, D., Wang, Y.Z., Li, Z.Y., 2008. Expression differentiation of CYC-like floral symmetry genes correlated with their protein sequence diver gence in Chirita heterotricha (Gesneriaceae). Dev. Genes Evol. 218, 341–351. Gaudin, V., Lunness, P.A., Fobert, P.R., Towers, M., Riou-Khamlichi, C., Murray, J.A., et al., 2000. The expression of D-cyclin genes defines distinct develop mental zones in snapdragon apical meristems and is locally regulated by the Cycloidea gene. Plant Physiol. 122, 1137–1148. Giurfa, M., Dafni, A., Neal, P.R., 1999. Floral symmetry and its role in plantpollinator systems. Int. J. Plant Sci. 160, S41–S50. Goebel, K., 1905. Organography of Plants. 1. General Organography; 2. Special Organography, Oxford University Press, Oxford. Goldschmidt, R., 1940. The Material Basis of Evolution, Yale University Press, New Haven.
132
H. CITERNE ET AL.
Go´mez, J.M., Bosch, J., Perfectti, F., Ferna´ndez, J.D., Abdelaziz, M., Camacho, J.P. M., 2008a. Spatial variation in selection on corolla shape in a generalist plant is promoted by the preference patterns of its local pollinators. Proc. R. Soc. Lond., B, Biol. Sci. 275, 2241–2249. Go´mez, J.M., Bosch, J., Perfectti, F., Ferna´ndez, J.D., Abdelaziz, M., Camacho, J.P. M., 2008b. Association between floral traits and rewards in Erysimum mediohispanicum (Brassicaceae). Ann. Bot. 101, 1413–1420. Go´mez, J.M., Perfectti, F., Camacho, J.P.M., 2006. Natural selection on Erysimum mediohispanicum flower shape: insights into the evolution of zygomorphy. Am. Nat. 168, 531–545. Gompel, N., Prud’homme, B., 2009. The causes of repeated genetic evolution. Dev. Biol. 332, 36–47. Gong, Y.B., Huang, S.Q., 2009. Floral symmetry: pollinator-mediated stabilizing selection on flower size in bilateral species. Proc. R. Soc. Lond., B, Biol. Sci. 276, 4013–4020. Gorelick, R., 2001. Did insect pollination cause increased seed plant diversity? Biol. J. Linn. Soc. Lond. 74, 407–427. Graham, S.W., Barrett, S.C.H., 1995. Phylogenetic systematics of Pontederiales: implications for breeding-system evolution. In: Monocotyledons: Systema tics and Evolution (Rudall, P.J., Cribb, P.J., Cutler, D.F., Humphries, C.J. (Eds.)), Royal Botanic Gardens, Kew, UK, pp. 415–441. Grant, V., 1949. Pollination systems as isolating mechanisms in angiosperms. Evolution 3, 82–97. Grimaldi, D., 1999. The co-radiations of pollinating insects and angiosperms in the Cretaceous. Ann. Mo. Bot. Gard. 86, 373–406. Grimaldi, D., Engel, M.S., 2005. Evolution of the Insects, Cambridge University Press, Cambridge, UK. Gu¨bitz, T., Caldwell, A., Hudson, A., 2003. Rapid molecular evolution of CYCLOIDEA-like genes in Antirrhinum and its relatives. Mol. Biol. Evol. 20, 1537–1544. Gustafsson, A., 1979. Linnaeus’ peloria: the history of a monster. Theor. Appl. Genet. 54, 241–248. Harder, L.D., Johnson, S.D., 2009. Darwin’s beautiful contrivances: evolutionary and functional evidence for floral adaptation. New Phytol. 183, 530–545. Helme, N., Linder, H., 1992. Morphology, evolution and taxonomy of Wachendorfia (Haemodoraceae). Bothalia 22, 59–75. Herrera, J., 2009. Visibility vs. biomass in flowers: exploring corolla allocation in Mediterranean entomophilous plants. Ann. Bot. 103, 1119–1127. Herrera, J., Arista, M., Ortiz, P.L., 2008. Perianth organization and intra-specific floral variability. Plant Biol. 10, 704–710. Hileman, L.C., Baum, D.A., 2003. Why do paralogs persist? Molecular evolution of CYCLOIDEA and related floral symmetry genes in Antirrhineae (Veroni caceae). Mol. Biol. Evol. 20, 591–600. Hileman, L.C., Cubas, P., 2009. An expanded evolutionary role for flower symmetry genes. J. Biol. 8, 90. Hileman, L.C., Kramer, E.M., Baum, D.A., 2003. Differential regulation of symme try genes and the evolution of floral morphologies. Proc. Natl. Acad. Sci. U.S.A. 100, 12814–12819. Howarth, D.G., Donoghue, M.J., 2005. Duplications in CYC-like genes from Dip sacales correlate with floral form. Int. J. Plant Sci. 166, 357–370. Howarth, D.G., Donoghue, M.J., 2006. Phylogenetic analysis of the “ECE” (CYC/ TB1) clade reveals duplications predating the core eudicots. Proc. Natl. Acad. Sci. U.S.A. 103, 9101–9106.
THE EVOLUTION OF FLORAL SYMMETRY
133
Howarth, D.G., Donoghue, M.J., 2009. Duplications and expression of DIVARICATA-like genes in Dipsacales. Mol. Biol. Evol. 26, 1245–1258. Hu, S.S., Dilcher, D.L., Jarzen, D.M., Taylor, D.W., 2008. Early steps of angiospermpollinator coevolution. Proc. Natl. Acad. Sci. U.S.A. 105, 240–245. Hubbard, L., McSteen, P., Doebley, J., Hake, S., 2002. Expression patterns and mutant phenotype of teosinte branched1 correlate with growth suppression in maize and teosinte. Genetics 162, 1927–1935. Jabbour, F., Damerval, C., Nadot, S., 2008. Evolutionary trends in the flowers of Asteridae: is polyandry an alternative to zygomorphy? Ann. Bot. 102, 153–165. Jabbour, F., Nadot, S., Damerval, C., 2009a. Evolution of floral symmetry: a state of the art. C. R. Biol. 332, 219–231. Jabbour, F., Ronse De Craene, L.P., Nadot, S., Damerval, C., 2009b. Establishment of zygomorphy on an ontogenic spiral and evolution of perianth in the tribe Delphinieae (Ranunculaceae). Ann. Bot. 104, 809–822. Jacob, F., 1977. Evolution and tinkering. Science 196, 1161–1166. Jesson, L.K., Barrett, S.C.H., 2002. The genetics of mirror-image flowers. Proc. R. Soc. Lond., B, Biol. Sci. 269, 1835–1839. Jesson, L.K., Barrett, S.C.H., 2003. The comparative biology of mirror-image flow ers. Int. J. Plant Sci. 164, S237–S249. Jesson, L.K., Kang, J., Wagner, S.L., Barrett, S.C.H., Dengler, N.G., 2003. The development of enantiostyly. Am. J. Bot. 90, 183–195. Kalisz, S., Purugganan, M.D., 2004. Epialleles via DNA methylation: consequences for plant evolution. Trends Ecol. Evol. 19, 309–314. Kalisz, S., Ree, R.H., Sargent, R.D., 2006. Linking floral symmetry genes to breeding system evolution. Trends Plant Sci. 11, 568–573. Kellogg, E.A., 2000. The grasses: a case study in macroevolution. Annu. Rev. Ecol. Syst. 31, 217–238. Kim, M., Cui, M.L., Cubas, P., Gillies, A., Lee, K., Chapman, M.A., et al., 2008. Regulatory genes control a key morphological and ecological trait transferred between species. Science 322, 1116–1119. Kirchoff, B.K., Lagomarsino, L.P., Newman, W.H., Bartlett, M.E., Specht, C.D., 2009. Early floral development of Heliconia latispatha (Heliconiaceae), a key taxon for understanding the evolution of flower development in the Zingiberales. Am. J. Bot. 96, 580–593. Krenn, H.W., Plant, J.D., Szucsich, N.U., 2005. Mouthparts of flower-visiting insects. Arthropod Struct. Dev. 34, 1–40. Kunze, H., 1985. Die Infloreszenzen der Marantaceen und ihr Zusammenhang mit dem Typus der Zingiberales- Synfloreszenz. Beitr. Biol. Pflanz. 60, 93–140. Lane, I.E., 1955. Genera and generic relationships in Musaceae. Mitt. Bot. Staats sammlung, Mu¨nchen 2, 114–131. Lehrer, M., 1999. Shape perception in the honeybee: symmetry as a global frame work. Int. J. Plant Sci. 160, S51–S65. Leppik, E.E., 1972. Origin and evolution of bilateral symmetry in flowers. Evol. Biol. 5, 49–85. Li, P., Johnston, M.O., 2000. Heterochrony in plant evolutionary studies through the twentieth century. Bot. Rev. 66, 57–88. Lide´n, M., 1986. Synopsis of Fumarioideae (Papaveraceae) with a monograph of the tribe Fumarieae. Opera Bot. 88, 1–133. Luo, D., Carpenter, R., Copsey, L., Vincent, C., Clark, J., Coen, E., 1999. Control of organ asymmetry in flowers of Antirrhinum. Cell 99, 367–376. Luo, D., Carpenter, R., Vincent, C., Copsey, L., Coen, E., 1996. Origin of floral asymmetry in Antirrhinum. Nature 383, 794–799.
134
H. CITERNE ET AL.
Manuel, M., 2009. Early evolution of symmetry and polarity in metazoan body plans. C. R. Biol. 332, 184–209. Marazzi, B., Endress, P.K., 2008. Patterns and development of floral asymmetry in Senna (Leguminosae, Cassinae). Am. J. Bot. 95, 22–40. Martı´n-Trillo, M. Cubas, P., 2009. TCP genes: a family snapshot ten years later. Trends Plant Sci. 15, 31–39. Mitchell, C.H., Diggle, P.K., 2005. The evolution of unisexual flowers: morphologi cal and functional convergence results from diverse developmental transi tions. Am. J. Bot. 15, 1068–1076. ¨ ber die Symmetrie der Pflanzen. Flora 20, 385–431. Mohl, H., 1837. U Mo¨ller, A.P., 1995. Bumblebee preference for symmetrical flowers. Proc. Natl. Acad. Sci. U.S.A. 92, 2288–2292. Mo¨ller, A.P., 2000. Developmental stability and pollination. Oecologia 123, 149–157. Mo¨ller, A.P., Eriksson, M., 1994. Patterns of fluctuating asymmetry in flowers: implications for sexual selection in plants. J. Evol. Biol. 7, 97–113. Mo¨ller, A.P., Shykoff, J.A., 1999. Morphological developmental stability in plants: patterns and causes. Int. J. Plant Sci. 160, S135–S146. Mo¨ller, A.P., Sorci, G., 1998. Insect preference for symmetrical artificial flowers. Oecologia 114, 37–42. Mondrago´n-Palomino, M. Theissen, G., 2008. MADS about the evolution of orchid flowers. Trends Plant Sci. 13, 51–59. Mondrago´n-Palomino, M. Theissen, G., 2009. Why are orchid flowers so diverse? Reduction of evolutionary constraints by paralogues of class B floral homeotic genes. Ann. Bot. 104, 583–594. Moylan, E.C., Rudall, P.J., Scotl, R.W., 2004. Comparative floral anatomy of Strobilanthinae (Acanthaceae), with particular reference to internal parti tioning of the flower. Plant Syst. Evol. 249(1–2), 77–98. Neal, P.R., Dafni, A., Giurfa, M., 1998. Floral symmetry and its role in plantpollinator systems: terminology, distribution, and hypotheses. Annu. Rev. Ecol. Syst. 29, 345–373. Ojeda, I., Francisco-Ortega, J., Cronk, Q.C., 2009. Evolution of petal epidermal micromorphology in Leguminosae and its use as a marker of petal identity. Ann. Bot. 104, 1099–1110. Ollerton, J., Alarcon, R., Waser, N.M., Price, M.V., Watts, S., Cranmer, L., et al., 2009. A global test of the pollination syndrome hypothesis. Ann. Bot. 103, 1471–1480. Ornduff, R., Dulberger, R., 1978. Floral enantiomorphy and the reproductive system of Wachendorfia paniculata (Haemodoraceae). New Phytol. 80, 427–434. Parkin, J., 1914. The evolution of inflorescence. J. Linn. Soc. 42, 511–563. Perez-Rodriguez, M., Jaffe, F.W., Butelli, E., Glover, B.J., Martin, C., 2005. Devel opment of three different cell types is associated with the activity of a specific MYB transcription factor in the ventral petal of Antirrhinum majus flowers. Development 132, 359–370. Poinar, G.O., Danforth, B.N., 2006. A fossil bee from Early Cretaceous Burmese amber. Science 314, 614–614. Prenner, G., Vergara-Silva, F., Rudall, P.J., 2009. The key role of morphology in modelling inflorescence architecture. Trends Plant Sci. 14, 302–308. Preston, J.C., Hileman, L.C., 2009. Developmental genetics of floral symmetry evolution. Trends Plant Sci. 14, 147–154. Preston, J.C., Kost, M.A., Hileman, L.C., 2009. Conservation and diversification of the symmetry developmental program among close relatives of snapdragon with divergent floral morphologies. New Phytol. 182, 751–762. Proctor, M., Yeo, P., Lack, A., 1996. The natural history of pollination, Harper Collins, London.
THE EVOLUTION OF FLORAL SYMMETRY
135
Ramsay, N.A., Glover, B.J., 2005. MYB-bHLH-WD40 protein complex and the evolution of cellular diversity. Trends Plant Sci. 10, 63–70. Rapp, R.A., Wendel, J.F., 2005. Epigenetics and plant evolution. New Phytol. 168, 81–91. Reardon, W., Fitzpatrick, D.A., Fares, M.A., Nugent, J.M., 2009. Evolution of flower shape in Plantago lanceolata. Plant Mol. Biol. 71, 241–250. Ree, R.H., Citerne, H.L., Lavin, M., Cronk, Q.C., 2004. Heterogeneous selection on LEGCYC paralogs in relation to flower morphology and the phylogeny of Lupinus (Leguminosae). Mol. Biol. Evol. 21, 321–331. Ren, Y., Li, H.F., Zhao, L., Endress, P.K., 2007. Floral morphogenesis in Euptelea (Eupteleaceae, Ranunculales). Ann. Bot. 100, 185–193. Renshaw, A., Burgin, S., 2008. Enantiomorphy in Banksia (Proteaceae): flowers and fruits. Aust. J. Bot. 56(4), 342. Rodriguez, I., Gumbert, A., de Ibarra, N.H., Kunze, J., Giurfa, M., 2004. Symmetry is in the eye of the “beeholder”: innate preference for bilateral symmetry in flower-naive bumblebees. Naturwissenschaften 91, 374–377. Ronse De Craene, L.P., 2003. The evolutionary significance of homeosis in flowers, a morphological perspective. Int. J. Plant Sci. 164, S225–S235. Ronse De Craene, L.P., 2010. Floral Diagrams—An Aid to Understanding Flower Morphology and Evolution, Cambridge University Press, Cambridge. Ronse De Craene, L.P., De Laet, J., Smets, E.F., 1998. Floral development and anatomy of Moringa oleifera (Moringaceae): what is the evidence for a Capparalean or Sapindalean affinity? Ann. Bot. 82, 273–284. Ronse De Craene, L.P., Smets, E., 1994. Merosity in flowers: definition, origin, and taxonomic significance. Plant Syst. Evol. 191, 83–104. Ronse De Craene, L.P., Smets, E., 2001. Floral developmental evidence for the sys tematic relationships of Tropaeolum (Tropaeolaceae). Ann. Bot. 88, 879–892. Ronse De Craene, L.P., Smets, E., Clinckemaillie, D., 2000. Floral ontogeny and anatomy in Koelreuteria with special emphasis on monosymmetry and septal cavities. Plant Syst. Evol 223, 91–107. Ronse De Craene, L.P., Soltis, P.S., Soltis, D.E., 2003. Evolution of floral structures in basal angiosperms. Int. J. Plant Sci. 164(5 Suppl.), S329–S363. Ronse De Craene, L.P., Yang, T.Y.A., Schols, P., Smets, E.F., 2002. Floral anatomy and systematics of Bretschneidera (Bretschneideraceae). Bot. J. Linn. Soc. 139, 29–45. Rosin, F.M., Kramer, E.M., 2009. Old dogs, new tricks: regulatory evolution in conserved genetic modules leads to novel morphologies in plants. Dev. Biol. 332, 25–35. Rudall, P.J., Bateman, R.M., 2002. Roles of synorganisation, zygomorphy and heterotrophy in floral evolution: the gynostemium and labellum of orchids and other lilioid monocots. Biol. Rev. Camb. Philos. Soc. 77, 403–441. Rudall, P.J., Bateman, R.M., 2003. Evolutionary change in flowers and inflores cences, evidence from naturally occurring terata. Trends Plant Sci. 8, 76–82. Rudall, P.J., Bateman, R.M., 2004. Evolution of zygomorphy in monocot flowers, iterative patterns and developmental constraints. New Phytol. 162, 25–44. Rudall, P.J., Bateman, R.M., 2010. Defining the limits of flowers: the challenge of distinguishing between the evolutionary products of simple versus com pound strobili. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 365, 397–409. Rudall, P.J., Bateman, R.M., Fay, M.F., Eastman, A., 2002. Floral anatomy and systematics of Alliaceae with particular reference to Gilliesia, a presumed insect mimic with strongly zygomorphic flowers. Am. J. Bot. 89, 1867–1883. Sargent, R.D., 2004. Floral symmetry affects speciation rates in angiosperms. Proc. R. Soc. Lond., B, Biol. Sci. 271, 603–608.
136
H. CITERNE ET AL.
Schmidt, R.J., Ambrose, B.A., 1998. The blooming of grass flower development. Curr. Opin. Plant Biol. 1, 60–67. Schumann, K., 1900. Musaceae. In: Das Pflanzenreich (Egler, H.G.A. (Ed.)), Engel mann, Leipzig, pp. 1–45. Sehr, E.M., Weber, A., 2009. Floral ontogeny of Oleaceae and its systematic implica tions. Int. J. Plant Sci. 170, 845–859. Smith, J.F., Funke, M.M., Woo, F.V., 2006. A duplication of gcyc predates diver gence within tribe Coronanthereae (Gesneriaceae): phylogenetic analysis and evolution. Plant Syst. Evol. 261, 245–256. Smith, J.F., Hileman, L.C., Powell, M.P., Baum, D.A., 2004. Evolution of GCYC, a Gesneriaceae homolog of CYCLOIDEA, within Gesnerioideae (Gesneria ceae). Mol. Phylogenet. Evol. 31, 765–779. Song, C.F., Lin, Q.B., Liang, R.H., Wang, Y.Z., 2009. Expressions of ECE-CYC2 clade genes relating to abortion of both dorsal and ventral stamens in Opithandra (Gesneriaceae). BMC Evol. Biol. 9, 244. Sprengel, C.K., 1793. Das entdeckte Geheimniss der Natur im Bau und in der Befruchtung der Blumen, Frederich Vieweg, Berlin. Sun, G., Ji, Q., Dilcher, D.L., Zheng, S., Nixon, K.C., Wang, X., 2002. Archaefruc taceae, a new basal angiosperm family. Science 296, 899–904. The Angiosperm Phylogeny Group classification for the orders and families of flowering plants: APG III. Bot. J. Linn. Soc.161,105–121 Theissen, G., 2000. Evolutionary developmental genetics of floral symmetry: the revealing power of Linnaeus’ monstrous flower. Bioessays 22, 209–213. Theissen, G., 2006. The proper place of hopeful monsters in evolutionary biology. Theory Biosci. 124, 349–369. Thorp, R.W., 2000. The collection of pollen by bees. Plant Syst. Evol. 222, 211–223. Tripp, E.A., Manos, P.S., 2008. Is floral specialization an evolutionary dead-end? Polli nation system transitions in Ruellia (Acanthaceae). Evolution 62, 1712–1736. Tsai, W.C., Kuoh, C.S., Chuang, M.H., Chen, W.H., Chen, H.H., 2004. Four DEFlike MADS box genes displayed distinct floral morphogenetic roles in Phalaenopsis orchid. Plant Cell Physiol. 45, 831–844. Tsou, C.-H., Mori, S.A., 2007. Floral organogenesis and floral evolution of the Lecythidioideae. Am. J. Bot. 94, 716–736. Tucker, S.C., 1996. Trends in evolution of floral ontogeny in Cassia sensu stricto, Senna, and Chamaecrista (Leguminosae, Caesalpinioideae, Cassieae, Cas siinae): a study in convergence. Am. J. Bot. 83, 687–711. Tucker, S.C., 1999. Evolutionary lability of symmetry in early floral development. Int. J. Plant Sci. 160, S25–S39. Tucker, S.C., 2002. Floral ontogeny in Sophoreae (Leguminosae: Papilionoideae). III. Radial symmetry and random petal aestivation in Cadia purpurea. Am. J. Bot. 89, 748–757. Tucker, S.C., 2003a. Comparative floral ontogeny in Detarieae (Leguminosae: Cae salpinioideae). III. Adaxially initiated whorls in Julbernadia and Sindora. Int. J. Plant Sci. 164, 275–286. Tucker, S.C., 2003b. Floral ontogeny in Swartzia (Leguminosae: Papilionoideae: Swart zieae): distribution and role of the ring meristem. Am. J. Bot. 90, 1271–1292. Tucker, S.C., Hodges, S.A., 2005. Floral ontogeny of Aquilegia, Semiaquilegia and Enemion (Ranunculaceae). Int. J. Plant Sci. 166, 557–574. Ushimaru, A., Dohzono, I., Takami, Y., Hyodo, F., 2009. Flower orientation enhances pollen transfer in bilaterally symmetrical flowers. Oecologia 160, 667–674. Ushimaru, A., Hyodo, F., 2005. Why do bilaterally symmetrical flowers orient vertically? Flower orientation influences pollinator landing behaviour. Evol. Ecol. Res. 7, 151–160.
THE EVOLUTION OF FLORAL SYMMETRY
137
van Kleunen, M., Meier, A., Saxenhofer, M., Fischer, M., 2008. Support for the predictions of the pollinator-mediated stabilizing selection hypothesis. J. Plant Ecol. 1, 173–178. Vincent, C.A., Coen, E.S., 2004. A temporal and morphological framework for flower development in Antirrhinum majus. Can. J. Bot. 82, 681–690. Wang, H.C., Meng, A.P., Li, J.Q., Feng, M., Chen, Z.D., Wang, W., 2006. Floral organogenesis of Cocculus orbiculatus and Stephania dielsiana (Menisper maceae). Int. J. Plant Sci. 167, 951–960. Wang, Z., Luo, Y., Li, X., Wang, L., Xu, S., Yang, J., et al., 2008. Genetic control of floral zygomorphy in pea (Pisum sativum L.). Proc. Natl. Acad. Sci. U.S.A. 105, 10414–10419. Wanntorp, L., Ronse De Craene, L.P., 2007. Flower development of Meliosma (Sabiaceae): evidence for multiple origins of pentamery in the eudicots. Am. J. Bot. 94, 1828–1836. Waser, N.M., 1998. Pollination, angiosperm speciation, and the nature of species boundaries. Oikos 82, 198–201. Weeks, E.L., Frey, F.M., 2007. Seed production and insect visitation rates in Hesperis matronalis are not affected by floral symmetry. Int. J. Plant Sci. 168, 611–617. Westerkamp, C., Classen-Bockhoff, R., 2007. Bilabiate flowers: the ultimate response to bees? Ann. Bot. 100, 361–374. Whitfield, J.B., Kjer, K.M., 2008. Ancient rapid radiations of insects: challenge for phylogenetic analysis. Annu. Rev. Entomol. 53, 449–472. Wignall, A.E., Heiling, A.M., Cheng, K., Herberstein, M.E., 2006. Flower symmetry preferences in honeybees and their crab spider predators. Ethology 112, 510–518. Wilson, A.C., Carlson, S.S., White, T.J., 1977. Biochemical evolution. Annu. Rev. Biochem. 46, 573–639. Winkler, H., 1930. Musaceae. In: Die natu¨rlichen Pflanzenfamilien (Engler, A. (Ed.)), Engelmann, Leipzig, pp. 505–541. Wolfe, L.M., Krstolic, J.L., 1999. Floral symmetry and its influence on variance in flower size. Am. Nat. 154, 484–488. Wood, T.E., Burke, J.M., Rieseberg, L.H., 2005. Parallel genotypic adaptation: when evolution repeats itself. Genetica 123, 157–170. Wydler, H., 1844. Einige Bemerkungen u¨ber die Symmetrie der Blumenkrone. Bota nische Zeitung 2, 609–611. Yuan, Z., Gao, S., Xue, D.W., Luo, D., Li, L.T., Ding, S.Y., et al., 2009. RETARDED PALEA1 controls palea development and floral zygomorphy in rice. Plant Physiol. 149, 235–244. Zhang, W., Kramer, E.M., Davis, C.C., 2010. Floral symmetry genes and the origin and maintenance of zygomorphy in a plant-pollinator mutualism. Proc. Natl. Acad. Sci. U.S.A. 107, 6388–6393. Zhou, X.R., Wang, Y.Z., Smith, J.F., Chen, R., 2008. Altered expression patterns of TCP and MYB genes relating to the floral developmental transition from initial zygomorphy to actinomorphy in Bournea (Gesneriaceae). New Phy tol. 178, 532–543.
Protein Turnover in Grass Leaves
LOUIS JOHN IRVING,*,1 YUJI SUZUKI,† HIROYUKI ISHIDA†
AND AMANE MAKINO†
*
Graduate School of Life and Environmental Science, University of Tsukuba, 1-1-1 Tennodai, Tsukuba 305-8572, Japan † Graduate School of Agricultural Science, Tohoku University, 1-1
Tsutsumidori Amamiyamachi, Aoba-ku, Sendai 560-0043, Japan
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cellular Regulation of Protein Synthesis . . . . . . . III. Protein Degradation. . . . . . . . . . . . . . . . . . . . . . . . . . A. Cellular Regulation of Protein Degradation. . . . . B. Plant Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . C. Chloroplast Stromal Protein Degradation . . . . . . D. Thylakoid-Associated Proteins . . . . . . . . . . . . . . E. Environmental Regulation of Senescence. . . . . . . IV. Whole-Leaf Regulation of Protein Content. . . . . . . . . V. Implications of Protein Turnover in Whole Plants . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1
... ... ... ... ... ... ... ... ... ... ... ... ... ...
.... .... .... .... .... .... .... .... .... .... .... .... .... ....
.... .... .... .... .... .... .... .... .... .... .... .... .... ....
140
141
141
146
146
149
151
157
159
162
168
171
172
172
Corresponding author: E-mail:
[email protected] Advances in Botanical Research, Vol. 54 Copyright 2010, Elsevier Ltd. All rights reserved.
0065-2296/10 $35.00
DOI: 10.1016/S0065-2296(10)54004-7
140
L. J. IRVING ET AL.
ABSTRACT
In this chapter, we discuss the processes of protein synthesis and degradation at the cellular, organ and whole-plant levels. In particular, we focus on the leaf protein Rubisco, which is important as both the most abundant form of N in most leaves and the carboxylating enzyme in photosynthesis. Chloroplasts contain the largest fraction of cellular N, divided approximately equally between soluble protein and thylakoidassociated N. Recently, small vesicles have been noted emanating from chloroplasts; however, there is considerable debate on the properties and regulation of these bodies. Similarly, recent investigations into the turnover of the D1 protein have questioned the orthodoxy view that D1 turnover is caused by oxidative fragmentation. The final two sections of this chapter look into the factors influencing the patterns of protein synthesis and degradation at the whole-leaf and whole-plant levels, and the implica tions that has for plant growth, development and productivity.
I. INTRODUCTION In most environments, nitrogen (N) is the predominant element limiting plant growth and, in agronomic systems, yield (Addiscott et al., 1991). The majority of N in most plants is stored in aboveground organs, primary among them are leaves. In cereals, the aboveground biomass may contain as much as 80% of plant N, with leaves accounting for approximately 60% of that. Leaf N concentration scales with photosynthetic capacity (Wright et al., 2004), and the N content of plants is closely related to their nutritional quality for animals, including humans. N remobilization is important as an N supply for new growth, with over 60% of new leaf N being derived from protein turnover in older leaves (Mae, 1986). A recent paper showed that the average period in which N had been in the plant prior to its incorporation into a new leaf was 14 days, compared to 3 days for C (Lattanzi et al., 2005). Similarly, remobilized old leaf N has also been shown to be the main N source for new spring growth in perennial grasses (Bausenwein et al., 2001). For these reasons, in this chapter we shall discuss the processes of N remobilization in plant leaves. We shall focus on grass plants although we will use other species where they provide a useful example. Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco; EC4.1.1.39) will be especially focused on for two reasons: (i) Rubisco is the carboxylating enzyme in photosynthesis; and (ii) due to its low enzymatic rate, Rubisco is the most abundant enzyme not only in most plant tissues but in the world (Ellis, 1979). A third factor making Rubisco interesting is its method of production, with its small subunits encoded in the nuclear genome and translated in the cytosol, while large subunit biosynthesis takes place in the chloroplast.
PROTEIN TURNOVER IN GRASS LEAVES
141
This chapter is split into the following main sections: Section II discusses the regulation of protein synthesis at the cellular level, while Section III does the same for degradation. Sections IV and V will look at the integration of synthesis and degradation at the whole-leaf level and the implications of differential N turnover at the whole-plant level, respectively. At this point, I would like to note that this chapter is not intended to be comprehensive in its treatment of this subject. The processes described in this chapter are becoming better and better investigated every day, and the volume of information available on this subject is simply too large for even a single book, let alone one review paper. This chapter would be better thought of as an introduction to the subject, providing readers with the opportunity to branch out and explore individual subjects at will.
II. PROTEIN SYNTHESIS A. CELLULAR REGULATION OF PROTEIN SYNTHESIS
Within the leaf, a vast majority of N, perhaps 70% or more, is found within the chloroplast. Of that, the majority, perhaps three-quarters, is associated with photosynthesis (Makino and Osmond, 1991). Endosymbiotic theory tells us that all eukaryotic cells result from ancestral bacteria taking up residence inside other cells, probably around a billion years ago. This accounts for the double-membrane structure of higher plant chloroplasts, with the inner membrane being derived from the ancestral bacteria and the outer membrane deriving from the host cell (some marine microalgae have a four-membrane structure, revealing that the original endosymbiont underwent a secondary endosymbiosis). In accordance with their bacterial ancestry, eukaryotic organelles, specifically mitochondria and chloroplasts, maintain their own genomes. These genomes are much reduced compared to extant bacteria, as gene functions, if not genes themselves, were trans ferred from the bacterial genome to the nuclear genome over evolutionary time. The chloroplast genome encodes approximately 100 chloroplast proteins, while nuclear genes encode a number probably 30 times higher (Aluru et al., 2006). The expression of the nuclear and chloroplast gen omes must be synchronized in their action during the cells lifetime, although it is not well understood how this is regulated. However, com munication between the chloroplast and the nucleus is known to be bidirectional. Bacteria use a class of proteins known as sigma factors to regulate the activity of RNA polymerase, and these sigma factors have also been found in the higher plant nuclear DNA (Lysenko, 2007). Free sigma
142
L. J. IRVING ET AL.
factors 1–3 do not bind to the plastid DNA, but rather they bind with RNA polymerase, which causes a conformational change allowing them to bind later with the plastid DNA. The mechanism of action of the sigma factors 4–6 remains unclear. Sig6 mutants are known to have decreased rbcL and psbA transcript abundances, suggesting that this gene may be important in nucleus to chloroplast signalling in developing leaves. The regulation of Rubisco synthesis, further discussed below, is a good example of chloroplast behaviour being modulated by the behaviour of the nucleus, while the downregulation of the nuclear light-harvesting complex b gene (Lhcb) and the Rubisco small subunit (RBCS) gene (RbcS) in response to tissue treatment with a carotenoid biosynthesis inhibitor (La Rocca et al., 2004), and in chloroplast defective mutants, demonstrates chloroplast to nucleus signalling (Aluru et al., 2006). Carotenoid biosynthesis mutants, such as the immutans mutant line of Arabidopsis, suffer photo-oxidative stress, leading to chloroplast bleaching, with sections of immutans leaves losing all their green chlorophyll colouration. We might also expect that nuclear genes which are responsive to light follow some kind of chloroplast to nuclear signalling. In grasses, new cells are produced during cell division, which takes place at the base of the leaf. New cells contain about 15 small spherical prochloroplasts, 1–2 mm in diameter. Mature cells contain up to 60 chloro plasts per cell, with a size range of approximately 6–8 mm. During this maturation process, chloroplast DNA increases rapidly, up to 7.5-fold per chloroplast in wheat, which is presumably important in the rapid produc tion of the vast quantities of protein which must be synthesized for photosynthesis. In illuminated barley leaves, the number of plastid DNA copies was maximal in cells at the base of the leaf and decreased as the cells matured. However, in leaves expanding in darkness, this decrease in plastid DNA copy number with cell age was not noted (Shaver et al., 2008). After this initial increase, the genome copy number rapidly decreases upon exposure to blue or white, but not red light (Oldenburg et al., 2006). In dividing cells, the rate of plastid transcription is very low; however, as the chloroplasts start to mature, transcription activity increases approximately 10-fold per plastid, and due to the increased number of plastids, approximately 30-fold per cell. This upregulation in transcription precedes increases in RNA levels, ribosome numbers and photosynthetic proteins (Baumgartner et al., 1989), and appears to be light-regulated (Dubell and Mullet, 1995). Sigma factors would seem to be implicated in this light signalling, with some sigma factors considerably upregulated upon illumination (Lysenko, 2007). Additionally, many sigma factors are only expressed in illuminated tissues, such as leaves, but not in roots.
PROTEIN TURNOVER IN GRASS LEAVES
143
Higher plant Rubisco is composed of eight large (Rubisco large subunit (RBCL)) and eight small subunits (RBCS). The large subunits are encoded by the chloroplast genome (plastome), whereas the small are encoded by the nuclear genome. The small subunit RNA is translated into protein in the cytosol and then imported into the chloroplast through the chloroplast membrane, losing its transit peptide in the process (Highfield and Ellis, 1978). rbcL mRNA is translated in the chloroplast, by bacterial type 70S ribosomes. RBCS protein entering the chloroplast binds with the RBCL, which is otherwise quickly degraded in the absence of RBCS. RBCL excess represses the translation of further rbcL, which seems generally to be in excess (Wostrikoff and Stern, 2007). While this system in which unassembled proteins are unstable and are rapidly degraded is common in many organ isms, unassembled cytochrome f protein is no less stable than the assembled protein; however, its transcription is massively decreased in the absence of its assembly partners (Choquet and Vallon, 2000; Choquet et al., 2003). This downregulation of protein synthesis is known as “control by epistasy of synthesis” (CES). CES appears to be very important in the control of thylakoid membrane protein synthesis. Many historical studies have used antisense mutant plants to investigate the subunit control of Rubisco biosynthesis (Hudson et al., 1992; Makino et al., 1997b; Quick et al., 1992) and its downstream effects on photosynth esis; however, only recently have the opposite rbcS-overexpressing plants been successfully produced. Suzuki et al. (2007), using rbcS-overexpressing plants, noted a 2.1–2.8-fold increase in rbcS, a 1.2–1.9-fold increase in rbcL and a 1.3-fold increase in Rubisco content. This suggests that Rubisco synthesis is regulated at several levels. rbcL levels were upregulated by rbcS overexpression; it seems likely that excess RBCS bound with RBCL which would be otherwise in excess, thus increasing the Rubisco protein levels and derepressing the production of rbcL. Rubisco concentrations increased by only 70% of the increase in rbcL transcript, however, suggest ing a further level of regulation, which has been suggested to be mainly controlled by leaf N-influx (Imai et al., 2005). Of course, the rate of N import into a leaf represents an upper limit to the total amount of N-containing molecules which can be produced by the leaf, and so perhaps this result is unsurprising. Tobacco rbcS mutants with only 5% of wild-type rbcS transcript exhibited wild-type levels of rbcL transcript, but only 5% of wild-type RBCL protein (Wostrikoff and Stern, 2007). Similar results have also been noted in other studies. For example, rbcS antisense plants produce significantly less rbcS mRNA than control plants, but the levels of rbcL transcript were unchanged (Rodermel et al., 1996). RbcS also utilizes post transcriptional control to regulate protein levels, with Chlamydomonas cells
144
L. J. IRVING ET AL.
unable to produce the large subunit due to a lack of 70S ribosomes exhibiting decreased rbcS mRNA levels (Mishkind and Schmidt, 1983). Rubisco rbc gene expression exhibits a strong diurnal fluctuation (Cheng et al., 1998). In Arabidopsis, rbc levels increase overnight, then decrease sharply upon illumination—especially those of rbcS 1B. Following this initial decline, rbc transcript abundances increase in an almost linear manner in rbcS 1A and 1B. Although several control mechanisms can be postulated for these patterns of transcript abundance, this linear phase suggests that transcription may take place in a largely constitutive manner. However, the regulation of rbcS 2B and 3B appears to be more complicated, although the reason for this remains unclear. Both the fluence rate and wavelength are important, with blue light shown to be important in the upregulation of rbcS transcript abundance, whereas red and white light were less important (Lopez-Juez et al., 2007; Sawbridge et al., 1994). Various classes of photo receptors are known to be active in plants, and these have been implicated in the regulation of both gene transcription (Spalding and Folta, 2005) and proteolysis (Huq, 2006). Grass leaves tend to green up and produce Rubisco as they exit from the previous leaf sheath (Gastal and Nelson, 1994) and it seems plausible that the mechanism promoting Rubisco synthesis in chloroplasts may be some light-mediated signal. Phytochromes make a good candidate as a major signalling mechanism for light-induced protein synthesis when they are transported to the nucleus upon photostimulation (Spalding and Folta, 2005). Indeed phytochrome-interacting factor 1 (PIF1) has been implicated in the greening of seedling plants, with PIF1 mutants accumulating a chlorophyll precursor which renders seedlings sensitive to light stress (Huq et al., 2004). Carbohydrate accumulation, especially glucose, represses the transcript abundances of several photosynthetic genes, including Rubisco. Indeed, sugar levels are well known to regulate transcription (Sheen, 1990). Recent evidence has shown a negative correlation between leaf glucose levels and rbcS transcript abundances, in both wild-type and ethylene-insensitive Nicotiana plants (Acevedo-Hernandez et al., 2005). Acevedo-Hernandez’s study exposed the cells to a range of sugar concentrations, noting that higher glucose availabilities rescued rbcS transcription after the initial reduction, implying multiple promoters and repressors, operating differentially under varying conditions. Interestingly, in the absence of glucose, abscisic acid (ABA) had a negative effect on rbcS transcript abundance, but a positive effect in the presence of glucose. Older studies, however, suggest that it is an intermediate in carbohydrate metabolism, rather than carbohydrate accu mulation per se which controls rbcS levels (Krapp et al., 1991, 1993). Although transcription is generally thought to be more important in
PROTEIN TURNOVER IN GRASS LEAVES
145
determining mRNA levels, the importance of mRNA turnover should not be underestimated, with this being an important yet poorly understood factor allowing nuclear control of plastid genes (Drager et al., 1998). Other studies have shown that carbohydrate status is important in regulating chloroplast biogenesis itself, with Arabidopsis grown with supplemental glucose failing to green up and exhibiting significantly decreased levels of hexadecatrienoic acid—a major constituent of chloroplast membranes (To et al., 2003). Similarly with ABA, cytokinins are also able to modify transcript levels, specifically upregulating them in barley leaves (Zubo et al., 2008). As discussed in more detail later, cytokinins are also able to retard proteolytic rates, while ABA has been shown to promote senescence. Zubo et al. (2008) used detached barley leaves in their experiments, showing that cytokinin application had the strongest effect on the apical, and therefore oldest, cells. Several genes were upregulated in response to exogenous cytokinin application, including the rbcL gene; however, interestingly the psbA and psbD genes, encoding the photosystem II (PSII) D1 and D2 proteins, respectively, were unaffected by cytokinin. Light appeared to be required for gene upregulation by cytokinins (Zubo et al., 2008). Nitrate supply to maize roots has been shown to lead to cytokinin transport to the leaves, which may act as a long-range signalling mechanism of N sufficiency (Saka kibara et al., 1998). Atmospheric CO2 concentration appears to have a strong influence on rbcS gene expression, with increased transcript abundances noted at low CO2 levels (Gesch et al., 1998). Rice cultivars grown under elevated CO2 levels and temperatures showed strong genotype-dependent changes in rbcS transcript abundance (Gesch et al., 2003). The CO2-mediated decrease in rbcS transcript was temperature-dependent; one variety showed a CO2 dependent decrease in rbcS only at 28˚C, and not at either 34˚C or 40˚C, while the second variety exhibited a decrease at 34˚C and 40˚C, but not 28˚C. Increased temperature caused a decrease in transcript abundance at 350 ppm CO2, but not in plants grown at 700 ppm CO2 (Gesch et al., 2003). Other studies have similarly shown a decrease in leaf Rubisco concentration by both elevated CO2 and temperature in both rice and soybean (Vu et al., 1997).The mechanism by which these decreases are affected was unclear, whether due to a decrease in Rubisco synthesis or due to an increase in degradation. Increased photosynthesis at elevated CO2 causes substantial increase in leaf sugar concentrations, especially sucrose and hexose (Vu et al., 2001), and sugars have been linked to increased proteolytic rates (Wingler et al., 2006). However, C status is only one part of the puzzle, with N status also having important implications for gene expression (Scheible et al., 2004) and,
146
L. J. IRVING ET AL.
ultimately, the levels of photosynthetic enzymes within plant leaves (Matt et al., 2002). Increasing N availability leads to increased leaf Rubisco contents in hydroponically grown rice plants (Makino et al., 1984b); however, it is less clear how this relates to cellular Rubisco contents because plants grown at higher N levels also have larger leaves and larger cells. Thus, changes in leaf Rubisco concentration may be buffered at intermediate N levels due to changes in leaf size and thickness. Scheible et al. (2004) showed that the supply of N to N-starved Arabidopsis seedlings upregulated genes involved in chlorophyll biosynthesis, the photosynthetic light reactions and also numerous genes associated with the Calvin cycle and starch biosynthesis.
III. PROTEIN DEGRADATION A. CELLULAR REGULATION OF PROTEIN DEGRADATION
In many eukaryotic cells, specific protein degradation in the cytosol and the nucleus is carried out in a ubiquitin-dependent pathway by the 26S proteo some. Ubiquitin conjugates with the protein to be degraded, which is then degraded by serine and cysteine proteases (CPs) in the proteosome (Buchanan-Wollaston et al., 2005). Protein degradation is generally considered to occur in a stepwise manner, with an initial rate-limiting cleavage by a specific enzyme followed by further degradation of the cleavage products by generalist proteases. Although the general assumption is of a single protease being the rate-limiting step, the degradation of a single protein may be regulated by multiple pathways (Callis, 1995). Protein degradation is generally an ATP-dependent process, despite being an exogonic reaction, which could theoretically proceed without the requirement for energy (ATP) supply. Leaf senescence is well regulated, with individual cells senescing in a well defined order. In grasses, senescence primarily starts from the tip, the oldest cells, working towards the base of the leaf, with the cells around the leaf veins taking longest to senesce (Wingler et al., 2004). In Arabidopsis, simi larly, a well-defined pattern of cell senescence can be seen, with cells around the veins taking longest to senesce, with similar patterns emerging in rice coleoptiles (Inada et al., 1998a, b). Since nutrient export from the leaves is dependent on the transfer of nutrients to new leaves through the phloem, cells bordering the veins would be expected to senesce last. An additional factor may be that cells bordering the phloem are more able to export C than those further away, since increased sugar levels in the cells can cause
PROTEIN TURNOVER IN GRASS LEAVES
147
senescence. Cells further away from the phloem may build up sugars more rapidly, promoting increased rates of senescence. However, as far as we are aware, this possibility has not been explored in detail, but would be interesting. Senescence timing and characteristics can vary greatly depending on the environmental conditions under which the plant lives. For example, Wingler et al. (2004) grew Arabidopsis plants on agar with varying levels of N and glucose. Plants growing at reduced N supply visibly accumulated anthocya nins—plant pigments expressed under light stress conditions, which appear to function as a photosynthetic “light shield” (Albert et al., 2009). Nonphotosynthetic quenching (NPQ) was shown to increase during senescence, while the dark-adapted fluorescence yield (Fv/Fm) was relatively unaffected. This tells us that while chlorophyll levels are remaining relatively stable, a smaller fraction of the incident energy is being used to drive photochemistry. Increased NPQ is indicative of light stress, while the liberation of freechlorophyll from the thylakoid membranes can drive the production of free-radicals of oxygen. Oxygen radicals are hugely dangerous for the cells, and complex biochemical pathways exist to mitigate their production (Asada, 2006). These mechanisms are not 100% effective, however, and the chloroplast becomes a more oxidizing environment through time (McRae and Thompson, 1983). We might hypothesize that these increases in active oxygen levels cause the vacuole rupture which precedes cell necrosis (Obara et al., 2001), although this is unknown. While the addition of glucose to low-N media leads to an increase in the rate of leaf senescence in Arabidopsis, plants grown on high-N media with supplemental glucose did not exhibit accelerated senescence and were dark green than the non-glucose supple mented plants (Wingler et al., 2004). Many genes coding for protein kinases and phosphatases, along with several genes involved in calcium metabolism, are upregulated during senes cence, indicating the functioning of kinase-signalling cascades (BuchananWollaston et al., 2005). There is some evidence linking increased calcium levels with cell death (Huang et al., 1997), with calcium being necessary for the functioning of some CPs (Callis, 1995). A vast array of genes involved in N metabolism and sugar, peptide, amino acid and cation membrane trans porters are also upregulated. Transcript abundances for genes involved in glutamate/glutamine metabolism are upregulated, including glutamate receptor proteins and glutamine synthetase (GS) (Buchanan-Wollaston et al., 2005). Several genes involved in ABA signalling were upregulated, while various cytokinin-induced genes were downregulated. This suggests an increase in ABA levels and a decrease in cytokinin levels in senescing leaves. Exogenously supplied cytokinins have been shown to retard chlorophyll loss
148
L. J. IRVING ET AL.
in excised Arabidopsis leaves (Sergiev et al., 2007), and cytokinin decrease has been hypothesized to be a major factor in shade-induced senescence (Causin et al., 2009). Similarly, a maize stay-green mutant exhibited increased levels of leaf cytokinins, while ABA levels were decreased com pared to an early senescent variety (He et al., 2005). Both cytokinin and ABA have been shown to have effects on leaf sugar regulation (Yang et al., 2002), with one study suggesting that a mutant’s stay-green phenotype resulted from decreased C transport from leaves (Yang et al., 2003) although it is unclear how much of the extra remobilization and transfer of stored C is directly hormonally mediated, and how much is simply a result of decreased amounts of recent photosynthate as ABA causes stomatal closure (Acharya and Assmann, 2009). Recent photoassimilate is generally important in sup plying C for growth (Lattanzi et al., 2005), with photosynthesis expressed per unit plant mass and relative growth rate showing a strong linear relation ship (Kruger and Volin, 2006). During starvation-induced senescence in cell suspension culture, many of the cytokinin-inducible genes downregulated under natural senescence were not expressed at all, while little change from control conditions could be seen under dark conditions. Since cytokinins tend to retard senescence, the absence of cytokinin-inducible genes presumably reflects an absence of cytokinins and increased proteolytic rates. Cytokinins appear to be impor tant in dark-induced senescence; however, the mechanism of dark-induced senescence is unclear, as the expression of cytokinin-induced genes is quite dissimilar to that in control of C-starved conditions (Buchanan-Wollaston et al., 2005). Various other hormones have been reported to be linked to leaf senes cence, and transcriptomic approaches have demonstrated increases in tran scripts related to, for example, jasmonic acid and salicylic acid metabolism (Van der Graaff et al., 2006). Jasmonic acid has been shown to increase over the period of senescence in the leaf (He et al., 2002), but like ABA both jasmonic acid and salicylic acid also accumulate during stomatal closure, and have been linked to that (Acharya and Assmann, 2009). On the other hand, there are some suggestions that senescence-associated genes (SAGs) are dependent on salicylic acid (Van der Graaff et al., 2006). Finally, two other classes of compounds, gibberellins and brassinosteroids, were identi fied by Van der Graaff et al. (2006) as potential hormones involved in senescence, despite direct evidence being relatively scarce for these two chemicals. Interestingly, along with all the other hormones discussed above, both gibberellins and brassinosteroids were identified by Acharya and Assmann (2009) as being involved in determining stomatal aperture. Thus, the exact relationship between stomatal aperture and senescence
PROTEIN TURNOVER IN GRASS LEAVES
149
remains unclear, although it is clear that a relationship exists. Ethylene is understood to be important in determining the rates of leaf senescence, with ethylene-insensitive mutants exhibiting increased leaf lifespan, including an increased duration of protein retention, compared to control plants (Grbic and Bleecker, 1995). Ethylene was also shown to increase the transcript abundances of SAGs, although the effects of ethylene were more pro nounced in old leaves than in younger leaves, suggesting age-dependent effects. However, like the control plants, ethylene-insensitive plants also upregulated SAGs in senescent leaves, although this was delayed compared to control plants (Grbic and Bleecker, 1995). This suggests rather that ethylene has a role in moderating the timing of leaf senescence, although ultimately the leaves senesced in both control and ethylene-insensitive varieties. The majority of proteolysis takes place in the vacuole, due to its low pH (Bassham, 2009), with both proteases (Yoshida and Minamikawa, 1996) and protein degradation products (Huffaker, 1990) found there. Proteins from the cytosol or chloroplasts must be taken into the vacuole by the process of autophagy (ATG). ATG was originally characterized in yeast (Tsukada and Ohsumi, 1993), but many ATG genes have also been identified in plants, especially Arabidopsis (Hanaoka et al., 2002; Inoue et al., 2006), but also in maize (Chung et al., 2009). Recently, Van der Graaff et al. (2006) noted that 19 out of 21 identified ATG genes were upregulated during natural senescence. Genes for cytosolic GS were not strongly expressed in dark-mediated senescence. These genes generally fix ammonia produced during photore spiration, which can be assumed to be negligible under dark conditions. However, other N-related genes are upregulated; for example, various trans aminases are increased under dark-induced senescence (BuchananWollaston et al., 2005).
B. PLANT PROTEASES
Plant cell endoproteases can be divided into four main groups: CPs, serine proteases, metalloproteases and aspartate proteases (Callis, 1995). A fifth group, consisting of those which cannot be identified as one of the above four, also exists. CPs generally have an acidic pH optimum and are generally located in the vacuole. However, as mentioned above, CPs have also been implicated in regulating protein degradation of chloroplast stromal constituents (Minamikawa et al., 2001; Prins et al., 2008), although the specific location
150
L. J. IRVING ET AL.
of degradation remains unclear. Certainly, the stromal pH should be too high for CPs to work efficiently. As mentioned previously, the 26S proteosome is a proteolytic complex found in the cytosol and nucleus of eukaryotic cells. The proteosome is a complex of generic proteases, which tend to have a neutral pH. Prior to degradation by the proteosome, ubiquitination of target proteins is required. The attachment of ubiquitin to the protein is an ATP-dependent process (Demartino and Slaughter, 1993) achieved through the action of three enzymes: the ubiquitin-activating enzyme (E1), the ubiquitin-conjugating enzyme (E2) and the ubiquitin-protein ligase (E3) (Moon et al., 2004). E3 protein ligases are substrate-specific, and over 90% of the approximately 1400 genes involved in the ubiquitin proteosome pathway code for E3 protein ligases (Smalle and Vierstra, 2004). Phytochrome-mediated light signals have been demonstrated to increase the activity of some E3 protein ligases, which may have an important role in photoacclimation or other regulatory processes (Huq, 2006). The proteosome is generally considered important in the cell cycle, with mutation in 26S proteosome genes prevent ing successful mitosis (Ghislain et al., 1993; Gordon et al., 1993). Genes involved in the ubiquitination of proteins for degradation were not greatly affected by the resupply of N to N-deficient Arabidopsis seedlings, a finding which may be surprising given that we might expect comprehensive changes in plant growth strategy under N-deficient and N-sufficient conditions (Scheible et al., 2004). Various stresses may also upregulate protease expression. For example, osmotic and salt stress has been shown to lead to an increase in mRNA transcript abundances, although this was not related to abscissic acid levels (Koizumi et al., 1993). CP mRNA levels increased in cold-treated tomato fruits (Schaffer and Fischer, 1988), while heat shock is well known to induce proteolytic activity in bacterial and yeast cells (Finley et al., 1987; Katz et al., 2009). Oxidative and UV-B stress leads to Rubisco fragmentation (Desimone et al., 1996; Ishida et al., 1997, 1998); however, these fragments accumulate to levels detectable by sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDS–PAGE), suggesting that either protease levels in the chloroplast or the cell or the rate of transport of the fragments to the site of degradation is insufficient to degrade the fragments before they start to build up. It seems likely that these fragments would be in a proteolytically suscep tible state, implying that even the generic proteases which would normally be involved in fragment degradation are limiting. Oxidative modification is known to enhance the binding of Rubisco protein to the chloroplast envel ope (Marin-Navarro and Moreno, 2006), presumably making vesicular transport from the chloroplasts to the vacuole by Rubisco-containing bodies
PROTEIN TURNOVER IN GRASS LEAVES
151
(RCBs) (discussed below) more likely. Other recent work has shown that oxidative stress can lead to an upregulation of macroautophagy in Arabidopsis, presumably as a mechanism to deal with oxidatively damaged proteins. RNA is perhaps particularly susceptible to degradation by UV-B, which may limit the de novo synthesis of proteases under these conditions (Takeuchi et al., 2002). Proteolysis is known to be important in seed germi nation, and often the proteases involved were synthesized during seed filling and drying. It is tempting to hypothesize that the production of these proteases is upregulated by water stress during seed desiccation. Vacuolar proteases have been shown to be able to degrade the Rubisco large, but not small, subunit (Yoshida and Minamikawa, 1996). This has led to the speculation that the majority of proteolysis must occur in the vacuole, with proteins being transported from other cellular locations into the vacuole. This will be discussed in more detail below, however, it should be pointed out here that the fact that the vacuole contains proteases, which are capable of degrading Rubisco, does not mean that Rubisco degradation is their primary, or even normal, function—many general proteases exist, able to degrade a wide range of proteins. Small subunit degradation is relatively understudied, although some reports exist (Miller and Huffaker, 1982). Miller and Huffaker used 14C-labelling of Rubisco and found three endo proteases with the ability to degrade the Rubisco, naming them EP1, EP2 and EP3. EP1 is a chloroplast-located metalloprotease, which primarily acts upon the RBCL. EP1 degrades Rubisco to a 36-kDa fragment in vitro, and it seems likely that EP1 also acts in a similar manner in vivo. EP2 primarily acted against the small subunits, while EP3 caused the autolysis of Rubisco purified by SDS–PAGE. Unassembled proteins, or those with mistakes, tend to be rapidly degraded in vivo, with unassembled 32S-labelled yeast fatty acid synthase a-subunit being almost completely turned over within 6 hr after the end of the labelling period (Baek and Choi, 2008), although that is outside the scope of this chapter. C. CHLOROPLAST STROMAL PROTEIN DEGRADATION
As leaves senescence, the protein concentration of the individual cells decreases. Early studies showed that protein concentration decreased at a much higher rate than the loss of chloroplasts within cells (Mae et al., 1984; Wardley et al., 1984). This suggests that chloroplast loss itself cannot be solely responsible for the noted decrease in leaf Rubisco concentration through time, suggesting that Rubisco and other chloroplast stromal
152
L. J. IRVING ET AL.
proteins must either be degraded within the chloroplast or be exported from the chloroplast then degraded. Chloroplast protein fragments have been isolated in the vacuole (Huffaker, 1990), implying that the vacuole is a major site of proteolysis in plants, and it is known that wheat chloroplasts shrink through time (Ono et al., 1995). Recent studies have shown chloro plasts budding off small spherical Rubisco-containing vesicles, dubbed RCBs, which are then degraded within the vacuole (Chiba et al., 2003; Ishida et al., 2008; Wada et al., 2009). While RCBs were named based on their concentration of Rubisco, it is clear that they also contain other stromal proteins, for example, Gln synthetase, or chloroplast-targeted green fluor escent protein (GFP) in its native form (Ishida et al., 2008), which may imply a lack of specificity for stromal proteins. RCBs do not, however, contain thylakoid membranes, with Wada et al. (2009) noting that chlorophyll was absent from the fluorescence emission spectra of RCBs; however, it is unclear whether the degradation products of thylakoid-associated proteins are transported in RCBs. However, immunolocalization studies failed to show any signal when RCBs were stained with antibodies for thylakoid proteins, such as LHCII or cytochrome (Chiba et al., 2003). In ATGdeficient mutant Arabidopsis plants, which lack the capacity to form RCBs, the rate of Rubisco degradation was unchanged, while chlorophyll catabolism was increased in the mutants (Wada et al., 2009). This implies the existence of other mechanisms of Rubisco degradation, probably stromal proteases, and may even suggest that RCB production is incidental and unnecessary for Rubisco degradation. On the other hand, Wada et al. (2009) showed the build up of GFP-labelled stromal proteins in the vacuoles of darkened control plant leaves treated with concanamycin A, which inhi bits vacuolar ATPase causing a pH shift repressing protease activity. This clearly suggests that stromal proteins are transported to the vacuole without the breakdown of GFP, and presumably other proteins also. Furthermore, the increased chlorophyll catabolism in the atg4s mutant leaves suggests elevated protease levels in the chloroplasts of these plants compared to control plants, further implying that the breakdown mechanism in the atg mutants is not the normal degradation pathway. The atg genes were dis covered initially in yeast; however, they were later noted in Arabidopsis and appear to be important in the production of RCBs. The atg mutants used in Ishida et al. (2008) exhibit increased stromule activity. Stromules are stromafilled protrusions of the chloroplast envelope, although their function is unclear (Holzinger et al., 2007). Interestingly, the rate of proteolysis is identical whether RCBs can be formed or not, which suggests that the rate of Rubisco degradation may be controlled by some factor preceding either export by RCBs or degradation by chloroplast proteases.
PROTEIN TURNOVER IN GRASS LEAVES
153
The structure and function of stromules remains unclear, although various functions have been postulated. However, as chloroplasts are derived from ancient free-living bacteria which formed an endosymbiotic relationship with proto-plant cells (Sagan, 1967), and bacteria routinely transfer genetic infor mation in the process of conjugation, stromules may be derived from this process. It seems plausible that RCBs could be derived from stromule protuberances, with RCB-sized vesicles being pinched off from stromules noted previously (Arimura et al., 2001; Waters et al., 2004; Gunning, 2005). Furthermore, similar to RCBs, these stroma-containing vesicles do not contain thylakoid material (Holzinger et al., 2007). RCB production within leaves is known to vary over time (Chiba et al., 2003). In Ishida et al. (2008), the number of RCBs in the vacuole of concanamycin A-treated cells was noted for the fourth, sixth, eighth and tenth leaves of Arabidopsis. While RCB number was low in the youngest leaves (10th) showing high rbcS 2B gene expression, which suggests active Rubisco synthesis, RCBs were present suggesting some turnover even in expanding leaves. RCB number increased approximately fourfold in the eighth leaf, then a further twofold in the sixth leaf. RCB numbers were approximately similar in the fourth and sixth leaves, although variability in the number of RCBs in the fourth leaves was higher (Ishida et al., 2008). This pattern of RCB numbers correlates with the rates of Rubisco throughout the leaves’ lifespan, which is considered to be relatively low during leaf expan sion, increasing around full leaf expansion and then generally high during senescence. Factors controlling RCB production remain unknown, yet these patterns of RCB production through time suggest strong ontogenic effects, although it is unclear whether these are genetic or environmental. RCB studies are generally conducted on darkened leaves and it is difficult to visualize RCBs in leaves which have not been incubated in the dark. Light acts both as the energy source for photosynthesis and via photoreceptor chemicals such as phytochrome as a signal in its own right. It is unclear, therefore, whether RCBs are produced to remobilize N from shaded leaves or RCB formation is controlled by leaf sugar status. Certainly, it is known that whole-darkened plants do not senesce (Weaver and Amasino, 2001). These plants similarly do not grow, so the lack of proteolysis may be due to either a reduction in the sink strength of developing leaves or a lack of C derived under normal circumstances from other leaves. ATG of whole chloroplasts takes place predominantly close to the end of the leaves lifespan. Wada et al. (2009) showed an increased rate of chlor oplast ATG in darkened Arabidopsis leaves. The Arabidopsis plants used had been bred with chloroplast–stroma-targeted GFP, allowing the visualization of RCBs by laser-scanning confocal microscopy (Fig. 1). While RCBs
154
L. J. IRVING ET AL.
Fig. 1. Visualization of RCBs in living leaf cells of Arabidopsis by stromal-targeted GFP and laser-scanning confocal microscopy. Stromal-targeted GFP is shown by pseudo green and chlorophyll fluorescence is by pseudo-red and the merged image is shown. Chloroplasts having both GFP and chlorophyll appear yellow, whereas RCBs having only GFP appear green. (See Color Insert.)
strongly exhibited the GFP signal, vacuolar-located chloroplasts exhibited fluorescence signals associated with chlorophyll fluorescence only, with no stromal proteins apparent. It is unclear how much stromal protein the chloroplasts contained at the time of transport, or whether stromal proteins are simply more proteolytically susceptible and were degraded upon the chloroplast being subsumed. There are some reports that the DNA content of chloroplasts decreases at a relatively early point during senescence, pre ceding large-scale proteolysis (Inada et al., 1999). Small Rubisco-containing vesicles have been noted by other investigators exploring senescence in tobacco, although reported as “senescence-associated vacuoles” (SAVs) (Martinez et al., 2008; Otegui et al., 2005). SAVs were noted to have a pH lower than the central vacuole, which in turn is lower than chloroplast pH values (Laisk et al., 1989; Oja et al., 1999). Martinez et al. (2008) used transformed tobacco plants with chloroplast-targeted cyan fluor escent protein (CFP). Leaves were induced to senesce and produce SAVs by detachment from the plant and incubation in continuous darkness and it is unclear in Martinez et al.’s (2008) study whether SAVs could be visualized
PROTEIN TURNOVER IN GRASS LEAVES
155
under normal conditions; however, RCBs cannot be seen under control condi tions, so it is unclear whether SAVs can be visualized under more normal conditions. However, contrary to the situation in RCBs, Martinez et al. (2008) noted that some tobacco SAVs exhibited a chlorophyll fluorescence signal; however, they did not note this in either Arabidopsis or soybean in an earlier study (Otegui et al., 2005). This may confirm either that RCBs and SAVs are indeed separate and independent organelles, or that the inclusion of chlorophyll in RCBs is species-specific, or that chlorophyll degradation products were found in true RCBs by Martinez et al. (2008). Dark-incubation, carbohydrate starva tion and natural senescence all seem to be separate processes, with very different transcript profiles for each, and it is possible that either RCB can contain chlorophyll under these conditions or SAVs are unique vacuoles associated with a wounding response to the leaf detachment. Interestingly, Martinez et al. (2008) noted that dark-incubated, excised tobacco leaves treated with ethylene produced approximately double the number of SAVs than control leaves, untreated with ethylene. The percentage of these containing the GFP signal was approximately the same as in the control plants, and it was suggested that this indicates a higher rate of degradation. It is unclear exactly how quantitative this work is though, and doubling the number of SAVs will not lead to a greater rate of protein degradation if the protein concentration in those SAVs is halved. Martinez et al. (2008) also noted high levels of chloroplastic GS2 in isolated SAVs as well as large quantities of Rubisco. Furthermore, they demonstrated the presence of proteases inside isolated SAVs, by showing that the addition of protease inhibitors in the culture media retarded the degradation of the RBCL. Martinez et al. (2008) went on to conclude that since SAVs contain both proteases and chloroplast proteins, they have an important role in mediating the loss of chloroplast proteins in tobacco. However, significantly, they failed to quantify the rates of protein loss from the chloroplasts or to provide evidence that SAVs have sufficient protease activity to achieve noted rates of protein degradation. In Wada et al.’s (2009) study, plants unable to produce RCBs suffered no retardation in their ability to degrade Rubisco—degradation rates were identical in control and RCB-less plants—suggesting that RCBs, and potentially SAVs too, are unnecessary to explain Rubisco degradation in plant leaves. Interestingly, Wada et al. (2009) did not note any small spherical bodies in ATG-deficient (atg) mutant plants, nor any accumulation of stromal targeted GFP in the vacuoles of concanamycin A-treated mutants, suggesting either that SAVs are not present in Arabidopsis or that SAV production is similarly dependent on the ATG gene, or that protein transported in SAVs is degraded en route to the vacuole. For SAVs and RCBs to be both ATG5 dependent would be a huge coincidence, which while not obviously silly, seems statistically implausible. However, both Martinez et al.’s (2008)
156
L. J. IRVING ET AL.
evidence of chlorophyll constituents in SAVs and Wada et al.’s (2009) evidence of the lack of chlorophyll fluorescence in RCBs are troublesome, and it is hard to reconcile these apparently contradictory reports other than by invoking the somewhat unsatisfying answer of “species-specificity”. SAVs differ from RCBs in a second key way—while RCBs exhibit a double-membrane structure, SAVs exhibit only a single membrane. The reason for this is again unclear, although it may be that while RCBs are enclosed in a second membrane, transported to the vacuole and assimilated in macroautophagy, SAVs may undergo microautophagy and be assimi lated directly into the vacuole. If this is true, it may suggest that SAVs and RCBs are in fact the same structures, with Martinez et al. (2008) simply noting either chlorophyll breakdown products in the vacuoles or the sensi tivity of their HPLC approach simply being far higher than the sensitivity of Wada et al.’s (2009) chlorophyll fluorescence approach. As Martinez et al. (2008) did not quantify the proteins noted, it is difficult to differenti ate between these possibilities. However, while the chlorophyll signals noted in Martinez et al.’s (2008) paper (Fig. 3b in Martinez at al. (2008)) are rather weak, the GFP signal is much stronger, and it is difficult to see any blue colouration in the merged image. Furthermore, no chlorophyll fluor escence can be noted in a SAV-enriched fraction. Isolated chloroplasts cannot degrade Rubisco completely, with only the initial cleavage of the RBCL occurring, yielding a 44-kDa fragment (Kokubun et al. 2002). Wada et al.’s (2009) data, however, clearly showed that RCBs are unnecessary for Rubisco degradation. This implies that Rubisco is not exported from the chloroplast, even by leakage or membrane transport, simply since we do not see leakage of GFP from the chloroplast, but instead that Rubisco (and GFP, and presumably other proteins) is degraded within the chloroplast. These two pieces of information taken together, the inability of isolated chloroplasts to degrade Rubisco coupled with the degradation of Rubisco within the chloroplast in mutants unable to produce RCBs, suggest a cytoplasmic origin of the majority of Rubisco proteases, agreeing with the schemes of both Prins et al. (2008) and Martinez et al. (2008). Isolated chloroplasts, treated with thermolysin to degrade contaminating proteases adhering to the chloroplast envelope, displayed the ability to degrade the RBCL, in a manner that was time-, temperatureand pH-dependent (Zhang et al., 2007). Contrary to Kokubun et al.’s (2002) data, Zhang et al. (2007) noted that the initial cleavage product was a 51-kDa fragment, although this may have been a result of the chloroplasts being dark induced to senesce. Zhang et al. (2007) noted that the cleavage site of their noted 51-kDa fragment was similar to that for a previously noted vacuolar protease. Since the experiment was conducted using isolated
PROTEIN TURNOVER IN GRASS LEAVES
157
chloroplasts, the presence of a vacuolar protease seems strange; however, it seems plausible that some proteases may be transported along with Rubisco and other proteins from the chloroplast to the vacuole in RCBs, where the proteases, but not the Rubisco, accumulate. Zhang et al.’s (2007) data further suggested that the vacuole is an unlikely venue for at least the initial few cleavages of the Rubisco subunits, although it seems likely that the later stages of degradation occur there. Recent work has illustrated the importance of CPs in the degradation of chloroplast proteins, including Rubisco and Rubisco activase (Minamikawa et al., 2001; Prins et al., 2008). Minamikawa et al. (2001) used antibodies raised against Rubisco and a CP in detached French bean leaves, noting that over the 8-day incubation period, chloroplasts were taken into the vacuole and degraded. Prins et al. (2008) overexpressed a rice cystatin in tobacco plants, which blocks the protease action of the CPs, leading to significantly higher leaf protein concentrations, and commensurately higher photosyn thetic rates, than control tobacco leaves, apparently due to a decrease in proteolysis. Unfortunately, since protein levels were not been quantified through time for individual leaves, it is unclear whether these increases in protein concentration represent the steady increase in leaf protein through time, or whether the majority of the proteins were produced during leaf expansion, with relatively little subsequent protein synthesis. Repression of CPs by cystatin led to an increase in total CP activity, which may suggest that CP expression and activity may be modulated by amino acid or protein levels. While the major protein degradation site is postulated to be the vacuole, cystatin is largely found in the cytosol, although was also found in the chloroplasts and vacuole. D. THYLAKOID-ASSOCIATED PROTEINS
Other proteins, for example, those associated with the thylakoid membranes do not appear to be exported from the chloroplast and degraded, but rather to be degraded in situ. The D1 protein may have one of the shortest half-lives of any protein in any living system and may be the most researched. The D1 protein forms a complex with the homologous D2 protein—both of which have a molecular mass of approximately 32 kDa. The D1–D2 complex forms a part of the PSII reaction centre. During illumination, the D1 complex is degraded, although the specific mechanism by which this occurs is disputed. Older literature with much current literature argues that D1 is initially cleaved by the action of oxygen radicals on the protein. This would seem to be borne out by evidence showing that D1 degradation rate is strongly dependent on light intensity, with degradation rates increasing rapidly
158
L. J. IRVING ET AL.
between 0 and 250 mmol photons m-2 sec-1, and more slowly thereafter (Edelman and Mattoo, 2008). However, it does raise the question of why D1 degradation is not progressively faster at higher irradiances. Other, more recent, investigations have raised alternate possibilities, such as the hypothesis that the primary action of reactive oxygen species is in inhibiting D1 synthesis, which negatively affects the repair cycle of D1 (Takahashi and Murata, 2008). Treatment with propyl gallate, a reactive oxygen species scavenger, has been reported to promote D1 synthesis (Edelman and Mattoo, 2008). Suitably high light will also cause damage to the D2 protein, which is also degraded under high levels of photo-oxidative stress. Unlike the D1 protein, however, two beta-carotene molecules bind to the D2 protein, quenching oxygen radicals, and protecting D2 from degradation (Telfer, 2005). Proteolysis of the D1 protein appears to follow a structural modification to the protein structure, possibly a covalent modification (Andersson and Aro, 1997). Although this conformational modification is generally light-dependent, detergent treatment of the PSII complex may induce D1 protein degradation in darkness (Nakajima et al., 1995, 1996). Similarly, the use of a PSII inhibitor, PNO8 (N-octyl-3-nitro-2,4,6 trihydroxybenzamide) facilitated the degradation of the D1 protein in dark ness. Furthermore, in Chlamydomonas, when a conformational change in D1 is suppressed by the interaction of the PQH2 ligand with the QB site of D1, the subsequent degradation of D1 is also prevented (Zer et al., 1994). These data, taken together, suggest that the enzymes required for the degradation of D1 are available at all times; however, the conditions for the conforma tional change under normal conditions generally only occur in the daytime, under high light conditions. Recent evidence has suggested a key role for the FtsH proteases in the turnover of the D1 protein (Kato et al., 2009), although it is unclear whether FtsH causes the primary cleavage of the D1 protein, or whether it is more important in its subsequent degradation. Mutant plants lacking the FtsH protease tend to accumulate higher levels of oxygen radicals than wild-type plants. ATP-dependent post-transcriptional phosphorylation appeared to be a second mechanism regulating the degradation of the D1 protein in higher plants (Kettunen et al., 1991); however, later studies, in which the break down of unphosphorylated control D1 protein and D1 protein which had been phosphorylated by exogenous addition of [g-32P]ATP, showed that phosphorylated D1 degradation occurs at a lower rate than unphosphory lated D1 (Andersson and Aro, 1997). Plants grown under low light conditions produce a greater quantity of the LHCII protein than those grown under high light. LHCII is a protein involved in the capture of photons during photosynthesis and is the most
PROTEIN TURNOVER IN GRASS LEAVES
159
abundant thylakoid protein. At high photon flux densities, high levels of LHCII are unnecessary and presumably would lead to increased light stress. Thus, after a lag period of approximately 2 days, chloroplast LHCII con tents are decreased. Thus it appears to be effected by the degradation of the outermost LHCII proteins, by an as-yet unknown ATP-dependent cysteine or serine protease (Yang et al., 1998). The rate of proteolysis is substrate limited. Furthermore, like the D1 protein discussed above, phosphorylation appears important in determining the rate of LHCII degradation with phosphorylated LHCII being a poor substrate for degradation, similar to the D1 protein. The major site for the degradation of thylakoid proteins appears to be the stroma-exposed membrane regions, rather than the appressed regions, where thylakoid membranes are “stacked” one on top of the other. This suggests that thylakoid protein degradation may be facilitated by stromal proteases. It is known, however, that neither Clp nor FtsH are responsible for the degradation of LHCII (Yang et al., 1998). E. ENVIRONMENTAL REGULATION OF SENESCENCE
Various environmental factors are well known to regulate the rate of leaf senescence, with many signalling pathways leading to chlorosis. For exam ple, sugars are understood to be implicit in the process, with high leaf sugar levels promoting protein degradation and leaf senescence (Wingler et al., 2006). Young expanding leaves are C deficient, as the processes of cell division, expansion and protein synthesis require large amounts of energy. However, as leaf expansion completes, and the chloroplasts mature, we would expect the leaf’s C budget to shift from being negative to positive, and for the leaf to accumulate sugars and become a net C source. Indeed, sugar contents of Arabidopsis leaves increase through time, with additional glucose further increasing the apparent rate of senescence, as determined by reductions in Fv/Fm (Wingler et al., 2006). Cold treatment can delay senescence, even despite the accumulation of sugars and ABA, factors which normally promote senescence (MasclauxDaubresse et al., 2007). The reasons for the accumulation of sugars in coldtreated plants are not well understood; however, it could simply be related to lower growth and respiration rates at reduced temperatures. Certainly, the cold-grown plants shown in Fig. 2A of Masclaux-Daubresse et al. (2007) were much smaller than the control plants. Reduced growth rates would almost certainly lead to a large divergence from the plants’ normal source— sink ratio. In some ways, this may be similar to the whole-darkened plants noted above, which do not grow, and do not show visible symptoms of senescence.
160
L. J. IRVING ET AL.
RCB
0.5 µm
RCB
2 µm
Fig. 2. Detection of RCB in naturally senescing wheat leaves by immunoelectron microscopy. Gold particles bind to RBCL (black spots). RCBs seem to be surrounded by autophagosome-like membrane structures.
Both water deficit and waterlogging can lead to premature senescence (Irving et al., 2007; Yang et al., 2002). Yang et al. (2003) noted that water-deficient wheat plants exhibited increased stem ABA, decreased cytokinin levels and increased rates of chlorophyll loss, when compared to water-sufficient control plants. While a direct hormonal role on pro tease levels or activity cannot be ruled out, it seems plausible that ABA effects are indirect. ABA plays an important role in regulating plant water status, causing stomatal closure under drought conditions (Acharya and Assmann, 2009), and has been directly implicated in the production of cellular reactive oxygen species (Zhang et al., 2001). Furthermore, stoma tal closure may limit photosynthetic rates, promoting N loss from the leaf like ammonia produced during photorespiration, and also further causing the production of oxygen radicals. Cytokinins, conversely, promote stomatal aperture and decrease stomatal sensitivity to ABA, which may help to maintain photosynthetic rates, preventing the production of oxygen radicals and retarding senescence (Criado et al., 2009). Cytoki nins may have a secondary effect, by promoting cell division and leaf growth, and causing an increase in protein synthesis. The process by which
PROTEIN TURNOVER IN GRASS LEAVES
161
waterlogging causes senescence remains unclear, but may be related to carbo hydrate build up in the stem (Castonguay et al., 1993), leading to a sugarbased repression of photosynthesis (Araya et al., 2006) and increased light stress. Senescence may, ironically, proceed somewhat along the same lines in both water-deficient and waterlogged plants. Interestingly, suppression of drought-induced senescence by the overexpression of an isopentenyltransfer ase gene in tobacco led to a superior drought resistance, with the plants maintaining the ability to photosynthesize during the drought treatment, and even improved survival, with the transgenic plants surviving a drought which killed the control plants (Rivero et al., 2007). Conversely, root water logging may lead to a pH decrease in the root medium, with leaf damage caused by reduced leaf pH as water is transpired through the plant. Other stresses may be important in determining protein levels; for example, supple mental ozone supply has been demonstrated to decrease Rubisco transcript abundances, particularly rbcL levels (Glick et al., 1995). Similarly, water deficit has been shown to cause dose-dependent decreases in both Rubisco gene transcript abundances and Rubisco concentrations, and it is unclear whether Rubisco degradation rates were upregulated or the reduced protein levels were solely a result of suppressed protein synthesis (Bauer et al., 1997). Many hundreds of SAGs have been identified over the years, although some of these have been identified under somewhat unnatural conditions, for example, in detached or dark-incubated leaves, and so some caution should be exercised (Buchanan-Wollaston et al., 2003; Chen et al., 2002; He et al., 2001). Developmental senescence (i.e. natural senescence) and starvation-induced senescence have been compared, and significant differ ences noted, particularly at the gene transcript level. Similarly, Phaseolus leaves when detached from the plant senescence more rapidly than attached leaves (Yoshida and Minamikawa, 1996). The physiological relevance of studies in which plants are placed under extreme condition such as these is questionable; however, because it is unclear whether the noted increases in senescence were related to leaf C balance, water con tent, light stress, were hormonally related, or for some other reason. Microarray analysis of transcript abundance in Arabidopsis leaves undergoing natural, starvation-induced or dark-induced senescence indi cated profound differences in the number and identities of genes upregu lated. While nearly 2000 genes were upregulated under dark-induced conditions, and nearly 1500 genes were upregulated under starvationinduced senescence, only 827 genes were upregulated under control conditions (Buchanan-Wollaston et al., 2005). This makes it likely that a huge number of SAGs, which have been identified under starvation- or
162
L. J. IRVING ET AL.
dark-induced senescence conditions, have no function in natural senes cence, and may suggest that studies using dark-induced senescence have limited relevance when considering natural leaf senescence. Of course, the specific mechanisms causing the senescence of leaves under natural con ditions remain unclear and may be the result of either genetic or environ mental factors—further complicating our evaluation of the importance and the role of individual SAGs under any given conditions. Another study reported the upregulation of approximately 1300 genes during natural senescence in Arabidopsis (Van der Graaff et al., 2006). Van der Graaff et al. (2006) also reported that approximately 2000 genes were regulated during stress-induced senescence, although some of these 2000 must have been downregulated. The reasons for the differences between these two studies are hard to reconcile but may be related to the different growth and sampling conditions used in the two studies. Irrespective of these inconsistencies, it is clear that senescence is a genome-wide event, involving the action of many genes, in networks which are currently almost completely obscure. Cell death is the final stage in the senescence of the leaf cell. During cell necrosis, the cell’s vacuole lyses (Obara et al., 2001), lowering the pH of the cells and releasing vacuolar proteases which degrade the cellular constituents for remobilization to other organs.
IV. WHOLE-LEAF REGULATION OF PROTEIN
CONTENT
However, regulation at the whole-leaf and whole-plant levels is more poorly understood, despite continued efforts at understanding the processes involved (Hidema et al., 1991; Imai et al., 2005, 2008; Ishizuka et al., 2004; Mae et al., 1983; Makino and Osmond, 1991; Makino et al., 1997b; 1984a, b; Suzuki et al., 2001, 2007). A recent paper developed a mathematical modelling process to help elucidate component processes and further understand the regulation of Rubisco synthesis at a whole-leaf level (Irving and Robinson, 2006). Rubisco synthesis takes place in a physiologically well-defined zone, 50–100 mm from the site of cell division (Gastal and Nelson, 1994). According to the Irving and Robinson (2006) model, through time Rubisco synthesis rate approximates a normal curve, correlating with leaf expansion rates, while degradation follows the first-order kinetic rules; degradation rates are initially low during leaf expansion increasing to a maximum just after full leaf expansion when Rubisco concentration is maximal. This is
PROTEIN TURNOVER IN GRASS LEAVES
163
followed by a period of senescence at a steadily decreasing rate, as the degradation rate decreases with decreasing Rubisco concentration, through to leaf death. Thus, through time, Rubisco content can be described by a log-normal curve. Rubisco turnover, as defined here, represents the sum of activity of both the biosynthesis and degradation of Rubisco within the leaf (i.e. faster turnover equals more synthesis and degradation, slower turnover less). A recent review expressed some scepticism over the validity of using a single decay constant in the Irving and Robinson (2006) model, citing the stabilizing effects of CA-1-P binding and Rubisco activation in preventing Rubisco degradation (Parry et al., 2008). This point of view may find some support in the data of Ishizuka et al. (2004), which suggest that Rubisco degradation rates are greatly decreased and cannot be described by a single decay constant. However, caution should be exercised in interpreting the data of Ishizuka et al. (2004) as they does not represent a true time course, in which they measured Rubisco contents in leaves at different positions, rather than making measurements on the same leaf of replicate plants through time. Furthermore, it is not clear whether Rubisco turnover kinetics is identical in all leaves, or whether the protein turnover rates change with plant age. Finally, we would point out that the degradation mechanism of Rubisco remains very unclear, we do not know, for example, whether Rubisco is primary degraded at night time or during the day, or whether it is continuous and invariant. Thus, it may be premature to attempt a quanti tative argument for the importance or otherwise of Rubisco activation or inhibitory compound on Rubisco turnover rates. Irrespective of the logic of Parry et al. (2008) that CA-1-P and other compounds should lead to a variable rate of Rubisco catalysis, isotopic labelling experiments such as those of Suzuki et al. (2001) return data, which are consistent with a single decay constant for Rubisco turnover. Furthermore, the Irving and Robinson (2006) model predicts Rubisco synthesis rates, which correlate very strongly with transcript abundances, and is completely consistent with Lattanzi et al.’s (2005) steady-state labelling experiment to understand N remobiliza tion in ryegrass plants. Leaf N content tends to be higher in plants grown at higher irradiances except in plants grown at very low N availabilities, although the allocation of N to Rubisco did not change with light intensity in rice leaves (Makino et al., 1997a). Chlorophyll content, however, appeared to vary with light intensity, with plants grown at low irradiances exhibiting increased chlorophyll levels. A similar lack of large differences in N allocation as a result of changes in the fluence rate has also been observed in both Chenopodium and Alocasia (Hikosaka and Terashima, 1996). Another,
164
L. J. IRVING ET AL.
more recent study showed that neither light intensity nor temperature affected the proportion of N invested in Rubisco; however, under condi tions likely to promote oxidative stress (low temperature and high light intensity), Plantago exhibited decreased chlorophyll levels (Hikosaka, 2005b). Light seems to have little effect on the partitioning of N within the leaf, yet is certainly necessary for the greening process, as leaves grown in the dark do not green up. Similarly grass leaves produce the majority of their Rubisco as they emerge from the previous leaf sheath, with even fairly large leaves containing little Rubisco before emergence. Whole-leaf Rubisco content increases rapidly during leaf expansion, followed by a peak around full leaf expansion and a subsequent decline in the leaf Rubisco concentration (Mae et al., 1983). The loss of Rubisco post-full leaf expansion follows an approximately exponential decrease, with the maximum rate of Rubisco loss occurring a few days after full leaf expansion. As the chloroplast numbers decrease only slightly during this period, it implies that the majority of Rubisco degradation and N loss during this period are attributable to the degradation of proteins within the chloroplast and export of proteins from the chloroplast by RCBs (Ishida et al., 2008). This was proposed to be a more economically viable method of facilitating the remobilization of proteins from senescent leaves to new tissues than whole-chloroplast senescence. Rubisco concentrations are often in excess for immediate photosynthetic requirements (Warren and Adams, 2004), a phenomenon which may have been exacerbated in the last century by increases in atmospheric CO2 partial pressures, and photosynthesis is limited by light interception and BuBP regeneration. However, this raises the question of why Rubisco, and presumably other proteins, is so strongly expressed to levels higher than those required for photosynthesis in expanding leaves. Some authors have hypothesized that “excess” Rubisco may represent a store of N for plants (Lawlor, 2002); however, one could argue that such a store would be suboptimal in terms of plant growth, since that N could be better used to increase the leaf area and thus energy interception by the plant. To minimize the expenditure of energy in protein synthesis, while still maintaining high photosynthetic rates, surely the optimal strategy would be for plants to express Rubisco at lower levels, but then maintain Rubisco contents in the leaves for a longer period, without the peak followed by rapid decline in Rubisco content around full leaf expansion notable in grass leaves. This would free up some N for growth, perhaps leading to larger leaf areas or increased tillering. However, neither in rice (Mae et al., 1983), wheat (Mae et al., 1989), barley (Friedrich and Huffaker, 1980), Arabidopsis (Wada et al., 2009) nor in ryegrass (Lolium perenne; LJ Irving and
PROTEIN TURNOVER IN GRASS LEAVES
165
C. Matthew, unpublished) do we see any sort of plateau, rather we see a sharp peak, followed by approximately exponential decline in Rubisco content. This suggests that Rubisco contents cannot be precisely regulated and that degradation proceeds at some predefined rate. Initial processing by some protease located inside the chloroplast may be responsible for this. Indeed, as mentioned above, isolated chloroplasts, incubated in the dark exhibited degradation of the RBCL (Kokubun et al., 2002). The quantity of degradation products increased in a time-dependent manner, suggesting a limiting concentration of the active agent, and temperature and pH dependence suggest a protease. Other investigations in tobacco suggest that a DNA-binding protease, CND41 (chloroplast nucleoid DNA binding protein), may be the initial Rubisco protease (Kato et al., 2004). In most species, N is remobilized from the lower, older leaves to younger leaves, yielding a canopy gradient in N content. However, tobacco CND41 antisense mutants exhibit an unusual N distribution, with the soluble protein levels approximately equal between young expanding leaves and older leaves. Furthermore, while the control plants showed a gradient in their leaf Rubisco contents, from youngest to oldest, the antisense plants did not exhibit these patterns, suggesting a suppression of even the initial stage of Rubisco degradation. This work highlights the importance of N remobilization from old leaves to younger leaves, with young CND41 antisense leaves having significantly lower protein levels than the control plants (Kato et al., 2004). The rate of proteolysis in isolated chloroplasts was estimated by Kokubun et al. (2002) at 4% of the starting concentration of the large subunit per day, very similar to Irving and Robinson’s (2006) estimates of 3.5% per day modelled from isotopic labelling studies. It is notable that in Kokubun et al. (2002), the Rubisco degradation products accumulated, suggesting that chloroplasts are unable to further degrade Rubisco, and presumably other proteins, by themselves. Given these facts, the Irving and Robinson (2006) model presented an alternate hypothesis to explain excess Rubisco; given that grasses seem unable to closely regulate their rate of Rubisco degradation, and that photosynthetic rates correlate with Rubisco concentrations, over investment in Rubisco may be a strategy by the plant to maintain maximal photosynthesis for a longer period of the leaf lifespan, and thus maximize photosynthetic gains per unit of C and N invested in a leaf. Rubisco is generally understood to be protected from degradation when it is in its catalytically active form, or when bound to CA-1-P (Khan et al., 1999). Kato et al. (2004) showed that while urea-treated inactive Rubisco could be readily degraded by the CND41 protease, activated Rubisco was less susceptible to degradation. The urea treatment noted would cause the
166
L. J. IRVING ET AL.
denaturation of inactive Rubisco, which presumably allowed its degrada tion. Interestingly, however, the optimal pH for the activity of CND41 is in the acidic range, while chloroplast stromal pH is typically slightly alkali. Pre-treatment of Rubisco in acidic media increased its degradation rate by CND41, and it is unclear quite how CND41 acts in the chloroplast, although changes in chloroplast redox state through time may play a part. Similarly, Martinez et al. (2008) showed that the pH of their SAVs was in the acidic range, around 5.8. Nutrient limitation has been shown to increase the rate of decline in leaf protein concentration. This is normally posited as an increase in the degradation rate; however, there is little evidence for this position. The net leaf protein concentration is determined by the rates of protein synth esis and degradation, and a change in either of these will affect the leaf protein concentration. Although some factors such as leaf sugar levels appear to have effects on RCB production, it seems likely that a part of these changes in protein concentrations is caused by declines in protein synthesis rates. Although nutrient limitation increases the net loss of Rubisco and other proteins, leaf lifespan has been shown to be increased for leaves of some species grown at lower N availabilities (Aerts, 1989), although opposite patterns, or a lack of difference, have been shown in other species (Aerts and Decaluwe, 1995). Oikawa et al. (2006) noted that Xanthium canadense plants grown at high N availabilities had a significantly longer leaf lifespan than plants grown under low N levels. All other things being equal, reduced leaf lifespan will reduce the amount of light captured by a leaf, and hence its total C gain. Older leaves tend to contain less N as it is remobilized through time, notably the leaf Rubisco content decreases rapidly after full leaf expansion (Mae and Ohira, 1981; Mae et al., 1983), and this correlates with the reductions in photosynthetic rate (Makino et al., 1983). Despite this, Oikawa et al. (2006) clearly showed that the cumulative net C gain of leaves correlates with time, implying that increase in leaf lifespan would lead to more C being fixed by that leaf. The mechanism by which N availability extends leaf lifespan is unclear; however, two potential general mechanisms can be posited. We know that C sufficiency can promote senescence, and both high and low N leaves should be able to equally accumulate C—if anything, high N leaves should accumulate more C due to their higher levels of photosynthetic proteins. This suggests that either the C/N ratio is important or N acts in some way to shift the balance towards protein synthesis, thus delaying senescence. The quantity of Rubisco synthesized by a plant depends on its environmental conditions, with high N plants producing more
PROTEIN TURNOVER IN GRASS LEAVES
167
Rubisco than their low N counterparts (Makino et al., 1984b). While the low N leaves in Makino et al.’s (1984b) study appeared to senescence earlier than the high N leaves, the curve shapes between the high, medium and low N leaves were very similar, suggesting that the control of leaf Rubisco content by N operates by regulating synthesis rather than degra dation. Removing N from the growth medium of wheat leaves led to significant decreases in both the leaf Rubisco content and the RNA transcript abundance (Crafts-Brandner et al., 1998). While direct C : N ratio sensing is not an obviously bad idea, in practice it seems harder to understand exactly how such a system would function. Early senescence of leaves grown under low N is easier to understand, especially if the proteolytic rates are fairly constant, where the leaves simply run out of protein to degrade earlier than higher N controls. Similarly, some system relying on leaf C status itself would seem relatively easy to imagine. N and other nutrients are transferred from older plant parts to newer ones during senescence. The proportion of N supplied for new leaf growth from the turnover of proteins in older leaves is generally considered to be between 60% and 100% in rice (Mae, 1986), although perhaps a little lower around 40% in ryegrass (Lattanzi et al., 2005). The remaining nutrients must be supplied by uptake from the roots. A recent review paper (Hikosaka, 2005a) suggests a method by which light intensity may indirectly affect the propor tion of nutrients under varying nutrient conditions. At high irradiances with high nutrient availability, growth is rapid and supported by both nutrient uptake and remobilization. At high irradiance, but low nutrient availabil ities, growth is rapid, but its nutrient demands are supported solely by remobilization, causing an increase in old leaf senescence rate. At low irradiances and high nutrient availabilities, the plants grow slowly, with the nutrient demand met by uptake, repressing or at least not promoting, old leaf senescence. Finally, at low irradiances and low nutrient availabil ities, growth is very slow, with the nutrient demand of the growing tissues being met by remobilization from older leaves and new assimilation being both energetically costly and unnecessary to support growth rates. Some evidence exists that plants growing under sufficient light but at low nutrients have a higher stomatal conductance, which is suggested to be a mechanism by which the plants can increase their nutrient capture (Cramer et al., 2008). It is unclear how much of this effect is purely due to the increased transpira tion rate, or and how much can be attributed to increases in photosynthesis fuelling increased membrane nutrient transporters in the root. Irrespectively, simply taking nutrient supply and growth rate into consideration, this could largely account for the behaviour that we see in plants grown under various nutrient regimes.
168
L. J. IRVING ET AL.
Shading of fully expanded leaves appeared to retard the rate of Rubisco degradation in fully expanded rice leaves (Hidema et al., 1991), suggesting a role for light in this process. This seems to run contrary to Hikosaka et al.’s (1996) work discussed below, which clearly showed that shading would tend to decrease the protein levels in vine leaves. There seem to be several possibilities for explaining Hidema et al.’s (1991) data: (i) plant response to light and shading is species-specific, with rice simply behaving in a different way to Ipomoea; (ii) light stress is an important causal factor for protein degradation in rice and (iii) unshaded leaves gained more C than shaded leaves, causing increased proteolytic rates. Unfortunately, it is not easy to decide between these possibilities although the third seems most likely. Interestingly, levels of other enzymes were also affected by the shading treatments. While NADP-G3PDH followed a similar pattern to Rubisco, with full light leaves exhibiting more rapid loss of protein than their shaded counterparts, cytochrome f contents were oppositely affected, with shaded leaves exhibiting a more rapid loss of protein than unshaded leaves. Chlor ophyll contents decreased more rapidly in unshaded plants. No difference in the proportion of N allocated to Rubisco was noted in the treatments, suggesting that the differences in leaf protein levels accurately reflected the total fluxes of N in the leaf. Deficiency of nutrients other than N can also alter the amount of Rubisco and presumably other proteins in plants. For example, S starva tion has been shown to lead to a large decrease in the amount of Rubisco, and a smaller decrease in the total protein levels, in rice leaves. An increase in light stress was also noted in S-deficient rice (Lunde et al., 2008). In Lemna minor fronds, S deficiency, but not the deficiency of other nutrients, including N, led to a large decrease in the Rubisco concentration (Ferreira and Teixeira, 1992).
V. IMPLICATIONS OF PROTEIN TURNOVER IN
WHOLE PLANTS
Protein turnover is understood to have an important role in recycling N and other nutrients within the plant in such a manner as to increase photosynthesis (Grindlay, 1997) and therein reproductive success. Some fraction of leaf N will be lost during senescence and remobilization, and that represents lost photosynthetic capacity; however, remobiliza tion of N to a new leaf can increase whole-plant photosynthesis, especially in dense canopies of homogenous plants, such as crops. Under N-limiting conditions, a leaf will senesce when the quantity of C it can
PROTEIN TURNOVER IN GRASS LEAVES
169
fix drops below the amount of C that could be fixed by that N in a new leaf, taking the C cost of N lost during senescence into account. Recent evidence has shown precisely this phenomenon in Xanthium canadense plants grown under low N conditions (Oikawa et al., 2006). This can lead to leaves senescing even when they are still able to maintain a positive photosynthetic rate. Under N-sufficient or N-excess conditions, however, such a strong requirement for N remobilization to maximize photosyn thetic gain is unnecessary, and indeed under N replete conditions plants seem to retain greater concentrations of N in senescent leaves than are necessary for the amount of photosynthesis performed by those leaves (Oikawa et al., 2008). In relation to C gain per unit N, plant canopies seem to pursue an optimal strategy, remobilizing N from lower, shaded leaves, to upper leaves where the light intensity is higher. There are two possible explanations for this, which I shall term deliberate optimization and inherent optimization. Deliberate optimization would be defined as the plant having some kind of mechanism by which it can, based on its internal conditions (N content, etc.), adjust and regulate leaf lifespan and N remobilization in such a way as to specifically and deliberately increase whole-plant photosynthesis. The mechanisms for this are obscure, but it is not implausible that such mechanisms could exist—we know from earlier sections that considerable plasticity exists in the regulation of protein synthesis and degradation. Inherent optimization suggests that optimization of canopy photosynthesis is simply an emergent property of the system, and that plants have limited capacity to adjust their leaf N contents to the ambient conditions after the termination of leaf expansion. Protein synthesis occurs during leaf expan sion, while degradation occurs throughout the leaves’ lifespan. Leaves deeper in the canopy (lower on the stem) are older than leaves nearer to the light, at least in grasses, and contain less N simply as a result of their age. This sets up a gradient in N concentration down through the canopy, approximately matching that for light. However, plants which have protein turnover rates allowing the older leaves to senescence and new leaves to grow in the pattern closely resembling the light gradient, thereby maximiz ing photosynthesis, will tend to outcompete plants with sub-optimal leaf properties and come to dominate the stand. Thus, creating the illusion of deliberate optimization of leaf N, as only the plants with optimal protein turnover rates for the environment will be present. Of course, as noted throughout the text, plant leaves certainly have some capacity to alter their rates of protein synthesis and degradation in respect to their environ mental conditions, but these can be explained largely as the changes of cellular behaviour based on their local conditions, rather than the
170
L. J. IRVING ET AL.
global changes to the whole plant. Indeed, in the vine Ipomoea tricolor grown horizontally to avoid shading, a developmental gradient in leaf N was noted in plants grown at low and intermediate N availabilities, with the youngest leaves having the highest N concentration, while the oldest leaves had the lowest concentrations. If the plants goal is the optimization of C fixation, then all other things being equal, N should have been distributed evenly among the leaves (Presland and McNaughton, 1984). On the other hand, shading of older leaves was able to exacerbate the N gradient evident at low N concentrations, while causing a gradient in older leaves grown at higher N availabilities. Stronger shading tended to produce stronger gradients, and inverse shading—where young leaves were shaded while old leaves were unshaded—produced an inverse N distribution, with old leaves behaving like young leaves and vice versa. Indeed, this clearly shows that both N and light can have strong effects on the N distribution between leaves, perhaps even more so than the devel opmental gradients. It is unclear whether other species share such plasticity with vines. A recent paper using a barley quantitative trait loci (QTL) mapping popu lation presented results indicating a positive relationship between the effi ciency of N remobilization, as characterized by the difference in N concentration between leaf maturity and leaf death, in the flag leaf and grain yield (Mickelson et al., 2003). Plants with large flag leaves were more efficient at N translocation, with leaf weight positively correlated with grain yield. The physiological basis for differences in N remobilization noted by Mickelson et al. (2003) are not currently understood, nor are the reasons for genotypic specificity. However, another paper from the same group looking at the effects of a QTL involved in determining seed N concentration found broadly similar results, with leaf N remobilization from leaves correlated with grain N levels (Jukanti and Fischer, 2008). Recent work has demonstrated a more causal link between leaf N turnover and plant growth (E Khaembah, PhD Thesis, Massey University, NZ). Leaf Rubisco content was quantified through time and modelled using the Irving and Robinson (2006) model for 135 ryegrass (Lolium perenne) genotypes in a QTL mapping population. The maximum Rubisco content correlated negatively with leaf length, tiller weight and plant dry weight. This suggests that plants which invest less N into Rubisco tend to have larger leaves, bigger tillers and a higher overall plant mass. Speculatively, plants investing less N into each leaf may be able to support more leaves, increasing light-use efficiency and total C fixed on a whole-plant basis in an N-limiting environment with low levels of competi tion. Where competition for light is higher, faster turnover rates and higher maximum Rubisco contents may be favoured. Negative relationships were
PROTEIN TURNOVER IN GRASS LEAVES
171
also noted between the maximum Rubisco content and the standard devia tion of the log-normal model and the peak time. Plants which had a high-peak Rubisco content reached that peak content early in the leaf’s lifespan, then exhibited a rapid decrease in the Rubisco content, while those which had lower, later peaks also had a more modest subsequent loss of Rubisco. Plant parasitism may have strong influences on the host-plant leaf con dition and on the nutritional status of the plant (Irving and Cameron, 2009). Parasites can be split broadly into two groups: phloem-feeding parasites and xylem-feeding parasites. Phloem-feeding parasites tend to be wholly dependent on their hosts for C (holoparasites), while xylem-feeding parasites tend to absorb water, hormones and nutrients, yet photosynthe size independently (hemiparasites). This was hypothesized to have differ ential effects on host tissues, and holoparasitized host plants often exhibit retarded leaf senescence (Hibberd et al., 1999), presumably because carbo hydrates do not accumulate in their leaves. Hemiparasites may be expected to promote leaf senescence, since they intercept N in the xylem before it reaches the host leaves. Indeed, parasitized Phleum plants had significantly less chlorophyll and a slight decrease in leaf Rubisco contents (Cameron et al., 2008).
VI. CONCLUSIONS N remobilization is hugely important for plant development and growth. In this chapter, we have attempted to give a broad overview of the processes of protein synthesis and degradation, and the factors which seem important in their determination. In grasses, protein synthesis mainly takes place during leaf expansion, with various factors important in reg ulating leaf protein concentration, specifically N or other nutrients and some hormones such as cytokinins responsible for an increase in leaf protein levels, while C and ABA downregulate synthesis. Light appears to be instrumental in the synthesis of many leaf proteins. The integrated and co-ordinated behaviour of the chloroplast and the nuclear genomes is important, and the mechanisms by which it is affected remain incompletely understood, although progress is being made. Protein degradation is a complex process, with multiple degradation pathways posited for many proteins. The loss of stromal proteins seems to be at least partly a result of vesicular export from the chloroplast, although the mechanism of this export remains deeply contentious. Similarly, the mechanism of degradation of the D1 protein remains contentious, as we have shown here. Reactive oxygen species seem important in D1 degradation and indeed can also cause the
172
L. J. IRVING ET AL.
degradation of Rubisco. While it is unclear how much of a role that reactive oxygen species have in natural senescence, it is clear that they certainly have the potential to bleach the cells in carotenoid-less plants, whether this is a result of a mutant gene (immutans Arabidopsis) or whether it is a result of the application of carotenoid-suppressing chemicals. The relative amounts of protein synthesized and degraded throughout the lifespan of the leaves determine the protein contents of the leaves. It is well documented that plants exhibit an apparently optimized N distribution in their canopy, although the mechanisms by which these are affected remain unknown. Studies using parasitized plants suggest that the behaviours noted in the plants can be explained by shifts in the C and N levels within the plants. Finally, several recent studies have shown evidence of a strong link between the rates of N remobilization within the plant and its productivity, a con clusion which reinforces the need for continued work in understanding the processes controlling plant N levels.
ACKNOWLEDGEMENTS LJ Irving would like to thank Shinya Wada and Wataru Yamori for many helpful discussions during the course of writing this chapter. LJ Irving is supported by a Japan Society for the Promotion of Science postdoctoral fellowship (Award number: PO8097).
REFERENCES Acevedo-Hernandez, G.J., Leon, P., Herrera-Estrella, L.R., 2005. Sugar and ABA responsiveness of a minimal RBCS light-responsive unit is mediated by direct binding of ABI4. Plant J. 43(4): 506–519. Acharya, B., Assmann, S., 2009. Hormone interactions in stomatal function. Plant Mol. Biol. 69(4): 451–462. Addiscott, T.M., Whitmore, A.P., Powlson, D.S., 1991. Farming, Fertilizers and the Nitrate Problem, C.A.B. International, Oxon. Aerts, R., 1989. The effect of increased nutrient availability on leaf turnover and above-ground productivity of 2 evergreen ericaceous shrubs. Oecologia 78(1): 115–120. Aerts, R., Decaluwe, H., 1995. Interspecific and intraspecific differences in shoot and leaf lifspan of 4 Carex species which differ in maximum dry-matter produc tion. Oecologia 102(4): 467–477. Albert, N.W., Lewis, D.H., Zhang, H., Irving, L.J., Jameson, P.E., Davies, K.M., 2009. Light-induced vegetative anthocyanin pigmentation in Petunia. J. Exp. Bot. 60(7): 2191–2202. Aluru, M.R., Yu, F., Fu, A.G., Rodermel, S., 2006. Arabidopsis variegation mutants: new insights into chloroplast biogenesis. J. Exp. Bot. 57(9): 1871–1881.
PROTEIN TURNOVER IN GRASS LEAVES
173
Andersson, B., Aro, E.M., 1997. Proteolytic activities and proteases of plant chlor oplasts. Physiol. Plant. 100(4): 780–793. Araya, T., Noguchi, K., Terashima, I., 2006. Effects of carbohydrate accumulation on photosynthesis differ between sink and source leaves of Phaseolus vul garis L. Plant Cell Physiol. 47(5): 644–652. Arimura, S., Hirai, A., Tsutsumi, N., 2001. Numerous and highly developed tubular projections from plastids observed in Tobacco epidermal cells. Plant Sci. 160(3): 449–454. Asada, K., 2006. Production and scavenging of reactive oxygen species in chloro plasts and their functions. Plant Physiol. 141(2): 391–396. Baek, K.H., Choi, D., 2008. Roles of plant proteases in pathogen defense. Plant Pathol. J. 24(4): 367–374. Bassham, D.C., 2009. Function and regulation of macroautophagy in plants. Bio chim. Biophys. Acta 1793(9): 1397–1403. Bauer, D., Biehler, K., Fock, H., Carrayol, E., Hirel, B., Migge, A., et al., 1997. A role for cytosolic glutamine synthetase in the remobilization of leaf nitrogen during water stress in tomato. Physiol. Plant. 99(2): 241–248. Baumgartner, B.J., Rapp, J.C., Mullet, J.E., 1989. Plastid transcription activity and DNA copy number increase early in barley chloroplast development. Plant Physiol. 89(3): 1011–1018. Bausenwein, U., Millard, P., Raven, J.A., 2001. Remobilized old-leaf nitrogen pre dominates for spring growth in two temperate grasses. New Phytol. 152(2): 283–290. Buchanan-Wollaston, V., Earl, S., Harrison, E., Mathas, E., Navabpour, S., Page, T., et al., 2003. The molecular analysis of leaf senescence – a genomics approach. Plant Biotechnol. J. 1(1): 3–22. Buchanan-Wollaston, V., Page, T., Harrison, E., Breeze, E., Lim, P.O., Nam, H.G., et al., 2005. Comparative transcriptome analysis reveals significant differ ences in gene expression and signalling pathways between developmental and dark/starvation-induced senescence in Arabidopsis. Plant J. 42(4): 567–585. Callis, J., 1995. Regulation of protein degradation. Plant Cell 7(7): 845–857. Cameron, D.D., Geniez, J.M., Seel, W.E., Irving, L.J., 2008. Suppression of host photosynthesis by the parasitic plant Rhinanthus minor. Ann. Bot. 101(4): 573–578. Castonguay, Y., Nadeau, P., Simard, R.R., 1993. Effects of flooding on carbohydrate and Aba levels in roots and shoots of Alfalfa. Plant Cell Environ. 16(6): 695–702. Causin, H.F., Roberts, I.N., Criado, M.V., Gallego, S.M., Pena, L.B., Rios, M.D., et al., 2009. Changes in hydrogen peroxide homeostasis and cytokinin levels contribute to the regulation of shade-induced senescence in wheat leaves. Plant Sci. 177(6): 698–704. Chen, W.Q., Provart, N.J., Glazebrook, J., Katagiri, F., Chang, H.S., Eulgem, T., et al., 2002. Expression profile matrix of Arabidopsis transcription factor genes suggests their putative functions in response to environmental stres ses. Plant Cell 14(3): 559–574. Cheng, S.H., Moore, B.D., Seemann, J.R., 1998. Effects of short- and long-term elevated CO2 on the expression of ribulose-1,5-bisphosphate carboxylase/ oxygenase genes and carbohydrate accumulation in leaves of Arabidopsis thaliana (L) Heynh. Plant Physiol. 116(2): 715–723. Chiba, A., Ishida, H., Nishizawa, N.K., Makino, A., Mae, T., 2003. Exclusion of ribulose-1,5-bisphosphate carboxylase/oxygenase from chloroplasts by spe cific bodies in naturally senescing leaves of wheat. Plant Cell Physiol. 44(9): 914–921.
174
L. J. IRVING ET AL.
Choquet, Y., Vallon, O., 2000. Synthesis, assembly and degradation of thylakoid membrane proteins. Biochimie 82(6–7): 615–634. Choquet, Y., Zito, F., Wostrikoff, K., Wollman, F.A., 2003. Cytochrome f transla tion in chlamydomonas chloroplast is autoregulated by its carboxylterminal domain. Plant Cell 15(6): 1443–1454. Chung, T., Suttangkakul, A., Vierstra, R.D., 2009. The ATG autophagic conjugation system in maize: ATG transcripts and abundance of the ATG8-lipid adduct are regulated by development and nutrient availability. Plant Physiol. 149(1): 220–234. Crafts-Brandner, S.J., Holzer, R., Feller, U., 1998. Influence of nitrogen deficiency on senescence and the amounts of RNA and proteins in wheat leaves. Physiol. Plant. 102(2): 192–200. Cramer, M.D., Hoffmann, V., Verboom, G.A., 2008. Nutrient availability moderates transpiration in Ehrharta calycina. New Phytol. 179: 1048–1057. Criado, M.V., Caputo, C., Roberts, I.N., Castro, M.A., Barneix, A.J., 2009. Cytokinin-induced changes of nitrogen remobilization and chloroplast ultra structure in wheat (Triticum aestivum). J. Plant Physiol. 166(16): 1775–1785. Demartino, G.N., Slaughter, C.A., 1993. Regulatory proteins of the proteasome. Enzyme Protein 47(4–6): 314–324. Desimone, M., Henke, A., Wagner, E., 1996. Oxidative stress induces partial degra dation of the large subunit of ribulose-1,5-bisphosphate carboxylase/oxyge nase in isolated chloroplasts of barley. Plant Physiol. 111(3): 789–796. Drager, R.G., Girard-Bascou, J., Choquet, Y., Kindle, K.L., Stern, D.B., 1998. In vivo evidence for 50 !30 exoribonuclease degradation of an unstable chlor oplast mRNA. Plant J. 13(1): 85–96. Dubell, A.N., Mullet, J.E., 1995. Differential transcription of pea chloroplast genes during light-induced leaf development – continuous far-red light activates chloroplast transcription. Plant Physiol. 109(1): 105–112. Edelman, M., Mattoo, A.K., 2008. D1-protein dynamics in photosystem II: the lingering enigma. Photosyn. Res. 98(1–3): 609–620. Ellis, R.J., 1979. The most abundant protein in the world. Trends Biochem. Sci. 4: 241–244. Ferreira, R.M.B., Teixeira, A.R.N., 1992. Sulfur starvation in Lemna leads to degradation of ribulose bisphosphate carboxylase without plant death. J. Biol. Chem. 267(11): 7253–7257. Finley, D., Ozkaynak, E., Varshavsky, A., 1987. The yeast polyubiquitin gene is essential for resistance to high temperatures, starvation, and other stresses. Cell 48(6): 1035–1046. Friedrich, J.W., Huffaker, R.C., 1980. Photosynthesis, leaf resistances, and ribulose1,5-bisphosphate carboxylase degradation in senescing barley leaves. Plant Physiol. 65: 1103–1107. Gastal, F., Nelson, C.J., 1994. Nitrogen use within the growing leaf blade of tall fescue. Plant Physiol. 105(1): 191–197. Gesch, R.W., Boote, K.J., Vu, J.C.V., Allen, L.H., Bowes, G., 1998. Changes in growth CO2 result in rapid adjustments of ribulose-1,5-bisphosphate car boxylase/oxygenase small subunit gene expression in expanding and mature leaves of rice. Plant Physiol. 118(2): 521–529. Gesch, R.W., Kang, I.H., Gallo-Meagher, M., Vu, J.C.V., Boote, K.J., Allen, L.H., et al., 2003. Rubisco expression in rice leaves is related to genotypic varia tion of photosynthesis under elevated growth CO2 and temperature. Plant Cell Environ. 26(12): 1941–1950. Ghislain, M., Udvardy, A., Mann, C., 1993. Saccharomyces cerevisiae 26S protease mutants arrest cell division in G2/metaphase. Nature. 366(6453): 358–362.
PROTEIN TURNOVER IN GRASS LEAVES
175
Glick, R.E., Schlagnhaufer, C.D., Arteca, R.N., Pell, E.J., 1995. Ozone induced ethylene emission accelerates the loss of Ribulose-1,5-bisphosphate carbox ylase/oxygenase and nuclear encoded mRNAs in senescing potato leaves. Plant Physiol. 109(3): 891–898. Gordon, C., McGurk, G., Dillon, P., Rosen, C., Hastie, N.D., 1993. Defective mitosis due to a mutation in the gene for a fission yeast 26S protease subunit. Nature 366(6453): 355–357. Grbic, V., Bleecker, A.B., 1995. Ethylene regulates the timing of leaf senescence in Arabidopsis. Plant J. 8(4): 595–602. Grindlay, D.J.C., 1997. Towards an explanation of crop nitrogen demand based on the optimization of leaf nitrogen per unit leaf area. J. Agric. Sci. 128: 377–396. Gunning, B.E.S., 2005. Plastid stromules: video microscopy of their outgrowth, retraction, tensioning, anchoring, branching, bridging, and tip-shedding. Protoplasma 225(1–2): 33–42. Hanaoka, H., Noda, T., Shirano, Y., Kato, T., Hayashi, H., Shibata, D., et al., 2002. Leaf senescence and starvation-induced chlorosis are accelerated by the dis ruption of an Arabidopsis autophagy gene. Plant Physiol. 129(3): 1181–1193. He, P., Osaki, M., Takebe, M., Shinano, T., Wasaki, J., 2005. Endogenous hormones and expression of senescence-related genes in different senescent types of maize. J. Exp. Bot. 56(414): 1117–1128. He, Y.H., Fukushige, H., Hildebrand, D.F., Gan, S.S., 2002. Evidence supporting a role of jasmonic acid in Arabidopsis leaf senescence. Plant Physiol. 128(3): 876–884. He, Y.H., Tang, W.N., Swain, J.D., Green, A.L., Jack, T.P., Gan, S.S., 2001. Networking senescence-regulating pathways by using Arabidopsis enhancer trap lines. Plant Physiol. 126(2): 707–716. Hibberd, J.M., Quick, W.P., Press, M.C., Scholes, J.D., Jeschke, W.D., 1999. Solute fluxes from tobacco to the parasitic angiosperm Orobanche cernua and the influence of infection on host carbon and nitrogen relations. Plant Cell Environ. 22(8): 937–947. Hidema, J., Makino, A., Mae, T., Ojima, K., 1991. Photosynthetic characteristics of rice leaves aged under different irradiances from full expansion through senescence. Plant Physiol. 97(4): 1287–1293. Highfield, P.E., Ellis, R.J., 1978. Synthesis and transport of small subunit of chlor oplast ribulose bisphosphate carboxylase. Nature 271(5644): 420–424. Hikosaka, K., 1996. Effects of leaf age, nitrogen nutrition and photon flux density on the organization of the photosynthetic apparatus in leaves of a vine (Ipo moea tricolor Cav) grown horizontally to avoid mutual shading of leaves. Planta 198(1): 144–150. Hikosaka, K., 2005a. Leaf canopy as a dynamic system: Ecophysiology and optim ality in leaf turnover. Ann. Bot. 95(3): 521–533. Hikosaka, K., 2005b. Nitrogen partitioning in the photosynthetic apparatus of Plantago asiatica leaves grown under different temperature and light con ditions: similarities and differences between temperature and light acclima tion. Plant Cell Physiol. 46(8): 1283–1290. Hikosaka, K., Terashima, I., 1996. Nitrogen partitioning among photosynthetic compo nents and its consequence in sun and shade plants. Funct. Ecol. 10(3): 335–343. Holzinger, A., Buchner, O., Lutz, C., Hanson, M.R., 2007. Temperature-sensitive formation of chloroplast protrusions and stromules in mesophyll cells of Arabidopsis thaliana. Protoplasma 230(1–2): 23–30. Huang, F.Y., PhilosophHadas, S., Meir, S., Callaham, D.A., Sabato, R., Zelcer, A., et al., 1997. Increases in cytosolic Ca2þ in parsley mesophyll cells correlate with leaf senescence. Plant Physiol. 115(1): 51–60.
176
L. J. IRVING ET AL.
Hudson, G.S., Evans, J.R., von Caemmerer, S., Arvidsson, Y.B.C., Andrews, T.J., 1992. Reduction of ribulose-1,5-bisphosphate carboxylase/oxygenase con tent by antisense RNA reduces photosynthesis in transgenic tobacco plants. Plant Physiol. 98: 294–302. Huffaker, R.C., 1990. Proteolytic activity during senescence of plants. New Phytol. 116(2): 199–231. Huq, E., 2006. Degradation of negative regulators: a common theme in hormone and light signaling networks? Trends Plant Sci. 11(1): 4–7. Huq, E., Al-Sady, B., Hudson, M., Kim, C.H., Apel, M., Quail, P.H., 2004. PHYTOCHROME-INTERACTING FACTOR 1 is a critical bHLH reg ulator of chlorophyll biosynthesis. Science 305(5692): 1937–1941. Imai, K., Suzuki, Y., Mae, T., Makino, A., 2008. Changes in the synthesis of rubisco in rice leaves in relation to senescence and N influx. Ann. Bot. 101(1): 135–144. Imai, K., Suzuki, Y., Makino, A., Mae, T., 2005. Effects of nitrogen nutrition on the relationships between the levels of rbcS and rbcL mRNAs and the amount of ribulose 1 center dot 5-bisphosphate carboxylase/oxygenase synthesized in the eighth leaves of rice from emergence through senescence. Plant Cell Environ. 28(12): 1589–1600. Inada, N., Sakai, A., Kuroiwa, H., Kuroiwa, T., 1998a. Three-dimensional analysis of the senescence program in rice (Oryza sativa L.) coleoptiles – Investiga tions by fluorescence microscopy and electron microscopy. Planta 206(4): 585–597. Inada, N., Sakai, A., Kuroiwa, H., Kuroiwa, T., 1998b. Three-dimensional analysis of the senescence program in rice (Oryza sativa L.) coleoptiles – Investigations of tissues and cells by fluorescence microscopy. Planta 205(2): 153–164. Inada, N., Sakai, A., Kuroiwa, H., Kuroiwa, T., 1999. Senescence program in rice (Oryza sativa L.) leaves: analysis of the blade of the second leaf at the tissue and cellular levels. Protoplasma 207(3–4): 222–232. Inoue, Y., Suzuki, T., Hattori, M., Yoshimoto, K., Ohsumi, Y., Moriyasu, Y., 2006. AtATG genes, homologs of yeast autophagy genes, are involved in consti tutive autophagy in Arabidopsis root tip cells. Plant Cell Physiol. 47(12): 1641–1652. Irving, L.J., Cameron, D.D., 2009. You are what you eat: interactions between root parasitic plants and their hosts. Adv. Bot. Res. 50, 87–138. Irving, L.J., Robinson, D., 2006. A dynamic model of Rubisco turnover in cereal leaves. New Phytol. 169(3): 493–504. Irving, L.J., Sheng, Y.B., Woolley, D., Matthew, C., 2007. Physiological effects of waterlogging on two lucerne varieties grown under glasshouse conditions. J. Agron. Crop Sci. 193(5): 345–356. Ishida, H., Nishimori, Y., Sugisawa, M., Makino, A., Mae, T., 1997. The large subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase is fragmented into 37 kDa and 16-kDa polypeptides by active oxygen in the lysates of chloroplasts from primary leaves of wheat. Plant Cell Physiol. 38(4): 471–479. Ishida, H., Shimizu, S., Makino, A., Mae, T., 1998. Light-dependent fragmentation of the large subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase in chloroplasts isolated from wheat leaves. Planta 204(3): 305–309. Ishida, H., Yoshimoto, K., Izumi, M., Reisen, D., Yano, Y., Makino, A., et al., 2008. Mobilization of rubisco and stroma-localized fluorescent proteins of chlor oplasts to the vacuole by an ATG gene-dependent autophagic process. Plant Physiol. 148(1): 142–155. Ishizuka, M., Makino, A., Suzuki, Y., Mae, T., 2004. Amount of ribulose-1,5 bisphosphate carboxylase/oxygenase (Rubisco) protein and levels of mRNAs
PROTEIN TURNOVER IN GRASS LEAVES
177
of rbcS and rbcL in the leaves at different positions in transgenic rice plants with decreased content of Rubisco. Soil Sci. Plant Nutr. 50(2): 233–239. Jukanti, A.K., Fischer, A.M., 2008. A high-grain protein content locus on barley (Hordeum vulgare) chromosome 6 is associated with increased flag leaf proteolysis and nitrogen remobilization. Physiol. Plant. 132(4): 426–439. Kato, Y., Miura, E., Ido, K., Ifuku, K., Sakamoto, W., 2009. The variegated mutants lacking chloroplastic FtsHs are defective in D1 degradation and accumulate reactive oxygen species. Plant Physiol. 151(4): 1790–1801. Kato, Y., Murakami, S., Yamamoto, Y., Chatani, H., Kondo, Y., Nakano, T., et al., 2004. The DNA-binding protease, CND41, and the degradation of ribulose-1,5-bisphosphate carboxylase/oxygenase in senescent leaves of tobacco. Planta 220(1): 97–104. Katz, C., Rasouly, A., Gur, E., Shenhar, Y., Biran, D., Ron, E.Z., 2009. Temperature-dependent proteolysis as a control element in Escherichia coli metabolism. Res. Microbiol. 160(9): 684–686. Kettunen, R., Tyystjarvi, E., Aro, E.M., 1991. D1 protein degradation during photoinhibition of intact leaves – a modification of the D1 protein precedes degradation. FEBS Lett. 290(1–2): 153–156. Khan, S., Andralojc, P.J., Lea, P.J., Parry, M.A.J., 1999. 20 -Carboxy-D-arabitinol 1-phosphate protects ribulose 1,5- bisphosphate carboxylase/oxygenase against proteolytic breakdown. Eur. J. Biochem. 266(3): 840–847. Koizumi, M., Yamaguchishinozaki, K., Tsuji, H., Shinozaki, K., 1993. Structure and expression of 2 genes that encode distinct drought-inducible cysteine pro teinasesin Arabidopsis thaliana. Gene 129(2): 175–182. Kokubun, N., Ishida, H., Makino, A., Mae, T., 2002. The degradation of the large subunit of ribulose-1,5- bisphosphate carboxylase/oxygenase into the 44 kDa fragment in the lysates of chloroplasts incubated in darkness. Plant Cell Physiol. 43(11): 1390–1395. Krapp, A., Hofmann, B., Schafer, C., Stitt, M., 1993. Regulation of the expression of Rbcs and other photosynthetic genes by carbohydrates – a mechanism for the sink regulation of photosynthesis. Plant J. 3(6): 817–828. Krapp, A., Quick, W.P., Stitt, M., 1991. Ribulose-1,5-bisphosphate carboxylase/ oxygenase, other Calvin-cycle enzymes, and chlorophyll decrease when glucose is supplied to mature spinach leaves via the transpiration stream. Planta 186(1): 58–69. Kruger, E.L., Volin, J.C., 2006. Reexamining the empirical relation between plant growth and leaf photosynthesis. Funct. Plant Biol. 33(5): 421–429. La Rocca, N., Barbato, R., Bonora, A., Valle, L.D., De Faveri, S., Rascio, N., 2004. Thylakoid dismantling of damaged unfunctional chloroplasts modulates the Cab and RbcS gene expression in wheat leaves. J. Photochem. Photo biol. B, Biol. 73(3): 159–166. Laisk, A., Oja, V., Kiirats, O., Raschke, K., Heber, U., 1989. The state of the photosynthetic apparatus in leaves as analyzed by rapid gas exchange and optical methods – the pH of the chloroplast stroma and activation of enzymes in vivo. Planta 177(3): 350–358. Lattanzi, F.A., Schnyder, H., Thornton, B., 2005. The sources of carbon and nitro gen supplying leaf growth. Assessment of the role of stores with compart mental models. Plant Physiol. 137(1): 383–395. Lawlor, D.W., 2002. Carbon and nitrogen assimilation in relation to yield: mechanisms are the key to understanding production systems. J. Exp. Bot. 53(370): 773–787. Lopez-Juez, E., Bowyer, J.R., Sakai, T., 2007. Distinct leaf developmental and gene expression responses to light quantity depend on blue-photoreceptor or
178
L. J. IRVING ET AL.
plastid-derived signals, and can occur in the absence of phototropins. Planta 227(1): 113–123. Lunde, C., Zygadlo, A., Simonsen, H.T., Nielsen, P.L., Blennow, A., Haldrup, A., 2008. Sulfur starvation in rice: the effect on photosynthesis, carbohydrate metabolism, and oxidative stress protective pathways. Physiol. Plant. 134(3): 508–521. Lysenko, E.A., 2007. Plant sigma factors and their role in plastid transcription. Plant Cell Rep. 26(7): 845–859. Mae, T., 1986. Partitioning and utilization of nitrogen in rice plants. Jpn. Agric. Res. Q. 20(2): 115–120. Mae, T., Kai, N., Makino, A., Ohira, K., 1984. Relationship between ribulose bisphosphate carboxylase content and chloroplast number in naturally senescing primary leaves of wheat. Plant Cell Physiol. 25(2): 333–336. Mae, T., Kamei, C., Funaki, K., Miyadai, K., Makino, A., Ohira, K., et al., 1989. Degradation of Ribulose-1,5-bisphosphate carboxylase/oxygenase in the lysates of the chloroplasts isolated mechanically from wheat (Triticum aestivum L.) leaves. Plant Cell Physiol. 30(2): 193–200. Mae, T., Makino, A., Ohira, K., 1983. Changes in the amounts of ribulose bispho sphate carboxylase synthesized and degraded during the life span of rice leaf (Oryza sativa L.). Plant Cell Physiol. 24(6): 1079–1086. Mae, T., Ohira, K., 1981. The remobilisation of nitrogen related to leaf growth and senescence in rice plants (Oryza sativa L.). Plant Cell Physiol. 22(6): 1067–1074. Makino, A., Mae, T., Ohira, K., 1983. Photosynthesis and ribulose-1,5-bisphosphate carboxylase in rice leaves – changes in photosynthesis and enzymes involved in carbon assimilation from leaf development through senescence. Plant Physiol. 73(4): 1002–1007. Makino, A., Mae, T., Ohira, K., 1984a. Effect of nitrogen, phosphorus or potassium on the photosynthetic rate and ribulose-1,5-bisphosphate carboxylate con tent in rice leaves during expansion. Soil Sci. Plant Nutr. 30(1): 63–70. Makino, A., Mae, T., Ohira, K., 1984b. Relation between nitrogen and ribulose-1,5 bisphosphate carboxylase in rice leaves from emergence through senescence. Plant Cell Physiol. 25(3): 429–437. Makino, A., Osmond, B., 1991. Effects of nitrogen nutrition on nitrogen partitioning between chloroplasts and mitochondria in pea and wheat. Plant Physiol. 96 (2): 355–362. Makino, A., Sato, T., Nakano, H., Mae, T., 1997a. Leaf photosynthesis, plant growth and nitrogen allocation in rice under different irradiances. Planta 203(3): 390–398. Makino, A., Shimada, T., Takumi, S., Kaneko, K., Matsuoka, M., Shimamoto, K., et al., 1997b. Does decrease in ribulose-1,5-bisphosphate carboxylase by antisense RbcS lead to a higher N-use efficiency of photosynthesis under conditions of saturating CO2 and light in rice plants? Plant Physiol. 114(2): 483–491. Marin-Navarro, J., Moreno, J., 2006. Cysteines 449 and 459 modulate the reductionoxidation conformational changes of ribulose 1.5-bisphosphate carboxy lase/oxygenase and the translocation of the enzyme to membranes during stress. Plant Cell Environ. 29(5): 898–908. Martinez, D.E., Costa, M.L., Gomez, F.M., Otegui, M.S., Guiamet, J.J., 2008. “Senescence-associated vacuoles” are involved in the degradation of chlor oplast proteins in tobacco leaves. Plant J. 56(2): 196–206. Masclaux-Daubresse, C., Purdy, S., Lemaitre, T., Pourtau, N., Taconnat, L., Renou, J.P., et al., 2007. Genetic variation suggests interaction between cold
PROTEIN TURNOVER IN GRASS LEAVES
179
acclimation and metabolic regulation of leaf senescence. Plant Physiol. 143(1): 434–446. Matt, P., Krapp, A., Haake, V., Mock, H.P., Stitt, M., 2002. Decreased Rubisco activity leads to dramatic changes of nitrate metabolism, amino acid meta bolism and the levels of phenylpropanoids and nicotine in tobacco antisense RbcS transformants. Plant J. 30(6): 663–677. McRae, D.G., Thompson, J.E., 1983. Senescence dependent changes in superoxide anion production by illuminated chloroplasts from bean leaves. Planta 158 (3): 185–193. Mickelson, S., See, D., Meyer, F.D., Garner, J.P., Foster, C.R., Blake, T.K., et al., 2003. Mapping of QTL associated with nitrogen storage and remo bilization in barley (Hordeum vulgare L.) leaves. J. Exp. Bot. 54(383): 801–812. Miller, B.L., Huffaker, R.C., 1982. Hydrolysis of ribulose-1,5-bisphosphate carbox ylase by endoproteinases from senescing barley leaves. Plant Physiol. 69(1): 58–62. Minamikawa, T., Toyooka, K., Okamoto, T., Hara-Nishimura, I., Nishimura, M., 2001. Degradation of ribulose-bisphosphate carboxylase by vacuolar enzymes of senescing French bean leaves: immunocytochemical and ultra structural observations. Protoplasma 218(3–4): 144–153. Mishkind, M.L., Schmidt, G.W., 1983. Post-transcriptional regulation of ribulose1,5-bisphosphate carboxylase small subunit accumulation in Chlamydomo nas reinhardtii. Plant Physiol. 72(3): 847–854. Moon, J., Parry, G., Estelle, M., 2004. The ubiquitin-proteasome pathway and plant development. Plant Cell 16(12): 3181–3195. Nakajima, Y., Yoshida, S., Inoue, Y., Ono, T., 1996. Occupation of the Q(B)-binding pocket by a photosystem II inhibitor triggers dark cleavage of the D1 protein subjected to brief preillumination. J. Biol. Chem. 271(29): 17383–17389. Nakajima, Y., Yoshida, S., Inoue, Y., Yoneyama, K., Ono, T.A., 1995. Selective and specific degradation of the D1 protein induced by binding of a novel photosystem II inhibitor to the Q(B) site. Biochim. Biophys. Acta 1230 (1–2): 38–44. Obara, K., Kuriyama, H., Fukuda, H., 2001. Direct evidence of active and rapid nuclear degradation triggered by vacuole rupture during programmed cell death in Zinnia. Plant Physiol. 125(2): 615–626. Oikawa, S., Hikosaka, K., Hirose, T., 2006. Leaf lifespan and lifetime carbon balance of individual leaves in a stand of an annual herb, Xanthium canadense. New Phytol. 172(1): 104–116. Oikawa, S., Hikosaka, K., Hirose, T., 2008. Does leaf shedding increase the wholeplant carbon gain despite some nitrogen being lost with shedding? New Phytol. 178(3): 617–624. Oja, V., Savchenko, G., Jakob, B., Heber, U., 1999. pH and buffer capacities of apoplastic and cytoplasmic cell compartments in leaves. Planta 209(2): 239–249. Oldenburg, D.J., Rowan, B.A., Zhao, L., Watcher, C.L., Schleh, M., Bendich, A.J., 2006. Loss or retention of chloroplast DNA in maize seedlings is affected by both light and genotype. Planta 225(1): 41–55. Ono, K., Hashimoto, H., Katoh, S., 1995. Changes in the number and size of chloroplasts during senescence of primary leaves of wheat grown under different conditions. Plant Cell Physiol. 36(1): 9–17. Otegui, M.S., Noh, Y.S., Martinez, D.E., Vila Petroff, M.G., Andrew Staehelin, L., Amasino, R.M., et al., 2005. Senescence-associated vacuoles with intense
180
L. J. IRVING ET AL.
proteolytic activity develop in leaves of Arabidopsis and soybean. Plant J. 41(6): 831–844. Parry, M.A.J., Keys, A.J., Madgwick, P.J., Carmo-Silva, A.E., Andralojc, P.J., 2008. Rubisco regulation: a role for inhibitors. J. Exp. Bot. 59(7): 1569–1580. Presland, M.R., McNaughton, G.S., 1984. Whole plant studies using radioactive 13N.2. A compartmental model for the uptake and transport of nitrate ions by Zea mays. J. Exp. Bot. 35(158): 1277–1288. Prins, A., Van Heerden, P.D.R., Olmos, E., Kunert, K.J., Foyer, C.H., 2008. Cysteine proteinases regulate chloroplast protein content and composition in tobacco leaves: a model for dynamic interactions with ribulose-1,5 bisphosphate carboxylase/oxygenase (Rubisco) vesicular bodies. J. Exp. Bot. 59(7): 1935–1950. Quick, W.P., Fichtner, K., Schulze, E.D., Wendler, R., Leegood, R.C., Mooney, H., et al., 1992. Decreased ribulose-1,5-bisphosphate carboxylase oxygenase in transgenic tobacco transformed with antisense RbcS.4. Impact on photo synthesis in conditions of altered nitrogen supply. Planta 188(4): 522–531. Rivero, R.M., Kojima, M., Gepstein, A., Sakakibara, H., Mittler, R., Gepstein, S., et al., 2007. Delayed leaf senescence induces extreme drought tolerance in a flowering plant. Proc. Natl. Acad. Sci. U.S.A. 104(49): 19631–19636. Rodermel, S., Haley, J., Jiang, C.Z., Tsai, C.H., Bogorad, L., 1996. A mechanism for intergenomic integration: Abundance of ribulose bisphosphate carboxylase small-subunit protein influences the translation of the large-subunit mRNA. Proc. Natl. Acad. Sci. U.S.A. 93(9): 3881–3885. Sagan, L., 1967. On origin of mitosing cells. J. Theor. Biol. 14(3): 225. Sakakibara, H., Suzuki, M., Takei, K., Deji, A., Taniguchi, M., Sugiyama, T., 1998. A response-regulator homologue possibly involved in nitrogen signal trans duction mediated by cytokinin in maize. Plant J. 14(3): 337–344. Sawbridge, T.I., Lopezjuez, E., Knight, M.R., Jenkins, G.I., 1994. A blue-light photoreceptor mediates the fluence rate dependent expression of genes encoding the small subunit of ribulose-1,5-bisphosphate carboxylase/oxy genase in light grown Phaseolus vulgaris primary leaves. Planta 192(1): 1–8. Schaffer, M.A., Fischer, R.L., 1988. Analysis of messenger RNAs that accumulate in response to low-temperature identifies a thiol protease gene in tomato. Plant Physiol. 87(2): 431–436. Scheible, W.R., Morcuende, R., Czechowski, T., Fritz, C., Osuna, D., PalaciosRojas, N., et al., 2004. Genome-wide reprogramming of primary and sec ondary metabolism, protein synthesis, cellular growth processes, and the regulatory infrastructure of Arabidopsis in response to nitrogen. Plant Physiol. 136(1): 2483–2499. Sergiev, I., Todorova, D., Somleva, M., Alexieva, V., Karanov, E., Stanoeva, E., et al., 2007. Influence of cytokinins and novel cytokinin antagonists on the senescence of detached leaves of Arabidopsis thaliana. Biol. Plant. 51(2): 377–380. Shaver, J.M., Oldenburg, D.J., Bendich, A.J., 2008. The structure of chloroplast DNA molecules and the effects of light on the amount of chloroplast DNA during development in Medicago truncatula. Plant Physiol. 146(3): 1064–1074. Sheen, J., 1990. Metabolic repression of transcription in higher plants. Plant Cell 2 (10): 1027–1038. Smalle, J., Vierstra, R.D., 2004. The ubiquitin 26S proteasome proteolytic pathway. Annu. Rev. Plant Biol. 55: 555–590. Spalding, E.P., Folta, K.M., 2005. Illuminating topics in plant photobiology. Plant Cell Environ. 28(1): 39–53.
PROTEIN TURNOVER IN GRASS LEAVES
181
Suzuki, Y., Makino, A., Mae, T., 2001. Changes in the turnover of Rubisco and levels of mRNAs of rbcL and rbcS in rice leaves from emergence to senescence. Plant Cell Environ. 24(12): 1353–1360. Suzuki, Y., Ohkubo, M., Hatakeyama, H., Ohashi, K., Yoshizawa, R., Kojima, S., et al., 2007. Increased Rubisco content in transgenic rice transformed with the “sense” rbcS gene. Plant Cell Physiol. 48(4): 626–637. Takahashi, S., Murata, N., 2008. How do environmental stresses accelerate photoinhibition? Trends Plant Sci. 13(4): 178–182. Takeuchi, A., Yamaguchi, T., Hidema, J., Strid, A., Kumagai, T., 2002. Changes in synthesis and degradation of Rubisco and LHCII with leaf age in rice (Oryza sativa L.) growing under supplementary UV-B radiation. Plant Cell Environ. 25(6): 695–706. Telfer, A., 2005. Too much light? How beta-carotene protects the photosystem II reaction centre. Photochem. Photobiol. Sci. 4(12): 950–956. To, J.P.C., Reiter, W.D., Gibson, S.I., 2003. Chloroplast biogenesis by Arabidopsis seedlings is impaired in the presence of exogenous glucose. Physiol. Plant. 118(3): 456–463. Tsukada, M., Ohsumi, Y., 1993. Isolation and characterization of autophagy defec tive mutants of Saccharomyces cerevisiae. FEBS Lett. 333(1–2): 169–174. Van der Graaff, E., Schwacke, R., Schneider, A., Desimone, M., Flugge, U.I., Kunze, R., 2006. Transcription analysis of Arabidopsis membrane transporters and hormone pathways during developmental and induced leaf senescence. Plant Physiol. 141(2): 776–792. Vu, J.C.V., Allen, L.H., Boote, K.J., Bowes, G., 1997. Effects of elevated CO2 and temperature on photosynthesis and Rubisco in rice and soybean. Plant Cell Environ. 20(1): 68–76. Vu, J.C.V., Gesch, R.W., Pennanen, A.H., Allen, L.H., Boote, K.J., Bowes, G., 2001. Soybean photosynthesis, Rubisco and carbohydrate enzymes function at supraoptimal temperatures in elevated CO2. J. Plant Physiol. 158(3): 295–307. Wada, S., Ishida, H., Izumi, M., Yoshimoto, K., Ohsumi, Y., Mae, T., et al., 2009. Autophagy plays a role in chloroplast degradation during senescence in individually darkened leaves. Plant Physiol. 149(2): 885–893. Wardley, T.M., Bhalla, P.L., Dalling, M.J., 1984. Changes in the number and composition of chloroplasts during senescence of mesophyll cells of attached and detached primary leaves of wheat (Triticum aestivum L.). Plant Physiol. 75(2): 421–424. Warren, C.R., Adams, M.A., 2004. Evergreen trees do not maximize instantaneous photosynthesis. Trends Plant Sci. 9(6): 270–274. Waters, M.T., Fray, R.G., Pyke, K.A., 2004. Stromule formation is dependent upon plastid size, plastid differentiation status and the density of plastids within the cell. Plant J. 39(4): 655–667. Weaver, L.M., Amasino, R.M., 2001. Senescence is induced in individually darkened Arabidopsis leaves but inhibited in whole darkened plants. Plant Physiol. 127(3): 876–886. Wingler, A., Mares, M., Pourtau, N., 2004. Spatial patterns and metabolic regulation of photosynthetic parameters during leaf senescence. New Phytol. 161(3): 781–789. Wingler, A., Purdy, S., MacLean, J.A., Pourtau, N., 2006. The role of sugars in integrating environmental signals during the regulation of leaf senescence. J. Exp. Bot. 57(2): 391–399. Wostrikoff, K., Stern, D., 2007. Rubisco large-subunit translation is autoregulated in response to its assembly state in tobacco chloroplasts. Proc. Natl. Acad. Sci. U.S.A. 104(15): 6466–6471.
182
L. J. IRVING ET AL.
Wright, I.J., Reich, P.B., Westoby, M., Ackerly, D.D., Baruch, Z., Bongers, F., et al., 2004. The worldwide leaf economics spectrum. Nature 428(6985): 821–827. Yang, D.H., Webster, J., Adam, Z., Lindahl, M., Andersson, B., 1998. Induction of acclimative proteolysis of the light-harvesting chlorophyll a/b protein of photosystem II in response to elevated light intensities. Plant Physiol. 118 (3): 827–834. Yang, J.C., Zhang, J.H., Wang, Z.Q., Zhu, Q.S., Liu, L.J., 2002. Abscisic acid and cytokinins in the root exudates and leaves and their relationship to senes cence and remobilization of carbon reserves in rice subjected to water stress during grain filling. Planta 215(4): 645–652. Yang, J.C., Zhang, J.H., Wang, Z.Q., Zhu, Q.S., Liu, L.J., 2003. Involvement of abscisic acid and cytokinins in the senescence and remobilization of carbon reserves in wheat subjected to water stress during grain filling. Plant Cell Environ. 26(10): 1621–1631. Yoshida, T., Minamikawa, T., 1996. Successive amino-terminal proteolysis of the large subunit of ribulose 1,5-bisphosphate carboxylase/oxygenase by vacuo lar enzymes from French bean leaves. Eur. J. Biochem. 238(2): 317–324. Zer, H., Prasil, O., Ohad, I., 1994. Role of plastoquinol oxidoreduction in regulation of photochemical reaction center II D1 protein turnover in vivo. J. Biol. Chem. 269(26): 17670–17676. Zhang, L.F., Rui, Q., Zhang, P., Wang, X.Y., Xu, L.L., 2007. A novel 51-kDa fragment of the large subunit of ribulose-1,5-bisphosphate carboxylase/ oxygenase formed in the stroma of chloroplasts in dark-induced senescing wheat leaves. Physiol. Plant. 131(1): 64–71. Zhang, X., Zhang, L., Dong, F.C., Gao, J.F., Galbraith, D.W., Song, C.P., 2001. Hydrogen peroxide is involved in abscisic acid-induced stomatal closure in Vicia faba. Plant Physiol. 126(4): 1438–1448. Zubo, Y.O., Yamburenko, M.V., Selivankina, S.Y., Shakirova, F.M., Avalbaev, A. M., Kudryakova, N.V., et al., 2008. Cytokinin stimulates chloroplast transcription in detached barley leaves. Plant Physiol. 148(2): 1082–1093.
AUTHOR INDEX
A
Aarts, M. G., 4, 9, 39
Acevedo-Hernandez, G. J., 144, 172
Achard, P., 77, 78, 81
Acharya, B., 148, 160, 172
Acosta, I. F., 69, 70, 78
Adams, M. A., 164, 181
Addiscott, T. M., 140, 172
Aerts, R., 166, 172
Aguilar-Martı´nez, J. A., 122, 128
Ainsworth, C., 55, 78
Albert, N. W., 147, 172
Alleman, M., 74, 78
Allwood, E. G., 26–27, 39, 45–46
Almeida, J., 88, 116, 125, 128, 131
Aluru, M. R., 141–142, 172
Amasino, R. M., 70–71, 79, 153, 179, 181
Ambrose, B. A., 76, 78, 94, 135
Andersson, B., 158, 173, 181
Andrew Staehelin, L., 179
Araya, T., 161, 173
Arens, N. C., 98, 130
Ariizumi, T., 4–5, 39
Arimura, S., 153, 173
Armbruster, W. S., 131
Aro, E. M., 158, 173, 177
Asada, K., 147, 173
Assmann, S., 148, 160, 172
Astrom, H., 26, 39
Aukerman, M. J., 71, 78
B
Baek, K. H., 151, 173
Banks, J. A., 54, 82
Barrett, S. C. H., 92, 111, 128, 132, 133
Basinger, J. F., 98, 128
Bassham, D. C., 17–18, 39, 52, 149, 173
Bateman, R. M., 89, 93–95, 101–102,
114, 135
Bauer, D., 161, 173
Baum, D. A., 123–124, 130, 132, 136
Baumgartner, B. J., 142, 173
Bausenwein, U., 140, 173
Baxter, C. E., 124, 128
Bensen, R. J., 63, 71, 78
Berg, R. L., 113, 128
Bernhardt, P., 100, 128
Biesmeijer, J. C., 112, 128
Blackmore, S., 4, 39
Bleecker, A. B., 149, 175
Boisson-Dernier, A., 12, 39
Bokoch, G. M., 24, 41
Borg, M., 3, 39
Borges, F., 36, 39
Bosch, M., 19, 40, 47
Botto-Mahan, C., 114, 128
Boualem, A., 57–59, 78, 81
Bowman, J. L., 71, 79
Braun, A., 89, 128
Broholm, S. K., 118, 120, 123, 128
Buchanan-Wollaston, V., 78, 146–149, 161,
173
Burgin, S., 92, 135
Busch, A., 88, 94, 118–120, 123, 128
C
Cai, G., 14, 30, 40, 49
Caldero´n-Urrea, A., 55, 62, 66–68, 79–80
Callis, J., 146–147, 149, 173
Camacho, L., 19, 40, 47
Cameron, D. D., 171, 173, 176
Capron, A., 12, 40
Cardenas, L., 31, 40
Carpenter, R., 88–89, 128–129, 133
Castonguay, Y., 161, 173
Causin, H. F., 148, 173
Certal, A. C., 19, 40
Chae, K., 11, 40
Chapman, M. A., 118, 120, 123–124,
129, 133
Charlesworth, D., 54, 79
Chen, C. Y., 12, 14, 40
Chen, T., 12, 36, 40–41, 51
Chen, W. Q., 161, 173
Chen, X., 41, 71, 79, 83
Chen, Y., 36, 40, 51
Chen, Y. F., 5, 40
Chen, Y. H., 12, 40
Cheng, P. C., 62, 79
Cheng, S. H., 144, 173
183
184
AUTHOR INDEX
Cheung, A. Y., 3, 8, 14, 19, 23, 29, 38, 40–41,
43, 46, 52
Chiba, A., 152–153, 173
Chittka, L., 112, 129
Choi, D., 151, 173
Choquet, Y., 143, 174
Chuck, G., 53–78, 79
Chung, T., 149, 174
Citerne, H. L., 85–128, 129, 135
Clapham, D. E., 25, 40
Clark, J. I., 102, 116–117, 129
Classen-Bockhoff, R., 108, 137
Cle´ment, C., 15, 40
Clifford, H. T., 62, 79, 129
Coe, E. H., 65, 80–81 Coen, E., 89, 128–129, 133
Coen, E. S., 59, 79, 101–102, 116–117, 119,
128–129, 136
Cohen, Y., 57, 80
Cominelli, E., 124, 129
Corley, S. B., 91, 117, 129
Costa, M. M., 116–117, 120, 124, 80, 128–129
Crafts-Brandner, S. J., 167, 174
Cramer, M. D., 167, 174
Crawford, B., 58, 79
Cremona, O., 24, 40
Crepet, W. L., 99–100, 129
Cresti, M., 14, 30, 40, 47, 49
Criado, M. V., 160, 173–174
Cronk, Q., 112–113, 129
Cronquist, A., 55, 79
Cubas, P., 88, 98, 116, 119, 121–124, 128–129,
132–133
D Dahlgren, R. M. T., 102–103, 129
Dai, S., 36, 40–41 Dale, E. E., 64–65, 81
Damerval, C., 85–128, 129, 133
Danforth, B. N., 98, 100, 134
Darwin, C., 87, 100, 111, 130
Dawe, R. K., 64, 79
Day, I. S., 29, 49
de Azevedo Souza, C., 5, 41
De Camilli, P., 24, 40
de Graaf, B. H., 19, 23, 40–41
De La Guardia, M. D., 56, 82
Decaluwe, H., 166, 172
Dellaporta, S. L., 55, 62, 66–68, 79–81, 83
DeLong, A., 64, 67, 79
Demartino, G. N., 150, 174
DerMardirossian, C., 24, 41
Desimone, M., 150, 174, 181
Dickinson, D. B., 15, 50
Dickinson, H. G., 6, 8, 41, 43, 49–50
Diggle, P. K., 94, 134
Dilcher, D., 99, 130
Dilcher, D. L., 98, 128, 132, 136
Dinesh-Kumar, S. P., 17, 46, 50
Dobritsa, A. A., 5, 41
Doebley, J., 121–122, 129–130, 132
Dong, X., 5, 41
Dong, Z. C., 93, 130
Donoghue, M. J., 91, 119, 122–123, 130, 132
Dorweiler, J., 74, 78, 80
Doughty, J., 7, 41, 43
Douglas, A. W., 92–93, 95, 130
Dowd, P. E., 24, 41, 52
Doyle, J. A., 97, 101, 130–131
Drager, R. G., 145, 174
Drews, G. N., 50, 73, 79
Du, H., 119, 124, 130
Du, Z. Y., 119, 130
Dubell, A. N., 142, 174
Dulberger, R., 92, 134
Dumas, C., 6, 42–43, 45–46, 49
Dutta, R., 32, 41
E
Eberwein, R., 89, 130
Edelman, M., 158, 174
Eichler, A. W., 89, 130
Elleman, C. J., 6, 41, 50
Ellis, R. J., 140, 143, 174–175
Endress, P. K., 91–95, 97, 101, 110–112, 114,
119, 130–131, 133, 135
Engel, M. S., 100, 132
Erhard, K. F., 75, 79
Eriksson, M., 113, 134
Escobar-Restrepo, J. M., 12, 41
F Faegri, K., 91, 113, 130
Fan, X., 28, 41
Faure, J., 24, 41
Feijo´, J. A., 31, 33, 39–40, 42, 44, 49
Feild, T. S., 98, 130
Feng, M., 93, 118–119, 137
Feng, X., 93, 118–119, 130
Fenster, C. B., 109, 112–113, 131
Fernando, D. D., 36, 42
Fiebig, A., 9–10, 42, 47
Filonova, L. H., 17, 42
AUTHOR INDEX Finlayson, S. A., 122, 131
Finley, D., 150, 174
Fischer, A. M., 170, 176
Fischer, R. L., 42, 150, 180
Fleming, T. H., 112, 131
Foissner, I., 26–27, 42
Folta, K. M., 144, 180
Foreman, J., 6, 42
Freeling, M., 64, 79–80
Frey, F. M., 113–114, 131, 137
Friedman, W. E., 100, 131
Friedrich, J. W., 164, 174
Friis, E. M., 98–99, 101, 131
Frohlich, M. W., 98, 131
Fu, Y., 26, 42, 52
Fujiki, Y., 18, 42
Fujioka, S., 64, 79
Fukuda, T., 123, 131
G Gaffal, K. P., 17, 42
Galego, L., 88, 116, 125, 128, 131
Gan, S., 70–71, 79
Gao, J.-Y., 92, 131
Gao, Q., 119, 131
Garzon, M., 9, 42
Gastal, F., 144, 162, 174
Gaude, T., 6, 42
Gaudin, V., 116, 131
Geitmann, A., 27, 42
Gesch, R. W., 145, 174, 181
Ghislain, M., 150, 174
Giurfa, M., 112, 128, 131, 134–135
Glick, R. E., 161, 175
Glover, B. J., 124, 134
Goebel, K., 93, 131
Goldman, M. H., 8, 42
Goldschmidt, R., 126, 131
Gomez, J. M., 113–115, 127, 131–132 Go´mez-Mena, C., 76, 80
Gompel, N., 126, 132
Gong, Y. B., 113, 132
Gordon, C., 150, 175
Gorelick, R., 100, 132
Gossot, O., 27, 42
Graham, S. W., 92, 132
Grant, V., 100, 132
Grbic, V., 149, 175
Grimaldi, D., 100, 112, 132
Grimball, P. C., 57, 82
Grindlay, D. J. C., 168, 175
Grossniklaus, U., 3, 44–45, 49
185
Gu, Y., 22, 35, 42, 44, 51
Guan, Y. F., 5, 42
Gu¨bitz, T., 123, 132
Gunning, B. E. S., 153, 175
Gustafsson, A., 87, 132
H
Haerizadeh, F., 36, 43
Hanaoka, H., 149, 175
Hancock, J., 6, 43, 48
Hannoufa, A., 9, 43, 47
Hansen, J. D., 9, 43
Hanson, M. R., 29, 49, 175
Harberd, N. P., 64, 78, 80–81
Harder, L. D., 114, 132
Harrison-Lowe, N. J., 18, 43
Hay, J. C., 24, 43
He, P., 148, 175
He, Y. H., 148, 161, 175
Heizman, P., 5–6, 43
Helentjaris, T., 63, 83
Helling, D., 24, 43
Helme, N., 92, 132
Hepler, P. K., 14–15, 37, 40, 42–43, 46–48, 51
Herrera, J., 109, 113, 132
Heslop-Harrison, J., 6, 43, 47
Heslop-Harrison, Y., 4, 43
Hibberd, J. M., 171, 175
Hidema, J., 162, 168, 175, 181
Higaki, T., 27, 43
Higashiyama, T., 11, 43
Highfield, P. E., 143, 175
Hikosaka, K., 163–164, 167–168, 175, 179
Hileman, L. C., 88, 119, 123–124, 132, 134,
136
Hillmer, S., 17, 47
Hiscock, S. J., 6–8, 41, 43, 47
Hodges, S. A., 93, 136
Hoekstra, F. A., 15, 43
Holdaway-Clarke, T. L., 32, 44
Hollick, J. B., 75, 79–80, 82
Holmes-Davis, R., 36, 44
Holweg, C., 29, 44
Holzinger, A., 152–153, 175
Honys, D., 13, 36, 44
Howarth, D. G., 122–123, 125, 132
Hu, S. S., 100, 132
Huang, F. Y., 147, 175
Huang, J.-C., 1–38
Huang, S., 28, 44, 81
Huang, S. Q., 113, 132
Hubbard, L., 122, 130, 132
186
AUTHOR INDEX
Huck, N., 12, 41, 44
Hudson, G. S., 143, 175
Huffaker, R. C., 149, 151–152, 164, 174, 176,
179
Hulskamp, M., 8–9, 44
Hultquist, J., 74, 80
Huq, E., 144, 150, 176
Hussey, P. J., 27, 44–46, 52
Hwang, J. U., 14, 33, 44
Hyodo, F., 109, 113, 136
I
Imai, K., 143, 162, 176
Inada, N., 146, 154, 176
Inoue, Y., 18, 44, 149, 176, 179
Irish, E., 65, 79
Irish, E. E., 55, 64–67, 80
Irving, L. J., 139–172, 173, 176
Ishida, H., 139–172, 173, 176–177, 181
Ishiguro, S., 50, 70, 80
Ishizuka, M., 162–163, 176
Itkin, M., 60, 80
Ito, T., 5, 44, 50, 76, 80, 83
Iwahori, S., 56, 80
J
Jabbour, F., 85–128, 133
Jackson, D., 65, 80
Jacob, F., 122, 133
Jauh, G.-Y., 1–38, 44, 48, 51
Jenks, M. A., 9, 44, 49
Jesson, L. K., 92, 111, 133
Jiang, P.-L., 15, 17, 44
Johnson, M. A., 10–12, 44
Johnson, S. D., 114, 132
Johnston, M. O., 94, 133
Johri, M. M., 65, 80
Jones, D. F., 66, 80
Jukanti, A. K., 170, 176
K
Kalisz, S., 111, 121, 133
Kamachi, S., 56–57, 80
Kandasamy, M. K., 7, 44, 51
Kaothien, P., 13, 44
Kater, M., 59, 76, 80
Kato, Y., 158, 165, 177
Katz, C., 150, 177
Kazanietz, M. G., 24, 52
Kellogg, E., 62, 81
Kellogg, E. A., 69, 81, 94, 133
Kenigsbuch, D., 57, 80
Ketelaar, T., 27, 44–45
Kettunen, R., 158, 177
Khan, S., 165, 177
Kim, J. C., 62, 80
Kim, M., 118, 120–121, 133
Kim, T. H., 11, 39
Kirchoff, B. K., 90, 133
Kjer, K. M., 112, 137
Klahre, U., 22, 43, 45
Klionsky, D. J., 17, 45
Knight, M. R., 6, 49, 180
Knox, R. B., 6, 8, 45, 47
Ko¨hler, C., 3, 45
Koizumi, M., 150, 177
Kokubun, N., 156, 165, 177
Koornneef, M., 9, 45
Kost, B., 14, 22, 24, 26, 43, 45
Kovar, D. R., 28, 45
Kramer, E. M., 88, 121, 132, 135, 137
Krapp, A., 144, 177–178
Krenn, H. W., 112, 133
Krstolic, J. L., 113, 137
Kruger, E. L., 148, 177
Kunst, L., 9, 45, 49
Kunze, H., 90, 133
L La Rocca, N., 142, 177
Laisk, A., 154, 177
Laloi, C., 6, 45
Lalonde, B. A., 7, 45, 51
Lane, I. E., 89, 133
Lattanzi, F. A., 140, 148, 163, 167, 177
Lauter, N., 72, 81, 129
Lawlor, D. W., 164, 177
Le Roux, L., 62, 81
Lebel-Hardenack, S., 69, 81
Lee, C. B., 11, 45
Lee, D. Y., 72, 81
Lee, J. Y., 36, 45
Lefebvre, B., 19, 45
Lehrer, M., 112, 133
Leppik, E. E., 88, 133
Levine, B., 17, 45–46 Li, D., 68, 81
Li, H., 22, 46
Li, P., 94, 133
Li, Z., 57, 81
Lide´n, M., 127, 133
Liebe, S., 29, 46
Linder, H., 92, 132
AUTHOR INDEX Liu, S., 60, 40, 81
Liu, Y., 17, 46
Lolle, S. J., 8, 46, 49
Lopez-Juez, E., 144, 177, 180
Lord, E. M., 10–11, 15, 42, 44–48
Lovy-Wheeler, A., 27, 33, 46
Lukens, L., 121, 130
Lunde, C., 168, 177
Luo, D., 89, 91, 115–116, 118, 129, 133, 137
Luu, D. T., 6–7, 43, 46
Lysenko, E. A., 141–142, 178
M
Ma, H., 11, 44, 46
Ma, J., 36, 46
MacMurray, A. L., 56, 81
Mae, T., 146, 151, 162, 164, 166–167, 173,
175–178, 180–181
Makino, A., 139–172, 173, 175–178, 180
Malcomber, S. T., 69, 81
Malho, R., 19, 40, 47
Manos, P. S., 113, 136
Manuel, M., 88, 133
Mara, C., 60, 81
Marazzi, B., 92, 133
Marin-Navarro, J., 150, 178
Martin, A., 58, 78, 81
Martinez, D. E., 154–156, 166, 178–179
Martı´n-Trillo, M., 116, 122, 133
Marton, M. L., 11, 46
Masclaux-Daubresse, C., 159, 178
Mathieu, J., 73, 81
Matsubayashi, Y., 5, 40, 46, 50
Matt, P., 146, 178
Matthew, C., 165, 176
Mattoo, A. K., 158, 174
Mattsson, O., 6, 47
Mayfield, J. A., 10, 42, 47
McCormick, S., 5, 12, 39, 47–48, 51–52
McInnis, S. M., 6, 43, 47
McKenna, S. T., 31, 40, 47, 51
McNaughton, G. S., 170, 179
McNevin, J. P., 9, 47
McRae, D. G., 147, 179
Meijer, H. J., 28, 47
Meinhard, M., 112, 47
Mena, M., 76, 81–82
Menzel, D., 29, 45–46
Meyerowitz, E. M., 59, 79, 80, 83, 116, 129
Mibus, H., 56–57, 81
Mickelson, S., 170, 179
Miki-Hirosige, H., 15, 47
187
Miku, V. E., 67, 81
Miller, B. L., 151, 179
Miller, C. M., 56, 81
Miller, D. D., 30, 47–48
Minamikawa, T., 51, 149, 151, 157, 161, 179, 182
Mishkind, M. L., 144, 179
Mitchell, C. H., 94, 134
Mohl, H., 89, 134
Mo¨ller, A. P., 112–114, 134
Mo¨ller, M., 123, 129
Mollet, J. C., 11, 40, 45, 47–48
Mondrago´n-Palomino, M., 125, 134
Monteiro, D., 27–28, 47
Moon, J., 150, 179
Moose, S. P., 72, 81
Morant, M., 5, 41, 47
Moreno, J., 150, 178
Mori, S. A., 97, 136
Moriyasu, Y., 17, 39, 44, 47, 176
Moscatelli, A., 19, 47
Moylan, E. C., 92, 134
Mullet, J. E., 142, 173–174
Munnik, T., 28, 47
Murata, N., 158, 180
Muschietti, J., 13, 48, 51
Mustyatsa, S. I., 67, 81
Myers, C., 32–33, 48
N Nadot, S., 85–127, 133
Nakajima, Y., 158, 179
Nakamura, S., 15, 47
Navarro, L., 77, 81
Neal, P. R., 100, 131, 134
Nebenfuhr, A., 29, 48
Neill, S. J., 6, 43, 48
Nelson, C. J., 144, 162, 174
Nelson, T. M., 55, 80
Neuffer, M. G., 60, 63, 65, 81
Nick, P., 29, 44
Nickerson, N. H., 63–65, 81
Niklas, K. J., 99–100, 129
Nishikawa, S., 5, 41, 48
Nobuta, K., 74, 81
Noda, T., 17, 48, 175
Noir, S., 36, 48
Nugent, J. M., 101–102, 119, 129, 134
O Obara, K., 147, 162, 179
Oeller, P., 56, 82
188 Ohira, K., 166, 178
Ohsumi, Y., 39, 42, 44, 48, 149,
176, 181
Oikawa, S., 166, 169, 179
Oja, V., 154, 177, 179
Ojeda, I., 112–113, 125, 129, 134
Okada, K., 12, 50, 80
Okamuro, J. K., 76, 82
Okuda, S., 11, 48
Oldenburg, D. J., 142, 179–180
Ollerton, J., 113, 134
Olsen, L. J., 18, 39, 43
Ono, T. A., 152, 179
Onodera, Y., 75, 82
Ornduff, R., 92, 134
Osmond, B., 141, 162, 178
Otegui, M. S., 154–155, 178–179
P
Pacini, E., 15, 40, 48
Palanivelu, R., 11, 48, 50
Pandey, K. K., 6, 48
Park, S. Y., 11, 45, 47–48, 81
Parkin, J., 101, 134
Parkinson, S. E., 75, 79–80, 82
Parry, M. A. J., 163, 177, 179
Parton, R. M., 19, 31, 48
Paxson-Sowders, D. M., 4, 48
Peng, J., 64, 82–83
Perez-Rodriguez, M., 116, 134
Perl-Teves, R., 56, 82
Phinney, B. O., 63, 79, 82
Phipps, I. F., 65, 82
Picton, J. M., 19, 48
Pierce, L. K., 56, 82
Pierson, E. S., 32, 48
Piffanelli, P., 5, 48
Pina, C., 36, 49
Poethig, R. S., 74, 83
Poinar, G. O., 99–100, 134
Poole, C. F., 57, 82
Potocky, M., 28, 49
Prado, A. M., 6, 49
Prenner, G., 101, 134
Presland, M. R., 170, 179
Preston, J. C., 88, 118–119,
122–124, 134
Preuss, D., 4, 8–10, 41–42, 47–52
Prins, A., 149, 156–157, 180
Proctor, M., 91, 113, 134
Prud’homme, B., 126, 132
Pruitt, R. E., 8, 44, 46, 49
AUTHOR INDEX Q Qin, G., 18, 49
Qin, Y., 37, 49
Quick, W. P., 143, 175, 177, 180
Quinn, J. A., 56, 83
R Raine, N. E., 112, 129
Ramsay, N. A., 124, 134
Rapp, R. A., 121, 134
Rashotte, A. M., 9, 49
Reardon, W., 121–122, 124, 134
Reddy, A. S., 29, 49
Ree, R. H., 124, 130, 133, 135
Reisen, D., 29, 49, 176
Ren, H., 27, 41, 49, 52
Ren, Y., 89, 135
Renshaw, A., 92, 135
Rentel, M. C., 6, 49
Rivero, R. M., 161, 180
Roberts, I. N., 6, 49–50, 173–174 Robinson, D., 162–163, 165, 170, 176
Robinson, K. R., 32, 41
Robinson, R. W., 56, 82
Rodermel, S., 143, 172, 180
Rodriguez, I., 112, 135
Romagnoli, S., 27, 49
Ronse De Craene, L. P., 89–90, 93, 95, 97,
133, 135, 137
Rood, S., 63, 82
Rosin, F. M., 88, 121, 135
Rotman, N., 12, 49
Rowland, O., 5, 9, 49
Rudall, P. J., 89, 93–95, 101–102, 114, 134–
135
Russell, S. D., 15, 46
S Sagan, L., 153, 180
Saito, S., 57, 82
Sakagami, Y., 5, 40, 46, 50
Sakai, H., 71, 78–79 Sakakibara, H., 145, 180
Samuels, A., 9, 45
Sandaklie-Nikolova, L., 3, 50
Sargent, R. D., 98, 133, 135
Sarker, R. H., 9, 50
Sawbridge, T. I., 144, 180
Schaffer, M. A., 150, 180
Scheible, W. R., 145–146, 150, 180
Schein, M., 10, 50
AUTHOR INDEX Schiott, M., 32, 50
Schmidt, G. W., 144, 179
Schmidt, R. J., 73, 78, 82, 94, 135
Schumann, K., 89, 135
Scott, R. J., 5, 50
Seay, M. D., 17, 50
Sehr, E. M., 89, 135
Sergiev, I., 148, 180
Shannon, S., 56, 82
Shaver, J. M., 142, 180
Sheen, J., 144, 180
Shimizu, K. K., 12, 50
Shinohara, H., 5, 46, 50
Shivanna, K. R., 4, 43
Shykoff, J. A., 113, 134
Sisco, P. H., 72, 81
Slaughter, C. A., 150, 174
Smalle, J., 150, 180
Smets, E., 95, 97, 135
Smith, J. F., 123–124, 136–137
Song, C. F., 119, 136
Sorci, G., 112, 134
Southworth, D., 15, 50
Spalding, E. P., 144, 180
Spray, C., 63, 82
Sprengel, C. K., 89, 136
Staiger, C. J., 27, 41, 44–45, 50
Stead, A. D., 6, 50
Steer, M. W., 19, 48
Stenmark, H., 23, 50
Stern, D., 143, 181
Sun, G., 89, 101, 136
Suzuki, G., 7, 50
Suzuki, T., 5, 7, 44, 50, 176
Suzuki, Y., 139–172, 176, 180
Swanson, R., 5, 9, 50
T
Takahashi, S., 158, 180
Takayama, S., 7, 50
Takeuchi, A., 151, 181
Tang, W., 13, 51
Tanurdzic, M., 54, 82
Tatlioglu, T., 57, 81
Taylor, L. P., 15, 37, 51
Telfer, A., 158, 181
Terashima, I., 163, 173, 175
Theissen, G., 125, 127, 134, 136
Thomas, C., 28, 51
Thompson, J. E., 147, 179
Thorp, R. W., 112, 136
To, J. P. C., 145, 181
Tonelli, C., 124, 129
Toyooka, K., 18, 51, 179
Trebitsh, T., 57, 82
Tripp, E. A., 113, 136
Tsai, W. C., 125, 136
Tsou, C.-H., 97, 136
Tsukada, M., 149, 181
Tucker, S. C., 90, 92–95, 130, 136
Twell, D., 13, 36, 39, 44
U Umbach, A. L., 7, 51
Updegraff, E. P., 10, 51
Ushimaru, A., 109, 113, 136
V
Vallon, O., 143, 174
Van der Graaff, E., 148–149, 162, 181
van der Pijl, L., 91, 113, 130
Van Gestel, K., 29, 51
van Kleunen, M., 113, 136
Vidali, L., 28, 40, 43, 47, 51
Vierstra, R. D., 150, 174, 180
Vincent, C. A., 93, 136
Volin, J. C., 148, 177
Vu, J. C. V., 145, 174, 181
W Wada, S., 152–153, 155–156, 164, 181
Wang, C.-S., 44
Wang, H., 76, 52, 82
Wang, H. J., 1–38, 51
Wang, W., 94, 119, 137
Wang, X., 28, 51, 136
Wang, Y., 36, 51
Wang, Y. Z., 118–119, 125, 131, 136–137
Wang, Z. Y., 118–119, 125, 130
Wanntorp, L., 90, 137
Ward, J. M., 76, 83
Wardley, T. M., 151, 181
Warren, C. R., 164, 181
Waser, N. M., 100, 134, 137
Wasternack, C., 69, 83
Waters, M. T., 153, 181
Wendel, J. F., 121, 134
Wengier, D., 13, 44, 51
Westerkamp, C., 108, 137
Whitfield, J. B., 112, 137
Wignall, A. E., 112, 137
Wilson, A. C., 121, 137
Wingler, A., 145–147, 159, 181
189
190
AUTHOR INDEX
Winkler, H., 90, 137
Winkler, R., 63, 83
Wolfe, L. M., 113, 137
Wolters-Arts, M., 8–9, 51
Wood, T. E., 126, 137
Wostrikoff, K., 143, 174, 181
Wright, I. J., 140, 181
Wu, G., 35, 42, 51, 74, 83
Wu, H. M., 3, 11, 14, 29, 36, 40–41, 52
Wu, J., 74, 81
Wu, X., 3, 14, 29, 36, 67–68, 51, 83
Wu, Y. M., 74, 130
Wydler, H., 89, 137
X
Xiang, Y., 27, 28, 49, 52
Xiong, Y., 17, 52
Y
Yamasaki, S., 56–57, 82–83
Yang, C., 24, 52
Yang, D. H., 159, 181
Yang, J. C., 148, 160, 181–182
Yang, W. C., 5, 41
Yephremov, A., 8, 52
Yin, T., 56, 83
Yoon, G. M., 32, 52
Yoshida, S., 149, 151, 161, 179
Young, T. E., 71, 83
Yu, H., 76, 80, 83
Yuan, Z., 121, 137
Z
Zachgo, S., 88, 94, 118–119, 120, 123, 128
Zer, H., 158, 182
Zhang, K., 5, 22, 40, 45
Zhang, L., 5, 22, 160, 40, 49, 130, 182
Zhang, L. F., 156–157, 182
Zhang, P., 156–157, 182
Zhang, W., 98, 118–120, 123, 137
Zhang, X., 160, 182
Zhang, Y., 5, 22, 52
Zhao, L., 71, 75, 83, 135, 179
Zhou, L., 32, 52
Zhou, X. R., 119, 122, 124–125, 137
Zinkl, G. M., 4, 6, 9, 48, 52
Zubo, Y. O., 145, 182
SUBJECT INDEX
A Abscisic acid (ABA), 144–145, 160
signalling, 147–148
Actin
associated motor proteins
vesicles/organelle movement, 29–31
binding activity of NtADF1, 27
in pollen tubes
distribution study, 26
dynamics by ABPs, 27
remodelling and pollen tube growth, 20–21
Actin-binding proteins (ABPs), 20–21
formin, 29
plant development and pollen tube
growth, 27
profilin, 28
remodelling and dynamics, 27–29
in tip region
Hþ and Ca2þ gradients, 30, 32
villins/gelsolins, 28
Actinomorphy, 89
AGAMOUS-related MADS box gene, 59
Alocasia, 163–164
1-Aminocyclopropane-1 carboxylic acid
synthase (ACS) genes, 56
a-Amylase in germinating lily pollen, 18
Amylogenesis/amylolysis, 15
Antirrhinum majus, 91, 115
APETALA2 (AP2) family of transcription
factors, 71
Apical region-excluded localization of
Hþ-ATPase, 19
Arabidopsis eceriferum, 144–147, 149, 152,
164–165
AGAMOUSLIKE MADS box
transcription factors, 73
AP2 gene, regulator of floral organ, 71
autophagy-related genes in
AtATGs, 18
capping activity, 28
carpel development in, 58
defective pollen coat in, 4
ECERIFERUM (CER) mutants, 8
cer-mutant pollen hydration, 9
genome, myosin-like genes in, 29
germinating pollen
organellar differentiation and
organization, 16
phenomenon for, 17
leaves of, 161
microspores in bicellular and tricellular
stages, 36
mutual consent (amc) study by, 12
mya2-1 and mya2-1 mutants, 29
pollen-specific calcium regulatory
proteins, 32
proteome mapping, 36
proteomic analysis of, 9–10
RAD-like genes (AtRLs), 124
RDR2 gene in, 74
receptor-like kinase (RLK) genes in, 13
RhoGEFs in polar tube growth, 22
SAG12 senescence-induced promoter, 70
SQUAMOSA PROMOTER BINDING
LIKE (SPL), 74
tip-focused Ca2þ gradient, 32
Asymmetric flowers
Cannaceae and Valerianaceae, 92
Zygogynum and Senna, 92
ATG. See Autophagy (ATG) Atmospheric CO2 concentration, 145–146 ATP-dependent process attachment of ubiquitin to protein, 150
post-transcriptional phosphorylation, 158
protein degradation, 146
Autophagy (ATG), 17–18, 149
atg mutants, 149, 152
B Basic helix–loop–helix domain (bHLH), 122
BFA. See Brefeldin A (BFA)
Bilateral symmetry, 87
Bournea leiophylla, 122
Brassica oleracea
acetone treatment, 5
pollen hydration in, 9
pollen–stigma cross-linking adhesion in, 7
Brefeldin A (BFA), 17
191
192
SUBJECT INDEX
C Cadia purpurea, 122
Calcineurin B-like (CBL) proteins, 32
Calcium-dependent protein kinases (CDPKs)
in signalling cascades, 25
Callose in vesicles, 15
Calmodulin-like protein (CML), 32
CA-1-P binding, 163
Capping proteins (CPs), 28
Carbohydrates
accumulation, 144
for pollen grain growth, 15
Carotenoid biosynthesis mutants, 142
CDPKs. See Calcium-dependent protein
kinases (CDPKs)
Cell death, leaf cell, 162
Cellular regulation
of protein
degradation, 146–149
synthesis, 141–146
Central Cell Guidance (CCG) gene, 12
Ceratopteris richardii
sex determination study in, 54
CER3, exine formation, 5
CES. See Control by epistasy of synthesis
(CES)
Chemocyanin, 11
Chenopodium, 163–164
Chlamydomonas, 158
Chloroplasts, 141
DNA, 142
genome, 141
stromal protein degradation, 151–157
Chloroplast-targeted cyan fluorescent protein
(CFP), 154
Chloroplast-targeted green fluorescent
protein (GFP), 152
Class A pollen coat proteins (PCP-A), 7
CML. See Calmodulin-like protein (CML)
Colchicine, 30
See also Microtubule-associated proteins
(MAPs)
Constitutive triple response 1 mutant, 77
Control by epistasy of synthesis (CES), 143
Cucumber. See Cucumis sativus L.
Cucumis melo
sex determination in
alleles, 57
carpel development and, 59
chromosome walk, 58
CmACS-7 genes, 57
Cucumis sativus L.
sex determination in
F and M genes cloning, 56
gibberellins (GA) and ethylene, 56
hormones controls, 56
loci, 56
CUM26 MADS box gene, 59
Cycloheximide-treated stigma, 9
CYCLOIDEA (CYC) genes, 115–116
Arabidopsis thaliana, 123
Asteraceae, 120
bHLH, 122
Chirita heterotricha, 119
CYC2 genes, 123–124
dorsal expression, 123
expression patterns of, 118
Gerbera hybrida, 120
Helianthus annuus, 124
I. amara case study for, 120
IaTCP1 in, 120
KEELEDWINGS 1 (KEW1), 119
Lamiales, 119
LOBED STANDARD 1 (LST1), 119
Malpighiales, 123
Mohavea confertiflora, 119
REP1 in rice, 120–121
studies, 118–119
TEOSINTE BRANCHED 1 (TB1), 122
See also Floral symmetry evolution
Cytochrome P450, exine formation, 5, 152
Cytokinins, 70–71, 145
inducible genes, 148
D Delphinieae, 94
See also Ranunculaceae, actinomorphic
flowers
DEX1, exine formation, 4
Diacyl glycerol (DAG)
apical region in growing pollen tubes,
targeting, 25
role in signalling transduction in tube
growth, 25
specific GFP marker and PtdIns-PLCs
activity, 24–25
transport, 20–21
DICHOTOMA (DICH) genes, 115–116
DIVARICATA (DIV) genes, 116
DNA-binding protease, 165
Double fertilization, 3
Double-membrane structure, plant
chloroplasts, 141
Dry stigmas, plants with, 4
hydration of pollen, 8
SUBJECT INDEX E Eastern gammagrass. See Tripsacum dactyloides Endocytosis, 14
Endomembrane trafficking, 20–21
exo/endocytotic vesicles in, 18
GFP-tagged cytoplasmic membrane
proteins, 19
in tip region, rapid and polar growth, 14
cellular modification and
differentiation, 15–18
Endosymbiotic theory, 141
Environmental regulation of senescence
protein degradation, 159–162
ER-localized HSP70 molecular chaperone
(BIP), 18
Escherichia coli polymorphisms in, 57
Ethylene, 149
as feminizing agent, 56
in monoecious and gynoecious plants,
57
Eukaryotic cells from ancestral bacteria, 141
Exine, pollen–stigma adhesion, 4
Exocytosis, 14, 18
F Faceless pollen-1, exine formation, 4
Fatty acyl-CoA synthetase, exine formation, 5
Feminization, factors involved in
an1, d1, d2, d3 and d5 mutants, 63
andromonoecious dwarves, 63
GA concentrations, 63–64
genetic proof, 63
Feronia (fer)/sire´ne (sir) mutant
role in signalling pathway for fertilization,
12
Floral symmetry evolution
angiosperms, distribution of
speciose taxa, 98
zygomorphy, 97–98
Asteridae, 95
bilateral symmetry, 87
Cariniana micrantha, 97
conservation and divergence
AtRLs, 124
DIV-like genes, 125
MADS-box transcription, 125
MYB genes, 124
Phalaenopsis equestris, 125
RAD-like genes, 124
SYMMETRIC PETALS 1 locus, 125
193
cyc-like genes
Arabidopsis thaliana, 123
Asteraceae, 120
bHLH, 122
Chirita heterotricha, 119
CYC2 genes, 123–124
dorsal expression, 123
expression patterns of, 118
Gerbera hybrida, 120
Helianthus annuus, 124
I. amara case study for, 120
IaTCP1 in, 120
KEELEDWINGS 1 (KEW1), 119
Lamiales, 119
LOBED STANDARD 1 (LST1), 119
Malpighiales, 123
Mohavea confertiflora, 119
REP1 in rice, 120–121
studies, 118–119
TEOSINTE BRANCHED 1 (TB1), 122
developmental trajectories, 95–96 floral constraints on
Aponogeton, 105
Asterids, 105, 108
Cleomaceae, 105
phylogenetic tree of Rosid families,
105–108
polyandry, 108
Resedaceae, 105
study, 105
floral zygomorphy, 97, 108
flowers and inflorescences, architecture of,
86
A. majus, 102
Apiaceae, 102
centroradialis mutant, 102
cymose inflorescences, 101
Maximum Parsimony use for, 102
monocots, mirror trees of, 102–104
morphogenetic gradients, 102
morphological analyses, 101
perianth symmetry, 100–101
racemose inflorescences, 101
terminal meristems, 101
terminal peloria, 102
zygomorphy, 102
genetic mechanisms underlying
Bournea leiophylla, 122
Cadia purpurea, 122
interspecific hybridization, 121
L. vulgaris, 121
Plantago lanceolata, 121
RAY locus, 121
194
SUBJECT INDEX
Floral symmetry evolution (Continued) growth and organ elaboration, impact of A. majus and Rehmannia angulata, 94
Gesneriaceae, 95
Poales sensu lato, 94
Sinningia cardinalis, 94
spurs are floral, 95
Stephania dielsiana, female flowers of, 94
Labiateae and Scrophulariaceae, 87
Linaria vulgaris, 87
molecular bases of
asymmetric expression in, 116
DEFICIENS and PLENA, 117
gene interactions, 117
gene regulatory network in Antirrhinum
majus, 115–117
RADIALIS (RAD) genes, 117
single-repeat MYB transcription, 117
peloria, 87
Anthirrhinum and Linaria, elucidation of, 88
principal types of, 87
rotational symmetry, 88
studies of, 88
TCP gene family in, 88
types of, 90
zygomorphy, emergence of, 98
bilateral symmetry, 99
plants and insects, coevolution, 100
timescale, 99
Turonian geological stage, 99
Floral zygomorphy, 97
Formin, 29
See also Actin-binding proteins (ABPs)
Fructans in vesicle/cytosol, 15
G GABA metabolism, pop2 mutant and, 11
Gene expression in pollen tubes, 36–37
Gerbera hybrida, 120
GFP-fused mouse Talin actin-binding domain
(GFP-mTalin)
actin indicator in living cells, 26
Gibberellins (GA)
biosynthesis, 76
induced germination
AGAMOUS-like genes, 76
LEAFY PETIOLE genes, 76
JA biosynthesis and, 76–77
as masculinizing agent, 56
b-1,3 Glucan synthases, exine formation, 4–5
Glucose in cytosol, 15
Glutamate receptor proteins, 147
Glutamine synthetase (GS), 147
Glycine max transcriptome analysis, 36
Glycine-rich proteins (GRPs)
oleosin domain and motif, 10
Glycosylated tobacco transmitting tissue-
specific (TTS) proteins, 11
Grasses, new cells production in, 142
Green fluorescent protein (GFP), 152
GRPs. See Glycine-rich proteins (GRPs)
GTPase-activating factors (GAPs), 20–21
intrinsic GTPase activity, 22
GTPase regulatory proteins, 22
Guanine nucleotide dissociation inhibitor
(GDI), 20–21
small GTPases and, 22
Guanine nucleotide-exchange factor
(GEF), 20–21
exchange of GDP to GTP, 22
H Haploid male gametophyte role in fertilization and crop production, 3
Helitron transposon insertions, 71
Hydration of pollens, 8–10
I
Iberis amara, 94
Inherent optimization, 169
Inositol 1,4,5-trisphosphate
(Ins[1,4,5]P3), 24
Isopentenyl transferase gene (IPT), 70–71
K b-Ketoacyl CoA synthase for synthesis of long chain lipids, 8
Kinase-signalling cascades, 147
KNS2 and LAP3, exine formation, 5
L Leaves
cytokinins, 148
nitrogen content in, 163
Rubisco, 170
concentration, 145–146
senescence, 146, 148
LePRK-mediated signal transduction, 13
SUBJECT INDEX Leucine-rich repeat receptor kinase (LRR-RK), 5
LHCII thylakoid proteins, 152, 159
Light-harvesting complex b gene (Lhcb), 142
Lilium longiflorum
in vivo-grown pollen tube, 37
Lily
germinating pollen
a-amylase in, 18
organellar differentiation and
organization in, 16
papain-type proteinase (SH-EP) in, 18
phenomenon for, 17
lily LIM1 (LlLIM1) expression, 26
pollen tubes, growth rates study, 14
Linaria vulgaris flower study, 87
Lipids
biosynthesis
cer mutants and, 9
for pollen grain growth, 15
Long-chain fatty acid omega-hydroxylase,
exine formation, 5
LORELEI gene, fertilization and, 12
LRR-RK. See Leucine-rich repeat receptor
kinase (LRR-RK) M MADS box genes, 59–60
Magatama mutants (maa1 and maa3), 12
Maize. See Zea mays L.
Maize ZmEA1 downregulation, 11
MALE STERILITY 2, exine formation, 4
MAPs. See Microtubule-associated proteins
(MAPs) Masculinization, genes involved in tassel seeds mutants
ts1 and ts2 genes, 64
ts4 and Ts6 genes, 65
Mediator of paramutation1 (mop1) mutants, 74
Melon. See Cucumis melo
Microtubule-associated proteins (MAPs)
pollen tube elongation, 30
Mitochondria, 141
Mohavea confertiflora, 119
Mutator transposons, 71
Myosin family, 29
N NADP-G3PDH, shading, 168
NEF1, exine formation, 4–5
Nicotiana alata
195
NaTTS from, 11
Nicotiana tabacum stigma exudates of, 8
Nitrogen remobilization, 140
Non-photosynthetic quenching (NPQ), 147
NPQ. See Non-photosynthetic quenching
(NPQ)
NtRab2 activity
downregulating by overexpression, 23
Nutrient limitation, 166
O Oblique zygomorphy, 89–90
Oryzalin, 30
See also Microtubule-associated proteins
(MAPs)
Oryza sativa gene expression, 36
Oscillatory calcium and proton gradients, 34–35
P Papain-type proteinase (SH-EP) in germinating lily pollen, 18
Paraboea rufescens, 92
Pectin methylesterase (PME)
PME–GFP fusion proteins, 19
Peloria, 87
Phloem-feeding parasites, 171
Phosphatidylinositol (4,5)-bisphosphate
(PtdIns[4,5]P2), 23
in apex of pollen tubes, 24
DAG and Ins[1,4,5]P3, 24
distribution, 20–21
downstream metabolism of, 24
in elongating pollen tubes, 24
NtPLC3 activity and, 24
and Rac-Rop GTPases, 24
as Rho GDI displacement factor
(RhoGDF), 24
Phosphatidylinositol monophosphate kinase
(PtdlnsPK), 23
Photoassimilate, 148
Photosynthesis, 141
enzymes in, 146
Photosystem II (PSII)
psbD genes, 145
rbcL gene and, 145
Phyllotaxis types, 93
Phytochrome-interacting factor 1 (PIF1), 144
Phytosulphokine-a (PSK-a)
pollen germination and, 5
196
SUBJECT INDEX
Plantago lanceolata, 121–122
chlorophyll levels, 164
Plants
cell endoproteases
aspartate proteases, 149
CPs, 149–150
metalloproteases, 149
serine proteases, 149
parasitism, 171
proteases, protein degradation, 149–151
Plastid DNA, 141–142
Plastids for storage of carbohydrates, 15
PME. See Pectin methylesterase (PME)
Pollens
adhesion on stigma
cer6-2 mutant, 6
“coat conversion,” 6
cross-linking, 5
“foot” structure, 5–6
initial, 4–5
membrane-like exinic outer layer
(EOL) of, 6
protease treatment, 6
protein pairs in, 7
role of SLG in, 7
hydration on stigma, 8–10
and female fertility of transgenic
STIG1-barnase plants, 8
fiddlehead mutant, 8
mentor effect, 8
NO synthesized, 6–7
population effect, 5
tube
endocytosis in, 19
endomembrane trafficking in, 20–21
FM4-64 staining study, 19
oscillatory growth pattern, 31–32, 34–35
small GTPases and actin remodelling,
20–21
tip, regions in, 14
tube growth in style
endocytosis and exocytosis, 14
GABA, role in, 11–12
30 hapless (hap) mutants, 12
hypotheses for, 10
LePRK1 and LePRK2, role in, 13
stages of, 10
Profilin, 28
See also Actin-binding proteins (ABPs)
Protein
degradation, 146
cellular regulation of, 146–149
chloroplast stromal, 151–157
environmental regulation of senescence,
159–162
plant proteases, 149–151
thylakoid-associated proteins, 157–159
synthesis, 160–161, 169
cellular regulation of, 141–146
turnover in whole plants, implications of,
168–171
Proteolysis, 149
Proteosome, 150
PsbA gene in chloroplast signalling in
developing leaves, 141
R Rac-Rho GTPases
oscillatory activity, 33–35
and pollen tube growth, 20–21
gain-of-function assays, 22
regulatory components of, 22
signalling, NtRac5 from, 22
and PtdIns[4,5]P2, 24
shank region-resident, 23
RADIALIS (RAD) genes, 117
Radial symmetry, 89
Ranunculaceae, actinomorphic flowers, 94
Reactive oxygen species (ROS)
cell signalling in, 6
Required to maintain repression 6 (rmr6) mutant, 75
RETARDED PALEA1 (REP1) in rice, 120–121
Rho GDI displacement factor (RhoGDF), 24
Ribulose-1,5-bisphosphate carboxylase/
oxygenase (Rubisco), 140, 166
activation, 163
concentration, 163
contents, 165
degradation, 151
in plant leaves, 155
fragmentation, 150
rbcL mRNA, 143
rbcS 1A, 144
rbcS antisense plants, 143
rbcS 1B, 144
rbcS 2B, 144
gene expression, 152
rbcS 3B, 144
rbcS genes expression, 145
rbcS mRNA, 143
rbcS-overexpressing plants, 143
RNA polymerase, 141–142
subunits, chloroplast genome, 143
SUBJECT INDEX synthesis, 144, 162
regulation of, 142
ROS. See Reactive oxygen species (ROS)
Rotational symmetry, 88
See also Symmetry
Rubisco. See Ribulose-1,5-bisphosphate
carboxylase/oxygenase (Rubisco)
Rubisco-containing bodies (RCB), 150–155
in living leaf cells of Arabidopsis, 154
Rubisco small subunit (RBCS) gene
(RbcS), 142
RUPTURED POLLEN GRAIN1, exine
formation, 5
S SAG. See Senescence-associated genes (SAG)
SCA. See Stigma/style cysteine-rich adhesin
(SCA)
Seed development, 3
Senecio squalidus SSP from, 6
Senescence, 161
Senescence-associated genes (SAG), 148–149
Senescence-associated vacuoles (SAV), 154–156
Sex determination for genetic fitness, 54
DELLA signalling pathway, 77
and floral homeotic regulators, 77
genes, genetic interaction between, 66–67
hormones
biosynthesis pathways, 55
involved in, 77
molecular data on, 55
signal
positional specificity of, 59–60 silkless1 (sk1) mutation, 66
Sig6 mutants, 142
Silene latifolia
sex determination study in, 54, 69
SLG. See S-Locus glycoprotein (SLG)
S-Locus glycoprotein (SLG)
pollen–stigma adhesion, role in, 7
S-Locus-related 1 (SLR1)
antisense suppression, 7
Small GTPases
and pollen tube growth, 20–21
Rop, exine formation, 5
SSP. See Stigma-specific peroxidase (SSP)
Starch in amyloplasts, 15
Starvation-induced senescence, 148
Stigma-specific peroxidase (SSP), 6
Stigma/style cysteine-rich adhesin (SCA), 11
Stromal proteins, 152
197
Stromules, chloroplast envelope, 152
structure and function, 153
Subapical-targeted exocytosis, 19
Sucrose in cytosol, 15
Symmetry, 91
asymmetric flowers, 92
defined, 88
floral symmetry and pollination
syndromes, 112
bee pollination syndrome, 113
functional groups of, 113
floral traits in zygomorphic and actinomorphic species, variability
Erysimum mediohispanicum, 114
flower–pollinator interaction, 115
fluctuating asymmetry, 114
genotype quality, 114
highly synorganized flowers, 114
integrative approaches, 113
low fluctuating asymmetry, 114
pollinator assemblages, 114–115
shape variations, 114
zygomorphic flowers, 114
and flower development
developmental trajectories, 95–96
establishment of, 93–94
growth impact, 94–95
and organ elaboration, 94–95
of gynoecium, 89
plant–pollinator interactions,
significance, 108
mating strategies promoting
outcrossing, 111
zygomorphy and outcrossing strategies,
109–111
pollinator preferences and perception of
bees, 112
insect, 112
See also Floral symmetry evolution
Synergid cells secreted defensin-like LUREs, 11
T Tassel seeds mutants lower floret abortion cytokinins involved with TS1 and TS2, 70–71 molecular identity of
dwarf and class I tasselseed mutants, 68
gynomonoecious sex form1 (gsf1)
mutant, 68
indeterminate spikelet1 (ids1), 72
supernumerary bract (snb) mutant, 72
ts1 gene, 69–70
198
SUBJECT INDEX
Tassel seeds mutants (Continued)
ts2 gene, 67, 69
ts4 gene, 71–73
Ts6 gene, 73–74
ts2 RNA expression, 68
zma-MIR172e, 72
pistil abortion in
microRNAs, 74
mop1 mutants, 74
rmr6 mutant, 75
SQUAMOSA PROMOTER BINDING
LIKE (SPL), 74
ts1 and ts2 genes, 64
ts4 and Ts6 genes, 65
TGN. See Trans-Golgi network (TGN)
Thylakoid-associated proteins, 157–159
Tobacco, 165
pollen tubes
actin filaments in, 26–27
growth rates study, 14
NtADF1overexpressing in, 27
PtdIns[4,5]P2-specific phospholipase C
(PtdIns-PLC) in, 24
rbcS mutants, 143
Tomato AGAMOUS-like gene
ACS2 promoter and, 60
Trans-Golgi network (TGN)
autophagosome formation, 17
Transverse zygomorphy, 90
Tripsacum dactyloides
ts2 gene function in, 68
Tropaeolum, 95
TTS proteins. See Glycosylated tobacco
transmitting tissue-specific (TTS)
proteins
U Ubiquitin-conjugating enzymes, 150
Ubiquitin-protein ligase (E3), 150
V
Vacuolar proteases, 151
Vesicle trafficking in living tubes, 18
Villin, actin-severing factor, 20–21
W Wachendorfia paniculata, 92
WAX2/YRE/FLP1 (At5g57800) gene function
in wax biosynthesis, 9
Wet stigmas, plants with, 4
hydration of pollen, 8
White campion. See Silene latifolia
Whole-leaf regulation of protein
content, 162–168
X
Xanthium canadense plants, 166, 169
Y Yellow fluorescent protein (YFP)-fused
C terminus of myosins, 29
Z Zea mays L. sex determination in AP2 genes and, 75–76 CYCLIN B expression, 62
ear spikelet, 61
floral meristem (FM), 62
florets, 60
glumes, sterile leaves, 62
gynoecial-specific nuclear degeneration, 62
inflorescence meristem (IM), 60
mop1 mutants, 74
rmr6 mutant, 75
in situ hybridization, 61
spikelet pair meristems (SPM), 60
SQUAMOSA PROMOTER BINDING
LIKE (SPL), 74
stamens and carpel primordia, 62
tassel primordium, 60
WEE1 regulator of mitosis, 62
wild-type male tassel
inflorescence, 61
Zygomorphy
in A. majus, 93
in Arachnites uniflora, 102
developmental processes, 93
in Gethyllis atropurpureum, 102
Lotus japonicus, 93
in Paphiopedilum appletonianum, 102
in Proteaceae, 95
in Tecophilaea cyanocrocus, 102
in Thismia americana, 102
transverse zygomorphy, 90
zygomorphic flowers
in Adonis and Nigella, 91
Geranium, 91
petals, combination of, 91
single symmetry axis, 89, 91
Solanum and Gladiolus, 92
