Biology of the Plant Cuticle (Annual Plant Reviews, Volume 23) [1 ed.] 140513268X, 9781405132688, 9781405171571

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Biology of the Plant Cuticle

Annual Plant Reviews A series for researchers and postgraduates in the plant sciences. Each volume in this series focuses on a theme of topical importance and emphasis is placed on rapid publication. Editorial Board: Professor Jeremy A. Roberts (Editor-in-Chief), Plant Science Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough, Leicestershire, LE12 5RD, UK; Dr David Evans, School of Biological and Molecular Sciences, Oxford Brookes University, Headington, Oxford, OX3 OBP; Professor Hidemasa Imaseki, Obata-Minami 2419, Moriyama-ku, Nagoya 463, Japan; Dr Michael T. McManus, Institute of Molecular Biosciences, Massey University, Palmerston North, New Zealand; Dr Jocelyn K.C. Rose, Department of Plant Biology, Cornell University, Ithaca, New York 14853, USA. Titles in the series: 1. Arabidopsis Edited by M. Anderson and J.A. Roberts 2. Biochemistry of Plant Secondary Metabolism Edited by M. Wink 3. Functions of Plant Secondary Metabolites and their Exploitation in Biotechnology Edited by M. Wink 4. Molecular Plant Pathology Edited by M. Dickinson and J. Beynon 5. Vacuolar Compartments Edited by D.G. Robinson and J.C. Rogers 6. Plant Reproduction Edited by S.D. O’Neill and J.A. Roberts 7. Protein–Protein Interactions in Plant Biology Edited by M.T. McManus, W.A. Laing and A.C. Allan 8. The Plant Cell Wall Edited by J.K.C. Rose 9. The Golgi Apparatus and the Plant Secretory Pathway Edited by D.G. Robinson 10. The Plant Cytoskeleton in Cell Differentiation and Development Edited by P.J. Hussey 11. Plant–Pathogen Interactions Edited by N.J. Talbot 12. Polarity in Plants Edited by K. Lindsey 13. Plastids Edited by S.G. Moller 14. Plant Pigments and their Manipulation Edited by K.M. Davies 15. Membrane Transport in Plants Edited by M.R. Blatt 16. Intercellular Communication in Plants Edited by A.J. Fleming 17. Plant Architecture and its Manipulation Edited by C. Turnbull 18. Plasmodesmata Edited by K.J. Oparka 19. Plant Epigenetics Edited by P. Meyer 20. Flowering and its Manipulation Edited by C. Ainsworth 21. Endogenous Plant Rhythms Edited by A. Hall and H. McWatters 22. Control of Primary Metabolism in Plants Edited by W.C. Plaxton and M.T. McManus 23. Biology of the Plant Cuticle Edited by M. Riederer

Biology of the Plant Cuticle Edited by MARKUS RIEDERER and CAROLINE MÜLLER Julius-von-Sachs-Institut für Biowissenschaften Universität Würzburg Germany

© 2006 by Blackwell Publishing Ltd Editorial Offices: Blackwell Publishing Ltd, 9600 Garsington Road, Oxford OX4 2DQ, UK Tel: +44 (0)1865 776868 Blackwell Publishing Professional, 2121 State Avenue, Ames, Iowa 50014-8300, USA Tel: +1 515 292 0140 Blackwell Publishing Asia, 550 Swanston Street, Carlton, Victoria 3053, Australia Tel: +61 (0)3 8359 1011 The right of the Author to be identified as the Author of this Work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. 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, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. First published 2006 by Blackwell Publishing Ltd ISBN-10: 1-4051-3268-X ISBN-13: 978-1-4051-3268-8 Library of Congress Cataloging-in-Publication Data Biology of plant cuticle / edited by Markus Riederer & Caroline Muller. p. cm. Includes bibliographical references. ISBN-13: 978-1-4051-3268-8 (hardback: alk. paper) ISBN-10: 1-4051-3268-X (hardback: alk. paper) 1. Plant cuticle. I. Riederer, Markus. II. Muller, Caroline. QK725.B575 2006 574.4′ 51–dc22 2005022952 A catalogue record for this title is available from the British Library Set in 10/12 pt Times by Newgen Imaging Systems (P) Ltd. Chennai, India Printed and bound in India by Replika Press Pvt, Ltd, Kundli The publisher’s policy is to use permanent paper from mills that operate a sustainable forestry policy, and which has been manufactured from pulp processed using acid-free and elementary chlorine-free practices. Furthermore, the publisher ensures that the text paper and cover board used have met acceptable environmental accreditation standards. For further information on Blackwell Publishing, visit our website: www.blackwellpublishing.com

Contents

Contributors Preface

1

2

Introduction: biology of the plant cuticle MARKUS RIEDERER 1.1 The evolution of the plant cuticle 1.2 Major functions of the plant cuticle 1.2.1 Transpiration control 1.2.2 Control of loss and uptake of polar solutes 1.2.3 Controlling the exchange of gases and vapours 1.2.4 Transport of lipophilic substances 1.2.5 Water and particle repellence 1.2.6 Attenuation of photosynthetically active and UV radiation 1.2.7 Mechanical containment 1.2.8 Separating agent in plant development 1.2.9 Interface for biotic interactions 1.3 Convergence with other integuments 1.4 Objectives of this book References The fine structure of the plant cuticle CHRISTOPHER E. JEFFREE 2.1 Introduction 2.1.1 The distribution of the plant cuticle 2.1.2 Definition and nomenclature of the plant cuticle 2.1.3 The pectin lamella 2.2 The structure of the cuticle proper 2.2.1 The procuticle 2.2.2 The cuticle proper 2.2.2.1 Lamellate substructure of the CP 2.2.2.2 Lamella position and orientation 2.2.2.3 What are the CP lamellae? 2.3 Cuticle polymers 2.3.1 Chemical types in Angiosperm and Gymnosperm cuticles 2.3.2 The algal cuticle

xv xvii 1 1 2 2 3 3 4 4 5 5 5 6 6 7 8 11 11 11 13 14 16 16 21 23 34 36 38 38 40

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3

CONTENTS

2.3.3 Chemical types in Bryophyte and Pteridophyte cutins 2.3.4 Ontogeny, composition and structure of the CL 2.3.5 Layering of the CL 2.3.6 Cutin cystoliths 2.4 Cuticle structural types 2.4.1 Cuticle types 1 and 2 2.4.2 Cuticle type 3 2.4.3 Cuticle type 4 2.4.4 Cuticle types 5 and 6 2.4.5 A seventh cuticle type? 2.5 Summary of the cuticle types 2.5.1 Cuticle structure/ecology 2.5.2 Cuticle thickness and environment 2.5.3 Cuticle structure and phylogeny 2.6 The epicuticular wax 2.6.1 Epicuticular wax types 2.6.2 Chemical and structural classification of EW 2.6.2.1 Granules 2.6.2.2 Filaments 2.6.2.3 Plates 2.6.2.4 Tube-type waxes 2.6.2.5 Rodlet-type waxes 2.6.3 The background EW film 2.7 Cuticular pores and permeability of the CM 2.7.1 Permeability of water and solutes 2.7.2 Wax secretion, cuticular pores and microchannels 2.7.3 Relative sizes of wax crystals and cuticle 2.8 Crystallisation studies on EW 2.8.1 The tube wax crystal 2.8.2 The single-compound hypothesis 2.8.3 How do wax crystals grow? 2.9 Crystal orientation and spatial patterning 2.10 Degradation of EW 2.11 Summary of cuticle ontogeny Acknowledgements References

40 41 45 49 49 49 50 51 52 53 54 54 54 55 56 57 71 75 75 75 79 86 88 91 91 92 94 95 97 100 101 102 104 107 110 110

The cutin biopolymer matrix RUTH E. STARK and SHIYING TIAN 3.1 Introduction: protective plant polymers 3.2 Biosynthesis 3.3 Monomer composition 3.4 Polymeric structure of intact cutin

126 126 127 127 129

CONTENTS

3.5

Molecular structure of cutin fragments 3.5.1 Oligomeric degradation products 3.5.2 Polymeric residues from chemical degradation procedures 3.6 Mechanical properties 3.6.1 Methodology 3.6.2 Measurements of surface elastic modulus 3.6.3 Measurements of bulk molecular dynamics 3.7 Thermodynamic properties 3.8 Summary and prospectus Acknowledgements References

4

5

Composition of plant cuticular waxes REINHARD JETTER, LJERKA KUNST and A. LACEY SAMUELS 4.1 Methods used for the chemical analysis of plant cuticular waxes 4.1.1 Wax extraction 4.1.2 Instrumental analysis 4.2 Chemical profiles of plant cuticular waxes 4.2.1 Ubiquitous constituents of cuticular waxes 4.2.2 Taxon-specific major constituents of cuticular waxes 4.2.3 Cuticular triterpenoids 4.2.4 Other minor constituents of cuticular waxes 4.2.5 Other compounds located at or near the plant surface 4.3 Spontaneous reactions of cuticular wax constituents 4.4 Quantitative composition of cuticular waxes 4.4.1 Chain length distributions within compound classes 4.4.2 Distribution of compound classes 4.4.3 Wax amounts 4.5 Dynamics of wax composition 4.6 Arrangement of plant cuticular waxes 4.6.1 Formation of epicuticular crystals 4.6.2 Chemical differences between epicuticular film and intracuticular wax 4.6.3 Crystalline arrangement of epi- and intracuticular wax molecules References Biosynthesis and transport of plant cuticular waxes LJERKA KUNST, REINHARD JETTER and A. LACEY SAMUELS 5.1 Introduction 5.2 Synthesis of very long-chain fatty acid wax precursors

vii 131 131 134 135 135 137 138 140 141 141 141 145

145 145 147 150 150 151 155 157 159 160 162 162 163 165 165 168 168 171 174 175 182 182 182

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CONTENTS

5.3

Biosynthetic pathways to monofunctional aliphatics 5.3.1 Synthesis of primary alcohols and wax esters 5.3.2 Synthesis of alkanes, secondary alcohols and ketones 5.3.3 Biosynthesis of β-diketones and alkan-2-ols 5.4 Triterpenoid biosynthesis 5.5 Regulation of wax biosynthesis 5.6 Wax biosynthesis and transport in the context of the epidermal cell 5.6.1 Intracellular sites of wax synthesis and trafficking of wax constituents 5.6.2 Transport of wax through the PM 5.6.3 Transport of wax through the cell wall to the cuticle 5.7 Concluding remarks References

6

Optical properties of plant surfaces ERHARD E. PFÜNDEL, GIOVANNI AGATI and ZORAN G. CEROVIC 6.1 Introduction 6.2 Methods to determine optical properties of plant surfaces 6.2.1 UV-visible absorbance spectrophotometry 6.2.2 Integrating sphere 6.2.3 Microfibre optics 6.2.4 UV-excited chlorophyll fluorescence 6.2.5 Fluorescence microscopy 6.2.6 Auto-fluorescence and reagent-induced fluorescence 6.3 Electronic absorption of radiation 6.3.1 Phenolics 4-Hydroxycinnamic acids Colourless flavonoids Anthocyanins 6.3.2 Betalains 6.3.3 Carotenoids 6.4 Non-absorptive optical properties 6.4.1 Reflectance Glabrous surfaces Glaucous surfaces Trichomes Water and salt 6.4.2 Optical effects of surface architecture Lens effect The sieve effect Structural colour

189 189 191 195 196 199 200 201 205 206 207 207 216 216 217 217 217 218 220 220 222 223 224 224 226 227 229 229 230 230 231 232 234 235 235 235 236 237

CONTENTS

7

8

ix

6.5 Concluding remarks Acknowledgements References

238 238 239

Transport of lipophilic non-electrolytes across the cuticle MARKUS RIEDERER and ADRIAN FRIEDMANN 7.1 Introduction 7.2 How do LNE enter the cuticle? 7.2.1 Sources 7.2.2 Points of entry 7.2.3 Partitioning 7.3 Transport through the cuticle 7.3.1 Laws of mass transfer 7.3.2 Measuring transport in cuticular fractions 7.3.3 Analysis of transport properties 7.3.4 Prediction of transport properties 7.3.5 Cuticular permeability in perspective 7.4 Enhancing the transport properties of the cuticle 7.4.1 Modes of action of uptake enhancers 7.4.2 Transport enhancing compounds 7.5 Conclusions References

250

Characterisation of polar paths of transport in plant cuticles LUKAS SCHREIBER 8.1 Introduction 8.1.1 Penetration of non-ionic, lipophilic molecules across cuticles 8.1.2 Penetration of polar and charged molecules across cuticles 8.1.3 Effect of air humidity on cuticular barrier properties 8.2 Results providing evidence for polar paths of transport 8.2.1 Factors influencing permeability of inorganic ions and charged molecules across plant cuticles 8.2.2 Ion permeability in comparison to permeability of uncharged molecules 8.3 Significance of polar paths of transport 8.3.1 Chemical nature of polar domains in cuticles 8.3.2 Relevance of polar paths of transport in cuticles References

250 250 251 252 254 258 258 259 262 265 266 267 267 269 272 273 280 280 280 281 281 281 283 284 286 287 289 289

x

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CONTENTS

Cuticular transpiration MARKUS BURGHARDT and MARKUS RIEDERER 9.1 Introduction 9.1.1 Definition of transport parameters 9.1.2 Experimental methods 9.2 Mechanisms of water transport through the cuticle 9.2.1 The lipophilic pathway 9.2.2 The hydrophilic pathway 9.2.3 Relationship between cuticular transpiration and cuticle structure and composition 9.3 Environmental effects on transpiration 9.3.1 Relative humidity 9.3.2 Temperature 9.4 Physiology of cuticular transpiration in relation to stomatal closure 9.5 The cuticular transpiration barrier as a mechanism of the drought avoidance strategy 9.6 Conclusions References

10 The cuticle and cellular interactions HIROKAZU TANAKA and YASUNORI MACHIDA 10.1 Introduction 10.2 Essential roles of the cuticle in post-embryonic development 10.2.1 The cuticle is required for prevention of organ fusion 10.2.2 Genetic screening for mutants with defective cuticles 10.2.3 Molecular identification of genes involved in the generation of a functional cuticle 10.3 Functions of the cuticle in plant reproduction and embryogenesis 10.3.1 The cuticle as an interface in stigma–pollen interactions 10.3.2 Role of the cuticle during embryogenesis 10.4 Regulators of epidermal differentiation and cuticle formation 10.4.1 Intercellular signalling and cuticle formation 10.4.2 Transcriptional control of cuticle production 10.5 Concluding remarks References

292 292 292 294 295 295 297 299 301 301 302 303 306 307 309 312 312 312 312 314 318 322 322 325 326 326 327 329 329

CONTENTS

11 Microbial communities in the phyllosphere JOHAN H.J. LEVEAU 11.1 Introduction 11.2 Methodologies in phyllosphere microbiology 11.2.1 Sampling techniques 11.2.2 Artificial inoculation 11.2.3 Microscopy 11.3 Getting to the phyllosphere (and leaving again) 11.3.1 Immigration 11.3.2 Adhesion 11.3.3 Emigration 11.4 Microbial communities in the phyllosphere 11.4.1 Composition 11.4.2 Abundance 11.4.3 Dynamics 11.5 Microbial perception of the phyllosphere 11.5.1 Topography 11.5.2 Physico-chemical parameters 11.5.3 Biological environment 11.6 Surviving (or not) in the phyllosphere 11.6.1 Concept of epiphitness 11.6.2 Adaptive strategies 11.6.3 Epiphitness genes 11.7 Microbial growth in the phyllosphere 11.7.1 Growth requirements 11.7.2 Types and sources of nutrients 11.7.3 Nutrient bioavailability 11.8 Microbial interactivities in the phyllosphere 11.8.1 Niche modification 11.8.2 Competition 11.8.3 Antibiosis 11.8.4 Communication 11.8.5 Gene exchange 11.9 Biocontrol in the phyllosphere 11.9.1 Phyllosphere diseases 11.9.2 What makes a plant pathogen? 11.9.3 Strategies for biocontrol 11.10 Future directions of phyllosphere microbiology References

12 Filamentous fungi on plant surfaces TIM L.W. CARVER and SARAH J. GURR 12.1 Introduction 12.2 Adhesion prior to germination

xi 334 334 334 334 338 339 341 341 342 344 344 344 345 347 347 347 348 349 349 349 350 351 352 352 353 353 354 354 355 355 355 356 356 356 357 357 358 359 368 368 370

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CONTENTS

12.2.1 The environment, leaf surface characteristics and spore shape 12.2.2 The challenges and nature of spore adhesion 12.2.3 Examples of passive and active adhesion and combinations of the two 12.2.4 Plant surface versus artificial substrata 12.2.5 Effects of ungerminated conidia on underlying host cells 12.3 Influence of leaf surface characteristics on spore germination 12.4 Directional emergence of fungal germ tubes 12.5 The special case of the PGT of B. graminis 12.6 AGT growth, appressorium differentiation and penetration 12.6.1 Adhesives associated with AGTs 12.6.1.1 Germ tube growth and appressorium differentiation by fungi that penetrate the host surface directly 12.6.1.2 Signal transduction in fungi that penetrate the host surface directly 12.6.1.3 Germ tube growth and appressorium differentiation by fungi that enter via stomata 12.7 Entry into the host leaf 12.7.1 Direct penetration of the host surface from appressoria 12.7.2 Entry via stomata 12.8 Conclusions Acknowledgements References

13 Plant–insect interactions on cuticular surfaces CAROLINE MÜLLER 13.1 Introduction 13.2 Access to the plant surface 13.2.1 Impeding attachment from the plant perspective 13.2.2 Attachment from the insect perspective 13.3 Recognition cues for insects 13.3.1 Deterrent properties 13.3.2 Attractive properties 13.4 Mimicry 13.4.1 Mimicry from the plant perspective 13.4.2 Mimicry from the insect perspective

370 371 372 374 375 376 377 379 382 383

384 386

387 389 389 391 391 392 392 398 398 399 399 400 400 401 410 412 413 413

CONTENTS

13.5 Methods of investigation 13.6 Application in biological pest management 13.7 Conclusion References Index Colour plates appear after p. 249

xiii 414 415 416 417 423

Contributors

Dr Giovanni Agati Instituto di Fisica Applicata ‘Nello Carrara’ – CNR, via Madonna del Piano, 50019 Sesto Fiorentino, Firenze, Italy, [email protected] Dr Markus Burghardt Julius-von-Sachs-Institut für Biowissenschaften, Universität Würzburg, Julius-von-Sachs-Platz 3, 97082 Würzburg, Germany, [email protected] Dr Tim L.W. Carver Plant Genetics and Breeding (PGB), IGER, Plas Gogerddan, Aberystwyth SY23 3EB, UK, [email protected] Dr Zoran G. Cerovic LURE-CNRS, BP34, Bâtiment 203, 91898 Orsay, France, [email protected] Dr Adrian Friedmann Syngenta Crop Protection AG, WMU-3120.2.06, Im Breitenloh 5, 4333 Münchwilen, Switzerland, [email protected] Professor Sarah J. Gurr Plant Sciences, University of Oxford, South Parks Road, Oxford OX1 3RB, UK, [email protected] Dr Christopher E. Jeffree Science Faculty Electron Microscope Facility, Daniel Rutherford Building, King’s Buildings, Mayfield Road, Edinburgh EH9 3JH, UK, [email protected] Professor Reinhard Jetter Departments of Botany and Chemistry, University of British Columbia, 3510-6270 University Blvd, Vancouver, BC V6T 1Z4, Canada, [email protected] Professor Ljerka Kunst Department of Botany, University of British Columbia, 6270 University Blvd, Vancouver, BC V6T 1Z4, Canada, [email protected] Dr Johan H.J. Leveau NIOO-KNAW, Centre for Terrestrial Ecology, P.O. Box 40, 6666 ZG Heteren, The Netherlands, [email protected] Dr Yasunori Machida Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan, [email protected] Dr Caroline Müller Julius-von-Sachs-Institut für Biowissenschaften, Universität Würzburg, Julius-von-Sachs-Platz 3, 97082 Würzburg, Germany, [email protected] Dr Erhard E. Pfündel Julius-von-Sachs-Institut für Biowissenschaften, Universität Würzburg, Julius-von-Sachs-Platz 3, 97082 Würzburg, Germany, [email protected] Professor Markus Riederer Julius-von-Sachs-Institut für Biowissenschaften, Universität Würzburg, Julius-von-Sachs-Platz 3, 97082 Würzburg, Germany, [email protected] Professor A. Lacey Samuels Department of Botany, University of British Columbia, 6270 University Blvd, Vancouver, BC V6T 1Z4, Canada

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CONTRIBUTORS

Professor Lukas Schreiber Ökophysiologie der Pflanzen, Botanisches Institut, Universität Bonn, Kirschallee 1, 53115 Bonn, Germany, [email protected] Professor Ruth E. Stark Department of Chemistry and Institute for Macromolecular Assemblies, City University of New York, College of Staten Island, 2800 Victory Boulevard, Staten Island, NY 10314-6600, USA, [email protected] Dr Hirokazu Tanaka College of Bioscience and Biotechnology, Chubu University, 120 Matsumoto-cho, Kasugai, Aichi 487-8501, Japan, [email protected] Dr Shiying Tian Department of Chemistry and Institute for Macromolecular Assemblies, City University of New York, College of Staten Island, 2800 Victory Boulevard, Staten Island, NY 10314-6600, USA, [email protected]

Preface During recent years the science of plant surfaces, and the cuticle in particular, has advanced at a fast pace, encompassing new fields of study along the way. Considerable progress has been made possible by the application of new concepts and techniques for investigating the biosynthesis, composition, structure and functional complexity of the plant cuticle. We now have an increased understanding of the microscopic and submicroscopic fine structure of the cuticular membrane as a whole, as well as of the cutin matrix and the associated wax deposits. By employing mutants and applying molecular biological techniques and advanced analytical tools, a much clearer image can now be drawn of the composition of cuticular waxes and the biosynthetic pathways leading to them. Intriguing variations can be found in the cuticular chemistry, morphology and function between and within plant species. Studies assessing the impact of UV radiation on plant life have emphasised the role of the cuticle and the underlying epidermis as optical filters for solar radiation. The field concerned with the diffusive transport of lipophilic organic non-electrolytes across the plant cuticle has reached a state of maturity, which makes it possible to quantitatively analyse and predict permeabilities based on physico-chemical predictors and to manipulate them in vivo. Recently, a new paradigm has been proposed for the diffusion of polar compounds and water across the cuticle. Within the context of plant ecophysiology, cuticular transpiration can now be considered in the perspective of whole-leaf water relations. New and unexpected roles have been assigned to the cuticle in plant development and in pollen–stigma interactions. Finally, much progress has been made in understanding the cuticle as a specific and extraordinary substrate for the interactions of the plant with microorganisms, fungi and insects. Since the early 1970s, three books on the plant cuticle have been published. Only the first addressed all aspects of the subject; the other two were multiauthor volumes arising from scientific meetings. Considering the progress made in this field, a book which deals comprehensively with plant surface characteristics and functions is overdue. The title, Biology of the Plant Cuticle, is intended to express the multidisciplinary and integrative approach to the subject. As functions are interconnected and rely heavily on the (bio)chemistry and properties of the cuticle, it is hoped that bringing together thus far disparate views of the subject will substantially advance the field of plant surface science. The book is also intended to provide a comprehensive overview and critical discussion of the current state of knowledge, paying close attention to the applied aspects of the field wherever appropriate. Biology of the Plant Cuticle is aimed at a broad audience, ranging from biologists working on the molecular and whole-organism level to industrial agrochemists.

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PREFACE

It is hoped that it will be of interest to phytochemists, plant (eco)physiologists, ecologists and environmental scientists, as well as to scientists and practitioners from the agricultural and horticultural sciences. In comparison with its predecessors, this book extensively considers the biological interactions occurring on plant surfaces and, therefore, is hoped to be of special appeal to scientists who, in the past, did not consider the plant surface a priori as a subject of prime importance. Thus, this volume is furthermore directed at phytopathologists, environmental microbiologists, entomologists and chemical ecologists. The editors are indebted to the chapter authors for an enjoyable collaboration on this project and for timely delivery of carefully prepared manuscripts. In addition, the editors gratefully acknowledge the encouragement, advice and support continuously provided by Graeme MacKintosh and David McDade of Blackwell Publishing. Markus Riederer Caroline Müller

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

1

Introduction: biology of the plant cuticle Markus Riederer

‘Does it make sense, and is it fun at all, to spend so much time with the outermost micrometer of a plant?’ This was the question a member of a search committee asked when the author applied for a job at a German university. As all scientists in this field know and deeply feel, it is fun indeed to study the plant cuticle and the plethora of processes related to it. The authors of this book hope that the reader will come to the conclusion that it is worthwhile to invest time, brains and funds into this endeavour. The cuticle has often been called the ‘skin’ of the primary parts of higher plants, and in fact, the Latin word from which this term is derived (cuticula) means ‘thin skin’. The term cuticle has undergone a kind of evolution and profound changes in meaning during the last two centuries. At the beginning, the whole primary integument tissue or epidermis of a plant was called ‘cuticle’ stressing the convergence with animal skin, which is also cellular in nature. The modern usage of the word, meaning ‘a superficial film formed of the cutinized outer layers of the superficial walls of the epidermal cells’ (Oxford English Dictionary Online) of a plant, goes back to A.P. de Candolle. In 1827, he restricted the use of the French term ‘cuticule’ to the meaning in which it is used today (Wagenitz, 1996). Thus, the word cuticle is no longer used for a cellular layer but for a continuous extracellular membrane. In 1852, the word appeared in English for the first time in Henfrey’s translation of H. von Mohl’s ‘Grundzüge der Anatomie und Physiologie der vegetabilischen Zelle’ (OED Online). It is this term which will accompany us throughout this book.

1.1 The evolution of the plant cuticle The cuticle as a structure has a very long history on the palaeobiological timescale. It is fortunate that the cuticle is a highly recalcitrant material which can easily resist decay for millions of years under favourable deposition conditions. It is fascinating that major chemical features like cutin composition are preserved over such prolonged periods of time (Ewbank et al., 1996; Edwards et al., 1997). Thus, essentially intact cuticles with clearly delineated epidermis cell silhouettes can be obtained from old sediments. The oldest remnants of plant cuticles date back to the boundary between the late Siluarian and the early Devonian (about 400 million years ago) periods. The earliest cuticles, in the modern sense of the term which is assigned to higher plants, were found dispersed in sediments and belong to sporangia of rhyniophytoids. These specimens lack the impression of stomata while beginning with the

2

BIOLOGY OF THE PLANT CUTICLE

basal Devonian period, preserved cuticles show imprints from guard and accessory cells that are comparable to the modern stomatal apparatus. For a recent review on this subject see Edwards et al. (1996). The finding that cuticles and stomata appear concomitantly in early kormophytes has profound impact on modern concepts of the evolution of vascular land plants and also on the interpretation of the selection pressure which acted on the evolution of stomata (Raven, 1977, 2002). Palaeoecophysiology has interpreted the simultaneous appearance of cuticles and stomata as evidence for the physiological adaptations to the colonisation of the land and thus for the relatively dry atmosphere by basal precursors of modern higher plants. Cuticles and stomata form a syndrome together with extended root systems, supracellular transport in vascular structures and the development of intercellular air spaces (Raven, 1977, 2002). These features are interpreted as necessary adaptations for photosynthesising homoiohydric life forms in an atmosphere with low water activity. The early cuticle probably also had additional functions equivalent to those of modern cuticles with defence against parasites, protection against ultraviolet (UV) radiation and water repellence being the most important ones.

1.2 Major functions of the plant cuticle The cuticle is a structure that incorporates numerous functions of essential importance for plant life (Kerstiens, 1996b). This book treats the major functions in detail and, in most cases, devotes separate chapters to each of them. Nevertheless, a short synopsis is included in this introduction because it appears necessary to make one point very clear: the cuticle is a non-living though highly multifunctional structure into which numerous functions have been integrated. As will be shown later, this integration is sometimes not ideal as some physiological demands are in conflict with each other.

1.2.1

Transpiration control

As mentioned earlier, one of the major exigencies of the terrestrial lifestyle of higher plants is to have control over water relations. In order to stay alive, which essentially means to be more or less turgescent, the plant has to maintain the equilibrium between transpirational water loss and root water uptake. Any pronounced disequilibrium will severely compromise the viability and thus the fitness of the plant. The control of transpiration from leaves, primary stems, flowers and fruits has two components: the stomata and the cuticle. Depending on the primary focus of scientific interest, the importance of either the stomata or the cuticle will be stressed by different authors. However, an effective control of transpiration is feasible only if the stomata and the cuticle act together in an optimised way. The low permeability of the cuticle makes it possible to control water loss by adjusting stomatal aperture.

INTRODUCTION

3

But control will only work in a satisfactory way if the water loss across the cuticular surface is lower than the residual water loss through stomatal pores at optimal closure of stomata. The cuticular permeability for water and transpiration confinement by the plant cuticle will be treated extensively in Chapters 8 and 9. This subject, of course, has met the interest of many researchers in the past. For general and early literature, the reader is referred to Stålfelt (1956), Schönherr (1982), and the textbooks by Larcher (2003) and Nobel (1991).

1.2.2

Control of loss and uptake of polar solutes

In principle, all organisms must have control over their inner milieu and therefore must have resistant integuments separating them from the environment. This is also true for terrestrial plants which would loose ions and polar organic solutes from the apoplastic solution unless they have a highly resistant cuticle that impedes the transport from the interior to the environment. Thus, the plant gains control over the loss of solutes and, at the same time, may modulate it by salt-excreting glands or hydathodes according to its specific needs. As the transport across the cuticle is symmetric, this membrane also hinders the uptake of polar substances from the outside. Control over solute loss and uptake is exerted by the same barrier properties of the cuticle as transpiration control. It might be speculated whether the need for controlling solute loss was an additional driving force in the evolution of the cuticular diffusion barrier. A new view of the cuticular permeability of polar substances is currently evolving. There is increasing evidence that ions and small polar solutes move across the cuticle via continuous polar pathways that bypass the wax-based cuticular transport barrier. Chapter 8 (and partially also Chapter 9) will present this new view of polar solute (and water) transport across the cuticle and will put it in perspective with older work implying that such pathways may exist. For a thorough review of the older literature and for a primarily horticultural point of view on the subject of solute loss from plants (leaching), refer to the review by Tukey (1970).

1.2.3

Controlling the exchange of gases and vapours

When stomata are closed (which, on the average, is the case for approximately 12 h a day), the cuticle completely limits the loss and uptake of gases and vapours across the plant–atmosphere interface. This is true not only for water vapour as treated earlier but also for gases like carbon dioxide, oxygen, inorganic air pollutants and volatile organic compounds like terpenes (Lendzian and Kerstiens, 1991; Kerstiens et al., 1992; Kerstiens, 1994). For highly lipophilic organic vapours, the cuticle is the preferred pathway of exchange even under conditions when the stomata are open (Riederer, 1995; Trapp, 1995). Exerting control over gas and vapour fluxes is, without any doubt, beneficial to the plant in most cases.

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BIOLOGY OF THE PLANT CUTICLE

However, there is a conflict between controlling volatile exchange and photosynthesis. It has been shown experimentally in intact leaves with artificially clogged stomata that while the cuticle allows small amounts of carbon dioxide and water vapour to pass through, it markedly discriminates against the transport of carbon dioxide (Boyer et al., 1997). A comparison with the properties of synthetic polymeric membranes like polyethylene, polycarbonate or polyester makes this property of the plant cuticle understandable. Woolley (1967) compared the permeabilities of plastic films to water and carbon dioxide and found that no synthetic material in existence has a higher permeability for carbon dioxide than for water. We can therefore conclude that intrinsic properties of a transport barrier against water and polar solutes confer low permeabilities to carbon dioxide and many other inorganic gases (Langowski, 2002). Evolution over the past 400 million years does not seem to have generated a membrane that can escape these physical constraints. This is the case even though a strong selective pressure acts towards a cuticle that allows photosynthesis during the light period but with the stomata closed.

1.2.4

Transport of lipophilic substances

The cuticle is the main aboveground interface for the exchange of lipophilic organic compounds between the environment and the interior of primary plant parts. All lipophilic compounds with low volatility or in solution have to cross the cuticle in order to enter or leave fruits, primary stems or leaves. The stomatal pathway is either not open to them (aqueous solutions of organic compounds) or is a very restricted route of exchange (semi-volatile compounds). The organic compounds in question may either be secondary metabolites of the plant or natural as well as anthropogenic compounds (pollutants, plant protection agents) occurring in the environment. From an applied point of view, the sorption and uptake of plant protection agents is of prime importance. Both the basic and applied aspects of this topic are discussed in Chapter 8. For publications covering the older literature, see these reviews and books: Van Overbeek (1956), Currier and Dybing (1959), Foy (1964), Sargent (1965), Bukovac (1976) and Hartley and Graham-Bryce (1980).

1.2.5

Water and particle repellence

After rains, many leaf surfaces are not covered by films of water and thus rapidly dry up. The cuticular surfaces of many plant species, at least their younger and pristine parts, are repellent to water and most water-based solutions. This is advantageous as water on the leaf surface may have several negative consequences for the plant; it (1) leads to leaching of ions and polar organic solutes from the plant’s interior, and (2) creates suitable conditions for the colonisation by potentially harmful microbes like phytopathogenic bacteria or parasitic fungi. The latter aspect will be covered in Chapters 11 and 12 where the current knowledge on microbial communities and filamentous fungi on plant surfaces will be discussed in detail.

INTRODUCTION

5

Certain plant surfaces may not only repel water and aqueous solutions but also microscopic particles like particulate aerosol, dust, spores and microbes. This is due to a self-cleaning mechanism based on the physico-chemical properties of some leaf surfaces and water droplets running off the surface taking along particles. This phenomenon has been termed Lotus effect and industrial applications have been explored (Barthlott and Neinhuis, 1997; Wagner et al., 2003; Otten and Herminghaus, 2004). The fine structure of the cuticle and the chemical composition of cutin and cuticular waxes are covered in Chapters 2, 3 and 4, respectively.

1.2.6

Attenuation of photosynthetically active and UV radiation

One of the main driving forces for the colonisation of the terrestrial environment by plants is the luxuriant availability of radiation in the wavelength range from 400 to 800 nm in most cases. However, photosynthesis depends on a highly complicated and sensitive arrangement of pigments, proteins and membrane-enclosed compartments. This complex is easily damaged by excessive light. One of the protective mechanisms involves the cuticle: a dense cover of epicuticular wax crystals enhances scattering and reflection to a degree making tolerable the intensity of the radiation which reaches the photosynthetically active tissues in the interior of the leaf. Another part of the electromagnetic spectrum hitting plant surfaces is UV radiation in the wavelength range from 280 to 400 nm. Excessive irradiation by UV results in damages in the photosynthetic apparatus and other vital parts of the plant cell. The cuticle, often together with the outer epidermal cell wall and the vacuoles of the epidermis, can contribute to an effective screening of UV radiation and thus to protecting the sensitive inner tissues. Chapter 6 covers the optical properties in the visible and UV range of the cuticle but also looks at properties of the epidermis and distinct sub-epidermal layers.

1.2.7

Mechanical containment

In a limited number of cases, the mechanical properties of plant cuticles support other structures like cell walls in maintaining the structural integrity of plant tissues. An economically important example for the mechanical importance of cuticles is fruit cracking in tomato and sweet cherry. In both cases, increasing internal pressure by uptake of water via roots or the fruit surface leads to the development of cracks. These cracks severely interfere with the economic and nutritional value of the fruits. Several studies have been performed on this issue either from an applied horticultural (Emmons and Scott, 1997; Bukovac et al., 1999; Knoche et al., 2002) or from a biomechanical (Wiedemann and Neinhuis, 1998; Matas et al., 2004) point of view.

1.2.8

Separating agent in plant development

The cuticle plays a crucial role in plant development also and may be compared to a separating agent in developmental processes. Mutants with defective cuticles

6

BIOLOGY OF THE PLANT CUTICLE

exhibit increased water loss and, at the same time, extraordinary morphological abnormalities such as the fusion of organs. The emerging knowledge on the role played by the cuticle in cellular interactions and plant morphogenesis is extensively covered in Chapter 10.

1.2.9

Interface for biotic interactions

The cuticle-covered surface of higher plants is the main locality for major aboveground interactions with small organisms. On a microscopic scale, it is the interaction of bacteria, yeasts and fungi with the plant that may profoundly be influenced by cuticular properties. Features of the cuticle may have effects on adhesion, host recognition and mineral and carbon nutrition of the microbes as well as on the availability of liquid water. In addition, the cuticle may provide mechanical protection against the invasion by microbes. Cuticle–microbe interactions are treated in Chapters 11 and 12. For extensive reviews on these matters refer to Blakeman (1981, 1982, 1993) and Beattie and Lindow (1995). On the macroscopic scale, the cuticle may interfere when insects or other arthropods interact with leaf surfaces. This may happen when a herbivore is searching for a suitable host for food or oviposition. Numerous cases have been reported where cuticular and leaf surface features in general influence herbivore behaviour and thus indirectly the integrity and fitness of the plant. This subject is reviewed in Chapter 13.

1.3 Convergence with other integuments Not only plants but many other terrestrial organisms face at least some of the problems listed earlier. Very often, the main challenge is the danger of desiccation due to living in a dry atmosphere. The long-term maintenance of water balance must be solved by any terrestrial organism irrespective of its habitat. In order to hold back the water obtained from their surroundings, animals like plants typically possess an outer integument that greatly reduces the rate of water loss (Hadley, 1981, 1991). In many species, the outer layers of the integument are covered and/or impregnated with more or less solid lipids which are primarily responsible for the observed waterproofing properties. This is especially true for plants and arthropods (insects and arachnids) both of which have a lipophilic matrix (cutin in plants, epicuticle in arthropods) with associated waxes. In both cases, these lipids are mixtures of long-chain aliphatic compounds as described for plants in detail in Chapters 3 and 4 and for insects in several reviews (Blomquist et al., 1987; de Renobales et al., 1991; Nelson and Blomquist, 1995). The cuticles of both arthropods and plants are continuous non-cellular membranes with multiple layers which cover the epidermis. In arthropods and in higher plants alike it is the physical structure, arrangement and composition of cuticular lipids that determines the waterproofing quality of the integument. Quantity and composition are species and age specific. For further

INTRODUCTION

7

details on the convergence of plant and arthropod waterproofing properties and their relationship to chemical composition and physical structure of the cuticle, see the reviews by Hadley (1981, 1989, 1991). This parallelism in the chemical, structural and physical properties of the outer layers of the integument is an outstanding example of convergent evolution in two widely divergent groups of organisms. The evolutionary success of wax-based transpiration barriers is extraordinary: if we take conservative species estimates as given by Wilson (1988, 1992), approximately 80% of all species on earth share a cuticlelike integument with low water permeability achieved by associated waxes. It may be speculated that two properties of mixtures of long-chain aliphatic molecules are responsible for this extraordinary evolutionary success: (1) waxes are plastic and can therefore follow growth and movements, and (2) they are multi-component, partly liquid or amorphous solids having self-healing properties which allow closing small defects inflicted on the integuments of plants or arthropods.

1.4 Objectives of this book The science of the plant cuticle has received increasing attention among plant scientists as hitherto unknown functions and properties of this fraction of the epidermis have been discovered. Palaeobiologists, ecologists and especially plant evolutionary biologists become increasingly interested in cuticular remains and what can be deduced from their occurrence and structure. The volume of literature on the plant cuticle is growing at an increasing rate. A search in the BIOSIS database shows that during the last ten years (that is the time interval since the last book on the cuticle has appeared), approximately 2300 publications concerning the cuticles of plants have appeared. The overall subject of this book has been treated in the past in several books. To the author’s knowledge, the first modern experiment-based and comprehensive treatment of the cuticle and the associated waxes was provided by Frey-Wyssling (1938). Twenty years later, Martin and Juniper (1970) published a book which was the first to be exclusively devoted to the cuticles of plants. Thereafter, two volumes each compiling the proceedings of meetings devoted to the plant cuticle were edited by Cutler et al. (1982) and by Kerstiens (1996a), respectively. A small booklet on the surfaces of plants oriented at a general scientific audience was published by Juniper and Jeffree (1983). Considering the progress made in this field since then, a new book covering the whole field of cuticular science appears overdue. Since Martin and Juniper (1970), the present book is the first written exclusively for this purpose and not derived from a scientific meeting. The title Biology of the Plant Cuticle has been chosen in order to express the multidisciplinary and integrative views of the subject. Cuticular functions are interrelated and heavily rely on the (bio)chemistry and the physical properties of the cuticle. Therefore, combining so far disparate views on this subject into a common perspective is expected to advance the field of plant surface

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BIOLOGY OF THE PLANT CUTICLE

science substantially. Bringing the different functions together will delineate where and under which conditions they are in accordance and in conflict with each other depending on the special needs of the plant. Obviously, the book is intended to provide a comprehensive and critical treatment of the current state of knowledge about plant surfaces and the cuticle in particular in its full depth and breadth. Recent developments having a pronounced impact on our understanding of the cuticle’s fine structure, biosynthesis, composition, physical and transport properties are extensively reviewed in this book. For the first time, a comprehensive overview of cuticular functions in plant morphogenesis is given. Part of the book is devoted to the rapidly evolving field of biotic interactions taking place on plant surfaces. Where appropriate, special attention has been paid to the applied aspects of the field, especially in agricultural chemistry.

References Barthlott, W. and Neinhuis, C. (1997) Purity of the sacred lotus, or escape from contamination in biological surfaces, Planta, 202, 1–8. Beattie, G.A. and Lindow, S.E. (1995) The secret life of foliar bacterial pathogens on leaves, Annual Reviews of Phytopathology, 33, 145–172. Blakeman, J.P. (1981) Microbial Ecology of the Phylloplane, Academic Press, London. Blakeman, J.P. (1982) Phylloplane interactions, in Phytopathogenic Prokaryotes (eds Z. Klement, M.S. Mount and G.H. Lacy), Academic Press, London, New York, pp. 307–333. Blakeman, J.P. (1993) Pathogens in the foliar environment, Plant Pathology, 42, 479–493. Blomquist, G.J., Nelson, D.R. and de Renobales, M. (1987) Chemistry, biochemistry, and physiology of insect cuticular lipids, Archives of Insect Biochemistry and Physiology, 6, 227–265. Boyer, J.S., Wong, S.C. and Farquhar, G.D. (1997) CO2 and water vapor exchange across leaf cuticle (epidermis) at various water potentials, Plant Physiology, 114, 185–191. Bukovac, M.J. (1976) Herbicide entry into plants, in Herbicides (ed. L.J. Audus), Academic Press, London, New York, San Francisco, pp. 335–364. Bukovac, M.J., Knoche, M., Pastor, A. and Fader, R.G. (1999) The cuticular membrane: a critical factor in rain-induced cracking of sweet cherry fruit, HortScience, 34, 549. Currier, H.B. and Dybing, D.C. (1959) Foliar penetration of herbicides – review and present status, Weeds, 7, 195–213. Cutler, D.F., Alvin, K.L. and Price, C.E. (1982) The Plant Cuticle, Academic Press, London. de Renobales, M., Nelson, D.R. and Blomquist, G.J. (1991) Cuticular lipids, in The Physiology of the Insect Epidermis (eds K. Binnigton and A. Retnakaran), CSIRO, pp. 240–251. Edwards, D., Abbott, G.D. and Raven, J.A. (1996) Cuticles of early land plants: a palaeoecophsiological evaluation, in Plant Cuticles: An Integrated Functional Approach (ed. G. Kerstiens), BIOS Scientific Publishers, Oxford, pp. 1–32. Edwards, D., Ewbank, G. and Abbott, G.D. (1997) Flash pyrolysis of the outer cortical tissues in Lower Devonian Psilophyton dawsonii, Botanical Journal of the Linnean Society, 124, 345–360. Emmons, C.L.W. and Scott, J.W. (1997) Environmental and physiological effects on cuticle cracking in tomato, Journal of the American Society for Horticultural Science, 122, 797–801. Ewbank, G., Edwards, D. and Abbott, G.D. (1996) Chemical characterization of lower Devonian vascular plants, Organic Geochemistry, 25, 461–473. Foy, C.L. (1964) Review of herbicide penetration through plant surfaces, Agricultural and Food Chemistry, 12, 473–476. Frey-Wyssling, A. (1938) Submikroskopische Morphologie des Protoplasmas und seiner Derivate, Gebrüder Borntraeger, Berlin.

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Hadley, N.F. (1981) Cuticular lipids of terrestrial plants and arthropods: a comparison of their structure, composition, and waterproofing function, Biological Reviews, 56, 23–47. Hadley, N.F. (1989) Lipid water barriers in biological systems, Progress in Lipid Research, 28, 1–33. Hadley, N.F. (1991) Integumental lipids of plants and animals – comparative function and biochemistry, Advances in Lipid Research, 24, 303–320. Hartley, G.S. and Graham-Bryce, I.J. (1980) Physical Principles of Pesticide Behaviour, Academic Press, London, New York, Toronto, Sydney, San Francisco. Juniper, B.E. and Jeffree, C.E. (1983) Plant Surfaces, Edward Arnold, London. Kerstiens, G. (1994) Air pollutants and plant cuticles: mechanisms of gas and water transport, and effects on water permeability, in Air Pollutants and the Leaf Cuticle (eds K.E. Percy, J.N. Cape, R. Jagels and C.J. Simpson), Springer, Berlin, pp. 39–53. Kerstiens, G. (1996a) Plant Cuticles – An Integrated Functional Approach, BIOS Scientific Publishers, Oxford. Kerstiens, G. (1996b) Signalling across the divide: a wider perspective of cuticular structure–function relationships, Trends in Plant Science, 1, 125–129. Kerstiens, G., Federholzner, R. and Lendzian, K.J. (1992) Dry deposition and cuticular uptake of pollutant gases, Agriculture Ecosystems and Environment, 42, 239–253. Knoche, M., Peschel, S. and Hinz, M. (2002) Studies on water transport through the sweet cherry fruit surface: III. Conductance of the cuticle in relation to fruit size, Physiologia Plantarum, 114, 414–421. Langowski, H.C. (2002) Flexible Materialien mit ultrahohen Barriereeigenschaften, Vakuum in Forschung und Praxis, 14, 297–302. Larcher, W. (2003) Physiological Plant Ecology, Springer-Verlag, Berlin. Lendzian, K.J. and Kerstiens, G. (1991) Sorption and transport of gases and vapors in plant cuticles, Reviews of Environmental Contamination and Toxicology, 121, 65–128. Martin, J.T. and Juniper, B.E. (1970) The Cuticles of Plants, Edward Arnold, London. Matas, A.J., Cobb, E.D., Bartsch, J.A., Paolillo, D.J. and Niklas, K.J. (2004) Biomechanics and anatomy of Lycopersicon esculentum fruit peels and enzyme-treated samples, American Journal of Botany, 91, 352–360. Nelson, D.R. and Blomquist, G.J. (1995) Insect waxes, in Waxes: Chemistry, Molecular Biology and Functions (ed. R.J. Hamilton), The Oily Press, West Ferry, pp. 1–90. Nobel, P.S. (1991) Physicochemical and Environmental Plant Physiology, Academic Press, San Diego. Oxford English Dictionary Online, entry cuticle, http://dictionary.oed.com/entrance.dtl, 2nd edn, 1989 with additions from 1992 and 1997, visited on 6 April 2005. Otten, A. and Herminghaus, S. (2004) How plants keep dry: a physicist’s point of view, Langmuir, 20, 2405–2408. Raven, J.A. (1977) The evolution of vascular land plants in relation to supracellular transport processes, Advances in Botanical Research, 5, 153–219. Raven, J.A. (2002) Selection pressures on stomatal evolution, New Phytologist, 153, 371–386. Riederer, M. (1995) Partitioning and transport of organic chemicals between the atmospheric environment and leaves, in Plant Contamination. Modeling and Simulation of Organic Chemical Processes, (eds S. Trapp and J.C. McFarlane), Lewis Publishers, Boca Raton, pp. 153–190. Sargent, J.A. (1965) The penetration of growth regulators into leaves, Annual Reviews of Plant Physiology, 16, 1–12. Schönherr, J. (1982) Resistance of plant surfaces to water loss: transport properties of cutin, suberin and associated lipids, in Physiological Plant Ecology (eds O.L. Lange, P.S. Nobel, C.B. Osmond and H. Ziegler), Springer-Verlag, Berlin, pp. 153–179. Stålfelt, M.G. (1956) Die cuticuläre Transpiration, in Handbuch der Pflanzenphysiologie, Bd. 3, Pflanze und Wasser (ed. W. Ruhland), Springer-Verlag, Berlin, Göttingen, Heidelberg, pp. 342–350. Trapp, S. (1995) Model for uptake of xenobiotics into plants, in Plant Contamination. Modeling and Simulation of Organic Chemical Processes (eds S. Trapp and J.C. McFarlane), Lewis Publishers, Boca Raton, pp. 107–151.

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Tukey, H.B. (1970) The leaching of substances from plants, Annual Reviews of Plant Physiology, 21, 305–324. Van Overbeek, J. (1956) Absorption and translocation of plant regulators, Annual Reviews of Plant Physiology, 7, 355–372. Wagenitz, D. (1996) Wörterbuch der Botanik, Gustav Fischer Verlag, Jena. Wagner, P., Furstner, R., Barthlott, W. and Neinhuis, C. (2003) Quantitative assessment to the structural basis of water repellency in natural and technical surfaces, Journal of Experimental Botany, 54, 1295–1303. Wiedemann, P. and Neinhuis, C. (1998) Biomechanics of isolated plant cuticles, Botanica Acta, 111, 28–34. Wilson, E.O. (1988) The current state of biological diversity, in Biodiversity (eds E.O. Wilson and F.M. Peter), National Academy Press, Washington, D.C., pp. 3–18. Wilson, E.O. (1992) The Diversity of Life, Allan Lane The Penguin Press, London. Woolley, J.T. (1967) Relative permeabilities of plastic films to water and carbon dioxide, Plant Physiology, 42, 641–643.

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

2

The fine structure of the plant cuticle Christopher E. Jeffree

2.1 Introduction 2.1.1

The distribution of the plant cuticle

The cuticle is a translucent film of polymeric lipids (Chapter 4) and soluble waxes (Chapter 5) located at the interface between a plant and its aerial environment. The cuticle is present on the outer surfaces of the epidermal cells at the aerial surfaces of vascular plants, and also on the sporophytes and sometimes the gametophytes of mosses, hornworts and liverworts. Only the epidermal cells of aerial organs are known to be capable of synthesising the constituents of the cuticle, and it is always absent from the root epidermis. The primary function of the cuticle is as a permeability barrier against water vapour loss from tissues (Schönherr and Mérida, 1981; Riederer, 1991; Schreiber et al., 1996), one of a system of innovations including stomata, intercellular spaces in a photosynthetic mesophyll and a vascular conducting system for water (xylem) and assimilates (phloem) that enable homoiohydry in land plants (Raven, 1977). The outer surface of the cuticle is coated with epicuticular waxes (EW) which confer water-repellency (Adam, 1963; Holloway, 1969a,b, 1970), keep the plant surface clean and dry (Barthlott and Neinhuis, 1997; Neinhuis and Barthlott, 1997), attenuate short-wave radiation (Chapter 6; Krauss et al., 1997) and discourage attachment of microorganisms (Chapters 11 and 12) and climbing by insects (Chapter 13; Eigenbrode, 1996) other than those with which a plant has symbiotic relationships (Markstädter et al., 2000). The cuticle may also have an important role in preventing developing organs from adhering to each other while developing in-bud (Chapter 10; Nawrath, 2002). On leaves the cuticle is present on both adaxial and abaxial surfaces, lines the stomatal apertures and covers the free inner epidermal cell surfaces of the substomatal cavity and intercellular spaces (Osborn and Taylor, 1990), but not the mesophyll. The cuticle is usually thickest above the anticlinal epidermal cell walls (CW), often forming pegs or spandrels by penetrating deeply between the anticlinal walls of adjacent epidermal cells (Figure 2.1b). The cuticular membrane (CM) thus bears an imprint of the epidermal cell pattern of the plant organs on which it was formed, which may survive as the only remaining fossil evidence of multicellular structure of the earliest land plants (Edwards et al., 1996). Epidermal cells of tomato fruits are also coated on internal and external surfaces, and intercellular cutinised layers may form as permeability barriers between stem and head cells in both absorptive and secretory glands (Mahlberg and Kim, 1991; Kim and Mahlberg, 1995).

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BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

Figure 2.1 Cuticular membranes with extended cuticular layer (CL) development. (a) The adaxial leaf cuticle of pear (Pyrus communis), stained en bloc with OsO4 , section stained with Reynold’s lead citrate. The cuticle proper (CP) is very electron-lucent and amorphous, and is terminated by a single outer electron-lucent lamella. Beneath the CP are two extensive CL zones, about 1 μm of chaotically lamellate material with no or few fibrillae (ECL), and an inner reticulate zone (ICL) with fewer lamellae. Note that in much of the ECL in this image, the lamellae run vertically. (b) The leaf cuticular membrane (CM) of lemon (Citrus limon) showing the peg between two adjacent epidermal cells. Holloway reported this CM as Type 4, reticulate throughout, but an amorphous CP covers the fibrils of the reticulate ECL (inset), making it Type 3. (c) The adaxial leaf cuticle of apple (Malus pumila) following isolation and methanol extraction, showing structure analogous to that in pear (see Figure 2.1a). The CP is not clearly demarcated from the ECL, but a single continuous very electron-lucent lamella terminates the CM. (d) The outer part of a stomatal complex still in the process of cell separation in a Phaseolus vulgaris leaf shows heavy cuticle development. Once again, a single electron-lucent lamella terminates the CM, and the ECL layer contains extensive lamellae (see inset). (a–c) Bars = 100 nm; (d) bar = 200 nm. Figure 2.1a reproduced from Holloway (1982a), Figure 2.1b–d by P.J. Holloway.

THE FINE STRUCTURE OF THE PLANT CUTICLE

13

A substomatal cuticle is unequivocally present on the surfaces of epidermal cells lining the substomatal cavity of many, if not all, species – in leaves of Gossypium hirsutum (Wullschleger and Oosterhuis, 1989), Malus pumila (Holloway, 1982a), Phaseolus vulgaris (Martin and Juniper, 1970) oak, Quercus velutina (Osborn and Taylor, 1990) and Cirsium horridulum (Pesacreta and Hasenstein, 1999). This carries the implication that the surfaces of epidermal cells do not evaporate significant amounts of water into the intercellular space system. However, an internal cuticle has also sometimes been reported on the surfaces of mesophyll and palisade cells, as in the mistletoe Phoradendron flavescens (Calvin, 1970). Seen in transmission electron microscope (TEM) sections, the internal cuticle may extend a short distance from the underside of the epidermal cell onto the surface of the palisade and spongy mesophyll cells, before tapering off to an amorphous, slightly osmiophilic layer on the mesophyll cell surfaces that has never been isolated or characterised chemically. Pure water beads with high contact angles can be condensed on the mesophyll cell surfaces indicating that they are hydrophobic (Sheriff, 1977; Willison and Pearce, 1983; Willison et al., 1984; Jeffree et al., 1987), but a trace of ethanol or wetting agent will cause the beads to wet (Lewis, 1945). On the other hand, the CW microfibrils are exposed at the mesophyll cell surface in freezefractured rye leaves, except in the vicinity of cell junctions, where they are overlain by amorphous material on which frozen water droplets were visible (Willison and Pearce, 1983). The aim of this chapter is to review currently available data on the structure, ontogeny and composition of the CM and the EW, and to integrate the observed variability in cuticle types into a structural and developmental model.

2.1.2

Definition and nomenclature of the plant cuticle

This chapter uses the terminologies of Wattendorff and Holloway (1980), Holloway (1982a) and Jeffree (1996) with additions where appropriate. Brongniart (1834) gave the name ‘cuticula’ to a superficial membrane isolated from the cabbage leaf epidermis by retting in water. He recognised that this first isolated cuticle contained fatty materials because it stained with Sudan dyes. More than a century later, Norris and Bukovac (1968) defined the limits of the cuticle as ‘all of the layers that can be separated from the underlying cellulose cell wall’, thus including the so-called cuticular, cutinised and cuticularised layers (sensu Esau, 1953; Crafts and Foy, 1962; Sitte and Rennier, 1963). The concept of a layered cuticle is attributable to von Mohl (1847), who regarded the cuticle or CM as consisting of two main zones at maturity, as has generally been accepted since then (Lee and Priestley, 1924; von Ziegenspeck, 1928; Crisp, 1963; Sitte and Rennier, 1963; Hülsbruch, 1966a,b; Chafe and Wardrop, 1973; Jarvis and Wardrop, 1974; Sargent, 1976a,b,c; Holloway, 1982a; Jeffree, 1996). The outermost cuticle layer appears outside the primary epidermal CW very early in organ ontogeny and was called the cuticle proper (CP) by von Mohl (1847), who defined it as being composed of soluble and polymeric lipids that can be saponified without leaving a cellulosic

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BIOLOGY OF THE PLANT CUTICLE

residue. This author accepts the positional aspect of his definition and the absence of cellulose, but the CP is not always completely saponifiable, especially in mature leaves (Schmidt and Schönherr, 1982; Domínguez and Heredia, 1999). Beneath the CP a zone of the primary cell wall (PCW) and then secondary cell wall (SCW) of variable thickness becomes progressively impregnated with cutin during organ development (Schieferstein and Loomis, 1959). Although Esau (1953) referred to this zone as the ‘cuticularised layer’ (see also Martin and Juniper, 1970), von Mohl (1847) called it the cuticular layer (CL) meaning the cuticular layer of the CW. This original term was preferred by Holloway (1982a), Wattendorff and Holloway (1984) and Jeffree (1996), and will also be used here, both because of its precedence and because the unspecified process and location of cuticularisation leads to ambiguity. Late in development the CL may undergo further modification from the outside inwards, so that in the CL of some species (e.g. Ficus elastica and Ilex integra; Hülsbruch, 1966a,b) embedded cellulose may no longer be identifiable, making it hard to distinguish from the CP by von Mohl’s definition (see Figure 2.2g, P.J. Holloway). Hülsbruch called this zone ‘Cuticularsaum’, but Jeffree (1996) referred to the internal and external zones of the CL layer as the internal cuticular layer (ICL) and external cuticular layer (ECL), respectively. Evolving concepts of the structure of the cuticle have been summarised in various diagrams (Roelofsen, 1952; Schieferstein and Loomis, 1959; Martin and Juniper, 1970; Holloway, 1971; Hadley, 1981; Holloway, 1982a; Hallam, 1982; Juniper and Jeffree, 1983; Wattendorff and Holloway, 1984; Jeffree, 1986, 1996; Schönherr et al., 1991; Holloway, 1993, 1994).

2.1.3

The pectin lamella

During early ontogeny, the CM (CP only at this stage) is underlain by a superficial layer of the PCW that behaves like a polyanion. Cytochemically it gives positive reactions with cationic dyes (ruthenium red, Alcian blue, Coriphosphine O, hydroxylamine-ferric chloride) and periodate-Schiff consistent with the presence of pectin. No doubt influenced by the observations of Sitte and Rennier (1963) and Hülsbruch (1966a,b), Sargent (1976a,c) interpreted the CM of Libertia elegans as entirely external to the CW, secreted from its junction with the underlying pectic layer, not as a cutin-impregnated region of PCW. She therefore referred to the CL as the ‘secondary cuticle’, interpreting polysaccharide fibrils extending into the secondary cuticle as extensions from the pectin lamella. Orgell (1954, 1955) reported the isolation of the cuticles of Convolvulus arvensis, Vinca major and Philodendron sp. with pectic enzymes. The pectin-rich layer beneath the cuticle of pear (Norris and Bukovac, 1968) and Citrus leaves (Baker and Procopiou, 1975) can be removed using pectinase, thus liberating the CM. In some species and at some stages of development, treatments with EDTA (Letham, 1958; Schneider, 1960), oxalic acid or ammonium oxalate (Huelin and Gallop, 1951) have the same effect, presumably chelating calcium ions in the egg-box pectins of the

THE FINE STRUCTURE OF THE PLANT CUTICLE (a)

15

(e)

(b)

(c) (f) (d)

(g)

Figure 2.2 (a–e) Stages in cuticular membrane (CM) development in Norway spruce (Picea abies). (a) A thin (approximately 24 nm) apparently amorphous cuticle proper (CP) is formed outside the 240-nm thick PCW. Arrow marks an electron-dense superficial region of the primary cell wall (PCW). (b) Globular deposits beneath the CP initiate construction of a cuticular layer (CL) (arrow). (c) The reticulate CL now exceeds 200 nm thickness. Its fibrils terminate abruptly (arrow) beneath the amorphous CP. (d) The CL, now 1 μm thick, shows outer parallel-striated external cuticular layer (ECL) and inner cross-linked reticulate internal cuticular layer (ICL) regions above a non-cutinised secondary cell wall (SCW), now thickened to 1 μm. (e) A section stained en bloc with KMnO4 , showing strong contrast between the fibrillar reticulum and the cutinous matrix of the CM. The abrupt termination of the CL fibrils beneath the CP is now very clear, but there is no trace of lamellae in the CP. (f, g) Two stages in CM development in Sitka spruce (Picea sitchensis). (f) A young Sitka spruce CM showing an amorphous CP (between arrowheads) demarcated by the abrupt termination of CL fibrillae, as in (c). (g) In an older leaf, the boundary between CP and the developing electron-lucent ECL layer is scarcely visible, its probable position indicated by the arrowhead. (a–g) Bars = 200 nm. Figures (a–e) from Tenberge (1992), Canadian Journal of Botany, 70, 1467–1487. Figures 2.2f,g by P.J. Holloway.

16

BIOLOGY OF THE PLANT CUTICLE

pectic lamella. Norris and Bukovac (1968) report that the basal region of the isolated pear leaf cuticle (ICL as defined here) stains pink with ruthenium red, suggesting that pectins are embedded in the cutin matrix in positions protected from pectinase extraction. This point could be resolved today using immunogold labelling with antibodies against specific pectic epitopes, but there has been no attempt to characterise the location and type of pectin present beneath and embedded in the CM during a sequence of CM development. Although some cuticles are released rapidly by pectinase, those of mature leaves tend to be resistant (Lendzian et al., 1986). In these cuticles the pectin lamella may become so impregnated with cutin that it is impossible to remove it using enzymes, and the CM is now attached to a region of the CW that is richer in cellulose. Combinations of cellulose and pectinase usually release such cuticles and are in routine use today (Schönherr and Riederer, 1986; Lendzian et al., 1986), but more aggressive chemistry involving acids or strong oxidants may be required to liberate the CM from CW layers that are heavily cutinised or impregnated with lignin, as in conifer leaves (Silva Fernandes, 1964; Holloway and Baker, 1968). Sargent (1976a,c) interpreted the pectin lamella as coterminous with the middle lamella. However, the middle lamella derives from the cell plate formed between daughter cells following cell division, and can only be formed internally between adjacent mother CW pairs, not at the outer periclinal wall of the epidermis. In order for the middle lamella to join a sub-cuticular lamella, a region of PCW would have to be removed (see Jeffree, 1996). The process of middle lamella formation has recently been discussed by Jarvis et al. (2003). Viougeas et al. (1995) report that isolating the substantial (approximately 2 μm thick; Holloway, 1982a; Table 2.1) ivy cuticle with pectinase 2% and cellulase 0.2% at pH 4.5 does no damage to fine structure, and the isolated CM can therefore be considered valid for both fine structural and permeability studies. However, at the other end of the thickness spectrum, it is not yet possible to isolate the CM of e.g. Arabidopsis thaliana or of wheat intact, and therefore catastrophic damage is being caused to both fine structure and permeability properties. Between these extremes, a spectrum of artefactual damage must be assumed to occur, thus caveat emptor.

2.2 The structure of the cuticle proper 2.2.1

The procuticle

The CP appears on aerial plant organs very early in epidermal cell development, e.g. on unexpanded leaves still in bud (Wattendorff and Holloway, 1980; Schmidt and Schönherr, 1982; Riederer and Schönherr, 1988; Tenberge, 1992). Sargent (1976a,c) proposed that the CP be called the ‘primary cuticle’, by analogy with the terms primary and secondary as applied to the CW. However, it is now clear that

THE FINE STRUCTURE OF THE PLANT CUTICLE

17

Table 2.1 Holloway’s six cuticle structural types. The definitions are as stated by Holloway (1982a), but the examples given are selected by this author, and do not necessarily coincide either with Holloway’s categorisation or the categories specified by other sources. Further examples and their source references are listed in Table 2.3 Type

Description

Examples

1

Leaves of Agave americana and Clivia miniata

4

Polylamellate outer region, sharply delineated against inner mainly reticulate region Outer region faintly lamellate, gradually merging with inner mainly reticulate region Outer region amorphous, inner region mainly reticulate All regions reticulate

5

All regions lamellate

6

Mainly amorphous

2

3

Leaves of Hedera helix and Ficus elastica

Leaves of Plantago major, Picea abies, Citrus limon and Quercus velutina Leaf of Hydrangea macrophylla, fruit of Lycopersicon esculentum Leaves of Beta vulgaris and Taraxacum officinale Leaves of Potamogeton crispus and Brassica oleracea var gemmifera

there is no secondary cuticle with the same external status as that of the CP, and the use of the terms primary and secondary cuticles now seems redundant. However the CP, as seen at maturity and in the youngest expanding leaves, is not the earliest manifestation of the cuticle. The very earliest epidermal cells in the shoot apices and leaf primordia are already covered by a highly water-repellent, osmiophilic film which forms an amorphous, electron-dense layer approximately 20 nm thick as seen in TEM sections (Figures 2.3a, 2.4a). In Cuscuta gronovii and Utricularia sandersonii an amorphous procuticle (Heide-Jørgensen, 1991) approximately 22 nm thick is present on the earliest protodermal surfaces (Figure 2.3a), increasing in thickness to approximately 30 nm, slightly reducing in electron-density to develop a globular transitional cuticle (Figure 2.3b) before transforming into a lamellate layer approximately 35–40 nm thick and containing two–three pairs of electrondense and lucent lamellae (Figure 2.3c) identifiable as the first manifestation of the lamellate CP. The procuticle is thus the first region of the cuticle to appear during cuticle ontogeny, quickly giving rise to the CP which is assembled externally to the PCW. In the cuticle of Iris germanica (Figure 2.5a, P.J. Holloway) the lamellate CP is devoid of any trace of fibrillar reticulum, which is first evident immediately beneath the basal electron-lucent lamella. Equally in cuticles with an amorphous zone representing the CP (Holloway’s Cuticle Type 3) the CP is always totally devoid of fibrillar material, the fibrils terminating at a sharp boundary (Sitka spruce; Holloway, 1982a; Wattendorff and Holloway, 1984; Norway spruce; Tenberge, 1992; Figure 2.2b–e). It may be tempting to think that there is some relationship between the procuticle

18

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

Figure 2.3 A sequence of stages in development of the cuticular membrane (CM) in the bladder primordium of Utricularia laterifolia (Droseraceae). (a) A uniformly electron-dense procuticle (arrow). (b) Transitional stage between procuticle and cuticle proper, showing globular, electron-dense material in a translucent matrix (arrow). The adjacent cell wall microfibrils (pectin lamella?) are electron-dense. (c) The procuticle has transformed into a lamellate cuticle proper (CP) (arrows), with an underlying electron-dense globular layer. (d) A reticulate cuticular layer (CL) is being constructed beneath the CP, which has more lamellae and is of greater thickness. (e) The reticulate CL is now as thick as the original primary cell wall (PCW), but the thickness of the uncutinised cell wall is undiminished. LCP = Lamellate cuticle proper. (a–e) Bar = 100 nm. Figures 2.3a–e reproduced with permission from Heide-Jørgensen (1991), Planta, 183, 511–519.

THE FINE STRUCTURE OF THE PLANT CUTICLE

(a)

19

(e)

(b)

(c)

(d) (f)

Figure 2.4 (a–d) Cuticles of mosses. (a) A three-week protonema of Sphagnum fimbriatum bears a superficial electron-dense layer analogous to the procuticle shown in Figure 2.3a. (b) Two sections of the cuticular membrane (CM) of ten-week protonemata of S. fimbriatum, showing a cuticle proper (CP) composed of lenticular electron-lucent lamellae embedded in electron-dense matrix. (c) The CM and outer epidermal cell wall of a Marchantia polymorpha gametophyte, fixed in GA/Os and stained with Reynolds lead citrate, shows a thin, apparently structureless CM. Staining with KMnO4 enhances the contrast of the CM and resolves it as two equal layers equivalent to a CP and cuticular layer (CL; later). (d) The outer ledge of a guard cell in the capsule of a Funaria hygrometrica sporophyte shows lamellate/reticulate structure in the CM adjacent to the pore (right) but lamellae do not appear in the outer cuticle (top). The fibrils may reach the surface. (e,f) Sections of Type 1 (e) and Type 3 (f) cuticles in which the epicuticular waxes are preserved, showing their sizes relative to the CP and CL layers. (e) Agave americana: the lamellae of the CP are not interrupted or diverted beneath the wax crystals (f) Sitka spruce (Picea sitchensis). The diameter of the tube waxes is about half the thickness of the CM, but no pore of this size is resolved. Figures 2.4a–f bars = 200 nm. Figures 2.4a,b reproduced from Cook and Graham (1998), International Journal of Plant Sciences, 159, 780–787. Figure 2.4c and inset by P.J. Holloway. Figures 2.4d from Sack and Paolillo (1983a) Protoplasma, 116, 1–13. Figure 2.4e by J. Wattendorff, unpublished. Figure 2.4f from Holloway (1982a).

20

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(e)

(c)

(d)

Figure 2.5 Transmission electron micrographs of Type 1 lamellate/reticulate cuticles. (a) Isolated cuticular membrane (CM) of Iris germanica, showing a thick lamellate cuticle proper (CP), and a coarsely reticulate cuticular layer (CL). Reticulations are mostly radial with horizontal crosslinks between. (b–e) Cuticles of Agave americana leaves. (b) A CM fixed in Ga/Os/Ruthenium red without section stain shows enhancement of the density of electron-dense lamellae in the CP.

THE FINE STRUCTURE OF THE PLANT CUTICLE

21

and the hydrophobic ‘internal cuticle’. However, the presence of a CM on the surfaces of cells with a gas exchange function would be paradoxical. In the author’s experience, no cuticle or procuticle sensu Heide-Jørgensen (1991) or Jeffree (1996) is present on mesophyll cell surfaces. It is relevant to note that the capacity to convert hydroxy-fatty acids into the polymer cutin is believed to be a specialised function of the epidermal cells alone, which is lacking in mesophyll (Kolattukudy, 1968, 1984, 1996; Hooker et al., 2002).

2.2.2

The cuticle proper

The earliest appearance of the lamellate CP in Phormium tenax (Jarvis and Wardrop, 1974) is strikingly similar to that in Clivia miniata in which the lamellate CP is the earliest cuticular structure observed in sections taken from the youngest regions at the leaf base (Riederer and Schönherr, 1988). At the stage of transition from the procuticle (Figure 2.6), the CP consists only of two pairs of electron-lucent/electrondense lamellae, and contains no detectable ester cutin. At 1–2 cm from the base the CP, now increased to about 70 nm thickness, has extended to 8–10 pairs of periclinal electron-dense and electron-lucent lamellae (Figure 2.6b), and contains ester cutin, although at this stage a non-saponifiable fraction is of greater mass. Leaf area increases by a factor of about 9 in the first 5 cm from the leaf base of C. miniata but the CP lamellae give no appearance of being stretched by this, maintaining the number of lamellae through to maturity. Indeed the thickness of the lamellae and thus of the CP is at its greatest during the period of most rapid expansion up to about 6 cm from leaf base (see also Gilly et al., 1997). During ontogeny of the CP in U. sandersonii, the osmiophilic procuticle begins to develop a flocculent structure prior to the appearance of lamellae on its surface (Figure 2.3b). This very electron-dense flocculent layer increases in thickness to about 100 nm while further lamellae are added to the stack, and begins to move into the outer layers of the CW (Figures 2.3c,d), representing the earliest phases of

(Figure 2.5) The periodicity of CP lamellae decreases towards the exterior, and lamellae may merge. Termination of an electron-lucent lamella is indicated by up-arrow. Reticulations in the external cuticular layer (ECL) extend to the base of the CP, and are interspersed with faint electron-lucent lamellae (diagonal arrow), (c) CP, ECL and part of internal cuticular layer (ICL) of a mid-aged leaf stained by the H2 SO4 -I2 /KI-Ag Proteinate reaction (Holloway et al., 1981) demonstrates the distribution of epoxide groups in the electron-dense lamellae, and in ECL. The ICL is most intensely stained. As in (d and e), splitting of the CP has occurred within an electron-lucent lamella. (d) A CP detached from the CL during processing (Ga/Os fixation, U/Pb section staining) has split within an electron-lucent lamella. Arrows show terminations of electron-lucent lamellae within electron-dense lamellae. Periodicity of the lamellae decreases from inner (left) to outer (right) surfaces. The electron-dense lamellae show tripartite structure, a central electron-lucent lamella separating two denser subunits. (e) Tripartite structure of the electrondense lamellae in Figure 2.5d is emphasised by digital foreshortening of the image. (a,d) Bar = 200 nm; (b,c) bars = 100 nm; (e) bar = 50 nm. Figure 2.5a by P.J. Holloway, unpublished, Figures 2.5b–e reproduced from Wattendorff and Holloway (1980), Annals of Botany, 46, 13–28.

22

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(i)

(e)

(f)

(g)

(h)

Figure 2.6 A sequence of developmental stages of the cuticular membrane (CM) of Clivia miniata (Amaryllidaceae) leaf (adaxial): (a) the leaf base: a simple cuticle proper (CP), with one–two pairs of lamellae. (b) The CP 1 cm from base has eight–nine pairs of lamellae, underlain by an electron-dense, probably pectinaceous, region of the outer primary cell wall. (c) The CP 2 cm from base: twelve–thirteen pairs of CP lamellae are now present, of greater and more variable thickness. The innermost and outermost lamellae are denser than in the central region. Electron-dense globules are accumulating beneath the CP, commencing the process of cuticular layer (CL) construction (arrow). (d) CP with twelve–thirteen pairs of lamellae, 3 cm from the leaf base. The electron-dense lamellae show tripartite structure. Almost 200 nm of reticulate CL layer has developed, electron-dense globules remaining in its outer layer (arrow). (e) The CP 4 cm from leaf base, in the cell-expansion phase, the CL has a finely reticulate external layer (ECL) and a coarsely reticulate inner layer (ICL). (f) The CP 5 cm from leaf base, late in the cell-expansion phase, the reticulum of the ECL extends the base of the CP. (g) The CP 6 cm from the leaf base, the reticulum in the ECL layer is still visible but with low contrast. (h) The CP 20 cm from the leaf base, the lamellae are of low contrast except at the base of the CP. Fibrils are no longer visible in the ECL, which now contains faint lamellae. Os; U/Pb, x 75,000; bar = 100 nm. (i) The CM of C. miniata in a leaf of unspecified age, showing parallel–periclinal lamellae in the CP, chaotically orientated lamellae at the boundary between CP and ECL, an electron-lucent ECL, with reticulate structure visible only towards its base, and the reticulate ICL beneath. Bar = 200 nm. Figures 2.6a–h reproduced with permission from Riederer and Schönherr (1988), Planta, 174, 127–138. Figure is 2.6i reproduced with permission from Holloway (1982a).

THE FINE STRUCTURE OF THE PLANT CUTICLE

23

construction of the CL layer by impregnation of the PCW. Additions of lamellae within the outermost parts of the PCW are often made in a less precisely periclinal orientation to those first secreted outside it in the CP (Figure 2.3d,e). Ultimately the lamellate region of the CP in Utricularia is about 80 nm thick, and is subtended by a reticulate CL that is more than 200 nm thick and which is now more electron lucent compared with the initial stages (Figure 2.3e). Micrographs by H.S. Heide-Jørgensen published in Jeffree (1996) showed a thick lamellate CP under construction in Hakea suaveolens with densely osmiophilic basal CP lamellae. Osmiophilic globules ranging from about 20 to 75 nm in diameter are enclosed in an electron-lucent lamella about 5 nm thick, which appears to unwrap from the surface of the electron-dense globule as it coalesces with the base of the CP. Closely analogous structures are also seen in suberised layers under development in the wall of a suberised epidermal idioblast in Cassytha pubescens (Heide-Jørgensen, in Jeffree, 1996; see also de Vries, 1968). The lucent lamellae in the CP are widely held to contain soluble wax, as discussed later. If composed of nonpolar soluble cuticular lipids (SCL), then the position of electron-lucent lamellae outside such hydrophobic globules is incompatible with the aqueous and polar environment of the apoplast. However, a thin (approximately 2.5 nm), dense border is present that may be consistent with an outer protein shell, or the polar functional groups of long-chain fatty acids. The dynamics of the movement of these globules through the CW matrix (if indeed they do move through it) is obscure, as is the composition of the electron-dense and electron-lucent components of this system, and of the construction process itself (self-assembly or protein mediated assembly?), offering fertile ground for fundamental research.

2.2.2.1 Lamellate substructure of the CP Probably in most species, the CP is of lamellate construction. In those species of cuticle Types 1 and 2 for which data are shown in Table 2.2 the mean thickness of the lamellate region is about 150 ± 32 nm, exceptionally more than 1 μm in thickness, as in H. suaveolens (Heide-Jørgensen, 1978b). For many other species, listed as Type 3 by Holloway (1982a; Table 2.2), the CP appears amorphous, as in Picea abies and Picea sitchensis. In these species the lamellate structure is either absent or composed of materials of similar electron density, and thus of low contrast. As noted by Wattendorff (1984) the CP lamellae in Agave americana show substructure within the basic alternating layers of light and dark [relative to a positive print from the EM negative, this translates into electron-lucent (¯eL) and electrondense (¯eD)]. Lamellar substructure is evident in many of his excellent TEM images of young and old leaves, and the reality of the observations is reinforced by the fact that images viewed both unstained and contrasted using various different staining and contrasting methods show the same pattern. The more e¯ D lamellae are revealed as composed of three units – two outer dense lines and an inner unit of intermediate density (Figures 2.5d and e). The e¯ D lamellae of the CL layer in A. americana also

Organ St Fibres SC, fringe layer Nucellus, at anthesis A ssc A Bladder LAd L L L L L L Bs S P P P S LAb LAd LAd LAd L L

Family or order

Poaceae Malvaceae Malvaceae Malvaceae Haloragaceae Fabaceae Fabaceae Lentibulariaceae Adiantaceae Adiantaceae

Agavaceae Agavaceae Agavaceae Agavaceae Liliaceae Chenopodiaceae Apiaceae Apiaceae Apiaceae Brassicaceae Myrsinaceae Myrsinaceae Chenopodiaceae Chenopodiaceae Iridaceae Buxaceae

Species

Avena fatua Gossypium hirsutum Gossypium hirsutum Gossypium hirsutum Myriophyllum verticillatum Phaseolus vulgaris Pisum sativum Utricularia laterifolia Adiantum capillus-veneris Adiantum raddianum var. raddianum Agave americana Agave americana Agave americana Agave lutea Allium cepa Anabasis articulata Apium graveolens Apium graveolens Apium sativum Arabidopsis thaliana WT Ardisia crenata Ardisia crenata Beta vulgaris Beta vulgaris Bobartia gracilis Buxus sempervirens 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

Pc Pc Pc Pc Pc Pc Pc Pc 1 1

Type

∼ ∼ 4000 750 760 1024 270 850 660 128 800 1600 88 160 2500 0

33 20 20 20 20 20 20 25 ∼ 1000

CM (nm)

∼ ∼ 450 238 160 134 49 130 85 64 160 250 30 120 500 0

∼ ∼ ∼ ∼ 20 ∼ ∼ ∼ ∼ ∼

CP (nm)

∼ ∼ 3550 513 600 890 221 720 575 60 640 1350 58 40 2000 0

33 ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼

CL (nm)

Crisp (1965) Wattendorff and Holloway (1980) Wattendorff and Holloway (1984) Holloway, unpublished Schönherr and Mérida (1981) Lyshede (1977b) Chafe and Wardrop (1973) Juniper and Cox (1973) Hallam and Juniper (1971) Sieber et al. (2000) Fisher and Bayer (1972) Fisher and Bayer (1972) Holloway, unpublished Holloway, unpublished Sargent (1976a,b) Gouret et al. (1993)

Kaufman et al. (1970) Ryser and Holloway (1985) Ryser et al. (1988) Ryser et al. (1988) Hallam (1982) Jeffree (1996) Jeffree (1996) Heide-Jørgensen (1991) Wada and Staehelin (1996) Archer and Cole (1986)

Reference

Table 2.2 Summary of the distribution of plant cuticle structural types, and the dimensions of the cuticular membrane, cuticle proper and cuticular layer. Allocation to type is based on the constraints on Holloway’s definitions (Holloway, 1982a; Table 2.1) discussed in this chapter, principally that ‘outer region’ equates strictly to the cuticle proper (CP) and ‘inner region’ equates to the cuticular layer (CL)

24 BIOLOGY OF THE PLANT CUTICLE

L L LAd L Stigma

L S T P L L L Capsule L L L Fibres L L T L L Young leaf Older leaf Expanded leaf P L L L L

Amaryllidaceae Amaryllidaceae Amaryllidaceae Amaryllidaceae Liliaceae

Pinaceae Convolvulaceae Droseraceae Apiaceae Myrtaceae Myrtaceae Moraceae Funariales Liliaceae Liliaceae Theaceae Malvaceae Proteaceae Proteaceae Proteaceae Araliaceae Araliaceae Araliaceae Araliaceae Araliaceae Moraceae Iridaceae Asteraceae Iridaceae Iridaceae

Clivia miniata Clivia miniata Clivia miniata Clivia nobilis Crocus chrysanthus

Cunninghamia lanceolata Cuscuta campestris Dionaea muscipula Eryngium rostratum Eucalyptus papuana Eucalyptus perriniana Ficus elastica Funaria hygrometrica Gasteria planifolia Gasteria verrucosa Gordonia axilaris Gossypium hirsutum Hakea leucoptera Hakea suaveolens Hakea suaveolens Hedera helix Hedera helix Hedera helix Hedera helix Hedera helix Humulus lupulus Iris germanica Lactuca sativa Libertia elegans Libertia ixioides 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

1 1 1 1 1

∼ 70 560 800 100 320 0 200 3000 180 2000 25 150 11500 1492 2500 0 40 210 700 ∼ 800 100 700 700

210 4500 6700 ∼ 900

∼ 30 95 130 45 177 0 130 130 180 160 25 ∼ 1500 62 100 0 40 70 80 ∼ 95 50 150 150

210 200 130 ∼ 100

∼ 40 465 670 45 143 0 70 2870 ∼ 1840 0 ∼ 10000 1430 2400 0 0 140 620 ∼ 700 50 550 550

∼ 4300 6570 ∼ 800

Holloway (1982a) Holloway (1982a) Mérida et al. (1981) Holloway (1982a) Heslop-Harrison and Heslop-Harrison (1975, 1982); Heslop-Harrison (1977) Sargent (1976b) Jeffree (1986) Sievers (1968) Chafe and Wardrop (1973) Hallam (1970a) Hallam (1982) Gouret et al. (1993) Sack and Paolillo, Jr. (1983a) Holloway (1982a) Heumann (1990) Sargent (1976a,b) Ryser (1985) Sargent (1976b) Heide-Jørgensen (1978b) Heide-Jørgensen (1980) Viougeas et al. (1995) Gouret et al. (1993) Gilly et al. (1997) Gilly et al. (1997) Gilly et al. (1997) Chafe and Wardrop (1973) Holloway et al. (1981) Holloway et al. (1981) Sargent (1976b) Sargent (1976a,b)

THE FINE STRUCTURE OF THE PLANT CUTICLE

25

Organ L, CF air F F L LAb L L Outer St ridge Lower St ridge L L L L Papillae L L L Gametophyte Pitcher, inside Pitcher, inside Pitcher, outside SP SP

L, Cuticle < 2 μm thick S LAb

Family or order

Magnoliaceae Rosaceae Rosaceae Rosaceae Rosaceae Solanaceae Solanaceae Fabaceae Fabaceae Fabaceae Agavaceae Agavaceae Agavaceae Agavaceae Anacardiaceae Plantaginaceae Pinaceae Psilotaceae Sarraceniaceae Sarraceniaceae Sarraceniaceae Poaceae Caryophyllaceae

Iridaceae Fabaceae Theaceae

Species

Liriodendron tulipifera Malus pumila Malus pumila Malus pumila Malus pumila Nicotiana glauca Nicotiana tabacum Phaseolus vulgaris Phaseolus vulgaris Phaseolus vulgaris Phormium tenax Phormium tenax Phormium tenax Phormium tenax Pistacia palaestina Plantago major Pseudotsuga menziesii Psilotum nudum Sarracenia psittacina Sarracenia psittacina Sarracenia purpurea Secale cereale Silene dioica

Sisyrinchium filifolium Spartocytisus filipes Symplocos paniculata

Table 2.2 Continued

1 1 1

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

Type

1500 ∼ 193

200 ∼ ∼ 380 980 250 260 444 20 120 333 1063 635 4300 250 230 150 375 122 43 770 26 330

CM (nm)

∼ ∼ 40

54 ∼ ∼ 40 52 46 35 143 ∼ 16 133 144 120 200 170 50 150 200 40 25 39 5 75

CP (nm)

∼ ∼ 153

146 ∼ ∼ 340 928 204 225 300 ∼ 104 200 919 515 4100 80 180 ∼ 175 82 18 731 21 255

CL (nm)

McQuattie and Rebbeck (1994) Linskens and Gelissen (1966) Hilkenbäumer (1958) Hoch (1979) Holloway (1982a) Mérida and Ogura (1987) Krüger et al. (1996) Holloway, unpublished Holloway, unpublished Holloway (1982a) Hallam (1982) Holloway, unpublished Jarvis and Wardrop (1974) Jarvis and Wardrop (1974) Sargent (1976b) Fisher and Bayer (1972) Sargent (1976b) Whittier and Peterson (1995) Joel and Heide-Jørgensen (1985) Joel and Heide-Jørgensen (1985) Joel and Heide-Jørgensen (1985) Heslop-Harrison (1977) Heslop-Harrison and Heslop-Harrison (1975, 1982); Heslop-Harrison (1977) Sargent (1976b) Lyshede (1978) Tegelaar (1990)

Reference

26 BIOLOGY OF THE PLANT CUTICLE

Tamarix pentandra Taraxacum officinale Ticoa harrisii Utricularia laterifolia Vanilla planifolia Cuscuta gronovii Abutilon striatum Anabasis articulata Ardisia crenata Arum maculatum Dionaea muscipula Eucalyptus papuana Ficus elastica Hedera helix Lactuca sativa Malus pumila Malus sp. Spinacia oleracea Narcissus pseudonarcissus Acacia senegal Acer saccharum Acer saccharum Arabidopsis thaliana wax2 Arum maculatum Arum maculatum bergenia purpurascens Berula erecta Cannabis sativa Chamaegigas intrepidus Citrus limon Cuscuta odorata Dianthus caryophyllus

Convolvulaceae Malvaceae Chenopodiaceae Myrsinaceae Araceae Droseraceae Myrtaceae Moraceae Araliaceae Asteraceae Rosaceae Rosaceae Chenopodiaceae Amaryllidaceae Fabaceae Aceraceae Aceraceae Brassicaceae Araceae Araceae Saxifragaceae Apiaceae Moraceae Scrophulariaceae Rutaceae Convolvulaceae Caryophyllaceae

Tamaricaceae Asteraceae Cycadeae Lentibulariaceae

L L Lad Bladder LAd L S P S L L T L L L L LAd LAd L Petal L LAd LAb S LAd LAb L Aerial L G L L S L 1 1 1 1 1 1,5 2 2 2 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 3 3 3 3 3

200 182 3270 313 0 260 ∼ 1120 1600 320 ∼ 100 ∼ 1957 250 ∼ 525 240 220 2000 4200 1875 209 320 220 175 161 200 ∼ 1385 ∼ 2000

200 30 820 78 0 50 ∼ 179 250 200 ∼ 45 ∼ 260 36 ∼ 250 35 40 55 1200 500 90 68 40 31 800 20 ∼ 30 ∼ ∼ ∼ 152 2180 235 0 210 ∼ 941 1350 120 ∼ 45 ∼ 1697 214 ∼ 275 205 180 1945 3000 1375 120 252 180 144 810 180 ∼ 1354 ∼ ∼

Sargent (1976b) Holloway (1982a) Archangelsky et al. (1986) Heide-Jørgensen (1991) Gouret et al. (1993) Heide-Jørgensen (1991) Chafe and Wardrop (1973) Lyshede (1977b) Fisher and Bayer (1972) Holloway (1982a) Sievers (1968) Hallam (1970a) Holloway (1982a) Holloway (1982a) Srivastava et al. (1977) Hoch (1975, 1979) Holloway, unpublished Holloway et al. (1981) Holloway, unpublished Wattendorff (1974) Gordon (1995) Gordon (1995) Sieber et al. (2000) Holloway, unpublished Holloway, unpublished Holloway, unpublished Frost-Christensen et al. (2003) Mahlberg and Kim (1992) Lehmann and Schulz (1976) Holloway (1982a) Weinert and Barckhaus (1975) Reed (1979)

THE FINE STRUCTURE OF THE PLANT CUTICLE

27

Organ G G L L LAd L L Glandular T L L SC Nucellus, post-anthesis L L L L, 2xO3 Aerial L Aquatic L L ab LAb LAb, Inner epidermal cuticle L SP Leaf

Family or order

Droseraceae Droseraceae Aspidiaceae Myrtaceae Fagaceae Moraceae Rosaceae Malpighiaceae Theaceae Malvaceae

Malvaceae

Malvaceae

Saxifragaceae Oleaceae Liliaceae Magnoliaceae Lobeliaceae Lobeliaceae Rosaceae Rosaceae Rosaceae Liliaceae Onagraceae

Fabaceae

Species

Dionaea muscipula Drosophyllum lusitanica Dryopteris filix mas Eucalyptus perriniana Fagus sylvatica Ficus lyrata Fragaria ananassa Galphimia brasiliensis Gordonia axilaris Gossypium hirsutum cv Green Lint Gossypium hirsutum cv Green Lint Gossypium hirsutum cv Green Lint Hydrangea macrophylla Ligustrum ovalifolium Lilium candidum Liriodendron tulipifera Lobelia dortmanna Lobelia dortmanna Malus pumila Malus sp. Malus sp. Muscari atlanticum Oenothera organensis

Phaseolus vulgaris

Table 2.2 Continued

3

3 3 3 3 3 3 3 3 3 3 3

3

3

3 3 3 3 3 3 3 3 3 3

Type

1357

727 770 500 150 952 938 980 812 2000 ∼ 250

∼

10055

90 300 138 500 309 2000 480 690 1900 156

CM (nm)

107

55 ∼ 150 54 220 220 52 205 ∼ ∼ 70

20

55

∼ 800 40 ∼ 43 ∼ 160 17 316 ∼

CP (nm)

∼

672 ∼ 350 96 732 718 928 667 ∼ ∼ 180

∼

10000

∼ 220 98 ∼ 266 ∼ 320 667 1560 ∼

CL (nm)

Holloway (1982a) Hallam and Juniper (1971) Maier (1968) McQuattie and Rebbeck (1994) Frost-Christensen et al. (2003) Frost-Christensen et al. (2003) Holloway (1982a) Holloway, unpublished Holloway, unpublished Holloway (1982a) Heslop-Harrison and Heslop-Harrison (1982) Hallam (1982)

Ryser et al. (1988)

Ryser and Holloway (1985)

Joel and Juniper (1982) Joel and Juniper (1982) Holloway (1982a) Hallam (1970a) Bussotti et al. (1998) Davis (1987) Holloway (1982a) Castro et al. (2001) Holloway, unpublished Ryser and Holloway (1985)

Reference

28 BIOLOGY OF THE PLANT CUTICLE

Phyllirea latifolia Phyllirea latifolia Phyllirea latifolia Phyllitis scolopendrium Picea abies Picea rubens Picea sitchensis Picea sitchensis Pinus sylvestris Pinus sylvestris Pinus strobus Pinus strobus Plantago major Plantago major Potamogeton crispus Prunus laurocerasus Prunus persica Pyrus communis Pyrus communis cv ‘Passe Crassanne’ Quercus velutina Quercus velutina Quercus velutina Quercus velutina Quercus velutina Quercus velutina Quercus velutina Ribes nigrum Rumex sp. Rumex conglomeratus Phyllitisscolopendrium Spartocytisus filipes

Mature T Young T Senescent T L L L L Young L Leaf, control Leaf UVB treated L, 2 × O3 L, CF air L L L L L LAb L LAb GC outer, SUN LAb GC outer, SHADE LAb GC outer, SUN LAb GC outer, SHADE LAd, SUN LAd, SHADE LAb L Glandular T Glandular T L S

Oleaceae Oleaceae Oleaceae Aspleniaceae Pinaceae Pinaceae Pinaceae Pinaceae Pinaceae Pinaceae Pinaceae Pinaceae Plantaginaceae Plantaginaceae Potamogetonaceae Rosaceae Rosaceae Rosaceae Rosaceae

Fagaceae Fagaceae Fagaceae Fagaceae Fagaceae Fagaceae Fagaceae Saxifragaceae Polygonaceae Polygonaceae Dspleniaceae Fabaceae 3 3 3 3 3 3 3 3 3 3 3 3

3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 871 661 145 355 2960 890 500 ∼ 350 650 77 17000

1154 219 500 ∼ 3600 370 2200 288 192 412 150 500 ∼ 275 95 188 ∼ 500 ∼ 97 48 ∼ ∼ ∼ ∼ ∼ ∼ 38 150 21 140

∼ ∼ ∼ ∼ 44 45 200 15–25 64 74 54 54 ∼ 38 ∼ 64 ∼ 100 ∼ 774 613 ∼ ∼ ∼ ∼ ∼ ∼ 313 500 56 17000

∼ ∼ ∼ ∼ 3556 325 2000 273–263 128 338 96 446 ∼ 238 ∼ 124 ∼ 400 ∼ Osborn and Taylor (1990) Osborn and Taylor (1990) Osborn and Taylor (1990) Osborn and Taylor (1990) Osborn and Taylor (1990) Osborn and Taylor (1990) Osborn and Taylor (1990) Holloway (1982a) Holloway (1982a) Chafe and Wardrop (1973) Holloway, unpublished Lyshede (1978)

Gravano et al. (1998) Gravano et al. (1998) Gravano et al. (1998) Holloway (1982a) Tenberge (1992) Percy et al. (1992) Holloway (1982a) Holloway (1982a) Laakso et al. (2000) Laakso et al. (2000) McQuattie and Rebbeck (1994) McQuattie and Rebbeck (1994) Crisp (1965) Holloway (1982a) Holloway, unpublished Holloway (1982a) Schneider and Dargent (1977) Holloway (1982a) Gouret et al. (1993)

THE FINE STRUCTURE OF THE PLANT CUTICLE

29

LAd L L L LAd L P L L L L C Stigma Petal G T T L Peel SC Fruit L LAd L G Td L L

Chenopodiaceae Caryophyllaceae Arecaceae Theaceae Theaceae Liliaceae Ericaceae Fabaceae Fabaceae Pinaceae Liliaceae Poaceae Brassicaceae

Ranunculaceae Moraceae Moraceae Moraceae Solanaceae Rutaceae Rutaceae Rutaceae Rubiaceae Rubiaceae Pinaceae Droseraceae Bignoniaceae

Spinacia oleracea Stellaria media Syagrus coronata Symplocos hallensis Symplocos paniculata Tulipa gesneriana Vaccinium reticulatum Vicia faba Vicia faba Abies balsamea Aloe arborescens Avena fatua Brassica oleracea var. botrytis Caltha palustris Cannabis sativa Cannabis sativa Cannabis sativa Capsicum annuum Citrus paradisi Citrus paradisi Citrus sinensis Coffea arabica Coffea arabica Cryptomeria japonica Drosera sp. Eccremocarpus scaber Galium aparine Genista aetnensis

Fabaceae

Organ

Family or order

Species

Table 2.2 Continued

4 4 4 4 4 4 4 4 4 4 4 4 4 4 4

3 3 3 3 3 3 3 3 3 4 4 4 4

Type

580 157 190 590 ∼ 1900 1000 ∼ 2000 1000 2000 130 250 ∼ 850

263 58 ∼ 2182 484 240 2100 110 188 1200 2200 1100 125

CM (nm)

∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼

110 15 ∼ 0 40 80 0 75 63 ∼ ∼ ∼ ∼

CP (nm)

580 157 190 590 ∼ ∼ ∼ ∼ 2000 1000 2000 ∼ ∼ ∼ 850

160 43 ∼ 2182 444 160 2100 25 125 ∼ 2200 1100 ∼

CL (nm)

Whatley (1984) Kim and Mahlberg (1995) Mahlberg and Kim (1991) Hammond and Mahlberg (1978) Gouret et al. (1993) Espelie et al. (1980) Espelie et al. (1980) Thomson and Platt-Alloia (1976) Holloway et al. (1981) Holloway, unpublished Sargent (1976b) Chafe and Wardrop (1973) Junker (1977) Gouret et al. (1993) Lyshede (1982)

Holloway, unpublished Holloway (1982a) Machado and Barros (1995) Tegelaar (1990) Tegelaar (1990) Holloway (1982a) Singh and Hemmes (1978) Holloway, unpublished Holloway, unpublished Chabot and Chabot (1977) Kluge et al. (1979) O’Brien (1967) Elleman et al. (1988)

Reference

30 BIOLOGY OF THE PLANT CUTICLE

Hakea leucoptera Ilex aquifolium Ilex aquifolium Ilex aquifolium Ilex aquifolium Ilex integra Lycopersicon esculenntum Lycopersicon esculentum Lycopersicon esculentum Magnolia grandiflora Malus pumila Nicotiana tabacum Phaseolus vulgaris Pinus nigra Pinus sylvestris Prunus laurocerasus Sisyrinchium filifolium Spartocytisus filipes Spartocytisus filipes Tamarix pentandra Tradescantia virginiana Triticum aestivum Tropaeolum majus Utricularia sandersonii Vicia faba Arabidopsis thaliana Atriplex semibaccata Beta vulgaris Beta vulgaris Eucalyptus cinerea Gordonia axilaris Plantago major

Proteaceae Aquifoliaceae Aquifoliaceae Aquifoliaceae Aquifoliaceae Aquifoliaceae Solanaceae Solanaceae Solanaceae Magnoliaceae Rosaceae Solanaceae Fabaceae Pinaceae Pinaceae Rosaceae Iridaceae Fabaceae Fabaceae Tamaricaceae Commelinaceae Poaceae Tropaeolaceae Lentibulariaceae Fabaceae Brassicaceae Chenopodiaceae Chenopodiaceae Chenopodiaceae Myrtaceae Theaceae Plantaginaceae

T LAd LAb L L L L F F L Ad. F T Inner epidermal cuticle L L L L, cuticle > 2 μm thick L, Mucilaginous cell Mucilaginous cell Papillae Petal Ovule Stigma Stolon, wart L Cutinase-expressing L L Epidermis, internal L L L Stomatal lip 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 5 5 5 5 5 5 5 ∼ 1880 28 ∼ 10000 ∼ ∼ 9000 2000 1800 10000 300 6 4700 ∼ ∼ 2500 110 ∼ ∼ 156 870 240 520 100 240 100 80 56 590 2000 1000

∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ 100 67 56 100 ∼ ∼

∼ ∼ ∼ ∼ ∼ ∼ ∼ 9000 2000 ∼ 10000 300 ∼ ∼ ∼ ∼ ∼ 110 ∼ ∼ ∼ 870 240 520 100 ∼ 0 15 0 490 ∼ ∼

Sargent (1976b) Holloway (1982a) Holloway (1982a) Holloway (1982a) Sargent (1976b) Hülsbruch (1966a,b) Gouret, Rohr, and Chamel (1993) Wilson and Sterling (1976) Rijkenberg et al. (1980) Postek (1981) de Vries (1968a) Akers et al. (1978) Holloway, unpublished Campbell (1972) Walles et al. (1973) Gouret et al. (1993) Sargent (1976b) Lyshede (1977a) Lyshede (1977a) Sargent (1976b) Holloway, unpublished Morrison (1975) Shayk et al. (1977) Heide-Jørgensen (1991) McKeen (1974) Sieber et al. (2000) Kolattukudy (1980) Holloway, unpublished Holloway (1982a) Hallam (1964) Sargent (1976a,b) Holloway, unpublished

THE FINE STRUCTURE OF THE PLANT CUTICLE

31

Organ Young Pitcher, outside N Leaf L Aquatic L L L L T L L L L Aerial L Aquatic L L LAb gametophyte Aerial L Aquatic L L

Family or order

Sarraceniaceae Malvaceae Brassicaceae Poaceae Apiaceae Brassicaceae

Brassicaceae

Brassicaceae

Moraceae Hydrocharitaceae Hydrocharitaceae Rosaceae Poaceae Acanthaceae Acanthaceae Lemnaceae Rosaceae Marchantiales Lamiaceae Lamiaceae Posidoniaceae

Species

Sarracenia purpurea Abutilon venosum Arabidopsis thaliana Avena fatua Berula erecta Brassica napus

Brassica oleracea var. gemmifera Brassica oleracea var. botrytis Cannabis sativa Elodea canadensis Elodea canadensis Fragaria ananassa Hordeum vulgare Hygrophila corymbosa Hygrophila corymbosa Lemna minor Malus pumila Marchantia sp. Mentha aquatica Mentha aquatica Posidonia australis

Table 2.2 Continued

6 6 6 6 6 6 6 6 6 6 6 6 6

6

6

5 6 6 6 6 6

Type

∼ 36 ∼ ∼ ∼ 52 3 ∼ ∼ 25 71 62 ∼

∼

∼

642 400 91 ∼ 25 ∼

CM (nm)

∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ 25 ∼ ∼ ∼

∼

∼

75 ∼ ∼ ∼ ∼ ∼

CP (nm)

∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ 0 ∼ ∼ ∼

∼

∼

567 ∼ ∼ ∼ ∼ ∼

CL (nm)

Hammond and Mahlberg (1978) Holloway, unpublished Crisp (1965) Holloway et al. (1981) Sargent and Gay (1977) Frost-Christensen et al. (2003) Frost-Christensen et al. (2003) Holloway (1982a) Holloway et al. (1981) Holloway, unpublished Frost-Christensen et al. (2003) Frost-Christensen et al. (2003) Kuo (1978)

Holloway (1982a)

Joel and Heide-Jørgensen (1985) Findlay and Mercer (1971) Sieber et al. (2000) Holloway (1982a) Frost-Christensen et al. (2003) Armstrong and Whitecross (1976) Reed (1979)

Reference

32 BIOLOGY OF THE PLANT CUTICLE

Potamogetonaceae Potamogetonaceae Potamogetonaceae Saxifragaceae Saxifragaceae Euphorbiaceae Saxifragaceae Fabaceae Liliaceae Lentibulariaceae Scrophulariaceae Scrophulariaceae Fabaceae Poaceae Zosteraceae Rosaceae Rosaceae Rosaceae Fabaceae Rutaceae Caprifoliaceae Caprifoliaceae

L Aquatic L Aquatic LAd LAb LAd L L T L Bladder T Aerial L Aquatic L GC L L LAd LAd LAb, Outer epidermal cuticle H Juice sac N N 6 6 6 6 6 6 6 6 6 6 6 6 6 6 6 7 7 7 ? ? ? ?

42 35 45 ∼ ∼ ∼ ∼ ∼ ∼ 71 91 28 72 ∼ 570 1200 1100 2400 ∼ 100 400 3000 ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ 88 42 ∼ ∼ ∼ ∼ ∼

∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ ∼ 1112 1058 ∼ ∼ ∼ ∼ ∼

Holloway (1982a) Frost-Christensen et al. (2003) Frost-Christensen et al. (2003) Holloway et al. (1981) Holloway et al. (1981) Holloway et al. (1981) Lehmann and Schulz (1976) Lyshede (1976) Holloway et al. (1981) Fineran and Lee (1975) Frost-Christensen et al. (2003) Frost-Christensen et al. (2003) Pallas and Mollenhauer (1972) Holloway (1982a) Barnabus et al. (1977) Holloway, unpublished Holloway (1982a) Holloway, unpublished Schnepf (1969) Espelie et al. (1980) Fahn and Rachmilevitz (1970) Fahn and Rachmilevitz (1970)

Abbreviations used: A = shoot apex, Ab = abaxial, Ad = adaxial, Bs = bulb scale, C = coleoptile, Ch = chambered CM, CM = cuticular membrane, CP = cuticle proper, CL = cuticular layer, ECL = external CL, ICL = internal CL, PL = pectin lamella, GC = guard cell, D = detached CM, EP = epidermal papillae, F = fruit, G = gland, H = hydathode, L = leaf, Mc = mucilaginous cell, O = ovule, N = nectary, P = petiole, Pc = procuticle, po = porous CM, S = stem, ssc = sub-stomatal cavity, SC = seed coat, SP = stigma papillae, St = stoma, T = trichome, Td = tendril, wr = wrinkled CM, ∼ = no data available.

Potamogeton crispus Potamogeton crispus Potamogeton spp. Ribes nigrum Ribes nigrum Ricinus communis Saxifraga granulata Spartocytisus filipes Tulipa gesneriana Utricularia monanthos Veronica anagallis-aquatica Veronica anagallis-aquatica Vicia faba Zea mays Zostera capensis Pyrus communis Pyrus communis Malus sp. Cicer arietinum Citrus paradisi Lonicera japonica Lonicera japonica

THE FINE STRUCTURE OF THE PLANT CUTICLE

33

34

BIOLOGY OF THE PLANT CUTICLE

show tripartite structure in sections stained with KMnO4 (Figure 2.5b, Holloway, 1982a; Figure 2.7b, Wattendorff and Holloway, 1984). CP and CL lamellae are of similar structure indicating that they are of similar composition. The thickness of the e¯ L lamellae is approximately 5–8 nm (Chafe and Wardrop, 1973), generally rather greater than for the e¯ L lamellae of suberin (3–6 nm; Heide-Jørgensen, 1980) although the ranges overlap. The e¯ L lamellae of cuticles are often reported to be more uniform in thickness at about 5 nm than the e¯ D lamellae (see e.g. Domínguez and Heredia, 1999). In youngest leaves of Hedera helix the CP has 3–5 lamellar units increasing to 14 with mean thickness of a lamellar unit (one e¯ D and one e¯ L layer) of 5.2 nm. Later in the expanded leaf the number of lamellar units reduces again to 9, and the thickness of the lamellar unit increases to 9.4 nm (Gilly et al., 1997). Graded periodicity in the thickness of the lamellae is commonly observed, with the closest spacing at the external surface as in Wattendorff’s images of A. americana (Figures 2.4e and 2.5b,d). Graded thickness of the CP lamellae in P. tenax is most pronounced in early CM development (Jarvis and Wardrop, 1974). The same can be said of many other species, including Gasteria planifolia, C. miniata, and so on (Riederer and Schönherr, 1988; Figures 2.6c–f). In general, the e¯ L lamellae are of more uniform thickness, while e¯ D lamellae towards the bases of the stacks are often particularly variable in thickness, especially where there is high convex curvature of the CM. However, there is considerable variability, making it hard to make consistent generalisations. In a set of Holloway’s negatives from the same Iris leaf it is possible to find locations where all these generalisations are contradicted, so that electronlucent lamellae may in places be larger and more variable. Clearly there may be changes in apparent thickness with local tilt relative to the beam (as demonstrated by Heide-Jørgensen, in his figure 2.8 in Jeffree, 1996). However, the following points lead me to conclude that lamellar thicknesses are intrinsically variable over a limited range: first, the commonly observed tapering from base to surface of the stack cannot be explained by variation in tilt angle, and second the contrast and edge sharpness of regions which demonstrate variability often both remain high, while tilting out of their plane of alignment with the beam degrades both.

2.2.2.2 Lamella position and orientation Usually the cuticular lamellae are superficial, located in the CP or outermost ICL, but many examples exist of lamellae extending throughout the CM. In some species with heavy cuticles such as A. americana (Figure 2.7a), the lamellate region extends fully to the base of the ICL. Most cuticles demonstrating lamellae in the CL layer also have a lamellate CP. Very occasionally, examples arise of CMs with lamellae only in the CL layer, and not in the CP, as in the adaxial leaf cuticles of pear (Pyrus communis, Figure 2.1a) and apple (M. pumila; Figure 2.1c; Holloway, 1982a), and there is no reason to suppose these are unique. Lamella orientation is strictly parallel to the epidermal cell surface in the CP layer of most species, although Hallam (1967) described the CP lamellae of Eucalyptus cinerea as anastomosing. The orientation of lamellae in the CL layer generally becomes increasingly chaotic

THE FINE STRUCTURE OF THE PLANT CUTICLE (b)

35

(a)

Figure 2.7 The cuticular membrane (CM) of Agave americana following exposure of the cut end of the membrane to KMnO4 , simultaneously fixing and staining en bloc. (a) The distance of penetration of the CM by KMnO4 from right to left of this image gives a graphical representation of the permeability to the salt in aqueous solution. The cuticle proper (CP) (top right) is scarcely penetrated. In the cuticular layer (CL) the polysaccharide reticulum appears to provide fast-track access to deeply located sites. The salt also moves more rapidly in the electron-dense lamellae of the CP than in the electron-lucent lamellae (b). The entire CL of this Type 1 CM is lamellate to its base. (a) Bar = 200 nm; (b) bar = 50 nm. Figures 2.7a,b from Wattendorff and Holloway (1984), Planta, 161, 1–11.

36

BIOLOGY OF THE PLANT CUTICLE

with depth in the CL. In Holloway’s image of P. communis adaxial leaf cuticle (Figure 2.1a; Holloway, 1982a) the CL lamellae are sometimes almost vertical, and traverse most of the thickness of the CL. If the permeability properties of the CM depend on orientation of the lamellae transverse to the diffusion pathway for water vapour then these are in a relatively unfavourable configuration.

2.2.2.3 What are the CP lamellae? The e¯ D lamellae in the CP and CL appear to be a fast pathway for periclinal penetration of KMnO4 (Wattendorff and Holloway, 1984). Their contrast is increased by section staining with KMnO4 , and they are also positive for the KI/H2 SO4 silver-proteinate reaction that localises epoxide groups in the cutin (see later). These findings are consistent with a polar cutin domain, while the e¯ L lamellae appear impermeable and unreactive to most EM stains, including osmium tetroxide, consistent with a non-polar hydrocarbon. It has been suggested that the e¯ L lamellae represent the location of SCL that have been extracted during specimen preparation and that these may constitute the main cuticular permeability barrier (Wattendorff, 1992). Gilly et al. (1997) discuss the concept that the e¯ L lamellae consist of soluble lipids and the e¯ D lamellae of cutin, citing Fisher and Bayer (1972), Heide-Jørgensen (1978b) and Wattendorff and Holloway (1980) (see also Hallam, 1964; Sitte, 1975; Schönherr and Mérida, 1981; Domínguez and Heredia, 1999). Fisher and Bayer (1972) clearly regarded the e¯ L lamellae of Plantago major and Ardisia crenata as equivalent to EW plates lying in horizontal orientation. The extraction of SCL from the e¯ L lamellae would account for their almost total non-reactivity to electronstains, but the presence of non-polar saturated hydrocarbons would give the same result. The CP of A. americana cuticles is prone to splitting along the e¯ L layer during exposure to the electron beam indicating that this layer is both mechanically weak and thermolabile (Wattendorff and Holloway, 1980; Figure 2.5c). The extraction of waxes from the e¯ L lamellae may account for this mechanical weakness, but again the same result may be obtained with SCL present. The polylamellate structure of suberin, which closely parallels that of cutin, is likewise thought to result from the self-assembly of alternating layers of a polyester with phenolic constituents and soluble waxes, predominantly long-chain alkanes and alcohols (Scott, 1994). There are a number of good reasons to doubt that the e¯ L lamellae are composed only of soluble, crystalline intracuticular waxes. The existence of CP lamellae is only known from EM treatments which normally expose them both to solvent extraction, and to potentially damaging temperatures close to the melting points of the SCL. If the e¯ L lamellae consisted entirely of SCL they would be extracted by many standard EM procedures, resulting in the abolition or more chaotic spacing and appearance of the lamellae. Wattendorff reported that after extraction in hot chloroform/methanol, the A. americana cuticle was penetrated equally by KMnO4 in anticlinal and periclinal directions (Wattendorff, 1984, 1992), but that the lamellar structure remained. CP lamellae may appear more prominent

THE FINE STRUCTURE OF THE PLANT CUTICLE

37

following extraction in chloroform (Mérida et al., 1981), consistent with the exposure of polar material to staining by the removal of obscuring SCL, but Viougeas et al. (1995) found that lamellae of H. helix CP were abolished by treatment with lipid solvents. Domínguez and Heredia (1999) state that ‘wax extraction from isolated cuticles followed by cutin depolymerisation still yields a solid residue with this bilayered pattern’ (see also Schmidt and Schönherr 1982 and Figures 2.8d,e). (a)

(d)

(b)

(c)

(e)

Figure 2.8 Clivia miniata leaf (adaxial): sections of enzyme-isolated cuticular membranes (CM) and polymer matrix membranes (PMX) that have been CHCl3 /methanol-extracted and acid hydrolysed (6 M HCl, 120◦ C, 24 h) (Tissue fixed and stained en bloc with GA/KMnO4 , sections stained with U/Pb). (a) CM of young leaf, 6.5 cm from base, showing a dense (polar) internal cuticular layer (ICL) and electronlucent (non-polar) external curicular layer (ECL). (b) CM of a mature leaf, 12.5 cm from base, with polar cutinised cystoliths in the inner ICL. (c) PMX, showing lamellate CP (inset). Cuticular layer (CL) are both reticulate and polylamellate. (d,e) PMX membranes de-esterified in BF3 -methanol. (d) 6.5 cm from base: the ECL is alkali-resistant (= non-ester cutin), but the ICL has been extracted (= ester cutin). Reticulate fibrils occur in the inner two-thirds of the ECL. (e) 12.5 cm from base, the non-ester cutin layer of the ECL has doubled in thickness. The outer two-thirds is now non-reticulate or weakly reticulate. The ICL has been extracted, but a polar, globular residue remains at its base, probably polysaccharide-rich. Insets to Figures 2.8d,e show that the lamellate structure of the cuticle proper (CP) survives the combination of solvent extraction, acid and alkaline hydrolysis. (a,b) Bars = 1 μm; (c–e) bars = 500 nm; insets, bars = 200 nm. Figures 2.8a–e reproduced from Schmidt and Schönherr (1982), Planta, 156, 380–384.

38

BIOLOGY OF THE PLANT CUTICLE

These observations indicate significant variability in response of cuticles to extraction treatments, which may either be attributable to differences in cuticle age and development or to intrinsic differences between species. While they are consistent with the involvement of SCL in the lamellate structure, a telling observation is that the CP of Allium cepa contains only 1 μg cm−2 of wax, equivalent at best to about 25% of the amount required to fit the alternating polymer/wax model (Schönherr and Mérida, 1981). This does not, however, negate the possibility that equivalent chemical species are involved in this lamellar structure. Simply that they are no longer soluble. A possible model would be that the waxes involved in the e¯ L lamellae of the CP are primary alcohols esterified to an ester cutin lamella, with their hydrocarbon chains co-aligned transverse to the plane of the lamella. Set against this is the key observation of the extreme sensitivity of cuticular permeability to solvent extraction (Schönherr and Mérida, 1981) which indicates that the permeability barrier is soluble. The lamellar structure is known to survive in the fossil record into the Eocene (Tegelaar, 1990) and early Cretaceous (Archangelsky et al., 1986). It is inconsistent at one point in a paper to be talking about the possible mobility of this class of compounds in epicuticular crystal self-assembly (see later), and at another to suppose that similar waxes will remain immobile during 200 million years of burial in sediment. This apparent paradox could be resolved if the e¯ L layer were composed of a crystalline array of carbon chains covalently bonded to a cutinous e¯ D lamella, by e.g. esterification of primary alcohols with cutin acids. Domínguez and Heredia (1999) argue that it may be sufficient for partial bonding to occur by cross-linking between fatty acids of the e¯ D and alcohols of the e¯ L lamella, to form a framework into which SCL (mostly alkanes) may self-assemble. In C. miniata, the main SCL constituents are approximately 38% C16 and C18 fatty acids and approximately 41% C19–C26 alkanes with C22 dominant. Their molecular model predicts the self-assembly of these constituents by hydrophobic interactions between the tails of the molecules to form into a molecular bilayer unit 4.9 nm thick, corresponding closely to the commonly reported e¯ L lamella dimension of about 5 nm (Riederer and Schönherr, 1988). This model seems to encompass the objections of resistance of the lamellae to extraction and the simultaneous vulnerability of their permeability barrier properties to solvents. The remaining challenge is to explain how it comes about that this framework is resistant to depolymerisation.

2.3 Cuticle polymers 2.3.1

Chemical types in Angiosperm and Gymnosperm cuticles

10,16-Dihydroxyhexadecanoic acid and its positional isomers are generally the major constituents of the cutins of modern Angiosperms and Gymnosperms (Holloway, 1982b) as in leaves and fruits of the Solanaceae and Citrus species, in leaves of Magnolia grandiflora, Liriodendron tulipifera and Prunus lusitanica,

THE FINE STRUCTURE OF THE PLANT CUTICLE

39

and notably as the only cuticle type in Gymnosperms. Mixed cuticles composed of C16 dihydroxy acids and C18 trihydroxy acids are very common in Angiosperms, e.g. in Vitis vinifera, A. cepa and I. germanica. C18 trihydroxy acids are rarely major constituents but occur in A. americana, Hyacinthoides non-scripta (Caldicott, 1973) and Spinacia oleracea (Holloway, 1974). Widely separated taxa often have similar cutin composition, and a single species may have different cutin compositions on different organs. Citrus leaf cuticles have a C16 type cutin, whereas in the cuticle of the inner seed coat of Citrus paradisi C18 acids predominate (Espelie et al., 1980). A completely new cuticle polymer ‘cutan’ is now known to occur in species as diverse as C. miniata (Amaryllidaceae), which contains both cutin and cutan, and sugar beet Beta vulgaris (Chenopodiaceae) in which cutan is the principal polymer (Tegelaar, 1990) and cutin may not be detectable (Holloway and Baker, 1970). The cuticle of B. vulgaris has a CP of about 100–200 nm thickness with a pronounced periclinal lamellate structure, composed of 5–15 lamellae (Type 5; Holloway, 1982a), and often lacks significant development of the CL. Histochemical reactivity of the opaque lamellae in C. miniata and A. americana suggests that polar molecules or substituted groups are present, at least in younger leaves. Thus lamellar contrast is reported by several authors to be enhanced by treatment with ruthenium red (Wattendorff and Holloway, 1980; Kruger et al., 1996; Figure 2.5d), and with KMnO4 or Ba(MnO4 )2 (Hoch, 1979; Mérida et al., 1981; Wattendorff and Holloway, 1982; Holloway, 1982a; Figures 2.4d, 2.5b) pointing to the presence of carboxyl groups. Further, the dense lamellae of A. americana react positively with H2 SO4 -I2 /KI–silver proteinate (Wattendorff and Holloway, 1980; Figure 2.5c), which is believed specifically to identify epoxy groups in the C18 type cutins by the formation of epoxide iodohydrin derivatives that reduce silver proteinate to metallic silver (Wattendorff and Holloway, 1980, 1982; Holloway et al., 1981). Schmidt and Schönherr (1982; Figures 2.8d,e) observed that following exhaustive extraction of enzyme-isolated C. miniata CM in chloroform/methanol (to remove SCL) and 6 M HCl, 120◦ C for 24 h (to hydrolyse proteins and polysaccharides), the lamellate structure of the CP of C. miniata was distorted but not destroyed by treatment with 20% BF3 -methanol, at 80◦ C for 24 h, which cleaves ester bonds in cutin (see also Domínguez and Heredia, 1999). Its appearance also predates the appearance of identifiable ester cutin in the membrane (Riederer and Schönherr, 1988). These findings have several implications. Thus, while the electron-lucent lamellae of the CP may be assembled from extractable SCL, and the electron-dense lamellae from cutin, these constituents are rapidly modified into an insoluble and non-saponifiable hydrocarbon structure. The CP in these species thus fails von Mohl’s (1874) definition that it should be completely saponifiable, and the possibility is now raised that either of two polymers, cutin and cutan, or various mixtures of them, may equally be involved in forming e¯ D lamellae that are morphologically indistinguishable.

40

2.3.2

BIOLOGY OF THE PLANT CUTICLE

The algal cuticle

Although the thallus of Chondrus crispus (Rhodophyta) is covered by a detachable film of lamellar, extracellularly secreted material that looks superficially like a cuticle or pellicle (Craigie et al., 1992), the structure is composed of proteins and polysaccharides with lipids only as minor components (Hanic and Craigie, 1969). The Green algae (Chlorophyta) and, in particular, the Charales with their multicellularity, tissue specialisation and biochemical affinity with land plants (Embryophytes) are regarded as their probable ancestors. Cook and Graham (1998) have suggested that the osmiophilic surface of the pellicle covering Coleochaete orbicularis may be analogous to the procuticle of higher plants, as defined here and in Jeffree (1996). The veracity of this interpretation depends ultimately on chemical analysis, and the detection of cutin or cutan polymers and waxes in these layers, which are currently of unknown composition and appear to lack lipids.

2.3.3

Chemical types in Bryophyte and Pteridophyte cutins

Monoclea gottschei (liverwort) and Notothylas orbicularis (hornwort) have osmiophilic surface layers analogous to a procuticle (Cook and Graham, 1998). The occurrence and functional significance of a hydrophobic cuticle of Marchantia and other liverworts in preventing water ingress through air pores into photosynthetic tissues was examined by Schönherr and Ziegler (1975). Holloway’s TEM image of the cuticle of Marchantia shows a simple uniformly electron-lucent layer comparable in thickness (25–30 nm) with the CP of typical higher plant cuticles (Figures 2.4c,e; Table 2.2). The structures of moss cuticles are analogous with those of vascular plants. Three-week old protonemata of Sphagnum fimbriatum bear a superficial e¯ D layer comparable with the procuticle of vascular plants (Figure 2.4a; Cook and Graham, 1998) that develops by 10 weeks into cuticle containing lenticular e¯ L lamellae analogous with the CP lamellae in vascular plants (Figure 2.4b; Cook and Graham, 1998). The sporophytes of mosses and hornworts have stomata and in Funaria cuticle structure closely parallels that of vascular plants (Figure 2.4d; Sack and Paolillo Jr., 1983a,b), as does cutin composition in the few mosses which have been examined. Sphagnum palustre contains 10,16-dihydroxy hexadecanoic acid identical to that which forms the major constituent of higher plant cutins (Caldicott and Eglinton, 1976). Although the CM is rarely well developed in moss gametophytes the Polytrichales are an exception, having both cuticles and EW on the edges of the photosynthetic lamellae of the leaves. The linear apertures between adjacent photosynthetic lamellae are functionally equivalent to stomata, and the lamellae to mesophyll. By contrast, the Hepaticae lack dihydroxyacids in their cutin, containing instead a unique class of (ω − 1)-hydroxymonobasic acids (Caldicott and Eglinton, 1976). Psilotum nudum also lacks dihydroxy acids, and contain hexadecane1,8,16-triol as major constituent (40%) together with 16-hydroxyhexadecanoic acid (Caldicott and Eglinton, 1975). The fern and lycopod cutins are chiefly composed of ω-hydroxymonobasic acids (Hunneman and Eglinton, 1972; Caldicott, 1973),

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41

regarded as primitive by Holloway (1982b). However the Isoetales and some members of the Selaginellales contain dihydroxyacids in substantial quantity (Caldicott, 1973).

2.3.4

Ontogeny, composition and structure of the CL

The cuticle was thought by Karsten (1857, 1860) and others to be formed by chemical transformation of the polysaccharides of the PCW, the so-called ‘metacrase’ theory. By contrast, von Mohl (1847) and de Bary (1871, 1884) saw it as superficial to the epidermal cells, lying entirely outside the CW. Many (van Wissenligh, 1895) thought of it as an oxidised lipid film, analogous to the films formed by drying oils such as linseed and poppy oils in varnish and paint, a concept that was prevalent in the twentieth century also (Lee and Priestley, 1924). It was already clear that the epidermis was the origin of the constituents of the cuticle (Damm, 1902; Lee and Priestley, 1924). Successive generations of scientists (von Ziegenspeck, 1928; Pohl, 1928; Martens, 1934; Weber, 1942) conceded that cuticle and wax originated in the epidermal cells, and searched in vain for pores that were considered necessary to transport cuticle constituents to the surface. Prior to the widespread availability of the electron microscope two further seminal observations were made. First that the CM could be thrown into folds and wrinkles on the surfaces of many leaves and petals (Figures 2.9g,h). Martens (1934) argued that these could only arise from over-secretion of cuticular materials, and not from CW modification, leading to the rapid demise of the metacrase theory. Second, Damm (1902) and later Anderson (1934) demonstrated that a cutinised layer could arise on the internal epidermal CW, and that direct external exposure to atmospheric oxygen was not essential for its formation. Epidermal cell wrinkling, seen widely in petal epidermis of angiosperms, including Tradescantia, Narcissus (Holloway; Figure 2.9h), Tropaeolum speciosum (Jeffree, 1986) and A. thaliana (Jenks et al., 1996; Chen et al., 2003; Figure 2.10i), and also in the leaf epidermises of e.g. Aesculus hippocastanum (Martin and Juniper, 1970), Syringa vulgaris (Holloway, 1971) and Acer pseudoplatanus (Holloway, 1971; Wilson, 1984), is often mistaken as a manifestation of wax structure (e.g. Dieffenbachia maculata; Sutter, 1985), but it is clear from sections of the epidermis (Figures 2.9g,h) that the cuticle itself is folded into ridges and ripples. Sargent (1976a,c) interpreted the formation of the CL or ‘secondary cuticle’ as a second secretory event on a fully expanded epidermis. The CP precedes the CL in all species [Eryngium rostratum (Chafe and Wardrop, 1973); Phormium tenax (Jarvis and Wardrop, 1974); A. americana (Wattendorff and Holloway, 1980); Utricularia laterifolia, Athanasia parviflora, C. gronovii (Heide-Jørgensen, 1991); C. miniata (Schmidt and Schönherr, 1982; Riederer and Schönherr, 1988); P. abies, (Tenberge, 1989, 1992); H. helix(Viougeas et al., 1995)], but the development of the CL begins much earlier than the completion of epidermal expansion. In P. abies, an amorphous CP (approximately 100 nm) is present on the protodermal cells in-bud (Phase I; Tenberge, 1992; Figure 2.2a). At this stage, the leaves bear no crystalline EW,

42

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

Figure 2.9 (a–d) Cuticle types of various aquatic vascular plants. (a) The thin cuticular membrane (CM) of Lemna minor shows at least three layers indicated by arrows in the inset. The outermost electron-lucent layer might have been overlooked if debris had not been deposited on top of it.

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43

which first appears on rapidly expanding leaves prior to the division of the guard cell mother-cells (Jeffree, 1974a; Figure 2.11a,b). Construction of the spruce CL starts in spring, the first accumulations of the CL entrapping polysaccharide fibrils (Figure 2.2b). In C. miniata, the CL starts to develop from about 3 cm above the leaf base, where area expansion is the most rapid (Schmidt and Schönherr, 1982), cutin deposition occurring within superficial layers of the PCW, immediately beneath the CP (Riederer and Schönherr, 1988; Figure 2.6d). The boundary of the CL with the CW is marked by globular cutin-rich cystoliths (Mérida et al., 1981) which appear not to migrate through the CW, since they are not observed throughout it, but to arise by accretion of cutin in interfibrillar space in the outer CW, the globules expanding until they coalesce. Consequently, the CL is permeated with electrondense fibrils of CW polysaccharide. Especially at the base of the CL these appear to be separated into bundles, confined to the interstices between the growing cutinous globules. In C. miniata, cell expansion is complete at about 5–6 cm from the leaf base; the amount of ester cutin in the membrane continues to increase, reaching a peak at 10–15 cm (Schmidt and Schönherr, 1982; Riederer and Schönherr, 1988). As in C. miniata (Schmidt and Schönherr, 1982; Riederer and Schönherr, 1988), the reticulate CL of spruce appears immediately beneath a clearly demarcated CP, which is amorphous in P. abies (Tenberge, 1992; Figures 2.2a–e) and P. sitchensis (Figures 2.2f,g; P.J. Holloway; Figure 2.4f, Holloway 1982a), but not necessarily so in all conifers (Table 2.2), and is formed by deposition of cutin within the outer CW. In the reticulate CL of spruce, the first cutin-embedded fibrils of the CL are carboxylated and alkali-soluble, not cellulose, and early CL development coincides with the disappearance of the pectin lamella as a distinct substratum (Tenberge, 1992; Figure 2.2b). The CL reticulum is PaTAgP-positive, indicating the presence of polysaccharides, and is labelled by cellulase-gold sol, indicating the presence of cellulose or xyloglucan (Tenberge, 1989, 1992; Figures by Tenberge in Jeffree, 1996). The reticulum also stains strongly with KMnO4 and hydroxylamine-ferric

(Figure 2.9) (b) Similarly the electron-lucent and amorphous CP of Elodea canadensis shows little contrast against the background, but is revealed beneath an attached microorganism. (c) The CM of Potamogeton crispus is very thin (approximately 50 nm), very electron-lucent and almost amorphous. (d) Aquatic and aerial leaves of Lobelia dortmanna have CM of similar thickness. The aquatic CM shown here (between arrowheads) has a faintly lamellate cuticle proper (CP) and reticulate cuticular layer (CL). (e) The CM of Red beet (Beta vulgaris) may show scant CL development, as here, but the CP is strongly lamellate. (f) The CP of dandelion (Taraxacum officinale) is strongly lamellate, grading into a sparsely lamellate and reticulate CL, but the inner CL layer lacks lamellae. The wrinkled CM of Brassica napus leaf (g) and Narcissus pseudonarcissus petal (h) are mainly amorphous, with reticulations most conspicuous beneath the wrinkles. However, faint lamellae are visible at arrows in N. pseudonarcissus (h). (i) The CL of Phaseolus vulgaris leaf CM is strongly reticulate. The fibrillae terminate beneath a sparingly lamellate CP that is terminated by a single continuous outermost electron-lucent lamella, again highlighted by attached superficial debris. (a–c and e–i) Bars = 100 nm; (d,g,h) bars = 200 nm. Figures 2.9a–c and e–i by P.J. Holloway. Figures 2.9d reproduced from Frost-Christensen et al. (2003), Plant, Cell and Environment, 26, pp. 561–569, with permission from Blackwell Publishing Ltd.

44 (b)

BIOLOGY OF THE PLANT CUTICLE

(a)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

Figure 2.10 (a–c) Cuticles of ferns: (a,b) cuticular membrane (CM) of Hart’s tongue fern (Phyllitis scolopendrium) fixed in Ga/Os and section stained with lead, shows reticulate cuticular layer (CL) structure beneath a faint amorphous cuticle proper (CP) (b, between arrows). (c) The cuticle of Dryopteris filix mas showing strongly reticulate structure of the CL beneath amorphous CP. (d) Fossil cuticle of Ticoa harissii (Cycadales) from the Cretaceous of Argentina, showing lamellate structure in the outer regions (en bloc staining with OsO4 , section staining with KMnO4 ). (e) The amorphous, wrinkled cuticle of the stem epidermis of wild-type Arabidopsis thaliana Columbia Col-0/gl1. (f) Cutinase-expressing transgenic A. thaliana Col-0/gl1. The cuticle shows lamellate structure throughout, interrupted by large voids or chambers. (g) Wheat (Triticum aestivum) stomatal complex, stained with Nile red and viewed by fluorescence microscopy. The guard cell ledges are selectively stained, and may represent exposed cutin. (h,i) Cuticles of A. thaliana ecotype C24 and wax2 mutant stem.

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45

chloride consistent with the presence of pectin, both reactions being abolished by extraction with ammonium oxalate (Tenberge, 1989, 1992). Contrary to Sargent’s interpretation that the lamellate CP constructs the EW and is consumed in the process (Sargent, 1976a,b), the CP in spruce remains constant in appearance and thickness throughout leaf expansion and CM development and during synthesis of the crystalline epicuticular layer. In H. helix, cuticles of young unexpanded leaves consist of a lamellate zone only (CP), sharply demarcated from the underlying PCW (Gilly et al., 1997). The thickness of the lamellate zone remains constant during leaf expansion, representing a third of the thickness of the CM in young leaves and about 11% in fully expanded ones, while the number of lamellae declines from 14 to 9 between these stages. Since the CP is neither thinned nor depleted during expansion of epidermal surface area, it follows that sufficient material is continuously added to the CP during leaf expansion to maintain its thickness and structural integrity, and therefore its assembly, while initiated earlier than the appearance of the CL, continues subsequently.

2.3.5

Layering of the CL

In many cuticles, the CL develops into two broad layers, differentiated by their staining reactions and evidence of differences in composition. This layering arises in part from the process of impregnation of the PCW and SCW layers, but also from a process of centripetal maturation of the deposited cutin. The ICL of the C. miniata CM stains strongly with KMnO4 , while in the ECL staining is less intense (Mérida et al., 1981). This distinction is clear at maturity, and first appears late in expansion phase of leaf growth, about 5–6 cm from the leaf base (Figures 2.6e,f; Figure 2.8). Mérida and Ogura (1987) included the CP in the ECL, but the ICL and ECL are subdivisions of the CL only as defined here, and do not include the CP (Wattendorff and Holloway, 1980; Schmidt and Schönherr, 1982). Two CL layers are also differentiated in A. americana (Wattendorff and Holloway, 1980) and H. helix (Holloway et al., 1981; Viougeas et al., 1995). The ECL may be noticeably denser than the ICL when stained with osmium, uranium and lead as in Nicotiana glauca (Mérida and Ogura, 1987). The ICL and ECL layers are also distinguishable in Avena sativa (O’Brien, 1967), Eryngium and Apium

(Figure 2.10) (h) The stem CM (between arrowheads) of wild-type C24 shows a CP with lamellate structure, and an electron-dense reticulate CL. CW = cell wall. (i) CM of wax2 stems are thicker, and the CL more electron-lucent than the wild type, but the CW and reticulum stains more strongly. Disorganised lamellae may be present. (a,c,e,f) Bar = 200 nm; (b) bar = 50 nm; (d) bar = 250 nm, (g) bar = 50 μm (h,i) bar = 100 nm. Figures 2.10a–c by P.J. Holloway. Figure 2.10d by Archangelsky et al. (1986), Botanical Journal of the Linnean Society, 92, 101–116. Figures 2.10e,f from Sieber et al. (2000), The Plant Cell, 12, 721–727 and Nawrath (2002). The biopolymers cutin and suberin, The Arabidopsis Book, American Society of Plant Biologists. Figure 2.10g from Collins et al. (2001), Physiological and Molecular Plant Pathology, 58, 259–266. Figures 2.10h and i from Chen et al. (2003), The Plant Cell, 15, 1170–1185.

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BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

Figure 2.11 Low-temperature scanning electron microscope (SEM) images of stages in the development of the epicuticular waxes of Sitka spruce (Picea sitchensis); all stages occur prior to bud-burst and may be observed on a single needle. (a) An almost wax-free cuticle, showing shallow depressions that will become the stomatal antechambers. (b) The earliest stage of wax crystal production occurs just prior to the division of the guard cell mother cells (GCMC), clusters of tubes appearing first on the stomatal accessory cells. (c) At the time of division of the GCMCs, all epidermal cells have a covering of epicuticular wax tubes. (d,e) The first stomatal apertures appear as the antechambers begin to fill with wax tubes, mainly contributed by the accessory cells.

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(Chafe and Wardrop, 1973); P. tenax (Jarvis and Wardrop, 1974); and L. elegans (Sargent, 1976a,b,c), although this was not recognised in these earlier investigations. Significant chemical differences between the ICL and ECL layers underlie the differences in staining intensity. The outermost 500 nm of the CM in expanding leaves of C. miniata withstand saponification in BF3 (Figures 2.8d,e), leaving a residue of ‘non-ester cutin’ (Schmidt and Schönherr, 1982) that is probably coterminous with the polymethylenic fraction referred to as cutan by Tegelaar (1990). The quantity of cutan in the CL of the C. miniata leaf increases as cell expansion and cutin deposition decline, so that cutan ultimately becomes more abundant than cutin, comprising about 60% of the cuticle (Schmidt and Schönherr, 1982; Riederer and Schönherr, 1988; Tegelaar, 1990). The layer of non-ester cuticle is more than 500 nm thick in mature C. miniata leaves (Figure 2.6i, Holloway, 1982a). This significant deposit of cutan does not accumulate in the innermost layers of the ICL, but in the ECL immediately beneath the CP, in which position any cutan precursors synthesised de novo would have to pass through the CL. Likewise, in P. abies, both the amorphous CP (45 nm) and the outer third of the CL (350 nm) appear to be alkali-insoluble, and can be detached as a unit from the underlying alkali-soluble CL (Tenberge, 1992). Evidence from CP/MS 13 C NMR spectroscopy and analytical pyrolysis suggests, however, that the non-saponifiable residue of spruce CM may be closely related to lignin rather than to cutan (KögelKnabner et al., 1994). Despite having been subjected to acid hydrolysis in 6M HCl at 120◦ C for 24 h, the saponification-resistant ECL remains permeated with electron-dense reticulum at its base, indicating that in this region the polysaccharide is closed for use as a polar transport pathway even for small polar molecules. Also,the alkali-extracted membrane is almost clear of reticulum in the outer ECL (70–80 nm) of the immature cuticle at the 6.5-cm position and reticulum is absent from the superficial 300–400 nm of the mature ECL, despite the fact that the reticulum extended fully to the base of the CP in the earlier stages of development. It is suggested (Nip et al., 1986; Tegelaar, et al., 1989; Tegelaar, 1990) that the cutan of A. americana is covalently bound to the reticulum polysaccharide. It might be supposed that cutan is intruded between the cutinised CL and the CP. However, Schmidt and Schönherr (1982) and Riederer and Schönherr (1988) observed that as ‘nonester cutin’ increases, ultimately to become the major polymer of the C. miniata CM, the ester cutin declines. Cutanisation of the ECL therefore probably does not arise from de novo synthesis and secretion of a new layer, but from a maturation process, involving the progressive modification of the previously deposited cutin and any embedded polysaccharide and waxes, in situ. The progressive reduction in

(Figure 2.11) (e) Tubular crystals form in all orientations from parallel to normal to the cuticle surface. (f) A proportion of the tubes show spiral striations or open-spiral structure, indicating that they are fundamentally related to chiral ribbons (see also Cerinthe major, Figure 2.14c). (a) Bar = 6 μm; (b–d) bars = 10 μm; (e) bar = 1 μm; (f) bar = 100 nm. Figures 2.11a–f by C.E. Jeffree, S. Swift and L. St-John Mosse.

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BIOLOGY OF THE PLANT CUTICLE

reactivity of all components of the ECL indicates that all types of polar functional groups are in the cutin/polysaccharide framework and are systematically eliminated during this maturation phase. The apparent absence of polysaccharide reticulum in the ECL of mature C. miniata leaves recalls von Mohl’s (1847) definition that the CP leaves no cellulose residue upon saponification. In C. miniata and P. abies however, two significant layers contain no cellulose, but neither is saponifiable, casting doubt on which layer it was that he observed. Although his definition may apply in very early C. miniata cuticles, it is rapidly defeated by changes in polymer composition in both CP and CL as the leaf develops and matures. It is relevant here also to recall the findings of Sitte and Rennier (1963) that the CL in F. elastica may be cellulose-free, and of Hülsbruch (1966a,b) that there may be both cellulosic and cellulose-free regions in I. integra. (Sargent, 1976a,c) interpreted the whole CL, or ‘secondary cuticle’, as interposed between the CP and the PCW in several species of the Iridaceae. Again the apparent absence of cellulose in the ECL probably influenced her interpretation. In many species (e.g. B. vulgaris) the CP is saponifiable early in development, but quickly becomes cutanised and unsaponifiable without losing structural integrity in the process. The CL layer is usually polysaccharide-rich at an early stage, but then becomes unsaponifiable and polysaccharide-depleted later. The CP, occurring as it does outside the CW, is not contaminated with polysaccharide. In A. americana, which also has a cuticle containing cutin and cutan (Tegelaar, 1990), the reticulate region extends to the base of the lamellate CP (Wattendorff and Holloway, 1980; Figure 2.7a). The reticulate region also terminates abruptly beneath the amorphous CP of P. abies and P. sitchensis (Figures 2.2d,f). In both A. americana and C. miniata, lamellae in the outer CL show a sharp transition from the periclinal–lamellate layering of the CP to anticlinally or randomly oriented lamellae within the CL. Polysaccharide fibrils ramify between these CL lamellae. The mature cuticles of C. miniata and A. americana show lamellation throughout a bilayered CL, much of which is also reticulate (Wattendorff and Holloway, 1982; Figure 2.7a; Schmidt and Schönherr, 1982; Figure 2.6h). Since CL lamellae precede and survive cutanisation, they are not a manifestation of the presence of cutan, and can occur in CM matrices of both cutin and cutan polymer types. Further layering of the CL can be recognised in species with heavy CMs. Tenberge (1992) recognised three sub-layers in the mature P. abies CL, differentiated chiefly by the form of the fibrils ramifying through them, and Osborn and Taylor (1990) observed an intermediate CL layer in the adaxial leaf cuticle of Q. velutina. The differences in the structure of the layers of the CL, which arise successively during development, result from two sources; first the fact that they are formed by cutin impregnation of successively deeper layers of the CW, first the pectin lamella, then the primary CW and later the secondary CW, and second from the progressive centripetal maturation of the CL by conversion to alkali resistant material. They do not signify the development of a third cuticle zone.

THE FINE STRUCTURE OF THE PLANT CUTICLE

2.3.6

49

Cutin cystoliths

The underside of the isolated CM often shows the rounded, pillow-like profiles of the cutinous globules from which the CL layer was constructed. These globules are exposed when the CM is isolated. In thick SCW of fruits, or in leaves of xerophytic species, or where cutin is deposited between epidermal cells to form spandrels, cutin may accumulate as granules separated from the CM to form cystoliths comparable morphologically and chemically to the innermost layers of the ICL. Although it is possible that cutin cystoliths are packets of cutin precursors en route from the cell membrane to the CL (Mahlberg and Kim, 1992), their entanglement in polysaccharide microfibrils indicates that they are outlying sites of accumulation of cutin within the CW.

2.4 Cuticle structural types Holloway (1982a) summarised the variation in cuticle structure by defining six types, as shown in Table 2.1, that are widely accepted as the standard descriptive tool for analysis of variation in cuticular structures. Many examples taken from the literature and elsewhere are listed in Table 2.2. Holloway’s definitions specify the inner and outer regions of the CM as lamellate, amorphous or reticulate, and do not refer explicitly to those regions as the CP and CL as defined here or in Jeffree (1996). However, in considering them further here the explicit connection is made between the outer region and the CP and the inner region and the CL. This makes it possible to resolve certain type conflicts in Holloway’s set, and leads to greater consistency in the allocation of cuticles to type.

2.4.1

Cuticle types 1 and 2

Of Holloway’s cuticle types, four out of six are concerned with the intensity of the images of lamellae and their allocation between inner (CL) and outer (CP) layers. The cuticles of Type 1 have a strongly lamellate CP in which the lamellar orientation is almost invariably parallel–periclinal. The type species include C. miniata (Schmidt and Schönherr, 1982) and A. americana (e.g. Wattendorff and Holloway, 1980, 1982, 1984) with thick, substantial cuticles (Figures 2.5b–e, 2.6, 2.7, 2.8), but many species with much thinner CMs also fall into Type 1, such as Utricularia (Figure 2.3). The cuticle of Apium graveolens petiole was shown by Juniper and Cox (1973) to have a conspicuously lamellate CP and reticulate CL (Type 1) and by Hallam and Juniper (1971) to have a faintly-lamellate CP with a reticulate CL, thus Holloway’s Type 2. However, in the same species, Chafe and Wardrop (1972) showed no evidence of a CP in A. graveolens, making it Type 4. This variation in the image contrast of the lamellae might derive either from intrinsic differences in the capacity of the lamellae to stain at different ages or developmental stages, or to differences in staining and other specimen preparation procedures between individual

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BIOLOGY OF THE PLANT CUTICLE

workers. Holloway (1982a) placed the adaxial CM of P. vulgaris in which only one– three pairs of lamellae are present (Figure 2.9i, P.J. Holloway) in Type 2, but on the same developing guard cell the upper cuticular ridge, which has clear lamellae in the CP and a reticulate CL (Figure 2.1d), could equally be in Type 1. However, the lower (internal) cuticular ridge covering the guard cell bordering the intercellular space (ICS) lacks lamellae entirely, and is therefore Type 3. Since these variations of cuticle type occur in functionally distinct regions of the same cell, it seems clear that the P. vulgaris CMs cannot be constructed on fundamentally different models but arise from variations in the expression of different features under the control of the same genome. Such intermediates between cuticle types occur frequently within and between closely related species resulting in frequent boundary conflicts between types. Similarly, the leaf cuticle of P. major normally shows no lamellae (Type 3), or only the faintest trace of lamellae on the CM of the general epidermal surfaces (Type 2), but Holloway’s image of the stomatal lip (not shown) contains extensive chaotic lamellation, demonstrating that the capacity to produce lamellae is present, even though it is not uniformly expressed over all of the epidermal surface.

2.4.2

Cuticle type 3

In the section of a mature Sitka spruce cuticle shown in Holloway’s 1982 paper, the CP is not clearly evident, the outer region showing low contrast with fibrillar material close to the surface. He therefore placed Sitka spruce in Type 4. However, in other negatives of leaves at earlier developmental stages (Figure 2.2f) a clearlydemarcated electron-lucent zone 15–25 nm thick is present outside the limit reached by the electron-dense CW-derived microfibrils, which are thus capped by this layer, and do not in fact reach the surface. I interpret this layer as the CP, and therefore allocate Sitka spruce to cuticle Type 3, rather than 4. Later, maturation of the CL layer converts the outer region to a low-density material in which fibrillar contrast is reduced, against which the CP cannot easily be distinguished (Figure 2.2g). Its earlier existence must however be acknowledged, and unless it can be demonstrated that the CP is lost at maturity in Picea, for which there is no evidence, it must (1) remain functionally significant as a permeability barrier and (2) continue to cap the ends of the polysaccharide microfibrils. Similarly, the images of P. abies cuticles by Tenberge (1992) show a clearly demarcated e¯ L layer of dimensions consistent with the CP lying on top of abruptly terminated e¯ D microfibrils. In the earliest stages of development, prior to the appearance of the CL layer in spruce, the CP stains more darkly than in maturity (Tenberge, 1992; Figure 2.2a), its electron density later reducing to match the density of the CL matrix (Figure 2.2c). In more mature spruce CMs the region clear of microfibrils increases to a few hundred nanometres thick, presumably by progressive cross-linking, cutanisation and modification of the polymers of cuticle matrix and embedded polysaccharides. The cuticle of Sitka spruce, like that of in P. abies, is therefore Type 3 and in all probability is Type 3 also in other spruce species, and the microfibrils do not in fact reach the surface. These amorphous CPs with very low electron density may easily be lost against the

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background, especially when photographed with slight over-exposure, giving the impression that the fibrils reach the surface. The lemon (C. limon) cuticle (Holloway, 1982a; Type 4) is closely similar to that of Sitka spruce, again showing a clearly demarcated layer capping the CW microfibrils embedded in the ECL and is therefore properly assigned to Type 3, not Type 4. The leaf cuticle of A. thaliana is normally shown to have a Type 3 or 4 CM with a reticulate CL and an indistinct or amorphous CP without visible lamellae. However, there are lamellae in the cuticle of the stem of this species (Jenks et al., 1996; Chen et al., 2003; Figure 2.10i), and in cutinase-expressing A. thaliana (Sieber et al., 2000) there is an exaggerated globular lamellate structure consisting of osmiophilic electron-dense globules and electron-lucent lamellae (Figure 2.10e,f). The layer is interrupted by large voids, presumably a consequence of cutinolytic activity, but in all essentials this assemblage strongly resembles the structure of lamellate cuticles and suberised lamellae during their assembly phases, and demonstrates that the same fundamental lamellate structural paradigm is present even when it is not normally visible or expressed in Type 3 cuticles. Even in submerged aquatics with Type 3 cuticles [Mentha aquatica, Lobelia dortmanna (Figure 2.9d), Berula erecta; Frost-Christensen et al., 2003], the microfibrils do not reach to the outer surface of the CM, but are capped by non-reticulate CP.

2.4.3

Cuticle type 4

By definition, Type 4 cuticles are all-reticulate, and therefore fibrils must reach the outer surface. This may apply to the cuticle of Funaria (Sack and Paolillo, 1983a; figure 6d). Since the CP is external to the CW by the definition used here, it is never reticulate. Logically then, fibrillar reticulum can only reach the surface of the CM if the CP is lost, or if the distribution of the CP is discontinuous, either from the start, or as a consequence of its failure to keep pace with area expansion of an organ. The question of whether a CP is present or absent has important functional implications for the CM as permeability barrier for water and polar, hydrophilic molecules (Chapters 8 and 9). In Hydrangea macrophylla (Type 4; Holloway 1982a) the polysaccharide fibrils appear to be close to the surface of the CM. However, in sections stained with HI AgP a continuous non-reactive zone of the dimensions of a typical CP is demarcated at the surface on top of a CL subdivided into an outer ECL and an inner ICL layer. As in P. abies and C. limon, H. macrophylla is therefore more appropriately categorised as Type 3. In the same family (Solanaceae), Mérida and Ogura (1987) and Kruger et al. (1996) detected lamellae in the CP of Nicotiana tabacum and Nicotiana glaucum, contradicting the assignment to Type 4 in Holloway (1982a) based on the data of Akers et al. (1978). The CP of I. integra was thought by Hülsbruch (1966a,b) to be lost after synthesis of the CL, which might give it the appearance of a Type 4 cuticle. Likewise, Sargent (1976a,c) noted the loss of the lamellate structure of the CP in the mature CM of Sisyrinchium filifolium. The CP

52

BIOLOGY OF THE PLANT CUTICLE

in Type 3 cuticles is often of very low electron-density, and in TEM exposures calculated for the dense CL layer or CW the brightness may saturate, thus losing it from the image. Most cuticles putatively of Type 4 are the cuticles of fruits, such as tomato (Lycopersicon esculentum), and in some examples the fibrils may come to the surface, perhaps accounting for their high permeability to water compared with cuticles of Type 1 (Schreiber and Riederer, 1996), but this hypothesis, and indeed confirmation of the existence of Type 4 cuticles, both remain to be confirmed.

2.4.4

Cuticle types 5 and 6

Holloway (1982a) included B. vulgaris (Figure 2.9e) and Taraxacum officinale (Figure 2.9f), among examples of Type 5 CMs. In B. vulgaris the tightly ordered periclinally lamellate region of the CM is similar in both size and appearance to the lamellate CP of Type 1 cuticles, such as that of I. germanica (Figure 2.5a), and is clearly a lamellate CP with only scant development of a CL layer. The CL in the B. vulgaris cuticle consists of a narrow region of disordered lamellae into which CW microfibrils penetrate, but only to a depth of a few tens of nanometres (Figure 2.9e). The B. vulgaris genome therefore codes for CL development, but it is not extensively expressed, and B. vulgaris therefore has no lesser case to be included in Type 1 than does A. americana. Interestingly, the Chenopod Spinacea oleracea closely related to B. vulgaris has a cuticle of Type 2 or 4, with no CP lamellae or only very faint traces. In S. oleracea, the CL layer is much further developed than in B. vulgaris, and contains a conspicuous, periodate-Schiff-positive reticulum in which occasional lamellae are also visible (Holloway, unpublished TEM images). The CM of A. americana is amplified to enormous thickness compared with that of B. vulgaris, and has the trademark lamellate CP of Type 1 cuticles, but the ECL and ICL are also both densely lamellate and reticulate almost to the boundary of the ICL with the PCW. Can the A. americana cuticle (Holloway Type 1) be excluded legitimately from Type 5? The predominant structural distinction between the two cuticles is not of structural type but of the extent of the development of the CL layer. As in A. americana, the Type 5 CM of T. officinale (Figure 2.9f) has not one but two main zones, the CP and the CL, each with distinctive structures. The outer region is a strongly periclinally lamellate CP, while in each case the inner shows lamellae running in less regular pattern, at a wider variety of spacings and angles. The lamellate region in T. officinale is therefore not equivalent with the lamellate CP of Type 1 CMs, but has lamellate CP and CL layers, nor is it identical structurally to the cuticle of B. vulgaris. The B. vulgaris CM is analogous to the outermost region of the T. officinale and A. americana cuticles, and may also be compared with an early developmental stage of the cuticle of Utricularia (HeideJørgensen, 1991; Figure 2.3c,d), while the thicker cuticle of T. officinale compares well with later developmental stages in Utricularia (Figure 2.3). In much thicker cuticles still, the two outer regions, CP and ECL, often correspond in appearance

THE FINE STRUCTURE OF THE PLANT CUTICLE

53

with the T. officinale cuticle, with lamellate CP and less ordered ECL, while the inner CL (ICL) is extensively reticulate. However, even in the innermost ICL there may be lamellae right down to the boundary with the PCW [A. americana (Wattendorff and Holloway, 1984; Figure 2.7)]. As in T. officinale, periclinally ordered lamellae are the signature of the CP, and the boundary between the CP and the CL is usually marked by a descent into more disordered and ultimately chaotic orientation of lamellae. Holloway (1982a) included Eucalyptus cinerea in Type 5 following Hallam (1964; Hallam figure 4.2 in Martin and Juniper, 1970) who described its cuticle as having anastomosing channels through which waxes might migrate to the surface (Hallam, 1964, 1967). In this CM the superficial three or four lamellae appear to be in tight order, and probably represent the CP, while the remainder show much more variable electron-dense lamella thickness and anastomoses between the electron-lucent lamellae and probably represent the ECL rather than the CP. Holloway (1982a) shows the Potamogeton cuticle (Figure 2.9c) as Type 6, but close inspection of his negatives indicates that it would also be possible to classify it as Type 3, since microfibrils penetrate the flocculent structure at the base, resulting in a weakly-developed reticulate CL. In Holloway’s images of Lemna (Figure 2.9a) and Elodea (Figure 2.9b) and Potamogeton (Figure 2.9c) attached epiphytic microorganisms demarcate the upper limit of the very electron-lucent CP layer, which is amorphous in Elodea, but contains one or two pairs of faint lamellae in Lemma minor. The cuticle of L. minor is thus an example of a minimally developed Type 1 cuticle, rather than Type 6. The caveats that apply to Type 5 CMs also apply to Type 6. Classifications must be made on uniform criteria. Holloway’s definitions of the cuticle types refer to ‘Inner’ and ‘Outer’ regions without explicitly connecting the outer region and the CP, although the two coincide in Type 1 cuticles for example. This makes it possible to include in Types 5 (all lamellate) and Type 6 (all amorphous) three categories of CMs that are radically different structurally and ontogenetically: cuticles that are all-CP with little or no CL development; cuticles that are all-CL without any evident CP; and cuticles with well-developed CP and CL. I therefore propose that this connection should be formally made, making it necessary to specify separately the structure of the CP and CL layers.

2.4.5

A seventh cuticle type?

In a variation that falls outside Holloway’s six types the adaxial leaf cuticles of M. pumila and P. communis both show a clear, amorphous CP with a chaotically lamellate ECL penetrated by CW microfibrils. Holloway (1982a) placed these CM in Types 2 (P. communis and M. pumila leaf adaxial cuticles; Figures 2.1a and c), but the presence of lamellae deep into the CL is an uncomfortable fit with either of these types. Lamellae are apparently absent in the abaxial leaf cuticles of the same species which are therefore Type 3 (see Holloway, 1982a).

54

BIOLOGY OF THE PLANT CUTICLE

2.5 Summary of the cuticle types The fine structure of cuticles varies predominantly in four parameters: the presence or absence of lamellae, the presence or absence of reticulum, the subdivision or otherwise of the CL layer into inner ICL and outer ECL layers and the presence or absence of any of the four layers (CP, CL, ECL, ICL). Probably, the major differences between cuticle structural types can be ascribed to the relative extent of the development of the CM and of the underlying CW, rather than to fundamental differences in mechanism, or biochemistry. Thus, for example, the simple lamellate CM of B. vulgaris (Holloway, 1982a) is structurally analogous to those of C. miniata (Figure 2.6b) and A. americana at the earliest stages of development which have been documented (Wattendorff and Holloway, 1980; Riederer and Schönherr, 1988). Indeed it is striking that across almost all cuticles for which adequate ultrastructural data are currently available the range of variation in structure displayed between fully developed CMs of increasing thickness, from example, Potamogeton to B. vulgaris. P. vulgaris and T. officinale through I. germanica, H. helix, C. miniata and A. americana, is paralleled by the changing complexity of the thickest of these cuticles during stages of their development (Jeffree, 1996). There are detailed variations in cuticle size, structure and composition between the cuticles of Bryophytes and vascular plants, between one plant family and another, and between plants adapted to contrasting ecological conditions and lifestyles; but it is also evident that they all contain elements of a common set of ultrastructural features. The cuticles of all land plants are therefore all variants on a common theme, based on the same fundamental paradigm, not on a set of six or seven different paradigms.

2.5.1

Cuticle structure/ecology

What if anything is the functional relevance of cuticle structural types? Contrasting ecological types, such as tropical rainforest species and temperate mesophytes can both have the same structural types. For example, cuticles of F. elastica and P. vulgaris and S. oleracea are all included in Holloway’s cuticle Type 2. Structural Type 1 also occurs in A. americana, Phormium, Iris, Hedera and Apium. Species with lamellate CP sensu lato (Types 1, 2, 5) occur in genera as diverse as Beta, Taraxacum, Humulus, Spartocytisus, Agave, Clivia, Pistacia, Pseudotsuga and Polytrichum, while contrasting types occur in related taxa: Abies spp. and Picea spp. are in structural Type 4 (Holloway, 1982a) but probably best allocated to Type 3 as here, since there is often a clearly demarcated amorphous CP layer (Table 2.1), while cuticles of Cunninghamia and Pseudotsuga leaves have a lamellate CP, and are thus Type 1.

2.5.2

Cuticle thickness and environment

Cuticle thickness, morphology and composition differ between plants cultivated under glass, in tissue culture and in the field (Martin and Juniper, 1970; van den Ende

THE FINE STRUCTURE OF THE PLANT CUTICLE

55

and Linskens, 1974). So also do the thickness and morphology of the cuticle differ between aerial and submerged leaves of amphibious plants (Frost-Christensen et al., 2003; Table 2.2). In Q. velutina, sun leaf CMs are thicker than shade leaf CMs due to massive amplification of the reticulate ICL (Osborn and Taylor, 1990). In the adaxial cuticles, Osborne and Taylor (1990) reported a thickened amorphous CP. However, at 0.76 μm thick this region is an order of magnitude or thicker than a typical CP as defined here, and is mounted atop a massive reticulate ICL at least 1.4 μm thick. This thick amorphous region is almost certainly not only the CP but includes an intensively cutanised ECL, and it is probable that a study of the ontogeny of this species would demonstrate the existence of an equivalent reticulate layer in expanding leaves that loses its reticulate appearance as the ECL matures. An amorphous CP, as appears to be present on abaxial leaf surfaces, would be hard to distinguish from the amorphous ECL, but its presence would again be revealed by an ontogenetic study. Different CM structures were observed by Osborne and Taylor (1990) on different cells of the abaxial epidermis, normal epidermal cells having an amorphous CP (Type 3), while subsidiary cells have reticulate CM that traverse most of the outer CM without a CP (thus Type 4). However, in my judgement, both of these CMs have a clearly distinguishable amorphous zone outside the termination zone of the CL fibrils, with dimensions within the typical range for a CP (Table 2.1). Both sun and shade outer guard cell CMs are therefore Type 3. The CM continues through the stomatal aperture to line the inner periclinal wall of the guard cells. Moreover an internal CM, which is only sparsely reticulate, lines substomatal chambers. The internal CM of sun leaves is thicker and extends considerably deeper into substomatal chambers than in shade leaves.

2.5.3

Cuticle structure and phylogeny

Norris and Bukovac (1968) commented that ‘because of technical difficulties, many early anatomical investigations were limited to plants with thick cuticles, such as Yucca, Dasylirion, Clivia, Agave and Ficus. No information is available to establish how well these early findings represent the cuticle of mesophytic economic crop plants’. The cuticles for which we have structural data remain a trivial subset of world species, and crop plant cuticles are relatively poorly sampled. Notably missing from Table 2.2 are records for rice, most grain and forage grasses, sugarcane and indeed the cuticle structures of the vast majority of the worlds top crops, and the dominant species of the world’s forests and other vegetation formations are either unknown or poorly characterised. Most of the generalisations about cuticle structure discussed so far appear to apply across the spectrum of extant vascular plants including the ferns although records for important basal groups such as the Lycopsida and Equisetopsida are lacking. Cuticles were an enabling innovation in the attainment of homoiohydry by land plants (Raven, 1977), appearing in the earliest axial pre-vascular and vascular land plants of the Silurian, although the ‘cuticles’ of some other coeval

56

BIOLOGY OF THE PLANT CUTICLE

land plants such as Nematothallus did not contain cutin or cutan (Edwards et al., 1996). The gametophytes and sporophytes of many mosses have both stomata and cuticles analogous in fine structure to those of Tracheophytes (see e.g. Funaria hygrometrica; Figure 2.4d). The gametophytes of the Polytrichales (Dawsonia, Polytrichum) also have waxy cuticles at the ends of the photosynthetic lamellae at their leaf bases. The cuticle fine structure of these has not been reported, although EW similar to those of vascular plants are present (Neinhuis and Jetter, 1995). By contrast, the cuticle of the advanced thalloid liverwort Marchantia (gametophyte, Marchantiopsida) is very thin, completely without lamellae or fibrillar/reticulate structure, and presumably falls into Type 6. The reaction is negative to staining with uranyl and lead salts and KMnO4 indicating that it is waxy, impermeable and non-polar. It seems likely that common biosynthetic pathways for the production of these compounds are present in the most primitive groups of extant land plants as well as in representatives of the most advanced, and that waxes from these widely distributed taxa can have closely comparable compositional profiles. The implication of this is that the biosynthetic pathways involved may already have been present in the most basal land plants of the Silurian, and that they have probably been strongly conserved during their subsequent evolutionary radiation.

2.6 The epicuticular wax The EW often form a visible waxy bloom on plant surfaces. They are readily isolated and examined by light and electron microscopy, and their properties, structure and chemistry are consequently much studied (Chapters 4, 5, 7–9; Baker, 1982; Barthlott and Frölich, 1983; Jeffree, 1986; Barthlott, 1990, 1993; Barthlott et al., 1998). Waxes occur on the surfaces of all land plants. EW of mosses and liverworts may contain identical compounds to those of Gymnosperms and Angiosperms, suggesting that they appeared early in land plant evolution, but although cuticles survive in the fossil record, waxes are fugitive. Plants with waxy bloom and thick cuticles are correlated in the minds of many with xeric habitats of deserts, coasts and dunes, but there is no straightforward relationship, and the Lotus effect, synonymous today with ultra-hydrophobicity caused by wax nano-structures (Barthlott and Neinhuis, 1997; Neinhuis and Barthlott, 1997; Furstner et al., 2000), is named after the sacred lotus Nelumbo nucifera, an aquatic species with glaucous leaves that uses waxes to repel water rather than conserve it. The ultrastructure of the EW and that of other species is both complex and varied. On grapes, where the EW layer is developed sufficiently to scatter light, the surface has a matt, bluish, glaucousness, contrasting with the glabrous bright green glossiness of species carrying little EW. The wax structures of sugarcane (Saccharum officinarum) are sufficiently large to be seen individually with a powerful hand lens or light microscope, and were accurately described by de Bary (1871, 1884), but on most species wax structure is only visible in the electron microscope.

THE FINE STRUCTURE OF THE PLANT CUTICLE

57

EW are often described as SCL, which does not distinguish between cuticular waxes embedded in the CM and EW located outside the CM. The terms ‘surface lipids’ and ‘superficial wax’ are also common in the literature. EW is defined here solely on the basis of its position on the leaf surface as wax, freely soluble or otherwise, deposited externally to the CP. This definition includes overtly crystalline and amorphous wax structures and any background wax film, structureless or otherwise, whether freely soluble or not. The question of whether EW is visible or otherwise in the scanning electron microscope (SEM; Neinhuis and Barthlott, 1996) is irrelevant. This definition is based on more than an arbitrary choice of boundary. There has long been evidence that wax embedded in the cuticle (intracuticular wax, CW) is chemically distinct from EW (Martin, 1960; Baker, 1977), and recent developments in methods of isolating the EW without contaminating the extract with CW have strengthened this view (Jetter et al., 2000). The quantity of EW may be as little as 1 μg cm−2 on leaves of some temperate annuals but the fan palm Copernica cerifera, the source of carnauba wax, and the ouricuri palm Syagurus coronata, both of arid north-eastern Brazil, have thick EW crusts weighing several milligrams per square centimetre that can be mechanically harvested by thrashing the dried leaves. Carnauba wax has the highest melting point of any natural wax, and is the most commercially important, used to give a hard, abrasion-resistant shine to products as diverse as cars, cabinets and candy. The Andean wax palms Ceroxylon spp. by contrast inhabit the cool, humid cloud forests at altitude in South America. Obligate submerged aquatic and amphibious plants have little or no EW, in which case their surfaces are fully wettable [e.g. Myriophyllum spp. (Hallam, 1982); Potamogeton crispus (Frost-Christensen, et al., 2003)]. Many mesophytic species of commercial importance including Lactuca sativa, P. vulgaris (Baker, 1982), B. vulgaris (Baker and Hunt, 1981) and N. tabacum (Jeffree et al., 1975; Jeffree, 1986) have no visible wax, but carry a measurable load nevertheless. Visible wax loads are absent on many weed species too, making them easy to wet with herbicide sprays (Rumex obtusifolia, Stellaria media and Myosotis arvensis; Baker and Bukovac, 1971). In the majority of species the EW layer is visible by electron microscopy (Juniper and Bradley, 1958; Amelunxen et al., 1967; Jeffree, 1986 and references therein). The wax loads on fruits are invariably heavier than those on the leaves of the same species, which in turn may carry different amounts of wax on their adaxial and abaxial surfaces. The chemical composition of the wax may differ from place to place on a plant, and the fine structure may vary correspondingly. Notable examples are the leaves of peas and wheat, which have different wax chemistry and wax ultrastructure on the adaxial and abaxial surfaces.

2.6.1

Epicuticular wax types

Classification of EW types on the evidence of SEM images was first attempted by Amelunxen et al. (1967), who extended de Bary’s original scheme (1871, 1884) based on observations using light microscopy (LM). Amelunxen et al. (1967)

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BIOLOGY OF THE PLANT CUTICLE

proposed five main classes: (1) (2) (3) (4) (5)

‘Wachskörnchen’ or grains, ‘Wachsstäbchen und –fäden’ or rods and filaments, ‘Wachsplättchen und –schuppen’ or plates and scales, ‘Wachsschichten und –krusten’ or films and crusts and ‘Flüssiger und schmieriger Wachsüberzug’ or fluid and greasy wax layers.

Each class contained two–five types making sixteen basic types. A further class dealt with massed accumulations of the grains, rods and filaments already included in their classes 1 and 2. Only waxes with discrete structures large enough to be resolvable in LM were included in de Bary’s (1871, 1884) classification. The fact that his classification stood the test of a technology capable of an order of magnitude better resolution is in one sense a remarkable testament to his powers of observation, but it is also attributable to the fact that simple paradigms for EW forms occur repeatedly at a range of size scales. Those gigantic rods he observed on S. officinarum are paralleled by ten-fold smaller rodlets on other species. It is now clear that many of the examples of rod and filament waxes (Rosa canina fruit, Tulipa gesneriana fruit and leaf, Eucalyptus globulus leaf) of Amelunxen et al. (1967) and Metcalfe and Chalk (1979) are in fact members of a more recently recognised tube-type (Hallam, 1967; Johnson and Jeffree, 1970). In the early SEM images of tube waxes, resolution was scarcely adequate to confirm the existence of a hollow centre in these tubes (see however Johnson and Jeffree, 1970; Jeffree, 1974a; Jeffree et al., 1976). Misclassification frequently arises out of inadequate image resolution (arguably the same is true for many reportedly non-lamellate cuticles), but there are many instances of clear misclassification in the literature even when resolution had been adequate, and classifications based on morphology alone are often unsound. Re-examination of figure 8 of Hallam and Juniper (1971) clearly shows that the ‘granules’ of wax on the surface of a fruit of Prunus sp. in a crisp micrograph of a TEM replica (supplied by D. Skene) are short curls and tubes overlying plate-like structures. The EW of ripe fruit of Prunus domestica contains about 48% of asymmetrical secondary alcohols, mostly 10-nonacosanol (Holloway et al., 1976), now known to produce hollow tubular epicuticular crystals (Jeffree, 1974a; Jeffree et al., 1975, 1976; Jetter and Riederer, 1994). The most commonly encountered crystal morphologies in plant EW remain amorphous films, grains or granules, plates (simple or crenate, polygonal or rounded or spiky, prostrate or erect), filaments, rods, generally terete in cross-section, but other shapes are possible: tubes with a hollow centre; elongated flattened ribbons with various forms of edge decoration; and plates (Table 2.3), and there are good reasons to take a conservative approach in classifying EW types. First, although attempts have been made to describe and classify these structures over many decades, our understanding of the morphological range is still limited by poor resolution of the morphology in many cases, and by a crude vocabulary for description of variations

59

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

Species

The main epicuticular wax morphological types, and their distribution among species Family or higher taxon

Amorphous films, or no visible wax Acer mono Aceraceae Aesculus hippocastanum Hippocastanaceae Arabidopsis thaliana Brassicaceae Arabidopsis thaliana Brassicaceae Beta vulgaris Chenopodiaceae Blechnum capense Blechnaceae Bromus interruptus Poaceae Chamerion angustifolium Onagraceae Chenopodium album Chenopodiaceae Chionochloa rigida Poaceae Chrysanthemum morifolium Asteraceae Citrus limon cv. Ranunculaceae ‘Adamopolou’ Clematis vitalba Ranunculaceae Coffea arabica Rubiaceae Dieffenbachia maculata Araceae Euphorbia cerifera Euphorbiaceae Fagus sylvatica Fagaceae Gaultheria depressa Ericaceae Gerbera jamesonii Asteraceae Gnetum gnemon. Gnetaceae Griselinia littoralis Cornaceae Heliconia densiflora Heliconiaceae Hydrangea hortensis Hydrangeaceae Hydrocotyle bonariensis Apiaceae Juncus inflexus Cyperaceae Lactuca sativa Asteraceae Lycopersicon esculentum Solanaceae Magnolia grandiflora L. Magnoliaceae Malus hupehensis Rosacaeae Myosotis arvensis Boraginaceae Myriophyllum spp. Haloragaceae Nicotiana tabacum Solanaceae Nothofagus solandri var. Nothofagaceae Cliffortioides Nothofagus spp. Nothofagaceae Phaseolus vulgaris Fabaceae Potamogeton crispus Potamogetonaceae Prunus laurocerasus Rosacaeae Prunus laurocerasus Rosacaeae Prunus persica cv. ‘Red Rosacaeae Haven’ Rhododendron ponticum Ericaceae Rumex obtusifolius Polygonaceae

Organ

Reference

LAb L L Sepal LAd L A P L LAd L L

Barthlott and Neinhuis (1997) Juniper and Bradley (1958) Jenks et al. (2002) Figure 2.12a Baker and Hunt (1981) Hall and Burke (1974) Jeffree, unpublished Figure 2.14f Baker and Bukovac (1971) Hall and Burke (1974) Sutter (1985) Jeffree et al. (1975); Jeffree (1986)

L LAd L L LAd LAd L L LAd LAd L L L L F, L LAd LAd L SL L LAd

Baker (1982) Silva Fernandes (1965a,b) Sutter (1985) Juniper and Jeffree (1983) Barthlott and Neinhuis (1997) Hall and Burke (1974) Sutter (1985) Barthlott and Neinhuis (1997) Hall and Burke (1974) Barthlott and Neinhuis (1997) Silva Fernandes (1965a,b) Barthlott et al. (1998) Juniper (1960) Baker (1982) Jeffree et al. (1975); Jeffree (1986) Barthlott and Neinhuis (1997) Baker and Hunt (1981) Baker and Bukovac (1971) Hallam (1982) Jeffree et al. (1975); Jeffree (1986) Hall and Burke (1974)

LAd L SL L LAd LAd

Hall and Burke (1974) Baker (1982) Hallam (1982) Davis (1971) Jetter et al. (2000) Jeffree et al. (1975); Jeffree (1986)

LAb L

Holloway and Jeffree (2005) Juniper and Bradley (1958); Baker and Bukovac (1971) Continued

60

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued

Species

Family or higher taxon

Organ

Reference

Spathiphyllum wallissii Spinacia oleracea Stellaria media Tamus communis Taraxacum officinale Tilia cordata Trifolium repens Ulex europaeus Vicia faba Vitis vinifera

Araceae Chenopodiaceae Caryophyllaceae Dioscoreaceae Asteraceae Tiliaceae Fabaceae Fabaceae Fabaceae Vitaceae

LAd L L L L LAd LAb S LAd LAb

Sutter (1985) Baker (1982) Baker and Bukovac (1971) Baker (1982) Baker and Bukovac (1971) Schreiber (1990) Holloway (1971); Baker (1982) Zabkiewicz and Gaskin (1978) Holloway and Jeffree (2005) Baker (1982)

Aegicerataceae Rutaceae

L F

Granules Aegiceras corniculatum Citrus limon cv. ‘Adamopolou’ Citrus paradisi cv. ‘Marsh seedless’ Conocephalum conicum Exomotheca bullata Hedera helix Helianthus annuus Lunularia cruciata Macaranga triloba Marchantia paleacea Marchantia polymorpha Pistacia vera Plagiochasma rupestre Rebouilia hemisphaerica Targiona hypophylla

Rutaceae

JV

Barthlott et al. (1998) Jeffree et al. (1975, 1976); Baker (1982) Fahn et al. (1974)

Bryophyta Bryophyta Araliaceae Asteraceae Bryophyta Euphorbiaceae Bryophyta Bryophyta Anacardiaceae Bryophyta Bryophyta Bryophyta

G G LAd LAd G S G G LAd G G G

Schönherr and Ziegler (1975) Schönherr and Ziegler (1975) Martin and Juniper (1970) Hallam and Juniper (1971) Schönherr and Ziegler (1975) Markstädter et al. (2000) Schönherr and Ziegler (1975) Schönherr and Ziegler (1975) Baker (1982) Schönherr and Ziegler (1975) Schönherr and Ziegler (1975) Schönherr and Ziegler (1975)

Soft waxes Crataegus prunifolia Malus sp.

Rosaceae Rosaceae

F F

Metcalfe and Chalk (1979) Metcalfe and Chalk (1979), Martin and Juniper (1970)

Barthlott et al. (1998) Figure 2.12b Frölich and Barthlott (1988) Meusel et al. (1999) Meusel et al. (1999) Frölich and Barthlott (1988) Meusel et al. (1999) Meusel et al. (1999) Meusel et al. (1999) Meusel et al. (1999); Frölich and Barthlott (1988) Neinhuis and Barthlott (1996) Neinhuis and Barthlott (1996)

Rodlets (a) Aristolochia Type: Transversely ridged rodlets Actinidia melanandra Ammophila arenaria Aristolochia Aristolochia gigantea Centranthus ruber Fritillaria meleagris Fritillaria pallidiflora Gypsophila acutifolia Laurus nobilis Leucojum aestivum

Actinidiaceae Poaceae Aristolochiaceae Aristolochiaceae Valerianaceae Liliaceae Liliaceae Caryophyllaceae Lauraceae Liliaceae

L LAd L L L L L L L L

Liriodendron sp. Magnolia sp.

Magnoliaceae Magnoliaceae

LAb LAb

61

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

Continued

Species

Family or higher taxon

Organ

Reference

Monodora sp. Nicotiana glauca Osmunda regalis Paeonia mlokosewitchii Paeonia officinalis Williamodendron quadrillocellatum Brassica napus Rigo mutant

Annonaceae Solanaceae Osmundaceae Paeoniaceae Paeoniaceae Lauraceae

LAb L L L L LAb

Neinhuis and Barthlott (1996) Meusel et al. (1999) Jetter and Riederer (1999b, 2000) Meusel et al. (1999) Meusel et al. (1999) Neinhuis and Barthlott (1996)

Brassicaceae

L

Brassica oleracea mutant gl2

Brassicaceae

L

Galanthus nivalis

Amaryllidaceae

L

Jeffree et al. (1975, 1976); Baker (1982) Jeffree et al. (1975, 1976); Baker (1982) Koch et al. (2004)

(b) Strelitzia Type: Longitudinally striated rods without transverse ridges Arundinaria sp. Benincasa hispida Carex flacca Colletia cruciata Dendrocalamus giganteus Eryngium sp Hedychium flavum Heliconia collinsiana Maranta leuconeura Musa paradisaica

Poaceae Cucurbitaceae Cyperaceae Rhamnaceae Poaceae Apiaceae Zingiberaceae Heliconiaceae Marantaceae Musaceae

S F L L L L L L LAd L

Phenacospermum guianense Phragmites australis Saccharum officinarum Sorghum bicolor Strelitzia reginae Syagrus coronata Typha angustifolia Typha elephantina Typha latifolia

Strelitziaceae Poaceae Poaceae Fabaceae Strelitziaceae Arecaceae Typhaceae Typhaceae Typhaceae

L L LS L LAb LAb L L L

Jeffree et al. (1976) Meusel et al. (1994) Frölich and Barthlott (1988) Barthlott et al. (1998) Frölich and Barthlott (1988) Meusel et al. (1994) Frölich and Barthlott (1988) Barthlott et al. (1998) Sutter (1985) Jeffree et al. (1975, 1976); Baker (1982) Barthlott et al. (1998) Jeffree, unpublished de Bary (1871); Jeffree et al. (1976) Blum (1975) Meusel et al. (1994) Machado and Barros (1995) Djebrouni (1989) Djebrouni (1989) Djebrouni (1989)

Butomaceae Hyacinthaceae Marantaceae Aizoaceae

L L L L

Frölich and Barthlott (1988) Frölich and Barthlott (1988) Sutter (1985) Barthlott et al. (1998)

Thymelaceae

L

Barthlott et al. (1998)

Rutaceae

JV

Fahn et al. (1974)

Epacridaceae

LAb

Figure 2.12f

(c) Simple Rod Type Butomus umbellatus Galtonia candicans Maranta leuconeura Sceletium compactum (d) Polygonal Rod Type Daphne tangutica Filaments (a) Simple filament or thread Citrus paradisi cv. ‘Marsh seedless’ Cyathodes colensoi

Continued

62

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued Family or higher taxon

Organ

Reference

Drosera burmanni Leptospermum laevigatum Pisum sativum

Droseraceae Rosaceae

L L

Barthlott et al. (1998) Hallam and Juniper (1971)

Fabaceae

LAb

Saelania glaucescens Solanum tuberosum Sorghum bicolor

Musci Solanaceae Poaceae

L Tu LS

Juniper and Bradley (1958); Juniper (1959); Martin and Juniper (1970); Jeffree et al. (1975, 1976) Haas (1982) Hayward (1974) Atkin and Hamilton (1982)

Species

(b) Flattened filament or ribbon Macaranga hypoleuca Pisum sativum

Euphorbiaceae Leguminosae

S LAb, P, St, T

Markstädter et al. (2000) Juniper (1960); Baker and Holloway (1971); Holloway (1971); Jeffree et al. (1975, 1976) Ihlenfeldt and Hartmann (1982)

Wooleya farinose

Mesembryanthemaceae

L

Euphorbiaceae

S

Markstädter et al. (2000)

(c) Chiral filament or ribbon Macaranga lamellate

(d) Filament or ribbon network Acer mono Acer pseudoplatanus

Aceraceae Aceraceae

LAb LAb

Fragaria ananassa cv. ‘Cambridge Favourite’ Fragaria ovalis Potentilla fruticosa Rosa sp.

Rosaceae

LAb

Neinhuis and Barthlott (1996) Holloway (1971); Wilson (1984) Mackerron (1976)

Rosaceae Rosaceae Rosaceae

LAd L LAb

Baker (1982) Rentschler (1971) Figure 2.14e

Brassica type, columns and plates Arabidopsis thaliana Brassicaceae

S

Brassica napus Brassica napus

Brassicaceae Brassicaceae

L L

Brassica oleracea var. capitata Brassica oleracea var. capitata Brassica oleracea var. gemmifera Bupleurum salicifolium Centranthus ruber Clarkia elegans

Brassicaceae

LAd

Brassicaceae

L

Brassicaceae

L

Apiaceae Valerianaceae Onagraceae

L L L

Crambe maritima

Brassicaceae

L

Teusink et al. (2002); Jenks et al. (2002) Holloway and Jeffree (2005) Armstrong and Whitecross (1976) Martin and Juniper (1970); Figures 2.5, 4.28 Martin and Juniper (1970); Hallam and Juniper (1971) Jeffree et al. (1975, 1976); Holloway et al. (1976) Barthlott et al. (1998) Baker (1982) Hunt et al. (1976); Baker (1982) Rentschler (1971)

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

63

Continued

Species

Family or higher taxon

Organ

Reference

Erythronium rostratum Polygala myrtifolia Thellungiella halophila Thellungiella parvula

Liliaceae Polygalaceae Brassicaceae Brassicaceae

L L LAd LAd

Frölich and Barthlott (1988) Rentschler (1971) Jenks et al. (2002) Jenks et al. (2002)

Hanover and Reicosky (1971) Jeffree et al. (1975, 1976); Holloway et al. (1976) Jeffree et al. (1975, 1976); Holloway et al. (1976) Rentschler (1971) Rentschler (1971); Holloway and Jeffree (2005) Neinhuis and Jetter (1995) Barthlott and Theisen (1995) Attiwill and Clayton-Greene (1984) Jeffree et al. (1975, 1976); Holloway et al. (1976) Jeffree et al. (1975, 1976); Holloway et al. (1976) Jeffree et al. (1975, 1976); Holloway et al. (1976) Rentschler (1971) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Silva Fernandes (1964); Jeffree et al. (1975, 1976); Holloway et al. (1976) Jeffree et al. (1975, 1976); Holloway et al. (1976) Neinhuis and Jetter (1995) Jeffree et al. (1975, 1976); Holloway et al. (1976); Rentschler (1973) Barthlott et al. (1996) Jeffree et al. (1975, 1976) Hanover and Reicosky (1971) Jeffree et al. (1975, 1976); Holloway et al. (1976) Hanover and Reicosky (1971) Hanover and Reicosky (1971) Jeffree (1974b); Jeffree et al. (1975, 1976) Hanover and Reicosky (1971)

Tubes (a) Asymmetrical secondary alcohol tubes Abies concolor Agathis australis

Pinaceae Pinaceae

L L

Aquilegia alpinum

Ranunculaceae

L

Aquilegia formosa Aquilegia vulgaris

Ranunculaceae Ranunculaceae

L L

Atrichum undulatum Berberis Callitris columnaris

Polytrichales Berberidaceae Cupressaceae

Cap L LAb

Chamaecyparis lawsoniana Chamaecyparis obtusa

Cupressaceae

L

Cupressaceae

L

Chelidonium majus

Papaveraceae

L

Cotinus coggygria Dawsonia beccarii Dawsonia beccarii Exochorda racemosa

Anacardiaceae Polytrichaceae Polytrichales Rosaceae

L Cap Cap L

Ginkgo biloba

Ginkgoaceae

L

Oligotrichum hercynicum Papaver somniferum

Polytrichaceae Papaveraceae

Cap L

Papaver somniferum Picea abies Picea glauca Picea pungens

Papaveraceae Pinaceae Pinaceae Pinaceae

Cap L L L

Picea pungens Picea pungens var. glauca Picea sitchensis

Pinaceae Pinaceae Pinaceae

L L L

Picea sitchensis

Pinaceae

L

Continued

64

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued

Species

Family or higher taxon

Organ

Reference

Pinus balfouriana Pinus nigra Pinus nigra var. maritima Pinus radiata

Pinaceae Pinaceae Pinaceae Pinaceae

L L L L

Pinus strobus Pinus sylvestris Pinus, 51 species Pogonatum belangeri Pogonatum urnigerum Pogonatum urnigerum Polytrichum commune Polytrichum formosum Polytrichum juniperinum Polytrichum juniperinum Prunus domestica

Pinaceae Pinaceae Pinaceae Polytrichales Polytrichaceae Polytrichales Polytrichales Polytrichales Polytrichaceae Polytrichales Rosaceae

L L L Cap Cap Cap Cap Cap Cap Cap L

Prunus domestica cv. ‘d’Agen’ Prunus sp.

Rosaceae

F

Jeffree, unpublished Hanover and Reicosky (1971) Campbell (1972) Rook et al. (1971); Leyton and Juniper (1963) Hanover and Reicosky (1971) Jeffree et al. (1975,1976) Yoshie and Sakai (1985) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Neinhuis and Jetter (1995) Jeffree et al. (1975, 1976); Holloway et al. (1976) Bain and McBean (1967, 1969)

Rosaceae

F

Pseudotsuga menziesii

Pinaceae

L

Rhus cotinus atropurpurea Rosa sp. cv. ‘Baccara’

Anacardiaceae

L

Rosaceae

S

Thalictrum flavum Tropaeolum majus

Ranunculaceae Geraniaceae

L L

Tropaeolum speciosum Tulipa gesneriana

Geraniaceae Liliaceae

L L

Tulipa kaufmanniana Waxes of apple, hawthorn and rose also contain 10-nonacosanol and other asymmetrical secondary alcohols

Liliaceae Rosaceae

L L

D. Skene in Hallam and Juniper (1971); Martin and Juniper (1970) Campbell (1972); Thair and Lister (1975); Lister and Thair (1981) Silva Fernandes (1964); Jeffree et al. (1975, 1976); Holloway et al. (1976) Silva Fernandes (1965b); Holloway (1971); Baker and Holloway (1971); Baker (1982) Barthlott et al. (1996) Rentschler (1971); Jeffree et al. (1975, 1976); Brunegger et al. (1982) C.E. Jeffree Rentschler (1971); Jeffree et al. (1975, 1976); Holloway et al. (1976) Johnson and Jeffree (1970) Wollrab (1969)

(b) Asymmetrical secondary alkandiol tubes Nelumbo nucifera

Nelumbonaceae

L

Barthlott et al. (1996); Barthlott and Neinhuis (1997); Neinhuis and Barthlott (1997)

Poaceae Myrtaceae

Gl, Lm L

Baum and Hadland (1975) Hallam (1967); Hallam and Chambers (1970); Hallam and Juniper (1971)

(c) β-diketone tubes 21 of 27 sp. of Avena 77 species of Eucalyptus

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

65

Continued

Species

Family or higher taxon

Organ

Reference

Andromeda polifolia Chrysanthemum frutecens Dianthus caryophyllus Eragrostis curvula Eucalyptus camaldulensis Eucalyptus glaucescens Eucalyptus globulus

Ericaceae Asteraceae Caryophyllaceae Poaceae Myrtaceae Myrtaceae Myrtaceae

LAb L L L L L L

Eucalyptus gunnii Eucalyptus nova anglica Eucalyptus perriniana Eucalyptus viminalis Festuca glauca Glumes and lemmas of grasses Hordeum vulgare

Myrtaceae Myrtaceae Myrtaceae Myrtaceae Poaceae Poaceae

L L LAd L L Gl, Lm

Jeffree (1986) Rentschler (1971) Jeffree et al. (1975, 1976) Leigh and Matthews (1963) Jeffree et al. (1975, 1976) Jeffree et al. (1975, 1976) Hallam (1967), Hallam and Juniper (1971); Jeffree et al. (1975, 1976) Jeffree (1974b) Jeffree et al. (1975, 1976) Hallam (1967) Jeffree et al. (1975, 1976) Jeffree et al. (1975, 1976) Baum and Hadland (1975)

Poaceae

F LS

Leymus arenarius Poa colensoi Triticum aestivum

Poaceae Poaceae Poaceae

LAd L FLAb

Vaccinium ashei

Ericaceae

L

von Wettstein-Knowles (1974); Jeffree et al. (1975, 1976); Baker (1982) Meusel et al. (2000) Hall et al. (1965) Netting and von Wettstein-Knowles (1973); von Wettstein-Knowles (1974); Jeffree et al. (1975, 1976) Freeman et al. (1979)

Boraginaceae

L

Jetter and Riederer (1999a)

Araceae Columelliaceae

L L

Barthlott et al. (1998) Barthlott et al. (1998)

Fumariaceae Hostaceae

L LAd

Barthlott et al. (1998) Figure 2.14d

Open spirals Buxus sempervirens Cerinthe major Chrysanthemum segetum

Buxaceae Boraginaceae Asteraceae

L L L

Lonicera korolkovii Stapf Papaver somniferum Pogonatum rubenti-viride Sedum telephium

Caprifoliaceae Papaveraceae Polytrichales Crassulaceae

L Cap Cap L

Barthlott et al. (1998); Meusel et al. (1999) Figure 2.14c Juniper and Bradley (1958); Martin and Juniper (1970); Barthlott et al. (1998); Meusel et al. (2000) Barthlott et al. (1998) Jetter (1993); Jetter and Riederer (1994) Neinhuis and Jetter (1995) Rentschler (1971)

Dendritic plates Allium cepa cv. ‘Ailsa Craig’

Liliaceae

L

Jeffree et al. (1975, 1976); Jeffree (1986)

(d) delta-lactone tubes Cerinthe minor (e) uncharacterized tubes Amorphophallus maximus Columellia oblonga Ruiz and Pav Corydalis cava Hosta sieboldiana Hook. Engl

Continued

66

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued

Species

Family or higher taxon

Organ

Reference

Allium porrum

Liliaceae

L

Brassica oleracea var. caulo-rapa Brassica oleracea var. gemmifera , high temp and humidity Brassica oleracea var. capitata Euphorbia lathyrus

Brassicaceae

L

Jeffree et al. (1975, 1976); Baker (1982) Burkhardt et al. (2001)

Brassicaceae

L

Baker (1974, 1982)

Brassicaceae

L

Figure 2.12d

Euphorbiaceae

L

Euphorbia peplus Lupinus albus Lupinus luteus

Euphorbiaceae Fabaceae Fabaceae

LAb LAd L

Jeffree et al. (1975, 1976); Jeffree (1986) Holloway (1971) Martin and Juniper (1970), Figure 8.3 Rentschler (1971)

Anarthia scabra Aristolochia duriar Aristolochia elegans Barbacenia tubulosa Carludovica palmata Colochasia esculenta Dawsonia superba Elegia verticciaris Equisetum arevense Gossypium hirstutum Habrop et alum dawei Hypericum hircinum Kensitia pillansii Kleinia articulata

Anarthriaceae Aristolochiaceae Aristolochiaceae Velloziaceae Cyclanthaceae Araceae Polytrichales Restionaceae Equisetaceae Malvaceae Dioncophyllaceae Clusiaceae Mesembryanthemaceae Asteraceae

L L L L L L GL L S L L L L LAd

Lecythis chartacea Nicolaia elatior Oxalis hedysaroides Paeonia suffruticosa Pandanus montanus Polygala myrtifolia Polytrichum juniperinum Prosopis spp.

Lecythidaceae Zingiberaceae Oxalidaceae Paeoniaceae Pandanaceae Polygalaceae Polytrichaceae Fabaceae

L L L L L L GL L

Rosa cv. ‘Baccara’

Rosaceae

L

Simmondsia chinensis Vitis vinifera cv. ‘Siebel’ Vitis vinifera cv. ‘Sultana’, ‘Kishmish’, ‘Thompson seedless’

Buxaceae Vitaceae Vitaceae

L F F

Simple plates (a) Uncharacterised plates Frölich and Barthlott (1988) Rentschler (1971) Rentschler (1971) Frölich and Barthlott (1988) Frölich and Barthlott (1988) Barthlott and Neinhuis (1997) Troughton and Sampson (1973) Frölich and Barthlott (1988) Baker and Holloway (1971) Hallam and Juniper (1971) Barthlott et al. (1998) Rentschler (1971) Ihlenfeldt and Hartmann (1982) Martin and Juniper (1970); Figure 4.24 Barthlott et al. (1998) Frölich and Barthlott (1988) Rentschler (1971b) Rentschler (1971b) Frölich and Barthlott (1988) Rentschler (1971b) Neinhuis and Jetter (1995) Bleckmann and Hull (1975); Hull et al.(1979) Silva Fernandes (1965b); Holloway (1971); Baker and Holloway (1971) Gülz and Hangst (1983); Gülz (1986) Jeffree et al. (1975) Possingham (1972); Baker (1982)

67

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

Continued Family or higher taxon

Organ

Reference

202 species of Eucalyptus

Myrtaceae

L

Acacia pychantha Acacia sp. Agropyron repens Avena fatua Avena, glumes of 15 species Avena sativa Avena sativa cv. ‘Black supreme’ Baptisia australis Bromus interruptus Calligonum comosum Chamerion angustifolium Chenopodium album Dactylis glomerata Eucalyptus cloeziana

Fabaceae Fabaceae Poaceae Poaceae Poaceae

LAb LAd LAd L Gl

Hallam (1967); Hallam and Chambers (1970); Hallam and Juniper (1971) Baker (1982) Neinhuis and Barthlott (1997) Coupland and Caseley (1975) Whitehouse et al. (1982) Baum and Hadland (1975)

Poaceae Poaceae

L L

Rentschler (1971b) Jeffree et al. (1975, 1976)

Fabaceae Poaceae Polygonaceae Onagraceae Chenopodiceae Poaceae Myrtaceae

L L S L L LAb L

Rentschler (1971b) Jeffree, unpublished Lyshede (1977b) Rentschler (1971); Figure 2.18b Taylor et al. (1981) Holloway (1971) Martin and Juniper (1970); Figure 4.26 Jeffree et al. (1975, 1976)

Species (b) Primary alcohol plates

Eucalyptus coccifera forma Eucalyptus microcarpa Eucalyptus miniata Eucalyptus pauciflora

Myrtaceae

L

Myrtaceae Myrtaceae Myrtaceae

L L L

Eucalyptus platypus Festuca arundinacea Hordeum sativum

Myrtaceae Poaceae Poaceae

28d L L LAd

Hordeum vulgare Hordeum vulgare cv.‘Proctor’ Laburnum sp. Oryza sativa Pisum sativum

Poaceae Poaceae

L L

Fabaceae Poaceae Fabaceae

LAd LAd LAd

Fabaceae

LAd

Poaceae Fagaceae Fagaceae Fabaceae

L LAb LAb LAd

Pisum sativum cv. ‘Meteor’, ‘Kelvedon Wonder’, ‘Alaska’ Poa nemoralis Quercus pubescens Quercus robur Robinia sp.

Attiwill and Clayton-Greene (1984) Hallam and Juniper (1971) Jeffree (1974a,b); Jeffree et al. (1975, 1976) Knight et al. (2004) Pitcairn et al. (1986); Figure 2.12c Martin and Juniper (1970) Figure 2.4; Juniper and Bradley (1958) Rentschler (1971) Jeffree et al. (1975, 1976) Neinhuis and Barthlott (1997) Mendgen (1996) Juniper (1959); Martin and Juniper (1970); Jeffree et al. (1975, 1976); Holloway et al. (1976) Juniper (1959); Martin and Juniper (1970); Jeffree et al. (1975, 1976); Holloway et al. (1976) Jeffree et al. (1975, 1976) Jeffree and van Gardingen (1993) Schreiber (1990) Neinhuis and Barthlott (1997) Continued

68

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued

Species

Family or higher taxon

Organ

Reference

Secale cerale cv. ‘Lovaspatonia’ Sorghum bicolor Trifolium pratense Trifolium repens

Poaceae

L

Jeffree et al. (1975, 1976)

Poaceae Fabaceae Fabaceae

LAd L LAd

Trifolium repens Trifolium repens

Fabaceae Fabaceae

LAd LAd

Triticum aestivum

Poaceae

L

Zea mays Zea mays

Poaceae Poaceae

LAd L

Atkin and Hamilton (1982) Juniper and Bradley (1958) Hall and Donaldson (1963); Hall (1967a,b); Rentschler (1973) Hall and Donaldson (1963) Holloway, in Martin and Juniper (1970) Figure 2.6 Netting and von Wettstein-Knowles (1973); von Wettstein-Knowles (1974); Jeffree et al. (1975, 1976) Martin and Juniper (1970) Figure 4.23 Rentschler (1971)

Fagaceae

L

Markstädter (1994)

Brassicaceae

L

Holloway and Jeffree (2005)

Brassicaceae

L

Baker (1974); Jeffree et al. (1975, 1976)

Liliaceae Liliaceae Euphorbiaceae Euphorbiaceae Euphorbiaceae Euphorbiaceae Euphorbiaceae Proteaceae Cucurbitaceae Oleaceae

L L LAb LAd L LAd LAb

F

Scott et al. (1958) Jeffree et al. (1975) Verdus (1973) Verdus (1973) Rentschler (1971) Holloway (1971) Verdus (1973) Barthlott et al. (1998) Barthlott et al. (1998) Baker (1982)

Anthericum liliago L. Alstromeria aurantiaca Anabasis articulata Asparagus retrofractus

Asphodelaceae Alstromeriaceae Chenopodiaceae Asparagaceae

L L S Phy

Bupleurum ranunculoides Burmannia biflora Calibanthus hookeri Camassia cusickii

Apiaceae Burmanniaceae Dracaenaceae Hyacinthaceae

L L L L

Chenopodium album Convallaria majalis

Chenopodiceae Convallariaceae

L L

(c) Aldehyde plates Fagus sylvatica (d) Alkane plates Brassica napus mutant nilla

(e) Symmetrical secondary alcohol plates Brassica oleracea mutant gl4 (f) Crenate and dendritic plates Allium cepa Allium cepa cv. ‘Ailsa Craig’ Andrachne telephioides Euphorbia characias Euphorbia milii Euphorbia peplus Euphorbia serrata Grevillea bipinnatifida Odosicyos sp. Olea europea (g) Convallaria plate types Frölich and Barthlott (1988) Barthlott et al. (1998) Lyshede (1977b) Frölich and Barthlott (1988); Barthlott et al. (1998) Figure 2.18c Frölich and Barthlott (1988) Frölich and Barthlott (1988) Frölich and Barthlott (1988); Barthlott et al. (1998) Taylor et al. (1981) Frölich and Barthlott (1988); Barthlott et al. (1998)

69

THE FINE STRUCTURE OF THE PLANT CUTICLE Table 2.3

Continued Family or higher taxon

Organ

Reference

Eriospermum triphyllum Hemerocallis citrina Herpolirion novae-zelandiae Herreria montevidensis Melaleuca hypericifolia Narthecium asiaticum

Eriospermaceae Hemerocallidaceae Doryanthaceae

L L L

Frölich and Barthlott (1988) Frölich and Barthlott (1988) Frölich and Barthlott (1988)

Herreriaceae Myrtaceae Melanthiaceae

L LAd L

Phormium cookianum

Phormiaceae

L

Prenia sladeniana Stypandra glauca Uvularia floridiana Watsonia bulbifera Yucca filamentosa

Mesembryanthemaceae Dianellaceae Colchicaceae Iridaceae Agavaceae

L L L L LAb

Frölich and Barthlott (1988) Neinhuis and Barthlott (1997) Frölich and Barthlott (1988); Barthlott et al. (1998) Frölich and Barthlott (1988); Barthlott et al. (1998) Ihlenfeldt and Hartmann (1982) Frölich and Barthlott (1988) Frölich and Barthlott (1988) Frölich and Barthlott (1988) Frölich and Barthlott (1988); Neinhuis and Barthlott (1997)

Species

(h) Orientated plates, various types Amherstia nobilis Erythroxylum coca Eucalyptus camaldulensis

Fabaceae Erythroxylaceae Myrtaceae

LAb L L

Eucalyptus pauciflora

Myrtaceae

L

Hypericum bucklei Acacia sp. Calliandra haematoma Bert Benth. Laburnum sp. Robinia sp. Quercus pubescens Quercus robur Trifolium repens

Hypericaceae Fabaceae Fabaceae

LAd

Fabaceae Fabaceae Fagaceae Fagaceae Fabaceae

LAd LAd L L L

Benthamia alyxifolia Nepenthes alata Nepenthes rufescens

Loranthaceae Nepenthaceae Nepenthaceae

Pt Pt

Prosopis tamarugo

Fabaceae

LAd

Tilia magnifica

Tiliaceae

LAb

Mixed tubes and plates 36 species of Eucalyptus

Myrtaceae

L

Neinhuis and Barthlott (1996) Rentschler (1971) Jeffree (1974a,b); Jeffree et al. (1975, 1976); Figure 2.12e Jeffree (1974a,b); Jeffree et al. (1975, 1976) Barthlott et al. (1998) Neinhuis and Barthlott (1997) Barthlott et al. (1998) Neinhuis and Barthlott (1997) Neinhuis and Barthlott (1997) Jeffree and van Gardingen (1993) Schreiber (1990) Hall and Donaldson, (1963); Hall (1967a); Rentschler (1973)

(i) Miscellaneous plate types Barthlott et al. (1998) Riedel et al. (2003) Juniper and Burras (1962); Martin and Juniper (1970); Juniper (1986) Bleckmann and Hull (1975); Hull et al. (1979) Neinhuis and Barthlott (1997) Hallam (1967); Hallam and Chambers (1970); Hallam and Juniper (1971) Continued

70

BIOLOGY OF THE PLANT CUTICLE

Table 2.3 Continued

Species

Family or higher taxon

Organ

Reference

Brassica napus Lupinus albus Malephora spp. Pinus sylvestris

Brassicaceae Fabaceae Aizoaceae Pinaceae

L LAd L L

Rhus cotinus atropurpurea Farinaceous Cheilanthus sp. Notholaena sp. Pityrogramma triangularis Primula spp. Pteris sp.

Anacardiaceae

S

Rentschler (1973) Juniper (1960) Ihlenfeldt and Hartmann (1982) Leyton and Juniper (1963); Martin and Juniper (1970); Figure 4.27 Baker (1982)

Adiantaceae Cryptogrammaceae Adiantaceae

L L L

Primulaceae Pteridaceae

LAb L

de Bary (1871); Wollenweber (1982) de Bary (1871); Wollenweber (1982) de Bary (1871); Chance and Arnott (1981) de Bary (1871); Blasdale (1947) de Bary (1871); Wollenweber (1982)

Arecaceae Arecaceae Arecaceae Arecaceae Arecaceae Cactaceae Asclepiadaceae Myrtaceae Euphorbiaceae Salicaceae Euphorbiaceae Euphorbiaceae Arecaceae Combretaceae

S L L L L L L L L S SC L LAd L

Martin and Juniper (1970) Martin and Juniper (1970) Martin and Juniper (1970) Barthlott et al. (1998) Martin and Juniper (1970) Barthlott et al. (1998) Barthlott et al. (1998) Knight et al. (2004) Juniper and Jeffree (1983) Juniper and Southwood (1986) Martin and Juniper (1970) Martin and Juniper (1970) Machado and Barros (1995) Barthlott et al. (1998)

Substantial wax crusts Ceroxylon andicola Ceroxylon andicola Copernica cerifera Copernica cowellii Copernica hospita Copiapoa cinerea Cynanchum sarcostemma Eucalyptus platypus Euphorbia cerifera Salix alba agentea Sapium sepiferum Sapium sepiferum Syagrus coronata Terminalia cf seyrigii

Abbreviations used: A = anther, Cap = capsule, F = fruit, FLAb = flag leaf abaxial surface, FLS = flag leaf sheath, G = gametophyte, Gl = glume, JV = juice vesicles, L = leaf, LAb = abaxial leaf surface, LAd = adaxial leaf surface, Lm = lemma, LS = leaf sheath, P = pod, Phy = phylloclade, Pt = pitcher, S = stem, SC = seed coat, St = stipule, T = trichome, Tu = tuber.

in the shape and size of objects in general. The range of types recognised by de Bary (1871, 1884) and Amelunxen et al. (1967) very nearly consumes the available words in the English language for describing such structures, reflecting the fact that we discriminate between comparatively few basic shapes. Second, our understanding of the relationships between wax morphology and the underlying chemical and genetic basis for specifying it is still incomplete, although substantial progress has been made in the past 35 years. Superficial classification based on morphological characters alone without understanding of the underlying chemistry will lead to errors in interpretation of taxonomy and phylogeny. A common trap, for example, is to assume that supposedly amorphous waxes are fundamentally different from

THE FINE STRUCTURE OF THE PLANT CUTICLE

71

obviously crystalline ones. It is therefore strongly recommended that morphologybased classifications should be used informally and with circumspection.

2.6.2

Chemical and structural classification of EW

Predominantly, EW are derivatives of n-acyl alkanes with chain lengths in the range C16–C35 (Chapter 4). The hydrocarbon chains may have substituted groups in terminal (fatty acids, primary alcohols, aldehydes) or mid-chain positions (β-diketones, secondary alcohols). With some exceptions (estolides of gymnosperm waxes; von Rudloff, 1959), and polymeric aldehydes in the waxes of S. officinarum (Lamberton and Redcliffe, 1959; Haas et al., 2001) and Nepenthes (Riedel et al., 2003) the constituents of plant EW are freely soluble in non-polar solvents such as hexane, benzene, chloroform and diethyl ether, but insoluble in water. The more polar constituents such as fatty acids are sparingly soluble in the polar solvents ethanol and ethyl acetate. The detailed description of the range and distribution of the compounds that have been detected in EW is outside the scope of this chapter, but the main constituent compound classes are summarised in Table 2.4. A detailed treatment of wax chemistry is given in Chapter 4. EW often fall into classes by virtue of having a single predominating constituent compound class, and it is commonly noted that such waxes have a characteristic structural type. The aliphatic constituents of the EW of most grass and Eucalyptus species fall into one of two types: those in which the primary alcohols hexacosanol or octacosanol predominate in the wax and those in which β-diketones hentriacontan-14,16-dione or tritriacontan-12,14-dione predominate (Tables 2.3 and 2.4). Sometimes as in wheat (Triticum aestivum) the waxes rich in primary alcohols (1-octacosanol) and β-diketones (tritriacontan-16,18-dione) may both be present, but are usually segregated into distinct regions of the flag leaf and other organs such as glumes and lemmas (Tulloch, 1973; Baum and Hadland, 1975; Baum et al., 1989). In these groups, the primary alcohol-rich waxes almost always have a simple plate morphology (Figures 2.12c,e and 2.13c) while the β-diketone-rich waxes are tubular, plates usually being absent (Figures 2.13a,b and 2.14d). Platetype waxes dominated by primary alcohols are widespread in the Fabaceae (Acacia, Trifolium, Pisum Figure 2.15a) and Myrtaceae (Eucalyptus, Figure 2.12e), and also appear frequently in the Poaceae (Triticum, Hordeum; Figure 2.13c, Poa, Zea, Elytrigia, etc.) and other monocotyledonous groups (Table 2.3). However, plates are not only correlated with primary alcohols. In grapes (V. vinifera) plate morphology is accompanied by the presence of oleanolic acid as a major constituent (Baker, 1982). Ketones are dominant compounds in the wax of Allium spp. and Euphorbia, which again have plate-like crystals, but usually of a more complex type than in primary alcohol species, with edges which are lobed or serrated or spiked (Jeffree et al., 1975; Baker, 1982, Figure 2.16a). The morphology is not always reliably distinguishable from the more common primary alcohol plate type. There is also some evidence that the dendritic structures (Figure 2.12d) sometimes observed in Brassica waxes are attributable to ketones (Baker, 1982).

Even

Odd

Even

Odd

Odd

Odd Odd

Even

Odd Even Even

Alkyl esters

Ketones

Aldehydes

Secondary Alcohols

Secondary alcohols

Secondary alkandiols β-Diketones

Primary alcohols

Hydroxy-β-diketones Fatty acids Fatty acids

Even

Odd

Alkanes

Polymeric aldehydes Polyesters δ-Lactones Triterpene acids

C-No.

Compound class

22–32

29–33 12–18 20–32

22–32

21–33 29–33

21–33

21–33

22–32

23–33

36–72

17–35

Range

Estolides 1,5-hexacosanolide Ursolic acid Oleanolic acid

Nonacosan-4,10-diol Hentriacontan-14,16-dione Tritriacontan-16,18-dione Hentriacontan-8,10-dione hexacosanol Octacosanol Triacontanol Hentriacontan-9-ol-4,16-dione Hexadecanoic acid Hexacosanoic acid Octacosanoic acid

Symmetrical Nonacosan-15-ol

Hexacosanal Octacosanal Asymmetrical Nonacosan-10-ol

Nonacosane Hentriacontane Octadecyl hexacosanoate Octacosyl octacosanoate Nonacosan-15-one

Common example

Brassica spp. Clarkia elegans Arabidopsis thaliana Nelumbo nucifera Eucalyptus globulus Triticum aestivum Buxus sempervirens Pisum sativum Triticum aestivum Trifolium repens Triticum aestivum Esterified in Copernica cerifera Esterified in Musa paradisaica Esterified in Musa paradisaica Saccharum officinarum Gymnosperms Cerinthe minor Malus, Prunus Vitis vinifera

Aquilegia vulgaris Tropaeolum majus Dawsonia superba

Brassica spp. Pisum sativum Copernica cerifera Musa paradisaica Brassica spp. Allium porrum Fagus sylvatica Citrus limon

Representative species

Leaf sheath, stem Leaf Leaf Fruit Fruit

Leaf Leaf Stem Leaf Leaf Flag leaf sheath Leaf Leaf adaxial Leaf Leaf Leaf abaxial Leaf Leaf

Leaf Leaf Sporophyte

Leaf adaxial Leaf abaxial Leaf Leaf Leaf Leaf Leaf Fruit

Organ

Columns, plates Columns, plates Columns, plates Tubes Tubes Tubes Open spirals Plates Plates Plates Tubes Crust Large rods Large rods Large rods ?Amorphous Tubes Plates, greasy film Plates

Tubes Tubes Tubes

Columns, plates Ribbons Crust Large rods Columns, plates Spiked plates Granular Granular

Associated wax type

Table 2.4 Principal classes of compounds in plant epicuticular waxes, with examples of representative constituents, plant species, organs and associated crystal morphologies

72 BIOLOGY OF THE PLANT CUTICLE

THE FINE STRUCTURE OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

73

Figure 2.12 Cuticular and epicuticular structure in various species. (a) Patterns of cuticular ridges such as these on a sepal epidermis of Arabidopsis thaliana are often mistaken for epicuticular wax, but are caused by cuticular wrinkling. No epicuticular wax structures are visible by SEM on this surface (cf. Figures 2.9g,h). (b) Aristolochia-type transversely ridged ribbons on the leaf adaxial surface of marram grass (Ammophila arenaria). (c) Simple plates of the primary alcohol type are frequently orientated in preferred orientations. On a leaf of Festuca arundinacea, the crystals prefer three orientations roughly 120◦ apart (see inset). (d) Dendritic structures growing from the tops of wax columns on a cabbage leaf may be related to the production of ketones. (e) Convallaria-type patterning of simple primary alcohol plates on a leaf of Eucalyptus camaldulensis. Rows of plates orientated transverse to the rows may traverse many different cells. (f) Looped filaments on the abaxial leaf surface trichomes of Cyathodes colensoi. (a,f) Bars = 10 μm; (c,e) bars = 2 μm; (b) bar = 1 μm; (d) bar = 5 μm. Figures 2.12a,b,d,f by C.E. Jeffree, Figure 2.12c from Pitcairn et al. (1986), Plant, Cell and Environment, 9, 191–196, Figure 2.12e from Jeffree et al. (1976).

74

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

(h)

(g)

(i)

(j)

(k) (l)

Figure 2.13 Structure and recrystallisation of epicuticular waxes, mainly of barley (Hordeum vulgare). (a) β-Diketone tube waxes on a barley flag leaf sheath and (b) on a barley lemma. (c) Primary alcohol plate wax on a barley leaf adaxial surface. (d) Recrystallised tube from barley leaf sheath wax.

THE FINE STRUCTURE OF THE PLANT CUTICLE

75

2.6.2.1 Granules The term granules [the ‘Körnchen’ of de Bary (1871, 1884) and Amelunxen et al. (1967)] is used as a bin for a range of indeterminate shapes that can be minute featureless particles (but usually with some kind of shape: round, polygonal, slightly flattened, elongated), or consist of compact aggregates of small rods, tubes or platelets or other sub-polygons. It is possible that granules are truly amorphous in some cases, but in others, such as those of Aegiceras corniculatum shown by Barthlott et al (1998) they are clearly aggregates of small crystals. Lack of crystallinity may not therefore be assumed, and there may be little common ground between the granular wax types either morphologically or chemically.

2.6.2.2 Filaments Massed, interwoven filaments 60–80 nm thick with lengths 3–10 μm occur in the tuber lenticels of potatoes (Solanum tuberosum; Hayward, 1974). The composition has not been reported, but filaments are often associated with triterpenes. Thread-like crystals of wax on the gametophyte of the moss Saelania glaucescens were found by Haas (1982) to contain the diterpenoid hydroxy kaurane as 94.6% of the wax. Wax crystals with fine thread and ribbon structures are also associated with the stems of several Macaranga species that are adapted to act as specific anti-climb barriers protecting their partner symbiotic ants from competition and predation (Federle et al., 1997). In all of the glaucous Macaranga species, triterpenoids were present in concentrations between 52 and 88%, containing epitaraxerol and taraxerone as major constituents, with smaller proportions of taraxerol, β-amyrin, and friedelin (Markstädter et al., 2000). However, in glossy species triterpenoids are absent, or present in much smaller quantities. A distinctive kind of structure loosely referred to here as a filament for want of a name occurs in Fragaria, Rosa (Figure 2.14e), and Potentilla and some other Rosaceae, and similar structures occur on the abaxial leaf surfaces of Acer spp. These structures often appear to have a triangular cross-section.

2.6.2.3 Plates Plate-type waxes vary considerably in shape, chemical composition and spatial pattern. The primary alcohol plate types represented by Eucalyptus, grasses and

(Figure 2.13) (e) Tubes recrystallised from the β-diketone fraction of barley leaf-sheath wax. (f) Plates recrystallised from barley leaf adaxial surface wax. (g,h) Replicas of recrystallised β-diketones (g) and hydroxy-β-diketones (h) from barley leaf-sheath wax, both showing tubular structure. (i) Negative-stained tubes recrystallised from barley leaf sheath whole wax. (j) Pt/C replica of tubes recrystallised from a solution of Ginkgo biloba wax in hexane. (k,l) Negative-stained tubes recrystallised from G. biloba wax showing evidence of spiral substructure. (a,b,d,e,j) Bars = 4 μm; (c,f,g) bars = 2 μm; (h) bar = 1 μm; (k,l) bar = 300 nm. Figures 2.13a,c,d,f from Jeffree et al. (1975, 1976). Figures 2.13g,j from Jeffree et al. (1976). Other images by C.E. Jeffree, E.A. Baker and P.J. Holloway.

76

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

Figure 2.14 Field emission low-temperature scanning electron micrographs of epicuticular wax structures of various species. (a) Quercus pubescens leaf abaxial surface, showing simple plate-type waxes. (b) Mahonia aquifolium young leaf adaxial surface, showing hollow tubes of the 10-nonacosanol type. (c) Spiral ribbons on a leaf of Cerinthe major purpurascens are intermediate between tubes and plates. (d) A dense layer of β-diketones tubes on a leaf of Hosta sieboldiana. (e) A filament network on a leaf abaxial surface of Rosa sp. The filaments appear to have a triangular cross-section. (f) A structured wax film on the surface of a trichome on the seed-pod of rosebay willowherb (Chamerion angustifolium). See also Figure 2.18b. (a,e) Bars = 1 μm; (b,c) bars = 200 nm; (d) bar = 3 μm; (f) bar = 500 nm. Figures 2.14a–f by C.E. Jeffree.

Fabaceae are probably the commonest morphological group. Other plate morphologies based on or modified by alkane, aldehyde, ester, ketone, symmetrical secondary alcohol and fatty acid constituents are comparatively poorly defined and there is still only weak discrimination between plate morphological variants or their chemical basis. This situation must be corrected before plate wax characters can be used safely

THE FINE STRUCTURE OF THE PLANT CUTICLE (a)

(b)

(c)

(d)

(e)

(f)

77

(g)

Figure 2.15 Recrystallisation of epicuticular waxes. (a) The adaxial leaf surface of Pisum sativum cv. ‘Meteor’ has a primary alcohol-rich plate-type wax that recrystallises in a plate form, as does (b) the primary alcohol fraction from the wax. (c) By contrast, the abaxial leaf surface, petioles, tendrils (shown here) and stipules bear elongated lobed ribbons. (d) The aldehyde fraction from the pea leaf abaxial surface approximates the morphology of the tendril wax. (e) Wax tubes on the leaf of Exochorda racemosa. The wax contains about 30% of asymmetrical secondary alcohols of which 93% is 10-nonacosanol. (f) Exochorda wax recrystallised in a ribbon morphology analogous to that of the pea tendril. However, short tubes are abundant at the bases of the ribbons (inset g). (a,c,d) Bars = 2 μm; (b,e,f) bars = 1 μm. (g) bar = 0.5 μm. Images by C.E. Jeffree, E.A. Baker and P.J. Holloway. Figures 2.15e, from Jeffree et al. (1975).

in taxonomy and phylogeny. I have struggled to restrain myself from including a special category for plate waxes shaped like ‘moose antlers’ (Prosopis; Hull et al., 1979); yet, there is frankly a lack of appropriate terminology to describe variation in plate wax forms. Most of the primary alcohol ‘simple plate type’ described here, and epitomised by the EW of the Eucalypts and Poaceae have distinctly lobed or crenate

78

BIOLOGY OF THE PLANT CUTICLE (a)

(b)

(c)

(d)

Figure 2.16 Structure and crystallisation of ketone waxes. (a) Spiked plates and filaments on the leaf surface of leek (Allium porrum). (b) Replica of the recrystallised ketone fraction from leek wax, showing plates with lobed margins. (c) Recrystallised leek wax, showing plates with spiked and lobed margins. In some instances, regularly grouped dendritic extensions are visible, analogous to the dendritic crystals of Brassica waxes (see Figure 2.12d). (d) The ketone fraction from Brussels sprout wax crystallises in much the same form as the ketone fraction from A. porrum (b) despite having different constituents. (a) Bar = 1 μm; (b,c) bars = 2 μm; (d) bar = 4 μm. Images by C.E. Jeffree, E.A. Baker and P.J. Holloway.

edges when viewed at sufficient resolution. But adequate resolution even to confirm their categorisation as a plate type is often not available from routine SEM images. From this perspective, the work of describing several tens of thousands of wax morphologies using high resolution SEM (Frölich and Barthlott, 1988; Barthlott et al., 1998) has been a major contribution to our understanding. Other wax types, such as the ketone rich plates of Allium porrum are chemically and structurally distinct from, for example, the primary alcohol rich plates of Trifolium repens, yet could be categorised with them in the absence of knowledge of their chemistry. There is a strong case, therefore, for reworking the wax morphological categories to take account of the correlation between wax chemistry and morphology.

THE FINE STRUCTURE OF THE PLANT CUTICLE

79

2.6.2.4 Tube-type waxes Tubes in general. Structures once taken to be rodlets in many species (Amelunxen et al., 1967) are now known to be tubes (Hallam, 1967, 1970b; Johnson and Jeffree, 1970; Jeffree et al., 1975, 1976). Plant surfaces bearing tube waxes are usually highly glaucous, strongly scattering of light enriched in short (blue) wavelengths. They are comparatively fragile (Barber, 1955), contrasting with the more mechanically robust plate type. Tube-type waxes form ultrahydrophobic surfaces, as in lotus N. nucifera (Barthlott et al., 1996; Barthlott and Neinhuis, 1997). Terminological confusion still remains at the boundaries between rods and tubes, and also between rods and filaments. Examples include the classification of the coiled crystals of Buxus as coiled rods by Barthlott et al. (1998) where on chemical considerations they are likely to be related closely to the well-recognised β-diketone tubes. Plant waxes do not always fall neatly quantitised into a tube bin or a plate bin. Intermediate forms occur, such as the mixtures of tubes and plates on the leaves of Pinus sylvestris (Leyton and Juniper, 1963) and Eucalyptus umbrawarrensis (Hallam and Juniper, 1971) and on the flag leaves of wheat. The term ‘tube’ is meaningless in these examples without further qualification of their chemical basis. Chemical analyses show that the commonest tube waxes fall into two distinct groups – those containing substantial amounts of asymmetrical secondary alcohols (predominantly 10-nonacosanol and its homologues; Table 2.5) and those containing substantial quantities of β-diketones, such as hentriacontan-14,16-dione in the Poaceae and tritriacontan-16,18-dione in Eucalyptus spp. (Horn and Lamberton, 1962; Horn et al., 1964; Hallam, 1967; Hallam and Chambers, 1970; Tulloch and Hoffman, 1971, 1974; von Wettstein-Knowles, 1972; Jeffree et al., 1975, 1976; Holloway et al., 1976; Baker, 1982; Tables 2.3 and 2.4). These two classes of compounds are never recorded as major constituents in the same species or even in the same family. Three further compound classes are associated with simple tubes, hydroxy-β-diketones, secondary alkanediols and delta lactones (see later). Secondary alcohol tubes. Tubes are the dominant wax crystal type in the Gymnosperm genera Ginkgo, Chamaecyparis, Taxus, Picea, Agathis, Pseudotsuga, Pinus, Abies and others, in which the asymmetrical secondary alcohol 10-nonacosanol is always an important constituent (Jeffree et al., 1975, 1976) although it can constitute less than a third of the EW load (Holloway et al., 1976). Asymmetrical secondary alcohols, of which 10-nonacosanol is the most important constituent, have also been recorded in a wide range of Angiosperms with tube waxes, mostly dicotyledons (Table 2.5), including Aquilegia alpinum, Chelidonium majus and Papaver somniferum, Rhus cotinus, Exochorda racemosa and P. domestica, Tropaeolum majus (Holloway et al., 1976). In Rosa cv. ‘Baccarra’, the variable distribution of tubes is positively correlated with the amounts of 10-nonacosanol present in the EW (Baker, 1982). Tubular crystals were also discovered on Tulipa kaufmanniana their hollow centres confirmed by negative staining, as were the tubes of P. sitchensis (Johnson and Jeffree, 1970; Jeffree, 1974a; Figure 2.17f). The composition as well

Exochorda racemosa

Aquilegia alpinum Papaver somniferum Chelidonium majus Prunus domestica

Pisum sativum

Clarkia elegans

12.5 12.5 3.0 3.0 3.0 5.5 5.5 5.5 57.0 65.7 66.0 48.3 48.3 48.3 48.3 30.1 30.1 30.1 30.1

12.5

Secondary alkanol fraction — Wax (%)

From Holloway et al. (1976) Brassica oleracea var. gemmifera

Species

tr

1.0

0.7 0.6 2.7 0.1 0.8 0.5 0.3 0.1 0.4 0.4 0.2

All

All 10-ol 10-ol 7-ol 8-ol 9-ol 10-ol 7-ol 8-ol 9-ol 10-ol

0.5

Fraction (%)

All

All

Isomer

0.40 0.39 1.78 0.05 0.39 0.24 0.14 0.03 0.12 0.12 0.06

0.06

tr

0.06

Wax (%)

C27 secondary alkanols

36.3 60.1 1.0 16.0 83.0 0.1 0.2 3.1 98.3 99.4 95.3 2.9 92.7 1.0 2.3 93.0 1.0

9-ol 10-ol 11-ol

0.9

Fraction (%)

14-ol 15-ol 13-ol 14-ol 15-ol 13-ol 14-ol 15-ol 10-ol 10-ol 10-ol 9-ol 10-ol 11-ol

13-ol

Isomer

0.69 27.99 0.30

4.54 7.51 0.03 0.48 2.49 0.01 0.01 0.17 56.03 65.31 62.90 1.40 44.77 0.48

0.11

Wax (%)

C29 secondary alkanols

9-ol 10-ol 11-ol

14-ol 15-ol 16-ol All All All All

All

All

Isomer

tr 1.0 0.9

2.4 37.1 51.8 tr tr tr 0.6

nd

0.9

Fraction (%)

0.30 0.27

0.13 2.04 2.85 tr tr tr 0.29

nd

0.11

Wax (%)

C31 secondary alkanols

Table 2.5 Asymmetrical secondary alkanols and alkandiols in tube waxes. The abundance of the major constituent as percentage of the secondary alkanol fraction is indicated in bold

80 BIOLOGY OF THE PLANT CUTICLE

16.0 61.0

From Neinhuis and Jetter (1995) Pogonatum urnigerum Pogonatum belangeri

nd = not determined; tr = trace; nd = not detectable.

Nelumbo nucifera Thalictrum flavum Papaver somniferum

From Barthlott et al. (1996)

Secondary alkanol fraction – wax (%) 28.0 57.0 61.0

All 10-ol

27.8 48.9

0.22 0.54

0.20 0.06

0.10 0.24 0.14 1.02

0.15

Secondary alkandiol fraction – wax (%) 57.0 12.0 11.0

0.8 1.1

1.0 0.6 —

All All All

Tulipa gesneriana

Tropaeolum majus

Picea sitchensis Picea pungens Chamaecyparis lawsoniana Agathis australis Ginkgo biloba

0.4

0.2 0.5 0.3 2.7

All

8-ol 9-ol 10-ol All

38.3

38.3 38.3 47.5 47.5 47.5 37.8 37.8 37.8 19.7 10.0 28.6

Rhus cotinus atropurpurea

C29 10-ol C29 10-ol

10-ol 10-ol

10-ol 11-ol 9-ol 10-ol 11-ol 9-ol 10-ol 11-ol 10-ol 10-ol 10-ol

9-ol

91.0 97.0

98.0 98.9

93.2 2.8 3.0 92.5 3.0 2.9 92.4 1.0 98.0 98.8 100.0

2.4

14.56 59.17

27.24 48.36

35.70 1.07 1.43 43.94 1.43 1.10 34.93 0.38 19.31 9.88 28.60

0.92

All All

1.2 —

1.0 0.6 —

tr

All

All All All

tr

tr

All

All

0.33

0.20 0.06

tr

tr

tr

THE FINE STRUCTURE OF THE PLANT CUTICLE

81

82

BIOLOGY OF THE PLANT CUTICLE

(a)

(b)

(c)

(d)

(e)

(f)

(g)

Figure 2.17 Pt/C replicas of the structure and crystallisation of Sitka spruce wax tubes. (a) Wax tubes on a current-season leaf. (b) A tube seen end-on, clearly demonstrating its hollow structure. (c) Epicuticular wax recrystallised from solution in benzene, showing tubular structure. (d) Tubular structure crystallised by slow-cooling of molten Sitka spruce wax. The tubes bear diagonal (spiral) striations at about 30◦ to the long axis. (e) The secondary alcohol fraction from Sitka spruce wax recrystallised in tube form. (f) A negatively stained Sitka spruce wax tube shows segments of stain in its lumen. (g) The remainder of the wax, depleted in secondary alcohols, forms large plates instead of tubes. (a) Bar = 1 μm; (b) bar = 200 nm; (c,e,f) bars = 500 nm; (d) bar = 400 nm; (g) bar = 2 μm. Figures 2.17a–e and g from Jeffree (1974a). Figure 2.17f from Johnson and Jeffree (1970), Planta, 95, 179–182.

THE FINE STRUCTURE OF THE PLANT CUTICLE

83

as the structure of the wax of T. kaufmanniana bears comparison with that of Ginkgo biloba or T. majus (Table 2.5). β-Diketone tubes. Hallam (1964, 1967) and Hallam and Chambers (1970) found that tubular wax structures and β-diketones in 113 Eucalyptus species were strongly correlated, as were plate waxes and primary alcohols in a further 238 eucalypts. In 36 of these species plates and tubes occurred together, and the wax contained both primary alcohols and β-diketones. Further β-diketone-rich waxes with tubular structures occur in the Poaceae (barley leaf sheath, wheat leaf abaxial surface; Netting and von Wettstein-Knowles, 1973; Jeffree et al., 1975, 1976) and Dianthus caryophyllus; Table 2.3; Figures 2.13a,b and 2.14e), and in the tubular waxes of Ericaceae (Rhododendron spp.; Evans et al., 1975; Vaccinum ashei, Freeman et al., 1979; Andromeda polifolia, Jeffree, 1986). In the Poaceae, as in the eucalypts, the leaf waxes fall into two contrasting types, sometimes to be found on different parts of the same plant. Secondary alcohol and β-diketone tubes are distinguishable from their morphology as well as their chemistry (see earlier). The secondary alcohol tubes are usually shorter and wider (0.3–1.1 μm long by 0.1–0.2 μm wide) than the β-diketone type (typically 2–3 μm long by 0.2–0.3 μm wide; Jeffree et al., 1975). Alkandiol tubes. More recently, it has become clear that secondary alkanols and secondary alkandiols often co-occur in tube wax species, forming about 12% of the wax of Thalictrum flavum and 11% of P. somniferum (Tables 2.3 and 2.4; Barthlott et al., 1996). Although they occur widely in secondary alcohol-rich waxes, usually as subsidiary compounds, it is rare for a secondary alkandiol to dominate. Currently the only known exception is in the tube wax of N. nucifera which contains 57% of secondary alkandiols [of which nonacosan-4,10-diol (38%), nonacosane-5,10-diol (38%) and nonacosan-10,13-diol (22%)] accompanied by 28% of secondary alkanols, with 10-nonacosanol being the major constituent (Jetter and Riederer, 1995; Barthlott et al., 1996). Similarly, β-diketones and hydroxy-β-diketones occur together in the tube waxes of many Poaceae, although not in dicots. However, there are no known examples of waxes containing both β-diketones and asymmetrical secondary alcohols. Delta-lactone tubes. Another completely new category of tube-wax forming compounds is the δ-lactones (1,5-alkanolides) reported in Cerinthe minor by Jetter and Riederer (1999a), the first example of this type that has been found. Tubes, curls, spirals and branches. Several of the images of tube waxes presented by Barthlott et al. (1998) show transitional forms between coils and tubes. Thus, in Lonicera korolkovii (their figure 20) the tubes appear to be constructed as tight coils, resulting in a characteristic twist at the distal end, and close spiral ridges down the side of the tube. Their images of Buxus sempervirens show small coiled rods (or are they coiled tubes? – more resolution is required to determine this) and much larger, clearly hollow tubes together with intermediate forms that appear to be curved plates.

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It is noteworthy that the wax of Cerinthe major purpurascens shows an open helical structure (Figure 2.14c), indicating a possible relationship between coiled ribbons and closed tubes. Curling of plates and coiling of tubes are also evident in some β-diketone waxes, first demonstrated in Chrysanthemum segetum by Juniper and Bradley (1958) in their paper on the then newly developed carbon replica technique. Recently, a similar morphology has been reported in B. sempervirens (Meusel et al., 2000). Both forms are associated with C31 β-diketones substituted in unusually asymmetrical 8,10 (Buxus) or 10,12 positions (Chrysanthemum) compared with the more nearly symmetrical 14,16 positions of the β-diketones in H. vulgare (Figures 2.13a,b), which produces tubes but no coiling. Brassica-type tubes. A third and less frequent category of tubes identified by Baker (1974, 1982) on leaves of Brassica spp. and stems of A. thaliana also appears in Clarkia elegans and Centranthus ruber together with comparable, though not identical, chemical compositions (Hunt et al., 1976). The tubes of Brassica wax are much larger than those described earlier, up to 8 × 1 μm, and have transverse striations visible in negative stained specimens (see Jeffree et al., Holloway, 1976) or in replicas (Martin and Juniper, 1970; Hallam and Juniper, 1971; Figures 2.18a,b). The Brassica wax type is characterised by mixtures of symmetrical secondary alcohols (n-nonacosan-15-ol), ketones (n-nonacosan-15-one) and alkanes (Netting et al., 1972; Holloway et al., 1977). Although short Brassica tubes appear to have hollow ends when seen in the SEM (e.g. Neinhuis et al., 1992; their figure 5) negative staining of the wax columns of Brassica does not reveal them to be hollow (Figure 2.18b), and it is therefore likely that they become filled as they grow. They are therefore not comparable with true tubes of the types formed by asymmetrical alkanols, alkanediols and β-diketones. Gomez-Campo et al. (1999) used the term ‘column’, and that seems a more appropriate term, distinguishing these characteristic structures from simpler terete rods. Because of the current importance of A. thaliana as a model plant it may be worthwhile to clarify some terminological issues concerning the EW morphologies of this species. Gomez-Campo et al. (1999) referred to three layers of crystals in Brassica waxes, a continuous sheet (i.e. background film), flat crystals and upright columns. Because the density of crystallites on Brassica leaves is very high, replicas of the intact surface crystal become too complex to interpret easily, and clarity of the images of individual crystallites is improved where areas of the wax have been damaged by abrasion, as in figure 4.28 in Martin and Juniper (1970) and figure 12 of Hallam and Juniper (1971). These images indicated two predominant crystal morphologies: lobed or crenate plates, often in low profile on the surface, and longitudinally fluted and transversely-ridged rods or tubes (Figure 2.18a). Recent experiments using a TEM with a goniometer tilting stage reveal that crystals that appear to be ‘plates’ at one angle resolve as fluted columns when tilted to another. The morphologies are therefore two ends of the spectrum of possibilities for column assembly using the same stacked planar modules. The crenate plates often contain holes, and there may be indications of a hole in images of tubes, but negative staining

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(b)

(c)

Figure 2.18 (a–d) Wax structures in Brassica spp.

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does not reveal a hollow core (Figure 2.18b). Successive plates diminish in size from base to apex of the rod, so that the rod usually tapers distally. However, in the gl2 mutant the direction of tapering is reversed (Figures 2.19e,f).

2.6.2.5 Rodlet-type waxes Baker (1982) pointed out the correlation between aldehydes in waxes of lemon C. limon fruits in which the wax is approximately 43% aldehydes, and short rods. Rods are again observed in the aldehyde-rich wax of sugarcane (S. officinarum), but in this species the aldehydes are not fully soluble, and are probably polymeric. Aldehydes, chiefly triacontanal, occur again as 43% in the specialised plate waxes of the Nepenthes pitcher (Martin and Juniper, 1970; Riedel et al., 2003). The Strelitzia rodlet type. Large wax rodlets of a readily recognisable type occur on the leaves of many Poaceae – S. officinarum (de Bary, 1871, 1884; Jeffree et al., 1975, 1976; Haas et al., 2001), Arundinaria spp. (Jeffree et al., 1976) and Typha spp. (Djebrouni, 1989) – and in Musa paradisaica and Strelitzia reginae and Heliconia (Meusel et al., 1994; Barthlott et al., 1998). These form co-aligned crystals (Kreger, 1949) aggregated into massive terete rodlets, or in Strelitzia and Heliconia cylindrical chimneys accurately surrounding each stomatal complex (Meusel et al., 1994; Barthlott et al., 1998). The massive wax films of carnauba palm (C. cerifera and C. cowellii) are huge accumulations of these co-aligned crystals. This type of wax structure appears to occur in the Monocotyledons almost exclusively in the Commelinidae and Arecacidae and to be absent from the Liliaceae and has been named the Strelitzia type (Barthlott and Frölich, 1983; Frölich and Barthlott, 1988; Meusel et al., 1994; Table 2.3). A similar wax type occurs in some Dicotyledons, including Wooleya farinosa and Malephora sp. (Ihlenfeldt and Hartmann, 1982) Eryngium (Frölich and Barthlott, 1988) and fruits of the wax gourd Benincasa hispida (de Bary, 1871, 1884; Barthlott, 1990; Meusel et al., 1994), but the list is not exhaustive. The waxes of the Strelitziaceae and Musaceae contain substantial quantities of alkyl esters, or of polymeric aldehydes (11% in Strelitzia;

(Figure 2.18) (a) Gold–palladium replica of the leaf surface of cabbage (Brassica oleracea var. capitata) showing stacks of flat-lobed plates, and tapering columns. The former appear to develop into the latter. The replica was made by detaching the sputter coating from a low-temperature SEM specimen using frozen ethanol. (b) Large simple wax plates formed transverse to cuticular ridges on a leaf of rosebay willowherb (Chamerion angustifolium). Some plates span two or more ridges. (c) Simple plates form a regular pattern of rows radiating from a stomatal aperture on the abaxial surface of a leaf of Bupleurum ranunculoides. (d) A negative-stained wax column from Brussels sprout (B. oleracea var. gemmifera) shows superficial transverse ridges, but no evidence of a hollow centre and is therefore not tubular. The pattern is analogous to the ‘Convallaria’-type waxes widespread in the monocotyledons (Barthlott et al., 1998), but the plates are also interspersed with transversely ridged rodlets of the Aristolochia type (see Meusel et al., 1999). (a) Bar = 1 μm; (b) bar = 2 μm; (c) bar = 5 μm; (d) the wax rod is 4.2 μm long. Figures 2.18a–c by C.E. Jeffree, Figure 1.12d from Jeffree et al. (1996).

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Figure 2.19 Structure and recrystallisation of Brassica oleracea waxes. (a) Recrystallised Brussels sprout wax, showing plates unlike those found on leaves. (b) The same wax delivered to a porous disc via a short chromatographic column, showing several crystal motifs characteristic of the epicuticular waxes of B. oleracea leaves. (c) Slow-crystallised B. oleracea wax, showing dendritic structures analogous to those shown on a cabbage leaf in Figure 2.12d. (d) Plate-like structures formed by recrystallisation of the secondary alcohol fraction from Brussels sprout wax. (e) Tapered columns on the leaf surface of B. oleracea gl2 mutant have an inverted taper compared with the normal wax, which is reproduced when the wax is recrystallised (f) together with other typical B. oleracea motifs, such as columns with a false hollow end. (a,b,d) bar 2 μm, (c,e,f) bar = 1 μm. Figures 2.19a, and c–e by C.E. Jeffree, E.A. Baker and P.J. Holloway. Figures 2.19b,f from Jeffree et al. (1975, 1976).

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Meusel et al., 1994), but the related wax morphology in B. hispida is associated instead with triterpene acetates (Meusel et al., 1994). The Aristolochia rodlet type. Another characteristic type of wax rodlet found in basal Angiosperm genera of the Magnoliidae such as Aristolochia, Liriodendron and Magnolia is referred to by Barthlott et al. (1998) as the Aristolochia type (Table 2.3). These substantial rodlets usually have pronounced transverse ridges, and often appear to be assemblages of transversely stacked platelets. The morphology of this group shows considerable overlap with the morphology of Brassica type waxes, some of the crystal motifs being shared between them. For example, it would be hard on morphological grounds alone to distinguish between the waxes of Brassica oleracea var.gemmifera and Fritillaria pallidiflora, and the waxes of N. glauca and Leucojum aestivum show the same type of construction of fluted columns from irregularly curled plates that occur in seakale (Crambe maritima). Analysis of these waxes reveals little common ground between them in terms of composition (Meusel et al., 1999). One group, comprising Aristolochia, Paeonia and Centranthus, to which could also be added the fern Osmunda regalis (Jetter and Riederer, 2000) has significant amounts of ketones, but others including Galanthus, N. glauca, L. aestivum and F. pallidiflora lack them completely, and instead have alkanes as about 70% or more of the wax together with alkyl esters and primary alcohols. In Paeonia the major ketone is the symmetrical palmitone (hentriacontan-16-one) and in Centranthus nonacosan-14-one, both with important quantities of triterpenoids. The major constituents in Laurus are primary alcohols (52%) and fatty acids (Meusel et al., 1999). It therefore appears that the waxes of these species have convergent wax crystal morphologies based on a variety of different chemical solutions. It is clear that in these species, and probably also in the Brassicaceae, a simplistic view of relationships between single major constituents and wax ultrastructure is not appropriate. Farina. On some ferns (Pityrogramma spp.; Wollenweber, 1982), and in many species of the Primulaceae the ‘wax’ or ‘farina’ found on the leaves is composed almost entirely of crystalline lipophilic flavonoids. In most of these species the farina forms dense assemblages of narrow rodlets.

2.6.3

The background EW film

Jeffree et al. (1975, 1976) postulated that the background layer underlying the EW crystallites was a continuous amorphous wax coating covering the CP. The use of the word ‘amorphous’ to indicate the absence of crystalline projections detectable by microscopy is inadvisable. In some instances (e.g. on the leaves of B. vulgaris, Clematis vitalba and P. vulgaris), there is little visible evidence of epicuticular ultrastructure at the modest resolutions achievable with the SEM of the 1970s. Evidence of planar structures in an otherwise featureless surface layer or in the spaces between crystallites was often visible in TEM replicas of B. oleracea var. capitata

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and Hyacinthus orientalis wax (Juniper and Bradley, 1958) and on the cuticle surface of cotton leaf (G. hirsutum; Hallam and Juniper, 1971), but confirmation of the existence of a discrete superficial wax layer by SEM is not usually possible. Jeffree (1996) showed a low-temperature SEM image of the surface of a Quercus pubescens leaf from which the wax was stripped at −196◦ C by using pentane as a cryo-adhesive. The image demonstrated that the EW crystallites arise from a continuous covering layer of wax about 0.2–0.3 μm thick. Likewise, confirmation that the apparently smooth surface of cuticles of Prunus laurocerasus nevertheless carries an EW film that is chemically distinct from the embedded cuticular wax has come from elegant experiments by Jetter et al. (2000), Ensikat et al. (2000) and Jetter and Schäffer (2001) in which the superficial wax layer was removed mechanically using ambient- or cryo-temperature adhesives. A striking observation is that despite the fact that EW must be transported through the cuticle, the soluble wax compositions of the intracuticular and epicuticular domains each appear to be uncontaminated with the other’s constituents. The data of Jetter et al. (2000) show that the mechanically harvested epicuticular layer of P. laurocerasus wax contains exclusively aliphatic compounds, while the triterpenoids ursolic and oleanolic acid represent almost two-thirds of the intracuticular wax. How this chemical segregation is achieved remains unknown. The EW appears to be extended as a film over all parts of the general epidermal surface to form the visible surface exposed between the EW crystallites. No pores or other structures that might correlate with points of origin of the crystallites are visible in the underlying cuticle. Whether the projecting crystals originate directly at the surface of the CP or are rooted somewhere within the EW film is still a matter of conjecture. In Phragmites australis leaves cryo-stripped with ethanol, what appear to be the stumps of wax rodlets remain attached. By contrast, Koch et al. (2004) found that the longitudinally ridged rodlets of Galanthus leaf surfaces were stripped completely with epoxy resin leaving a featureless smooth surface. Neinhuis and Barthlott (1997) stated that the EW penetrate the cuticle to build up films, crusts or distinct surface structures. The word ‘penetrate’ was probably used here in the sense of ‘enter while being transported across’. It is not suggested that epicuticular crystals are rooted deep in the CM, and images of the CM structure beneath EW crystals in various species (Figures 2.4e,f) lend no support to this, implying that the crystal structures originate entirely externally to the CP layer. Although the background wax film appears amorphous in the SEM, there are good reasons to doubt this, and the consensus of opinion is that the film may be crystalline, even if it does not appear to be so in EM images. Images of the structure of wax films regenerating on plant surfaces have recently been made using Atomic Force Microscopy (AFM) in three species – Euphorbia lathyrus, Galanthus nivalis and Ipheion uniflorum (Koch et al., 2004; Figures 2.20a–d). In this groundbreaking study, the first ever direct observation of a time-course of wax regeneration in a living leaf, it is clear that progressively the surface is covered completely with monomolecular and then bimolecular layers of crystalline wax. The images (Figure 2.20a,b) show rodlets growing from the cuticle independently,

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(a)

(b)

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Figure 2.20 Atomic force microscopy (AFM) images of epicuticular wax regenerating on a living Galanthus nivalis leaf adaxial surface, following removal of the earlier-secreted wax with epoxy resin glue. The images show changes in a 6 × 6 μm area during an 80-min time frame. Flat, lobed plates extend over the whole surface to form first a monolayer (light grey) (a,b) and then bilayered structures (dark grey) (c,d). Independently, rod-like crystals arise directly from the cleaned cuticle proper (CP) surface, extending by growth at their distal ends, not from the base. Diagonal arrows (a,b) and vertical arrows (c,d) mark reference points that remain in fixed positions during rod extension growth. The black arrow in (a) marks a crystal that in (b) has been removed by the AFM tip. (a–d) bars = 1 μm; reproduced with permission from Koch et al. (2004), Journal of Experimental Botany, 55, 711–718.

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beginning from bare areas of cuticle prior to the completion of the coverage of the background film. Only rarely is the CP exposed at the surface. Fungi, particularly the rusts, respond to cutin monomers by increasing the germination rate or frequency. Magnaporthe grisea (rice blast) differentiation is induced by the addition of cutin monomers (Gilbert et al., 1996; de Zwaan et al., 1999) and the fungus produces cutinase during infection (Sweigard et al., 1992). These monomers may be released directly from the cutin polymer, by cutinase secreted by the spores and hyphae, or there may be a sufficient amount of the molecules free in the covering wax film. The responses of germ tubes to stomata indicate that they can be sensed by hyphae, and there is some evidence that cutin is exposed at the stomatal lip of broad bean and cowpea leaves since it stains locally with Nile red (Collins et al., 2001; Figure 2.10g).

2.7 Cuticular pores and permeability of the CM 2.7.1

Permeability of water and solutes

The reticulum of polysaccharide microfibrils ramifying through the CL offers a potential pathway for diffusion (but not mass flow) of water and solutes through the CL (for a functional discussion see Chapters 8 and 9). Although the reticulum demonstrably traverses the entire CL in A. americana (Wattendorff and Holloway, 1980) and also in P. abies (Tenberge, 1989, 1992) it stops at the base of the ECL in mature C. miniata cuticles,which have notably low permeability to water (Schmidt et al., 1981; Schmidt and Schönherr, 1982). The ultrastructure of the pathway for diffusion of water and other molecules across a lamellate CP is probably at intermolecular scale. It has never been observed, and may never be directly by currently available technologies. Periclinal penetration of permanganate is two–four times faster in the electron-dense than in the electron-lucent lamellae of the CP. In the CL, penetration by the fibrillar pathway is some ten times or faster than via the cuticular polymer matrix (Wattendorff and Holloway, 1984). The lamellate cuticle covering the guard cells of F. hygrometrica sporophytes (Sack and Paolillo, 1983a,b) is penetrated apparently to the surface by polysaccharide fibrils (Figure 2.4d) which may evaporate water from their tips, but in almost all cuticles putatively of Type 4, the ends of the microfibrils are capped with a thin amorphous CP. Since our understanding of the ontogeny of the CM is that the construction of the procuticle and

(Figure 2.20) (e,f) SEM images of wheat leaf wax recrystallised from chloroform on glass forming an amorphous film (e) and on freshly cleaved highly ordered pyrolytic graphite (HOPG; PLANO GmbH, Wetzlar, Germany) (f), producing highly ordered wax crystals; reproduced with permission from Kerstin Koch (see also Koch, K., Barthlott, W., Koch, S., Hommes, A., Wandelt, K., Mamdouh, H., De-Feyter, S. and Broekmann, P. Structural analysis of wheat wax (Triticum aestivum, c.v. ‘Naturastar’ L.): from the molecular level to three dimensional crystals, Planta, in press). This figure is produced in colour in the colour plate section, which follows page 249.

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CP outside the CW always precedes the process of CW impregnation that produces the CL layer, the only ways that the fibrillar reticulum can become uncapped, and the fibrils exposed at the surface, are by the systematic disassembly or sloughing of the CP, or by the failure of the CP to keep pace with massive area expansion of an organ. There is currently no convincing evidence for any of these in ordinary leaves, but it is possible that in fruit cuticles the CP is discontinuous or lacking. In cuticles of higher plants, evidence for discrete pores, capable of functioning as pathways for fluid mass flow, or wax delivery is practically confined to secretory cells, including the receptive stigmatic papillae. In glandular hairs of many species, secreted materials accumulate between the PCW and the CM which becomes detached from the CW. Release of secretions may require rupture of these cuticles. A comparatively thick specialised Type 1 stigma cuticle of Crocus chrysanthus has a chambered structure in which the CL layer is eroded to the base of the CP to form ovoid chambers containing a protein–polysaccharide secretion. At maturity, the entire CM is underlain with this secretion, separating it from the PCW and permitting the growth of pollen tubes within the layer of secreted material (Heslop-Harrison and Heslop-Harrison, 1982). These processes presumably involve enzyme-mediated local reduction of the CM. In other species, cuticles covering glandular trichomes [e.g. Drosera (Chafe and Wardrop, 1973) stigmatic papillae] or absorptive cells [the pitcher epidermis of Sarracenia (Joel and Heide-Jørgensen, 1985)] and trap gland cells of Utricularia monanthos (Fineran and Lee, 1975) may be discontinuous or of highly variable thickness, or contain pores or slits through which secretion and absorption may occur. The pre-secretory cuticle covering the stigma of rye, Secale cereale, is thrown into ridges at the apex of which thinning occurs until mechanical rupture of the cuticle occurs at maturity, releasing a mucilaginous polysaccharide secretion (Heslop-Harrison and Heslop-Harrison, 1982). In Abutilon venosum, nectar droplets pass through a cuticle 0.4 μm thick perforated with pores 0.15–0.25 μm across at the tip of each nectary hair (Findlay and Mercer, 1971; Table 2.2). These pores seemed to have valve-like action, periodically releasing nectar accumulated between the cuticle and the CW. Gaps in the CM are reported for other gland types. In many cases the evidence points to mechanical rupture at weak points in the CM. Thus, the thin, discontinuous cuticles of Drosophyllum lusitanica (Joel and Juniper, 1982; Joel et al., 1983) rupture at predetermined points of weakness under the tension forces caused by area expansion of the underlying PCW, which reduces in thickness by a factor of more than 2. No gaps or pores were found in the cuticles of glandular trichomes of Phyllyrea latifolia (Gravano et al., 1998). The authors therefore thought that the secretory products diffused through the cuticle. In the glandular hairs of Galphimia brasiliensis the Type 3 cuticle comes very close to being Type 4, the CP zone free of visible microfibrils being less than 20 nm thick on top of approximately 0.7 μm of reticulate CL.

2.7.2

Wax secretion, cuticular pores and microchannels

The ontogeny of the EW layer (see also Chapters 4 and 5) has been debated for almost 150 years. de Bary (1871, 1884) proposed that wax was exuded to the surface via

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pores in the cuticle, and in the same year, Weisner (1871) proposed that the wax was transported to the surface in solution in a volatile solvent. Before the advent of SEM, pores were reported in the cuticles of S. officinarum (Wijnberg, 1909) and M. paradisaica (Mueller et al., 1954), both species with very large rodlets observed by LM by de Bary (1871, 1884). Mueller et al. (1954) proposed that the wax was extruded under pressure through these pores as a soft paste (see also Schieferstein and Loomis, 1959). Hall and Donaldson (1962, 1963) and Hall (1967a,b) claimed the existence of pores in direct and secondary TEM replicas of leaves of T. repens, Trifolium pratense, B. oleracea and Poa colensoi. Their images showed a network of putative pores on Trifolium leaves in locations which might correspond to the positions of plate wax crystals. Fisher and Bayer (1972) also claimed the existence of pores of 2.5 nm diameter traversing the entire CM in sections of P. major CM. More recent images of the CM of the same and many other species by Holloway (1982a and unpublished) lend no support to this. Von Wettstein-Knowles (1974) proposed that wax tubes on barley leaves and leaf-sheaths could be extruded from pores, with the wax precursors polymerising in contact with the air. It should be noted that with the exception of the estolides of gymnosperm waxes (von Rudloff, 1959), and the polymerisation of aldehydes in the waxes of S. officinarum (Lamberton and Redcliffe, 1959; Haas et al., 2001) and Nepenthes (Riedel et al., 2003), polymeric constituents are almost unknown in EW of Angiosperms, and the wax tubes of barley are now known to consist of freely soluble β-diketones and hydroxy-β-diketones. The forms of wax rods on leaf sheaths of S. officinarum and nodes of Arundinaria, the leaves of Strelitzia and the stomatal chimneys of Heliconia with their longitudinal striae (Frölich and Barthlott, 1988; Meusel et al., 1994) are suggestive of extruded structures, and Schieferstein and Loomis (1959) thought of them as bundles of extruded wax threads. But it is known from X-ray crystallographic data and electron diffraction studies (Kreger, 1949; Reynhardt and Riederer, 1991, 1994; Reynhardt, 1997; Meusel et al., 2000) that these wax forms are bundles not of extruded threads but of longitudinally aligned crystals. Nevertheless, the issue of wax extrusion through cuticular pores has again been raised in S. coronata, the ouricuri palm of Brazil, which bears Strelitzia-type waxes assembled into massive epidermal wax crusts, but the true identity of the observed structures by Machado and Barros (1995) cannot be confirmed because of poor image resolution. The morphologies of most other EW types listed in Table 2.3 do not lend themselves to an extrusionbased origin. Tubular crystals would require a complex pore with a central spigot, while crystals with crenate edges, or the transversely ridged structures seen on Bupleurum (Figure 2.18d), Actinidia (Barthlott et al., 1998) and Brassica species (Figures 2.18a,b) would require pores with fluctuating dimensions. Miller (1982, 1985, 1986) reported the ‘anticlinally-oriented transcuticular canals’ in leaves of more than 50 species, but his observations made with the light microscope failed to confirm the identity of such features. Electron microscopy has also consistently failed to reveal transcuticular pores in the sense of open transcuticular channels via which wax, or other materials, might pass freely (Roelofsen, 1952; Mueller et al., 1954; Juniper, 1959, 1960; Schieferstein and Loomis, 1959; Crisp, 1965; Jarvis and Wardrop, 1974; Jeffree, 1974a).

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Hallam (1964, 1967) suggested that wax might migrate to the surface through the anastomosing channels between the lenticular cuticular (CP) lamellae of Eucalyptus cinerea, and that the ultimate pattern of wax arrangement may be partly a result of the exit pathway through the cuticle. But he had already obtained and reported in the same works evidence of their crystalline properties that seemed more persuasive. In a light TEM and SEM study of the adaxial surface of primary needles of ten-weekold Pinus radiata (Franich et al., 1977; Wells and Franich, 1977), rows of wax tufts across cuticular ridges consisting of aggregates of wax tubes were observed, and were interpreted as covering clusters of pores from which they were exuded. Well before bud-break, Sitka spruce needles already have a CP and a CL, and EW are crystallising as tubes on top of the CM, which must therefore be permeable to 10-nonacosanol and other constituents of the EW (Figure 2.11). No structure corresponding to a pathway for the wax is visible by SEM or TEM, however (Jeffree, 1974a). The developing surfaces in-bud are completely dry, and there is no sign of the presence of a solvent. The mechanism by which the EW is delivered to the plant surface is currently unknown. AFM images of regenerating wax layers on leaves of various species show extension of mono- and bimolecular films of wax occurring without any visible evidence of a source (Koch et al., 2004). In the petiole of Vaccinium reticulatum, and in sections of leaf cuticles of P. major and A. crenata (Fisher and Bayer, 1972), the CL is penetrated by a ‘channel-like reticulum which emanates from the primary CW’, consisting of polysaccharide microfibrillar bundles embedded in cutin (Singh and Hemmes, 1978). Pectic radiating strands in the reticulate CL of Spartocytisus filipes were referred to as ‘microchannels’ by Lyshede (1978) and in H. suaveolens by Heide-Jørgensen (1978a,b). These structures, which correspond with massively enlarged forms of the tree-like reticulum in the spruce cuticle (Tenberge, 1989, 1992; and Figure 2.2), are light in TEM negatives, consisting of material more electron-dense than the matrix. They are thus inconsistent with the concept of open pores or channels. The reticulum of polysaccharide fibrils might be thought to represent a possible wax pathway, but their polar reaction, and the evidence of Wattendorff and Holloway’s (1984) experiments with KMnO4 suggesting they are permeable to water and solutes are incompatible with the view that they are pathways for wax mass transport.

2.7.3

Relative sizes of wax crystals and cuticle

Let us also be clear about the relative dimensions of wax crystals, the cuticle and the underlying CW. Images of the EW and cuticle structure usually feature one without the other, making it hard to appreciate the relative scale of wax crystals compared with the thickness and lamellar dimensions of the CP. However, the ghosts of wax crystals sometimes appear in TEM images of the cross-sections of the CM, as in the work of Hallam (1964, 1967, 1970), Hallam and Chambers (1970; tube and platetype Eucalyptus waxes), Jarvis and Wardrop (1974; P. tenax), Holloway (1982a; Figure 2.4f, P. sitchensis) and Wattendorff (1974; Figure 2.4e; A. americana). In a

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micrograph of Eucalyptus papuana epidermal cells by Hallam in Martin and Juniper (1970, his figure 1.1), the β-diketone wax tubes are as long or longer than the entire thickness of the CW, and the diameter of the tubes approximates the full thickness of the CM. In the daffodil petal cuticle (P.J. Holloway, Figure 2.9g) the cuticular ridges are approximately 0.6–0.9 μm wide. Similarly sized cuticular wrinkles on the leaf of willowherb (Chamerion angustifolium; Figure 2.18c) bear wax crystals that span two or more wrinkles. If pores were responsible for wax extrusion in these cuticles they would have to span the wrinkles. Two important conclusions can be drawn from these images: first the lengths (height) of the wax crystallites is typically greater than the thickness of the CP, and in some instances thicker than the entire CM, and second no trace of any pores of dimensions corresponding with the crystallites is ever visible. Pores large enough to act as paths for wax extrusion could not hide undetected. They would be conspicuous, their diameter up to an order of magnitude larger than the thickness of the entire CP, and moreover accumulations of yet-to-be extruded material might be expected to appear beneath them. In the CP of A. americana shown in Figure 2.4e, the CP lamellae are in orderly periclinal alignment from side to side of the image. They are not interrupted anywhere, they do not deviate relative to the positions of the wax crystallites or take any account of them whatever, and in the spruce image (Figure 2.4f), the CW microfibrils come close to the surface of the CM, but are much more than an order of magnitude smaller than the diameter of the tubes and their distribution shows no relationship with those of the tubes (see also Jeffree, 1974a). If wax moves through the structure of the CP, as it must, it does so without altering the ordered structure of the CP.

2.8 Crystallisation studies on EW Although de Bary (1871, 1884) proposed exudation of wax to the cuticle surface, he used the word ‘Krystalloid’ to describe the wax structures on S. officinarum and other plant surfaces and must have regarded their structure as fundamentally crystalline. In an analysis of the optical anisotropy of the waxes from C. cerifera, P. sylvestris and P. australis, Weber (1942) proposed a layered structure in the superficial wax film with vertical orientation of the carbon chains. For terete rodlet waxes he proposed a radial packing arrangement in which various molecular species such as long-chain acids, alcohols and esters might collaborate. Kreger (1949) confirmed by X-ray diffraction studies that EW rods of the type seen in S. officinarum, Arundinaria and Strelitzia are bundles of long crystals, and that their structure must be related to their chemical composition. Prior to the emergence of Bradley and Juniper’s groundbreaking replica technique (Juniper and Bradley, 1958), and the later advent of SEM little further progress was possible, but even then the matter remained unsettled for many years. Weisner (1871) suggested that wax precursors pass through the cuticle in a solvent carrier which evaporates, permitting the wax to crystallise at the surface,

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and by simple analogy with related compounds in petroleum products there is no doubt that the constituents of EW are capable of self-assembly into microcrystals, and that the EW structures on plant surfaces are crystalline, as indicated by Kreger (1949). The crystalline behaviour of long-chain hydrocarbons is of technological and economic importance, but although petroleum products include hydrocarbons similar to those in SCL, the results have not been widely assimilated by plant scientists. Many wax alkanes, acids and alcohols exhibit polymorphic behaviour, and slow evaporation of wax solutions in benzene or xylene can result in more than one type of crystal morphology (Amelinckx, 1955). Birdwell and Jessen (1966) reported that ‘pure n-paraffins exhibit a tabular or platy crystal habit, the most common forms of which are thin hexagonal and rhombic plates’. Homologous series of alkanes generally crystallised compatibly, but EW in which alkanes are dominant constituents show multiple contrasting crystal morphologies, suggesting that the relative proportions of different homologues, or relationships between alkanes and other constituents may be influential. That the structure of EW crystallites is predominantly a process of self-assembly, with the crystal morphology determined by dominant chemical constituents, is now well established both from correlative and genetic observations (Hallam, 1967; von Wettstein-Knowles, 1974), and by experimental analysis of crystal formation in many species (Jeffree, 1974a,b, 1986; Jeffree et al., 1975, 1976; Holloway et al., 1976; Lister and Thair, 1981; Jetter and Riederer, 1994, 1995; Jetter et al., 1996). Whitecross (1963) was the first worker to examine Weisner’s hypothesis (1871) of wax crystallisation from solution experimentally. He found that crystalline structures could be obtained from solutions of Brassica wax in organic solvents such as acetone. Later, Hallam (1967) and Hallam and Chambers (1970) who had observed a general correlation between chemical composition and morphology in Eucalyptus waxes, demonstrated that the primary alcohol-rich plate wax of Eucalyptus ovata and the β-diketone rich tube wax of E. globulus would recrystallise from solution in acetone with crystal forms corresponding to the in vivo morphology, and that the rate of solvent evaporation influenced the crystal structure obtained. Jeffree (1974a,b) developed a method for crystallising waxes and their constituent fractions by feeding solutions of them to a porous evaporating surface using a wick, enabling the crystallisation of waxes from a wide range of species and morphological types, together with isolated and purified constituents of the waxes (Jeffree et al., 1975, 1976; Baker, 1982). These experiments confirmed that the plate-type waxes of eucalypts (Jeffree, 1974a,b), grasses and peas, and the primary alcohol fractions isolated from them, could recrystallise in vitro as plates, while the remainder of the wax failed to reproduce the characteristic plate morphology. (Figures 2.13c,f, 2.15a,b). Jeffree (1974a) used this method to recrystallise the tube wax of Sitka spruce in its original tubular form from solution in chloroform (Figure 2.17c). The chromatographically isolated asymmetrical secondary alcohols from Sitka spruce, comprising 19.7% of the wax, of which 98% is 10-nonacosanol (Holloway et al., 1976), also recrystallised in tube form (Figure 2.17e; Jeffree, 1974a, 1986; Jeffree et al., 1975, 1976), while the remaining 80.3% of the wax lost the capacity to

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recrystallise in tube form (Figure 2.17g). Tube waxes rich in the asymmetrical secondary alcohol 10-nonacosanol isolated from leaves of Chamaecyparis lawsoniana, G. biloba (Figure 2.13k,l), Picea pungens, C. majus, Aquilegia vulgaris, T. majus all recrystallise in tube forms as do the isolated asymmetrical secondary alcohol fractions from these waxes. The secondary alcohol-depleted residues did not form tube crystals. Tube-wax of E. racemosa (Figure 2.15e), containing approximately 30% asymmetrical secondary alcohol (Holloway et al., 1976), recrystallised in a Strelitzia or Brassica-like form, with tubes present but plate-like structures predominating (Figures 2.15f,g), while Agathis australis wax (approximately 28% asymmetrical secondary alcohol) recrystallised as stubby plates without visible tubes (Jeffree et al., 1975). The secondary alcohol fractions isolated from these waxes by preparative scale thin-layer chromatography nevertheless always formed tubular crystals. Important recent data has been added by Jetter and Riederer (1995), showing that a range of alkanediols, present in the waxes of many secondary alcohol tube-forming species, also have tube-forming capability. The crystal sizes formed by x,10-nonacosan-diols, where x is 3, 5, 7, 9 and 13, were found by Jetter and Riederer to be uniform and closely comparable with the diameters of the tubes formed in the waxes of their origin, suggesting that they may be capable of collaborating in wax tube formation with 10-nonacosanol, as suggested earlier for R. cotinus by Hunt and Baker (1979). Similarly, both the tubular β-diketone-rich whole waxes of E. globulus, Eucalyptus glaucescens, D. caryophyllus, T. aestivum, Festuca glauca and H. vulgare, and the β-diketone and hydroxy-β-diketone fractions isolated from them crystallise in tube forms. The hollow form of the recrystallised secondary alcohol and β-diketone tube crystals was confirmed by negative staining (Jeffree et al., 1976; Figure 2.13i,j) as it had been earlier for the natural leaf waxes of Sitka spruce and T. kaufmanniana (Johnson and Jeffree, 1970; Figure 2.17f). Not only is the morphology reproduced in recrystallisation experiments, but also the sizes of the crystals are generally closely similar to the corresponding crystals found on plant surfaces (Hallam, 1967, 1970b; Jeffree et al., 1975; Jetter and Riederer, 1994, 1995).

2.8.1

The tube wax crystal

Hallam (1967) viewed the wax crystals of Eucalyptus cepahalocarpa leaves as tubular, reporting a complete cover of compound branching tubes produced in a 24-h period after the crystals had been removed by rubbing. ‘Two hours after rubbing it was apparent that the tubes were originating from the cuticle as narrow plates, these appearing to roll over and fuse their edges to form a tube’. Tubular crystals in Pseudotsuga menziesii wax probably develop by tight coiling of a narrow filament (Lister and Thair, 1981). Open coils or spirals can sometimes be observed (von Wettstein-Knowles, 1974). Spectacular examples, in which both tubular and helical morphologies occur side-by-side are shown by Jetter (1993) and Jetter and Riederer (1994) from the capsule of P. somniferum, by Neinhuis and Jetter (1995)

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on the gametophyte of Pogonatum rubenti-viride and by Barthlott et al. (1998) in L. korolkovii. Discussing the phenomenon of plates rolling to fuse along their free edges into hollow structures, Hallam (1967, 1970a) advocated use of the term ‘tube’ as more appropriate than ‘rod’ or ‘fibril’. This sequence of events is hard to observe in a dynamic sense, but it is clear today that Hallam was fundamentally correct. There is a spectrum of morphologies between straight ribbon, curled ribbon and tube in all types of tube-forming species, both secondary alcohol and β-diketone types and also in a new category of tube-forming waxes dominated by δ-lactones in C. minor (Jetter and Riederer, 1999a; see also C. major Figure 2.14c). Furthermore, in some species the tubes themselves may become curled or super-coiled as in B. sempervirens and in C. segetum (Meusel et al., 1999). Micrographs at adequate resolution show a spiral line on the surfaces of nonacosanol tubes at about 30◦ to the long axis (Jeffree, 1974a; Figure 2.17d) probably representing the edge seam. A notch is also common at the top of the tube, representing the terminus of the coiling ribbon. Current data suggests that the amount of asymmetry in the crystal-forming compounds is responsible for the tightness of the spiral structure. Branching of the tubes also occurs in many species. In crystals e.g. of quartz, this branching is known as ‘twinning’ and is observed more commonly in β-diketone-tube waxes (e.g. in several species of Eucalyptus; Hallam, 1967) than in the secondary alcohol types. In the experiments carried out by Jeffree, Baker and Holloway (1975, 1976) the isolated alkane fractions and pure alkanes generally formed crystals of a plate type, as also observed by Chambers, Ritchie and Booth (1976), even when their origin was from a wax that appeared amorphous, as in the wax of N. tabacum leaves and lemon fruit (C. limon). Jeffree et al. (1975) found that aldehydes tended to crystallise as elongated plates or ribbons, and suggested that their presence in the alkane-dominated ribbon wax of the abaxial leaf surfaces of peas (P. sativum) might be influential in shifting the crystal morphology away from the isodiametric plate type. This kind of domination of the wax morphology of some conifer waxes was already known in P. pungens, e.g. where 10-nonacosanol is less than 10% of the extractable wax (Holloway et al., 1976) but nevertheless determines that the dominant wax morphology is tubular. However, an alternative and more probable interpretation of the situation in P. sativum is suggested by the findings of Chambers et al. (1976), who showed that mixtures of hydrocarbon, C32 and C20 alkanes with C20 primary alcohol and C12 ketone crystallised as elongated ribbons, distinct from the crystal morphologies of the pure compounds indicating that new crystal morphologies can arise by co-crystallisation of compatible hydrocarbon constituents. The ketone fractions from A. porrum wax (Figure 2.16a), which is 36% palmitone, hentriacontan-16-one (Baker, 1982), recrystallise in a crenate plate form with lobed and spiked edges that approximates the wax type found on the plant surface, lending support to the hypothesis that the wax crystals of A. porrum are predominantly composed of palmitone. Barthlott et al. (1998) referred to the wax of Brassica, Clarkia and Centranthus as ‘polymorphic’, implying that the wax

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contains crystals of several different morphological types, which can be expressed in different proportions when Brussels sprout plants, for example, are grown under contrasting environmental conditions (Whitecross and Armstrong, 1972; Baker, 1972, 1974, 1982). Brassica wax polymorphism may arise from the presence in the wax of several compound classes, each capable of forming distinctive crystal morphologies, but the evidence also suggests the possibility that new crystal forms are specified by co-crystallisation of specific mixtures of these compounds. The wax contains dominant alkanes (nonacosane 46%) and C29 secondary alcohols, in B. oleracea var. gemmifera the symmetrical isomer, 15-nonacosanol being approximately 7.5%, together with about 4.5% of asymmetrical nonacosan-14-ol (Holloway et al., 1976). Both of these compound classes crystallise in plate-like forms in vitro that are not expressed on Brassica leaves (e.g. Figure 2.19d). Ketones, of which 95% is nonacosan-15-one in Brassica (Baker, 1982), account for about 30% of the wax of B. napus and B. oleracea (Shepherd et al., 1995) and the isolated ketone fraction recrystallises to form characteristically spiky-edged plates or dendrites (Figure 2.16d). Furthermore, the leek ketones often form dendritic structures (Figure 2.16c, top left) which recall the dendritic crystals of cabbage leaves (Figure 2.12d) and suggest (Jeffree et al., 1975, Jeffree et al., 1976) that ketones may be involved in the dendritic waxes of Brassica plants grown at high radiant energy and temperature, conditions which promote an increase in ketone production. Certain waxes fail to recrystallise in their in vivo morphology. The most problematic cases are of waxes that have apparently amorphous structure in vivo, but readily form crystallites in vitro, and complex waxes of the Brassica type that recrystallise correctly only under certain circumstances. The whole wax of B. oleracea var. gemmifera recrystallises to produce highly variable (polymorphic) results, even within a single experiment. The resultant morphologies appear to respond to variation in local and global conditions during recrystallisation which are not even crudely understood, such as the carrier solvent, rate of solvent feed, concentration of the wax, porosity of the evaporation surface, rate of evaporation from the surface (cf. Figures 2.19a,c). In our experiments, the most successful reproductions of in vivo morphology from recrystallised Brassica waxes occurred when wax solutions in hexane, chloroform or benzene were delivered to a porous surface via a chromatographic column (silica gel G) that partially separated the constituents in order of polarity, the least polar (alkanes) being delivered first (Jeffree et al., 1975, 1976). Under these circumstances various characteristic elements of the Brassica wax morphology were reproduced successfully (Figure 2.19b). The most significant observation is that to date it has not been possible to reproduce the characteristic fluted columns or the lobed plates by recrystallisation of any of the purified individual compounds or compound classes isolated from Brassica waxes. Similarly, recrystallisation experiments with the Aristolochia-type waxes showed that they could all be recrystallised in vitro from the whole wax with morphologies similar to or approximating the epicuticular morphology of the plant surface in some recrystallisation regime. However, in no instance was it possible to identify a wax

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constituent capable of re-creating any element of the Aristolochia-type morphology (Meusel et al., 1999).

2.8.2

The single-compound hypothesis

Barthlott et al. (1998) stated that ‘Dominant wax components are normally considered to be responsible for the formation of wax crystalloids’. Meusel et al. (1994) stated that ‘the dominant constituent(s) may be responsible for a particular ultrastructure of the crystalloids’. Neinhuis and Jetter (1995) reported that 10nonacosanol is the dominating component in cuticular waxes of gymnosperms and moss sporophytes. Dominant in what sense? The asymmetrical secondary alkanols and alkanediols form tubes on their own, and are decisive in determining the functional morphology of tube waxes, and in some cases (Aquilegia with approximately 56% 10-nonacosanol and Nelumbo with approximately 57% secondary alkandiols, Table 2.5) dominant constituents are responsible for the observed morphology. However, the 10-nonacosanol waxes of gymnosperms rarely contain more than 20– 30% of nonacosanol; so the compound class cannot be said to be quantitatively dominant even in those conifers with emphatic tube wax morphology. Why does the remaining 70–80% not have a say in determining crystal morphology? Individually, there is no reason why several other compound classes present in nonacosanol tube waxes should not form distinctive crystals, but together, in approximately the proportions found on the leaf surface, these compounds appear mutually to suppress the appearance of crystal morphs. This sort of trick applied to metals is the metallurgist’s Holy Grail. Waxes of conifers contain 16-hydroxyhexadecanoic acid (juniperic acid) and 12-hydroxydodecanoic acid (sabinic acid) interpolymerised into sparingly soluble estolides or etholides. These can be a significant proportion of the soluble material isolated from the leaves of conifers (von Rudloff, 1959) but they are not known to be involved in wax crystal formation. In general, across a wide variety of inorganic and organic materials, crystallisation can only take place by segregation of different components of mixtures, co-crystallisation occurring only between compounds with closely similar molecular dimensions and characteristics. In water containing KCl and NaCl in solution, rapid freezing will cause all three components to crystallise separately. Jeffree (1974a) and Jeffree et al. (1976) argued that in common with other crystalline substances, EW will usually crystallise in pure form from mixtures, and that the EW crystals on spruce leaves are likely to be composed of a single compound or closely similar compounds and isomers. Holloway et al. (1976) showed that the secondary alcohol fractions of tube wax species did not usually contain a single compound but homologous series of secondary alcohol compounds and many also contain asymmetrical diols. Likewise, β-diketone waxes also contain homologous series of related compounds, and most waxes of grasses also contain hydroxy-β-diketones that also have the capacity to crystallise as tubes as a purified fraction (Jeffree et al., 1976; Figure 2.13h). EW mechanically detached from E. globulus leaves by simulated rain contained exclusively C33 β-diketone, which is only 55% of the

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solvent-extracted wax from the same leaves (Baker and Hunt, 1986). This may be taken as evidence that the tube waxes of E. globulus are essentially pure crystals of the tritricaontan-14,16-dione or are significantly enriched in it. Equally, the C31 alkane hentriacontane predominated in the material detached from pea leaf abaxial surfaces, suggesting that it is the predominant constituent of the mechanically vulnerable wax ribbons on this species. It is also relevant in this respect that the loss of rodlet (tube) structure was reported to coincide with depletion of the wax in β-diketones when leaves of V. ashei are weathered (Freeman et al., 1979). By contrast with these examples, the Aristolochia-type and Brassica-type waxes represent a suite of types which are capable of self-assembly into crystals, but in which the morphology appears to be determined not by a single dominant constituent or constituent class, but by co-crystallisation of special combinations of compatible compounds (Meusel et al., 1999). Relative proportions of alkanes, secondary alcohols and ketones in rain-detached waxes of Brassica (Baker and Hunt, 1986) matched closely the proportions of the three principal compound classes in the solvent-extracted leaf wax. This may be taken as evidence that the three principal compound classes comprising 92% of B. oleracea wax are mutually responsible for the observed crystalline structure. It seems probable that the same situation applies in the Aristolochia-type species, but in these species it is also likely that the mix of constituents in the crystals differs between species, despite their analogous morphologies. The Brassica and Aristolochia waxes are therefore the first EW classes to be identified in which the crystallite morphologies are not directly produced by individual compound classes, as in the basal tube and plate types, but by a convergent series of different chemical combinations in different plant groups (Meusel et al., 1999).

2.8.3

How do wax crystals grow?

An important issue which is not resolved by these crystallisation experiments is the mechanism by which wax is delivered to the inner surface of the cuticle (see Chapter 5), and its route through the CM. This is part of a much wider series of unanswered questions about the mechanism by which relatively large hydrocarbons with high melting points can move from place to place – both through the developing CM, and then into their final positions in projecting EW crystallites. There is no evidence to support a solvent-carrier based mechanism as proposed by Weisner (1871), but the possibility cannot either be conclusively eliminated on current evidence. Jeffree et al. (1975) considered the idea that mobility of wax molecules through the CM was facilitated by a solvent, such as isoprene. However, isoprene turned out to be a poor solvent for the constituents of secondary-alcohol-type waxes, and attempts to recrystallise them using isoprene failed. Other considerations militate against a solvent-assisted model, not least the facts that many developing plants secrete very little of any likely candidate volatiles, and that wax crystal development is in many species, such as Sitka spruce, already well advanced inside multiple enclosing layers of bud scales from which evaporation of volatiles would be severely

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retarded. EW of B. oleracea varieties gemmifera and capitata also develop while the surfaces are enclosed in layers of older leaves that would strongly inhibit solvent evaporation. As Hallam (1982) pointed out, only very non-polar solvents, such as benzene, hexane and chloroform dissolve EW of most species. These solvents, e.g. chloroform, are also cytotoxic, and present an unsatisfactory model for a carrier system in essentially aqueous plant tissues. Hallam (1982) proposed that wax might be transported through the CW enclosed in an envelope of lipo- or glyco-proteins, which would have hydrophilic external surfaces compatible with the aqueous environment of the CW. Evidence in favour of this hypothesis is accumulating rapidly. Acyl transfer proteins occur in the EW of broccoli (B. oleracea; Pyee et al., 1994; Pyee and Kolattukudy, 1995), and the cer5 gene in A. thaliana encodes a protein that locates to the plasmalemma exclusively in epidermal cells. Disruption of cer5 results in the accumulation of sheets of lipid in the epidermal cytoplasm, accompanied by reduction in the wax loads on the surface. cer5 thus appears to be an ABC (adenosine phosphate binding cassette) transporter responsible for the export of wax from the cell to the plant cuticle (Pighin et al., 2004; Chapter 5; see also Kunst and Samuels, 2003). Are the molecules added to growing tubes at their apices or their bases? Von Wettstein-Knowles (1974) envisaged newly exuded wax accreting to the base of the wax structures, pushing the older polymeric material away from the plant surface. However, for the first time, Koch et al. (2004) have been able to image the development of individual wax crystals by AFM on the living surface of a Galanthus nivalis leaf. Consecutive images of an elongating wax rod (Figures 2.20a–d) show clearly that the extension is occurring at its distal end, not from its base. This finding appears to put the extrusion hypothesis finally to rest, but simultaneously raises again the question of how long-chain hydrocarbon molecules can migrate up the inner or outer surfaces of tubes or plates, to accrete at their distal ends. What form is the wax in – is it mobilised by a solvent, and what is the energy source for such transport?

2.9 Crystal orientation and spatial patterning In a great many plant species both the crystal orientation and the position of the wax crystals relative to surface features such as stomata, may be patterned in various ways. Spatial patterning in wax crystals implies non-randomness, which in turn carries the implication that both the position and orientation of crystals are under some form of control. Tufted, clustered tube wax crystallites are common on the leaves of conifers e.g. Sitka spruce (Jeffree, 1974a; Jeffree et al., 1976) and on developing leaves of T. majus (Rentschler, 1971). These aggregates of crystals give the appearance of arising from a common point of origin. Consequently it is tempting to search for pores beneath. However, the origin of the clustering is ambiguous. Young leaves are expanding, and this area expansion may separate an even distribution of crystals

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into islands, the bare spaces between rapidly filling with new wax (Jeffree, 1974a). Clustered or tufted tubes are also common in β-diketone type tube waxes, e.g. of eucalypts (E. globulus, Hallam, 1967; Eucalyptus camaldulensis, Jeffree et al., 1976). Clustering of plates is commonly observed with clusters of at least three types, those which aggregate into clusters side-by-side (Hypericum bucklei), those which aggregate into radial clusters (Erythroxylum coca, Rentschler, 1971; Calliandra haematoma) and those which aggregate into mutually adherent groups that have the appearance of granules (A. corniculatum, Barthlott et al., 1998). In Prenia sladeniana platelets are orientated transverse to the stomatal aperture, and are co-aligned at various angles in domains about the size of the epidermal cells (Ihlenfeldt and Hartmann, 1982). The most dramatic such patterning occurs in Convallaria majalis, where evenly spaced plates may show parallel alignment over large areas, typically at 90◦ to the stomatal apertures (Barthlott and Frölich, 1983; Barthlott et al., 1998). Frölich and Barthlott (1988) indicate that the Convallaria type wax is ‘more or less confined to the Monocotyledons’, but Dicotyledonous examples are also widespread. Marked alignment of plate type crystals is reported in Chenopodium album leaves (Taylor et al., 1981) and is present in the Apiaceae (Bupleurum ranunculoides; Figure 2.18d). Analogous patterning occurs in plate-forming eucalypts such as E. camaldulensis in which stripes of transversely aligned plates can traverse multiple epidermal cells (Jeffree et al., 1976; Figure 2.12e). Carr et al. (1985) described several types of orientated arrays in Eucalyptus, some of which are closely analogous to the Convallaria type while others form lines, swirls and rosettes of platelets. In many other primary alcohol plate-type species, such as P. sativum, T. repens and Quercus spp., and the Poaceae generally (e.g. Poa nemoralis, Jeffree et al., 1976; and Festuca arundinacea, Figure 2.12c and inset) the spatial distribution is highly regular over large areas and the crystals tend to occur in preferred orientations approximately 120◦ apart. Dramatic wax crystal patterning occurs in the rodlet-type waxes of Saccharum, Musa, Strelitzia and Heliconia where aggregates of co-aligned crystals form massive terete rodlets, the cylindrical chimneys accurately surrounding each stomatal complex in Heliconia being a notable example (de Bary, 1871; Barthlott et al., 1998). In most of these examples the basis of the wax patterning is obscure. Carr et al. (1985) raised the possibility of the involvement of ectodesmata, but their punctate distribution is at odds with formation of large-scale ordered crystal arrays. Kreger (1949) recognised that the direct determination of orientation by the cell membrane must be rejected because of the complexity of the pathway between it and the epicuticle. The orientation of new upright crystals could be influenced by crystal orientation of their neighbours, or by a large-scale crystal orientation in a basal wax film. Kreger (1949) discussed the possible nucleation of radial crystal groups by the first-secreted alcohol molecules, which might be expected to orientate head-to-head on the cuticle surface at 120◦ apart. This model would indicate random orientation of the nuclei themselves, perhaps spawning small-scale ordered domains. However,

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the fact that in Convallaria plate orientation does not occur at random with respect to large-scale surface features such as stomata, around which the crystals orientate like field lines round a magnet (Barthlott et al., 1998), indicates that random seeding of the pattern must be rejected. Wax crystals are always transverse to guard cells, not in alternative orientations. It is worth considering the possibility that wax crystals may epitax onto ordered or semi-ordered cutin polymer in the substrate CP. Chambers et al. (1976) were aware of the possibility of epitaxial growth of crystals on an orientated substrate, but appeared to dismiss the possibility on the grounds that the cuticle is a polymer. However, a striking finding in polymer technology is that transverse epitaxial crystal growth occurs on aligned polyethylene chains (Mackley, 1978). The resulting ‘shish-kebab’ crystals bear tantalising similarity to the linear tracks of primary alcohol crystals on Eucalyptus leaves, which may arise from an analogous process. Since the epidermis is still growing, the cuticle is still undergoing area expansion while wax secretion is taking place. The CM polymer may well be under mechanical stretching during growth, causing molecular alignment of the polymer chains of the CP, on which epitaxial growth of wax crystals can occur. If this hypothesis is true, then the distribution and orientation of the plate-type wax crystals on leaves maps out the distribution and orientation of the underlying cutin polymer chains, and therefore also the stress pattern on the cutin polymers during leaf area expansion. Experimental evidence for the ordered growth of crystals of wheat wax and its major constituent 1-octacosanol on ordered substrates (highly ordered pyrolytic graphite) has now been obtained by Koch et al. who found that wheat wax was reluctant to form crystals on amorphous surfaces such as glass (Figure 2.20e), but crystallised readily on crystalline substrates, the orientation of the wax crystals following the crystal orientation (Figure 2.20f). Crystallisation of wheat wax on isolated C. majalis cuticles produced the typical wax platelets of wheat, arranged in the complex patterns characteristic of C. majalis (Koch et al., in press). These observations strongly indicate that EW crystal orientation is under epitaxial control by an ordered CM structure.

2.10

Degradation of EW

Because of the strong linkage between chemistry and EW crystal morphology it must be assumed that any environmental impacts on wax chemistry will alter the wettability and reflectivity (Chapter 6) of leaves. There has been widespread concern that pollutant impacts accelerate degradation of the EW of conifers, and that this may be responsible for forest decline in Europe and elsewhere. Aldehydes (alkanals) comprising about 35% of the EW of newly expanded leaves of beech (Fagus sylvatica) are susceptible to atmospheric photo-oxidation and decline in amount, alkanoic acids of the same chain length correspondingly increasing (Gülz et al., 1992; Markstädter, 1994). With this exception, most common constituents of EW are very resistant to chemical change in vivo during the normal lifetime of plant organs.

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The leaves of evergreen conifers are long-lived, normally surviving for four– six years in Norway and Sitka spruce, and they must presumably be photosynthetically competent throughout this period. The ancestors of the angiosperms generally had long-lived leaves, and the Lotus effect (Barthlott and Neinhuis, 1997; Neinhuis and Barthlott, 1997) may have been more significant for them than it is today in most mesophytic annuals. In their discussion of the crystal morphology of secondary alkanols and alkandiols, Jetter and Riederer (1995) suggest that the tubular crystals of 10-nonacosanol should spontaneously, but slowly, transform to planar aggregates. If the structure of the tubes is intrinsically metastable, degrading continuously from leaf maturity onwards, this would have important implications for stomatal gas exchange in conifers where leaves persist for 4–6 years, and must presumably earn their keep. There is a large body of observational data in the public domain that documents the progressive weathering and degradation of the EW covering of conifers. Wells and Franich (1977) reported that the density of tubiform wax tufts on P. radiata leaves was greatest near the stomatal pores where the morphology of the wax changed from tubiform to amorphous as the primary needle matured. The tubular structure of conifer leaves is observed gradually to be degraded by a variety of impacts, mechanical from wind, abrasion and ballistic impacts of rain and detritus, chemical from deposited loads of pollutants, and biotic from the activities of epiphytic microorganisms and insects. Degradation of wax morphology has been reported in Pinus strobus and P. banksiana, resulting in the fusion of tubes into plate-like sheets (Riding and Percy, 1985). The images, however, show wax structures consistent with mechanical abrasion as described by Pitcairn et al. (1986) and van Gardingen et al. (1991). Crossley and Fowler (1986) reported a systematic increase in wax tube diameters on Scots pine leaves from approximately 130 nm at time 0 to 220 nm at 30 months leaf age. They also noted that undamaged tubes remained beneath a superficial layer of damaged tubes, suggesting that the change was a consequence of external effects. EW crystals are detached from leaves by repeated droplet-impacts of rain (Baker and Hunt, 1986). The fragile, long, narrow crystal rods, tubes and dendrites of species such as E. globulus and Brassica spp. (the ‘non-structural’ waxes of Barber, 1955) are more susceptible to such damage than the comparatively robust primary alcohol-type plates of grasses (Z. mays, H. vulgare). As waxes degrade, and wettability increases, weathered epicuticular surfaces of conifers may be colonised by fungal hyphae (Chapters 11 and 12) and green algae (principally Chlorococcus) that can begin to be visible to the naked eye from the second year onwards reducing photosynthesis of the whole tree by between 4 and 9% (Peveling et al., 1992). The layers of cells can be sufficiently dense to obscure the abaxial surface of three- and four-year-old Sitka spruce needles almost completely (Figure 2.21f). This epiphytic biofilm forms a surrogate abaxial leaf surface that intercepts radiation, and has properties such as wettability, water absorption capacity and albedo that are different from those of the original leaf surface. However, the tubular wax structures in the stomatal antechamber that function as an antitranspirant mechanism (Jeffree et al., 1971; Figures 2.21a,d,e) remain intact throughout the

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(b)

(c)

(d)

(e)

(f)

Figure 2.21 Low-temperature scanning electron microscope (SEM) images of the stomatal antechamber wax of Sitka spruce (Picea sitchensis).

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lifetime of Sitka spruce leaves (Figures 2.21b,c,e). Therefore the wax tubes do not spontaneously degrade or convert to other forms during physiologically relevant periods of time. Secondary alcohols in plant waxes may be expected to show long-term chemical stability in the natural environment, although Jetter et al. (1996) demonstrated that 10-nonacosanol can be oxidised by strong oxidants such as nitrogen dioxide (NO2 ) to yield the corresponding ketone, 10-nonacosanone and ultimately alkanoic acids that may not have tubular crystal morphologies. Although much higher NO2 concentrations are required than are experienced in the natural environment, the possibility exists that a chemical transformation of this major structure-determining component of the wax may indeed provide the long-sought explanation for the observed degradation of the tube-crystals on the long-lived leaves of conifers growing in polluted environments. The required acceleration of the reaction at physiologically relevant NO2 concentrations may turn out to be accounted for by interactions with other factors in the environment, such as water or particulate matter.

2.11

Summary of cuticle ontogeny

Summarising the process of development of plant cuticles, it is evident that an amorphous procuticle (Figures 2.3a, 2.22a), positioned outside the CW of the earliest epidermal surface, becomes flocculent in appearance (Figures 2.3b, 2.22b) and differentiates into the CP (Figures 2.3c,2.6b and 2.22c). The CP is most often lamellate, but may appear amorphous (Figures 2.2a–g; Figures 2.22c–g, left). The early CP may be subtended by a pectinaceous layer or pectin lamella (Figures 2.6a,b) bonding it to the PCW. Quite rapidly, before leaves are fully expanded, the cutin of the CP may be converted to the non-saponifiable polymer cutan without much, if any, detectable change in ultrastructure. In most species, cutinised layers of the CW begin to develop by impregnation of successive CW layers with cutin polymer. During the impregnation phase, globules of electron-dense cutin precursors enclosed in an electron-lucent lamella accumulate at the base of the CP and appear to fuse laterally first at the surface of the pectic lamella and PCW, and later within successive layers of SCW (Figures 2.6c,d, 2.22e). During the phase of CW cutinisation and at

(Figure 2.21) (a) The completed epicuticular wax of a fully expanded leaf, showing the fully developed antitranspirant wax plug in pristine condition. (b,c) Images of the plug structure in a four-year-old leaf, showing (b) tubular structure retained in the centre of the plug. (c) A view of the base of the plug in a four-year-leaf as seen looking up through the stomatal aperture. The vertical mid-line coincides with the stomatal aperture. (d,e) A vertical cryo-fracture through the wax plug and stomatal complex showing (d) the plug is attached to the accessory cell walls, but not to the guard cells, allowing them to move (e) uniform structure of the intermeshed wax tubes. (f) A dense biofilm of epiphytic algae and fungal hyphae forming a surrogate abaxial (upper in spruce) leaf surface. (a) Bar = 10 μm; (b,c) bar = 1 μm; (d) bar = 5 μm; (e) bar = 3 μm; (f) bar = 100 μm. Figures 2.21a–f by C.E. Jeffree, Figure 2.21d from van Gardingen et al. (1991), Plant, Cell and Environment, 14, 185–193.

108 (a)

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PROCUTICLE PCW

(c)

(d)

CP PCW

(e)

EWF CP CL PCW

EWC EWF CP

(f)

CL PCW SCW EWC EWF

(g)

CF ECL

CL

ICL

CYS

SCW

Figure 2.22 Summary diagrams of the development of the plant cuticle. (a) A uniformly electron-dense procuticle is present on the surface of the primary cell wall (PCW). (b) In the procuticle–cuticle-proper (CP) transition phase, the procuticle becomes flocculent or globular in appearance and (c) is converted to a simple electron-lucent amorphous or lamellate CP positioned outside the cell wall, subtended by a pectinaceous lamella. (d) Further construction of the amorphous or lamellate CP may involve accumulation of electron-dense globular lipids in the PCW beneath the CP. (e) The CP/CL (cuticular layer) transition. Within the primary cell wall polysaccharide matrix, globules of electron-dense lipids coated with electron-lucent shells construct the CP and the CP/CL transition zone.

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maturity, the CL may contain either paradermal or more random lamellate structure closest to the CP, but lamellae may extend to any depth within the CL (Figure 2.22g). The lamellae apparently derive from the electron-lucent envelope that encloses the electron-dense globules. Soluble epicuticular lipids begin to accumulate and crystallise at the CP surface throughout the process of cutinisation (Figures 2.11a–d) while the leaf is still not fully expanded. The transport of wax to the surface occurs across the continuously lamellate or amorphous CP pores without disturbing the ordered appearance, and no pores can be detected by microscopy. The outer CL layers (ECL) undergo maturation of the polymer matrix (Figures 2.6e–h, 2.2g, 2.4a–e), involving intensive cross-linking of the existing cutin and its progressive conversion, together with any embedded polysaccharide to the aliphatic polymer cutan, which may be impregnated with SCL. Within thick SCW, further accumulation and accretion of CL-like material may occur at locations detached from the main CL deposit (Figures 2.1c and 2.2g) as cutin cystoliths, which may range from the basic size of the pre-cutin globules to several hundred nanometres in diameter. Later stages of development of the CW may involve deposition of inorganic material (silica and calcium salts) in and beneath the CL (Postek, 1981; Davis, 1987; Tenberge, 1989, 1992), and progressive lignification of the underlying SCW (as exemplified by Picea; Tenberge, 1989, 1992). The EW layer is seen as forming a continuous basal film covering all of the cuticle surface that may be crystalline even if the crystallites cannot be resolved by microscopy. Crystallites of wax of various morphologies may emerge from the basal wax layer. They are capable of growth at their distal tips and edges, though other possibilities are not discounted. Crystallites form by self-assembly,

(Figure 2.22) The globules appear to assemble in situ beneath the CL, and are not observed traversing the PCW or SCW (secondary cell walls). Lamellae may become less regular within the PCW. An apparently amorphous epicuticular wax film is now present on the surface of the CP, which may be terminated by a single, continuous electron-lucent lamella. (f) Incorporation of PCW in the CL. The pectin lamella is no longer present. Reticulations are light in the outer CL and predominantly radial, reaching as far as the base of the CP. Lamellae may be absent, or occur to any depth and orientation within the CL, which is globular at its base. Epicuticular wax crystals begin to form before cell expansion has ceased. Thickness of the CP is maintained during rapid cell expansion. (g) Following the cessation of cell expansion, external and internal layers of the CL (ECL and ICL) may become chemically and structurally distinct. In the ECL, electron density of the matrix may diminish, and the boundary with the CP may lose contrast. Polysaccharide fibrils marking the CP/ECL boundary may also disappear. Reticulate polysaccharide formed in the PCW/SCW transition is usually more massive, denser and more three-dimensional than in the ECL. Crystalline epicuticular wax layer is well developed. In the mature cuticular membrane, the CP may be amorphous or any grade from weakly to strongly lamellate. The ECL may be nonreticulate, or contain reticulum throughout. The CL may contain lamellae extending to any depth from the CP. Cutinisation of the SCW may occur at points detached from the main body of the CM, forming cutin cystoliths (CYS). This developmental scheme, and variations on it, particularly with respect to the thickness of the various layers, appears to apply to the cuticles of all land plants. Further elaborations in mature cuticles include the impregnation of the CL with additional materials such as silica, calcium oxalate or lignin.

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their morphology determined by their chemical composition. Individual crystallites may in some instances be pure crystal domains of an individual constituent of the EW. In other instances particular combinations of two or more compounds may collaborate to form a given crystal morphology. The orientation of wax crystallites may be controlled by their mutual interaction and by epitaxial growth on a CP in which the polymer structure has become aligned by stretching during cell expansion.

Acknowledgements This chapter would not have been possible without the generosity of Peter Holloway in donating to me his entire collection of electron microscope negatives of plant cuticles. Grateful thanks are also extended to the following for permission to use their images to illustrate this review: Sergio Archangelsky, Martha Cook, Henning FrostChristensen, Matthew Jenks, Henning Heide-Jørgensen, Kerstin Koch, Christiane Nawrath, Nick Read, Markus Riederer, Fred Sack, Jörg Schönherr, Patrick Sieber, Klaus Tenberge and Joachim Wattendorff.

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Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

3

The cutin biopolymer matrix Ruth E. Stark and Shiying Tian

3.1 Introduction: protective plant polymers The leaves, fruits, and primary stems of higher plants are covered by a cuticular membrane that occupies approximately the outer 0.1–10 μm of the aerial plant surface. Chemically, the membrane consists of a variety of waxes that serve as waterproofing and the biopolyester cutin that functions as a densely networked structural support (Figure 3.1). As detailed in Chapter 4, the waxes are organized as both epicuticular lipids coating the outer surface of the cuticle and intracuticular lipids embedded within the cutin matrix. The cuticle controls the interactions of the plant with the environment, protects against water loss (Chapters 8 and 9), functions as the plant’s primary protective barrier against pathogenic attack (Chapters 12 and 13), and is thought to be necessary for plant organ development (Chapter 10; Kolattukudy, 1980, 1984; Baker, 1982; Holloway, 1982; Kolattukudy and Espelie, 1989; Walton, 1990; Sieber et al., 2000; Heredia, 2003). Moreover, the ω-hydroxy fatty acid monomeric constituents of cutin are thought to be involved in plant– pathogen interactions (Chapter 12; Heredia, 2003). In terms of agricultural impact, the breakdown of cuticular membranes by bacterial and fungal microorganisms contributes to an estimated 20% annual loss from crop damage worldwide (Oerke et al., 1994). Epicuticular surface coated with waxes

Removed cell walls

Cutin support and intracuticular lipids Figure 3.1 The architecture of the cuticular membrane and spatial organization of cuticular lipids, as illustrated by scanning electron micrographs of tomato fruit cuticle before and after dewaxing (left and right panels, respectively).

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Among the protective plant polymers, cutin is distinguished from cutan, suberin, and lignin by several criteria. Lignins are the dominant biopolymers in vascular plants and woody tissues where their accumulation enhances the structural integrity of cell walls (Davin and Lewis, 1992); suberins share some protective and waterproofing functions with cutins but are found in underground roots, in cork, and at wound-healing tissue surfaces. Cutin is the polymeric support for membranes that cover the aerial parts (leaves, fruits, primary stems) of higher plants, whereas cutan refers to the resistant polymeric residue from dewaxing and saponification treatments (Boom et al., 2005). Whereas lignin and suberin have substantial phenylpropanoid character, both cutin and cutan are primarily aliphatic polymers containing long methylene chains.

3.2 Biosynthesis Both the biosynthesis of cutin monomers within cells and their extracellular assembly to form the cutin biopolymer have been the subject of longstanding study. Through studies of radio-labelled precursors and intermediates, the biochemical pathways for both C16 and C18 monomers were established (Figure 3.2). For instance, it was shown in Vicia sativa that oleic acid undergoes ω-hydroxylation and enantioselective epoxidation of the double bond and then hydroxylation of the latter functional group (Kolattukudy, 2001). Catalysis of the proposed transformations was shown to involve an epoxide hydrolase and a cytochrome P450-dependent ω-hydroxylase (Pinot et al., 1992, Heredia, 2003). It has been three decades since the assembly of these monomers to form the cutin biopolymer was shown to be catalyzed by enzymes associated with epidermal cells and required both ATP and CoA (Kolattukudy, 1984), but the putative hydroxyacyl-CoA, cutin transacylase, that accomplishes the transformations has proven elusive. More recently, a new cutin acyltransferase and its corresponding gene from Agave americana leaves have been described in a preliminary report (Reina and Heredia, 2001).

3.3 Monomer composition The monomeric constituents isolated from both leaf and fruit cutins consist predominantly of saturated hydroxylated aliphatic acids, usually a mixture of C16 and C18 homologues. In some cutins a single chain length is predominant: C16 in tomato fruit (Lycopersicon esculentum) and broad bean leaf (Vicia faba), C18 in spinach leaf (Spinacia oleracea). Other species such as apple fruit (Malus pumila) include primarily unsaturated C18 monomers, similar to the suberin constituents of potato tubers and tree bark. No cutins have been detected in algae or fungi; cutins that lack C18 monomers are in general characteristic of gymnosperms and lower plants. Surprisingly, similar chemical constituents may predominate in plants that belong to very different families.

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Acetyl-CoA + 7 Malonyl-CoA NADPH CH3-(CH2)14-CO-S-ACP

CH3-(CH2)16-CO-S-ACP

CH3-(CH2)14-COOH

NADPH/O2

NADPH/O2 CH2-(CH2)14-COOH

CH3-(CH2)7-CH=CH-(CH2)7-CO-S-ACP

OH H2O

NADPH/O2

CH2-(CH2)5-CH-(CH2)6-COOH OH

CH3-(CH2)7-CH=CH-(CH2)7-COOH

OH

NADPH/O2 CH2-(CH2)7-CH=CH-(CH2)7-COOH OH

ATP, CoA NADPH/O2

CH2-(CH2)7-CH-CH-(CH2)7-CO-SCoA OH

O H2O

CH2-(CH2)7-CH-CH-(CH2)7-CO-SCoA OH

OH OH

Figure 3.2 Biosynthetic pathways for the formation of C16 and C18 hydroxyacid families of cutin monomers. Adapted from Kolattukudy, 1984.

In both C16 and C18 acids ω-hydroxylation is very common, and additional hydroxy or epoxy substituents are often present at midchain position(s). For instance, common monomers include 9,16- and 10,16-dihydroxyhexadecanoic acids and 9,10-epoxy-18-hydroxyoctadecanoic acids. Moreover, aldehyde, ketone and carboxyl groups have been reported as additional substituents and chain lengths up to 20 have been observed (Holloway, 1982; Ray et al., 1995; Tian, 2005). Small quantities of coumaric and ferulic acids have also been reported (Baker et al., 1975;

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Riley and Kolattukudy, 1975), and both glycerol and a series of glyceryl esters have been found recently in cutins from diverse leaf and fruit cuticles (Graca and Pereira, 2000; Graca et al., 2002). Cutin content may vary with the membrane thickness of the plant. Amounts range from approximately 10 μg cm−2 for membranes 0.1 μm thick to approximately 500 μg cm−2 for membranes 5 μm thick; typically 40–80% by weight of the cuticle consists of cutin (Kolattukudy, 1980; Walton, 1990; Heredia, 2003). Fruit cuticles in particular may exceed these amounts, for example, some apple cultivars can contain 1.5 mg cm−2 at maturity. Cutin chemical composition within a given plant can vary with organ (leaf or fruit) and location (adaxial or abaxial surfaces of the same leaf). Although many investigators have found only minor differences in cutin composition as leaves and fruits develop, others have noted the dominance of one or the another monomer at different stages of maturity, for instance, in Vicia faba leaves and M. pumila fruits (Holloway, 1982). Although the trends above are well established, several pitfalls in the determination of cutin monomer composition should be noted. The typical chemical degradation reagents (methanolic KOH, LiAlH4 , BF3 –CH3 OH, etc.) produce soluble products but also often leave behind at least 20% by weight of nondepolymerized residue, so that the resulting soluble monomers specify cutin composition in an incomplete and potentially misleading manner. For instance, it has been proposed that nonester cutin structures or heavily cross-linked regions are resistant to chemical degradation (Riederer and Schönherr, 1988; Ray et al., 1998). Conversely, incomplete dewaxing of the cutin may lead to identification of monomers that do not come from the biopolymer at all. Moreover, the combination of gas chromatography and electron-ionization mass spectrometry that became customarily available in the 1970s to identify the soluble compounds may fail for species that are nonvolatile or prone to fragmentation (Holloway, 1982, 1984; Kolattukudy, 1984).

3.4 Polymeric structure of intact cutin Plant cutins were identified as polyesters at least three decades ago based on their susceptibility to fungal degradation by hydrolytic enzymes (Kolattukudy, 1984) and consistent with the ester-forming ability of hydroxyacids. Even without direct evidence from oligomers or mature polymers, the fact that half of the C16 monomers contain a free midchain hydroxyl group suggests that they are involved in either cross-linking or branching (Holloway, 1982; Kolattukudy, 1984). Moreover, it has been suggested that ether bonds, serving as possible cutin cross-links, could protect plants from fungal invasion (Blée, 2002). Nonetheless, even the most complete information on monomer composition falls short of determining the polymeric architecture that is a likely key to the function of the cutin as a protective plant membrane. For intact (insoluble) cutins, chemical functionalities may be identified using solid-state nuclear magnetic resonance (NMR) or infrared (IR) spectroscopies.

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(CH2)n

CH CH2

CO

22

20

18

16

14

12 13

10

80

60

40

20

0

–20

C chemical shift (ppm)

Figure 3.3 75 MHz cross polarization-magic-angle spinning (CPMAS) 13 C NMR spectrum of dry dewaxed tomato fruit cutin, obtained with an 8-kHz spin rate and referenced to tetramethylsilane via external hexamethylbenzene. The major NMR resonances are identified with the designated carbon functionalities in the biopolymer structure. From Batteas and Stark, 2005, by permission from Woodhead Publishing.

For instance, the cross polarization-magic angle spinning (CPMAS) 13 C NMR spectrum (Schaefer and Stejskal, 1979) of tomato fruit cutin (Figure 3.3) contains resonances from chain methylenes (30 ppm), CH2 O and CHO groups (64 and 72 ppm), and carboxyl groups (168–173 ppm), where the assignments are made provisionally from reference data on similar chemical compounds. Additional aromatics and/or olefinic signals appear at 105–150 ppm in the spectrum of lime (Citrus aurantifolia) fruit cutin (Zlotnik-Mazori and Stark, 1988), where the relative numbers of various carbon types have also been determined from NMR spectra acquired under different cross-polarization and decoupling conditions (Table 3.1). In both cases, the structural heterogeneity and amorphous nature of the biopolymers limit the resolution of the rather broad spectral lines, though cross-link sites may be identified because their motional restriction produces characteristically inefficient spin relaxation in the solid state. In a complementary fashion, Fourier transform infrared (FTIR) spectra also serve to identify and quantify molecular groupings in bulk cuticular samples (Luque et al., 1995; Ramirez et al., 1992). For instance, a recent report used this methodology to demonstrate augmented cross-linking in mature (ripe) tomato fruit cuticles (Benitez et al., 2004). The spectral assignments described above should be viewed as limited: as noted, they are made simply by analogy with similar chemical compounds; moreover they can only identify small molecular moieties. These shortcomings have been addressed using high-resolution magic-angle spinning (HR-MAS), a hybrid NMR technique

131

THE CUTIN BIOPOLYMER MATRIX Table 3.1 Chemical composition and dynamics of lime fruit cutin from solid-state NMR1

Carbon type (CH2 )n CH2 OCOR CHOCOR Aromatics, alkenes CH2 OCOR CHOCOR C=O

% Mobile2

%Rigid3

45

32 1.7 7.1 3.8 0.9 4.9 0.7

T1 (C) for rigid carbons (ms) 145 122 >7000 ∼1000 ∼100 ∼1700

1 Adapted from Zlotnik-Mazori and Stark, 1988 and Garbow and Stark, 1990. 2 From comparison of integrated intensities for spectra with direct-

polarization/low-power decoupling and cross-polarization/high-power decoupling (zero contact time), both acquired with magic-angle spinning. 3 From extrapolation of cross-polarization 13 C NMR signal intensities to zero contact time.

in which solid samples are swelled in organic solvents to enhance molecular mobility and slow MAS produces high-resolution spectra (Keifer et al., 1996; Millis et al., 1997). It may then be possible to confirm directly or remotely bound 1 H–13 C pairs within a functional group, establish covalent connectivities between monomer units, and discern which groupings are close to one another in space. This approach is illustrated for lime fruit cutin in Figure 3.4, where the MAS-assisted 2D 1 H–13 C correlation (HMQC) NMR spectrum of the intact lime cutin polymer displays resolution approaching the soluble monomers and oligomers (discussed later) and reveals directly bonded C–H pairs that discriminate between resonances of, for example, CH3 , (CH2 )n , CH2 C=O, and CH2 O groups in the various materials (Hurd and John, 1991; Fang et al., 2001; Tian, 2005). Moreover, HMQC in conjunction with 2D 1 H– 1 H Total Correlation SpectroscopY (TOCSY) has provided evidence in tomato fruit cutin for α-branched carboxylic acids and esters, midchain (secondary) alcohols, and olefinic groups (Deshmukh et al., 2003). Thus, in addition to offering a direct assessment of cutin molecular structure, the HR-MAS NMR strategies confirm the presence of both expected monomeric building blocks and a small number of novel cross-linking elements that are likely resistant to chemical degradation.

3.5 Molecular structure of cutin fragments 3.5.1

Oligomeric degradation products

A complementary approach to understanding the protective functions of fruit cuticular polymers stakes out a middle ground between the identification of cutin hydroxylated fatty-acid building blocks from depolymerization products (Kolattukudy, 1984) and the examination of chemical functionalities and cross-link

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BIOLOGY OF THE PLANT CUTICLE

1

Monomer with oxo group

Monomer with hydroxyl group

2

2 2 3

3

4

4

4

1

Trimer

Cutin

1

2

2

1H

Chemical shift (ppm)

3

2

3

3

DMSO 3

4

5

5 70

60

50

40

30

20 13

70

60

50

40

30

20

C Chemical shift (ppm)

Figure 3.4 600 MHz MAS-assisted two-dimensional NMR of lime fruit cutin and its molecular building blocks, illustrating the characteristic spectral signatures of each material dissolved or swelled in DMSO at 50◦ C (Tian, 2005). These 1 H-13 C heterocorrelated multiple-quantum coherence (gHMQC) data were obtained with 2-kHz spinning and referenced to tetramethylsilane. The spectral peaks in boxes 2, 3, and 5 of the contour plots signify single-bond interactions for three chemically distinct types of methylene groups.

constraints for intact materials: mild enzymatic and chemical procedures are used to produce soluble oligomers that nevertheless retain many essential covalent linkages within the cuticular support structure. The first report of such results (Osman et al., 1995) used mild alkaline hydrolysis and proposed a primary ester of two ω-hydroxyhexadecanoic acid molecules but noted that dimers and higher oligomers are difficult to confirm by mass spectrometry (MS) because of their low abundance and modest volatility. In our laboratory, commercial lipases, iodotrimethylsilane, low-temperature HF, and methanolic KOH have each been used along with HPLC separations to generate oligomeric fragments from lime fruit cutin (Ray and Stark, 1998; Ray et al., 1998; Stark et al., 2000; Fang et al., 2001). In addition to the need for specialized MS methodologies, all of these studies confront challenges that include crude product mixtures in which monomers dominate but the numerous

133

THE CUTIN BIOPOLYMER MATRIX OH | -CHCH2CH2-

1.0

-CH2COO-

2.0 2.5

-CH2C(O)CH2-

3.0

1H

chemical shift (ppm)

1.5

CH3OC(O)-

3.5 4.0

-CH2OC(O)220

200

180

160

140 120 100 80 13C chemical shift (ppm)

60

40

20

0

Figure 3.5 600 MHz two-dimensional NMR of a soluble lime fruit cutin trimer (7) from KOH hydrolysis, containing two units of 10,16-dihydroxyhexadecanoic acid and one unit of 10-oxo-16-hydroxydecanoic acid (Tian, 2005). The contour map from this 1 H-13 C gradient-assisted heterocorrelated multiple-bond correlation (gHMBC) experiment (Rinaldi and Keifer, 1994) primarily displays three-bond interactions between carbon–proton pairs.

information-rich oligomers are rare, similar polarities that make separation of the various oligomers difficult, and functional groups that lack chromophores suitable for UV detection. The NMR and MS spectroscopic identification protocols used for our lime cutin oligomers are illustrated in Figures 3.5 and 3.6. In contrast to the MAS-assisted 2D 1 H–13 C correlation (gHMQC) NMR strategy described earlier, 1 H–13 C multiplebond correlation spectroscopy (gHMBC; Rinaldi and Keifer, 1994) carbon– hydrogen pairs identifies that are separated by three or four chemical bonds, making it an invaluable tool for the definitive identification of nonprotonated ester carbons and connections between monomer units. Either electrospray or atmospheric pressure chemical ionization MS (Smith, 1999) yield the molecular weight of each purified oligomer, confirming information from 1 H NMR integrations. The order in which the monomers are connected can occasionally be determined from fragments obtained by MS, and the presence of mixtures containing both C16 and C18 units may be deduced. Whereas a unique pentamer linked exclusively by secondary esters (1) (Figure 3.7) was obtained from reaction of lime fruit cutin with porcine pancreatic lipase, most of the isolated oligomers have primary ester linkages and midchain hydroxyl or oxo groups; all possible combinations of these units are found in the dimers and trimers (Figures 3.7 and 3.8). Although it is possible to obtain more and

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BIOLOGY OF THE PLANT CUTICLE

858

100

Relative intensity (%)

80

60 863 40

841

20

0 820

840

860

880

900

920

940

m/e (amu) Figure 3.6 ESI/MS spectrum of 7, showing molecular ions corresponding to M + NH+ 4 (m/e 858), M + Na+ (m/e 863), and M + H+ (m/e 841) (Tian, 2005). The 13 C isotopic peak intensity is about 55%, consistent with a trimer structure containing 49 carbons.

larger oligomers using an apparatus that permits quenching of the degradative reaction (Tian, 2005), the resulting products are numerous and similar in polarity, thus difficult to purify in amounts sufficient for full structural characterization. Only the products of alkaline hydrolysis have been found to contain any secondary hydroxyl groups or secondary esters, despite prior reports of such functional groups in intact cutin from lime and tomato fruits, respectively (Zlotnik-Mazori and Stark, 1988; Deshmukh et al., 2003). The overall scarcity of such connectivities among all oligomers could stem from either light cross-link density or inefficient hydrolysis of the densest polymeric regions.

3.5.2

Polymeric residues from chemical degradation procedures

After exhaustive degradation of leaf cutin by alkaline hydrolysis or transesterification there remains a cutan residue thought to consist of cutin monomers connected by nonester bonds. Application of a variety of physical methods has suggested that this nonsaponifiable material contains an ether-linked network of methylene chains, double bonds, and carboxyl groups (Villena et al., 1999). However, treatment of lime fruit cutin with iodotrimethylsilane or with HF at low temperature leaves behind unreacted residues in which esters of secondary alcohols are evidenced by resonances in their respective NMR spectra (data not shown).

135

THE CUTIN BIOPOLYMER MATRIX O

HO O

OH

I

I O

O I 1

3

O O

O

CH3

O

O

CH3 O

OH O (CH2)13CH3

O

O I O O

I

OH O

4

O

I

O

CH3

I

I O

O

O

O

O O 2

O

O CH3 O

Figure 3.7 Oligomers obtained from partial degradation of lime fruit cutin with porcine pancreatic lipase (1, Ray and Stark, 1998) and with iodotrimethylsilane (2–4, Ray et al., 1998), respectively.

3.6 Mechanical properties 3.6.1

Methodology

As for synthetic polymeric materials, it is expected that the mechanical behavior of plant cuticular biopolymers will influence adsorption, diffusion, and cracking – with practical agricultural consequences for the foliar application of chemicals, control of water loss, and maintenance of fruit appearance, respectively. Various physical methods offer complementary views of the surface and bulk mechanical properties of plant cuticular membranes. As discussed in Chapter 1, atomic force microscopy (AFM) provides the opportunity for in situ characterization of the surface – regarding both topology and mechanical response of plant tissues – but it lacks true chemical

136

BIOLOGY OF THE PLANT CUTICLE O HO

O

OH

O

O

5

O

6

CH3 O

O CH3

CH2

O

O

O

O

O

O

O

O

O

O HO

HO

O

O OH

OH

7

8

O

O

CH3

CH3

O O

O

OH O

O

O O

O O

OH OH

HO O

9

O CH3

O O

O

O O

Figure 3.8 Oligomers obtained from partial degradation of lime fruit cutin: a dimer (5, Fang et al., 2001) and a trimer (6) from low-temperature HF treatment; trimers esterified through primary alcohols (identified provisionally as 7 and 8); a trimer from methanolic KOH that displays mid-chain esterification through a secondary alcohol (9) (Tian, 2005). Arrows designate 1 H-13 C connectivities deduced from HMBC NMR experiments.

THE CUTIN BIOPOLYMER MATRIX

137

specificity. Applying NMR provides only average (bulk) properties but can reveal detailed information on molecular composition and architecture (described earlier) and offer insight into the dynamics of the component molecules of plant tissues, which may be related in turn to the bulk mechanical behavior of the system. Both of these techniques can complement and extend the stress-strain approaches reported by Petracek and Bukovac (1995), as illustrated for tomato and lime fruits later. Although AFM is becoming well established as a tool to provide the three-dimensional nanoscale architecture of biological materials directly under physiological conditions (Kirby et al., 1996; Mechaber et al., 1996; Round et al., 2000; Batteas and Stark, 2005) its different imaging modes also yield mechanical parameters such as surface viscoelastic response. In contrast to bulk rheology, which reflects elastic response parallel to a membrane, the AFM tip in a nanoindentation experiment senses the mechanical resistance to deformation in a direction perpendicular to the surface, the route taken by many pathogens. The Hertz model of contact mechanics (Hertz, 1882) is then used to deduce the Young’s modulus of elasticity from plots of resistive force versus indentation distance, plotted for both the sample and an ideal hard surface (silicon). Alternatively, lateral force microscopy probes the torsional response of a cantilever as it is scanned over the surface of interest, reflecting both tip-sample adhesive interactions and shear stress. Finally, surface viscoelasticity may be examined by setting the modulation amplitude and frequency of the AFM tip against a surface and then mapping changes in these settings with position. Whereas the NMR spectra described earlier for solid cutin polymers and the corresponding monomers and oligomers in solution can identify chemical groupings and establish molecular architecture, it is NMR spin relaxation that adds complementary molecular-level detail to AFM-based assessments of cuticular surface mechanical properties. Spectral linewidths and spin-relaxation rates reflect supramolecular organization and site-specific polymer dynamics on several timescales, both of which serve to link molecular structure and bulk mechanical properties (North, 1975; Schaefer and Stejskal, 1979; Bovey and Mirau, 1996). For instance, 1 H NMR linewidths in a two-dimensional wide-line-separation (WISE) experiment (Schmidt-Rohr et al., 1992) measure the extent of motional narrowing for each carbon type, whereas rotating-frame relaxation rates probe cooperative motions and cuticular resiliency as well as the organization of cutin and wax domains (Schaefer and Stejskal, 1979; Garbow and Stark, 1990).

3.6.2

Measurements of surface elastic modulus

In order to examine how the elasticity of the cuticular membrane changes as a function of environmental variables such as humidity, force-distance measurements were conducted on dewaxed and native tomato cuticle samples under controlled hydration conditions (Round et al., 2000). The resulting elastic moduli (resistances to deformation) are summarized in Table 3.2. For dry samples, the lipid-covered cuticle surface has an elastic modulus five times smaller than the dewaxed cutin

138

BIOLOGY OF THE PLANT CUTICLE Table 3.2 Surface elastic moduli of tomato fruit cuticles from atomic force microscopy Relative humidity 0% 30% 60% Water

Dewaxed cuticle (E, MPa)1

Native cuticle (E, MPa)

32 ± 11 9.2 ± 1.5 5.5 ± 1 5.8 ± 2

6.2 ± 1.7 — — 5.7 ± 2.5

1 From Round et al., 2000.

polymer. Whereas the modulus of cutin drops fivefold upon hydration (Round et al., 2000), the elasticity is invariant to hydration state for native cuticle. There are several plausible explanations for this last observation: AFM does not sense the underlying polymeric support; wax mitigates the effects of hydration on the cuticle; or the moduli cannot be determined precisely for heterogeneous materials such as lipid-covered cuticular surfaces. As reported previously for amorphous synthetic polymers (Galuska et al., 1997; Lee et al., 1997), AFM-based lateral force measurements also provide a means to evaluate the impact of temperature on the mechanical properties of the cuticular surface in situ (Round, et al., unpublished observations). The ‘drag’ of the AFM tip as it slides across the surface is found to increase with temperature as described previously (Batteas and Stark, 2005). The onset temperature of these changes (approximately 28◦ C) is consistent with a phase transition observed previously at 30◦ C for extracted epicuticular lipids (Luque and Heredia, 1997), though the surface becomes too soft above 45◦ C to make additional AFM measurements.

3.6.3

Measurements of bulk molecular dynamics

NMR relaxation measurements on intact lime and tomato fruit cuticles have also proven to be a rich source of molecular-level dynamic information, which is linked in turn to (bio)polymer mechanical properties (North, 1975) and complementary to determinations of bulk and surface rheology. Each of the major polyester functional groups identified in the CPMAS 13 C solid-state NMR spectrum of dry tomato cutin (Figure 3.3) may be characterized in terms of its molecular motion. One measure of the dynamic properties of the various carbon segments uses the WISE experiment described earlier (Schmidt-Rohr et al., 1992) and illustrated in Figure 3.9. Chain methylenes resonating at 30 ppm have motionally narrowed lines as compared with ester-linked methylenes at 72 ppm; even among the bulk methylene groups the 1 H linewidths discriminate between mobile and rigid components that may correspond to chains and cross-links, respectively. In parallel with the surface elasticity determinations made under stress conditions involving hydration, abrasion, and temperature, NMR relaxation has been used to understand the AFM trends in terms of dynamic behavior at specific molecular

139

THE CUTIN BIOPOLYMER MATRIX

0

20

40

80

100

120

13

C chemical shift (ppm)

60

140

160

180

200

–

–

–

–

– 1

0

10

20

30

40

50

H linewidth (kHz)

Figure 3.9 Contour plot from a 2D 1 H-13 C WISE NMR spectrum of dry tomato fruit cutin. The chain methylenes (30 ppm 13 C) exhibit superimposed 1 H spectral patterns with linewidths of 5 and 35 kHz, whereas the ester-linked methylenes (72 ppm 13 C) have a corresponding 1 H linewidths of ∼25 kHz.

sites of the cutin biopolymer. Table 3.3 shows how the linewidths and spin relaxation rates vary with relative humidity and temperature for both dewaxed and native tomato cuticle samples. Upon hydration of the dewaxed samples, both the diminished linewidths of the (CH2 )n resonances and greater proportion of the narrow spectral component indicate enhanced segmental motions at rates ≥50 kHz, linked previously to a corresponding decline in the resistance to deformation (North, 1975). An analogous trend of increasing segmental motions with rising temperature is evident from the 13 C spin-lattice relaxation rates. These motional trends are also in accordance with the drops in surface and bulk moduli observed with AFM and rheological methods. Moreover, even though water may displace hydrogen-bonded

140

BIOLOGY OF THE PLANT CUTICLE

Table 3.3 Flexibility of tomato fruit cuticle from solid-state NMR spectral measurements Conditions

Dewaxed cuticle

Native cuticle

Room Temperature 1 H Linewidth (kHz)1

Dry (0% RH) Saturated

Dry (0% RH) Saturated Ambient (50% RH) 253 K 263 K 273 K 283 K

Broad component 38.2±3.82 31.3 ± 3.12 Cutin (CH2 )n 3.6 ± 0.1 3.68 ± 0.02 3.00 ± 0.2 3.38 ± 0.2 3.66 ± 0.2 3.86 ± 0.2

Narrow component 5.3 ± 0.52 4.7 ± 0.52

% Narrow 9 ± 22 18 ± 42

Broad component 30 ± 5 27 ± 5

Room temperature R1 (C) (sec−1 )3 Wax (CH2 )n Cutin (CH2 )n — 3.2 ± 0.2 — 1.5 ± 0.2

Narrow component 5.5 ± 0.5 5.0 ± 0.5

% Narrow 25 ± 5 15 ± 3

Wax (CH2 )n 3.0 ± 0.2 3.1 ± 0.1

Variable temperature R1 (C) (sec−1 )4

1 Proton linewidths corresponding to the bulk methylene groups resonating at 30–35 ppm. 2 From Round et al., 2000. 3 R (C) ≡ 1/T (C) is the average spin-lattice relaxation rate for each carbon type. Values were derived from the 1 1 short-time behavior (10–320 ms) of 13 C magnetization observed after a CP inversion-recovery pulse sequence (Torchia,

1978). The line broadening used to apodize the NMR data was reduced to 15 Hz in order to resolve cutin and wax resonances at 30 and 33 ppm, respectively.

4 Values of R (C) were determined as described above using a nominally identical tomato cutin sample. 1

cross-links within the cutin polyester, the NMR results show that remote chain segments become more flexible in hydrated tomato cutin. But as deduced from AFM measurements, no water-induced plasticizing effects are evident in the NMR data for native cuticles; in fact, both the proportion of narrow 1 H components and rates of 13 C spin-lattice relaxation evidence less efficient segmental dynamics for the chain methylenes. Physically, it is likely that strong acyl-chain interactions between the cutin and wax constituents both block the plasticizing effects of water and attenuate the chain segmental motions.

3.7 Thermodynamic properties As noted above, phase transitions have been monitored for cuticular lipids as well as intact cuticle (Luque and Heredia, 1997; Casado and Heredia, 2001b; Matas et al., 2004). Glass transition temperatures (Tg ) were found to be 23◦ C for both tomato cuticle membranes and the cutin itself, but saturation of the membranes with water reduced Tg to about 16◦ C (Matas et al., 2004). Differential scanning calorimetry (DSC) also revealed an anomalously high value of specific heat (2.0–2.5 J K−1 g−1 ) for tomato cutin (Luque and Heredia, 1997; Casado and Heredia, 2001a,b), suggesting a thermoregulatory role in leaves and fruits. Given the potentially deleterious effects of heat waves and frosts on fruit viability, it is

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141

important to develop a molecular understanding of associated changes in cuticular surface structure, segmental motion, and cutin-wax domain architecture that may degrade the mechanical behavior of the cuticle so that cracking of the polymeric veneer is promoted and the tissue below becomes susceptible to pathogenic attack (Garbow and Stark, 1990).

3.8 Summary and prospectus Beginning with the identification of plant cutin monomers and the investigation of their biosynthesis in the 1970s, our molecular-level understanding of these protective biopolymers has been augmented by recent spectroscopic studies of intact cutins and their oligomeric degradation products. With these latter investigations, which rely on specialized methods including FT-IR, solid-state NMR, HR-MAS NMR, and electrospray MS, the ester linkages typical of extended cutin chains and several types of cross-links in tomato and lime fruit cutins have been established. A more comprehensive picture of the cross-link elements and their prevalence within the biopolymer structure would deepen our understanding of how cutin functions as a support structure for the coverings of aerial plant surfaces. A complementary physical view of the cutin biopolymer and its embedded waxes is also being assembled using DSC, AFM, and NMR relaxation methods. Importantly, these assessments of bulk and surface cuticular mechanical behavior may be made under environmental stress conditions including hydration, abrasion and extreme temperatures. For instance, in tomato fruit cuticles it has been possible to correlate temperature-induced changes in surface elastic modulus with the phase behavior of the epicuticular waxes and to associate hydration-induced plasticization of dewaxed cutin with the flexibility identified in acyl chain segments of the cutin biopolymer. Such studies can lay the groundwork for future crop protection strategies based on molecular and micromechanical principles.

Acknowledgements We gratefully acknowledge support of this work by the US Department of Agriculture, National Research Initiative (00-35100-9462), and the National Science Foundation (Grants MCB/IBN-9728503 and MCB-0134705). Dr James Batteas provided experimental data and critical feedback for the AFM experiments, which were conducted by A. Round, S. Dang and J. Saccardo. Coworkers for the chemical and NMR experiments included J.R. Garbow, A.K. Ray, Z. Chen, B. Yan, X. Fang, X. Cheng and H. Wang. Dr C. Soll acquired the MS data at the Hunter College Mass Spectrometry Facility. S. Dang and J. Saccardo were supported by NSF Research Experiences for Undergraduates stipends and Deans’ Fellowships at The College of Staten Island.

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Kolattukudy, P.E. (2001) Polyesters in higher plants. Advances in Biochemical Engineering Biotechnology, 71, 1–49. Kolattukudy, P.E. and Espelie, K.E. (1989) Chemistry, biochemistry, and function of suberin and associated waxes, in Natural Products of Woody Plants (ed J.W. Rowe), Springer, Berlin, Heidelberg, New York, pp. 304–367. Lee, W.-K., Tanaka, K., Satomi, N. et al. (1997) Relationships between lateral force and viscoelastic properties for amorphous polymer films based on lateral force microscopy. Polymer Bulletin, 39, 369–376. Luque, P., Bruque, S. and Heredia, A., 1995. Water permeability of isolated cuticular membranes. Archives of Biochemistry and Biophysics, 317, 417–422. Luque, P. and Heredia, A. (1997) The glassy state in isolated cuticular membranes: differential scanning calorimetry of tomato fruit cuticular membranes. Plant Physiology Biochemistry, 35, 251–256. Matas, A.J., Curatero, J. and Heredia, A. (2004) Phase transitions in the biopolyester cutin isolated from tomato fruit cuticles. Thermochimica Acta, 409, 165–168. Mechaber, W.L., Marshall, D.B., Mechaber, R.A., Jobe, R.T. and Chew, F.S. (1996) Mapping leaf surface landscapes. Proceedings of the National Academy of Sciences, USA, 93, 4600–4603. Millis, K., Maas, W.E., Singer, S. and Cory, D.G. (1997) Gradient, high-resolution, magic-angle spinning nuclear magnetic resonance spectroscopy of human adipocyte tissue. Magnetic Resonance in Medicine, 38, 399–403. North, A.M. (1975) Molecular Behavior and the Development of Polymeric Materials. Wiley, New York. Oerke, E.-C., Dehne, H.-W., Schonbeck, F. and Weber, A. (1999) Crop Production and Crop Protection. Estimated Losses in Major Food and Cash Crops, Elsevier Science, Amsterdam. Osman, S.F., Gerard, H.C., Fett, W.F., Moreau, R.A. and Dudley, R.L. (1995) Method for the production and characterization of tomato cutin oligomers. Journal of Agricultural and Food Chemistry, 43, 2134–2137. Petracek, P.D. and Bukovac, M.J. (1995) Rheological properties of enzymatically isolated tomato (Lycopersicon esculentum Mill.) Fruit Cuticle. Plant Physiology, 109, 675–679. Pinot, F., Salaun, J.P., Bosch, H., Lesot, C., Mioskowski, C. and Durst, F. (1992) Omega-Hydroxylation of Z9-octadecenoic, Z9,10-epoxystearic and 9,10-dihyroxystearic acids by microsomal cytochrome P450 system from Vicia sativa, Biochem Biophys Res Commun, 184, 183–193. Ramirez, F.J., Luque, P., Heredia, A. and Bukovac, M.J. (1992) Fourier transform IR study of enzymatically isolated tomato fruit cuticular membrane. Biopolymers, 32, 1425–1429. Ray, A.K., Chen, Z. and Stark, R.E., 1998. Chemical depolymerization studies of the molecular architecture of lime fruit cuticle. Phytochemistry, 49, 65–70. Ray, A.K., Lin, Y.Y., Gerard, H.C. et al. (1995) Separation and identification of lime cutin monomers by high performance liquid chromatography and mass spectrometry. Phytochemistry, 38, 1361–1369. Ray, A.K. and Stark, R.E. (1998) Isolation and molecular structure of an oligomer produced enzymatically from the cuticle of lime fruit. Phytochemistry, 48, 1313–1320. Reina, J.J. and Heredia, A. (2001) Plant cutin biosynthesis: the involvement of a new acyltransferase. Trends in Plant Science, 6, 296. Riederer, M. and Schonherr, J. (1988) Development of plant cuticles: fine structure and cutin composition of Clivia miniata reg. leaves. Planta, 174, 127–138. Riley, R.G. and Kolattukudy, P.E. (1975) Evidence for covalently attached p-coumaric acid and ferulic acid in cutins and suberins. Plant Physiology, 56, 650–654. Rinaldi, P.L. and Keifer, P.A. (1994) The utility of pulsed-field-gradient HMBC for organic structure determination. Journal of Magnetic Resonance, A 108, 259–262. Round, A.N., Yan, B., Dang, S., Estephan, R., Stark, R.E. and Batteas, J.D. (2000) The influence of water on the nanomechanical behavior of the plant biopolyester cutin as studied by AFM and solid-state NMR. Biophysical Journal, 79, 2761–2767. Schaefer, J. and Stejskal, E.O. (1979) High-resolution 13 C NMR of solid polymers., in Topics in C-13 NMR Spectroscopy, 3 (ed. G.C. Levy), pp. 283–324.

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Schmidt-Rohr, K., Clauss, J. and Spiess, H.W. (1992) Correlation of structure, mobility, and morphological information in heterogeneous polymer materials by two-dimensional wideline-separation NMR spectroscopy. Macromolecules, 25, 3273–3277. Sieber, P., Schorderet, M., Ryser, U. et al. 2000. Transgenic Arabidopsis plants expressing a fungal cutinase show alterations in the structure and properties of the cuticle and postgenital organ fusions. Plant Cell, 12, 721–738. Smith, R.M., 1999. Understanding Mass Spectra: A Basic Approach. John Wiley and Sons, New York. Stark, R.E., Yan, B., Ray, A.K., Chen, Z., Fang, X. and Garbow, J.R. (2000) NMR studies of structure and dynamics in fruit cuticle polyesters. Solid State NMR, 16, 37–45. Tian, S. (2005) Molecular Structures of Natural Polymers: Cutin, Suberin, and Melanin. Ph.D. Dissertation, City University of New York. Torchia, D.A. (1978) The measurement of proton-enhanced carbon-13 T1 values by a method which suppresses artifacts. Journal of Magnetic Resonance, 30, 613–616. Villena, J.F., Dominguez, E., Stewart, D. and Heredia, A. (1999) Characterization and biosynthesis of non-degradable polymers in plant cuticles, Planta, 208, 181–187. Walton, T.J. (1990) Waxes, cutin and suberin. Methods in Plant Biochemistry, 4, 105–158. Zlotnik-Mazori, T. and Stark, R.E. (1988) Nuclear magnetic resonance studies of cutin, an insoluble plant polyester. Macromolecules, 21, 2412–2417.

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

4

Composition of plant cuticular waxes Reinhard Jetter, Ljerka Kunst and A. Lacey Samuels

Plant cuticles contain, beside cutin (Chapters 2 and 3), wax as the second major chemical component. Cuticular wax plays pivotal physiological and ecological roles in the interactions between plants and their abiotic (Chapters 5 and 8) and biotic environments (Chapters 10–12), respectively. Usually, the mixture of compounds that can be obtained by surface extraction of intact plant organs with organic solvents of low polarity is designated as ‘wax’. This is typically a mixture with characteristic composition for each species and organ. Thus, in this chapter, to distinguish between diverse mixtures originating from different sources, we will use the plural ‘waxes’, while the singular ‘wax’ will refer to the mixture from one particular source. It should be noted that our use of the term wax differs from the definition used in purely chemical contexts, where it is usually restricted to long-chain alkyl esters. A number of reviews have previously addressed the chemical composition of plant cuticular waxes. They mostly emphasised selected aspects, such as analytical procedures (Walton, 1990), qualitative wax composition (Tulloch, 1976), epicuticular compounds (Baker, 1982) or the critical assessment of experimental approaches (Riederer and Markstädter, 1996). Much new information on the composition and structure of plant cuticular waxes has accumulated over the last decade. Here we attempt to summarise our knowledge on the chemical structure of plant cuticular waxes, integrating aspects from all these previous reviews and the literature published since they appeared. To this end, we will briefly describe the experimental procedures used for wax analysis (Section 4.1), summarise the qualitative composition of waxes (Section 4.2) and review our knowledge on spontaneous reactions occurring within waxes (Section 4.3). Then the quantitative chemical composition of wax mixtures (Section 4.4) and their dynamic changes will be described (Section 4.5). Due to space constraints, this review of chemical structures and of quantitative compositions, as well as the corresponding references, cannot be comprehensive, but must instead be attempted by depicting overall patterns and using individual examples. Finally, we will summarise our current understanding of the arrangement of wax constituents in epicuticular and intracuticular layers within the cuticle (Section 4.6).

4.1 Methods used for the chemical analysis of plant cuticular waxes 4.1.1

Wax extraction

Plant cuticular waxes are typically prepared for chemical analysis by surface extraction of intact organs. Extraction has to be carried out under controlled conditions

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in order to give reproducible results. The choice of solvent, the solvent volume, its temperature and the duration of contact between the solvent and the plant surface have to be chosen carefully. In most cases, an exhaustive extraction of wax is desirable, as the total amounts present will be the best basis for comparisons, for example, between species, organs, developmental stages and tissues grown under varying conditions. Solvents of intermediate polarity should be used to maximise solubility of all wax constituents, including the extremely hydrophobic hydrocarbons and the much more polar compounds containing (multiple) functional groups (Riederer and Schneider, 1989; Stammitti et al., 1996). Chloroform proved to have ideal properties and gave particularly high, reproducible wax yields for various species (Holloway, 1984). In some studies, other solvents, for example, methylene chloride (Arrendale et al., 1988) or petrol ether (Salasoo, 1983), have been used. In all of these solvents, most wax compounds have solubilities in the range of 1–10 mg/ml, and hence the volume needed to cover the tissue entirely is generally sufficient to dissolve all the wax present. In a number of investigations, n-hexane has been used to extract cuticular waxes and has given satisfactory results (Rashotte et al., 2001). However, it should be noted that this solvent, due to its very low polarity, can dissolve only limited amounts of the more polar wax constituents, so that relatively large volumes have to be applied for exhaustive extraction. The extraction efficiency can be improved by increasing the extraction temperature. This is generally not necessary if suitable solvents are used, and therefore extraction is usually performed at room temperature. To the authors’ knowledge, only in one case where oligomeric and/or polymeric constituents (derived of aldehyde monomers) were present, was it necessary to extract with warm solvent to release the wax compounds (Riedel et al., 2003). The duration of extraction has varied widely in previous studies, in part to distinguish between wax exposed near the surface or embedded deep inside the cuticle (Silva Fernandes et al., 1964; Holloway, 1974; Baker et al., 1975; Baker and Procopiou, 1975). The time required for solvent molecules to enter into the cuticle, to mobilise wax and to wash it out must depend on the thickness and molecular structure of the cuticle, depending in turn on the plant species and organ investigated. Therefore, the minimum time necessary for exhaustive extraction would have to be determined prior to each investigation. For a large number of tissues, however, it was found that extraction is complete after 5–20 s (Holloway, 1984). Therefore, extraction is usually not optimised for new objects, but instead standard protocols comprising two extraction steps of 30 s are employed and assumed to yield total wax mixtures. It should be noted that for some plant species it was shown that, even though the superficial extraction appeared exhaustive, the cuticle contained more soluble wax components that could only be released by soxhlet extraction of isolated cuticular membranes (Riederer and Schneider, 1989). In many cases extraction of cuticular wax has been performed by immersion of entire organs into large volumes of solvent. While this method of preparation is adequate for unifacial tissues, it is by nature unable to distinguish between different

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sides of the same organ, for example, the adaxial and abaxial sides of leaves, which may have diverse compositions. Extraction of single tissue sides has been attempted by running solvent over the surface (Holloway et al., 1977b; Eigenbrode et al., 1998), by sweeping the tissue with solvent-saturated glass wool (Wen and Jetter, unpublished results) and by containing the solvent in glass cylinders pressed onto the surface that is to be extracted (Jetter et al., 2000). Only the last procedure allows control of extraction conditions, such as extraction time, solvent volume and temperature. All these methods must be validated by repeated extraction to prove exhaustive yields, and by comparison of single-side yields with the multi-side yields determined by dipping experiments. Special care has to be taken to avoid contamination of samples by internal lipids, possibly extracted by solvent entering through surface lesions or the cut areas created during harvest of the material. Extraction of internal lipids is usually indicated by the presence of chlorophyll, visible as green colour of the extract, and by phospholipids detected chemically. Extraction of fresh and healthy tissue in most cases yields wax preparations that are free of internal lipids. It has not been systematically studied whether the solvent might also enter through stomata, and therefore the extracts might contain soluble lipids originating from substomatal cavities or intercellular spaces. For comparisons of chemical data between studies, and correlations of wax composition with functional and structural parameters, the wax quantities have to be expressed in absolute values. Although amounts have sometimes been given in units of wax mass per tissue dry weight, it is more useful to quantify wax mass per surface area. This value corresponds to a layer thickness, hence allowing direct interpretation together with microscopic results and data on the transport of water and organic compounds across the cuticle. Wax quantities have to be determined by comparison with known amounts of an internal standard. An artificial compound that is similar to the most common wax constituents should be chosen as standard, and must be added to the sample shortly before or after extraction. Further sample processing is straightforward and little sample loss usually occurs before instrumental analysis. Sensitivity of analytical instrumentation has gradually improved by the introduction of gas chromatography (GC), capillary GC, and flame ionisation detectors (FID) or mass spectrometers (MS). Consequently, for most plant species and organs, extraction of 1–10 cm2 of surface area will yield sufficient wax for single analyses. In some instances, even the wax mixtures from areas smaller than 1 cm2 have been used for complete compound identification and quantification.

4.1.2

Instrumental analysis

Due to the complexity of mixtures and the relatively low amounts available, cuticular wax extracts can best be analysed by GC. To improve peak shape and thus give better resolution, functional groups containing active hydrogen should be derivatised. To this end, hydroxyl groups of alcohols and carboxylic acids are

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typically transformed into the corresponding trimethyl silyl (TMSi) ethers and esters, employing silylation reagents like bis-N ,O-trimethylsilyltrifluoroacetamide (BSTFA). It has been shown that this derivatisation reaction goes to completion, and that the resulting compounds can be very easily identified and accurately quantified (Deas et al., 1974; Little, 1999). Using these derivatives, compounds with different functional groups and/or varying carbon numbers have, in all cases, been separated by GC (Figure 4.1a). After GC separation, individual compounds in most studies have been quantified using FID detection. The FID has especially high sensitivity and broad range of proportionality. Selected aliphatic compounds are usually employed as internal standards for quantification. Unfortunately, the FID is destructive and does not yield chemical information; so separate GC–MS analyses have to be performed for compound identification. Currently, the spectra of only a few wax constituents are represented in the published electronic MS libraries. Authentic standards of many wax constituents are commercially available, or can easily be prepared in single-step semi-syntheses, and are important for structure confirmation by coelution experiments and spectral comparisons. Mass spectral fragmentation patterns are very distinct for most compound classes present in waxes, especially for TMSi derivatives (Figures 4.1b and d). Under normal conditions, electron impact MS also gives information on the molecular weight of the compound, thus allowing assignments of the chain lengths of aliphatic compounds. Therefore, a qualitative analysis of a cuticular wax mixture, including the assignment of all the previously known compounds, can usually be performed within a few hours. In many instances, isomers could also be baseline separated by GC. This is true for skeletal isomers of diverse triterpenoids and for positional isomers of alkanediols (Franich et al., 1979). Baseline separation cannot be achieved for isomeric alkyl esters, i.e. compounds with the same overall carbon numbers but with differences in the alcohol and fatty acid moieties (Sümmchen et al., 1995b). Also, secondary alcohols with subtle differences can typically not be separated, for example, the C29 isomers nonacosan-14-ol and nonacosan-15-ol in the stem wax of Arabidopsis thaliana (Figure 4.1c; Jenks et al., 1995). In contrast, the more pronounced geometrical differences between asymmetric alcohols like the C33 isomers tritriacontane-10-ol, -12-ol and -14-ol suffice for partial GC separation (Figure 4.1e). To circumvent problems caused by decomposition of aldehydes during derivatisation with BSTFA, an additional step in sample preparation has been employed in one study (Nass et al., 1998). In this protocol, solid phase extractions on short silica columns were used to separate wax mixtures into two fractions that were analysed independently. More sophisticated pre-separation procedures have also been employed to prepare larger quantities of individual compound classes for structure elucidation. These included one- and two-dimensional thin-layer chromatography (TLC) as well as (repeated) column chromatography (Tulloch, 1975; Holloway, 1984). While these techniques are of great value for qualitative investigations, special care has to be taken when employing them in quantitative analyses (e.g. the use of multiple internal standards).

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(a)

(b)

(c)

(d)

(e)

Figure 4.1 Gas chromatographic (GC) separation and mass spectrometric (MS) identification of plant cuticular wax components. (a) GC–FID trace of the wax mixture extracted from inflorescence stems of Arabidopsis thaliana ecotype Columbia. (b) Mixed mass spectrum of trimethyl silyl (TMSi) derivatives of nonacosan-14-ol and nonacosan-15-ol [corresponding to the GC peak at 23.5 min in (a)] showing characteristic α-fragments at m/z = 285, 299, 313. (c) GC–MS traces of the α-ions, demonstrating that the isomeric 14- and 15-alcohols cannot be separated under the GC conditions used. (d) Mixed mass spectrum of TMSi derivatives of tritriacontan-10-ol, tritriacontan-12-ol and tritriacontan-14-ol (from leaf wax of Myricaria germanica) showing characteristic α-fragments at m/z = 229, 257, 285 and m/z = 369, 397, 425, respectively. (e) GC–MS traces of the α-ions, demonstrating that the isomeric 10-, 12- and 14-alcohols can be partially separated under the GC conditions used. This figure is produced in colour in the colour plate section, which follows page 249.

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4.2 Chemical profiles of plant cuticular waxes 4.2.1

Ubiquitous constituents of cuticular waxes

A number of aliphatic compounds have been identified in the wax mixtures of virtually all the plant species investigated to date (Figure 4.2). This standard set of compounds is characterised by unbranched, fully saturated hydrocarbon backbones that may carry one primary oxygen-containing functionality, i.e. a terminal hydroxyl, carbonyl or carboxyl group. Therefore, plant cuticular waxes are typically mixtures of primary n-alcohols, n-aldehydes and fatty acids as well as n-alkanes, each of these compound classes comprising a homologous series of chemicals with chain lengths ranging from 20 to almost 40 carbons. A species-specific array of these compounds alone may thus comprise more than 50 different chemical structures. Additionally, esters of C16 –C34 fatty acids and C20 –C36 primary alcohols have been identified, giving rise to homologous compounds with chain lengths ranging from C36 up to C70 , and to dozens of isomers for each chain length (Gülz et al., 1994; Shepherd et al., 1995; Sümmchen et al., 1995a). In many instances the esters contain alkyl and acyl moieties with chain lengths similar to the free alcohols and acids present in the wax mixture (Purdy and Truter, 1963; Allebone and Hamilton,

Figure 4.2

Structures of components ubiquitously occurring in plant cuticular wax mixtures.

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1972). Consequently, these alkyl esters can be classified as dimeric products, while the corresponding alcohols and acids (as well as aldehydes and alkanes) represent monomeric wax constituents.

4.2.2

Taxon-specific major constituents of cuticular waxes

In addition to the ubiquitous constituents, the wax mixtures of individual plant taxa may contain relatively high percentages of specific compounds or compound classes. The majority of structures involved may be summarised as follows: the compounds mostly have unbranched, fully saturated aliphatic chains with 29 or 31 carbons and alcohol or keto groups (Table 4.1). In contrast to the standard wax constituents, the taxon-characteristic components frequently have secondary functional groups, giving rise to the possibility of positional isomerism. Some of the taxon-specific compounds also contain two or more secondary functional groups. Both the chain lengths and the positions of the functional groups can be easily determined by GC–MS, by interpreting the characteristic patterns of α-fragmentation (Ubik et al., 1975; Holloway et al., 1976). Secondary alcohols have been identified in a number of plant waxes, and can be classified into three groups (Table 4.1; Figure 4.3). The first is characterised by homologous series of secondary alcohols with a relatively broad chain length distribution and a vast array of positional isomers, containing hydroxyl groups typically between positions C-4 and C-12. Typical examples have been described for the cuticular waxes from strawberry leaves (Baker and Hunt, 1979), apple fruits (Verardo et al., 2003) and rose flowers (Mladenova and Stoianova-Ivanova, 1977). A second class of secondary wax alcohols is defined by the very strong predominance of a single chain length (in most cases C29 or C31 ) and the presence of one isomer that has the hydroxyl group at the central carbon of the chain. It is usually accompanied by a second isomer carrying the functionality on the carbon adjacent to centre. Accordingly, the leaf wax of Pisum sativum contains large amounts of the C31 secondary alcohols hentriacontan-16-ol and hentriacontan-15-ol, together with traces of homologous secondary alcohols (Macey and Barber, 1970; Kolattukudy, 1970; Holloway et al., 1977b). The waxes from leaves of Clarkia elegans (Hunt et al., 1976), Brassica oleracea (Baker, 1974) and Brassica napus (Holloway et al., 1977a), as well as from stems of A. thaliana (Rashotte et al., 1997) contain the C29 secondary alcohols nonacosan-15-ol and nonacosan-14-ol, together with small amounts of secondary alcohols with other chain lengths. Finally, a third class of secondary alcohols from plant cuticular waxes, again dominated either by C29 or by C31 homologues, is characterised by the position of the hydroxyl function exclusively on carbon-10 or -12. Nonacosan-10-ol is the predominant constituent of leaf waxes from Papaveraceae (Jetter and Riederer, 1996), Gymnospermae (Riederer, 1989), Rosaceae (Wollrab, 1969) and other diverse taxa of vascular plants (Holloway et al., 1976), as well as on the sporophytes of some moss species (Neinhuis and Jetter, 1995). Hentriacontan-12-ol is found at high concentration in the leaf wax of Myricaria germanica (Jetter, 2000). These asymmetric

C31 C29

C31 C29 C29 C29 C31 C29

C29 C31

C31 C31

C29 –C33 C27 –C31

C29 –C33 C27 –C31 C27 –C31 C27 –C31 C29 –C33 C27 –C31

C27 –C31 C27 –C33

C29 –C35 C27 –C33

C27 –C33

β-Diketones

Hydroxy-β-diketones

Oxo-β-diketones

sec.|sec. Ketols

sec.|sec. Alkanediols

C31

C29

C27 –C31

Ketones

C31

C29 –C33

Dominating chain length

C26 –C30 C29

Range of chain lengths

Homologous series

C21 –C35 C27 –C31

sec. Alcohols

Compound class

C-14,16

C-8,10 C-14,16

C-10 C-14,16

C-16 C-10 C-14 C-10 C-12 C-14

C-12 C-15

C-10

C-15 and C-16

C-4 to C-12 C-14 and C-15

Position of 1st functional group

— C-4 to C-9 or C-25 C-4 to C-9 or C-25

C-4 or C-5 —

— — C-15 C-3 to C-16 C-4 to C-16 C-15

— —

—

—

— —

Position of 2nd functional group

Isomer mixture

Poaceae

Gymnospermae Poaceae Eucalyptus sp. Buxus sempervirens Poaceae

Gymnospermae Papaveraceae Myricaria germanica Arabidopsis thaliana Brassica oleracea Aristolochia gigantea Osmunda regalis Brassica oleracea Nelumbo nucifera Myricaria germanica Brassica oleracea

Rosa damascena Arabidopsis thaliana Brassica oleracea Pisum sativum

Plant taxon

Leaves

Needles Leaves Leaves Leaves Leaves

Needles Leaves Leaves Inflor, stems Leaves Leaves Fronds Leaves Leaves Leaves Leaves

Refrences

Tulloch et al., 1980

Mladenova and Stoianova-Ivanova, 1977 Rashotte et al., 1977 Baker, 1974 Hollway et al., 1977b Macey and Barber, 1970 Kolattukudy, 1970 Riederer, 1989 Jetter and Riederer, 1996 Jetter, 2000 Rashotte et al., 1977 Baker,1974 Meusel et al., 1999 Jetter and Riederer, 2000 Holloway and Brown, 1977 Barthlott et al., 1996 Jetter, 2000 Holloway and Brown, 1977 Holloway et al., 1977a Riederer, 1989 Tulloch, 1981 Horn et al., 1964 Dierickx, 1973 Tulloch et al., 1980

Examples

Petals Inflor, stems Leaves Leaves

Organ

Table 4.1 Typical composition of prominent compound classes with secondary functional groups in plant cuticular waxes

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Figure 4.3

Examples of compounds with secondary functional groups, accumulating to high concentrations in the wax mixtures of certain plant taxa.

5

COMPOSITION OF PLANT CUTICULAR WAXES

153

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alcohols contain a stereogenic centre, and an excess of the S enantiomer has been reported for nonacosan-10-ol from Picea pungens (Jetter and Riederer, 1994). Wax secondary alcohols of the second and third categories described earlier are frequently accompanied by the corresponding ketones, at widely varying concentrations. In leaf wax of C. elegans (Hunt et al., 1976), B. oleracea (Baker, 1974) and B. napus (Holloway et al., 1977a), as well as on inflorescence stems of A. thaliana (Rashotte et al., 1997) the C29 ketone nonacosan-15-one is present in relatively high concentrations. The symmetric C31 ketone hentriacontan-16-one (=palmitone), but not the corresponding symmetric secondary alcohol hentriacontan-16-ol, has been reported in the leaf waxes of Allium porrum (Maier and Post-Beittenmiller, 1998) and diverse dicots including Aristolochia gigantea and Paeonia officinalis (Meusel et al., 1999). It is noteworthy that Asparagus officinalis leaf wax has been found to contain the C27 ketone heptacosan-14-one, a symmetric ketone with relatively short chain length (Scora et al., 1986), while Centhrantus ruber wax contains nonacosan14-one (Meusel et al., 1999). Finally, corresponding to the class of asymmetric secondary alcohols with high regio- and chain-length specificity, nonacosan-10-one has been described as a constituent of the wax on leaves of Encephalartos spp. (Osborne and Stevens, 1996), as well as on fronds of Osmunda regalis (Jetter and Riederer, 2000). Also associated with secondary alcohols, alkanediols and ketols have been described as cuticular wax constituents. They are characterised by a strong predominance of the C29 homologue, and can be classified into two groups according to the position of functionalities. In those diols and ketols typically detected together with corresponding symmetric secondary alcohols, one functionality is located on the central carbon and the second on the adjacent carbon. Thus, nonacosane-14,15-diol, 15-hydroxynonacosan-14-one and 14-hydroxynonacosan-15-one have been identified in leaf wax of Brassica species (Holloway and Brown, 1977; Holloway et al., 1977a). A second group of diols and ketols, associated with the corresponding asymmetric alcohols, has one functional group on carbon number 10 and a second group on one of the carbons between C-3 and C-16. Respective nonacosanediol isomers have been described for wax mixtures from diverse Gymnospermae (Franich et al., 1979), Papaveraceae (Jetter and Riederer, 1996), and are especially abundant in the wax of Nelumbo nucifera (Barthlott et al., 1996). In some of these species, 10-hydroxynonacosan-4-one and 10-hydroxynonacosan-5-one have been reported together with the diols. It is noteworthy that all the ketones, ketols and diols thus identified as important components of cuticular waxes showed a chain length or isomer pattern closely matching those of the secondary alcohols present either in the same species or in other taxa. Based on these purely chemical observations it has been suggested that the secondary alcohols are biosynthesised first and that the other compounds are derivatives formed by ensuing pathway steps (Franich et al., 1979; Jetter and Riederer, 1996). In the leaf waxes of numerous species of the Poaceae (Tulloch, 1981) and of the genus Eucalyptus (Horn et al., 1964), large percentages of aliphatic β-diketones have

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been described. This compound class is characterised by unbranched hydrocarbon chains with 27–33 carbons and two mid-chain carbonyl groups in 1,3-position to each other. The β-diketones usually show strong predominance of single homologues and isomers in the wax of a given species, but can vary substantially between taxa. The most widely distributed representative, the C31 compound hentriacontane-14,16-dione, is the dominant compound in the waxes of various organs of well-investigated species such as Hordeum vulgare (Jackson, 1971; von Wettstein-Knowles and Netting, 1976a) and Triticum aestivum (Bianchi et al., 1979). In contrast, leaf wax of Buxus sempervirens was reported to contain mainly the isomeric hentriacontane-8,10-dione (Dierickx, 1973). In many cases further derivatives of β-diketones have been described, including hydroxy- or keto-β-diketones with an additional alcohol or carbonyl function on carbons C-4 to C-9 or C-25 (Tulloch et al., 1980). These derivatives occur as species-specific mixtures including various isomers that all share the same basic chain length and position of the β-diketo functionality. Thus, similar to the relation between secondary alcohols and alkanediols and ketols, the chemical structures suggest that β-diketones are the parent compounds from which further derivatives are biosynthesised through hydroxylation and hydroxyl oxidation.

4.2.3

Cuticular triterpenoids

In the cuticular wax mixtures of many plant species, triterpenoid constituents have been detected. While they occur at trace levels in most cases, including model species such as A. thaliana (Jenks et al., 1995) and B. oleracea (Baker and Holloway, 1975), they can accumulate to very high concentration in the waxes of specific taxa. Cuticular triterpenoids can be classified according to the carbon backbone structure, according to the number, nature, and position of functional groups and according to the derivatives formed through bonding between the triterpenoid and other compounds. Out of the more than 200 basic triterpenoid carbon skeletons described to date (Xu et al., 2004), only a small number has been detected in plant cuticular waxes (Figure 4.4). The large majority of these cuticular constituents are pentacyclic triterpenoids, characterised by the presence of five condensed carbon rings. Triterpenoids with fernane structures and related carbon backbones have been detected mostly on the surfaces of fern fronds (Tanaka et al., 1992). In the cuticular wax mixtures of seed plants, lupane, oleanane and ursane derivatives have been encountered most frequently (Walton, 1990). For example, in wax from Prunus laurocerasus leaves, oleanane and ursane structures co-occur as major components (Jetter et al., 2000), while oleananes predominate in waxes on grapes (Vitis vinifera; Radler, 1970), plums (Prunus domestica; Ismail et al., 1977) and olives (Olea europaea; Bianchi et al., 1992), on shoots of Euphorbia lathyris (Hemmers et al., 1989b), and on leaves of Tilia tomentosa (Gülz et al., 1988). Reports on single species from other taxa documented diverse variations of this standard set of triterpenoids. For instance, in wax of tomato fruits (Lycopersicon esculentum),

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Figure 4.4

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Examples for triterpenoids occurring in plant cuticular waxes.

oleanane and ursane structures have been found associated with equivalent amounts of δ-amyrin (Vogg et al., 2004). In relatively few taxa, including Cirsium arvense (Tulloch and Hoffman, 1982) and Ricinus communis (Guhling and Jetter, unpublished results), ursane or lupenane derivatives predominate, respectively. In other groups of plants, wax mixtures dominated by other triterpenoid skeletons including taraxerane in Macaranga spp. (Markstädter et al., 2000) and glutinane in Euphorbia cyparissias (Hemmers et al., 1989a) have been described. The large majority of the triterpenoids identified in cuticular waxes of various plant species to date contain a hydroxyl function in the 3β position. This finding can be explained with the generally accepted concepts for triterpenoid biosynthesis (Chapter 5), according to which 3β-alcohols are the immediate products of the first steps of the pathways, and are hence ubiquitously present in triterpenoids of higher plants. In many cases, the 3β-triterpenols are accompanied by the corresponding ketones, sometimes accumulating to relatively high percentages. For example, in stem cuticular waxes of several Macaranga species taraxerone was found (Markstädter et al., 2000), while glutinone was one of the prominent compounds of Euphorbia cyparissias wax (Hemmers et al., 1989a) and friedelin was reported for grapefruit (Citrus paradisi) waxes (Nordby and McDonald, 1994). Whereas thousands of triterpenoid structures with different numbers of functional groups on the carbon backbone have been identified from internal plant tissues, only very few of them have so far been detected in cuticular waxes. For the lupenane, oleanane and ursane structures, some of the C-28 oxidation products have been reported, including the diols erythrodiol, uvaol (Griffiths et al., 2000) and betulin (Wollenweber et al., 1999) as well as the hydroxyacids oleanolic and ursolic acids

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(Jetter et al., 2000). It may be speculated that other oxygen-containing triterpenoid derivatives might not be present in cuticular waxes because they are too polar to partition into such a lipophilic mixture. Finally, various derivatives of the triterpenoids described above have been identified in cuticular waxes. These include alkyl ethers and acyl esters of the triterpenoid alcohols as well as alkyl esters of the triterpenoid acids. Hence, the triterpenols lupeol, β- and α-amyrin tend to be associated with the corresponding methyl ethers (Smith and Martin-Smith, 1978), acetate esters (Manheim and Mulroy, 1978) or long-chain (C16 and C18 ) as well as very long-chain (>C20 ) fatty acid esters (Gülz et al., 1988). Ursolic and oleanolic acid have been found accompanied by their methyl esters (Bakker et al., 1998). While these examples demonstrate some emerging patterns in the distribution of triterpenoid structures in cuticular wax mixtures, our knowledge is still very sketchy. It relies on relatively few analyses of species selected with a strong bias towards certain plant taxa, and was performed by only a few research groups. The identification of cuticular triterpenoids is largely based on GC–MS data used to separate and delineate isomers with relatively subtle differences in structure. Fragmentation patterns and MS peak intensities are less predictable than for aliphatic wax constituents. Hence, it is very important that triterpenoid structures, after MS interpretation, be confirmed by comparison with authentic standards. Unfortunately, only a few triterpenoids, representing little diversity of carbon backbones, are commercially available or can easily be prepared and authenticated from plant material. This hampers structure identification of novel cuticular triterpenoids, creates a bias towards repeated description of known structures, and might thus give the misleading impression of only a few structures being present in the cuticular waxes of diverse plant taxa. Our chemical knowledge of cuticular triterpenoid diversity and distribution, bearing important biological implications on cuticle structure and function, will likely change substantially as more species are investigated and more standards for identification become available.

4.2.4

Other minor constituents of cuticular waxes

Beside the compounds described earlier, a large number of wax constituents have been identified over the past decades, and the following description can only highlight a few examples for specialty components of plant cuticular waxes. Some structural diversity arises from modifications in the hydrocarbon chains of aliphatic wax constituents. In a few cases, a double bond is present, giving rise to unsaturated compounds that are analogous to the predominating saturated wax constituents. Double bonds with Z-configuration seem to predominate, and they have frequently been located between carbon numbers 2 and 3 or 9 and 10. As a second modification, methyl branches have been detected on the hydrocarbon chain of wax alkanes, primary alcohols, fatty acids and their alkyl esters. Typically, homologous series of both iso- and anteiso-isomers are present, that is, compounds with methyl branches located either one or two carbons away from the alkyl chain terminus, respectively.

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One example for the relatively prominent presence and wide arrays of branched structures is the leaf wax of Brassica species, where primary alcohols and fatty acids with iso- and anteiso-branching have been reported as free compounds and in the corresponding alkyl esters (Baker and Holloway, 1975). Branched hydrocarbons have been identified in petal waxes of various species, including Antirrhinum majus (Goodwin et al., 2002) and Papaver rhoeas (Stransky and Streibl, 1969). Aliphatic acids may be found esterified with methanol, resulting in series of homologous fatty acid methyl esters with acyl chain lengths from C20 to C34 . In a similar way, C20 –C30 wax primary alcohols are found esterified with acetic acid in some cases. Another class of trace compounds worth mentioning are esters formed between very long chain fatty acids and secondary alcohols. In the wax mixtures of some plant species that were otherwise characterised by the presence of β-diketones, such esters were formed by medium-chain alkan-2-ols. Their composition and distribution in the waxes of various plant organs has been intensively studied for Hordeum vulgare (von Wettstein-Knowles and Netting, 1976b; Mikkelsen, 1984; von Wettstein-Knowles and Madsen, 1984) (Chapter 5). Relatively recently, a vast range of aliphatics with novel structures containing two functional groups has been identified. They are generally characterised by C22 –C32 hydrocarbon chains, the presence of a primary oxygen-containing functional group on one chain terminus and of a second oxygen-functionality at specific distances inside the chain. Variations of these primary and secondary functional groups give rise to homologous series of alkanediols in the wax of Papaver species (Jetter et al., 1996b), Osmunda regalis (Jetter and Riederer, 1999b), M. germanica (Jetter, 2000) and R. communis (Vermeer et al., 2003), together with hydroxyaldehydes for R. communis (Vermeer et al., 2003) and ketoalcohols for O. regalis (Jetter and Riederer, 1999b). Corresponding hydroxy fatty acids were found in the form of δ-lactones on the leaves of Cerinthe minor (Jetter and Riederer, 1999a). Most remarkably, the substitution was in all cases on odd-numbered carbons, resulting in bifunctional compounds of 1,3-, 1,5-, 1,7- and so on. It was suggested that this geometry of functional groups might arise from polyketide-type elongation, explaining the mid-chain functionality as a product of modified fatty acid elongation cycles. Various reports have described cuticular wax constituents that combine structural features of the standard aliphatic wax structures with those of other plant metabolites. Most notably, the latter may contain aromatic ring structures mainly originating from the phenyl propanoid metabolism. For example, mixed aliphatic–aromatic esters may be formed by wax fatty acids and cinnamyl alcohol in wax of Vicia faba flowers (Griffiths et al., 1999), as well as hydroxyphenyl propanol, and hydroxyphenyl butanol in Taxus baccata (Jetter et al., 2002). Other aromatic compounds have been detected in plant cuticular waxes, including homologous series of very long-chain fatty acids esterified with benzyl alcohol or phenylethyl alcohol (Gülz and Marner, 1986), as well as of alkyl benzoates (Gülz et al., 1987). Finally, series of 5-alkyl resorcinols accumulate to relatively high amounts in the wax on seeds of H. vulgare (Garcia et al., 1997).

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The waxes of various gymnosperms, but also of a few other plant species, contain two classes of characteristic compounds. The first of them, called estolides, are oligomeric structures formed by ester linkages between two or more hydroxy fatty acids, α,ω-diols, as well as primary and secondary alcohols (Tulloch and Bergter, 1981). The second class of compounds, frequently associated with estolides, are characteristic triglycerides containing two normal fatty acyl moieties esterified to the glycerol positions C-1 and C-3, while having a hexanoic or octanoic acyl moiety bonded to C-2 (Tulloch and Hoffman, 1982). Finally, it should be noted that in a number of plant species, tocopherols have been reported as wax constituents (Gülz et al., 1992). The three most abundant representatives of these compounds in internal lipids, α-, γ- and ε-tocopherol, are usually occurring together in the plant cuticle (Griffiths et al., 2000).

4.2.5

Other compounds located at or near the plant surface

In surface extracts of many plants, a number of phytosterols, in most cases including β-sitosterol and stigmasterol, have been identified (Holloway, 1971). These compounds have, for example, been reported in leaf waxes of Poaceae species (Avato et al., 1990). Based on their overall polarity, but also based on structure similarities with the pentacyclic triterpenoids commonly found in cuticular waxes, it seems plausible that the phytosterols might be actively exported towards the cuticle and/or passively partitioned into the cuticular waxes. On the other hand, it should be noted that the phytosterols reportedly accumulate in specific membrane compartments such as plasma membranes (Borner et al., 2005). This must lead to relatively concentrated pools of internal phytosterols, which might cause substantial contamination of surface extracts when solvent molecules accidentally reach inner parts of the tissue even if only locally. Consequently, it cannot be judged at present whether phytosterols are in fact present in cuticular wax mixtures, and if so what their cuticular concentrations are. Similarly, other compounds like alkaloids and free C16 and C18 fatty acids might be extraction artefacts. To date, the alkaloids detected in surface extracts of fruit capsules of diverse Papaver species are the only nitrogen-containing cuticle constituents (Jetter and Riederer, 1996). It has been pointed out that for each poppy species only those alkaloids carrying methoxy groups instead of hydroxyls, that is, with the lowest polarities, were found in the extracts. Even though this observation suggests passive partitioning of alkaloids into the cuticular wax, direct proof for their presence in the cuticle is missing. The same holds true for palmitic and stearic acids, abundant fatty acids in all tissues that have repeatedly been reported in plant surface extracts (Walton, 1990). Even though it seems plausible that these compounds accumulate in cuticular waxes, their localisation in or on the cuticle needs to be confirmed by in situ investigations. Finally, in some special cases, compounds other than waxes may be present at the plant surface. The diterpenoids exuded by glandular trichomes on leaves of Nicotiana species may serve as one example (Severson et al., 1984). Even though

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these compounds are produced locally, they spread across the entire leaf surface and cover it with a fairly thick, homogenous film, probably consisting of polymerised oxidation products. As a second example, the surface of some fern fronds (Wollenweber, 1982; Wollenweber et al., 1993) and of diverse angiosperm leaves, especially in the Primulaceae and Lamiaceae (Wollenweber and Dietz, 1981), are dusted over by a thick farinose layer of flavonoids, deposited as relatively large particles. Due to their comparatively high polarity and the molecular structure that is clearly differing from typical wax compounds, it seems unlikely that these flavonoids are exported together with wax.

4.3 Spontaneous reactions of cuticular wax constituents Large portions of the wax mixtures consist of hydrocarbon chains, constituting the alkyl moieties of the diverse compound classes described earlier. As the reaction repertoire of these parts of the cuticular waxes is very limited, virtually no spontaneous chemical changes are to be expected on the hydrocarbon chains. On the other hand, the functional groups of the wax constituents might engage in spontaneous reactions within the cuticle, and could thus lead to gradual modifications of the wax composition. Based on purely chemical arguments, a number of reactions seem feasible for wax constituents. These include ester bond formation or cleavage, photodimerisation of compounds with C=C double bonds, hemiacetal and acetal formation, aldol and Claisen condensations, as well as redox transformations involving alcohols, aldehydes/ketones and fatty acids. As plant cuticles are in some cases exposed to extreme climatic conditions for long periods of time during the lifetime of individual tissues, speculations on such chemical reactions have repeatedly been put forward. It is a well-documented fact that plant surfaces frequently experience relatively high temperatures, intense irradiation of (partially high energy) light and substantial concentrations of singlet oxygen as an oxidizing agent (Jones, 1992). On the other hand, various other parameters necessary to describe the chemical environment, for example, the concentration of water and other reagents, or the presence of catalysts like metal ions, acids or bases have not been monitored accurately to date. It is therefore impossible to predict whether spontaneous reactions of cuticular waxes are likely to occur. From a practical point of view, spontaneous wax transformations seem to be rare. In the numerous wax analyses performed to date, systematic changes in wax composition as a function of time have scarcely been reported. Instead, fairly constant amounts of waxes and their constituents were found when the same species and organ was repeatedly investigated, in many cases using mature tissue of likely widely varying age. This is especially true for model species used in cuticle research, such as T. aestivum, H. vulgare, and A. thaliana. It has to be added, though, that due to the complex mixtures and substantial biological variability in their quantitative

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composition (see Section 4.4), subtle changes caused by chemical reactions would likely have been missed in these studies. Self-reaction products of very long-chain aldehydes have been reported for the wax mixtures of Nepenthes alata (Riedel et al., 2003), Oryza sativa and Saccharum officinarum (Haas et al., 2001), and flower wax of some decorative roses (Mladenova et al., 1976). Both indirect evidence employing Fourier Transform Infrared (FTIR) spectroscopy and direct chromatographic evidence showed that these wax aldehydes are present as oligomeric or polymeric acetal forms. Although the corresponding monomers are readily released in hot solvent, it has not been possible to date to transform them back into acetals (Riedel et al., 2003). This result suggests that special conditions exist during formation, export or accumulation of wax in the plant cuticle that allow reaction between aldehyde molecules. One key prerequisite for this reaction might be that adjacent molecules in the wax mixture are sufficiently aligned to bring the carbonyl groups into close contact. To test the hypothesis of (possibly subtle) spontaneous reactions between wax components and the atmosphere, either analyses closely monitoring wax composition over time or artificial reaction set-ups mimicking the natural situation have to be performed. The first approach has been taken in only a few studies. As an example, the wax on upper surfaces of P. laurocerasus leaves was found to change dramatically during organ development (Jetter and Schäffer, 2001). To distinguish between ontogenetic changes and possible chemical reactions, in this case, the second approach was also used. The wax mixture was removed from young leaves without altering the molecular arrangement, deposited on a glass surface and exposed to weathering for two weeks. While the native plant surface experienced dramatic modifications in this period of time, the artificial set-up remained unchanged. In two other investigations, artificial set-ups were also used to test specific reactions of individual wax compounds. In the first study, asymmetric secondary alcohols, being the major constituents in the (epi-)cuticular wax of gymnosperm needles, were exposed to various concentrations of air pollutants such as ozone, NO2 and SO2 (Jetter et al., 1996a). Even though these gases are strong oxidizing agents, implied in direct oxidative damage to the plant surface, they did not react with the wax alcohols under ambient conditions. Only when applied in more than 1000-fold higher doses, could the expected oxidation products be detected (ketones and carboxylic acids generated by oxidative α-bond cleavage). In the second study, possible UV-induced photoreactions of cuticular phenylpropanoids were monitored using homologous series of cis- and trans-coumarates from beech leaf wax (Riederer and Markstädter, personal communication). The expected 2+2 cyclisation products were detected in the resulting mixture in very small concentrations, but only upon irradiation with high doses of UV radiation. In summary, all the experimental evidence currently available argues against the hypothesis that spontaneous reactions occur within the bulk of cuticular wax mixtures at any substantial scale. In a TOF-SIMS (Time-Of-Flight Secondary Ion Mass Spectrometry) analysis of the top layer of molecules on P. laurocerasus leaves, some signals could not be explained with the known epicuticular wax constituents (Perkins et al., 2004).

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This raised the possibility that spontaneous transformations might be occurring in a very thin layer near the cuticle surface. Reactions thus restricted to the outermost surface would be of great biological importance, but have yet to be confirmed in more detailed studies.

4.4 Quantitative composition of cuticular waxes 4.4.1

Chain length distributions within compound classes

Most of the wax constituents described in the previous sections contain an alkyl moiety with one or more functional groups. Even in early chemical analyses of plant cuticular waxes, it was noticed that compounds within one compound class, for example, the primary alcohols and fatty acids, comprise complete series of compounds differing only in the number of methylene units (Chibnall et al., 1931; Kreger, 1958; Juniper, 1959). These homologous series had characteristic patterns in the relative amounts of constituents. The compositional patterns have two aspects, relating to the ratio between structures with odd and even carbon numbers, on one hand, and to the overall chain length distribution on the other. Similar patterns have been found in the waxes of all plant species investigated to date and for all alkyl-containing wax compound classes alike (Baker, 1982). In all the homologous series of wax constituents, a strong predominance of compounds with either odd or even carbon numbers has been found. Compound classes with preference for even carbon numbers are fatty acids, aldehydes and primary alcohols. Similarly, the acyl moiety of diverse esters (e.g. methyl, phenyl propyl, benzyl esters and alkyl esters) as well as the alkyl moiety of diverse other esters (e.g. alkyl acetates) is largely dominated by even-numbered homologues (for references see Section 4.2). Compound classes with preference for odd carbon numbers include alkanes, secondary alcohols and ketones as well as β-diketones and their hydroxy- and keto-derivatives. In summary, compound classes with a terminal functional group have a predominance of even-numbered homologues, whereas those lacking the primary functionality tend to have odd-numbered chains. This chemical classification likely reflects the biosynthetic relationships between wax compound classes, grouping at least fatty acids and primary alcohols along one pathway (Chapter 5). Loss of the terminal functional group together with its carbon atom leads to alkanes, secondary alcohols and ketones along another pathway. Interestingly, the same patterns are found for multifunctional compound classes. For example, alkanediols and ketols were reported to have either even or odd carbon numbers depending on the position of their functional groups. Where both functionalities are located on mid-chain carbons, diols and ketols are largely dominated by odd-chain compounds (Holloway and Brown, 1977; Franich et al., 1979). In contrast, diol and ketol series with one primary and one secondary functional group show a strong predominance of homologues with even carbon numbers (Jetter et al., 1996b; Vermeer et al., 2003).

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The overall chain length distribution of wax compound classes is best summarised giving the range of chain lengths, the most abundant chain length and its relative portion in the class. The characteristics of homologous series in cuticular waxes can vary widely, to the extreme with one compound largely predominating and several other homologues being present only in trace amounts. As an example for this situation, the wax on inflorescence stems of A. thaliana shows a strong predominance of C29 homologues of alkanes, secondary alcohols and ketones (Rashotte et al., 1997). In other cases, relatively even distributions of five and more chain lengths have been reported. For example, the seed coat wax of Fagus sylvatica contains an especially wide array of alkane, primary alcohol and fatty acid homologues, each of them accounting for less than 35% of the fractions, respectively (Gülz et al., 1989). In many plant species, the chain length distribution was found to match between compound classes, both in their chain length range and in relative homologue contributions. Consequently, the overall wax composition is in many cases dominated by a single chain length. Most frequently, wax mixtures with a pronounced preference for C26 , C28 , C29 , C30 or C32 compounds have been encountered. The homologue pattern within compound classes and the resulting chain length profile of the overall wax mixture play important roles in the models for the physical structure of the cuticle. Two hypotheses have been put forward that describe the crystalline arrangement of wax molecules, either inside the cuticle or on its surface. The first hypothesis predicts that the chain length distribution of aliphatics influences the size and geometry of crystalline domains in the wax, and consequently limits the water flow across the cuticle (Riederer and Schreiber, 1995; Chapter 6–8). The primary physiological function of the cuticle is thus linked to the chain length distribution in the homologous series of wax constituents. The second hypothesis proposes that the accumulation of individual compounds near the cuticle surface (Baker, 1982), triggered by the strong predominance of a single compound class and homologue, causes the formation of epicuticular wax crystals (see Section 4.6.1). The ecological functions associated with these surface crystals (Chapters 10–12) are therefore also linked to the chain length distribution in the wax. In conclusion, the homologue pattern is a central parameter in our current understanding of the causal relationship between the chemical composition, the physical properties and the biological functions of the cuticle.

4.4.2

Distribution of compound classes

To further summarise wax chemical datasets beyond homologue distributions, the relative amounts of compound classes within the wax mixture are usually given as percentages. To this end, all the known compounds, as described in Section 4.2, can be categorised according to their functional groups. The wax constituents that remain unidentified should also be quantified and given as a separate class of compounds to complete the analysis. For most plant species, more than 80% of the mass of compounds within the cuticular wax mixture have been identified.

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Relatively broad ranges of compound classes have been reported for cuticular waxes from some species, with no class predominating. For example, the wax on leaves of Zea mays was found to contain 9–42% each of alkanes, aldehydes, primary alcohols, fatty acids and alkyl esters (Bianchi et al., 1984). It should be noted that waxes containing such a broad range of compounds, all at relatively small percentages, are exclusively found on tissues that do not have epicuticular wax crystals on their surface. In contrast, the wax mixtures from many other plant species were reported to contain high percentages of a single compound class. This situation may be illustrated by the composition of H. vulgare leaf wax, as it was found to contain 89% of primary alcohols together with only 0.2–9.2% of alkanes, aldehydes, fatty acids and alkyl esters (Giese, 1975). Interestingly, the leaf surfaces of this species are covered with epicuticular wax crystals, leading to the assumption that the high percentage of alcohols causes the formation of the surface structures. Our current knowledge relies on analyses of relatively few plant species, and a description of compound classes grouped by plant taxa must necessarily be biased. Only a few selected examples may therefore suffice to illustrate the specific predominance of individual compound classes in diverse plant groups. Alkanes are the major constituents on leaves of Cactaceae [e.g. Opuntia engelmannii (Wilkinson and Mayeux, 1990)], and Brassicaceae [e.g. B. oleracea (Baker, 1974)], and in the abaxial leaf wax of P. sativum (Holloway et al., 1977b). Asymmetric secondary alcohols dominate the wax composition of all gymnosperm needles (Riederer, 1989), of Papaveraceae leaves and fruit capsules (Jetter and Riederer, 1996) and of diverse Liliaceae, Ranunculaceae and Rosaceae (Holloway et al., 1976). Triterpenoids tend to accumulate in the wax mixtures of a range of taxa, including Euphorbiaceae, Crassulaceae and Rosaceae (Walton, 1990). Diverse species of Fabaceae have leaf waxes with high percentages of primary alcohols (Baker, 1982), Eucalyptus leaf waxes are dominated by β-diketones (Horn et al., 1964) and many Poaceae species have large quantities of either of these compound classes on various organ surfaces (Baum and Tulloch, 1982). The particularly well-studied grass species in the genera Triticum, Hordeum and Secale may thus serve as fine examples to illustrate the complexity of cuticular wax compositions, varying widely between surfaces of awns, fruits, stems, leaf sheaths and blades, and even depending on the position and side of the leaf. Finally, it should be noted that some compound classes, even though they are ubiquitous constituents of waxes, including the free fatty acids and aldehydes, have rarely been found in high percentages. Analyses of single compound classes that lack information on the relative contribution of these compounds to the overall wax mixture have frequently been published. As the percentages of individual compound classes can vary widely, it cannot be determined whether these reports deal with major wax constituents or only with trace compounds. The resulting data therefore cannot contribute to our knowledge on cuticular wax composition, even though substantial effort was invested on these analyses of various interesting species.

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Wax amounts

On a third level of the chemical datasets, absolute quantities of cuticular wax loads have to be considered. They are usually given in units of micrograms per square centimetre referring to the mass of wax per area of cuticle surface. Corresponding values have the advantage that they can be directly compared between species and organs. It has been shown that solvent molecules can enter deep into the cuticle within short times of immersion (Jetter et al., 2000). Consequently, both epi- and intracuticular waxes are likely released, and the resulting mixtures must contain the total waxes rather than only the epicuticular portion. Even though many previous reports stated loads of epicuticular waxes, they should now be re-interpreted as giving the loads of total cuticular waxes instead. Depending on the plant species and organ analysed, wax loads range from one to several hundred micrograms per square centimetre of surface area. In many cases, fruit cuticles have been found to contain higher amounts of waxes than the corresponding leaf waxes of the same species. For example, tomato fruits showed a wax coverage of 15 μg/cm2 , while leaves had only 3 μg/cm2 of total cuticular wax (Vogg et al., 2004). Numerous analyses of cuticular waxes also revealed that tissues possessing epicuticular wax crystals tend to have especially high wax loads. In many of these cases, 50–100 μg/cm2 of wax have been reported, and it must be concluded that a large portion of the wax mass is located in the surface crystals of these species. Implicitly, values for the wax loads also give information on the thickness of the wax layer. This interpretation is based on the fact that all major wax constituents consist mainly of methylene units, causing fairly constant densities of approximately 0.8–1.0 g/cm3 for the different compounds as well as the resulting mixture. A load of 1 μg/cm2 of pure wax would consequently correspond to a layer thickness of 10 nm. However, this estimate is directly applicable only to epicuticular wax films, as these are the only compartments containing pure wax. For species and on organs where a large portion of the wax is localised in the epicuticular wax film, the comparison of wax loads gives a direct comparison of layer thickness. For all other tissues, wax loads have to be interpreted in perspective with the amounts of cutin present. For a number of species, both wax and cutin amounts were quantified, and cutin was found to make up 40–80% of the cuticle mass (Schreiber and Riederer, 1996).

4.5 Dynamics of wax composition In the previous sections of this chapter, the composition and quantity of plant cuticular waxes has been described as a fairly constant character of given plant species and tissues. This static view seems overly simplistic and has frequently been questioned. As plant cuticular waxes play pivotal physiological and ecological roles, it might be advantageous to adapt their composition and properties to fluctuating environmental

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conditions. Such dynamic changes might occur on a number of levels, including individual compounds, compound classes or entire wax mixtures, and affect their relative percentages or absolute amounts. Variations might for example be visible in individual leaves according to the growth conditions during organ formation, or induced by environmental conditions after organogenesis. These possible dynamic effects must be distinguished from changes in wax composition caused by ontogenetic programs, that is, differential regulation of biosynthetic pathways during development. Before dynamic effects can be evaluated, the intraspecific variability has to be assessed. In diverse studies, tissues have been systematically harvested from various plant individuals and analysed. Sample sizes typically ranged from n = 3 to 10, and in most cases the reported standard deviations were 5–20% of the average absolute amounts of individual wax compounds. Similar variabilities have been reported for the surface loads of compound classes as well as entire wax mixtures for a number of plant species. A few studies have investigated the ontogenetic development of plant cuticular waxes using organs of different age on the same plant. For example, a comparison of leaves along a single branch of Hedera helix showed that the cuticular wax contained high percentages of alkanes, aldehydes, primary alcohols and fatty acids early on, and increasing amounts of alkyl esters during seasonal development (Hauke and Schreiber, 1998). In the most detailed ontogenetic study available to date, the epicuticular wax film on adaxial surfaces of Prunus laurocerasus leaves was investigated (Jetter and Schäffer, 2001). While the overall amount of waxes increased steadily over the 60 d period of leaf growth and maturation, the relative composition changed three times very drastically (Figure 4.5). In the first 10 d of leaf development alkyl acetates dominated the mixture and then declined gradually. They were replaced by larger amounts of primary alcohols that in turn diminished after approximately 10 d, when finally alkanes started accumulating to result in the mature surface composition. Very similar chemical developments have been found in P. laurocerasus plants grown in different years under various conditions. It must be concluded that the dynamic changes seen in this system reflect an ontogenetic programme that strictly regulates the biosynthesis and accumulation of surface compounds. Unfortunately, comparable data for other plant species are lacking, and it is therefore not clear whether P. laurocerasus shows exceptional dynamic changes during development, or whether similar effects are more widespread. Further evidence for dynamic changes comes from experiments in which cuticular waxes on living tissue have been disturbed at one time and investigated again later. Due to the pronounced developmental effects of its leaf cuticular waxes, P. laurocerasus was again used as a model species. When the epicuticular wax film was removed from the leaves at various stages of development, it was in all cases regenerated within a few days and was found to have the composition typical for that next time in development (Jetter and Schäffer, unpublished results). This shows that regeneration is more than mere re-organisation of preformed compounds, but involves de novo biosynthesis of compounds according to the preset ontogenetic

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Figure 4.5 Development of the epicuticular wax layer on adaxial Prunus laurocerasus leaf surfaces. In the course of leaf growth and maturation, the composition of the epicuticular wax changes repeatedly. Initially alkyl acetates dominate, whereas primary alcohols accumulate to high concentrations between days 14 and 28 of the time course, and alkanes reach highest amounts much later.

program. In an SEM survey, regeneration of surface waxes has also been observed for certain plant species, either restricted to the still growing tissues or continued even after expansion had ceased (Neinhuis et al., 2001). This might point to similar ontogenetic programmes acting in species other than P. laurocerasus. But this generalisation has to be cautious, because the microscopic results are only qualitative. They must be supported by quantitative information, e.g. chemical analyses comparing wax composition before the experiment, immediately after wax removal and after regeneration. Finally, dynamic effects in cuticular wax composition induced by environmental conditions can be discussed. Extensive studies have been carried out to test the effects of light, temperature and air humidity during growth of leaves of B. oleracea (Baker, 1974). In this model system, growth conditions largely influenced the wax composition of the growing tissue. Most interestingly, both increased temperature and decreased relative humidity caused an increase in overall wax amounts, and a shift in percentages from alkanes and ketones to aldehydes and primary alcohols. Changes in irradiant light energy had comparatively little effect. These results are of great interest, as they suggest a feedback mechanism, maybe causing increased

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wax accumulation under transpiration stress. Unfortunately, the transpiration barrier in B. oleracea leaves has not been characterised, and therefore the hypothesis that the physiological properties of the waxes of this species are affected by changes in composition cannot be tested at present. Additional studies have addressed the effects of growth conditions on cuticular wax composition in other species as well. In one example, it was reported that drought caused a slight decrease in the percentage of alkanes in leaf wax of Rosa x hybrida, but no change in the overall wax amounts (Jenks et al., 2001). Even though this effect was only very small, it is similar to the results for B. oleracea and might hence reflect a more general phenomenon. In another study, reduced light was found to decrease average chain lengths in various wax compound classes of H. vulgare leaves (Giese, 1975). Overall, relatively small effects of growth conditions have been found for plant species other than the Brassica species. As summarised in Section 4.3, the wax compositions and quantities found on surfaces of diverse species are fairly stable characters, even though plants had been grown under various (uncontrolled) conditions. It therefore seems that, under normal circumstances and for many species, wax properties are largely controlled by genetic programmes rather than by environmental factors. The very limited information available to date suggests that the Brassica leaf wax composition is exceptionally responsive to changes in environmental conditions. It will be most interesting to study the effect of growth conditions on the regulation of wax biosynthesis once the necessary molecular tools become available, focusing on comparisons between Brassica and other model species.

4.6 Arrangement of plant cuticular waxes In recent years, much progress has been made in combining chemical (see Sections 4.4 and 4.5) with structural (Chapter 2) information on plant cuticles, so that we can now clearly distinguish three layers within the cuticular wax defined as intracuticular wax, epicuticular wax film, and epicuticular wax crystals protruding from this film. All the available evidence suggests that molecules of wax components spontaneously self-arrange and segregate into the three layers. In the following section, we will summarise our current knowledge on (1) the composition of epicuticular wax crystals, (2) the processes leading to separation of epicuticular film and intracuticular wax and (3) the crystalline arrangement of wax molecules within all three cuticular compartments.

4.6.1

Formation of epicuticular crystals

Scanning electron microscopy surveys of various plants revealed a fascinating diversity in the shape, size and arrangement of epicuticular wax crystals (Jeffree, 1986; Barthlott et al., 1998). Due to this diversity, questions arose regarding the composition, organisation, biogenesis and function of these surface structures. In an

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169

initial effort to answer these questions, comparative studies were performed over more than three decades. They revealed good correlations between characteristic shapes and the predominance of certain compounds in the wax mixture. For example, the presence of platelet-shaped crystals in many cases coincided with high concentrations of primary alcohols in the total wax, while epicuticular tubules were correlated with nonacosan-10-ol, and threads with triterpenoids (Figure 4.6). In a second approach, in vitro crystallisation experiments were performed with solutions of pure compounds or wax fractions containing them. They yielded structures that closely matched those on the plant surfaces in shape, size and arrangement (Jeffree et al., 1975; Jetter and Riederer, 1994; Meusel et al., 1999), showing that the presence of these components in high concentrations was the only and sufficient prerequisite for formation of the epicuticular wax crystals. These studies also confirmed that the characteristic shapes of crystals were due to the presence of certain compounds (Jeffree, 1986). Alternative hypotheses, explaining the shapes of epicuticular wax by its extrusion through pores of characteristic geometries (Hall and Donaldson, 1962), were ruled out. In order to gain direct evidence for composition of surface wax crystals, methods for the selective sampling of epicuticular material had to be developed. Most of the previous experiments had employed surface extraction of the intact tissue with organic solvents. The solvents, however, do not only mobilise epicuticular wax, but also enter into deeper layers of the cuticle where they release intracuticular wax (Jetter et al., 2000). Thus, extractive methods can only give information on total wax composition, and it was not clear whether the composition of the epicuticular wax structures differed from the intracuticular wax layer underneath. Jeffree (1996) and Ensikat et al. (2000) reported that frozen droplets of polar liquids can be used to mechanically remove epicuticular wax crystals from plant surfaces and transfer them onto artificial substrates. This new sampling method was then employed to probe the smooth epicuticular wax film on P. laurocerasus leaves (see Section 4.6.2), and for the first time epi- and intracuticular wax constituents could be quantified separately (Jetter et al., 2000). To date, mechanical sampling has been applied only in one study of epicuticular crystals – focusing on surfaces of N. alata pitchers (Riedel et al., 2003). Parts of the inner walls of these pitchers are covered with wax platelets, creating a microscopically rough surface that is assumed to be slippery for insect feet and helps to catch prey in these carnivorous plants (Knoll, 1914). Previous analyses had been based on extraction of total waxes, and consequently did not allow the identification of the wax constituents involved in the formation of these crystals (Juniper et al., 1989). The selective removal of epicuticular wax with frozen water as a cryo-adhesive demonstrated that the wax platelets consisted predominantly of the C30 aldehyde triacontanal (Riedel et al., 2003). This investigation presents the first direct evidence that the accumulation of a single compound is responsible for the formation of unique surface wax structures. At the same time, the results showed that wax platelets that are virtually identical in shape can be formed on different plant species either by primary alcohols or by aldehydes (Figure 4.6).

Ricinus communis∗

H

O

Lupeol HO

O H3C C O

acetate

β-Amryrin

Hentriacontane-14,16-dione

CH3 (CH2)12 C (CH2) C (CH2)14 CH3

O

Nonacosan-10-ol

CH3 (CH2)8 CH (CH2)18 CH3

OH

CH3 (CH2)28 C Triacontanal

CH3 (CH2)14 C (CH2)14 CH3 Hentriacontan-16-one O

CH3 (CH2)24CH2 Hexacosanol O

OH

Main crystal component

β-Amryrin

Macaranga tanarius∗ HO

Dudleya brittonii

Hordeum vulgare

Eucalyptus globulus

Taxus baccata∗

Pogonatum urnigerum

Nelumbo nucifera

Picea pungens

Papaver somniferum

Nepenthes alata∗

Allium porrum

Hordeum vulgare

Pisum sativum

Species

(unpublished)‡

Hobl, Guhling and Jetter

Markstädter et al., 2000

Manheim and Mulroy, 1978

and Netting, 1976a

Horn et al., 1964 von Wettstein-Knowles

Jeffree et al., 1975¥ Jetter and Riederer, 1994¥ Barthlott et al., 1996 Neinhuis and Jetter, 1995 Wen and Jetter (unpublished)

Riedel et al., 2003‡

Rhee et al., 1998

Giese, 1975

Macey and Barber, 1970

Selected references

Figure 4.6 Examples for characteristic shapes of epicuticular wax crystals caused by their main components. Platelets are formed either by primary alcohols, ketone or aldehydes, tubules are formed by asymmetric secondary alcohols (nonacosan-10-ol) or by β-diketones, while threads are formed by triterpenoids. The SEMs shown are examples from the species marked by an asterisk (*). Evidence for crystal-forming compounds is provided by correlation (unmarked references), by in vitro reconstitution experiments (references marked by ¥ ), or by selective crystal sampling and analysis (references marked by ‡ ).

Threads

Tubules

Platelets

Crystal shape

170 BIOLOGY OF THE PLANT CUTICLE

COMPOSITION OF PLANT CUTICULAR WAXES

171

In the epicuticular wax of N. alata pitchers, small amounts of primary alcohols with a broad chain length distribution and tetracosanoic acid (C24 fatty acid) were also detected. The same compounds were present in the underlying intracuticular wax layer, albeit in much higher concentrations. In contrast, aldehydes were found at much lower concentration in the intracuticular wax than in the epicuticular crystals of N. alata (Riedel et al., 2003). In summary, gradients of aldehydes and countergradients of alcohols and fatty acids exist between intra- and epicuticular wax of this species. All the current results taken together, crystal formation can be interpreted as a spontaneous physical process of phase separation within the cuticular wax mixture (Jetter and Riederer, 1994). Once the crystal-forming compound accumulates above a critical concentration in the mixture, it starts to form a separate solid phase. From comparative studies of related plant species with/without epicuticular wax crystals, the threshold concentrations for some crystal-forming compounds can be assessed. Jetter and Riederer (1996) accordingly estimated that nonacosan-10-ol must be accumulated beyond 40% before tubular crystals start to form. Above that threshold composition, crystal-forming compounds will accumulate in the crystals. The crystals consequently differ dramatically from the bulk wax composition. In a number of cases, a clear correlation between single dominating wax constituents and the shape of epicuticular crystals could not be established (Figure 4.7). Conflicting evidence has, for example, been reported (1)

for B. oleracea, where three different compound classes together form the surface structures, (2) for B. oleracea and Arabidopsis thaliana, as they have very distinct surface features but share similar wax composition, (3) for transversely ridged rodlets that have very similar appearance on the surfaces of diverse plant species, even though they consist either of alkanes, or of symmetrical or asymmetrical ketones and (4) for one of these compounds, hentriacontan-16-one, as it can alternatively also form platelet-shaped crystals on Allium porrum (Rhee et al., 1998).

Based on this information, it seems likely that crystals with the characteristic shapes of dendrites, longitudinal bundles of rodlets and transversely ridged rodlets are formed as a consequence of local crystallisation conditions together with chemical composition. To date, there is no direct evidence for the composition of these epicuticular structures, and it cannot be judged whether they differ from the underlying intracuticular layer.

4.6.2

Chemical differences between epicuticular film and intracuticular wax

The compositional gradients between surface crystals and the underlying wax mixture led to speculations that similar differences might exist between epicuticular wax films and the intracuticular layer. Therefore, a distinction between intra- and

Osmunda regalis*

Gypsophila acutifolia

Aristolochia gigantea Liriodendron tulipifera

(CH 2) 13 CH 3

O

Nonacosan-15-ol

CH 3 (CH 2) 13 CH (CH 2) 13 CH 3

OH

Nonacosan-15-one

CH 3 (CH 2) 13 C

O

Nonacosan-15-ol

CH 3 (CH 2) 8 C (CH 2) 18 CH 3 Nonacosan-10-one

O

CH 3 (CH 2) 29 CH 3 Hentriacontane

CH 3 (CH 2) 14 C (CH 2) 14 CH 3 Hentriacontan-16-one

Nonacosane

OH CH 3 (CH 2) 13 CH (CH 2) 13 CH 3

CH 3 (CH 2) 27 CH 3

Arabidopsis thaliana*

Nonacosane

(CH 2) 13 CH 3

Nonacosan-15-one

CH 3 (CH 2) 13 C

O

Main crystal component

CH 3 (CH 2) 27 CH 3

Brassica oleracea*

Species

Jetter and Riederer, 2000

Meusel et al., 1999¥

Meusel et al., 1999¥ Gülz et al., 1992

Jenkset al., 1995 Rashotteet al., 1997 Rashotteet al., 2001

Baker, 1974 Holloway et al., 1977a

Selected references

Figure 4.7 Examples for characteristic shapes of epicuticular wax crystals caused by their main components together with the (local) crystallisation conditions. The same compounds form different crystal shapes on Brassica oleracea leaves and Arabidopsis thaliana inflorescence stems, while various compounds form similar crystals in the shape of transversely ridged rodlets on the leaf surfaces of Aristolochia gigantea, Gypsophila acutifolia and Osmunda regalis. The SEMs shown are examples from the species marked by an asterisk (*). Evidence for crystal-forming compounds is provided by correlation (unmarked references), or by in vitro reconstitution experiments (references marked by ¥ ).

2 µm

Transversely ridged rodlets

2 µm

Longitudinal bundles of rodlets

2 µm

Dendrites

Crystal shape

172 BIOLOGY OF THE PLANT CUTICLE

COMPOSITION OF PLANT CUTICULAR WAXES

173

epicuticular wax was attempted, initially by variations of solvent extraction methods (Silva Fernandes et al., 1964; Baker and Procopiou, 1975). Wax released by very brief (surface) extraction of the intact tissue was considered as epicuticular, while thorough extraction of isolated cuticular membranes was performed in order to analyse the intracuticular wax constituents. Later, films of cellulose acetate or nitrocellulose were employed to remove waxes mechanically from the plant surface (Baker et al., 1983; Haas and Rentschler, 1984). In all these studies, gradients in the percentages of individual compounds were detected, suggesting chemical differences between the intracuticular wax and the epicuticular film. But, unfortunately, even the mechanical probing protocols involved organic solvents that probably led to the (partially) mixed extraction of intra- and epicuticular waxes. Therefore, none of the methods was sufficiently selective to allow a reliable quantification of the compositional gradients between wax layers. The development of the cryo-adhesive technique (see Section 4.6.1) allowed for the first time the sampling of epicuticular wax films from plant surfaces with high selectivity. This new method was employed to probe the smooth epicuticular wax film on P. laurocerasus leaves and to quantify its constituents (Jetter et al., 2000). Steep gradients between the epi- and intracuticular wax layers were detected, proving that the mechanical sampling technique has a much higher selectivity than the previously used extractive protocols. Based on these quantitative results, the selectivity of other mechanical sampling techniques were tested, and aqueous solutions of gum arabic established as a second adhesive for the selective removal of epicuticular waxes (Jetter and Schäffer, 2001). Employing cryo-adhesive sampling, the epicuticular wax load on adaxial cuticles of mature P. laurocerasus leaves was found to be 13 μg/cm2 (Jetter et al., 2000). With a total wax coverage of 28 μg/cm2 and a cutin matrix of 333 μg/cm2 (Schreiber and Riederer, 1996), the epicuticular wax film accounted for less than 4% of the cuticular material. The thickness of the epicuticular layer could be estimated based on the amount of material removed (approximately 130 nm), and the result agreed well with the thickness measured using SEM ( 18 : 1 > 18 : 0 > 14 : 0; A. thaliana mutant phenotype

Acyl-ACP thioesterase FATB class (FATB)

1

Molecular evidence

Biochemical evidence

Enzyme and abbreviation

Reaction in Figure 5.1

Table 5.1 Summary of enzymatic steps involved in cuticular wax biosynthesis. Current knowledge of the enzymes catalysing individual reactions is provided, and the information on cloned genes encoding these enzymes, wherever available, is given.

186 BIOLOGY OF THE PLANT CUTICLE

Solubilised 58 kDa protein from P. sativum microsomes catalyses (C16 ) alcohol formation in vitro; expression of the jojoba FAR gene in B. napus and E. coli resulted in alcohol formation

Expression of the jojoba WS gene in A. thaliana embryos resulted in the formation of alkyl esters

The presence of a relatively constant proportion of free VLCFAs in wax mixtures suggests a requirement for a thioesterase

Solubilised 28 kDa protein from P. sativum microsomes catalyses (C16 ) aldehyde formation in vitro

Conversion of aldehydes to alkanes with release of CO demonstrated in vitro using P. sativum microsomes

Enzyme activity shown in leaf tissue of B. oleracea by assay for hydroxylation of radiolabelled alkane yielding secondary alcohols

Enzyme activity shown in leaf tissue of B. oleracea by assay for oxidation of radiolabelled secondary alcohols yielding ketones

Alcohol-forming fatty acyl reductase (FAR)

Wax synthase (WS)

Putative thioesterase

Putative aldehyde-forming fatty acyl reductase

Putative decarbonylase

Putative hydroxylase

Putative oxidase

7

8

9

10

11

12

13

Molecular identity of the proposed oxidase is unknown

Kolattukudy et al., 1973

Kolattukudy et al., 1973

Molecular identity of the proposed hydroxylase is unknown

Lardizabal et al., 2000

Metz et al., 2000 Vioque and Kolattukudy, 1997

Cheesbrough and Kolattukudy, 1984

WS (S. chinensis)

FAR (S. chinensis)

Molecular identity of the proposed decarbonylase is unknown

Molecular identity of the proposed reductase is unknown

Molecular identity of the proposed thioesterase enzyme/gene is unknown

Demonstrated in vivo role in biosynthesis of alkyl esters in S. chinensis (jojoba) embryos

Demonstrated in vivo role of NADPH-dependent alcohol-forming reductase in biosynthesis of fatty alcohols in S. chinensis (jojoba) embryos

BIOSYNTHESIS AND TRANSPORT OF PLANT CUTICULAR WAXES

187

188

BIOLOGY OF THE PLANT CUTICLE

Unlike the condensing enzymes, the other three enzyme activities of the FAE are presumably identical in all plant VLCFA elongases. They have been suggested to have broad substrate specificities and generate acyl products of diverse chain lengths used to make different classes of lipids (Millar and Kunst, 1997), including cuticular waxes, seed triacylglycerols (TAGs) and sphingolipids. This hypothesis has not been tested directly until recently because the genes encoding the two reductases and the dehydratase have not been available. Although the dehydratase remains unknown, the cloning and characterisation of the β-ketoacyl reductase (Beaudoin et al., 2002) and enoyl reductase (Kohlwein et al., 2001) genes from S. cerevisiae allowed the identification of the corresponding genes from plants. The maize GLOSSY8 (GL8) gene was previously suggested to encode a reductase involved in fatty acid elongation (Xu et al., 1997). Evidence presented by Beaudoin et al. (2002) and additional characterisation of the maize glossy8 mutant (Xu et al., 2002) confirmed that the maize GL8 functions as a β-ketoacyl-reductase that is required for wax production. Further analyses of the maize genome revealed that there is another GL8-like gene present in maize (Perera et al., 2003; Dietrich et al., 2005). Both genes, named GL8A and GL8B, are not expressed exclusively in the epidermis, but also in internal tissues of a number of organs, suggesting that they may not be involved only in wax production. In addition, attempts to generate double mutants by crossing gl8a × gl8b failed because embryos carrying both mutations were not viable. This information that β-ketoacyl-reductase activity is essential for normal development of maize kernels demonstrates that it has a more general role than wax biosynthesis (Perera et al., 2003). It was suggested that reduced sphingolipid levels and altered composition of sphingolipids detected in the double mutant may be responsible for embryo lethality (Dietrich et al., 2005). A BLAST query of the A. thaliana genome database using the β-ketoacyl reductase sequence from S. cerevisiae (Beaudoin et al., 2002) resulted in the identification of two putative homologues. One of these A. thaliana sequences was previously reported to be an orthologue of the maize GLOSSY8 (GL8) gene, and therefore suggested to encode a reductase involved in fatty acid elongation (Xu et al., 1997). Further molecular and biochemical characterisation of these genes/enzymes is required to determine their biochemical and biological functions. An A. thaliana single-copy gene was identified as an enoyl-CoA reductase (ECR) candidate also based on its similarity to the yeast tsc13 gene encoding the ECR (Kohlwein et al., 2001). Heterologous expression of the putative plant ECR gene rescued the temperature-sensitive lethality of yeast tsc13-1elo2 cells (Gable et al., 2004) demonstrating that it functions as an ECR. The A. thaliana ECR gene is ubiquitously expressed, and the protein physically interacts with the ELO2P and ELO3P condensing enzymes when expressed in yeast (Gable et al., 2004). The A. thaliana ECR was shown to be identical to CER10 (Zheng et al., 2005) – the gene defective in one of the original eceriferum A. thaliana mutants identified by Koorneef et al. (1989). Biochemical analysis of the cer10 mutant demonstrated that the ECR gene product is involved in VLCFA elongation required for the synthesis

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189

of all the VLCFA containing lipids, including cuticular waxes, seed TAGs and sphingolipids (Zheng et al., 2005). Surprisingly, knockout mutations in the single A. thaliana ECR gene are not lethal. Furthermore, in the absence of the ECR gene product, cer10 mutants still accumulate 40% of wild-type cuticular wax, as well as VLCFAs in sphingolipids and seed TAGs. It is possible that the ECR identified on the basis of sequence similarity to the yeast TSC13P is not the only ECR in A. thaliana. Alternatively, unknown enzymes functionally similar to the ECR may complement the CER10 deficiency by supporting VLCFA synthesis (Zheng et al., 2005).

5.3 Biosynthetic pathways to monofunctional aliphatics 5.3.1

Synthesis of primary alcohols and wax esters

Primary alcohols are found in cuticular wax mixtures either as free alcohols or in the form of esters of fatty alcohols and fatty acids (Figure 5.1). From the time they were discovered in cuticular wax, primary alcohols were assumed to be produced by the reduction of fatty acyl-CoA. However, evidence for such a reduction was not available until the 1970s when the biosynthesis of alcohol from the fatty acyl-CoA using cell free preparations from Euglena gracilis was demonstrated (Kolattukudy, 1970b). Although the alcohol-generating reduction presumably proceeds through an aldehyde intermediate, a free aldehyde was not released, but could be demonstrated by chemical trapping (Kolattukudy, 1970b). In higher plants, the reduction of fatty acyl-CoAs to fatty alcohols was first reported for B. oleracea (Kolattukudy, 1971). Partial purification of reducing activities from B. oleracea leaves led to a proposal that primary alcohol production is a two-step process carried out by two separate enzymes – an NADH-dependent acyl-CoA reductase required for a reduction of fatty acids to aldehydes, and an NADPH-dependent aldehyde reductase required for a further reduction of aldehydes to primary alcohols (Kolattukudy, 1971). Initially, support for the involvement of two reductases in the conversion of fatty acids to alcohols came from the analyses of the gl5 mutant of Z. mays in which a block in production of primary alcohols results in high accumulation of aldehydes (Bianchi et al., 1978). However, further analyses of the gl5 maize line revealed that it actually carried two mutations at different loci, subsequently designated GL5 and GL20 (von Wettstein-Knowles, 1995), and only lines homozygous for both mutations were glossy. Thus far wax analyses of the gl5 and gl20 single mutant lines have not been performed, and the mutated genes have not been identified. Without this information it is not possible to rationalise the biochemical phenotype observed in the original gl5gl20 double mutant. Important contributions to the study of alcohol forming fatty acyl-CoA reductases (FAR) have come from the study of developing jojoba seeds (Simmondsia chinensis). This is an unusual plant which produces wax esters comprised of longchain alcohols and fatty acids as a seed lipid energy reserve. Biochemical studies by Pollard et al. (1979) indicated that in jojoba embryos alcohol formation from

190

BIOLOGY OF THE PLANT CUTICLE

fatty acid precursors requires a single FAR, and that it proceeds without the release of the free aldehyde. In support of this notion, Vioque and Kolattukudy (1997) purified two distinct reductase activities from pea (Pisum sativum) and demonstrated that the 58 kDa enzyme generates fatty alcohols without any detectable accumulation of fatty aldehyde intermediates, as first described in E. gracilis (Kolattukudy, 1970b). These results were verified by solubilisation and biochemical characterisation of a FAR from the jojoba embryos and cloning of the corresponding cDNA. Expression of the jojoba FAR cDNA in Escherichia coli and Brassica napus resulted in a functional FAR enzyme and the accumulation of fatty alcohols in E. coli, as well as fatty alcohols together with wax esters in transgenic B. napus oil (Metz et al., 2000). The pea and the jojoba alcohol-forming fatty acyl reductases are both integral membrane proteins, which appear to be associated with the ER and have similar molecular masses in the range of 56–58 kDa (Vioque and Kolattukudy, 1997; Metz et al., 2000). FAR-related sequences from maize (Z. mays), rice (Oryza sativa), cotton (Gossypium hirsutum) and B. napus found in the public databases, and a family of eight FAR-like proteins in A. thaliana (Kunst and Samuels, 2003) suggest that alcohol-generating FARs are ubiquitous in plants. One of the eight FAR-like A. thaliana sequences is likely the CER4 gene, given that the cer4 mutant exhibits major decreases in primary alcohols and wax esters, and slightly higher levels of aldehydes, alkanes, secondary alcohols and ketones (Hannoufa et al., 1993; Jenks et al., 1995). The cer4 biochemical phenotype has been attributed to a lesion in an aldehyde reductase of the acyl-reduction pathway (Hannoufa et al., 1993; Jenks et al., 1995). In view of the new evidence indicating that primary alcohols are produced from fatty acids without release of an aldehyde intermediate, the cer4 phenotype is most likely due to a mutation in the alcohol-forming acyl-CoA reductase, not the aldehyde reductase. The observed increase in aldehyde, alkane, secondary alcohol and ketone levels in the cer4 mutant can then be explained by an increased flux of fatty acyl precursors into the alkane biosynthetic pathway. The primary alcohols generated in the epidermal cells are further used for the synthesis of wax esters (Figure 5.1). Biochemical studies of wax ester formation in B. oleracea leaves (Kolattukudy, 1967a) and jojoba (S. chinensis) seeds (Wu et al., 1981) suggested that this reaction involves the transfer of an acyl chain from fatty acyl-CoA to a fatty alcohol that is catalysed by a membrane-bound acyltransferase (wax synthase, WS). Partial purification of a WS polypeptide from jojoba embryos allowed the identification of a cDNA encoding the WS enzyme (Lardizabal et al., 2000). The identity of the cloned cDNA was demonstrated by activity assays in developing A. thaliana embryos, expressing the jojoba WS cDNA. The transgenic A. thaliana embryos also accumulated high levels of wax esters. Hydropathy analysis of the deduced protein sequence revealed seven to nine transmembrane domains (TMD) suggesting that jojoba WS is an integral membrane protein (Lardizabal et al., 2000). The jojoba WS has considerable similarity with twelve A. thaliana sequences (Lardizabal et al., 2000; Kunst and Samuels, 2003). Several of these twelve putative genes are differentially expressed in inflorescences and seeds of

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191

A. thaliana but to date their enzymatic functions have not been assessed, nor have their biological roles been determined.

5.3.2

Synthesis of alkanes, secondary alcohols and ketones

Alkanes present in cuticular wax are approximately twice as long as the fatty acids of the cellular membranes. This finding, together with the discovery that alkanes, secondary alcohols and ketones of identical chain lengths and same location of their functional groups within their carbon chain occur in wax mixtures isolated from a certain tissue, led to a proposal that alkane molecules are generated by a head-tohead condensation between two fatty acids (Channon and Chibnall, 1929). Such a condensation reaction with decarboxylation of one of them would yield a ketone, which would be reduced to produce a secondary alcohol, then dehydrated and further reduced to generate an alkane. However, labelling experiments with 14 C and/or 3 H showed that head-to-head condensation reaction is not involved in alkane production in B. oleracea (Kolattukudy, 1966), P. sativum or S. oleracea (Kolattukudy, 1968), and that the entire carbon chains of C16 fatty acids were incorporated into alkanes without decarboxylation, that is, without the loss of the carboxyl carbon. This result was further substantiated by the demonstration that fatty acids of various chain lengths (C10 –C18 ) were all incorporated into C29 alkane n-nonacosane with the longer ones being incorporated much more efficiently (Kolattukudy, 1966). Furthermore, the exogenously added labelled C29 ketone nonacosan-15-one could not be converted into C29 secondary alcohol (nonacosan-15-ol) or alkane (n-nonacosane) by B. oleracea leaf disks, peeled epidermis or leaf homogenates (Kolattukudy, 1966). On the basis of this new experimental evidence, Kolattukudy (1966, 1967b) proposed an elongation–decarboxylation pathway for alkane biosynthesis. According to this pathway, C16 fatty acid is elongated by the addition of C2 units and, when it reaches the appropriate length (C30 –C32 ), it gets decarboxylated. The generated alkane can then be hydroxylated to secondary alcohol, which can give rise to a ketone by oxidation. Conversion of 3 H-labelled C29 alkane to C29 secondary alcohol and ketone, and 3 H-labelled secondary alcohol to C29 ketone by B. oleracea leaves confirmed that this sequence of reactions was feasible (Kolattukudy and Liu, 1970). Similarly, 3 H-labelled C30 fatty acid was incorporated into C29 alkane by B. oleracea leaves (Kolattukudy et al., 1972), 3 H-labelled C32 fatty acid into C31 alkane by cell free extracts from P. sativum leaves (Khan and Kolattukudy, 1974) and 3 H-labelled C24 fatty acid (tetracosanoic acid) to C23 alkene (n-tricosane) by Allium porrum (Cassagne and Lessire, 1974). These data, together with the isolation of mutants with lesions in elongation or decarboxylation that also inhibit alkane production (von Wettstein-Knowles, 1993), provided additional support for the elongation–decarboxylation hypothesis of alkane synthesis. Progress leading to proposals for the biochemical steps involved in the production of alkanes, secondary alcohols and ketones has not been matched by identification and biochemical characterisation of enzymes catalysing the implicated reactions. Early experiments, designed to examine the mechanism of fatty acid conversion to

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alkane, demonstrated that cell-free preparations from P. sativum required oxygen and ascorbate for alkane synthesis and that alkane production was strongly inhibited by metal ion chelators (Khan and Kolattukudy, 1974). Subsequent work showed that C18 –C32 fatty acids can serve as substrates for alkane formation, and that all of these substrates generated alkanes two carbons shorter than the parent fatty acid. The discrepancy between this result and the observed predominance of alkanes with odd numbers of carbons found in cuticular wax mixtures was suggested to be due to differences between alkane generating reactions occurring in vitro and in vivo. In vitro α-oxidation of fatty acids gives rise to n-2 alkanes, whereas in vivo aldehydes formed from an acyl-CoA by a reductase could be decarbonylated to n-1 alkane (Bognar et al., 1984). The aldehyde had not been previously recognised as an intermediate in alkane biosynthesis because a chemical mechanism for conversion of an aldehyde to an alkane was not known until the discovery that porphyrin-coordinated ruthenium complexes can catalyse such a decarbonylation reaction (Domazetis et al., 1981). That an analogous reaction can generate alkane and CO from an aldehyde was then shown using P. sativum microsomes (Cheesbrough and Kolattukudy, 1984). However, in these experiments the decarbonylation was not assayed using the natural C30 aldehyde as substrate, but the conversion of C18 aldehyde to C17 alkane was monitored instead. In addition, because the amount of CO released was too low to allow direct chemical detection, the CO was trapped using a rhodium complex, which forms a stable adduct with CO. The recovery of CO from C18 aldehyde was slightly lower than the stoichiometric amount of alkane formed. In order to verify the involvement of the decarbonylase in alkane biosynthesis, dissect the mechanism of conversion of an aldehyde to an alkane and determine the subcellular site of alkane production, it is necessary to isolate the enzyme catalysing this reaction. The purification of the decarbonylase was attempted from P. sativum leaves (Schneider-Belhaddad and Kolattukudy, 2000). However, only partial purification was achieved and the quantity of purified protein was so low that reliable characterisation of the enzyme was not possible. For example, it had not been determined if a porphyrin was present, and the nature of the metal ion, believed to be involved in the synthesis of alkanes because of its sensitivity to metal chelators, remains uncertain. Similarly, mechanistic details of the conversion of aldehydes to alkanes still need to be elucidated. If aldehydes are indeed direct precursors of alkanes, their production requires a FAR. A 28-kDa enzyme that catalyses the conversion of fatty acyl-CoA to an aldehyde in vitro has been solubilised and purified to homogeneity from P. sativum leaves (Vioque and Kolattukudy, 1997). The biochemical identity of this putative reductase, however, needs to be confirmed in planta, and its substrate specificity and cellular compartmentation remain to be determined. Radioactive labelling studies (Kolattukudy and Liu, 1970; Kolattukudy et al., 1972; Khan and Kolattukudy, 1974) and genetic evidence indicating that a block in alkane synthesis results in an absence of secondary alcohols and ketones (McNevin et al., 1993) support the biochemical pathway for the synthesis of secondary alcohols and ketones from alkanes (Kolattukudy and Liu, 1970). At present, however,

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the individual steps in this pathway remain hypothetical, as the enzymes involved in catalysing the proposed reactions have not been identified and characterised. The only available information pertaining to these enzymatic steps dates back to the labelling experiments of Kolattukudy et al. (1973), who showed that 3 H-C29 alkane n-nonacosane was incorporated into two different isomers of C29 secondary alcohol (nonacosan-14-ol and nonacosan-15-ol) in B. oleracea in a ratio similar to that found in the natural mixture, whereas the ketone was almost exclusively nonacosan-15-one. This work also revealed that these reactions require molecular oxygen and are severely inhibited by phenanthroline, a metal chelator. Curiously, in the presence of phenanthroline, incorporation of 3 H into ketones was much higher than into the secondary alcohols. Based on the fact that this inhibition was reversed by Fe2+ , it was suggested that hydroxylation of the C29 alkane n-nonacosane may involve a mixed function oxidase (Kolattukudy et al., 1973), but in the absence of the purified enzyme, this hypothesis could not be experimentally tested. It has also not been determined if hydroxylation in vivo occurs at the alkane level, or whether a hydroxyl group can also be inserted into a fatty acyl or aldehyde precursor. From the earlier discussion, it is clear that progress in our understanding of the biosynthetic pathway leading to the formation of alkanes, secondary alcohols and ketones awaits identification of the enzymes catalysing the formation of these major cuticular wax constituents. Attempts using traditional biochemical approaches have had limited success since wax biosynthetic enzymes are membrane-bound, and because the sufficient amount of epidermal tissue and the substrates required for enzyme activity assays are difficult to obtain. Even though genetic approaches in A. thaliana and Z. mays seem better suited for such a task, they have not yet been fruitful in identifying the genes encoding these enzymes. Only three A. thaliana mutants, cer1, cer22 and wax2, have been reported to have altered levels of alkanes, secondary alcohols and ketones in their stem wax (Hannoufa et al., 1993; McNevin et al., 1993; Lemieux et al., 1994; Jenks et al., 1995; Chen et al., 2003; Kurata et al., 2003; Rashotte et al., 2004). Cer1 and cer22 both exhibit increases in aldehyde levels and dramatic reductions in the accumulation of alkanes, secondary alcohols and ketones suggesting a block in the conversion of aldehyde to alkane. Based on this biochemical phenotype, it was proposed that CER1 may encode an aldehyde decarbonylase (Hannoufa et al., 1993; McNevin et al., 1993). This suggestion was subsequently questioned, because cer1 mutation also decreased the primary alcohol levels in stem wax (Jenks et al., 1995). A simple block in the aldehyde decarbonylase step of the pathway could conceivably result in a greater flux of carbon towards primary alcohols, but reduced accumulation of primary alcohols would not be expected. The cloning of the cer1 gene (Aarts et al., 1995) did not reveal the biochemical identity of the CER1 protein, but the deduced amino acid sequence of CER1 showed that it contains three histidine-rich motifs common with fatty acyl desaturases, alkane hydroxylase and xylene monooxygenase – three enzymes that catalyse similar reactions (Fox et al., 1994; Shanklin et al., 1994). CER1 also has some sequence similarity with proteins of unknown function: EPI23, a Kleinia odora

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epidermis-specific protein, GLOSSY1 protein of Z. mays (Hansen et al., 1997; Sturaro et al., 2005) and WAX2 of A. thaliana (Chen et al., 2003). CER22 was also predicted to be the C30 aldehyde decarbonylase, or a protein regulating its activity in A. thaliana stems (Rashotte et al., 2004), because the mutation in CER22 blocks the conversion of the C30 aldehyde triacontanal to C29 alkane n-nonacosane. Unlike cer1, the cer22 line does not have substantially altered primary alcohol levels, but does exhibit a dramatic increase in the levels of wax esters [300% of the wildtype (WT); Rashotte et al., 2004], presumably due to a redirection of carbon towards primary alcohols. The CER22 gene has not yet been isolated; so the hypothesis that it may encode a protein involved in the conversion of aldehydes to alkanes remains to be confirmed. Wax2/yore-yore/faceless pollen1 mutant of A. thaliana, identified independently by three research groups (Ariizumi et al., 2003; Chen et al., 2003; Kurata et al., 2003), has a glossy bright green stem due to an approximately 80% decrease in total wax load. Aldehydes, alkanes, secondary alcohols and ketones that jointly comprise up to 90% stem wax in WT A. thaliana were severely reduced in this mutant, with corresponding increases detected in the levels of fatty acids, primary alcohols and esters. Cloning of the WAX2 gene indicated that the predicted WAX2 polypeptide is an integral membrane protein sharing the highest sequence similarities with the GLOSSY1 of Z. mays (Sturaro et al., 2005), a rice sequence annotated as a GLOSSY1 homologue (Hansen et al., 1997), K. odora EPI23 (Hansen et al., 1997) and A. thaliana CER1 (Aarts et al., 1995). The authors (Chen et al., 2003; Kurata et al., 2003) speculate that WAX2 may function as an aldehyde-generating acyl-CoA reductase; however, based on the limited amount of information currently available, it is not possible to deduce the biochemical role of WAX2 in wax biosynthesis. Our inability to identify genes encoding enzymes involved in the production of alkanes and their derivatives using genetic approaches may be due to several reasons. First, A. thaliana genome sequencing revealed that only 35% of the predicted protein encoding sequences are unique, while 37% belong to families with more than five members (The Arabidopsis Genome Initiative, 2000). The lack of obvious phenotypes in the vast majority of single gene knockout mutants suggests that this genetic redundancy might be a serious obstacle in the functional characterisation of A. thaliana genes. Second, the standard visual screens for wax-deficient mutants that rely on a change in epicuticular wax crystal density, generating an alteration in surface reflectance, will consistently detect only mutants with pronounced phenotypes, most likely those with lesions in regulatory functions, or early wax biosynthetic steps. Mutants with defects causing more subtle changes in wax load or composition that do not result in changes in surface reflectance (see Chapter 6) would escape visual detection. Therefore, more accurate screening methods, such as gas chromatography, will need to be employed to identify them (Rashotte et al., 2004). Third, it is conceivable that some of the enzymes catalysing the biosynthetic reactions proposed on the basis of radioactive tracer and inhibitor studies are known, but that their involvement in these reactions has not been established,

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because the type of reaction carried out by these enzymes and the mechanism of catalysis is not understood. Rigorous biochemical work on proteins implicated in the production of alkanes, secondary alcohols and ketones, such as CER1 or WAX2, is therefore required to elucidate their precise biochemical function and the nature of the chemical reaction that they catalyse. Finally, the suggested biosynthetic steps are based on studies of alkane, secondary alcohol and ketone formation in leaves of several plant species and may not reflect the situation in all plants and tissues. Verification of this pathway in additional plant species is clearly an important future goal.

5.3.3

Biosynthesis of β-diketones and alkan-2-ols

β-diketones are a major component of the cuticular wax in some Eucalyptus, Festuca and Agropyron species, and are also found in Dianthus and Rhododendron species as well as in cereals including barley (H. vulgare), wheat (Triticum aestivum), oats (Avena sativa) and rye (Secale cereale) (von Wettstein-Knowles, 1972; Tulloch, 1976; Walton, 1990). Occasionally they are accompanied by short esterified alkan2-ols, as in Agropyron, Eucalyptus and barley. In Sorghum bicolor, however, alkan2-ols have been detected in the absence of β-diketones (von Wettstein-Knowles, 1995). The biosynthesis of these lipids is carried out by enzyme complexes with features of both FAE and polyketide synthases. Our present understanding of the β-diketone pathway is based on genetic and biochemical studies of cer-cqu mutants of barley (reviewed by von Wettstein-Knowles, 1993, 1995). Initial genetic analyses of cer-c, cer-q and cer–u mutants showed that these three complementation groups were very tightly linked and that double and triple mutants were present relatively frequently (von Wettstein-Knowles and Søgaard, 1980). Subsequent reversion studies demonstrated that these double and triple mutants could revert to WT as a result of single mutational events (von Wettstein-Knowles, 1992). These results are consistent with a single CERCQU gene coding for a polypeptide with three functional domains, CER-C, CER-Q and CER–U (von Wettstein-Knowles and Søgaard, 1981). Wax analyses of the spikes of the cer-c, cer-q and cer-u barley mutant revealed that the CER-CQU gene product was involved in the production of β-diketones, hydroxy- and oxo-β-diketones, methyl ketones and alkan-2-ols (von WettsteinKnowles, 1976, 1993). CER-Q mutations blocked the synthesis of β-diketones and all their derivatives, indicating that the CER-Q activity was required early in the pathway, and was likely involved in β-ketoacyl elongation (Mikkelsen and von Wettstein-Knowles, 1978; Mikkelsen, 1979). Later biochemical investigations demonstrated that the cer-q domain has β-ketoacyl synthase activity (Mikkelsen, 1984). CER-C mutations blocked β-diketone formation, and resulted in elevated levels of alkan-2-ols, but the activity specified by the CER-C domain is not known, although it is probably involved in two successive rounds of elongation (von Wettstein-Knowles, 1992). CER-U mutations caused the decrease in hydroxyβ-diketone accumulation with a corresponding increase in the β-diketone levels,

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suggesting that the CER-U protein domain functions as a hydroxylase introducing the hydroxyl group into β-diketones (von Wettstein-Knowles, 1972). Further studies using radioactively labelled precursors and tissue slices prepared from the cer-c, cer-q and cer-u mutants provided additional evidence that the β-ketoacyl-CoA compound generated by the β-ketoacyl elongase is the key intermediate in the synthesis of β-diketones and alkan-2-ols (Figure 5.2; Mikkelsen, 1984). This research demonstrated that the β-ketoacyl-CoA intermediate can be converted to a methylketone by a putative decarboxylase, and an alkan-2-ol by a putative methylketone reductase. Finally, the alkan-2-ol can give rise to an ester by reacting with an acyl-CoA (Figure 5.2). Alternatively, the β-ketoacyl-CoA intermediate can undergo successive condensations by the CER-Q β-ketoacyl synthase, followed by a number of chain elongations and decarboxylation or decarbonylation, resulting in the formation of a β-diketone. A hydroxyl group can then be introduced and oxidised to yield a hydroxy-β-diketone and a keto-β-diketone, respectively (von Wettstein-Knowles, 1993). The proposed biosynthetic pathway for the production of β-diketones and alkan-2-ol esters has not yet been verified by molecular and biochemical characterisation of the enzymes that catalyse the suggested reactions. This is clearly an essential next step in understanding the production of these wax constituents.

5.4 Triterpenoid biosynthesis Triterpenoids exhibit a striking diversity of carbon structures and of derivatives with varying functional groups (Xu et al., 2004). Based on rapidly accumulating chemical information on these structures, Ruzicka et al. in the 1950s proposed the concept of the biogenetic isoprene rule (Ruzicka, 1959). It provides the framework describing the biosynthetic pathways and mechanisms leading to the triterpenoid backbones (Eschenmoser et al., 1955), from which the final products are derived by modification reactions. The predictions of the biogenetic isoprene rule have since been confirmed by numerous chemical, biochemical and molecular biological investigations. We thus have a fairly good knowledge of the biosynthetic processes leading to triterpenoid formation in general. Even though published reports on the biosynthesis of cuticular triterpenoids are lacking to date, it is very likely that much of the information available on triterpenoid biosynthesis in general can be applied to these plant surface specific compounds. Most plant triterpenoids are, like steroids, derived from enzymatic cyclisation of 2,3-oxidosqualene, which is therefore the last common precursor in two separate pathways of primary and secondary metabolism (Abe et al., 1993). Protonation leads to epoxide ring opening and creates a carbocation that can be attacked by π-electrons from one of the double bonds in the squalene molecule. A series of these intramolecular addition reactions leads to formation of four fused carbon cycles, the six-membered rings A, B and C in chair conformation and the fivemembered D-ring. The dammarenyl cation thus formed is transformed by widening

Figure 5.2 Biosynthetic pathways for the formation of β-diketones, alkan-2-ols and their derivatives. C14 and C16 fatty acyl-ACPs formed by the fatty acid synthase (FAS) in the plastid are hydrolysed by an (1) acyl-ACP thioesterase (FATB), and during transport through the plastid envelope esterified to CoASH by a (2) long-chain acyl-CoA synthetase (LACS). C14 and C16 fatty acyl-CoAs undergo a (3a) condensation reaction with malonyl-CoA to yield β-ketoacyl intermediates, which can then be converted into methylketones, alkan-2-ols and alkan-2-ol esters. Alternatively, C14 and C16 acyl-CoAs can undergo two successive condensations carried out by one or two KCSs (3a and 3b), followed by several rounds of FAE to yield C32 or C34 β-diketoacyl intermediates used for the production of β-diketones, hydroxy- and keto-β-diketones. Enzymatic steps (1)–(6) shown in this figure are described in more detail in Table 5.1. Chemical formulas for one representative compound from each lipid class are shown.

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of the D-ring, followed by formation of the E-ring. The product of this reaction sequence (the lupenyl cation) is deprotonated in many cases, leading to lupeol as one of the most widespread triterpenoids. Alternatively, it can be rearranged in a series of 1,2-hydride and/or 1,2-methyl shifts, and the resulting cations can be deprotonated to produce the various other triterpenoid structures. The products are isomeric structures C30 H50 O, either with a hydroxyl function on C-3 and a C==C double bond in specific positions in the molecule, or with a carbonyl function at C-3 and a saturated carbon backbone. It has been shown that all the reaction steps from oxidosqualene to the deprotonation products can be catalysed by a single enzyme – a triterpene synthase (Herrera et al., 1998; Kushiro et al., 1998). A number of triterpene synthases have been cloned and characterised from various plant species. In most cases, they form predominantly one product, for example β-amyrin or lupeol. The mechanism of the cyclisation reaction has been investigated in some detail in these two enzyme groups. Active site residues involved in the protonation of the epoxide substrate, and in the stabilisation of carbocation intermediates have been identified (Tanaka et al., 2002). In addition, amino acids involved in determination of product specificity have been established, making it possible to switch a β-amyrin synthase into a lupeol synthase (and vice versa) by site directed mutagenesis of single residues (Kushiro et al., 2000). The only other product-specific triterpene synthase presently known is an isomultiflorenol synthase from Luffa cylindrica (Hayashi et al., 2001). Two multifunctional triterpene synthases have been described and are the only enzymes currently shown to form α-amyrin (Husselstein-Muller et al., 2001). Besides, they generate mixtures of either taraxasterol, -taraxasterol, multiflorenol, tirucalla7,21-dienol, butryrospermol and baurenol, or of δ-amyrin, germanicol, taraxasterol, -taraxasterol and butryrospermol. The species-specific triterpenoid compositions may therefore be formed by multiple single-product enzymes in some cases, and by one (or a few) multi-product enzyme(s) in others. The triterpenoid compositions have not been reported for all the plant species from which relevant genes have been cloned; thus it is not possible to assess in all cases whether the set of in vitro characterised genes is sufficient to explain in planta product formation. The same carbon structures, mostly with the hydroxyl function on C-3, have been reported for the cuticular triterpenoids of diverse species (Chapter 4, Section 4.2.3). For this reason it seems likely that very similar enzymes are involved in the biosynthesis of surface triterpenoids. Unfortunately, the spatial and temporal expression patterns of the triterpenoid synthase genes cloned to date have not been reported. The cuticular wax compositions of the species from which these genes were cloned have also not been investigated. Consequently, it is not clear whether the triterpenoid products of the enzymes characterised so far might be partially or completely exported to the cuticle. While the cuticle-relevance of the known genes/species is being tested, homologous genes from other species with reportedly high concentrations of cuticular triterpenoids should also be investigated.

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5.5 Regulation of wax biosynthesis Differences in plant resistance/susceptibility to environmental stresses, pathogens or insects have been linked to variations in wax accumulation (load) and wax composition among plant species (Eigenbrode and Espelie, 1995; Post-Beittenmiller, 1996; Chapters 11 and 12). The mechanisms by which plants control wax accumulation and composition are therefore of considerable interest. At present, however, the knowledge of these regulatory mechanisms is extremely limited. Cer mutants of A. thaliana and glossy mutants of Z. mays have been valuable resources for the identification and isolation of genes associated with plant cuticular wax production. Four of the genes cloned from these mutant collections, CER2, CER3, GL2 and GL15, have been proposed to encode regulatory proteins (Tacke et al., 1995; Hannoufa et al., 1996; Moose and Sisco, 1996; Negruk et al., 1996; Xia et al., 1996). Of those, only the GL15 was suggested to encode a transcription factor that regulates leaf epidermal cell identity, whereas the identities of CER2, CER3 and GL2 gene products could not be deduced from their primary sequences, and their predicted functions in regulation of wax deposition remain to be confirmed. Detailed expression analyses have been carried out for only two genes encoding wax biosynthetic enzymes, CER2 and CER6 (Xia et al., 1997; Hooker et al., 2002). Both studies documented that these genes were transcriptionally regulated in response to developmental cues. In addition, CER6 transcription was induced by light and osmotic stress (Hooker et al., 2002), environmental factors known to stimulate wax accumulation in a few species, for example B. oleracea (Baker, 1974). Somewhat unexpectedly, CER2 expression was not light-, heat-, cold- or wound-inducible, and was unaffected by osmotic stress (Xia et al., 1997). Despite the available evidence that wax production is under transcriptional control, transcriptional activators upregulating the expression of wax biosynthetic genes during development and in response to environment have not yet been identified. Recently, however, two groups reported the isolation and characterisation of WIN1/SHN1, an A. thaliana transcriptional regulator of the AP2/EREBP family which, when overexpressed, dramatically enhances wax accumulation in A. thaliana leaves and stems, and results in a strikingly glossy leaf phenotype (Aharoni et al., 2004; Broun et al., 2004). A detailed examination of the molecular mechanisms underlying the wax hyperaccumulation in the leaves of 35S:WIN1/SHN1 transgenic plants demonstrated that WIN1/SHN1 overexpression resulted in the induction of several wax-related genes, including CER1, CER2 and KCS1 (Broun et al., 2004), and consequently in a dramatic increase in leaf alkane, secondary alcohol and ketone levels (Aharoni et al., 2004; Broun et al., 2004). Extreme leaf glossiness and increased wax deposition on leaves and stems have also been detected in transgenic plants overexpressing the WIN1/SHN1 paralogues SHN2 and SHN3 (Aharoni et al., 2004; Broun et al., 2004). In addition, RNA-blot and microarray analyses showed that other genes predicted to encode lipid biosynthetic enzymes, and proteins involved in cellular trafficking, including an ABC transporter were

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up-regulated in the 35S:WIN1/SHN1 overexpressors (Broun, 2004). Similarly, WXP1, an AP2/EREBP domain transcription factor from Medicago truncatula, increases leaf cuticular wax accumulation and results in a glossy leaf phenotype when overexpressed under the control of the 35S promoter in alfalfa (Medicago sativa; Zhang et al., 2005). Unlike in A. thaliana, however, where SHN transcriptional regulators activate the production of alkanes and their derivatives, the wax load increase in alfalfa is mainly due to a greater production of the C30 primary alcohol triacontanol, the major component of the alfalfa leaf wax. Even though high levels of expression of SHN transcription factors under the control of the 35S promoter had pleiotropic effects on A. thaliana growth and development, and their exact in planta role could not be determined in the absence of loss-of-function phenotypes or fully silenced transgenic lines, it is tempting to speculate that the major function of the SHN clade of transcription factors in A. thaliana, and related AP2/EREBP domain transcription factors in other plants is the activation of the wax biosynthetic and export machinery in epidermal cells.

5.6 Wax biosynthesis and transport in the context of the epidermal cell During formation and growth of aboveground plant organs, their surface area typically expands first due to division and later due to controlled anisotropic expansion of epidermal pavement cells. Cuticle formation and growth have to be synchronised with this expansion in order to completely coat the surface for protection at all times. Investigations into the development of cuticle structure and composition have in many cases found that the major cuticular components, for example, the epicuticular wax crystals on the tissue surface, are in place already at the earliest time points studied, as well as during ensuing surface expansion. During growth, large quantities of cutin and wax have to be deposited on the periclinal surface of the growing cells. It is generally accepted that the cuticular components originate in the epidermal pavement cells, starting from ubiquitous precursors such as acetyl-CoA and involving common fatty acyl intermediates. Consequently, the epidermis cells have to strike and maintain a delicate balance of biosynthetic activities, coordinating the large flux of lipophilic products towards the cuticle with other lipid generating pathways. The large quantities of wax intermediates and products might also pose structural problems for the pavement cells, as they must handle saturated aliphatic molecules that, due to their lipophilic nature and very long-chain geometry, may interfere with the integrity of cell membranes (Ho et al., 1995; Hamilton, 1998; Millar et al., 1998). By analogy to numerous lipid transport processes in diverse eukaryotes, two fundamental mechanisms for handling the wax precursors and products may be distinguished. Both will likely be involved in channelling the molecules between biosynthetic enzymes and the cuticle. One possibility is that wax components are embedded in the membrane bilayers, spanning nearly their entire thickness. In that case special membrane microdomains or presently uncharacterised proteins might

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help to accommodate them to prevent disruption of cellular membrane functions. Alternatively, special proteins might serve to solubilise wax compounds in the cytoplasm, possibly by binding them as single molecules at the site of synthesis, and shuttle them across aqueous compartments on their way to the cuticle. En route to aliphatic wax compounds, 16:0-ACP and 18:0-ACP are synthesised in the plastids of pavement cells and hydrolysed by acyl-ACP thioesterases (see Section 5.2). During or after transfer through the plastid membranes, the free fatty acids are transformed into long-chain acyl-CoAs in the plastid envelope. Currently, nothing is known about the mechanism of acyl-CoA transport from the epidermal plastids to the ER, but at least two hypotheses may be considered. In many plant species and cell types, domains of the ER have been found located near the plastids, without apparent fusion or mixing of bilayers (Staehelin, 1997). This proximity may facilitate fatty acid transfer to the ER, involving non-vesicular mechanisms such as spontaneous desorption, diffusion and absorption. Alternatively, lipid transport from plastids to the ER could be facilitated by acyl-CoA binding proteins (ACBPs) (Johnson et al., 2002). This class of proteins has been described in a wide variety of eukaryotes (Chye et al., 2000), and a S. cerevisiae mutant lacking ACBP was found to have increased levels of C18:0 and reduced C26:0 . This result suggests that in yeast ACBP is involved in transport of acyl-CoAs towards the ER, where they are elongated from C18:0 to C26:0 for production of sphingolipids (Gaigg et al., 2001). In plants, both a cytoplasmic ACBP from A. thaliana (Engeseth et al., 1996; Leung et al., 2005) and membrane associated ACBPs (Li and Chye, 2003) have been described. Although it is postulated that their role is to maintain an acyl-CoA pool in the cytosol (Engeseth et al., 1996) or transport palmitoyl-CoA or oleoyl-CoA from the plastid to the ER (Leung et al., 2005), their in vivo functions remain to be investigated. As the ACBP1 and ACBP2 were located in the plasma membrane (PM) rather than the cytoplasm (Li and Chye, 2003; Leung et al., 2005), it is unlikely that they are trafficking lipids towards the ER. However, this does not preclude the possibility that other ACBP orthologues are involved in this transport process.

5.6.1

Intracellular sites of wax synthesis and trafficking of wax constituents

Multiple lines of evidence support the model that enzymes catalysing the elongation of long chain to VLCFAs are associated with the ER (Figure 5.3, scenario 1). In cell fractionation studies, fatty acid elongation activity was most enriched in microsomal fractions (Lessire et al., 1985; Bessoule et al., 1989), i.e. membrane preparations dominated by ER material. After the genes encoding VLCFA biosynthetic enzymes had been isolated, some of these enzymes have also been localised using fluorescent tags. For example, an A. thaliana β-ketoacyl-CoA synthetase (CER6) (Kunst and Samuels, 2003) and ECR (CER10) (Zheng et al., 2005) were fused to the green fluorescent protein (GFP) and expressed in the corresponding mutants. Both enzymes complemented the mutant phenotypes, indicating that they were functional, and were localised throughout the ER of epidermal pavement cells (Figure 5.4).

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Wax

Cutin Cell wall

4a

4a

4b

Plasmamembrane

3

3

Cytoplasm

4b

2a 2b

Endoplasmic reticulum 1

2c

Golgi

Figure 5.3 Hypothetical scenarios for export of aliphatic wax components to the cuticle. (1) C16 and C18 fatty acyl-CoAs are elongated by the fatty acid elongase (FAE, shown as a square) in the endoplasmic reticulum (ER). Subsequent modifications of very long-chain acyl precursors into alkanes, primary or secondary alcohols, or ketones are carried out by unknown enzymes and are, therefore, not shown. From the ER, the VLCFAs or their derivatives could be transported to the plasma membrane (PM) by three possible routes: (2a) direct ER to PM transport mediated by unknown proteins (shown as a diamond), delivering lipid either directly to an ABC transporter (shown as a hexagon) or into the PM bilayer; (2b) unknown lipid binding proteins could mediate transfer of VLCFAs to the ABC transporter or into the PM bilayer; (2c) cuticular lipids could move along the endomembrane system from ER to Golgi and on to the PM, either free in the lipid bilayer or in ‘lipid rafts’. Once at the cell surface, wax components could be pulled out of the bilayer by ABC transporters (3) and either (4a) transferred directly through the cell wall or (4b) carried by non-specific lipid transfer proteins (LTPs) to the cuticle.

In contrast, when the A. thaliana CER10::GFP fusion was heterologously expressed in yeast, CER10 was localised only to one particular domain of the ER, the nuclear–vacuolar junction (Zheng et al., 2005). This finding is in very good accordance with previous reports showing that the S. cerevisiae VLCFA ECR is also localised in this ER subdomain (Kohlwein et al., 2001). In contrast, other components of the yeast FAE complex are located throughout the ER (and nuclear envelope). It may be speculated that the restriction of both native and foreign ECRs to one subdomain of the ER in yeast is required for the production of sphingolipid VLCFA precursors. To date, there is no evidence for a similar sub-compartmentation of lipid biosynthesis within the ER of plant epidermal cells, but the diverse pathways in this organelle might be spatially organised to a certain degree. The first committed steps of the pathways for the production of sphingolipids, cutin monomers and waxes presumably all co-occur in the epidermal ER. It remains to be determined

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Figure 5.4 Green fluorescent protein (GFP) experiments demonstrate the location of the proteins involved in wax production. These images represent a projection of optic sections collected with the laser scanning confocal microscope. The enoyl-CoA reductase (CER10-GFP) component of the elongase was found in the endoplasmic reticulum of pavement cells in the leaf of Arabidposis thaliana. ABC transporter CER5 was localised to the plasma membrane in A. thaliana stems (GFP-CER5). Magnification bar = 10 μm. This figure is produced in colour in the colour plate section, which follows page 249.

how the enzymes catalysing these branch point reactions compete for 16:0-CoA and 18:0-CoA as substrates, and how their activities are regulated in space and time. Following elongation, VLCFAs are modified by various alternative pathways to form aliphatic wax components (see Section 5.3). Very little is known about the cellular localisation of the enzymes catalysing the corresponding reactions. Thus, it cannot be assessed whether compounds leave the ER as very long-chain acyl CoA intermediates, or as downstream wax constituents. In one extreme scenario, all the modifying enzymes might be located in the ER, and the entire mixture of wax constituents would be generated there. In this case the ensuing steps in the export towards the cuticle would have to accommodate a wide range of compounds, and the mechanisms should have little specificity. In the other extreme, the very longchain acyl CoAs might be selectively trafficked from the ER to the PM for further modification and export. The fact that conventional sample preparation techniques for electron microscopy (EM) extract lipophilic compounds (Reed, 1982) has made studies in lipid transport very difficult. For this reason, the mechanisms for transport of wax molecules (intermediates or products) from the ER to the PM are currently unknown. However, a number of alternative hypotheses for this aspect of cuticle formation

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have been put forward (Figure 5.3), based either on circumstantial evidence or on analogies with other intracellular lipid transport processes (Kunst and Samuels, 2003; Schulz and Frommer, 2004). As one possibility, direct molecular transfer between both membranes could occur at sites where cortical ER is in close proximity to the PM (Figure 5.3, scenario 2a) (Staehelin, 1997). The second more speculative possibility is that small aggregates of wax components could also traffic from the ER to the PM, surrounded by oleosin-like proteins, similar to those surrounding oil bodies in oilseeds and the tapetum (Kim et al., 2002; Jolivet et al., 2004). Such small (>75 nm) lipid complexes might have eluded transmission electron microscopic investigations, because the structures are in the size range of one section thickness (70 nm typically) and also because the saturated components of the wax would not react with EM stains. As a third alternative, lipid-binding proteins similar to ACBPs have to be considered (Figure 5.3, scenario 2b), even though experimental evidence for their involvement is missing (see earlier). As a fourth possibility, saturated wax precursors could also be accommodated and transported by vesicles moving along the secretory pathway from ER to Golgi to the PM. As the VLCFA and their derivatives would influence the membrane fluidity and rigidity, they might be contained within specialised membrane microdomains, that is, ‘lipid rafts’. Based on their detergent-resistance, characteristic microdomains of membranes have been isolated from plant cells growing in cultures. These ‘lipid rafts’ were especially sphingolipid- and sterol-rich (Mongrand et al., 2004) and were associated with specific, often GPI-anchored, proteins (Borner et al., 2005). Furthermore, those fractions of the rafts were enriched for PM proteins (Borner et al., 2005). This finding was in good accordance with yeast and mammalian cells, where ‘lipid rafts’ have been postulated to function in sorting of membrane proteins and clustering of components involved in signalling (Brown and London, 2000; Mayor and Rao, 2004). It has been suggested that VLCFA-containing lipids, for example, sphingolipids, are transported in vesicles from the ER via the Golgi to the PM (Moreau et al., 1988). Similarly, if wax precursors or components associate with sterols and/or sphingolipids in the ‘lipid rafts’, then they could be transported to the PM by vesicle traffic involving the Golgi (Figure 5.3, scenario 2c). The epidermal cell structure of most plant species is not polarised, that is, the Golgi or the ER do not predominate under the periclinal surface of the cell in a way that would suggest a subcellular mechanism of handling and exporting the hydrophobic wax molecules. One exceptional case is the spectacular cork cells of Sorghum bicolor, which secrete long filaments of epicuticular wax. When studied with transmission EM, these cells have abundant ER on the periclinal side of the cytoplasm, facing the cuticle; however they contain few Golgi stacks (Jenks et al., 1994). Correlated with wax secretion, these specialised cork cells have large vesicles with dark contents called ‘osmiophilic globules’. Osmiophilic particles have also been observed in the cytoplasm of epidermal cells in deepwater rice during cuticle formation. They were observed fusing with the PM during cuticle synthesis and their production could be inhibited with monensin, indicating that a functional trans-Golgi structure is required for their synthesis (Hoffmann-Benning

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and Kende, 1994; Hoffmann-Benning et al., 1994). However, it is not clear what the ‘osmiophilic’ material represents. When probed with enzyme-gold particles and antibodies, the rice osmiophilic granules were found not to contain characteristic components of the epidermis surface, such as pectin or cutin. And while they tested positive with a proteinase-gold probe, proteins that might be involved in lipid export, such as lipid transfer proteins (LTPs), could not be detected (Hoffmann-Benning et al., 1994). Thus, even though their presence correlated with cuticle synthesis, direct positive evidence for the role of these osmiophilic structures in cuticle formation could not be provided. In summary, support for the hypothesis that wax components traffic through the ER–Golgi–PM route comes by analogy with sphingolipid transport and due to the presence of the apparently Golgi-derived but enigmatic osmiophilic vesicles. The argument against Golgi-mediated transfer is that there is no change in Golgi morphology such as increased number of vesicles associated with the stacks, as is typically observed during secretion.

5.6.2

Transport of wax through the PM

During the final step of intracellular trafficking, wax molecules (intermediates or products) have to enter into the PM. Again, there is currently no experimental evidence that would indicate the mechanisms involved in this process. Two fundamental possibilities can be distinguished – one involving direct loading of lipid molecules into specific PM proteins, and the other one involving initial partitioning of molecules into the membrane, followed by their association with proteins involved in further transport (Figure 5.3, scenario 3). These mechanisms might require energy for active detachment of lipid molecules from previous (protein or vesicle) complexes, and/or active loading onto PM proteins. One recent advancement in understanding the mechanism of wax export came from the identification of an A. thaliana ATP binding cassette (ABC) transporter, CER5 (Pighin et al., 2004). Evidence that this transporter is involved in wax export comes from the analysis of the cer5 mutant in which wax components are reduced on the cuticle surface but instead accumulate inside the cells (Pighin et al., 2004). Using a GFP tag, CER5 was localised to the PM of epidermal cells (Figure 5.4). The CER5 gene encodes an ABC transporter related by sequence homology to the human ABCG subfamily, a group of proteins that transport lipids (Lorkowski and Cullen, 2002), and the white–brown complex (WBC) of Drosophila melanogaster, which transports eye pigment precursors (Ewart et al., 1994). A survey of all genes containing ATP-binding cassettes in the A. thaliana genome identified the WBC-type subfamily as the largest single group with 29 members within this ABC transporter superfamily (Sanchez-Fernandez et al., 2001). The CER5 protein (annotated as WBC12 in the A. thaliana ABC transporter survey) is the first ABC transporter of the WBC subfamily to have a biological role assigned to it in plants. CER5 is a half-transporter, as its protein structure is predicted to contain one ATP binding cassette and transmembrane domain (TMD), rich in membrane spanning

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α-helices. In all prokaryotic and eukaryotic ABC transporters known, the functional ABC transporter consists of two ABC domains and two TMDs (Higgins and Linton, 2004). The binding partner for CER5 could be either another CER5 molecule to form a homodimer, or another ABC half-transporter to form a heterodimer. If CER5 is directly exporting wax components out of the cell, judging by the cer5 biochemical surface wax phenotype, it likely transports a variety of wax components. This is not unusual, as many ABC transporters have the ability to pump a wide variety of substrates. Previously, plant ABC transporters have been characterised primarily in heavy metal detoxification and secondary metabolite transport (Rea et al., 1998; Jasinski et al., 2001, 2003); however in mammals and bacteria, ABC transporters have been identified as lipid transporters in outer membrane lipid A export, in bile secretion, cholesterol homeostasis in macrophages, and exporters of β-sitosterol (Pohl et al., 2005). The ABC transporter CER5 is one candidate protein that could either directly extract lipids from cytosolic transport proteins, or indirectly accept them from within the PM. If the wax components are embedded in the PM, they could enter via a side port in the pore formed by the TMD, by analogy to lipid ABC transporters from bacteria and humans (Chang and Roth, 2001). For both the direct and the indirect mechanism, ATP hydrolysis can provide the energy required for loading molecules into the ABC transporter. Consequently, the next step in the process, that is, export into the cell wall, may be energetically promoted already at the stage of detaching molecules from the last intracellular transport unit, be it the PM or a cytosolic transport protein.

5.6.3

Transport of wax through the cell wall to the cuticle

Once wax components have been exported from the epidermal cells, they must traverse the cell wall. As the highly lipophilic wax molecules have very low solubility in the aqueous environment of the cell wall, this transport process has to be facilitated in some way. One possibility is that special structures exist in the periclinal cell wall of epidermal cells, characterised by relatively non-polar internal surfaces that help guide the wax molecules to the cuticle (Figure 5.3, scenario 4a). As such domains have not been characterised, wax transport through the cell wall has usually been attributed to lipid transfer proteins (Figure 5.3, scenario 4b) (Sterk et al., 1991; Moreau et al., 1998), although it must be stressed that there is also no direct experimental evidence confirming LTP participation in this process. In principle, LTPs are small enough to diffuse through the pectin matrix of the primary cell wall and the emerging data describing their properties make them interesting candidates. Non-specific LTPs (nsLTPs) of plants are part of a larger superfamily of small, basic proteins that move phospholipids between lipid bilayers in vitro (Kader, 1996; Arondel et al., 2000; Rogers and Bankaitis, 2000). A few representatives of the protein family have been localised in the epidermal cell wall (Thoma et al., 1993) as well as in the cuticle (Pyee et al., 1994). LTPs have been grouped into two classes, nsLTP1 (molecular weight, 9 kDa) and nsLTP2 (7 kDa) (Kader, 1996).

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NMR spectroscopy and X-ray crystallography have been used to determine nsLTP1 protein structures from rice, barley and wheat. They contain four α-helices, stabilised by disulfide bonds between eight conserved cysteines, surrounding a flexible hydrophobic pocket (Douliez et al., 2000a). Various techniques such as displacement of fluorescent analogues and intrinsic tyrosine fluorescence microscopy have been used to assess the interactions of LTPs with lipids. While nsLTP1 have been shown to bind to linear acyl lipids (Douliez et al., 2000b), including cutin components such as ω-hydroxyhexadecanoic acid (Douliez et al., 2000b, 2001), nsLTP2s showed lower affinity for fatty acids and phospholipids and higher affinity to sterols, and have been postulated to be important in plant defence (Blein et al., 2002; Cheng et al., 2004). None of these studies have tested the binding of VLCFAs or other wax components to LTPs. In summary, nsLTP1s are candidates for transport of cutin and wax monomers through the cell wall, but experiments directly testing their involvement have not been performed to date. When wax molecules finally reach the cuticular layer, they can easily dissolve into the wax domains there. In case they arrive there as complexes bound to LTPs, the hydrophobic cargo could partition into the cuticle, freeing an LTP for another cycle of delivery. Within the hydrophobic environment of the cuticle, wax components self-arrange into intracuticular layers, epicuticular films and epicuticular crystals (see Chapter 4, section 4.6).

5.7 Concluding remarks Formation of cuticular waxes requires two types of pathways: those involved in the biosynthesis of wax precursors and those for modifying them into diverse aliphatic lipid classes. Whereas cellular sites of production of VLCFA wax precursors and the biochemical reactions involved in this process have now been established, and at least one gene/enzyme for each step characterised, with the exception of the β-hydroxyacyl-CoA dehydratase, the majority of the proposed reactions leading to the formation of aliphatic wax components still await confirmation, and their subcellular location remains obscure. Information on the identity of the enzymes catalysing various steps of wax biosynthesis and their regulation, currently emerging from molecular and genomic studies in A. thaliana and maize, is essential to our overall understanding of cuticular wax deposition, and should provide a framework for rational modification of plant cuticles by genetic engineering to enhance the stress resistance of important agricultural commodities.

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Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

6

Optical properties of plant surfaces Erhard E. Pfündel, Giovanni Agati and Zoran G. Cerovic

6.1 Introduction Plant organs are composed of many tissue types containing specialised cells differing in chemical composition and structure. This complexity results in complicated and variable interactions between radiation and plant matter. Among plant organs, the optical behaviour of leaves is probably the most studied because of its importance to the photosynthetic process (Terashima, 1989; Björn, 1992; Vogelmann, 1993; Smith et al., 1997; Evans, 1999; Carter and Knapp, 2001; Ustin et al., 2001; Vogelmann and Evans, 2002; Evans and Vogelmann, 2003), and for the interpretation of remote sensing data (Buschmann and Nagel, 1993; Zwiggelaar, 1998; Carter and Knapp, 2001; Ustin et al., 2004). This chapter reviews the optical characteristics of plant surfaces. As all three authors of this review are interested in various aspects of photosynthesis in higher plants, we primarily focus on surface optics of green leaves but also briefly discuss the surface optics of fruits and flower petals. In this review, we define ‘plant surface’ as any peripheral layer with the potential to influence radiation conditions inside the photosynthetic mesophyll of leaves; that is, the cuticle, epidermis and, in some cases, sub-epidermal layers. We consider natural ultraviolet (UV) radiation, which comprises of UV-B (280–315 nm) and UV-A spectral range (315–400 nm), because UV radiation can not only drive photosynthesis (McLeod and Kanwisher, 1962; Mantha et al., 2001) but is also able to damage various components of the photosynthetic machinery (Day and Neale, 2002; Hollósy, 2002; Jordan, 2002; Kakani et al., 2003). Naturally, we also discuss surface optics in the visible wavelength range (400–700 nm) which is the main energy source for photosynthesis: this spectral range can also damage components of the photosynthetic machinery if light energy is absorbed in excess of photosynthetic capacity (Osmond et al., 1997; Huner et al., 1998; Niyogi, 2000; Krieger-Liszkay, 2005). In this review, we restrict the use of the term ‘light’ to visible radiation only. We briefly describe current methods for analysing surface optics and then discuss, in detail, not only optical surface properties arising from electronic absorption of radiation by pigments but also those resulting from non-absorptive interaction of radiation with matter: plant surface optics, of course, integrates both of these optical aspects. Rather than provide complete coverage of the literature we have opted to concentrate on several especially interesting facets of surface optics in plants, and we apologise to those workers whose important works have not been cited. Consistent

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with our restricted focus, we refer to relevant classical papers and recent papers but also include a number of reviews containing broader and more extensive literature compilations.

6.2 Methods to determine optical properties of plant surfaces 6.2.1

UV-visible absorbance spectrophotometry

Solutions of (epi)cuticular matter obtained by short-term extraction of plant surfaces with chloroform, or aqueous/methanolic extracts of epidermal strips peeled off plant organs are often measured spectrophotometrically to assess absorption properties of plant surfaces (see Section 6.3.1). These measurements are suitable for classifying the principal absorbing molecules present, and to reveal changes in their concentration, for example, during acclimation processes. Because optical conditions within plant surfaces often differ fundamentally from those in solutions, spectrophotometric data of extracts should not be used uncritically to evaluate the absorption of these compounds in vivo. Stripping off the epidermis is not possible in many plant species; consequently, whole leaf extracts have been analysed spectrophotometrically to identify and classify epidermal UV-screening compounds (for a review see Searles et al., 2001). The limitations of this method obviously include not only the different optical behaviour of pigments in solutions and in vivo, but also the difficulty of allocating variations in concentration of UV-absorbing molecules between upper and lower epidermis and leaf mesophyll (Kolb and Pfündel, 2005). UV-Vis spectrophotometry has also been employed to analyse mechanically isolated epidermal strips or enzymatically prepared cuticles (see Section 6.3.1). Because isolation and subsequent handling can disrupt these layers, such spectroscopic studies should be accompanied by parallel microscopic investigations to confirm the intactness of the preparations. Furthermore, these layers scatter the measuring beam so that a part of the transmitted radiation, leaving the sample at oblique angles, misses the photodetector in standard spectrophotometers; therefore, spectrophotometric measurements of scattering samples tend to underestimate the sample transmittance. It is possible to reduce this error by placing the sample close to the photodetector, thereby increasing the angle of acceptance, and also by completely diffusing all radiation transmitted by the sample with an opal quartz plate (Butler, 1964). On the other hand, fluorescence excited by the measuring radiation might result in overestimation of transmittance particularly with highly absorbing samples.

6.2.2

Integrating sphere

Integrating spheres are hollow globes with highly reflective inner surfaces. In the so-called ‘external’ integrating sphere, the sample forms a defined area of the inner

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surface. Also, an aperture through which photons can reach the detector is located on the inner surface. Orientation of the sample relative to the aperture is arranged to prevent photons originating in the sample reaching the photodetector directly, but photons at all angles of emergence can reach the detector after multiple reflection with comparable probabilities. Therefore, integrating spheres realise a very large acceptance angle and, hence, overcome the problem of photons missed by scattering in normal spectrophotometers. In principle, transmittance of samples is calculated from the detector signal obtained with the interior illuminated through the sample, divided by the signal measured with direct illumination of the inner sphere. For reflection measurements, the sample is flipped over and illuminated from the inside of the sphere: by relating the signal obtained under these latter conditions with that recorded from a highly reflecting reference surface a value for reflectance is obtained. Absorptance of a sample is calculated according to: absorptance (%) = 100 (%) − reflectance (%) − transmittance (%)

(6.1)

Equation 6.1 illustrates that reflectance, which determines the optical appearance of plant organs viewed from the illuminated side, is determined by absorption and transmission properties. We emphasise that absorptance is not to be confused with absorbance: in the absence of reflection, the logarithmic relationship between these two terms is given by: absorbance = log{100/[100 − absorptance (%)]}

(6.2)

Integrating spheres are frequently employed to study intact leaves. At wavelengths of high electronic absorption by the leaf, information on surface reflectance can be derived from intact material (see Section 6.4.1). With plant species allowing the preparation of sufficiently large epidermal patches, optical properties of plant surfaces can be studied over the entire wavelength range of interest (Figure 6.1; Grammatikopoulos et al., 1999; and references in Section 6.4.1).

6.2.3

Microfibre optics

Information on plant surface optics is also available from measurement of gradients of radiation intensities inside leaves using fibre microprobes (Vogelmann and Björn, 1984; Vogelmann and Haupt, 1985). These gradients are obtained with a fibre optics probe having a thickness of a few micrometres by advancing it stepwise through the tissue and recording the radiation accepted by the probe using a spectroradiometer. The steepness of intensity curves below the illuminated surface and the depth of penetration of the radiation are both key parameters from which absorption properties of surfaces can be derived. This technique has been useful in obtaining information on UV screening of leaf surfaces (Bornman and Vogelmann, 1988; Bornman and Vogelmann, 1991; Day et al., 1992; Cen and Bornmann, 1993; Ålenius et al., 1995; Turunen et al., 1999; Olsson et al., 2000; Liakoura et al., 2003).

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(a) 100

0 Reflectance

40

tance

40 Intact leaf Mesophyll Epidermis

Transm it

Transmittance, %

60

60

Reflectance, %

20

80

80

20 Transmittance

100

0 (b) 100 Mesophyll

Absorptance, %

80

60

Intact leaf

40

Epidermis

20

0 400

500

600

700

Wavelength , nm Figure 6.1 Some optical properties of leaf preparations from Tulipa spec. Reflectance and transmittance spectra (a), and absorptance spectra, calculated from the former according to Equation 6.1, (b) were recorded using whole leaves (labelled: intact leaf), leaves with the epidermis stripped off (mesophyll) and epidermal strips (epidermis). Each spectrum corresponds to the mean of four independent measurements of outdoor-grown material recorded with an external integrating sphere (1800-12 integrating sphere, Li-Cor, Lincoln, Nebraska) in combination with an LI-1800 portable spectrometer (Li-Cor). Panel (a) illustrates important aspects of surface optics: (1) removal of the epidermis decreases reflectance at wavelengths greater than 400 nm; (2) reflectance of intact leaves and epidermis is lowest below 400 nm; and (3) reflectance of the intact leaf and epidermis coincide when intact leaf or mesophyll transmittance is minimal between 350 and 480 nm and near 680 nm (see Section 6.4.1 for a discussion of these phenomena). Epidermal transmittance (a) is minimal below 400 nm, which is almost certainly due to the presence of colourless flavonoids located in epidermal cell vacuoles (see Section 6.3.1). The calculated absorptance spectra (b) show that the mesophyll absorbs radiation better than the intact leaf at wavelengths greater than 400 nm, and also, show efficient epidermal absorbance at wavelengths lesser than 400 nm (Jana Leide and E.E. Pfündel, unpublished data).

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UV-excited chlorophyll fluorescence

UV fluorimetry allows the assessment of epidermal absorptance of intact leaves in the UV spectral range. The basis of the method relies on the efficient absorption of UV by chlorophyll (Cerovic et al., 1999) and on the fact that a certain fraction of leaf chlorophylls, excited by absorption of radiation, emits red fluorescence when returning to the ground state (Dau, 1994). In intact leaves, UV excitation is reduced when epidermal UV screening is high and, under such conditions, relatively weak chlorophyll fluorescence is observed (Cerovic et al., 1993; Sheahan, 1996; Mazza et al., 2000). Bilger et al. (1997) introduced the ratio of UV-excited fluorescence to that excited by blue–green light as an accurate measurement for epidermal UV screening with the rationale that visible radiation is not absorbed by the epidermis but fluorescence excitation in the visible and in the UV is subject to the same optical peculiarities of the individual sample. Consequently, this fluorescence excitation ratio (FER) method cancels out specific optical properties of the sample except that of epidermal UV screening. Recently, two portable devices have been developed: the Dualex leaf clip (Goulas et al., 2004) and the UV-A-PAM fluorimeter (Bilger et al., 2001, Krause et al., 2003; Kolb et al., 2003, 2005). Both permit repeated measurements of epidermal UV screening of the same sample under field conditions. The Bilger approach has been strongly supported by the parallel relationship observed between UV transmittance data from spectrophotometric measurements of isolated epidermal strips and FER data (Markstädter et al., 2001). The formation of anthocyanins, however, interferes with excitation by blue light and can, therefore, restrict the use of fluorimetry in red leaves (Barnes et al., 2000). This problem can be overcome by choosing another reference excitation wavelength in the red (650 nm in Dualex), which is outside the anthocyanin absorption range (Goulas et al., 2004). In addition, fluorescence excited at wavelengths within the anthocyanin absorption range when divided by fluorescence excited at wavelengths of high epidermal transparency can also be used to determine the epidermal screening due to anthocyanins in vivo (Agati et al., 2005).

6.2.5

Fluorescence microscopy

Fluorescence microscopy can complement data obtained with the earlier techniques by obtaining two- and even three-dimensional information on the arrangement of structures and compounds of plant surfaces. This technique can localise absorbing substances by fluorescence: such data were useful in providing an understanding of the origin of fluorescence signals of entire leaves and, also, for the interpretation of remote fluoro-sensing data. Epi-fluorescence microscopy is widely used to analyse leaf tissues and it has been much improved in the last decade due to progress in digital imaging, confocal laser scanning and multiphoton microscopy (Blancaflor and Gilroy, 2000). Increased performance of personal computers permitted the application of image restoration

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221

by algorithms to remove out-of-focus signals in wide-field fluorescence microscopy (McNally et al., 1999). Compared to wide-field fluorescence microscopy, which works well with relatively thin specimen ( 42 kJ mol−1 ). Water transport mediated in a hydrophilic environment mediated by polar pores penetrating a lipid membrane is characterised by a comparably low activation energy (Ea < 25 kJ mol−1 ) and the activation energy for the self-diffusion of water amounts to 19 kJ mol−1 (Elmoazzen et al., 2002). The low activation energy of cuticular water permeability at temperatures below the phase transition can be taken as further evidence that water movement preferably takes place in a more or less aqueous environment. The steep increase of cuticular permeances above the phase transition temperature is probably associated with structural changes of the cuticular waxes, since for polymer matrix membranes no phase transition in this temperature range was observed (Schönherr et al., 1979; Schreiber, 2002). Correspondingly, the regression lines of Arrhenius plots for the water permeance of cuticular and matrix membranes tend to converge at high temperatures. The intersection of both lines represents

303

CUTICULAR TRANSPIRATION –8 Hedera helix Camellia sinensis Pyrus communis Liriodendron tulipifera

ln PCM [m s–1]

–9 –10 –11 –12 –13 –14 3.0

3.1

3.2

3.3

3.4

3.5

T –1 × 1000 [K–1]

Figure 9.2 Arrhenius plots of cuticular water permeability (recalculated from Riederer and Schreiber, 2001). The natural logarithm of the vapour-phase-based cuticular permeance (PCM ) is plotted versus the reciprocal value of the absolute temperature (T ). Table 9.7 Phase transition temperature (TP ) and activation energy (Ea ) for the permeation of water across isolated cuticular membranes (Riederer and Schreiber, 2001) Species Hedera helix Camellia sinensis Pyrus communis Liriodendron tulipifera

TP [˚C]

Ea [kJ mol−1 ] below TP

Ea [kJ mol−1 ] above TP

37 36 32 38

28 25 18 22

97 87 76 98

the temperature where the effectiveness of the wax barrier is eliminated. This effect occurs for cuticular membranes from leaves of Juglans regia at a temperature of 69˚C (Figure 9.3). This value is in good correspondence with the temperature range (66–74˚C) where complete melting of the cuticular wax was observed by visible detection using a melting point microscope (Schreiber and Riederer, 1996). In addition to the onset of wax melting at the phase transition temperature (Merk et al., 1998), an increased volume expansion of the cutin polymer was assumed to cause defects in the wax barrier, which may contribute to an enhancement of the cuticular water permeability with increasing temperature (Schreiber and Schönherr, 1990).

9.4 Physiology of cuticular transpiration in relation to stomatal closure So far, this chapter has concentrated on the water permeability of the plant cuticle. The cuticle, however, is part of a system to which the stomata also belong. Therefore,

304

BIOLOGY OF THE PLANT CUTICLE

–5 MX

–6

ln P [m s–1]

–7 –8 –9 –10

CM

–11 3.0

3.1

3.2

3.3

–1

–1

T

3.4

3.5

× 1000 [K ]

Figure 9.3 Arrhenius plots for the cuticular membrane (CM) and the polymer matrix membrane (MX) of leaves from Juglans regia. The natural logarithm of the permeance (P ) is plotted versus the reciprocal value of the absolute temperature (T ) (Burghardt, unpublished results).

it should be asked what role the cuticle plays in this system as far as drought resistance of the leaf or the whole plant is concerned. Minimum leaf conductances at maximum stomatal closure can be obtained from leaf drying curves by measuring the mass loss of detached leaves as a function of time under controlled environmental conditions. Since even a low fraction of open stomatal pore area may contribute significantly to the overall transpiration rate, a clear differentiation between cuticular permeances of stomata-free systems (adaxial leaf surfaces of hypostomatous leaves) and minimum conductances should be made (Kersties, 1996a,b). Transpiration rates of detached leaves are closely related to the leaf water potential and the minimum transpiration rate is reached only at the turgor loss point, which can be considered as the main indicator for maximum stomatal closure (Burghardt and Riederer, 2003). The minimum transpiration rate remains nearly constant up to a relative water deficit equivalent to the symplastic water content (Figure 9.4). In some cases a slight decrease of the minimum conductance with declining leaf water content has been observed, which may be attributed to a decrease of the hydration state of the cuticular membrane itself (Van Gardingen and Grace,1992; Hoad et al., 1996). For a number of species the cuticular permeances of the adaxial leaf surfaces and the minimum conductances of leaves detached from the whole plant were not significantly different (Figure 9.5). H. helix is the only exception since minimum conductance is three-fold higher than cuticular permeance (Burghardt and Riederer, 2003). This deviation may be caused by residual transpiration due to incomplete stomatal closure. Model calculations show that stomata may contribute significantly to the minimum conductance even when stomatal pores visibly appear completely

305

CUTICULAR TRANSPIRATION

(a)

3.5

Leaf drying curve MX

J × 103 [g m–2 s–1]

3.0 2.5 2.0 1.5 1.0

CM

0.5 0.0 (b)

5

Pressure–volume curve

–Ψ–1 [MPa–1]

4

3

2

1

0 0.0

0.2

0.4

0.6

0.8

RWD Figure 9.4 (a) Representative leaf drying curve for Vinca minor. The transpiration rate (J ) is plotted versus the relative water deficit (RWD). (b) Pressure–volume curve of V. minor. The reciprocal value of the leaf water potential (Ψ) is plotted versus the relative water deficit (RWD) (data from Burghardt and Riederer, 2003, and unpublished results).

closed (Kerstiens, 1996b). An alternative explanation was recently offered by the development of a new technique for the measurement of water permeability of stomatous cuticular membranes. In this study the cuticular water permeability of the abaxial side of leaves from H. helix was measured to be eleven-fold higher than the permeability of the adaxial side (Santrucek et al., 2004). This finding may be related to early reports that the cuticle covering the guard cells may have a higher water permeability (‘peristomatal transpiration’) than the cuticle of the epidermis (Maier-Maercker, 1983).

306

BIOLOGY OF THE PLANT CUTICLE

–4.0

log gmin [m s–1]

Quercus

Fagus Juglans Vinca Nerium

Acer

–4.5

–5.0

Hedera Ilex

–5.5 –6.0

–5.5

–5.0

–4.5

–4.0

log PCM [m s–1] Figure 9.5 Plot of the minimum conductance (gmin ) versus the cuticular permeance (PCM ) (Acer campestre, Fagus sylvatica, Quercus petraea, Ilex aquifolium, Hedera helix: Burghardt and Riederer, 2003; and Juglans regia, Nerium oleander, Vinca minor, unpublished results).

Though under extreme experimental conditions (0% relative humidity, darkness, leaf water potential below the turgor loss point) residual stomatal transpiration does not contribute significantly to the overall transpiration of detached leaves, the question remains open, if, under realistic environmental conditions in the field, attached leaves of intact plant behave in the same way. The lowest conductances of plants observed in the field (Körner, 1994) are generally higher than minimum conductances and cuticular permeances (Kerstiens, 1996a,b). Comparison of daytime and nighttime conductances indicate that some species maintain substantial stomatal conductances at night (Snyder et al., 2003). This implies that under typical environmental conditions plants are only rarely forced to a strict drought avoidance strategy. However, so far there is no study available considering cuticular permeances, minimum conductances and lowest conductances under controlled experimental conditions in an integrative approach.

9.5 The cuticular transpiration barrier as a mechanism of the drought avoidance strategy A survey of cuticular permeances of a broad spectrum of plant species has been published recently (Riederer and Schreiber, 2001). Cuticular permeances of fruits (median: 8.6 × 10−5 m s−1 ) were on the average 15-fold higher than permeances of leaves (median: 5.8×10−6 m s−1 ). Analysis of cuticular water permeability of leaves with regard to lifeform and climate of origin shows that the highest permeances are

CUTICULAR TRANSPIRATION

307

found within the group of mesomorphic leaves of deciduous species growing in temperate climates. Cuticles from evergreen leaves from epiphytic plants growing in tropical climates have the lowest permeances. However, the classification is not conclusive in all cases. The group of xeromorphic leaves of evergreen species growing in mostly mediterranean non-tropical climates has cuticular permeances ranging from 10−4 to 10−6 m s−1 , which covers nearly the whole spectrum of leaf cuticular permeances measured so far. Since cuticular transpiration was generally examined at 25˚C and at 0% relative humidity, a more differentiated picture may emerge, if water permeability is measured under environmental conditions which mimic the real climatic conditions of the natural habitats. If the water supply from the soil is restricted, plants reduce the transpirational water loss by midday or even by permanent stomatal closure (Larcher, 1972). Under such conditions survival depends on the efficacy of the transpiration barrier to conserve the scarce water resources and to counteract the inevitable decline of the leaf water content. The survival time has been introduced as a useful parameter in order to assess the ability of leaves to avoid drought stress (Pisek and Winkler, 1953): Survival time =

(RWDSLD − RWDSC )SU Jmin

(9.15)

The available water content depends on the sublethal water deficit (RWDSLD , relative water deficit where irreversible leaf damage occurs), the relative water deficit at maximum stomatal closure (RWDSC ) and the degree of succulence (SU , total amount of water per leaf area). The minimum transpiration rate (Jmin ) is given according to Equation 9.2 by the cuticular permeance (or minimum conductance) and the driving force of transpiration. Model calculations for 50% relative humidity and 25˚C (Figure 9.6) yield survival times in the range from 12 (Fagus sylvatica) to 19 h (Acer campestre) for deciduous leaves growing in temperate climates and 51 h for the xeromorphic leaf of Nerium oleander growing in mediterranean-type climates. The highest survival times with 120 h (H. helix) and 130 h (Ilex aquifolium) are obtained for evergreen leaves growing in temperate climates (Figure 9.6; Burghardt and Riederer, 2003).

9.6 Conclusions The mechanism of water transport in cuticular membranes is still a matter of debate. Co-permeation of water with lipophilic organic compounds has been interpreted as evidence that water movement predominately takes place across the lipophilic cuticular pathway (Schönherr and Riederer, 1989; Niederl et al., 1998; Schreiber, 2002). However, permeation of water does not fit the prediction models established for lipophilic organic compounds according to the mechanism of a sorption–diffusion process and the free-volume theory (see Section 9.2.1 earlier). Further studies on a quantitative basis are necessary in order to decide if water permeation rather is related to the lipophilic cutin–wax region or primarily to aqueous polar pores as it

308

BIOLOGY OF THE PLANT CUTICLE

(a)

7 6

gmin [m s–1]

5 4 3 2 1 (b)

0 140

Survival time (h)

120 100 80 60 40 20 Ile x

H

ed

er a

m N

er

iu

er Ac

ns la

us Fa g

Ju g

Q

ue

rc

us

0

Figure 9.6 (a) Minimum conductance (gmin ) and (b) survival time of deciduous leaves (Quercus petraea, Fagus sylvatica, Juglans regia, Acer campestre) and evergreen leaves (Hedera helix, Ilex aquifolium) growing in temperate climates and the xeromorphic leaf of Nerium oleander growing in mediterranean-type climates (data from Burghardt and Riederer, 2003, and unpublished results).

was described for calcium salts (Schönherr, 2000). This chapter presents evidence from diverse experimental approaches that the predominance of the latter pathway is probable. A further open question is how the contribution of the lipophilic and hydrophilic pathway to the overall water permeance of the cuticle varies among different plant species. From a review of pooled data of plants growing in different climatic regions it was concluded that complete stomatal closure is rather an exception (Körner, 1994; Kerstiens, 1996a,b). Therefore, the role of the cuticular transpiration barrier as an adaptive strategy with regard to drought avoidance is restricted under

CUTICULAR TRANSPIRATION

309

standard growing conditions except for astomatous fruits. The relative importance of the cuticular transpiration barrier increases when stomatal closure increases due to turgor loss under prolonged drought conditions. Therefore, leaf cuticular permeances, minimum conductances and lowest conductances should be compared in a coordinated experimental approach in order to assess under which environmental conditions complete stomatal closure is achieved and to what extent cuticular transpiration and residual stomatal transpiration contribute to the overall transpiration in times of water stress.

References Abraham, M.H. and McGowan, J.C. (1987) The use of characteristic volumes to measure cavity terms in reversed phased liquid chromatography, Chromatographia, 23, 243–246. Becker, M., Kerstiens, G. and Schönherr, J. (1986) Water permeability of plant cuticles: permeance, diffusion and partition coefficients, Trees Structure and Function, 1, 54–60. Beyer, M., Lau, S. and Knoche, M. (2005) Studies on water transport through the sweet cherry fruit surface: lX. Comparing permeability in water uptake and transpiration, Planta, 220, 474–485. Burghardt, M. and Riederer, M. (2003) Ecophysiological relevance of cuticular transpiration of deciduous and evergreen plants in relation to stomatal closure and leaf water potential, Journal of Experimental Botany, 54, 1941–1949. Dominguez, E. and Heredia, A. (1999) Water hydration in cutinised cell walls: a physico-chemical analysis, Biochimica et Biophysica Acta, 1426, 168–176. Elmoazzen, H.Y., Elliott, J.A.W. and Mc Gann, L.E. (2002) The effect of temperature on membrane hydraulic conductivity, Cryobiology, 45, 68–79. Geyer, U. and Schönherr, J. (1990) The effect of the environment on the permeability and composition of Citrus leaf cuticles. l. Water permeability of isolated cuticular membranes, Planta, 180, 147–152. Hauke, V. and Schreiber, L. (1998) Ontogenetic and seasonal development of wax composition and cuticular transpiration of ivy (Hedera helix L.) sun and shade leaves, Planta, 207, 67–75. Hoad, S.P., Grace, J. and Jeffree, C.E. (1996) A leaf disc method for measuring cuticular conductance, Journal of Experimental Botany, 47, 431–437. Hoad, S.P., Grace, J. and Jeffree, C.E. (1997) Humidity response of cuticular conductances of beech (Fagus sylvatica L.) leaf discs maintained at high relative water content, Journal of Experimental Botany, 48, 1969–1975. Jeffree, C.E. (1996) Structure and ontogeny of plant cuticles, in Plant Cuticles: an Integrated Functional Approach (ed. G. Kerstiens), Bios Scientific Publishers Ltd., Oxford, pp. 33–82. Kerler, F. and Schönherr, J. (1988a) Accumulation of lipophilic chemicals in plant cuticles: prediction from octanol/water partition coefficients, Archives of Environmental Contamination and Toxicology, 17, 1–6. Kerler, F. and Schönherr, J. (1988b) Permeation of lipophilic chemicals across plant cuticles: prediction from partition coefficients and molar volumes, Archives of Environmental Contamination and Toxicology, 17, 7–12. Kerstiens, G. (1994) Air pollutants and plant cuticles: mechanisms of gas and water transport, and effects on water permeability, in Air Pollutants and the Leaf Cuticle (eds K.E. Percy, J.N. Cape, R. Jagels and C.J. Simpson), Springer-Verlag, Berlin, pp. 39–52. Kerstiens, G. (1996a) Diffusion of water vapour and gases across cuticles and through stomatal pores presumed closed, in Plant Cuticles: an Integrated Functional Approach (ed. G. Kerstiens), Bios Scientific Publishers Ltd., Oxford, pp. 121–134.

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Kerstiens, G. (1996b) Cuticular water permeability and its physiological significance, Journal of Experimental Botany, 47, 1813–1832. Kerstiens, G. and Lendzian, K.J. (1989) Interactions between ozone and plant cuticles. ll. Water permeability, New Phytologist, 112, 21–27. Knoche, M., Peschel, S., Hinz, M. and Bukovac, M.J. (2000) Studies on water transport through the sweet cherry fruit surface: characterisation conductance of the cuticular membrane using pericarp segments, Planta, 212, 127–135. Körner, C. (1994) Leaf diffusive conductances in the major vegetation types of the globe, in Ecophysiology of Photosynthesis. Ecological Studies 100 (eds E.D. Schulze and M.M. Caldwell), Springer-Verlag, Berlin, pp. 463–490. Larcher, W. (1972) Physiological Plant Ecology, Springer-Verlag, Berlin. Lendzian, K.J. and Kerstiens, G. (1991) Sorption and transport of gases and vapors in plant cuticles, Reviews of Environmental Contamination and Toxicology, 121, 65–128. Luque, P., Gavara, R. and Herédia, A. (1995) A study of the hydration process of isolated cuticular membranes, New Phytologist, 129, 283–288. Maier-Maercker, U. (1983) The role of peristomatal transpiration in the mechanism of stomatal movement, Plant, Cell and Environment, 6, 369–380. Marga, F., Pesacreta, T.C. and Hasenstein, K.H. (2001) Biochemical analysis of elastic and rigid cuticles of Cirsium horridulum, Planta, 213, 841–848. Merk, S., Blume, A. and Riederer, M. (1998) Phase behaviour and crystallinity of plant cuticular waxes studied by Fourier transform infrared spectroscopy, Planta, 204, 44–53. Mitragotri, S. (2003) Modeling skin permeability to hydrophilic and hydrophobic solutes based on four permeation pathways, Journal of Controlled Release, 86, 69–92. Niederl, S., Kirsch, T., Riederer, M. and Schreiber, L. (1998) Co-permeability of 3 H-labeled water and 14 C-labeled organic acids across isolated plant cuticles, Plant Physiology, 116, 117–123. Nobel, P.S. (1991) Physicochemical and Environmental Plant Physiology, Academic Press, London. Pearcy, R.W., Ehleringer, J.R., Mooney, H.A. and Rundel, P.W. (1991) Plant Physiology Ecology, Chapman and Hall, London. Pisek, A. and Winkler, E. (1953) Die Schliessbewegung der Stomata bei ökologisch verschiedenen Pflanzentypen in Abhängigkeit vom Wassersättigungszustand der Blätter und vom Licht, Planta, 42, 253–278. Riederer, M. and Schneider, G. (1990) The effect of the environment on the permeability and composition of Citrus leaf cuticles. ll. Composition of soluble cuticular lipids and correlation with transport properties, Planta, 180, 154–165. Riederer, M. and Schönherr, J. (1984) Accumulation and transport of (2,4-dichlorophenoxy)acetic acid in plant cuticles: l. Sorption in the cuticular membrane and its components, Ecotoxicological and Environmental Safety, 8, 236–247. Riederer, M. and Schönherr, J. (1990) Effects of surfactants on water permeability of isolated plant cuticles and on the composition of their cuticular waxes, Pesticide Science, 29, 85–94. Riederer, M. and Schreiber, L. (1995) Waxes – the transport barriers of plant cuticles, in Waxes. Chemistry, Molecular Biology and Functions (ed. R.J. Hamilton), The Oily Press, West Ferry, pp. 131–156. Riederer, M. and Schreiber, L. (2001) Protecting against water loss: analysis of the barrier properties of plant cuticles, Journal of Experimental Botany, 52, 2023–2032. Santrucek, J., Simanova, E., Karbulkova, J., Simkova, M. and Schreiber, L. (2004) A new technique for measurement of water permeability of stomatous cuticular membranes isolated from Hedera helix leaves, Journal of Experimental Botany, 55, 1411–1422. Schönherr, J. (1976a) Water permeability of isolated cuticular membranes: the effect of pH and cations on diffusion, hydrodynamic permeability and size of polar pores in the cutin matrix, Planta, 128, 113–126. Schönherr, J. (1976b) Water permeability of isolated cuticular membranes: the effect of cuticular waxes on diffusion of water, Planta, 131, 159–164. Schönherr, J. (2000) Calcium chloride penetrates plant cuticles via aqueous pores, Planta, 212, 112–118.

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Schönherr, J. and Baur, P. (1994) Modelling penetration of plant cuticles by crop protection agents and effects of adjuvants on their rates of penetration, Pesticide Science, 42, 185–208. Schönherr, J. and Huber, R. (1977) Plant cuticles are polyelectrolytes with isoelectric points around three Plant Physiology, 59, 145–150. Schönherr, J. and Lendzian, K. (1981) A simple and inexpensive method for measuring water permeability of isolated plant cuticular membranes, Zeitschrift für Pflanzenphysiologie, 102, 321–327. Schönherr, J. and Riederer, M. (1989) Foliar penetration and accumulation of organic chemicals in plant cuticles, Reviews of Environmental Contamination and Toxicology, 108, 1–70. Schönherr, J. and Schmidt, H.W. (1979) Water permeability of plant cuticles. Dependence of permeability coefficients of cuticular transpiration on vapor pressure saturation deficit, Planta, 144, 391–400. Schönher, J., Eckl, K. and Gruler, H. (1979) Water permeability of plant cuticles: the effect of temperature on diffusion of water, Planta, 147, 21–26. Schreiber, L. (2001) Effect of temperature on cuticular transpiration of isolated cuticular membranes and leaf discs, Journal of Experimental Botany, 52, 1893–1900. Schreiber, L. (2002) Co-permeability of 3 H-labelled water and 14 C-labelled organic acids across isolated Prunus laurocerasus cuticles: effect of temperature on cuticular paths of diffusion, Plant, Cell and Environment, 25, 1087–1094. Schreiber, L. and Riederer, M. (1996) Determination of diffusion coefficients of octadecanoic acid in isolated cuticular waxes and their relationship to cuticular water permeabilities, Plant, Cell and Environment, 19, 1075–1082. Schreiber, L. and Schönherr, J. (1990) Phase transitions and thermal expansion coefficients of plant cuticles, Planta, 182, 186–193. Schreiber, L., Skrabs, M., Hartmann, K.D., Diamantopoulos, P., Simanova, E. and Santrucek, J. (2001) Effect of humidity on cuticular water permeability of isolated cuticular membranes and leaf discs, Planta, 214, 274–282. Snyder, K.A., Richards, J.H. and Donovan, L.A. (2003) Night-time conductance in C3 and C4 species: do plants lose water at night? Journal of Experimental Botany, 54, 861–865. Van Gardingen, P.R. and Grace, J. (1992) Vapour pressure deficit response of cuticular conductance in intact leaves of Fagus sylvatica L., Journal of Experimental Botany, 43, 1293–1299. Verkman, A.S. (2000) Water permeability measurement in living cells and complex tissues. Journal of Membrane Biology, 173, 73–87. Vogg, G., Fischer, S., Leide, J. et al. (2004) Tomato fruit cuticular waxes and their effects on transpiration barrier properties: functional characterization of a mutant deficient in a very-long-chain fatty acid β-ketoacyl-CoA synthase, Journal of Experimental Botany, 55, 1401–1410.

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

10 The cuticle and cellular interactions Hirokazu Tanaka and Yasunori Machida

10.1

Introduction

The epidermis of the aerial parts of plants plays an important role in water retention (Chapter 9), defence against pathogens (Chapters 11, 12 and 13) and gas exchange. The cuticle covering the epidermis is believed to be important for such epidermal functions and to protect the plant from the outer environment. In addition, genetic as well as biochemical studies have suggested that the cuticle is also essential for proper organ formation and fertilisation. In these processes, the cuticle appears to be involved, either positively or negatively, in communication between plant cells. In this chapter, advances in our understanding of the roles of the cuticle in such cellular communications and the genes required for proper cuticle generation are described.

10.2

Essential roles of the cuticle in post-embryonic development

The aerial parts of plants are mainly derived from the shoot meristem, which maintains itself and successively generates organ primordia. In the shoot meristem and the floral meristem, boundaries between primordia for lateral organs such as leaves and the floral organs are established (for a review see Fleming, 2005). Subsequently, despite contact between adjacent organ primordia during their growth, each primordium remains separate. In the past decade, identification and characterisation of mutants and responsible genes have shed light on the mechanisms underlying this prevention of lateral organ fusion during development.

10.2.1 The cuticle is required for prevention of organ fusion To date, a number of mutants that produce fused organs have been described in Arabidopsis thaliana (Table 10.1), maize and other plants. In this section, the recent progress in identifying A. thaliana mutants and their phenotypic characterisation will be reviewed. Lolle et al. (1992) reported the characterisation of the fiddlehead ( fdh) mutant, which generates fused floral organs and leaves. Histological analysis indicated

313

THE CUTICLE AND CELLULAR INTERACTIONS Table 10.1

Arabidopsis thaliana mutants producing fused organs

Mutant name

abnormal leaf-shape1 acetyl-CoA carboxylase1/ pasticinno3/gurke airhead arabidopsis homologue of Crinkly4 bulkhead conehead deadhead eceriferum10 fiddlehead hothead/adhesion of caryx edges lacerata permeable leaves1 permeable leaves2 permeable leaves3 pothead thunderhead wax1 wax2/yore-yore/pel6

Abbreviation

Locus id or chromosome

References∗

Organ fusion Leaves

Flowers

Ovules

ale1 acc1/pas3/gk

At1g62340 At1g36160

Yes Yes

No Yes

No N.D.

12, 15 1, 2, 4, 16

ahd acr4

chr. 5 At3g59420

No No

Yes No

No Yes

10 3, 5, 13, 17

bud cod ded cer10 fdh hth/ace

chr. 1 chr. 2 chr. 1 chr. 3 At2g26250 At1g72970

No Yes Yes No Yes No

Yes Yes Yes Yes Yes Yes

Yes No Yes No Yes Yes

10 10 10 10 9, 11, 19 7, 10

lcr pel1 pel2 pel3 phd thd wax1 wax2

At2g45970 chr. 1 chr. 1 chr. 5 chr. 5 chr. 3 N.D. At5g57800

Yes Yes Yes Yes No Yes Yes Yes

Yes Yes Yes Yes Yes Yes Yes Yes

N.D. N.D. N.D. N.D. No Yes N.D. N.D.

18 14, 15 14, 15 14, 15 10 10 6 6, 8, 14

Notes: N.D., not determined. ∗ 1. Baud et al. (2003), 2. Baud et al. (2004), 3. Cao et al. (2005), 4. Faure et al. (1998), 5. Gifford et al. (2003), 6. Jenks et al. (1996), 7. Krolikowski et al. (2003), 8. Kurata et al. (2003), 9. Lolle et al. (1992), 10. Lolle et al. (1998), 11. Pruitt et al. (2000), 12. Tanaka et al. (2001), 13. Tanaka et al. (2002), 14. Tanaka et al. (2004), 15. Tanaka et al. unpublished observations, 16. Torres-Ruiz et al. (1996), 17. Watanabe et al. (2004), 18. Wellesen et al. (2001), 19. Yephremov et al. (1999).

that the epidermal cell layers remained intact and that fusion occurred without cytoplasmic union (Lolle et al., 1992). It was also reported that when leaves and inflorescences of wild-type and fdh plants are immersed in alcoholic solution, the rate of chlorophyll leaching is higher in fdh plants (Lolle et al., 1997). Based on this observation, the authors speculate that the higher rate of chlorophyll release is due to altered properties of the cell wall and/or the cuticle. Mutations in several other loci that cause phenotypes similar to that of fdh have since been identified by large-scale genetic screening (approximately 15 000 M1 family) and phenotypic characterisation of eceriferum mutants (ded, bud, hth, fdh, cod, cer10, thd, ahd, phd; see Table 10.1, Lolle et al., 1998). In all of these mutants, except pothead (phd), the rates of chlorophyll leaching are elevated. The correlation between chlorophyll permeability and organ fusion in these mutants is consistent with the hypothesis that changes in permeability play a role in the acquisition of fusion competence by epidermal cells (Lolle et al., 1998). Because organ fusion takes place at the organ surface and because the cuticle might be involved in the restriction of chlorophyll leaching, it is possible that cuticular properties are altered in these mutants, although this notion remains to be tested.

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BIOLOGY OF THE PLANT CUTICLE

(a) C CW CP

(b)

Leaf A Leaf B

CP CW C CW CP

Figure 10.1 Schematic representation of fused leaves and typical cuticle structure. (a) Plant with normal cuticle in which leaf fusion does not occur. (b) Plant with defective cuticle and fused leaves. Schematic diagrams of gross morphology (left), leaf sections (middle) and ultrastructure of outer epidermal surface (right) are shown. Abbreviations: c, cuticle; cw, cell wall; cp, cytoplasm.

It is also noteworthy that the permeability of a water-soluble dye (toluidine blue, TB) across the leaf surface of fdh and cer10 plants differs significantly from that of wild-type; leaves from both mutants allow incorporation of the dye whereas wild-type leaves do not (Tanaka et al., 2004). In principle, this method (TB test) allows the detection of cuticular defects and thus cuticle permeability and organ fusion competence appear to correlate in these cases. In addition, transgenic A. thaliana plants expressing a fungal cutinase also exhibit higher rates of chlorophyll leaching and organ fusion (Sieber et al., 2000), again corroborating the correlation between organ fusion and cuticle properties. Correlation between defects in the leaf cuticle and fusion of organs is also evident in cutinase-expressing plants as well as in lacerata (lcr) and abnormal leaf shape1 (ale1) mutants; in these transgenic plants and mutants, the electron-dense leaf cuticle is reduced and leaves fuse with one another (Figure 10.1; Sieber et al., 2000; Wellesen et al., 2001; Tanaka et al., 2001). In the wax2/yore-yore/pel6 mutant, the ultrastructure of the cuticular membrane in the stem is altered (Chen et al., 2003), wax contents of aldehydes and alkanes are markedly reduced (Chen et al., 2003; Kurata et al., 2003) and fusion between leaves and between floral organs is observed (Chen et al., 2003; Kurata et al., 2003; Tanaka et al., 2004). Taken together, these observations provide evidence that the cuticle plays an essential role in the prevention of organ fusion.

10.2.2 Genetic screening for mutants with defective cuticles Several groups have attempted to systematically identify A. thaliana mutants possessing an altered cuticle using criteria such as organ glossiness and physiological

THE CUTICLE AND CELLULAR INTERACTIONS

315

properties of the cuticle. Screening based on different criteria has resulted in the identification of partly overlapping but distinct classes of mutant loci. The reduction in the epicuticular waxes of the stems of A. thaliana leads to a readily visible glossy appearance and 89 independently arising eceriferum mutants, which resulted from 21 loci (CER1 through CER20 and TT5), have been identified (Dellaert et al., 1979; Koornneef et al., 1989). With a few exceptions (e.g. cer18), these mutants differ from wild type in the amount and/or morphology of epicuticular waxes on inflorescence stems. Most of the ECERIFERUM (CER) mutations do not result in organ fusion, but some (CER1, CER2, CER3, CER6, CER8 and CER10) affect fertility (Koornneef et al., 1989; Preuss et al., 1993). Interestingly, the cer10 mutant produces fused floral organs (Lolle et al., 1998). Biochemical analysis of wax contents revealed that cer mutants are associated with altered or reduced wax constituents in stems and leaves (Hannoufa et al., 1993; Jenks et al., 1995). Three other loci (CER22 through CER24) have been identified by genetic screening of 1,229 M2 individual plants derived from EMS-mutagenised A. thaliana plants based on non-visual screening by gas chromatography (Rashotte et al., 2004). These cer mutants also have drastic morphological defects. It should be noted, however, that such wax reductions having no effect on plant morphology do not necessarily demonstrate that waxes are dispensable in preventing organ fusion and plant growth as none of the cer mutants completely lacks epicuticular waxes. Jenks et al. (1996) performed a genetic screening of an A. thaliana population mutagenised by T-DNA insertions for mutants with increased leaf surface glaucousness or glossiness. Six glaucous mutants (designated knb and bcf) and two glossy mutants (designated wax1 and wax2) were identified. Wild-type A. thaliana leaves lack epicuticular wax crystals, while knb and bcf mutants produce flake-like wax crystals on the leaf surface. The wax1 and wax2 mutants produce fused organs and glossy inflorescence stems with reduced wax crystals (Jenks et al., 1996; Chen et al., 2003). Further characterisation revealed that the wax2 mutant is defective in both cuticular membrane structure and wax composition (Chen et al., 2003). Thus, it has been proposed that WAX2 might be involved in both cutin and wax production (Chen et al., 2003). Given that mutants producing fused organs include those with cuticle defects, it is reasonable to speculate that a properly functioning cuticle is required for prevention of organ fusion during the early development of leaves. Then, which constituents or properties of the cuticle are responsible for prevention of organ fusion? Tanaka et al. (2004) reported a simple method designated the TB test to detect leaves with altered cuticle properties, allowing a water-soluble molecule like TB to permeate across the defective cuticle. In principle, when leaves are dipped into TB solution, the solution is repelled if the leaf surface is covered with a hydrophobic barrier (e.g. wild-type leaf cuticle). However, if the cuticle is permeable to the solution, the dye, having affinity for the cell wall, stains the cell walls in the cuticledefective regions. The cotyledons of the cuticle-defective mutant ale1 were intensely stained with TB (Tanaka et al., 2004), while in some cer mutants (Table 10.2),

Increased b Increased a

Increased b Increased b Increased b Not significantly increased a Not significantly increased a Not significantly increased a Increased a Not significantly increased a Increased a, b Increased a Increased a Increased a Increased a, b Increased b

chr. 5 At3g59420

chr. 1 chr. 2 chr. 1 At1g02205

At1g68530

At1g72970

ahd acr4

bud cod ded cer1

cer2

cer3

cer5

cer6/pop1

cer10 cer12 cer14 cer19 fdh

th/ace

eceriferum2

eceriferum3

eceriferum5

eceriferum6/pop1

eceriferum10 eceriferum12 eceriferum14 eceriferum19 fiddlehead

hothead/adhesion of caryx edges

chr. 3 chr. 4 chr. 2 chr. 1 At2g26250

At1g51500

At5g02310

At4g24510

Increased a N.D.

chr. 2 At1g36160

ale1

abnormal leaf-shape1

ale2 acc1/pas3/gk

Permeability of leaf cuticle or cell wall

abnormal leaf-shape2 acetyl-CoA carboxylase1/ pasticinno3/ gurke airhead arabidopsis homologue of Crinkly4 bulkhead conehead deadhead eceriferum1

Locus id or chromosome

Increased a

Abbreviation

At1g62340

Mutant name

Yes

Yes N.D. N.D. N.D. Yes

N.D.

N.D.

N.D.

N.D.

No Yes Yes N.D.

Yes N.D.

N.D. N.D.

N.D.

Hydration of wild-type pollen on leaves

Table 10.2 Arabidopsis thaliana mutants defective in the generation of a functional cuticle

Fertile

Reduced Fertile Fertile Fertile Fertile

Sterile d, e

Fertile

Sterile d, e

Sterile d, e

Fertile Fertile Reduced Sterile d, e

Fertile Fertile

Fertile N.D.

Fertile

Male sterility

No

Yes Yes Yes Yes No

Yes

Yes

Yes

Yes

No No Yes Yes

No No

No N.D.

No

Glossy appearance

Related to alpha-type E3 ubiquitin-protein ligase Similar to the ABC transporters Similar to KCS and FATTY ACID ELONGASE ? ? ? ? Similar to KCS and FATTY ACID ELONGASE1 Similar to a group of FAD-containing oxidoreductases

? ? ? Similar to the sterol desaturase family proteins A novel protein

? Receptor-like protein kinase

Similar to subtilisin-like serine proteases ? Acetyl-CoA carboxylase

Gene product

13, 17

12, 17, 23, 27 12, 23, 27 12, 23, 27 12, 23, 27 15, 16, 22, 27, 33

6, 12, 18, 21

12, 20, 27

8, 9, 11, 12, 27

1, 19, 21, 27, 32

17 17 17 1, 9, 12, 27

17 7, 26, 30

28 2, 3, 5, 29

25, 28

References

316 BIOLOGY OF THE PLANT CUTICLE

lacs2

pel1

pel2

pel3

pel4

pel5

phd

thd

wax1

wax2

lacs2

peameable leaves1

peameable leaves2

peameable leaves3

permeable leaves4

permeable leaves5

pothead

thunderhead

wax1

wax2/yore-yore/pel6

N.D.

Increased b

Increased a Increased a Increased a Increased a Increased a No b Increased b N.D. Increased a, b, c

At2g45970

At1g49430

chr. 1

chr. 1

chr. 5

chr. 2

chr. 1

chr. 5

chr. 3

N.D.

At5g57800

N.D.

N.D.

Yes

No

N.D.

N.D.

N.D.

N.D.

N.D.

Yes

Yes

Stelile e

Sterile

Fertile

Fertile

Fertile

Fertile

N.D.

N.D.

N.D.

Fertile

N.D.

Yes

Yes

No

No

No

No

Yes

Yes

Yes

No

N.D.

Similarity to the sterol desaturase family and the short-chain dehydrogenase/ reductase family, homologous to Arabidopsis CER1 and maize GL1

?

?

?

?

?

?

?

?

Long-chain acyl-CoA synthetase

Fatty acid ω-hydroxylase (CYP86A8)

4, 10, 14, 27

10

17

17

27

27

27, 28

27, 28

27, 28

24

31

Notes: Mutants that produce fused organs are indicated in bold letters. N.D., not determined; a peameability of TB across leaf cuticle that was examined by the TB test; b rate of chlorophyll leaching from leaves in alcoholic solution; c rate of water loss; d defective in pollen hydration; e the male sterility can be overcome in high humidity. 1. Aarts et al. (1995), 2. Baud et al. (2003), 3. Baud et al. (2004), 4. Chen et al. (2003), 5. Faure et al. (1998), 6. Fiebig et al. (2000), 7. Gifford et al. (2003), 8. Hannoufa et al. (1996), 9. Hülskamp et al. (1995), 10. Jenks et al. (1996), 11. Jenks et al. (2002), 12. Koornneef et al. (1989), 13. Krolikowski et al. (2003), 14. Kurata et al. (2003), 15. Lolle et al. (1992), 16. Lolle et al. (1997), 17. Lolle et al. (1998), 18. Millar et al. (1999), 19. Negruk et al. (1996), 20. Pighin et al. (2004), 21. Preuss et al. (1993), 22. Pruitt et al. (2000), 23. Rashotte et al. (2004), 24. Schnurr et al. (2004), 25. Tanaka et al. (2001), 26. Tanaka et al. (2002), 27. Tanaka et al. (2004), 28. Tanaka et al. unpublished observations, 29. Torres-Ruiz et al. (1996), 30. Watanabe et al. (2004), 31. Wellesen et al. (2001), 32. Xia et al. (1996), 33. Yephremov et al. (1999).

lcr

lacerata

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317

318

BIOLOGY OF THE PLANT CUTICLE

the leaves were stained weakly or in specific regions (e.g. the petiole). Using the TB test as an assay for defective cuticles, a genetic screening of EMS-mutagenised A. thaliana population (descended from 3305 M2 plants) for mutants was performed and a total of 19 mutants exhibiting alterations in leaf surface properties were identified. The results of genetic mapping, examination of allelism and phenotypic classification revealed that these mutations represent at least seven loci [FDH, PERMEABLE LEAVES (PEL1 through PEL6)]. The pel6 mutant contained a mutation in the WAX2/YORE-YORE (YRE) gene, suggesting that PEL6 is a new mutant allele of WAX2/YRE. The genes responsible for the PEL1 through PEL5 mutations have not been cloned to date. Among the mutants identified, pel1, pel2, pel3 and fdh, which are more intensely stained than pel4 and pel5, produced fused leaves. The PEL4 and PEL5 mutations do not affect gross morphology of the plants and appear to be distinct from typical cer mutants; the glaucous appearance of inflorescence stems in pel4 and pel5 is indistinguishable from that of wild type. The pel6 mutant exhibits predominant staining of the trichomes, which also fuse to one another (Tanaka et al., 2004). The correlation between the intensity and/or pattern of staining and fusion events implies that the permeability of the cuticle is a critical factor that can determine the competency for organ fusion. Further phenotypic analysis of these mutants and identification of responsible genes as well as understanding of the biochemical roles of the gene products will deepen our understanding of the molecular basis for generation of cuticle as a permeability barrier.

10.2.3 Molecular identification of genes involved in the generation of a functional cuticle Genes responsible for the mutations described earlier have been identified (Table 10.2). In this section, the structural features and predicted functions of encoded proteins, particularly those apparently involved in the biosynthesis or export of cuticular constituents, are described. The FIDDLEHEAD (FDH) gene was cloned using the transposon-tagging procedure (Yephremov et al., 1999; Pruitt et al., 2000). The predicted FDH protein sequence exhibited significant sequence similarity to various condensing enzymes and most closely resembles those encoded by the FATTY ACID ELONGASE (FAE) family, such as β-keto-acyl-CoA synthase (KCS) from Simmondsia chinensis (jojoba) (Lassner et al., 1996), and FAE1, KCS1 and CER6/CUT1 from A. thaliana (James et al., 1995; Todd et al., 1999; Millar et al., 1999; Fiebig et al., 2000). There is increasing evidence suggesting that these factors are involved in the generation of very long-chain fatty acids (VLCFA; carbon-chain length >18). VLCFA are formed by a microsomal fatty acid elongation (FAE) system (for review see Somerville et al., 2000; Jenks et al., 2002). Each elongation reaction involves the addition of a two-carbon unit from malonyl-coenzyme A (CoA) to an acyl primer (e.g. fatty acyl-CoA in various chain length), followed by reduction, dehydration and a final reduction.

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319

Mutations in the A. thaliana FAE1 locus result in reduced VLCFA levels in seeds (James and Dooner, 1990; Lemierx et al., 1990; James et al., 1995), suggesting that FAE1 is involved in in vivo. Overexpression of FAE1 in tissues of A. thaliana and tobacco results in the accumulation of derivatives of VLCFA, as if FAE1 is the rate-limiting enzyme that catalyzes lipid chain elongation in vivo (Millar and Kunst, 1997). Using the Brassica napus low erucic (docos-13,14-enoic) acid rapeseed (LEAR) cultivar, which is defective in the FAE system, Lassner et al. (1996) demonstrated that the expression of jojoba KCS cDNA in the LEAR plants restored KCS activity, which is involved in the first step of the FAE system. CER6/CUT1 is required for the elongation of fatty-acid derivatives (Hannoufa et al., 1993; Preuss et al., 1993; Millar et al., 1999; Fiebig et al., 2000). Thus, the strong similarity in the amino acid sequences of FDH, KCS and FAE1 suggest that FDH may also be involved in the lipid metabolism required for the generation of a proper cuticle. In addition, consistent with the notion that FDH is involved in metabolism of cuticle-related lipids, FDH mRNA is detected in epidermis-related tissues such as the L1 layer of inflorescences (Yephremov et al., 1999). The LACERATA (LCR) gene encodes a cytochrome P450 family protein (CYP86A8) (Wellesen et al., 2001). This raises the possibility that LCR acts as an oxygenase, which is involved in the generation of a functional cuticle. Wellesen et al. (2001) further demonstrated that microsomes purified from yeast expressing LCR displayed significant activity in the ω-hydroxylation of fatty acids. Among several substrates examined, the most effective were hexadecanoate (C16:0) and octa-9,10-decenoate (C18:1). Considering that the chief monomers of cutin are 10,16-dihydroxyhexadecanoic acid, 18-hydroxy-9,10-epoxyoctadecanoic acid and 9,10,18-trihydroxyoctadecanoic acid, fatty acid hydroxylation is likely required for the biosynthesis of cutin monomers (Kolattukudy, 1980, 2001), and thus it is plausible that LCR is directly involved in cutin biosynthesis. The WAX2/YORE-YORE (WAX2/YRE) gene encodes a predicted protein of 632 amino acids. The sequence of this protein exhibits the highest sequence similarity to putative proteins encoded by Senecio odora cDNA EPI23 (Hansen et al., 1997) and the rice GLOSSY1 homologue (Hansen et al., 1997) and it is also related to A. thaliana CER1 (Aarts et al., 1995) and maize glossy1 (Hansen et al., 1997), which is required for wax generation. The WAX2 protein sequence is related to the sterol desaturase family at the amino terminal region and possesses the three His-rich motifs that are conserved in this family. The sequence near the carboxy terminus of this protein also exhibits similarity to the signature sequence found in the shortchain dehydrogenase/reductase (SDR) family. Such similarities are not found in GLOSSY1 or CER1. Based on the similarity with metabolic proteins, WAX2/YRE is believed to be involved in a certain metabolic process for the modification of fatty-acid derivatives. Phenotypic analysis has suggested that the wax2/yre mutant is defective in generation of aldehydes and alkanes as well as in cutin formation. Because the two domains are each related to different metabolic enzymes, an interesting hypothesis is that WAX2/YRE is a fusion of two functional proteins and this might explain

320

BIOLOGY OF THE PLANT CUTICLE

the defects of the wax2 mutant in both the generation of the cuticular membrane and cuticular wax. Consistent with the predicted role of WAX2/YRE in cuticle generation, the WAX1/YRE gene is specifically expressed in the protoderm and the epidermis (Kurata et al., 2003). Kurata et al. (2003) reported a synergistic effect of CER1 and YRE mutations on the morphology of trichomes, suggesting partially overlapping functions of CER1 and WAX2/YRE. The major constituents of the cuticle, cutin and waxes, are derived from fatty acids (carbon-chain length 16 and 18) generated in chloroplasts. Elongation of fatty acyl-CoA catalyzed by the FAE system requires malonyl-CoA as a carbon source (for reviews, see Somerville et al., 2000; Jenks et al., 2002). Acetyl-CoA carboxylase is known to catalyze the generation of malonyl-CoA using acetyl CoA as a substrate (Somerville et al., 2000). Recently, gurke/pasticino3 (gk/pas3) mutants of A. thaliana were found to have mutations in the ACETYL-COA CARBOXYLASE1 (ACC1) gene (Baud et al., 2004). Whereas strong GK/PAS3/ACC1 mutations cause a specific defect in the embryonic pattern such that cotyledons and shoots fail to develop (Baud et al., 2003), weak GK/PAS3 mutations have been reported to cause the fusion of lateral organs (Torres-Ruiz et al., 1996; Faure et al., 1998). Thus, it is possible that ACC1 is involved in generating the malonyl-CoA pool, which is used for biosynthesis of cuticle-related lipids such as waxes, although it remains unclear whether the gk/pas3/acc1 mutant is actually defective in cuticle properties. The plant cuticle is composed of derivatives of fatty acids, which are generated de novo in chloroplasts. The final products of fatty acid synthesis in the chloroplasts are 16:0-ACP, 18:0-ACP and 18:1-ACP. Before export from chloroplasts, they are cleaved from ACP by thioesterases. Subsequently, long-chain acyl-CoA synthetases (LACSs) catalyze the conversion of free fatty acids to fatty acyl-CoAs, which are used for general membrane lipid synthesis as well as wax biogenesis (Chapter 5). A. thaliana contains nine LACS isozymes (Shockey et al., 2002). Schnurr et al. (2004) demonstrated that one of these nine isozymes, LACS2, is involved in cuticle formation. First, using the LACS2 promoter-GUS reporter gene, LACS2 expression is specifically detected in the epidermis. Second, in vitro assays show that 16-hydroxyhexadecanoate is an excellent substrate for LACS2. Third, the lacs2 mutant exhibits defects in epidermal surface functions, such that the rate of chlorophyll leaching from leaves in alcoholic solution increases and mutant leaves, particularly on abaxial surfaces, support pollen germination. In addition, ultrastructural analysis, visual inspection of inflorescence stems and investigation of wax load and composition have suggested that the cutin layer in the abaxial surface of leaves of lacs2 mutants is thinner than that in the wild type, but the LACS2 mutation does not substantially affect wax generation (Schnurr et al., 2004). The lacs2 phenotype is similar to that of lcr mutants and transgenic plants expressing a fungal cutinase. The latter appear to have a defect in the cutin layer, although the phenotypes of lacs2 are distinct from those of LCR and cutinase-expressing plants, which also generate fused organs. The reasons for these phenotypic differences in terms of organ fusion are unclear; they may be the result of a relatively

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321

weak effect of the LACS2 mutation on cutin formation, as it seems that cutin is still sufficiently formed in the lacs2 mutant, particularly on the adaxial leaf surface. The HOTHEAD (HTH) gene was identified based on its chromosomal location and a candidate approach. It is predicted to encode a protein with 594 amino acids having similarity to the flavoprotein form of α-hydroxynitrile lyase and other oxidoreductases (Krolikowski et al., 2003). Considering that the hth mutant exhibits the organ fusion phenotype (Lolle et al., 1998; Krolikowski et al., 2003), the HTH gene might be directly or indirectly involved in the generation of a functional cuticle. Further biochemical characterization of the cuticle in the hth mutant would shed light on the role of HTH. The A. thaliana genome (The Arabidopsis Genome Initiative, 2000) contains seven other HTH-related genes, but the functions of these genes remain to be elucidated. Several eceriferum (cer) mutants have been shown to affect the properties of cuticle as indicated by the TB test (Tanaka et al., 2004; Table 10.2); most plants homozygous for CER5, CER10, CER12, CER14 and CER19 exhibited leaf staining with an aqueous solution of TB. This suggests that permeability of water-soluble molecules through the cuticle is increased in these mutants. The gene responsible for the CER5 mutation has been cloned (Pighin et al., 2004) and detailed phenotypic analysis involving TEM and cryo-SEM of the cer5 revealed an unusual inclusion of cytoplasm in the vacuole. Furthermore, unusual sheet-like inclusions appear to be accumulated in this cytoplasm in epidermal cells in the stem. Nile red staining indicated that these inclusions are lipidic and that the cuticular wax load is significantly reduced in the cer5 mutant, while total epidermal wax (surface plus intracellular) of wild-type and cer5 tissues do not differ significantly. Thus, cer5 appears to be defective in the export of lipids related to wax, not in wax biogenesis (Pighin et al., 2004). The CER5 gene has been isolated by a map-based approach combined with analysis of insertion mutants. The predicted CER5 protein contains domains characteristic of ABC transporters, including Walker A and B boxes, the C motif for nucleotide binding and six transmembrane domains. Thus, CER5 is predicted to be a so-called half-transporter and presumably dimerisation is required for its function. An attractive hypothesis is that CER5 is directly involved in the transport of wax-related lipids (see Chapter 5). Consistent with this notion, the CER5 gene is expressed in epidermal cells and the GFP-CER5 fusion protein is localised in the cell surface, presumably in the plasma membrane (Pighin et al., 2004). CER5 transcripts have been found in all organs examined, including inflorescence stems, leaves and roots. Incomplete loss of cuticular wax in the stems and leaves in the cer5 mutant suggest the presence of factors with overlapping function. The A. thaliana genome contains 129 genes encoding putative ABC transporters (Sánchez-Fernández et al., 2001). It would be of interest to test whether other ABC transporters, particularly those with significant sequence similarity to CER5 (e.g. At3g21090 and At1g51460), have functions overlapping with those of CER5 in vivo.

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BIOLOGY OF THE PLANT CUTICLE

10.3

Functions of the cuticle in plant reproduction and embryogenesis

10.3.1 The cuticle as an interface in stigma–pollen interactions Successful fertilisation involves pollen adhesion onto the stigma surface, hydration, pollen germination, pollen tube penetration of the stigma surface and pollen tube elongation towards the micropyle of ovule. In plants with a ‘dry’ stigma (HeslopHarrison, 1977), stigmatic surfaces are covered with a cuticle and, in such plants, it appears that the pollen breaches the cuticle to proceed into stigmatic cell walls (Dickinson and Lewis, 1973; Elleman et al., 1992). On the basis of ultrastructural observation, the electron-dense cuticle disappears prior to penetration of the pollen tube into the stigmatic cell wall, at least in plants including Raphanus and A. thaliana (Dickinson and Lewis, 1973; Elleman et al., 1992). These observations support the idea that the cuticle might be enzymatically degraded prior to penetration (Figure 10.2), and several physiological experiments reinforce this notion. Heslop-Harrison and Heslop-Harrison (1975) treated stigmata of Agrostemma githago with enzymatic solutions and examined their effect on pollen tube germination and entry. After treatment of stigma surfaces with pronase and subsequent pollination, pollen grains germinated but failed to penetrate implying that the stigmatic cell surface includes a protein that possesses cutinase activity or that may be involved in cutinase activation. Hiscock et al. (1994) reported that the pollen of B. napus contains an active cutinase. The protein sequence of this cutinase has not been determined to date; however, using an antibody raised against fungal cutinase, they also examined the localisation pattern of the cross-reactive substance in pollen. In their immunocytological experiments, signals were obtained at the site where the pollen tube emerges (Hiscock et al., 1994). This is consistent with the hypothesis that cutinase is localised at the tip of the pollen tube and that it may be involved in penetration of the stigmatic cuticle, although direct evidence is lacking. Apart from the location of the hypothetical cutinase, both the stigmata and pollen of B. napus may possess cutinase, as shown by a biochemical assay using the synthetic substrate p-nitrophenyl butyrate (PNB; Hiscock et al., 2002). In addition, such esterase activities were blocked by inhibitors of fungal cutinase, such as diisopropyl fluorophosphates (DIPF) and ebelactone B (Hiscock et al., 2002). Using such inhibitors, a physiological experiment was performed in order to investigate the causal relationship between inhibition of esterase (most likely cutinase) and pollen penetration through the stigmatic surface. Treatment of the B. napus stigma with either DIPF or ebelactone B does not seriously impair pollen germination but inhibits penetration by the pollen tube of the dry cuticle-covered Brassica stigma (Hiscock et al., 2002). As a result, Hiscock et al. (2002) proposed that cutinase is required for the penetration of the stigmatic cuticle by the pollen tube in B. napus. Thus, the results of ultrastructural analysis and physiological experiments suggest that the cuticle covering dry stigmatic surfaces functions to prevent pollen tube penetration. However, it is unclear whether degradation of stigmatic cuticle is sufficient for pollen tube penetration.

m

Recognition of a signal

m

m

m

Co-pollination of wild-type and mutant pollen

(d)

H2O

m

Polen tube grows

Successful fertilization of the mutant pollen

Pollen Pollen germination hydration and degradation of stigmatic cuticle

H2O

Recognition of a Hydration of both signal from wildwild-type and type pollen mutant pollen

m

Adhesion of Recognition of wild-type pollen a signal (lipids?) on stigma surface

(b)

Accumulation of callose in the papillae Pollen hydration

Stigmatic papillae

Pollen

Figure 10.2 Schematic diagram of successive events in compatible pollination. (a) Diagram of stigma of Arabidopsis thaliana. (b) Sequential events following compatible pollination. (c) Events following pollination by mutant pollen defective in pollen–stigma recognition. (d) Restoration of mutant. Abbreviation: m, pollen fertility by co-pollination with wild-type pollen. This figure is produced in colour in the colour plate section, which follows pages 249.

Adhesion of mutant pollen on stigma

(c)

(a)

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323

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BIOLOGY OF THE PLANT CUTICLE

The predicted role of the stigmatic cuticle described above is ‘protective’; it appears as though the cuticle protects the stigma from invasion by pollen. Molecular genetic studies, however, suggest that cuticle-related materials are actively involved in the recognition of compatible pollen by the stigma. In Brassicaceae, compatible pollination results in successful penetration of pollen through the cuticle and in the subsequent progression and fertilisation. In contrast, incompatible pollen is able to germinate and the stigmatic cuticle disappears upon pollen attachment, but stigmatic papillae cells accumulate callose and the pollen fails to progress (Dickinson and Lewis, 1973). Preuss et al. (1993) isolated a male sterile A. thaliana mutant defective in pollen–pistil interactions ( pop1, also known as cer6-2). Pollen generated by pop1 homozygous plants is able to attach to stigmatic papilla cells but fails to hydrate and germinate on the stigmatic surface. Similarly to incompatible pollination in Brassica, pollination with pop1 pollen induces accumulation of callose in stigmatic cells. Interestingly, pop1 pollen, when co-pollinated with wild-type pollen, is able to germinate and successfully complete fertilisation (Preuss et al., 1993; Figure 10.2). The reason for this is that in compatible pollination (as in wild-type A. thaliana), stigmatic cells perceive a signal from compatible pollen and alter their properties to support pollen hydration. In this scenario, pop1 pollen is defective in the generation of such a signal. Although the molecular nature of this signal is unknown, it is reasonable to speculate that cuticle-related lipids are involved in such a signalling event (Figure 10.2), as in addition to a defect in fertility, the pop1 mutant also exhibits a striking defect in the production of long-chain lipids in inflorescence stems and pollen (Preuss et al., 1993). The POP1 mutation is allelic to CER6 and the CER6/POP1 gene was found to be identical to the CUT1 gene, which encodes a putative condensing enzyme (Millar et al., 1999; Fiebig et al., 2000). Thus, the CER6/POP1/CUT1 protein appears to be directly involved in the production of VLCFA, corroborating a causal relationship between loss of VLCFA and the pop1 phenotype. Analysis of intragenic suppressors of cer6 mutants has demonstrated that low amounts of VLCFA are sufficient for pollen hydration and germination (Fiebig et al., 2000). The cer2 mutant which has been shown to have a defect in the generation of long-chain lipids also exhibits defects in pollen hydration (Preuss et al., 1993). Hülskamp et al. (1995) performed genetic screening of an EMS-mutagenised A. thaliana population (15 000 M2 plants) for male-sterile mutants defective in pollen–stigma recognition and identified five independently arising mutants. All five mutants were associated with a bright green stem phenotype, which is typical of eceriferum mutants. Based on the results of allelism tests and genetic mapping, these mutants were found to be allelic to CER1 (three alleles), CER3 (one allele) and CER6 (one allele) (Hülskamp et al., 1995). Pollen from these mutants fails to hydrate when placed on the stigma and these defects were restored by co-pollination with wild-type pollen (Hülskamp et al., 1995). The hydration of mutant pollen was facilitated even when wild-type pollen was treated with formaldehyde to prevent germination. Taken together, these results suggest that CER1, CER2, CER3 and

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CER6 genes in the male parent appear to be involved in pollen recognition by the stigma, and the subsequent transfer of water from the stigma to pollen (Figure 10.2; Table 10.2). It has been shown that cer1, cer2, cer3 and cer6 are defective in various classes of long-chain lipids (Hannoufa et al., 1993; Preuss et al., 1993; Jenks et al., 1995). Nonacosane (C29) is commonly lacking in these mutants, which is consistent with the notion that this compound and/or compounds related to it are required for pollen– stigma recognition. Another class of lipids – triacylglycerides – is implicated in pollen hydration, as application of trilinolein (a cis-unsaturated triacylglyceride) enables pollen grains obtained from cer1, cer3 and cer6 to produce tubes that penetrate the stigma (Wolters-Arts et al., 1998). A. thaliana mutants with defective leaf cuticle provide opportunities for testing the causal relationship between the cuticular barrier and the capacity for pollen hydration on the leaf surface. Pollen does not germinate when adhering to the surface of wild-type A. thaliana leaves. Interestingly, Lolle and Cheung (1993) reported that pollen attached to leaves of the fiddlehead mutant is capable of hydration. In addition to fdh, various mutant plants have been reported to allow adhered pollen to hydrate and germinate (reviewed in Lolle and Pruitt, 1999; Wellesen et al., 2001; Chen et al., 2003; Schnurr et al., 2004; Table 10.2). The genes responsible for these mutations include those encoding for a putative KCS (fatty acid elongase), fatty acid ω-hydroxylase and LACS, which appear to be directly involved in the biosynthesis of the cuticle (Table 10.2). Therefore, the cuticle seems to function as a barrier to pollen hydration in leaves. The lcr and lacs2 mutants are primarily defective in cutin biogenesis, suggesting that the cutin layer is essential for preventing pollen hydration on the leaf surface. The cuticle components or structures required for restricting pollen hydration remain elusive in other mutants.

10.3.2 Role of the cuticle during embryogenesis During embryogenesis, the cuticle is generated to surround developing embryos. A cuticle can be detected in plant embryogenesis as early as the zygote stage in the case of Citrus jambhiri (Bruck and Walker, 1985). In other plants, such as maize and A. thaliana, electron-dense cuticles, as detected by transmission electron microscopy, apparently are generated on the surface of globular-stage embryos prior to the generation of organ primordia. Why is the cuticle generated so early in embryogenesis? One clue comes from observations of an A. thaliana mutant defective in cuticle formation during embryogenesis. The abnormal leaf shape1 (ale1) mutant exhibits a defect in the generation of a continuous cuticle during embryogenesis and in seedlings. In addition to the cuticular defects, embryos of the ale1 mutant adhere to the endosperm cells that surround the developing embryos (Tanaka et al., 2001). In addition, it has been reported that mutation in A. thaliana ACC1 gene also causes adhesion between embryos and endosperm (Baud et al., 2004). Because ACC1 is probably involved in the generation of cytosolic malonylCoA, which is required for the biosynthesis of VLCFA, it is possible that the adhesion

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between embryos and endosperm is caused by a defect in cuticle formation (Baud et al., 2004).

10.4

Regulators of epidermal differentiation and cuticle formation

10.4.1 Intercellular signalling and cuticle formation The abnormal leaf-shape1 (ale1) mutant is characterised by the partial loss of an electron-dense cuticle on embryos and juvenile plants, increased permeability of TB solution across the leaf surface and sensitivity of seedlings to low humidity. The ALE1 gene encodes a protein related to subtilisin-like serine proteases (Tanaka et al., 2001). The sequence of the predicted ALE1 protein includes a signal sequence, which may direct the protein into a secretory pathway. Transcripts from the ALE1 gene are predominantly accumulated within endosperm cells surrounding developing embryos. This raises the possibility that, during embryogenesis, cuticle formation requires communication between the protoderm and extra-embryonic tissue, and that ALE1 might be involved in such a biological process. A number of subtilisin-like proteases in yeast and animals have been reported to catalyze the cleavage of various signalling molecules and this cleavage, in combination with other modifications, results in either activation (e.g. a-factor, insulin) or inactivation (e.g. neurotransmitter) of the signalling molecules. It would be of interest to determine whether ALE1 functions in a similar manner. Maize crinkly4 (cr4) mutants exhibit various abnormalities in the leaf epidermis such that epidermal cells are deformed and leaves fuse together (Becraft et al., 1996). In the cr4 mutant, the electron-dense cuticle is reduced (Jin et al., 2000). The CR4 gene encodes a receptor-like therine/threonine kinase with a putative extracellular domain containing novel repeats of 39 amino acids and a cysteine-rich region similar to that of the tumornecrosis factor receptor (TNFR) of mammals. Based on these results, CR4 is believed to transmit an extracellular signal into the cytosol and such signal transduction may promote differentiation of epidermal cells. Later, it has been demonstrated that a role in epidermal differentiation for the CR4-related receptorlike protein kinases is conserved between maize and A. thaliana (Gifford et al., 2003; Watanabe et al. 2004). An A. thaliana homologue of CRINKLY4 (ACR4) has been identified by degenerate PCR on the basis of the similarity to the maize CRINKLY4 gene (Tanaka et al., 2002). The A. thaliana genome contains five genes encoding proteins with putative CR4-related extracellular domains. Among these, only ACR4 contains all of the domains found in CR4 (Tanaka et al., 2002; Cao et al., 2005). ACR4 is preferentially expressed in epidermis-related tissues (Tanaka et al., 2002; Gifford et al., 2003) and acr4 mutants exhibit defects in the organization of cell layers in ovule integuments and the endothelium (Gifford et al. 2003; Watanabe et al., 2004) and in the generation of an electron-dense cuticle in the ovule (Watanabe et al., 2004). Furthermore, cuticle permeability in the ovule as well as in the leaves of acr4 is altered, as demonstrated by TB test.

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Epidermal defects in the leaves of acr4 are frequently observed in leaf petioles and the abaxial surfaces of leaf lamina as well as in unusual protrusions of epidermal cells on leaves (Watanabe et al., 2004). However, the ACR4 mutation has little or no effect on overall leaf morphology. The epidermal defects of the ALE1 mutant, originally identified in the Landsberg erecta (Ler) ecotype, are restored when introduced into the Wassilewskaja (Ws) ecotype by genetic crossing (Watanabe et al., 2004). The acr4 ale1 double mutant generated using single mutants crossed into the Ws background exhibited severe epidermal defects in the cotyledons and leaves with eventual deformation of overall shoot morphology (Watanabe et al., 2004). These results demonstrate that ALE1 and ACR4 are collectively essential for proper cuticle formation as well as organ development in A. thaliana (Watanabe et al., 2004).

10.4.2 Transcriptional control of cuticle production A number of genes involved in the biosynthesis or secretion of cuticle-related substances have been identified, as described earlier. Consistent with previous observations that biochemical activities related to cuticle biosynthesis are detected within the epidermis (Croteau and Kolattukudy, 1974; Evenson and Post-Beittenmiller, 1995), a number of these genes are reported to be predominantly expressed in epidermal and related tissues, such as the meristem L1 layer. It seems likely that differentiation of the epidermis and appropriate gene expression in epidermal cells is a prerequisite for cuticle generation. Two distinct classes of transcription factors that are probably involved in the expression of genes for cuticle biosynthesis have recently been described (Abe et al., 2003; Aharoni et al., 2004; Broun et al., 2004). The PROTODERMAL FACTOR1 (PDF1) gene of A. thaliana encodes for a putative extracellular proline-rich protein that is exclusively expressed in the L1 layer of shoot apices and the protoderm of organ primordia (Abe et al., 1999). A 1.5-kb region upstream of the PDF1 gene has been shown to direct L1-layerspecific expression of the β-glucuronidase (GUS) reporter gene. By analysis of progressive deletions of the PDF1 gene promoter, Abe et al. (2001) identified an 8-bp cis-regulatory element [designated L1 box, 5′ -TAAATG(C/T)A-3′ ] that is conserved in various L1-specific genes (e.g. FDH: Yephremov et al., 1999; SCR; WysockaDiller et al., 2000; ATLTP1; Thoma et al., 1994) and is critical for L1 layer-specific expression of the GUS reporter gene under control of the PDF1 promoter (Abe et al., 2001). It has been shown that two related homeo domain proteins, ATML1 and PDF2, can bind to a 21bp DNA containing the L1-box sequence in vitro and a mutation in the L1-box sequence abolishes this binding (Abe et al., 2001; Abe et al., 2003). Transcripts from ATML1 and PDF2 predominantly accumulate in the L1 layer and the protodermal cells of embryos (Lu et al., 1996; Abe et al., 2003). In addition, Abe et al. (2003) further demonstrated that A. thaliana plants doubly homozygous at T-DNA insertion of ATML1 and PDF2 alleles failed to generate typical epidermal pavement cells and to express the L1-layer-specific genes such as PDF1 and ACR4, although single mutants did not exhibit obvious defects.

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In transgenic A. thaliana plants with PDF2 co-suppression, fusion between sepals along their edges is often observed (Abe et al., 2003). This suggests that ATML1 and PDF2 have a redundant function and regulate the expression of L1and protoderm-specific genes in an L1-box-dependent manner. Considering the tight correlation between organ fusion and cuticle properties described earlier, organ fusion in the PDF2 co-suppressed plants might have been caused by altered expression of the genes involved in cuticle formation, although further experimental data are necessary. In this regard, the presence of the L1 box in the FDH promoter is of particular interest because loss of FDH expression would result in the fusion of organs, including sepals. Several other genes responsible for preventing organ fusion also include the L1 box and related sequences in upstream regions (Tanaka et al., unpublished results). The production of epicuticular wax involves various genes that are predominantly expressed in epidermis-related tissues (Chapter 4). For example, transcripts of the CER2, CER5, CER6/POP1 and WAX2/YRE/PEL6 genes are preferentially detected in the L1 layer of the shoot apical meristem, protodermal cells of organ primordia and epidermal cells, and mutations in these genes result in apparent modification of and/or reductions in wax content (Xia et al., 1996; Millar et al., 1999; Kurata et al., 2003; Pighin et al., 2004). In addition, using the β-glucuronidase (GUS) reporter, it has been demonstrated that genomic DNA fragments containing the 5′ -upstream regulatory sequences from genes such as CER2, CER6, YRE and CER5 (Xia et al., 1996; Hooker et al., 2002; Kurata et al., 2003; Pighin et al., 2004) are sufficient to direct the expression of the reporter gene in an epidermis-specific manner. These results indicate that the production of epicuticular wax is controlled at the transcription level, although other controlling mechanisms may also be involved. In an effort to functionally characterise A. thaliana transcription factors, Broun et al. (2004) performed functional screening of transgenic plants harbouring genes for transcription factors under the control of the CAMV 35S promoter. They identified a phenotype typical of wax overproduction in transgenic plants overexpressing the full-length sequence of an ERF/EREBP-type transcription factor gene, AT1G15360 [named WAX INDUCER1 (WIN1)]. In these plants, leaves were glossier than those of control plants and more alkanes were accumulated in both the leaves and the stem, while marked changes in nonacosane in leaves and stems and hentriacontane in leaves were seen. Primary alcohol content was lower in leaves and higher in stems when compared with the wild type. Thus, WIN1 overexpression results in a preferential increase in the products of the decarbonylation pathway. The fact that WIN1 is a putative transcription factor whose overexpression caused increased production of alkanes led the authors to examine the expression of genes potentially involved in wax biosynthesis. Among the genes examined, transcripts of CER1, KCS1 and CER2 were more abundantly accumulated in the leaves of transgenic plants expressing WIN1 than in wild-type leaves. Thus, it seems that WIN1 either directly or indirectly promotes the expression of these genes. Aharoni et al. (2004) independently identified WIN1/SHAIN1 (SHN) overexpressing plants

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by screening for an activation tagging line. In leaves of WIN/SHN-overexpressing plants, the contents of alkanes, secondary alcohols and ketones, which are produced by the decarbonylation pathway, were increased when compared with wild-type leaves (Aharoni et al. 2004). The CER1 protein appears to be positively involved in the production of alkanes, as the cer1 mutant is defective in alkane production (Hannoufa et al., 1993; Jenks et al., 1995), although compounds such as primary alcohols are also decreased in the cer1 mutant. Induction of CER1 expression by win1 and accumulation of alkanes is thus consistent with the observed phenotype and the hypothesis that CER1 facilitates decarbonylation. Transcriptional factors regulating the expression of the metabolic enzymes responsible for cuticle biogenesis have been emerging for some time, as describe earlier. Cuticle biogenesis is tightly linked with the differentiation of the protoderm and the epidermis during development as well as with the outer environment. Elucidation of target genes as well as the regulation mechanisms for the expression and activation of these transcription factors would help our understanding of cuticle biogenesis during the plant life cycle.

10.5

Concluding remarks

Recent genetic and biochemical analyses have highlighted the physiological roles of the cuticle in intercellular and intertissue communication, including the prevention of fusion between organs and tissues, and the hydration and penetration of pollen during fertilisation. There is increasing evidence supporting the notion that permeability of molecules across the cuticle and a proper cutin layer are essential for preventing organ fusion, although it is still unclear to what extent wax is involved in these traits. The genes involved in the various steps of functional cuticle formation have been identified, including those apparently involved in the differentiation of epidermis-related tissues, transcriptional control, cuticle biogenesis and export of cuticle-related lipids. The plant genome presumably contains a family of genes involved in the biogenesis and export of cuticle-related lipids. Although unclear at present, the genetic and biochemical relationships between the regulators and the genes themselves appear to be directly involved in cuticle biogenesis. Further efforts using genetic materials as well as the information obtained to date, may reveal the causal relationship between loss of specific constituents or physiological alteration of cuticle and cellular interactions as well as the molecular mechanisms regulating cuticle biogenesis.

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Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

11 Microbial communities in the phyllosphere Johan H.J. Leveau 11.1

Introduction

The term phyllosphere was coined by Last (1955) and Ruinen (1956) to describe the plant leaf surface as an environment that is physically, chemically and biologically distinct from the plant leaf itself or the air surrounding it. The term phylloplane has been used also, either instead of or in addition to the term phyllosphere. Its twodimensional connotation, however, does not do justice to the three dimensions that characterise the phyllosphere from the perspective of many of its microscopic inhabitants. On a global scale, the phyllosphere is arguably one of the largest biological surfaces colonised by microorganisms. Satellite images have allowed a conservative approximation of 4 × 108 km2 for the earth’s terrestrial surface area covered with foliage (Morris and Kinkel, 2002). It has been estimated that this leaf surface area is home to an astonishing 1026 bacteria (Morris and Kinkel, 2002), which are the most abundant colonisers of cuticular surfaces. Thus, the phyllosphere represents a significant refuge and resource of microorganisms on this planet. Often-used terms in phyllosphere microbiology are epiphyte and epiphytic (Leben, 1965; Hirano and Upper, 1983): here, microbial epiphytes or epiphytic microorganisms, which include bacteria, fungi and yeasts, are defined as being capable of surviving and thriving on plant leaf and fruit surfaces. Several excellent reviews on phyllosphere microbiology have appeared so far (Beattie and Lindow, 1995, 1999; Andrews and Harris, 2000; Hirano and Upper, 2000; Lindow and Leveau, 2002; Lindow and Brandl, 2003). This chapter aims to provide a brief but broad and updated synthesis of research activities and findings related to phyllosphere microbiology. As the next chapter in this volume (Chapter 12) will focus in greater detail on phyllosphere fungi, the present chapter is biased, in some sections at least, towards bacterial colonisers of the leaf surface.

11.2

Methodologies in phyllosphere microbiology

11.2.1 Sampling techniques Sampling the phyllosphere is at the basis of many observations or experiments in leaf surface microbiology, and thus deserves its own and in-detail section

Publication 3593 NIOO-KNAW Netherlands Institute of Ecology.

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at the beginning of this chapter. Several reviews are available that describe the advantages and disadvantages of different methods for sampling and quantification of microorganisms from the phyllosphere (Donegan et al., 1991; Jacques and Morris, 1995; Dandurand and Knudsen, 1996). One of the simplest techniques is leaf printing (Corpe, 1985), by which a plant leaf is pressed onto the surface of an agar plate for a defined period of time and carefully removed again. The agar plate is then incubated to allow growth of the microorganisms that transferred from the leaf surface. Depending on the medium and the abundance and composition of the microflora, the result is a collection of bacterial and fungal growth foci (‘colonies’) that often follow the contours of the leaf. Obvious disadvantages of the method are its low resolution of observation and its limited ability to provide a quantitative appreciation for the phyllosphere community composition. However, leaf printing can provide a first indication that the distribution of microorganisms on leaf surfaces is not uniform (Leben, 1998). For example, more growth often occurs where the veins of the leaf touch the agar, suggesting that these structures are more densely populated areas on the leaf (Manceau and Kasempour, 2002). Furthermore, leaf printing has been an extremely useful and popular laboratory experiment for teaching purposes (Holland et al., 2000). The outcome of a leaf printing experiment will depend in large part on the medium that is used to prepare the agar plate. Medium composition can be varied and exploited to look only at a subset of the microbial community on the leaf. By incorporation of compounds with antifungal (e.g. cycloheximide) or antibacterial (e.g. penicillin) activity, fungal or bacterial colonisers, respectively, can be excluded from the analysis. As microorganisms differ in their nutritional capabilities, it is also possible to include specific nutrient sources in the medium to select for a nutritionally defined subpopulation of the phyllosphere. For example, agar plates that contain methanol as the sole source of carbon have been used to demonstrate the ubiquity of methylotrophic bacteria on plant leaf surfaces (Corpe, 1985). More quantitative than leaf printing is the method of leaf washing. One or more leaves are placed in a tube or flask containing a specified volume of wash solution (e.g. saline solution or phosphate buffer), and microbes are removed from the cuticular surface by vortexing and/or sonication. The number of bacteria in the wash solution is then determined by the most probable number (MPN) technique (Oblinger and Koburger, 1975) or by plating, directly or after dilution of the wash solution, onto nonselective or selective solid medium to determine the number of colony-forming units (CFUs). The plating can be done using, for example, a Drigalski spatula, glass beads or a spiral plater (Gilchrist et al., 1973). The latter has the advantage that quantitation of CFUs is possible without prior dilution of the leaf wash sample. As with the leaf printing technique, medium composition of the agar plate determines what subpopulation of the phyllosphere will be analysed. This analysis should also take into consideration the fact that different organisms from the leaf surface grow at different rates, so that those bacteria or fungi that grow fast and appear on agar plates first will be more likely to be included in the analysis

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than those that grow slowly and appear late or not at all due to overgrowth by fast appearing strains. From the number of CFUs per plate the number of bacteria that were present on the leaf can be estimated quantitatively by taking into account the volume of the leaf washing solution, the dilution factor and the volume of the aliquot that was spread onto the agar plate or used to inoculate the medium in the MPN method. CFU numbers are usually expressed as the number of bacteria per leaf, per gram of leaf weight or per square centimetre of leaf surface. Often the number is expressed as 10 log(CFU), as this allows comparison of population counts from different leaves, which are often distributed not normally but lognormally (Hirano et al., 1982; Kinkel et al., 1995; Woody et al., 2003). The leaf washing method can be modified to obtain answers to different questions. For example, by not vortexing or sonicating the leaf in a wash solution, bacteria that occupy the leaf surface as aggregates can be removed as intact aggregates and be separated by filtration from those bacteria that live solitary. In this way, Morris et al. (1998) were able to estimate the fraction of bacteria that live in bacterial aggregates on the leaf surface. Incidentally, this aggregated lifestyle of bacteria may be a reason for underestimation of total bacterial populations on leaf surfaces: an aggregate of two or more bacterial cells will produce a single CFU on plates, which stresses the need for breaking up of these aggregates as much as possible (Miller et al., 2000). Another reason for underestimating microbial populations from leaf washing data is the resistance of some microorganisms to be readily removed from the leaf surface by leaf washing (Romantschuk, 1992). A good example is the group of pink-pigmented facultatively methylotrophic bacteria (Holland, 1997). This is probably one of the reasons why these particular bacteria are often missed in phyllosphere composition studies (Holland and Polacco, 1994). Also, plant-pathogenic fungi, such as downy and powdery mildews, produce structures that anchor the fungal hyphae into the plant leaf making them hard to remove (Section 11.3.2, and Chapter 12). To check the efficiency of any leaf washing method, one can use microscopy (Section 11.2.3) to validate the degree of removal from the leaf surface, for example, by staining with the fluorescent DNA stain 4,6-diamidino-2phenylindole (DAPI). In one instance this was done to check the removal efficiency of a less common sampling method which involves placing a leaf on sterile water, rapidly freezing the water, then carefully removing the leaf and collecting the ice which contains the transferred leaf microflora (Heuser and Zimmer, 2002). Whereas washing by vortexing and sonication usually leaves the cuticular surface covered by residual microorganisms (Jacques and Morris, 1995; Heuser and Zimmer, 2002), the freezing method almost completely removes microorganisms from the leaf (Heuser and Zimmer, 2002). A method with similar removal efficiency is leaf maceration, by which a single leaf or a collection of leaves is macerated, for example, with a mortar and pestle, in a defined volume of buffer, which is then analysed by plate counting. However, since leaf maceration liberates in addition to those microorganisms that live on

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the leaf surface also those that grow inside the plant tissue, phyllosphere population sizes may be overestimated by this method. One final point of consideration when using methods such as MPN or plate counts is that they provide estimates only for the culturable subset of the microbial population. Microorganisms that resist growing under laboratory conditions are not included in these analyses (also see Section 11.4.1). Furthermore, viablebut-not-culturable-microbial cells (VBNC) that experience prolonged periods of starvation may enter a physiological state which renders them not-culturable, that is, they cannot form a colony on an agar plate, but they are still viable (Bogosian and Bourneuf, 2001). This concept also applies to cuticular surface colonisers. For example, Wilson and Lindow (1992) showed that up to 75% of the bacteria on inoculated bean leaves were VBNC, demonstrating that plate counts can grossly underestimate the viable fraction of bacteria on a leaf surface. All of the sampling techniques described so far are in essence destructive, that is, the population size of a single leaf can be measured only once, at which time the experiment for that leaf ends. This makes it impossible to follow the temporal dynamics of a microbial population for a single leaf, only for leaves picked at different time points. But because the variation among leaves at any time point can be substantial (Hirano et al., 1982; Jacques and Morris, 1995; Hirano and Upper, 2000), sample sizes necessarily have to be large (i.e. many leaves should be analysed for each time point) in order to obtain statistically significant and precise estimates for changes in microbial population sizes over time. To circumvent the collection of large sample sizes, a semi-destructive sampling method can be used such as the one developed by Woody et al. (2003) for studying the temporal population dynamics of the phyllosphere yeast Aureobasidium pullulans. Its success is based on the prerequisites that (1) sampling of a leaf segment does not affect the yeast population on other parts of the same leaf, and (2) the distribution of yeast cells is similar for different segments of the same leaf. These prerequisites cannot a priori be assumed to be true for every other combination of microorganism and plant (Woody et al., 2003). Microorganisms that are sampled from the phyllosphere may be used not only for the purpose of estimating phyllosphere population sizes; leaf washings have also been used to inoculate BIOLOG EcoPlates to obtain a carbon-metabolism profile for the microbial community as a whole (Yang et al., 2001). Microorganisms washed from a leaf may be examined microscopically (Section 11.2.3), for example, after staining with DAPI or another DNA dye such as acridine orange, after live/dead staining (Monier and Lindow, 2003a) or after hybridisation with rRNA-specific probe(s) using fluorescent in situ hybridisation (FISH) (Joyner and Lindow, 2000; Brandl et al., 2001; Leveau and Lindow, 2001). Microorganisms that have been modified to express the green fluorescent protein (GFP) can be analysed for individual GFP content using fluorescent microscopy and image analysis (Joyner and Lindow, 2000; Brandl et al., 2001; Leveau and Lindow, 2001; Miller et al., 2001) or flow cytometry (Axtell and Beattie, 2002; Marco et al., 2005). Leaf washings may also be examined for the activity of other reporter genes, such as inaZ (Miller

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et al., 2001). Furthermore, leaf washings may be the starting material for the isolation of microbial DNA for a culture-independent assessment – either qualitative or quantitative – of leaf surface microbiology (Yang et al., 2001; Heuser and Zimmer, 2002, 2003).

11.2.2 Artificial inoculation Experimental manipulation of phyllosphere composition and abundance has contributed a great deal to our understanding of the cuticular surface as a microbial habitat. It is achieved by artificial inoculation of plants in the field, greenhouse or laboratory with defined suspensions of bacteria, yeasts or fungal spores. The most common purposes of artificial inoculation are: (1)

to determine the ability of the inoculated microorganism(s) to survive and thrive in the phyllosphere; (2) to follow the growth and/or activity of the introduced strain(s) in relation to factors such as plant and environment; (3) to assess the effect of the introduced strain(s) on the abundance and activity of other microorganisms in the phyllosphere; or (4) to exploit the introduced microorganism(s) as a biological indicator of the physical, chemical or biological factors that govern the phyllosphere. In the greenhouse or laboratory, inoculation of plants is achieved by immersing whole plants in a dilute suspension of microorganisms with a known titer, or by spraying them with such a suspension using a device that produces a fine mist or spray. After inoculation, plants are incubated in one of many ways, depending on experimental design and research question. To create an environment that is conducive to microbial growth, it is most common to incubate under conditions of high relative humidity, for example, by covering the plants in bags or placing them in controlled-environment chambers. Plants have been exposed to diverse conditions in order to see if and how these conditions change microbial population sizes and activities. Some examples are different light intensities (O’Brien and Lindow, 1989), low relative humidity (O’Brien and Lindow, 1989; Andersen et al., 1998), or increased ultraviolet (UV) exposure (Kadivar and Stapleton, 2003; Jacobs et al., 2005). In the field, plants are spray-inoculated using knapsack sprayers or scaled-up devices. Most field inoculations are performed to test or exploit the ability of the microorganism(s) in the inoculum to suppress symptoms of disease or microbe-induced freezing (Section 11.9.3). To some extent, environmental conditions in the field can be manipulated, for example by covering plants with meshes to test the effect of rainfall momentum or exposure to sunlight (Upper and Hirano, 2002). Inoculation of plants can be done with single strains or with mixtures of strains. By mixing two strains in a 1 : 1 ratio and using this mixture as an inoculum, the two strains in the mixture can be directly compared for their relative fitness in

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the phyllosphere (Section 11.6.1). By this approach small but significant differences in behaviour between two species can be revealed better than when the two strains are inoculated separately (Lenski, 1992). Mixtures may also be used to test the effect of strains on each other’s performance in the phyllosphere. So-called de Wit replacement series, whereby two strains are mixed in various proportions at a constant total inoculum density, have provided important insights into the relative competitive abilities of microbial epiphytes, niche differentiation and resource utilisation (Wilson and Lindow, 1994a, 1995). Factors that have been shown to contribute significantly to the phyllosphere performance of inoculated microorganisms are preparation of the inoculum and inoculum density (Wilson and Lindow, 1994b). Each of these factors should be taken into consideration during the planning of artificial inoculation experiments and interpretation of their results. One special type of inoculum involves microbial bioreporters for habitat exploration (Leveau and Lindow, 2002). Bioreporters are whole-cell indicators of a specific microbial activity. They are usually bacteria that have been manipulated to carry a reporter gene such as the one coding for GFP or ice nucleation protein (InaZ) downstream of an inducible promoter. This promoter determines the usefulness and specificity of the bioreporter strain. For example, Leveau and Lindow (2001) introduced into the epiphytic bacterium Erwinia herbicola a fusion of a fructose-inducible promoter and the gene for GFP. The resulting strain of E. herbicola responds to the availability of fructose by the synthesis of GFP and emission of green fluorescent light. Inoculation of this strain onto plants made it possible to explore the leaf surface for the availability for fructose (see also Section 11.5.2). Several other bioreporters, with different specificities, have been applied to the phyllosphere (Joyner and Lindow, 2000; Miller et al., 2001; Axtell and Beattie, 2002), and each has provided unique insights into the microbial perception of the cuticular surface (Sections 11.5.2 and 11.7.3). The concept of bioreporting has also been used for the identification of genes that contribute to the fitness of epiphytic bacteria in the phyllosphere (Sections 11.6.1 and 11.6.3). Marco et al. (2003) developed a screening method for phyllosphereinducible promoters of Pseudomonas syringae which is based on the insertion of random genomic DNA fragments upstream of a promoterless but essential locus metXW for methionine biosynthesis. When a mixture of bacteria, each containing a different gene fusion, is inoculated onto plants only those bacteria that carry a phyllosphere-inducible promoter express the metXW genes and thus will survive. In other words, by inoculation onto plant leaves, bacteria with phyllosphereinducible promoters are enriched. This approach has been very useful in the identification of genes that are involved in the adaptation of bacterial colonisers to the phyllosphere (Marco et al., 2005; also see Section 11.6.3).

11.2.3 Microscopy The openness of the phyllosphere makes it extremely suitable for direct observation by microscopy. In this respect, the phyllosphere differs dramatically from

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the rhizosphere, which essentially is a hidden environment. Microscopical analysis can be done on whole leaves, leaf sections or even isolated leaf cuticles. The highest magnification, that is, 10 000× or more, is achieved by electron microscopy, and both transmission and scanning electron microscopy have revealed interesting details of phyllosphere life at the (sub)micrometre scale (Beattie, 2002). Environmental scanning electron microscopy is a modification of the latter technique which allows ‘wet-mode’ imaging of phyllosphere samples (Monier and Lindow, 2004). Atomic force microscopy has also been applied to the phyllosphere (Mechaber et al., 1996) to create three-dimensional, high-resolution surface maps of the leaf (see Section 11.5.1). More accessible to most phyllosphere researchers is light and fluorescence microscopy which generally have a maximum magnification of 1000× and a resolution of about 0.2 micrometres. One of the most exciting developments in phyllosphere research has been the combination of fluorescent microscopy and fluorescent proteins such as GFP (Brandl et al., 2001; Leveau and Lindow, 2001; Axtell and Beattie, 2002; Brandl and Mandrell, 2002; Monier and Lindow, 2003a) but also red-fluorescent protein (Brandl and Mandrell, 2002). Bacteria or fungi that have been modified to express GFP constitutively are easily recognised in situ on the leaf surface by their green fluorescence. Thus, colonisation patterns can be related to leaf surface structures. Furthermore, the activity of GFP-based bioreporters (as discussed in Section 11.2.2) can be interpreted in light of their location on the leaf surface (Leveau and Lindow, 2001). With confocal laser scanning microscopy (CLSM), individually sliced views in the z-axis can be viewed and stacked. CLSM has been used quite successfully to reconstruct the three-dimensional arrangement of microorganisms on the leaf surface (Figure 11.1). For epiphytic microorganisms that occur naturally on leaves or that have not been modified to express fluorescent proteins, different stains or dyes are available

Figure 11.1 View of a colonised bean leaf using confocal laser scanning microscopy. Individual bacterial colonisers producing GFP are visible as green fluorescent dots. The centre shows a stoma; the cuticular top of an epidermal plant cell is visible on the left top corner while at the bottom right the picture slices into the leaf’s palisade parenchyma. Chlorophyll is coloured red in this picture. This figure is produced in colour in the colour plate section, which follows page 249.

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to facilitate their visualisation. Most practical are those stains that can be used in combination with fluorescence microscopy. These include DNA stains like DAPI and acridine orange, and propidium iodide which stains dead bacteria red (Monier and Lindow, 2003a, 2004). A more specific type of dye is represented by fluorescently labelled oligonucleotides or probes, which are designed to target the rRNA in ribosomes in a procedure that is referred to as FISH (Amann et al., 1995). FISH probes can be labelled with different fluorophores, but carboxytetramethylrhodamine (TAMRA) has been most commonly used in phyllosphere research. TAMRA fluoresces red and its detection is compatible with that of GFP (Joyner and Lindow, 2000; Brandl et al., 2001; Leveau and Lindow, 2001).

11.3

Getting to the phyllosphere (and leaving again)

11.3.1 Immigration When a leaf appears on the plant, it is generally not sterile, that is, free of microbial inhabitants. Seeds that are naturally or artificially inoculated with bacteria have been shown to develop into plants that carry substantial populations of these bacteria on their leaves (Saettler et al., 1989; Lilley et al., 1997; Upper and Hirano, 2002). Thus, seeds and, indirectly, soil may be potential sources of phyllosphere colonisers. However, most microorganisms in the phyllosphere of naturally occurring plants are considered to arrive only after leaves have emerged by immigration from the air surrounding the leaf (Lindemann and Upper, 1985; Lindow and Andersen, 1996; Kadivar and Stapleton, 2003). The atmosphere is a rich and diverse source of potential microbial colonisers of the phyllosphere. In one study (Lighthart and Shaffer, 1995), the 10-m layer of air over a grass field in Oregon contained an average value of 121 and a peak value of 1369 bacterial CFUs per cubic metre. Plate counts fluctuated greatly between long-term (early, mid and late summer), short-term (morning, afternoon, evening, night) and even very short-term (intervals of 2 min) sampling times. Much of the short- and long-term fluctuation could be attributed to the temporal variation in meteorological conditions (convective wind sweepings, cleansing sea breezes) as they were recorded during the course of the day or season. Several mechanisms have been proposed for microbial immigration from the air onto the leaf surface (Lindow, 1996), including deposition by aerosol, wind, rain or insects (Lilley et al., 1997). Several studies have produced quantitative data for these processes, which allow a better appreciation for the contribution of immigration to phyllosphere biology. For bean plants in a Wisconsin field, Upper and Hirano (2002) reported an average phyllosphere immigration rate of 11 P. syringae bacteria per bean leaf on a rain-free day. Rates of immigration were clearly dependent on the presence of and distance from source plants (i.e. plants carrying high population sizes of P. syringae). From airborne insects that were captured in the same field, bacteria could be recovered in numbers that varied from undetectable to 10 000 CFU

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per insect (Upper and Hirano, 2002). Dispersal of bacteria by airborne insects appeared to be most effective when leaves were wet and accounted dispersion over up to tens of metres. Leaf-to-leaf migration may also occur through run-off or splash during rains, but this mechanism of dispersal obviously operates at a smaller, local scale within the plant leaf canopy. Bacterial cells and fungal spores may not arrive on the leaf surface as individuals, but rather in aggregates of more than one individual. The close proximity of these early immigrants to each other on the leaf surface and the interaction that follows between them probably is a first determinant in the fate of these immigrants during the next stages of leaf colonisation. Much of the success of an immigrant to the phyllosphere also depends on the initial physico-chemical conditions of the cuticular surface (Section 11.5.2; Chapter 4). For example, it has been shown that under conditions of low relative humidity, the chances of survival of P. syringae on bean leaves is reduced dramatically compared to conditions of high relative humidity (Monier and Lindow, 2003a). Furthermore, immigrants to a previously colonised leaf surface probably face an environment that is quite different from that of an uncolonised leaf surface. On the one hand, they might enter an environment that, through active modification by the residing microflora, has been made more conducive to their own survival or growth (Section 11.8.1). On the other hand, much of the initially available nutrients on colonised leaves may have been used up by earlier immigrants, leaving less substrate to utilise and to grow on for later immigrants (Section 11.7.3). This principle has been used to explain the relative success of early immigrants in dominating the phyllosphere (Upper and Hirano, 2002) and has been exploited as a biocontrol strategy for the suppression of certain plant pathogens (Section 11.9.3).

11.3.2 Adhesion Adhesion of epiphytic microorganisms to cuticular surfaces plays several ecological roles. First, it prevents removal from the leaf surface, either immediately after immigration or in the subsequent course of colonisation. It has been suggested, however (Andrews and Buck, 2002), that for many bacteria and yeasts the contribution of adhesion to phyllosphere colonisation is relatively small if the fraction of cells that resists removal and remains on the leaf surface is able to quickly re-populate the leaf surface. A second role of attachment is that it serves as the initial step in the formation of microcolonies, aggregates and biofilms (Andrews and Buck, 2002), which has been implicated in enhanced success of leaf colonisation, for example through niche modification (Section 11.8.1). The surfaces of leaves and fruits are covered by a hydrophobic cuticle which consists of a complex mix of long-chain aliphatic compounds (Chapters 1 and 4). Many fungi adhere to hydrophobic surfaces, including cuticular surfaces, much better than to hydrophilic surfaces (Clement et al., 1994; Buck and Andrews, 1999; Tucker and Talbot, 2001; Beattie, 2002), although exceptions exist (Buck and Andrews, 1999). Underlying the hydrophobicity mediated adhesion of yeasts, fungi and fungal spores

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are hydrophobic interactions between cuticular surface waxes and hydrophobic components in the cell wall such as water-insoluble glycoproteins (Tucker and Talbot, 2001). When leaf waxes are removed, for example, with chloroform, hydrophobicity dependent attachment is reduced (Young and Kauss, 1984). Attachment may also be mediated by fungal-produced components including extracellular polysaccharides (Andrews et al., 1994), hydrophobic mucilage (Hamer et al., 1998) and cutinases and esterases (Deising et al., 1992). The synthesis of such components is often influenced by the availability of water and nutrients (Andrews and Buck, 2002) or temperature (Tucker and Talbot, 2001). Adhesion to the cuticular surface may also be indirect: among phyllosphere yeasts, it is a common trait to be able to attach to fungal spores or hyphae (Allen et al., 2004). Quite a rigorous mode of fungal adhesion to the cuticular surface is through cuticle penetration by specialised hyphae (Chapter 12). Penetration is achieved by secretion of enzymes and/or by sheer physical force. With the subsequent formation of fungal structures inside the leaf (e.g. haustoria), fungal attachment is semi-permanent. In contrast to fungi, bacteria do not seem to engage in hydrophobic interactions with cuticular surface waxes. Instead, they rely on pili and extracellular polymeric substances for adhesion (Beattie, 2002). Most research has focused on the role of pili (Romantschuk, 1992), which are thin protein tubes originating from the cytoplasmic membrane and which are found in almost all gram-negative bacteria. Mutants of P. syringae lacking pili were washed more easily from leaves than the wildtype (Suoniemi et al., 1995), which suggests a role for pili in resisting removal by runoff rain. In field experiments, mutants of P. syringae pv. tomato lacking type IV pili achieved slightly lower population sizes on leaves of tomato plants than those of the parental wild-type strain DC3000 (Roine et al., 1998). Adhesion of piliated P. syringae was not affected by removal of the cuticular surface waxes, suggesting that waxes are not necessary for pili attachment (Romantschuk et al., 1993). Besides pili binding, several other mechanisms have been proposed to explain bacterial adhesion to plant leaf surfaces. Many bacteria produce extracellular polysaccharides on leaves, often as a matrix for biofilms (Romantschuk et al., 1996; Morris et al., 1997; Beattie and Lindow, 1999). Acting as glue, extracellular polysaccharides may help reduce the probability of removal by rain or wind. The same has been suggested for the production of cellulose by Pseudomonas fluorescens SBW25 (Gal et al., 2003). Bacterial attachment to leaves varies with bacterial species and with cuticular surface properties. Immediately after spraying suspensions of Pantoea agglomerans, Clavibacter michiganensis or P. syringae onto maize leaves, all three bacterial species resisted to some extent the removal from the leaf by sonication and vigorous vortexing, but the fraction of non-removable P. agglomerans cells was up to ten-fold higher than that of the other two species (Beattie and Marcell, 2002). Furthermore, it was shown with cuticular maize mutants that P. agglomerans was less influenced by changes in cuticular wax-dependent surface properties than C. michiganensis (Beattie and Marcell, 2002), suggesting that different bacteria use different mechanisms for adhesion. It is unclear whether these differences in mechanisms correlate

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with the relative importance of adhesion in the life cycle of any given bacterial epiphyte.

11.3.3 Emigration Many of the mechanisms by which microorganisms exit the surfaces of leaves are the reverse of those described for immigration (Section 11.3.1). Emigration may account for a substantial reduction in phyllosphere abundance. For example, rainfall may remove as many as 105 bacteria from a single bean leaf in the field in a period of 15 min (Lindemann and Upper, 1985). In this process, leaf cuticle properties may play an important role. For example, mutants of corn that differ in wax composition show less retention of sprayed bacteria when cuticular surfaces are more hydrophobic (Beattie, 2002). Microorganisms that run off leaves through rainfall onto the soil usually do not persist there for long. However, they might attach to seeds and end up on the leaves of newly emerging plants (Section 11.3.1).

11.4

Microbial communities in the phyllosphere

11.4.1 Composition To study the composition of microbial communities in the phyllosphere, the most common procedure is to spread leaf washings onto nonselective or selective solid media (Section 11.2.1), and identify the bacteria, yeasts or fungi that appear on the basis of morphological, physiological or phylogenetic features. Thus, composition analysis is generally biased towards culturable microbial epiphytes only, and in most cases it is not known how representative this subset is for the entire phyllosphere population. The bias towards culturables probably also underlies the common notion that populations of culturable bacteria on leaves are dominated by only a few genera including Pseudomonas, Erwinia and Xanthomonas (Jurkevitch and Shapira, 2000; Lindow and Brandl, 2003). All of these are readily culturable bacteria that grow rapidly on laboratory media. Pink-pigmented facultative methylotrophs of the genus Methylobacterium are some of the most abundant bacteria on plants but they are often overlooked due to their comparatively slow growth and special nutrient requirements: they are only found when searched for, and so far, they have been encountered on nearly every plant investigated (Corpe, 1985; Holland and Polacco, 1994). Very few studies have used culture-independent methods to avoid the bias towards culturables, and their outcome, not surprisingly, shows a much greater diversity in phyllosphere composition (Weidner et al., 2000; Yang et al., 2001; Kadivar and Stapleton, 2003). For example, by denaturing gradient gel electrophoresis (DGGE) of PCR-generated 16S ribosomal RNA gene products, Yang et al. (2001) identified 17 unique sequences in DNA collected from the ‘Valencia’ orange phyllosphere. Only four of these corresponded to bacteria that had been found in

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the phyllosphere before, that is, Acinetobactor sp., Bacillus pumilus, Enterobacter agglomerans and a Cytophagales species. In contrast, 16S rRNA sequences obtained from leaf washings that were first cultured in BIOLOG EcoPlates showed between 97 and 100% similarity to those of known phyllosphere bacteria such as Pseudomonas, Erwinia and Acinetobacter species, clearly demonstrating that culturing favours the culturable. Many composition analyses have targeted plants from a single geographical area (Inacio et al., 2002; Pereira et al., 2002) or a single plant species (Austin et al., 1978; Ercolani, 1991; Jurkevitch and Shapira, 2000; de Jager et al., 2001). While such studies may seem rather anecdotal, also given the fact that composition is highly variable (Section 11.4.3), they are instrumental in demonstrating the enormous diversity of microbial life in the phyllosphere. Snap bean plants were shown to carry more than 78 bacterial species representing 37 bacterial genera on their leaves (Beattie and Lindow, 1999). An inventory of phyllosphere fungi and yeasts from Mediterranean plants included a total of 1029 strains of filamentous fungi and 540 strains of yeasts, representing at least 36 and 46 distinct species, respectively (Inacio et al., 2002). Jacques and Morris (1995) presented an overview of bacterial species, representing 29 genera, isolated in 13 independent studies from plants in temperate zones. For at least 3 plants (Lolium perenne, Olea europaea and Ranunculus penicillatus), 3–27% of the bacteria that were isolated and cultured from the phyllosphere could not be identified, showing that our understanding of the microbial composition of the phyllosphere, even for culturable representatives, is still far from complete. Yet, this realisation also comes from culture-independent studies: 5 of the 17 rRNA sequences identified by Yang et al. (2001) had a similarity to database entries below 90%, suggesting that they represent previously undescribed taxa. The degree to which plants can determine the microbial composition in their phyllosphere remains unknown. Using DGGE (Yang et al., 2001), it was shown that the microbial community structure on plants such as green bean, cotton, sugar beet and orange, but not corn, were similar on different individuals of the same plant species, but different from all other plant species. If selection takes place it would probably be on the basis of leaf characteristics such as the presence or absence of protective sites (Section 11.6.2) or the availability of plant-specific nutrients (Section 11.7.3), in combination with environmental factors such as weather conditions or geographical location.

11.4.2 Abundance In general, microbial abundance in the phyllosphere refers to one of the following: (1) the total average number of bacteria, yeasts or fungi on a leaf, (2) the average number of a specified subset of bacteria, yeasts or fungi on a leaf, (3) the most abundant species of bacteria, yeasts or fungi on a leaf, or (4) the number of bacteria, yeasts or fungi on a dimensional scale smaller than that of a single leaf. As it is the case for composition studies, most estimates for microbial abundances on leaf

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surfaces are made on the basis of counts of culturables (Section 11.2.1), although some recent reports have used culture-independent methods such as real-time PCR (Heuser and Zimmer, 2002, 2003) and direct microscopic counting (Andrews and Buck, 2002). To be able to compare between plants, microbial abundances are best described as densities, for example, numbers of microorganisms per square centimetre of leaf surface, but they have also been expressed per leaf or per gram of leaf tissue. Of all microbial epiphytes, bacteria are generally considered to be the most abundant on leaves (Lindow and Brandl, 2003). A typical leaf may contain up to 106 –107 bacteria per square centimetre of surface. Scaled up to human dimensions, this translates into population densities that are 10- to 100-fold higher than the most densely populated countries in the world. The number of yeasts and yeast-like organisms may reach 107 CFU per gram (fresh weight) of leaf material on some plant species. The most abundant yeasts and yeast-like fungi on the phylloplane of temperate plants are Aureobasidium pullulans, Sporobolomyces species, Rhodotorula species and Cryptococcus species (McCormack et al., 1994). Microbial abundances in the phyllosphere are characterised by high degrees of variability, a factor 1000 and sometimes more, among leaves from the same plant or field (Hirano et al., 1982; Jacques and Morris, 1995; Hirano and Upper, 2000). Leaf densities of microorganisms are therefore generally described and compared after a 10 log transformation. The degree of leaf-to-leaf variation may differ between plant species or change as a result of environmental conditions (Kinkel et al., 2000). It has recently been suggested (Woody et al., 2003) that leaf-to-leaf variability is not due to asynchronous temporal changes in population sizes on individual leaves, but rather due to the variation among individual leaves in their ability to promote survival or sustain microbial growth (Sections 11.6 and 11.7). Variation in population densities is not restricted to the dimensional scale of single leaves. Even on different sections from the same leaf there can exist substantial variation in microbial numbers. For example, among 9-mm2 sections of potato leaves total bacterial populations varied by over 100-fold (Kinkel et al., 1995), and on bean leaf sections of 44-mm2 , P. syringae populations even varied 100 000-fold (Monier and Lindow, 2004). Also at the microscopic level bacteria, yeasts and fungi are not uniformly distributed across the leaf surface (Monier and Lindow, 2004). The most commonly and heavily colonised sites on naturally colonised plant leaves are the bases of trichomes, stomata, epidermal cell wall junctions and the grooves along veins (Beattie and Lindow, 1999). It has been hypothesised that these sites are ‘protective’ sites where microbial growth is favoured more than on other sites on the leaf surface, for example, because of higher nutrient availability (Section 11.7.3). From microscopy studies, it has also become clear that many bacteria occupy the leaf surface in aggregates (Morris et al., 1997). In fact, aggregates can constitute between 10 and 40% of the total bacterial population on leaves of certain plant species (Morris et al., 1998). On leaves from the field, these aggregates are often mixed populations of different species (Morris et al., 1998). Laboratory studies of aggregate formation by phyllosphere bacteria have revealed that aggregate sizes were distributed with a strong right-hand-skewed frequency (Monier and Lindow, 2004). While large

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aggregates on these bean leaves were not frequent, they accounted significantly for the majority of cells present on a leaf.

11.4.3 Dynamics Many studies have documented differences in microbial abundance and composition on leaves from the same plants or field at different points in time or after a defined treatment (O’Brien and Lindow, 1989; Ercolani, 1991; Ellis et al., 1999; Hirano and Upper, 2000; de Jager et al., 2001; Kadivar and Stapleton, 2003). This demonstrates that microbial populations on cuticular surfaces are not static. Changes in abundance or composition are ultimately a sum of immigration, growth, death and emigration, and each of these is directly or indirectly influenced by external factors such as rain, UV radiation exposure and leaf age. Rains have been shown to trigger rapid multiplication of epiphytic bacteria (Hirano and Upper, 2000), probably due to increased nutrient availability as a result of leaching (Section 11.7.2). Exposure of plants to conditions of low relative humidity usually causes bacterial populations to decrease and/or change in composition (O’Brien and Lindow, 1989; Hirano and Upper, 2000). Weather conditions may also determine the most abundant colonisers of leaves: for example, on bean leaves in the field, PPFMs were most abundant when the weather was hot and dry, whereas P. syringae was quite abundant during periods of wet and warm weather (Hirano and Upper, 2000). Changes in UV exposure have been shown to affect microbial populations on leaf surfaces (Newsham et al., 1997; Sundin and Jacobs, 1999; Jacobs and Sundin, 2001, 2002; Hughes et al., 2003; Jacobs et al., 2005). In one instance, UV-exposed phyllosphere samples showed an increase in bacterial diversity (Kadivar and Stapleton, 2003). Leaf age also has a clear effect on microbial composition (Ercolani, 1991). While bacteria, yeasts and filamentous fungi are often found to colonise the same leaf surface, there can be succession in microbial composition as the leaf gets older. In general, bacteria are the pioneers on young leaves, yeasts may dominate as the leaf becomes older and filamentous fungi up to leaf senescence (Blakeman, 1985). Other factors that may influence the density and diversity of microbial populations in the phyllosphere are plant species (Heuer and Smalla, 1997), air pollution (Brighigna et al., 2000), carbon dioxide levels (Magan and Baxter, 1996), leaf position in the canopy (Andrews et al., 1980; de Jager et al., 2001), acid rain (Helander et al., 1993) and insects (Stadler and Müller, 2000).

11.5

Microbial perception of the phyllosphere

11.5.1 Topography As the dimensions of humans and microorganisms differ by about six orders of magnitude, only a 1 000 000-fold magnification could offer us the best possible impression of leaf surface topography from the perception of an individual microbial

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epiphyte. Electron microscopy commonly achieves a 60 000-fold zoom, while light and fluorescence microscopy maximally reaches 1000-fold, offering (only) a bird’s eye view of the habitat. There are relatively few studies that provide quantitative data on the topography of leaf surfaces (Mechaber et al., 1996; Monier and Lindow, 2004). This is surprising, since leaf surface structures have been implicated to have a large influence on the microbial biology of the phyllosphere. Structurally and functionally different features of the leaf such as stomata, veins, trichomes and epidermal cells may vary dramatically in water retention capacity, cuticle thickness or release of plant compounds; so their individual ability to harbour and sustain microbial populations may also differ. Monier and Lindow (2004) scanned adaxial surfaces of bean leaves and found them to consist 74% of undifferentiated epidermal cells, while stomates, veins and trichomes accounted for 17, 7 and 2%, respectively. Using atomic force microscopy, Mechaber et al. (1996) mapped the adaxial surfaces of cranberry leaves in three dimensions. Young leaves showed a regular pattern of broad plateaus on the surface of individual epidermal plant cells with drops in elevation of up to 3 μm between the cells. Old leaf surfaces on the other hand appeared much rougher and with a less regular pattern. These differences between young and old leaves indicate that leaf surface topography and thus physical properties are highly variable with age, and the same is true for leaves of different plant species.

11.5.2 Physico-chemical parameters The phyllosphere represents a unique habitat in terms of physical and chemical parameters (Burrage, 1971; Chapter 4). Water may be available in the form of rain, fog or dew. In addition, the cuticle may in some cases be covered by aqueous deposits which are a result of the interaction between hygroscopic salt crystals on the cuticular surface and water vapour from the atmosphere and the stomata (Burkhardt et al., 1999). The wetness of individual leaf surfaces can be quantified by measuring electrical conductivity between two electrodes clipped to a living plant leaf surface (Klemm et al., 2002). To assess water availability at a smaller, that is, micrometre scale, bacterial bioreporters have been used (Axtell and Beattie, 2002). Cells of E. herbicola carrying a proU-gfp gene fusion produce GFP in a quantitative manner in response to low water potential. After inoculation onto bean leaves, GFP expression profiles indicated that cells experienced increased osmolarity, probably as a result of evaporation of the available water. However, there was considerable variation among individual bacteria on the leaf surface: those that were located close to veins were less fluorescent, suggesting that such leaf structures can locally increase water retention. Other physico-chemical parameters are UV exposure and temperature. UV radiation from the sun reaching the phyllosphere consists on the average 95% of UV-A (320–400 nm) and 5% UV-B (290–320 nm; Jacobs et al., 2005). The effects of these types of radiation on microbial epiphytes differ: UV-A exposure leads

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to the (sub)lethal formation of reactive oxygen species, whereas UV-B directly damages the DNA. UV exposure of individual leaves differs with geography, climate and canopy structure, but UV-protective measures are common among many phyllosphere microorganisms (Section 11.6.2). Temperatures can differ dramatically in the course of a single day, but also on a single leaf, for example, from the centre to the edge. Temperature may have a direct effect on the growth of microorganisms, but may also act indirectly, for example, through accelerated or delayed evaporation of available water.

11.5.3 Biological environment The area of the leaf surface that is actually covered by microorganisms varies depending on many factors. In a laboratory experiment with P. syringae, bean leaves were covered for up to 12% of their surface area with bacteria, whereas some 60% of the cells were located in an aggregate of 100 cells or more (Monier and Lindow, 2004). This means (1) that any new bacterial immigrant to these leaves would have a relatively low probability (up to 12%) of landing next to other bacteria, but (2) that any already present inhabitant has a high probability (60% or more) to be surrounded by many others. Bacteria in an aggregate are presented with a biological environment which is very different from that of a solitary cell and which probably sets the scene for fierce competition for food and space, and also opens the possibility for collaboration through density-dependent communication (Section 11.8.4) or for plasmid exchange (Section 11.8.5). In laboratory studies with single bacterial strains, aggregates mostly or exclusively consist of clonal individuals, whereas in nature, aggregates on leaves are most commonly mixes of different bacterial species (Morris et al., 1998). Surprisingly little is known about if and in what way bacteria on the one hand and fungi or yeasts on the other interact in the phyllosphere. Most of what is known comes from biocontrol studies (Section 11.9.3), which leaves many questions on the biology and ecology of bacteria/fungi interactions in the phyllosphere still unanswered.

11.6

Surviving (or not) in the phyllosphere

11.6.1 Concept of epiphitness Epiphytic fitness, or ‘epiphitness’ as it shall be referred to here, can be defined as the ability of a microorganism to survive and thrive on plant cuticular surfaces. Typically, this ability is demonstrated experimentally in the laboratory by inoculation of plants and subsequent monitoring of changes in population sizes. Epiphitness is therefore often expressed as a population size after a defined period of time after inoculation, or as an x-fold increase compared to population sizes immediately after inoculation. Generally, epiphitness is used as a relative measure: if under the same experimental circumstances the population size of one strain is lower than that of another, it is

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said that the former has a lower epiphitness than the latter. An example is the comparison of near-isogenic strains that differ in one gene or more – often a mutant versus the wildtype – to assess the role of that gene in phyllosphere competence (Section 11.6.3).

11.6.2 Adaptive strategies The cuticular surface is considered a harsh environment not only because of the relative extremes to which epiphytic microorganisms are exposed (e.g. water stress, nutrient availability, UV radiation), but perhaps even more so because of the rapidity with which such conditions change, for example, even within the time it takes a bacterium to double. Beattie and Lindow (1999) proposed two not mutually exclusive strategies that allow epiphytic bacteria to survive and thrive on plant cuticular surfaces. The first is a strategy of tolerance, which is based on traits that protect microorganisms against and help them to deal with the harsh environment of the phyllosphere. A good example is UV-protective pigmentation which is quite common among phyllosphere bacteria (Goodfellow et al., 1976; Dickinson, 1986; Lindow and Brandl, 2003; Jacobs et al., 2005), as are repair mechanisms for UV-induced damage to the DNA (Kim and Sundin, 2000; Sundin et al., 2000; Zhang and Sundin, 2004). In response to nutrient limitation in the phyllosphere, the bacterium P. syringae reduces its size (Björklöf et al., 2000; Monier and Lindow, 2003b), which is an active process, probably to optimise surface-to-volume ratio and nutrient uptake capacity (Monier and Lindow, 2003a). The phyllosphere fungus Epicoccum nigrum accumulates solutes such as glycerol and arabitol in response to water stress, which may assist in its survival and establishment on leaf surfaces (Pascual et al., 2003). The ever-changing conditions in the phyllosphere demand a high degree of plasticity from microbial epiphytic colonisers. There are several lines of evidence that suggest such plasticity in phyllosphere bacteria. For example, cells of P. syringae that were recovered from bean leaf surfaces and re-applied to uninoculated leaves showed a higher epiphitness than cells that were applied to leaves from a plate or broth culture (Wilson and Lindow, 1993). It has been proposed that nutrient limitation, in particular starvation for carbon, makes bacterial cells more adapted to the stressful conditions of the phyllosphere (Monier and Lindow, 2003a). Bacterial plasticity is probably correlated to the ability to go from a solitary lifestyle to life in an aggregate (see later), and vice versa (Boureau et al., 2004). Inability to tolerate the phyllosphere probably results in death or transition into a VBNC state. This may be the fate of many new microbial immigrants to the leaf or to those that are not part of a protective environment such as described later. It is unknown whether VBNCs can recover from their state, and under what conditions such recovery would occur. The second strategy proposed by Beattie and Lindow (1999) is a strategy of avoidance, which is based on so-called protective sites on the leaf surface. Some foliar pathogens avoid many of the leaf surface stresses by escaping to the interior of the leaf. This niche poses its own challenges for survival (Manceau and

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Kasempour, 2002), but many bacteria have learned to cope with these. The abaxial side of a leaf may protect better against the damaging effects of UV than the adaxial side (Sundin and Jacobs, 1999). Trichomes or other leaf structures may also represent protective sites as they offer shading from UV radiation, or retain water. Bacteria living in aggregates or biofilms in fact also employ a strategy of avoidance, as they create a local environment that is more conducive to growth and survival (see Section 11.8.1).

11.6.3 Epiphitness genes The common definition of epiphitness genes is that they confer a measurable advantage to a bacterium, yeast or fungus in colonising a leaf surface. Most epiphitness genes have been identified in knock-out studies: the underlying hypothesis is that mutations in epiphytic genes would reduce the epiphitness of its carrier. Lindow (1993) identified 82 transposon mutants of P. syringae with altered behaviour on leaf surfaces. Several of these were not able to withstand desiccation stress (Lindow et al., 1993a). None of the mutants were disrupted in their utilisation of 31 different carbon compounds, suggesting that P. syringae has a broad spectrum of carbon sources it can use or has redundant genes for utilisation of the most important carbon sources. Several mutants showed decreased motility, osmotolerance and extracellular polysaccharide production suggesting that these are important traits for establishment in the phyllosphere. Mutants of P. syringae lacking the gac regulon were less fit on bean plants in the field (Hirano et al., 1997). The gac regulon consists of the two-component regulatory genes gacA and gacS, which in P. syringae regulate, among other things, swarming, the synthesis of quorum sensing signal molecules called N-acyl homoserine lactones (AHLs; see Section 11.8.4) and production of the extracellular polysaccharide alginate, all of which have been considered to play a role in the epiphytic lifestyle of this and other epiphytic bacteria (Yu et al., 1999). Genes of the hrp regulon also have been shown to contribute significantly to epiphitness in field situations (Hirano et al., 1999). Both gac and hrp genes were initially identified as being involved in pathogenicity, suggesting a close link between epiphitness and being a successful pathogen (Section 11.9.2). Several other examples of epiphytic genes that have been identified through mutation analysis are available (e.g. Andersen et al., 1998; Brandl and Lindow, 1998; Roine et al., 1998; Sundin, 1999); they code for such traits as pili formation, alginate synthesis, production of the plant hormone indole-3-acetic acid and AHL production. A well-recognised problem with knock-out approaches is their limited effectiveness in identifying genes with incremental contribution to leaf surface fitness (Marco et al., 2003). An alternative approach is the use of reporter gene technology, which aims to answer what genes are specifically expressed during colonisation of the phyllosphere. Cirvilleri and Lindow (1994) used random insertion of a promoterless lux gene into the genome of P. syringae to identify transposon mutants that were bioluminescent on leaf surfaces. This analysis revealed that about 3% of all P. syringae genes are expressed on the leaf surface. Independently,

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Marco et al. (2003) came to a similar estimate using in vitro expression technology by which random fragments of the P. syringae genome were screened for their ability to complement a conditionally lethal phenotype on bean leaves. Using this strategy, called HIRS for habitat-inducible rescue of survival, Marco et al. (2005) were then able to describe a number of phyllosphere-induced loci with anticipated or presumed epiphytic functions such as water stress tolerance or utilisation of organic sulfur. Interestingly, several virulence-associated genes were also identified by the HIRS method, indicating that P. syringae expresses virulence factors during leaf colonisation. To some loci, no function could be assigned on the basis of sequence homology of DNA or protein, suggesting that yet-to-identify epiphitness traits may exist.

11.7

Microbial growth in the phyllosphere

11.7.1 Growth requirements Individual epiphytic microorganisms have different nutritional requirements. For example, it does not take much for a bacterium to duplicate: it was calculated that the epiphytic bacterium E. herbicola needs 0.3 pg of sugar to double (Leveau and Lindow, 2001). But growth yield, that is, the efficiency with which food is converted into biomass, is not the only factor that determines the success of a leaf coloniser. There are three additional factors to be considered, namely versatility, affinity and growth rate. Microorganisms differ in the range of nutrients, for example, carbon sources that they can use for growth (Ji and Wilson, 2002). Comparison of the nutrient utilisation spectrum of microbial species with the presence or absence of specific nutrients on leaf surfaces could, in theory, predict whether a bacterium or fungus has the potential to grow in the phyllosphere. By comparing nutrient utilisation spectra of two or more different microbial species, it is possible to derive a so-called nutritional niche overlap index (Wilson and Lindow, 1994a; Ji and Wilson, 2002). It may be used as a predictor of co-existence or competition. A low index value, for instance, indicates that species vary greatly in their nutritional spectrum, so that there is a low probability for competition for the same nutrients and a high probability for co-existence. This has also been referred to as nutritional resource partitioning (Wilson and Lindow, 1994c). It should be noted that niche overlap indices may be influenced by external factors including water availability and temperature (Lee and Magan, 1999). Another factor determining growth in the phyllosphere is affinity, which divides microorganisms into those that can use nutrients at very low concentrations and those that cannot. This dichotomy probably overlaps for a large part with the separation of microbial epiphytes into so-called K- and r-strategists (Andrews, 1984). The K-strategists reproduce more slowly and tend to be successful in resource-limited situations, whereas r-strategists are characterised by a high growth rate and dominate in situations where nutrient resources are abundant.

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11.7.2 Types and sources of nutrients One of the most obvious sources of nutrients on the plant leaf and fruit surface is the cuticle. It consists of the cutin polymer and associated waxes with high carbon and energy contents, and so would seem a logical growth substrate. However, there is no evidence to support the theory that cuticle components are used by microorganisms for growth (Beattie, 2002). Exogenous nutrients may be available fortuitously in the form of pollen, honeydew, dust, air pollution or microbial debris (Stadler and Müller, 2000; Leveau, 2004). However, the major source of nutrients to microbial colonisers is represented by plant metabolites that leach from the leaf’s interior to the surface. Leaching is a passive process, and is stimulated by the presence of water on the leaf, for example, in the form of rain drops or fog. Leaf leachates contain a variety of compounds (Tukey, 1970), but the most abundant are photosynthates such as glucose, fructose and sucrose (Fiala et al., 1990; Mercier and Lindow, 2000; Leveau, 2004). Uninoculated bean plants in the greenhouse carry 0.2–10 micrograms of sugars per leaf (Mercier and Lindow, 2000), enough to support bacterial populations of 107 per leaf. The carrying capacity, that is, the maximum microbial population which a given leaf can support, is often correlated to the availability of sugars. This is in agreement with the observation that the availability of carbon, not nitrogen or phosphate, is generally limiting the sizes of bacterial populations in the phyllosphere (Wilson and Lindow, 1994a; Wilson et al., 1995; Mercier and Lindow, 2000). Other perhaps less known examples of carbon sources in the phyllosphere are methanol and methylamine which are plant waste products that are preferentially used by PPFMs (Holland and Polacco, 1994), and the plant hormone indole-3-acetic acid which also has been shown to be a substrate for growth by phyllosphere bacteria (Leveau and Lindow, 2005).

11.7.3 Nutrient bioavailability There are several lines of evidence to suggest that nutrients are not equally available on different leaves from the same plant or field, nor that they are evenly distributed across the surface of a single leaf. Uninoculated bean leaves contain on average 2.5 μg of surface sugar per gram of leaf (Mercier and Lindow, 2000), but this may vary by about 25-fold from leaf to leaf. The variation has been explained due to differences between leaves in, for example, cuticle leakiness (Mercier and Lindow, 2000), photosynthesis rates, for example, as a result of positional effects (Fiala et al., 1990), leaf age or plant nutrition. After inoculation and incubation of bean plants with the bacterium P. fluorescens, original sugar abundances were decimated, suggesting (1) that bacteria utilise sugars and (2) that any new immigrants would face nutritional conditions different from those the first immigrants did. Interestingly, a residual amount of sugars remained on the leaf surface, even after the bacterial populations reached carrying capacity (Mercier and Lindow, 2000). This has been explained by the heterogeneous distribution of sugars, that is, some sugars remain unavailable to the bacteria. By using the size of individual cells as an

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indicator for trophic status, it has been shown that P. syringae bacteria on a single bean leaf experience very different nutrient bioavailabilities (Monier and Lindow, 2003b). Bacteria near glandular trichomes or veins were larger than those located elsewhere, suggesting that such sites offer more nutrients. Glandular trichomes have been shown to secrete a number of plant compounds such as sugars, proteins, oils, secondary metabolites and mucilage, all of which may contribute to microbial growth. In addition, their ability to retain water would favour local leaching of nutrients. A pattern of high heterogeneity in sugar availability has been demonstrated with bacterial bioreporters for fructose. It was estimated that newly arrived E. herbicola bacteria on a bean leaf were exposed to local initial fructose abundances ranging from less than 0.15 pg to more than 4.6 pg fructose (Leveau and Lindow, 2001). Uneven leaf surface distribution applies not only to fructose but also to other nutrients such as for example sucrose (Miller et al., 2001) and iron (Joyner and Lindow, 2000). One of the ways that epiphytic bacteria deal with low iron availability is by the production of siderophores (Loper and Buyer, 1991). These low-molecular-weight molecules chelate ferric ion and after recognition by specialised receptors are taken up by the bacteria. The molecular biology of siderophore production and recognition has been exploited to construct iron-responsive bioreporter strains of Pseudomonas species (Loper and Lindow, 1994; Joyner and Lindow, 2000). Using the inaZ reporter gene, it was possible to estimate that the average bacterial cell in the phyllosphere experienced low-iron conditions (Loper and Lindow, 1994). A similar result was obtained with a gfp-based iron bioreporter (Joyner and Lindow, 2000), but because GFP allows for the interpretation of reporter activity in individual cells, it was possible to show that actually there existed substantial microscale heterogeneity in iron availability in the phyllosphere.

11.8

Microbial interactivities in the phyllosphere

11.8.1 Niche modification Niche modification represents one of the microbial strategies to change local conditions away from the harsh environment that characterises the phyllosphere. The production of the plant hormone indole-3-acetic acid (Brandl et al., 2001) or the phytotoxin syringomycin (Lindow and Brandl, 2003) by bacterial leaf colonisers is thought to stimulate the localised release of nutrients by the plant. Another strategy to increase nutrient availability is by altering cuticular surface permeability (Schreiber et al., 2005), or by decreasing the contact angle of sessile droplets on the cuticular surface (Knoll and Schreiber, 2000). The latter stimulates leaf wetting, for example, by the production of surfactants (Bunster et al., 1989), thereby facilitating the leaching of nutrients (Knoll and Schreiber, 2000; Chapters 7 and 8). Extracellular polysaccharides are quite commonly used by phyllosphere bacteria to firmly attach to the cuticular surface. They may in addition serve to improve

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living conditions, for example, to protect from desiccation, trap nutrients, as a barrier against chemical, biological or environmental stresses or as a matrix for the communication via small diffusible molecules such as AHLs (see Section 11.8.4).

11.8.2 Competition Competition for nutrients is thought to be most fierce when nutrients are scarce. Evidence for nutrient competition in the pyllosphere comes from studies that demonstrate the principle of pre-emptive competitive exclusion (Lindow and Leveau, 2002), that is, the ability of an established microbial epiphyte to inhibit the development of a population of a second strain on leaves (Kinkel and Lindow, 1993). This principle is based on nutrient depletion and has been applied as a biocontrol strategy (Section 11.9.3). So-called r-strategists (Section 11.7.1) are more affected by competition than K-strategists, and strategies based on pre-emptive competitive exclusion are thought to be more effective in controlling the former than the latter (Marois and Coleman, 1995).

11.8.3 Antibiosis Several epiphytic yeasts have been shown to produce antibacterial compounds (McCormack et al., 1994), and antifungal activities have been attributed to epiphytic bacteria (Giesler and Yuen, 1998; Nair et al., 2002; Collins et al., 2003; Daayf et al., 2003). Antibiosis, however, has not been explicitly demonstrated to be a major mechanism in the interaction between bacteria on leaf surfaces, despite observations that antibiosis of bacterial epiphytes can be demonstrated in the laboratory (Lindow, 1988). It could well be that under such conditions production of and sensitivity towards antibiotics is different than under phyllosphere conditions.

11.8.4 Communication Many plant-associated bacteria produce quorum sensing signals, such as AHLs (Cha et al., 1998). These allow for the indirect sensing of population density and density-dependent control of gene expression (Juhas et al., 2005). In the epiphytic bacterium P. syringae, the production of AHLs is regulated in a complex and hierarchical manner (Quiñones et al., 2004), involving the GacS/GacA two-component system (Section 11.6.3). In the model proposed by Quiñones et al. (2004), the ahlI gene, which codes for the synthesis of 3-oxo-C6-homoserine lactone, is positively regulated by the AhlR protein in combination with 3-oxo-C6-homoserine lactone, resulting in a typical positive feedback (auto-induction). The enhanced survival of bacterial cells in densely packed aggregates on leaf surfaces seems to suggest that many epiphitness traits may be controlled in a cell-density-dependent manner. An ahlI− ahlR− double mutant of P. syringae had a reduced epiphitness on dry leaves compared to the wildtype (Quiñones et al., 2004), suggesting a role for quorum sensing in withstanding desiccation stress in the phyllosphere.

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11.8.5 Gene exchange Many bacterial epiphytes have been shown to carry plasmids (Kobayashi and Bailey, 1994; Sundin et al., 2004). Together with transposons, these constitute the so-called horizontal gene pool (Bailey et al., 2002). This gene pool often confers traits that promote survival in the phyllosphere, including virulence factors (Sundin et al., 2004) and resistance to antibiotics such as tetracycline which are sprayed in apple orchards (Schnabel and Jones, 1999). Cuticular surfaces are hotspots for gene exchange and have been called ‘breeding grounds for microbial diversity’ (Lindow and Leveau, 2002). The aggregated nature of bacterial cells on the cuticular surface is believed to play a key role in the efficiency of plasmid transfer. Rates of plasmid transfer on bean leaf surfaces were 30-fold higher than on membrane surfaces (Normander et al., 1998). Using a reporter gene system that is based on de-repression of GFP expression in plasmid recipients, plasmid transfer has been observed in situ on leaf surfaces (Normander et al., 1998). With this bioreporter, it became clear that plasmid exchange occurs not randomly, but primarily in junctures between epidermal cells and in substomatal cavities. Apparently, plasmid exchange did not require the bacteria to be metabolically active (Normander et al., 1998), although other environmental factors such as water availability do seem to matter (Björklöf et al., 2000).

11.9

Biocontrol in the phyllosphere

11.9.1 Phyllosphere diseases Examples of phyllosphere microorganisms that harm plants are plenty. Some, such as Pseudomonas and Erwinia species, cause frost injury through biological ice nucleation (Lindow, 1983). Bacterial ice nucleation has been shown to be mediated by bacterially produced proteins that serve as nucleators for ice formation at subzero temperatures. Genes encoding ice nucleation activity (ina) have been isolated from P. syringae, P. fluorescens, E. herbicola, E. ananas and X. campestris among others. Sequence analyses suggest that ina genes have a common ancestor, but the selective advantage of ice formation to bacteria is still unknown; perhaps they benefit for some unidentified reason from frost injury to plants (Hirano and Upper, 2000). Several bacteria and fungi can produce phytohormones which may disrupt normal plant functioning and cause growth deformation, such as leafy gall on sweet pea caused by Rhodococcus fascians (Vandeputte et al., 2005). Other bacteria affect plant productivity by the formation of leaf spots or lesions, or by inducing leaf blight or curling (Agrios, 1997). Some examples of foliar fungal diseases (and their causative agents) are rice blast disease (Magnaporthe grisea), downy mildew of grape (Plasmopara viticola) and powdery mildew affecting all kinds of plants (different fungal species). Interestingly, no yeasts are known to cause foliar disease (Agrios, 1997). Most common bacterial pathogens of the phyllosphere include members of the genera Erwinia, Pseudomonas and Xanthomonas. Different pathovars of

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P. syringae cause leaf spots and blights on tobacco, cucumber, bean, lilac or tomato, while X. campestris pathovars affect bean, cotton, rice cereals, tomato or pepper. A growing concern is the occurrence of microorganisms such as Salmonella and Campylobacter species in the phyllosphere of produce. By definition, these bacterial residents of cuticular surfaces are not epiphytic, as they are usually not capable of multiplying, yet they are apparently quite able to survive for prolonged periods of time on leafy vegetables (Brandl and Mandrell, 2002), posing a health danger to human consumers.

11.9.2 What makes a plant pathogen? There are many different ways in which pathogenic microorganisms attack plants. Common themes, however, are invasion of the plant tissue, nutrient acquisition and counteracting plant defence reactions (Agrios, 1997). The mechanisms that underlie pathogenicity involve, among others, degradation of plant structural components (e.g. cuticle, cell walls) by enzymatic activity (e.g. cutinase, pectinase, cellulase), the production of phytotoxins (e.g. tabtoxin, syringomycin), injection of effector molecules (e.g. type III secretion in certain bacteria) and modulation of phytohormone levels (e.g. indole-3-acetic acid) to exploit plant physiology. In bacteria, pathogenicity factors such as production of plant growth hormones, phytotoxins and other virulence genes are often plasmid-borne (Bailey et al., 2002), suggesting a role of the horizontal gene pool in the evolution and spreading of disease. Many bacterial plant pathogens differ from non-pathogenic epiphytes in having the ability to colonise also the interior of leaves, thereby avoiding the stresses associated with the cuticular surface. These internal populations are generally believed to be responsible for disease induction, and the larger the internal population size, the more likely it is that disease symptoms occur (Beattie and Lindow, 1999). The bacterium Xanthomonas campestris avoids the leaf surface and actively seeks the interior of the leaf (Hugouvieux et al., 1998) where it reaches high population sizes that may egress to the leaf surface. Other bacterial pathogens, such as P. syringae generally establish large surface populations before they ingress into the leaf’s interior and proliferate there (Hirano and Upper, 2000). For the latter type of pathogens, leaf surface population sizes are predictive of the probability of disease occurrence (Beattie and Lindow, 1999). For both types of pathogens, these external populations are probably important sources of inoculum for emigration to other plant leaf surfaces (Upper and Hirano, 2002).

11.9.3 Strategies for biocontrol Strategies for biocontrol of foliar diseases are often based on the prevention of establishment of the pathogen in the phyllosphere. Spraying leaves with spores of common phyllosphere fungi such as Alternaria, Cochliobolus, Septoria and Phoma has been shown to reduce fungal foliar diseases (Agrios, 1997). Yeasts have been successfully used as antagonists of pathogenic fungi (Fiss et al., 2000; Avis and

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Bélanger, 2002; Buck, 2002; Urquhart and Punja, 2002) and bacteria (Assis et al., 1999), while bacteria have been shown to control disease symptoms caused by fungi (Kucheryava et al., 1999; Zhang and Yuen, 1999; Nair et al., 2002; Collins et al., 2003) and bacteria (Volksch and May, 2001; Stromberg et al., 2004). Some mixtures of biocontrol agents show more anti-fungal activity than the single strains alone (Guetsky et al., 2002). The mechanisms underlying some of these strategies are not always clear. Mycoparasitism, that is, fungal attack on fungi, is an effective biocontrol mechanism which is best described for Trichoderma species (Bélanger and Avis, 2002). Some biocontrol agents have been shown to induce systemic resistance in plants (Bargabus et al., 2002). Others produce antibiotics (Giddens et al., 2003), but antibiosis in the laboratory is not necessarily a guarantee for success in the field (Lindow, 1988). Another strategy for biocontrol is based on the principle of pre-emptive competitive exclusion which assumes that growth in the phyllosphere is limited by the availability of nutrients. When plants are deliberately inoculated with a nonpathogen (the biocontrol agent) which will use up most of the nutrients, any immigrating pathogen will find itself unable to grow and form sufficiently large population sizes to cause damage. Naturally occurring non-ice-nucleating (Ice− ) strains of P. syringae and E. herbicola applied pre-emptively to plants prevented successful colonisation by Ice+ P. syringae strains and reduced the severity of frost injury (Wilson and Lindow, 1994a). Crucial to success of this strategy is the fact that the nutrient overlap indices of the pathogen and the biocontrol agent are quite similar. However, nutrient-overlap indices are not always predictive for the ability of a non-pathogen to control disease in the laboratory or in the field (Ji and Wilson, 2002).

11.10 Future directions of phyllosphere microbiology Historically, much of the research on microbial communities in the phyllosphere has been driven by the desire to understand the biology and ecology of microorganisms that are harmful to aerial plant parts. This trend is obvious from the bias, even in this chapter, towards our knowledge and understanding about plant pathogens. Clearly, the attention is slowly shifting in favour of non-pathogenic inhabitants of primary plant surfaces, not only because they have the potential to affect the function of phyllosphere pathogens, but also because they hold many undiscovered traits of adaptation to life on cuticular surfaces. The use of reporter genes such as gfp for the first time has opened up the possibility to study microbial epiphytes as individuals, and has fuelled the realisation that microorganisms operate at micrometre scales, and that the cuticular surface at that level of magnification is a highly heterogeneous environment. The ability of studying epiphytic individuality needs to be exploited much more than it has been already: it may help to explain phyllosphere-related phenomena for which there are currently no good explanations if one continues to stick to the traditional, that is, macroscale, view of single leaves as operational units.

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Furthermore, as more and more microbial interactivities in the phyllosphere are revealed and in ever more detail, their interconnected complexity will soon reach (or has already reached) a level where the interpretation of experimental results through linear thinking is no longer realistic. Instead, phyllosphere microbiologists will need to rely more on predictive and interpretive modelling, for which at this point much more quantitative data is needed, both on leaf surface structures and properties and on microbial (inter)activities in the phyllosphere. Also, the cautious embrace by phyllosphere microbiologists of culture-independent methods will need to mature into a full-scale exploitation of the molecular toolbox, including the application of such exciting new technologies as metagenomics analysis. As a final point, phyllosphere microbiology has entered the genomic era with the sequencing of whole-genomes of several plant-pathogenic, epiphytic lifestyle microorganisms. Many more genomes of phyllosphere microorganisms will become available in the next decade, and it will be a huge and incredibly exciting challenge to explain this enormous wealth of sequence information in the light of the experimental data from the 50 years of pre-genomic phyllosphere microbiology research.

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Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

12 Filamentous fungi on plant surfaces Tim L.W. Carver and Sarah J. Gurr

12.1

Introduction

Huge numbers of filamentous fungi spend at least part of their lives on the subterranean or aerial surfaces of the myriad plant species in existence. Almost nothing is known about most of these relationships, and little is known about the majority of the remainder. Here, therefore, we shall focus on relatively few of the better-studied interactions and restrict our consideration to relationships between leaf surface characteristics and pathogenic leaf-infecting fungi, concentrating on obligate biotrophic and hemi-biotrophic fungi. Further, we shall restrict our thoughts to those fungi which have evolved relatively sophisticated relationships with their hosts, infecting them via specialised infection structures known as appressoria. Key features of mature germlings developed from the unicellular spheroidal conidia of Blumeria graminis (the cereal and grass powdery mildew fungus), Erysiphe pisi (the pea powdery mildew fungus) and the multicellular, pyriform conidia of Magnaporthe grisea (the rice blast fungus) are illustrated in Figure 12.1. We shall draw both on our own experiences researching early interactions involving B. graminis and on the wealth of information from additional systems including pea powdery mildew (E. pisi), grape black rot (Phyllosticta ampelicida), rusts of monoand dicots (Puccinia and Uromyces spp.), rice blast (M. grisea) and diseases caused by Colletotrichum species. Compared to the relatively consistent subterranean environment, fungi that infect leaves are likely to have to survive rapid and dramatic fluctuations in temperature, humidity and light as well as having to avoid being dislodged or damaged by direct and indirect physical forces imposed by wind and rain. Plant surface characteristics are vitally important during the earliest stages of pathogen development before disease becomes established, and although they may have some influence thereafter, in most cases this is likely to be relatively small and has been little studied. We shall concentrate, therefore, on the influence on leaf surface features during the phase from spore deposition up to the time when the fungus enters the leaf. The resources carried by fungal spores are finite and must be used effectively and efficiently during the time taken to germinate, differentiate infection structures, invade the host and gain access to host-derived nutrient sources. The fungal germling/leaf surface interaction presents something of a paradox. On one hand, the cuticle and underlying epidermal cell wall present an immediate and formidable barrier to infection. On the other hand, many fungi have apparently evolved dependence upon physical, topographical and chemical information contained within their

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Figure 12.1 Cryo-SEM images of mature appressoria formed by the single-celled conidia of Blumeria graminis (on oat) and Erysiphe pisi (on pea) and the multicelled conidium of Magnaporthe grisea (on rice). Note, conidia of B. graminis and E. pisi show domed and ribbon-like surface projections, respectively. (a) B. graminis conidia (C) uniquely form a short primary germ tube (PGT) at around 1 h after inoculation before the appressorial germ tube (AGT) emerges and elongates to ca 40 mm before differentiating a simple, hooked appressorium (App) by 8–10 h. The PGT and App frequently form on different epidermal cells. (b) The AGT of E. pisi grows little before differentiating a multilobed appressorium by around 4 h. (c) The conidium and AGT of M. grisea collapse as the cytoplasm migrates into the domed appressorium. Micrograph kindly supplied by N.J. Talbot and G. Wakely (Exeter University). Note, the rice leaf has many wart like protuberances on which conidia are often deposited.

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host plant surface to stimulate activities and provide the signals regulating fungal responses driving the complex developmental processes which are pre-requisite to infection.

12.2

Adhesion prior to germination

12.2.1 The environment, leaf surface characteristics and spore shape Spores are not only dispersed by wind and rain but they also may be displaced by their action after delivery to potential infection sites. Water can wash away deposited spores and rain drop impact causes radial air movements that may briefly exceed 150 km h−1 over a short distance (Hirst and Stedman, 1963). Due to the boundary layer, direct wind effects are likely to be small close to leaf surfaces but wind shake can generate considerable force (Bainbridge and Legg, 1976; Wright et al., 2002a). It is imperative for survival, therefore, that ungerminated spores become rapidly attached to their host, and plenty of evidence indicates that this occurs. Gross effects of plant architecture (erect or prostrate habit) and canopy density can obviously affect spore deposition, but minute topographic leaf surface features may also influence spore adhesion and retention considerably. At a relatively crude level, leaf surfaces vary considerably in the abundance and form of trichomes and other epidermal protuberances, and becoming trapped on these structures can prevent spore deposition on the epidermal surface proper. Further, the outer epidermal cell wall is generally not flat but is convex in at least one dimension while anticlinal cell wall junctions form troughs. At the ultrastructural level, leaves are generally covered in hydrophobic epicuticular waxes that are frequently crystals which present minute outer tips or edges (see Chapter 1). Thus, a relatively large spore may be supported on the extremity of very few wax crystals so that the actual interface between surfaces is miniscule (Figure 12.2a). The geometry of the spore-surface interface is also important. Spore shape varies enormously from single-celled spheroids (e.g. B. graminis conidia) to more complex multicelled structures (e.g. M. grisea conidia). Some, e.g. Colletotrichum graminicola conidia, are sickle-shaped (falcate), and this can increase potential interfacial contact with plant cell surfaces. The concave profile of these conidia would ‘fit’ better to the convex plant cell surface, its convex profile would fit best when deposited across the trough between adjacent epidermal cells and its more linear, lateral profile would fit a flatter surface such as offered by the long axis of a graminaceous epidermal cell. In any case, axial rotation of a falcate spore settling from a dispersing water film would maximise its contact area with the plant surface. Although some spores appear smooth (e.g. C. graminicola), many show pronounced surface ornamentation such as the surface projections of B. graminis conidia or the spines of rust urediniospores. In these cases the interface may be restricted to a few minute points of contact between spore surface ornaments and leaf wax crystals (Figure 12.2a). In such cases, extremely efficient means of adhesion must come into play.

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(a)

(b)

Co

Le

1 μm

5 μm (d)

(c)

5 μm

10 μm

Figure 12.2 Cryo-SEM images of Blumeria graminis conidia on host leaves and an artificial substratum. (a) After 60 min on a barley leaf with epicuticular wax in place. Conidium (Co) to left and leaf (Le) to right. Contact is limited to the tips of a few conidial surface projections touching the raised edges of leaf wax crystals (white arrows) and intervening spaces are clear (from Carver et al., 1999). (b) After 1 min incubation on a de-waxed leaf the cryofixed spore was displaced by micromanipulation to reveal the original contact site (white arrows) where minute droplets of extracellular material (ECM, inset) were released from conidial surface projections that made leaf surface contact (from Wright et al., 2002b). (c) If conidia are not displaced, the minute droplets of ECM cannot be discerned at the contact site (black arrow) even after 15 min incubation on a de-waxed leaf. (d) On planar, hydrophobic artificial substrata (here silanized plastic), copious ECM is released and within 15 min fills the entire conidium/substratum interface.

12.2.2 The challenges and nature of spore adhesion Foliar pathogens must adhere to the hydrophobic, waxy leaf surface, and many studies using artificial substrata show a strong relationship between the occurrence and/or strength of adhesion and hydrophobicity, while spores of many species fail, or show much reduced capacity, to stick to hydrophilic surfaces (e.g. Hamer et al., 1988; Clement et al., 1993; Terhune and Hoch, 1993; Kuo and Hoch, 1996). As Jones (1994) noted, and discussed in some detail by Kuo and Hoch (1996), both fungal spores and plant surfaces carry a negative charge and electrostatic repulsion must result. This may be counteracted by various means including hydrophobic interaction, van der Waal’s attraction forces and hydrogen bonding. These forces might simply help spores remain in place over the short term. After a brief period in contact with a leaf, however, spores of many phytopathogenic fungi release adhesive extracellular materials (ECM) and relevant work has been reviewed in

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detail (Nicholson, 1996; Epstein and Nicholson, 1997; Kunoh et al., 2001, 2004; Tucker and Talbot, 2001). Nicholson (1996) points out that with the exception of the powdery mildew fungi most spores commence their host association in the presence of free-water in which case adhesives must be water-insoluble or rapidly become so after release. The precise chemistry of spore adhesives varies greatly between fungi although they commonly contain glyco-proteins, lipids and polysaccharides and in some cases enzymes such as cutinase. Jones (1994) considered there to be two basic strategies for spore adhesion, namely ‘passive’ and ‘active’. Most workers have adopted this distinction, which regards passive adhesion as resulting from the presence of pre-formed material on or within the spore, while active adhesion requires the de novo production of adhesive.

12.2.3 Examples of passive and active adhesion and combinations of the two The timing of adhesion merits attention because it conveys information about the passive or active nature of underpinning adhesion mechanisms. An example of extremely rapid attachment by passive means is shown by P. ampelicida pycnidiospores which, in an acidified aqueous environment, adhere firmly to hydrophobic but not hydrophilic artificial substrata within milliseconds, even if they are killed before inoculation (Kuo and Hoch, 1996). In this case, the role of spore ECM is unclear although the extracellular conidial sheath appears to mediate initial attachment and probably plays a role in selection for substratum specificity (Kuo and Hoch, 1995; Shaw and Hoch, 1999). However, a classic case of passive adhesion due to the release of ECM is shown by conidia of M. grisea (Hamer et al., 1988; Howard, 1997). In dry conidia, a large periplasmic deposit lies within the cell wall at the tip of the apical cell of the pyriform conidium, and this is released instantly upon hydration as ‘spore tip mucilage’. It contains α-linked-mannosyl and glucosyl residues, protein and lipids, and adheres immediately and robustly to Teflon (hydrophobic) though less strongly to glass (relatively hydrophilic). This requires no post-deposition energy expense, and if (see later) the material makes plant surface contact, it may be expected to convey fast and effective spore retention. Scanning electron microscopy of C. graminicola conidia shows that these also release ECM where their tip contacts maize leaves and along their entire region of contact with artificial hydrophobic substrata (Mercure et al., 1995). This ECM contains glycoprotein (Sugui et al., 1998) that was shown to play a role in active adhesion since it was abolished by addition of pronase and impeded by inhibitors of glycoprotein transport and protein synthesis (Mercure et al., 1994). Although reasonably rapid, maximum adhesion was not attained until around 30 min after contact, presumably reflecting the time taken for de novo synthesis and accumulation of the adhesive. Urediniospores of the rust fungi probably depend on a sequence of passive and then active mechanisms to effect increasingly strong adhesion as time passes. Thus, evidence from Clement et al. (1994) suggests that the spine-like ornaments of U. viciae-fabae urediniospores are invested in a hydrophobic lipid-containing

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sheath that promotes immediate, passive adhesion to hydrophobic substrata. This hydrophobic interaction is sufficient for spore retention during the more lengthy processes involved in imbibition of water that is a prerequisite for rust urediniospore germination. Furthermore, Clement et al. (1997) showed that although imbibition by spores is almost instant in the presence of free water, in its absence, even at 100% relative humidity, spore hydration awaits accumulation, by capillary condensation, of water that forms a ‘condensation pad’ at the spore–substratum interface, and this may take many hours. In the early stages, this condensate can be removed entirely by freeze drying, but capillary force generated by the developing condensation pad may itself confer adhesive force to support the hydrophobic interaction. However, as the spore hydrates the pad cannot be removed by freeze-drying, indicating the accumulation of non-volatile components in a structure termed the ‘adhesion pad’. Deising et al. (1992) had shown previously that while pads form beneath living and autoclaved U. viciae-fabae spores, only living spores adhere strongly to a leaf surface. They provided good evidence that this was due to the presence of a cutinase and two non-specific serine esterases present on the surface of living spores. Evidently, these enzymes were released into the adhesion pad following spore hydration to interact with the host surface and cause adhesion. This, they considered, represents an active process. The simple single-celled conidia of the powdery mildew fungi favour humid conditions but unlike the cases discussed so far, the spores can be damaged by free water. Amongst the powdery mildews, adhesion by B. graminis has been relatively well studied but it remains unclear whether this is solely passive or also incorporates active processes. The native spore surface is hydrophobic (Nicholson et al., 1993) and so hydrophobic interactions may be expected to lead to immediate passive adhesion. However, when on a true leaf surface, the conidia almost instantaneously release minute droplets of ECM that appears to emanate from the few surface projections (ornamentations that cover the spore surface) making leaf surface contact (Figure 12.2b; Wright et al., 2002b). Centrifugation studies (Wright et al., 2002a) show that within 10 min of deposition (the quickest it was possible to make a test) about 80% of conidia remained attached to leaves even when subject to about 4.2 × 10−9 N force, which is far greater than likely to be generated by wind-shake of crop leaves. Such strong adhesion was attributed to effects of the conidial ECM since it is known to contain non-specific esterases and a cutinase (Nicholson et al., 1988; Pascholati et al., 1992), isolated ECM is able to degrade host surface features (Kunoh et al., 1990), and similar enzymes are involved in adhesion by U. viciae-fabae (earlier). The uncertainty of whether this enzyme-associated adhesion is simply a passive process involving the release of pre-formed ECM components stems from data (Nicholson et al., 1988) showing that non-specific esterase release on artificial substrata is a two-phase process. The first occurs within 2 min of contact and is unaffected by metabolic inhibitors but the second, occurring after about 15 min, is inhibited by cycloheximide indicating de novo synthesis. It now seems possible that this second phase is not induced on true plant leaves. First, Wright et al. (2002b) obtained no evidence for increase in the number or size of

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ECM deposits from the first minute up to 12 h after deposition. Second, Wright et al. (2002a) found little evidence for increased adhesion to barley leaves between 10 and 30 min after spore deposition, whereas, by contrast, on hydrophilic glass there was a substantial increase in adhesion during this time and the quantity of deposited ECM continued to increase between 1 min and at least 3 h (Wright et al., 2002b). These observations indicate, therefore, that whereas contact with an artificial substratum may induce the second phase of ECM release, only the first phase, involving release of preformed ECM, occurs on the host leaf surface. This would indicate passive adhesion to leaves.

12.2.4 Plant surface versus artificial substrata While artificial substrata offer a valuable means of simplifying systems for experimental studies, the previously mentioned experiences using B. graminis indicate the potential danger of extrapolating from observation of spore behaviour on artificial substrata to the natural situation. In fact, previous studies of B. graminis had already indicated a problem of using artificial substrata to study conidial ECM release. Thus, while it was impossible even by low-angle, low-temperature SEM to resolve ECM deposits beneath in-place conidia on leaves (Figure 12.2a), even with epicuticular wax removed (Figure 12.2c), extraordinarily large accumulations were evident after as little as 15-min incubation on the silanized surface (hydrophobic) of planar glass or plastic (Figure 12.2d; Carver et al., 1999). In part this might be explained by the geometry of the interface: on the planar surface a larger number of ECM-releasing conidial surface projections would make contact with the substratum than when the ellipsoidal spore is in contact with the convex epidermal cell surface (Wright et al., 2002b). The additional capillary forces brought into play on the planar surface might then draw the spore closer to the surface, bringing more conidial projections into contact, causing more ECM to be released. The idea that interfacial geometry is important is supported by subsequent observations showing copious accumulation of ECM on flat, isolated leaf cuticle (Fujita et al., 2004a). A further case indicating the necessity of considering the nature of the true leaf surface came from recent observations of M. grisea on rice leaves (Koga and Nakayachi, 2004). As described previously, earlier studies (Hamer et al., 1988) indicated the potential of M. grisea spore tip mucilage to adhere to hydrophobic surfaces and this is seen as a classic example of passive spore adhesion. This fungus has a wide host range and it may be true that on many host species this mucilage acts as a spore adhesive. However, Koga and Nakayachi (2004) pointed out that the leaf surface of rice (Oryza sativa L., the most economically important host of M. grisea) is densely covered by ‘wart-like’ protuberances of the epidermis (Figure 12.1c). They found that spores were generally deposited on these protuberances and so had no contact with the underlying surface while their tip cell often had no contact with any plant structure. Thus, when spore tip mucilage was released upon hydration, it failed to make plant surface contact, conidia failed to adhere and for at least 1 h after deposition virtually all could be washed away simply by dipping

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into water. In this circumstance, adhesion of ungerminated spores to the rice leaf is at best very weak and certainly trivial compared to that conferred when spore tip mucilage is released onto an artificial planar surface. Strong adhesion to the rice leaf apparently depends on later contact by the fungal germ tube (Koga and Nakayachi, 2004).

12.2.5 Effects of ungerminated conidia on underlying host cells Few studies of spore ECM have considered its potential effects on underlying host cells although several early reports from the powdery mildew/barley system have suggested this possibility (see Fujita et al., 2004b). Direct evidence for an effect was provided recently by Fujita et al. (2004a) who allowed barley epidermal cells to have contact for less than 1 h with conidia of the non-pathogenic powdery mildew species E. pisi (the pea mildew pathogen) or B. graminis f.sp. tritici (a pathogen of wheat but not barley). During this time, conidia released ECM onto epidermal cells but they were removed before germinating. Epidermal cells that received ECM showed ‘induced inaccessibility’ being far more resistant to later attack by the pathogenic barley mildew fungus (B. graminis f.sp. hordei). Importantly, however, ECM from pathogenic conidia (B. graminis f.sp. hordei), had no detectable effect. Thus, this study showed perception and response of plant cells to component(s) of ECM released by the non-pathogenic fungi, but also indicated either the absence of inductive component(s) in the true pathogen’s ECM or the presence of a suppressive factor. It also implied that fungal ECM component(s) move into the plant cell. This could be facilitated by modification of the plant cell surface not only by the action of cuticledegrading enzymes (mentioned earlier) but also by cell-wall-degrading enzymes known to be present in B. graminis conidial ECM (reviewed by Kunoh et al., 2004). We now have evidence for such movement. As mentioned by Kunoh et al. (2004), Thomas and colleagues (unpublished) used immunogold labelling to trace distribution of a monoclonal antibody (MAb AF-BBG1) raised against surface washings of Aspergillus flavus (by M.F. Dewey, Oxford) and applied to sections of B. graminis conidia incubated for 30 min on oat leaves. The exact chemistry of the target antigen remains to be determined, but MAb AF-BBG1 has been shown to recognise commercially available cutinase and component(s) of B. graminis conidial cytoplasm, cell wall and ECM; importantly, it recognises no constituent of healthy cereal leaves (T. Gjetting, personal communication). Figure 12.3 shows that within 30 min of inoculation the antigen was present not only in the spore and its ECM released at the site of leaf contact, but also within the wall and cytoplasm of the plant epidermal cell. It is unknown whether this particular component directly affects plant cell activity, but the observation demonstrates the principle that a spore component may pass into plant cells prior to germination. It would be surprising if this had no effect on those cells. Clearly, consideration of spore ECMs should include not only their role in adhesion but also their possible influences on plant cells which may express enhanced resistance (or perhaps susceptibility?) to subsequent attempted infection.

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ECM

CCW

PCW

300 nm Figure 12.3 Transmission electron micrograph of the interaction site between a Blumeria graminis conidium and an underlying oat leaf epidermal cell 30 min after inoculation and following immunogold labelling with MAb AF-BBG1. The antigen is present not only within the conidium, its cell wall (CCW) and ECM, but gold label indicates that it has moved into the plant cell wall (PCW) and underlying cytoplasm (circled gold particles).

12.3

Influence of leaf surface characteristics on spore germination

There is little evidence of whether leaf surface characteristics directly influence germination by spores of obligate biotrophic and hemi-biotrophic fungi. For P. ampelicida, whose spores germinate in an aqueous environment, Kuo and Hoch (1996) showed that spore adhesion to a substratum was a mandatory prerequisite for germination. However, this requirement has not been clearly demonstrated for any other fungus and it is unknown whether leaf surface features may impede adhesion by P. ampelicida spores and hence influence germination. It has also been shown that germination by C. gloeosporoides is stimulated by contact with surface wax of avocado (Podila et al., 1993) but the specificity and basis of this effect is unclear. For the cereal powdery mildew fungi that favour a water-free environment, a number of early studies (e.g. Carver and Adaigbe, 1990 and work cited therein) suggested that leaf age, position and host genotype may all affect germination by B. graminis, but these relatively small effects remain unexplained. However, leaves of oat, barley and wheat are all covered by crystalline plate-like waxes and it is possible that the nature of these waxes changes as leaves age (e.g. Riederer and Markstädter, 1996) and varies between genotypes, and it is possible that these factors influence B. graminis germination. Thus, Carver and Thomas (1990) showed that removal of these waxes prior to inoculation can significantly reduce germination frequency. Although this relatively crude approach suggested an influence of epicuticular wax, removal of the crystalline plates would have altered the fine details of the contact interface between spore and leaf surface (see Figure 12.2a–c), and the physical consequences of this are unknown.

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A far more subtle evaluation of epicuticular wax influence was recently undertaken by Gniwotta et al. (2005) in the pea (Pisum sativum L.)–powdery mildew (E. pisi) system. Under identical environmental conditions, germination by E. pisi conidia was significantly higher on the adaxial (80%) than abaxial (57%) leaf surface although even here germination was higher than on hydrophilic or hydrophobic glass (49% germinated). This indicates not only that germination is specifically promoted by plant surface signals, but also that the efficacy of these signals is greater on the adaxial surface. Furthermore, Gniwotta et al. showed differences in both the structure and chemistry of waxes from the different surfaces. On the adaxial surface they formed plate-like crystals and contained mainly primary alcohols (71% of total content), being dominated by 1-hexacosanol. By contrast, the abaxial waxes formed ribbon-like crystals that consisted mainly of alkanes (73%) with little primary alcohol content. From many studies it is clear that, as for E. pisi, conidia of B. graminis germinate on a range of artificial substrata (reviewed by Green et al., 2002), and in this case germination may be a non-specific response to substratum contact, stimulated perhaps by ECM release. Nevertheless, Gniwotta’s observations of E. pisi (mentioned earlier) indicate that plant surface components can promote germination, although the details of how this may be mediated are unknown. However, Nielsen et al. (2000) showed hydrolytic enzyme activity in the region of B. graminis conidial ECM within 3 min of spore deposition. Further, they showed that prior to germination, anionic low-molecular weight compounds with physical properties similar to cutin monomers could be taken up by conidia incubated on host leaves. This suggests that fungal enzymes present in the conidial ECM (Nicholson et al., 1988; Pascholati et al., 1992) may degrade components of the plant surface/cuticle (Kunoh et al., 1990) to release breakdown products that can then be taken up by the spore as part of a signalling system involved in germination. What evidence is there for this? Transcript abundance assays (Perfect, 2005) for B. graminis cutinase 1 gene from conidia deposited on host leaf or on glass or plastic overlaid with cutin, revealed elevated transcription of the gene within 30 min of deposition. By contrast, no equivalent transcript induction was detectable for conidia deposited on clean glass or plastic surfaces on which normal germling development is not induced. Clearly, further experimentation is needed to resolve the details of this system in the powdery mildews and to search for analogous systems in other fungal/plant associations.

12.4

Directional emergence of fungal germ tubes

Germ tubes must make contact with the host leaf in order to function and since spores are three-dimensional bodies lying on an undulating leaf surface, the likelihood of germ tube contact will be greatly increased if they emerge from the spore at a point close to the leaf surface contact site. Although this idea is obvious, it received little attention until Wright et al. (2000) considered the primary germ tube (PGT) of B. graminis. As noted earlier, the PGT is invariably short (it ceases growth at

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Figure 12.4 Cryo-SEM image showing a Blumeria graminis conidium on a barley leaf epidermal strip where it was incubated for 15 min before it was rolled over so that its original site of contact with the leaf (region marked by dashed line) faced away from the leaf. Repeated light microscope observation of the living spore showed that the first and second (1st, 2nd) formed germ tubes then emerged close to the original contact site, demonstrating that initial contact had programmed these sites of emergence. Eventually, the conidium responded to the new site of contact so that its third (3rd) germ tube made leaf contact (from Wright et al., 2000).

around 5–10 μm) and this clearly restricts its chances of fortuitous contact with a substratum. Wright et al., (2000) developed a geometric model taking account of the relationships between the ellipsoidal conidium and curved host cell surface. From this they calculated that according to circumstances (spore orientation, epidermal cell size), only between 8 and 22% of PGTs would make contact if they emerged at a random point on the spore surface. However, in reality, observation shows that at least 80% of PGTs make host surface contact. This clearly indicates existence of a precise system controlling PGT emergence very close to the contact site. The accuracy of this control becomes even more remarkable when one remembers that the actual area of spore/leaf surface contact consists of the interface between very few conidial surface projections with the fine edges of epicuticular wax crystals (Figure 12.2a). The extraordinary sensitivity of the perception system is underlined by the ability of many conidia to recognise even the minute contact made with a 0.5 μm diameter spider’s suspension thread (Wright et al., 2000). Wright et al. (2000) also showed that recognition of contact is extremely fast. By micromanipulation, deposited B. graminis conidia were rolled over before they germinated so that their original site of contact then faced away from the leaf. Repeated observation of these living spores showed that even when they were rolled within 1 min of deposition, the majority of tubes eventually emerged very close to the original site of leaf contact and so grew upwards, away from the leaf (Figure 12.4). How could such a rapid response to contact be engaged? Present knowledge, from B. graminis and more recently from E. pisi, implicates conidial ECM release which, as discussed earlier, is known to occur within seconds of spore deposition onto artificial and plant surfaces. Details of the signalling system(s) are not understood, but in B. graminis this must apparently involve some form of non-specific response since most PGTs emerge very close to contact sites with

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all artificial substrata tested although the proportion is significantly higher if they are hydrophobic than hydrophilic (Carver et al., 1999; Wright et al., 2000). Furthermore, for E. pisi, substratum hydrophobicity promotes not only the frequency of emergence close to the contact site but also the speed of conidial ECM release and the speed of germ tube emergence (Fujita et al., 2004b). This suggests that it may be the simple act of localised ECM release that mediates non-specific response to contact. However, in E. pisi the response to surface contact can be overcome by positive phototropic effects. Thus, even when incubated on hydrophobic substrata or barley leaf epidermis, if light is supplied from directly above conidia, most germ tubes emerge from the surface facing away from the underlying surface and grow away from it (Fujita et al., 2004b). This indicates not only the existence of a photoreceptor system within ungerminated E. pisi spores, but also that light can have a more powerful influence on germ tube emergence than surface contact. In B. graminis, however, no such photoreceptor system appears to exist (Fujita et al., 2004b). Moreover, the frequency of B. graminis PGT contact is greater on leaf cells than on any artificial substrata even though geometric relationships predict a lower frequency than on planar substrata (Wright et al., 2000). This implies that specific characteristics of the host surface augment the non-specific response to surface contact. This may involve the release of leaf surface components by ECM enzymes and their subsequent uptake by conidia (Nielsen et al., 2000). If so, it is possible that differences in plant surface chemistry (due to genotype or species) may affect the signalling system and impair control of directed germ tube emergence although this possibility has never yet been tested.

12.5

The special case of the PGT of B. graminis

Although the multicellular spores of some fungi (e.g. M. grisea and Colletotrichum spp.) may produce a germ tube from more than one of their cells, this appears to be an unregulated phenomenon. Even among the powdery mildew fungi, B. graminis is to our knowledge unique in invariably producing a PGT before forming an appressorial germ tube (AGT). Why B. graminis should have evolved dependence on its PGT is unknown, but the vital functions it plays in pathogenesis illustrate the intimacy of relations between a fungus and its host surface. On contacting the leaf surface, the PGT secretes ECM that adheres rapidly and tenaciously to the leaf (Wright et al., 2002). Unlike conidial ECM, the ECM secreted by PGTs is not water soluble although it does contain protein, and its adhesion is so strong that if the germ tube is displaced the region of underlying cuticle is often torn away from the leaf cell wall (Carver et al., 1995; Figure 12.5). After attaching, the PGT forms a short penetration hypha, the ‘cuticular peg’, which penetrates the plant cuticle but not the cell wall (Edwards, 2002), probably aided by secretion of cutinase (Francis et al., 1996). Through its PGT the germling can absorb host water (Carver and Bushnell, 1983) which allows survival and continued germling growth under arid conditions that would otherwise desiccate and kill it. From the host plant’s

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(a)

(b) C

AGT

PGT ECM 5 μm

10 μm

ECM

Figure 12.5 Cryo-SEM images showing Blumeria graminis ECM and strength of adhesion revealed by stress displacement of the fungus during cryo-fixation. Leaves were de-waxed before inoculation so as to reveal ECM. (a) During fixation, the conidium and its PGT lifted away from the leaf surface, but strong adhesion at the PGT tip tore away the cuticle leaving traces of peripheral ECM around the wound site. (b) During fixation the appressorial germ tube of this young colony rolled back from the leaf surface exposing a track of ECM that is particularly thick around the appressorium contact site. Note: the fungal penetration peg is evident in the centre of this site (from Carver et al., 1995).

point of view, PGT contact triggers concomitant responses that include up-regulated transcription of many genes (Eckey et al., 2004; Hein et al., 2004), hydrogen and calcium ion efflux from the apoplast (Felle et al., 2004) and a series of cytological responses culminating in the deposition of a small apoplastic papilla (cell wall apposition) beneath the PGT tip (Zeyen et al., 2002). This early passage of signals through the plant cell wall apparently primes the capacity of cells to defend against later attack. Thus, a number of host genes are differentially transcribed according to the single gene-controlled resistance the plant carries (Hein et al., 2004), and, even where they possess no such resistance, cells contacted by a PGT show enhanced penetration resistance to later attack from the appressorium (Woolacott and Archer, 1984). Perhaps the most intriguing function of the PGT lies in its ability to recognise host surface features. Indeed this recognition is an absolute requirement since it drives elongation of the AGT that subsequently emerges from the conidium, and elongation of this germ tube is in turn a pre-requisite for appressorium differentiation. Why should the fungus have this requirement for energy- and time-consuming elongation of the AGT? One clear benefit is that growth away from the spore increases the likelihood that the appressorium will eventually form on an epidermal cell that has not been primed towards resistance by direct prior contact with either the conidial ECM or the PGT. The PGT’s function of recognising host surface features was first demonstrated (Carver and Ingerson, 1987) by the observation that although conidia suspended on spider’s thread or deposited on agar will germinate, they form only a series of short germ tubes none of which elongates and differentiates an appressorium. However, if germlings with a single short germ tube are transferred from spider’s thread to lie with their spore on agar but the germ tube tip placed in contact with a host epidermal strip, then the next germ tube to emerge is most likely to elongate.

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Subsequent studies have identified certain host surface characteristics that are recognised by the PGT. The presence of epicuticular wax crystals is not important because their physical removal does not reduce inductivity (Carver and Thomas, 1990). However, even after removal of epicuticular wax, leaves remain highly hydrophobic (Wright et al., 2000), presumably because of intracuticular wax and cutin, and substratum hydrophobicity is a key factor recognised by PGTs. Thus, although contact with clean, hydrophilic glass or agar does not induce AGT elongation, contact with biologically inert hydrophobic substrata is inductive, albeit less so than contact with the host leaf surface (Carver et al., 1996; Wright et al., 2000). Nevertheless, although leaf waxes are highly hydrophobic, not all components of leaf wax are recognised by the PGT and quite subtle differences in chemistry may have a relatively large effect. This was shown by Tsuba et al. (2002) who found that C26 chain length aldehydes (hexacosanal) present in barley leaf wax induce elongation of the AGT while C30 chain length aldehydes (triacontanal) in cabbage leaf wax (a non-host plant) are far less effective and the alcohol forms of these molecules are even less inductive. Furthermore, it has been known for some time that PGTs recognise various cellulose-containing membranes, since they stimulate AGT elongation, even though their surface is highly hydrophilic (Kobayashi et al., 1991; Carver et al., 1996). It is now established, however, that various cellulose degrading enzymes are produced by young germlings (Suzuki et al., 1998) and cellobiohydrolase I is released specifically at the PGT tip (Pryce-Jones et al., 1999) suggesting that enzymatic activity may generate cellulose breakdown products that act as signalling molecules perceived via the PGT. Similarly, B. graminis cutinase may release cutin monomers that can act as additional signalling molecules to fungi (Kolattukudy, 1996). Thus, Francis et al. (1996) showed that although glass coated with cutin monomers remained relatively hydrophilic, this treatment substantially enhanced induction of AGT formation. Conversely, treating leaves with cutinase inhibitor greatly reduced AGT formation. The current evidence therefore suggests that the plant leaf offers multiple factors recognised by the PGT. First it encounters leaf waxes and a combination of their hydrophobicity and chemical constitution can stimulate response. Then, as the peg emerges to penetrate the cuticle and make contact with the underlying cellulose matrix of the cell wall, fungal enzymes may release products that act as supplementary signal molecules. The multiplicity of these factors makes the plant surface more inductive than any artificial substratum possessing a single inductive characteristic (Carver et al., 1996). Molecular plant pathologists have recently started attempting to identify the intracellular signalling processes involved in germling morphogenesis, and in B. graminis it is becoming clear that signal transduction relay is complex. A simple signal is not sufficient to trigger germling morphogenesis. Indeed, multiple signals are needed to promote true differentiation and these are likely relayed by several signal transduction cascades, notably cAMP/PKA (protein kinase A) (Hall et al., 1999; Hall and Gurr 2000; Kinane et al., 2000), PKC (protein kinase C) (Zhang et al., 2001) and MAPK (mitogen activated protein kinase) (Bindslev et al., 2001; Zhang and Gurr, 2001; Kinane and Oliver, 2003). Of these, most is

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known about the cAMP/PKA pathway. Here, endogenous concentrations of cAMP, monitored in populations of conidia and germlings differentiating on host leaves or artificial surfaces (non-inductive glass or partially inductive cellulose membrane), revealed peaks in cAMP levels prior to both PGT and AGT emergence. Furthermore, this biphasic flux pattern was mirrored by changes in PKA activity (Kinane et al., 2000). Collectively, the model emerging from these studies is that upon landing, PGT emergence is mediated by a transient release of cAMP and a burst of PKA activity. Thereafter, surface perception by the PGT drives a second release of cAMP activity engaging MAPK cascade(s) (Kinane and Oliver 2003) and invoking AGT emergence. However, recent evidence points to appressorium formation being driven by the MAPK pathway alone (Kinane and Oliver 2003). Much remains to be unmasked, not least the precise link between host perception and signal transduction relay. Interest in understanding PGT/plant surface interactions arises from the possibility of plant breeding to alter the leaf surface so as to interfere with germling growth, and there is reason to believe this may be possible. For example, failure of PGTs to recognise leaf surface characteristics is not only a feature of interactions with certain non-hosts (Tsuba et al., 2002) but also occurs in species of the grass genus Lolium which are true hosts of the fungus (Carver et al., 1990). Here, PGTs recognise properties of the adaxial leaf surface on which the fungus forms functional appressoria and establishes thriving infection. In complete contrast, however, on the abaxial surface of Lolium leaves no infection develops and this is largely because most conidia fail to form an AGT. The reason for this is unknown although it obviously relates to the nature of the abaxial epicuticular waxes since their removal by chloroform washing restores inductivity. The physical appearance of the ab- and adaxial epicuticular waxes is quite different. On the abaxial surface they form amorphous overlapping sheets whereas those on the adaxial surface form crystalline plates similar in appearance to those seen on both surfaces of host cereal leaves. As would be predicted, some preliminary evidence (Carver et al., 1996) indicates differences in the chemistry of the waxes found on the two surfaces, but precise analyses and tests of individual components have not yet been performed. It seems possible, however, that some component(s) of the abaxial wax actively inhibit signal perception by the PGT because the intact surface is highly hydrophobic and the hydrophobicity of inert artificial substrata is sufficient to cause response (Carver et al., 1996, 1999; Wright et al., 2000). Understanding the basis of this effect may reveal means to develop novel forms of resistance that impede the pathogen’s early stages of development.

12.6

AGT growth, appressorium differentiation and penetration

According to their infection strategy, germ tubes of some fungi differentiate an apical appressorium relatively rapidly after emerging from the spore, while for others the germ tube may elongate considerably over a relatively long period before doing so. All available data indicate that appressorium differentiation is driven by perception of host-derived signals, that this depends upon intimate contact between elongating

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germ tube and substratum surface and that this in turn requires adhesion of the germ tube to the surface (Staples and Hoch, 1997; Tucker and Talbot, 2001; Apoga et al., 2004).

12.6.1 Adhesives associated with AGTs Adhesion to the hydrophobic leaf surface is pivotal to the disease process, both in anchoring the germling and as a prerequisite for full appressorium differentiation. In many cases, adhesion has been attributed to ECM released by the germ tube, although, as with the ECM associated with ungerminated spores (Section 12.2), the abundance of ECM varies greatly between species and for most there is little information on its chemical constitution or mode of action. For example, abundant ECM is visible by cryo-EM beneath the AGT of B. graminis (Figure 12.5b) and its release coincides with extremely strong adhesion (Wright et al., 2002a), but other than the fact that some components are water insoluble and that it contains protein, little is known of its chemistry (Carver et al., 1999). By contrast, the ECM associated with germ tubes of P. ampelicida is barely discernible by cryo-SEM although it is revealed by staining (Kuo and Hoch, 1995). Several attempts to characterise fungal components involved in adhesion have used lectins which bind to specific glycoproteins, and antibodies which recognise carbohydrate epitopes. Application of these lectins and antibodies blocks adhesion of numerous fungi e.g. C. graminicola (Mercure et al., 1994; Sugui et al., 1998), C. lindemuthianum (Hughes et al., 1999) and P. ampelicida (Kuo and Hoch, 1995) indicating the role of glycoproteins in adhesion. One class of proteins capable of mediating germ tube attachment are the hydrophobins, and these are key in M. grisea. Hydrophobins comprise a family of small cysteine-rich peptides which self-assemble into an amphipathic film at hydrophilic– hydrophobic interfaces, with the hydrophobic nature of the assembly aiding binding to the hydrophobic cuticle (Tucker and Talbot, 2001). Furthermore, hydrophobins may also play a role in sensing contact with a hydrophobic surface (Talbot et al., 1993; Beckerman and Ebbole, 1996). Thus, in M. grisea, deletion of the small, secreted hydrophobin gene mpg1 gave mutants showing reduced adhesion which in turn led to impaired appressorium formation and reduced pathogenicity (Talbot et al., 1996). More recently, attention has turned to the so-called ‘adhesins’, a family of ECM proteins carrying the RGD (arginine-glycine-aspartate) tripeptide motif, which are recognised by integrins on the host cell surface (Mellersh and Heath, 2001). This protein family is known to play key roles in cell adhesion, cytoskeleton scaffolding organisation and in intracellular signalling in a range of animal and microbial systems (Hostetter, 2000). Appressorium induction of Uromyces appendiculatus is blocked by exogenous application of RGD peptides, suggesting the presence of integrin-like molecules on the fungus or its host (Phaseolus vulgaris) leaves (Corrêa et al., 1996). Furthermore, in Oidium neolycopersici, the tomato powdery mildew fungus, application of RGDS (arginine-glycine-aspartate-serine) peptides

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leads to aberrant appressorial differentiation (Jones et al., 2001). However, various data cast doubt on a universal adhesion mechanism involving integrin-like proteins. Thus, RGD-peptide treatment does not affect germination or germ tube differentiation in B. graminis and whilst exogenous application of RGD peptides leads to enhanced adhesion this is directly attributable to the sticky surface residue left by the peptides (Perfect, 2005). Moreover, adhesion and morphology of the Oomycete Phytophthora megasperma f.sp. glycinea was unaffected by RGD-peptide treatment (Ding et al., 1994). Clearly, though vital to germling/host surface interaction, the mechanisms of germ tube adhesion are various but remain poorly understood.

12.6.1.1 Germ tube growth and appressorium differentiation by fungi that penetrate the host surface directly The germ tubes of many hemibiotrophic fungi differentiate appressoria after growing for only a short distance so that the appressorium forms very close to the mother spore. This implies rapid perception/response to features of the underlying surface. Speedy response not only maximises the likelihood that finite resources carried within the propagule will suffice to support appressorium formation and attempted penetration, but also minimises the time that the fungus is directly exposed to potentially adverse environmental influences (e.g. the presence or absence of free water, temperature extremes) before it penetrates the host to access the more stable internal environment of the leaf and establishes parasitism to support its nutritional requirements. For example, the germ tubes of C. graminicola differentiate appressoria within 3 h of emergence and differentiation is triggered by only 4.5 μm of continuous contact with a hard, hydrophobic artificial surface (Apoga et al., 2004). A comparable time-course of germination and appressorium formation is evident in other Colletotrichum spp. (C. coccodes, C. dematium and C. lagenarium; pathogens of red pepper) although here appressoria differentiate simply as a response to contact with a hard surface irrespective of its hydrophobicity (Ahn et al., 2003). In all these cases, therefore, the observations imply that a relatively simple perception system controls rapid differentiation. It is not possible to generalise on ‘simplicity’ within this fungal genus, however, because a much more complex perception system appears to operate in C. gloeosporoides. Here, contact with non-host waxes apparently inhibits appressorium formation whereas particular components of the host (avocado fruit) surface wax strongly stimulate appressorium differentiation (Kolattukudy et al., 2000). Relatively complex recognition systems also operate in M. grisea and P. ampelicida. The fully differentiated germ tubes of M. grisea formed on rice leaves and Teflon (hydrophobic surfaces) are generally very short, but differentiation is also stimulated on cellophane and glass (hydrophilic) although on these substrate germ tubes elongate far more before appressoria form (Jellito et al., 1994). Similarly, on its host grape leaves, P. ampelicida germ tubes also remain short, growing only about 5 μm before appressoria begin differentiating to become fully formed by 3–6 h after spore deposition (Kuo and Hoch, 1995). Here, differentiation also appears to involve hydrophobicity of the leaf surface since appressoria are induced

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on hydrophobic artificial substrata (Kuo and Hoch, 1996) although even on these substrata the germ tubes grow considerably longer (up to 40 μm) before differentiating. Thus, data from both M. grisea and P. ampelicida suggest that various signals can cause appressorium differentiation and that the true host may provide multiple cues that drive more rapid response. Among the obligate biotrophic fungi, AGT development and differentiation has been studied most intensively in E. pisi and B. graminis. The need for substratum contact as a driver of appressorial differentiation by E. pisi is easily demonstrated from the observation that the single germ tube formed by conidia suspended on spider’s suspension elongates to great length through the air (>100 μm) but never differentiates an appressorium (Carver et al., 1996), whereas, when incubated in darkness on pea leaves, the AGT swells and differentiates within a few microns (Ayres, 1983). As with M. grisea, a relatively simple surface perception system is indicated by published evidence. Apparently, substratum contact leads to a high frequency of appressorium differentiation irrespective of whether the surface is hydrophilic or hydrophobic (Ayres, 1983; Fujita et al., 2004b) although, like M. grisea, on hydrophilic surfaces far longer germ tubes are formed before appressoria differentiate (Carver et al., 1996). Nevertheless, because germ tube emergence close to the spore/substratum contact site is promoted on hydrophobic substrata (Section 12.4) frequencies of germ tube contact are higher on a hydrophobic artificial surface and here response is induced rapidly so that most appressoria differentiate within 4 h of spore deposition. A greater level of complexity is indicated, however, by very recent evidence that pea leaf wax constitution can also influence recognition by E. pisi germ tubes (Gniwotta et al., 2005). Thus, Gniwotta et al. (2005) found a far higher rate of appressorium differentiation on the adaxial (70%) than abaxial (49%) leaf surface, and this correlated to difference in the chemical constitution of waxes present on the different surfaces (see Section 12.3). Even further complexity in the E. pisi response system is demonstrated by the fact that lighting environment also has a marked effect: when incubated in lateral light, germ tubes differentiate appressoria only after growing to approximately three times the length of those formed in darkness (Ayres, 1983; Fujita et al., 2004b). Apparently, light superimposes a regulatory effect over germ tube response to contact stimuli. To some extent this is also true in M. grisea, although here germ tubes grow longer in darkness than in light (Jelitto et al., 1994). Although no such effect of light is seen in B. graminis (Fujita et al., 2004b), appressorial differentiation by the fungus can be driven by a variety of surface features implying that it possesses a number of recognition systems. These, however, do not induce rapid appressorial differentiation because, compared to e.g. Colletotrichum spp., M. grisea, P. ampelicida and E. pisi, on its host leaf B. graminis AGTs always elongate considerably (to around 40 μm) before differentiating appressoria. Evolution of this characteristic suggests an advantage to B. graminis pathogenesis despite the fact that growth must have the disadvantages of expending resources and taking time (around 6–8 h after germ tube emergence). It is likely that the benefit arises from the fact that elongation increases the probability of

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appressoria eventually forming on epidermal cells that have not been primed for defence by response to prior contact with PGTs of the fungus (Section 12.5). Studies (reviewed by Carver et al., 1996; Green et al., 2002) using artificial substrata indicate that the surface factors recognised by the B. graminis PGT (Section 12.5) are also recognised by elongating AGTs and induce their differentiation. Thus, surface hydrophobicity is inductive while simple contact with a hard surface is not. Breakdown products of leaf cutin and cellulose, released by fungal enzyme activities, are also implicated in different signalling systems. Together, these provide cues that act additively or interactively to stimulate appressorium differentiation with far greater efficiency on host leaf surfaces than on any artificial surface with a single inductive characteristic. Again, however, studies of artificial substrata do not reveal the full subtlety of interactions occurring with leaves, and recent studies indicate the influence that relatively small variations in plant surface chemistry may have on appressorial differentiation. It is well established that for B. graminis the physical presence of epicuticular leaf wax crystals is not required because appressoria differentiate normally even if the crystals are removed to expose the underlying cuticle proper before inoculation (Carver and Thomas, 1990). However, displacement of powdery mildew germ tubes from intact leaves reveals a distinct track where epicuticular waxes are missing, suggesting the possibilities that they either adhere to (and are lost with) the detached germ tube, or that they are ‘dissolved’ or degraded by components of the germ tube ECM (Staub et al., 1974). Support for the latter possibility comes from observations that B. graminis conidial ECM is capable of modifying the appearance of barley leaf surface waxes and rendering the surface more hydrophilic (Kunoh et al., 1990; Nicholson et al., 1993). If they are indeed degraded by fungal activity, their breakdown products may act as signals to the fungus. The ability of different wax components either to promote or to inhibit appressorium formation by C. gloeosporoides is well established (reviewed by Kolattukudy, 1996; Kolattukudy et al., 2000) and for B. graminis the importance of wax chemistry is indicated by a somewhat reduced frequency of appressorial differentiation on barley genotypes carrying certain eceriferum (wax) mutations (Yang and Ellingboe, 1972; Rubiales et al., 2001). The significance of relatively small differences in chemistry is further suggested from the finding that synthetic longchain aldehydes are more inductive of B. graminis appressorium differentiation than equivalent alcohols, and that the C26 chain length is more inductive than C30 chain length (Tsuba et al., 2002). The significance of wax chemistry in driving appressorial differentiation by plant pathogens clearly deserves further study; manipulation of leaf wax chemistry through plant breeding may offer a form of resistance that can disrupt the early stages of pathogenesis.

12.6.1.2 Signal transduction in fungi that penetrate the host surface directly The nature of signal transduction pathways engaged during appressorium formation has received a little attention. Recent evidence suggests that cAMP dependent (PKA) and MAPK pathways play a central and intertwined role driving differentiation of

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the appressorium in phytopathogenic fungi (reviewed by Lee et al., 2003). Most comprehensive amongst these studies are the data from M. grisea concerning the importance of cAMP signalling during appressorium differentiation in mutants lacking mac1 adenylate cyclase (Choi and Dean 1997; Adachi and Hamer 1998), mpg1 hydrophobin (Talbot et al., 1993), heterotrimeric G proteins components Gα, magB (Liu and Dean 1997), Gβ subunit, mgb1 (Nishimura et al., 2003) and cPKA (Xu and Hamer, 1996): all can be ‘rescued’ or ‘defect suppressed’ to varying degrees, by addition of cAMP. There is also growing evidence of the conservation of core elements of the MAPK signal pathway in pathogenic fungi and of interplay with the cAMP signal transduction cascade. For example, exogenous cAMP restores hooking and tip flattening in M. grisea  pmk1 mutants but not appressorial function. Far less is known, however, about signal transduction via PKC in phytopathogenic fungi. The signalling cascades mentioned may be linked to perception. However, it is extremely difficult to separate such signalling cascades from those involved in other aspects of fungal development. Thus, cAMP is known to influence growth and morphogenesis (Bencina et al., 1997) and infection structure differentiation, including appressorium formation in M. grisea (Lee and Dean, 1993). Indeed, in only one documented case is there a direct link between surface perception and signalling. This involves a lipid induced protein kinase (LIPK) from Colletotrichum trifolii, which is specifically and rapidly induced by purified plant cutin or longchain fatty acids that are monomeric constituents of cutin (Dickman et al., 2003). Gene replacement studies indicate that LIPK also plays a central role in triggering infection structure formation – deficient mutants were unable to form appressoria on hard surfaces and could not penetrate intact tissues, whereas over-expressors formed multiple abnormal appressoria. LIPK shares catalytic domain identity with PKCs and merits more study. Calcium/calmodulin-dependent signalling systems are involved in many biological systems and, amongst its functions within the fungi, Ca2+ is known to be required for appressorium formation by both P. ampelicida and C. trifolii (Warwar and Dickman, 1996; Shaw and Hoch, 2000). In M. grisea, exogenous application of calcium modulators, EGTA (a chelator) and calmodulin antagonists all inhibit appressorium formation although they do not affect germination. Indeed, EGTAinduced inhibition of appressorium formation is reversed by addition of CaCl2 (Lee and Lee, 1998). Similarly, in B. graminis calcium channel blockers, chelators and calmodulin inhibitors impeded appressorium differentiation (Hall, 1999). Collectively, these data attest to a broad role for Ca/calmodulin dependent signalling in appressorium formation.

12.6.1.3 Germ tube growth and appressorium differentiation by fungi that enter via stomata An immediate problem for fungi that enter the host through its stomata is that spore deposition occurs randomly and stomatal pores occupy a very small percentage of the leaf area: the probability of a chance encounter is extremely small if germ

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tube growth is unregulated. Much early work (reviewed by Staples and Macko, 1984; Hoch and Staples, 1991) focusing on this problem revealed that urediniospore germ tubes of many rust fungi show thigmotropic response to junctions between plant anticlinal cell walls. It became clear that germ tube growth orientates at right angles to these junctions and this maximises the likelihood of encountering a stoma. Even so, the germ tube may have to elongate for a great distance (up to several hundred micrometres) before either a stoma is located or reserves are exhausted. In some cases, leaf wax characteristics influence the ability of germ tubes to locate or recognise stomata. Thus, Puccinia hordei rarely forms appressoria over stomata of Hordeum chilense leaves, and this is thought due to their heavy encrustation with wax (Rubiales and Niks, 1996). In a more subtle way, certain eceriferum mutations in barley also affect P. hordei development, and this has been ascribed to influence surface wax properties and the ability of germ tubes to recognise anticlinal cell wall junctions and orient their growth (Rubiales et al., 2001). Strong evidence indicates that this thigmotropic sensing is contingent upon firm adhesion to the leaf surface and for Uromyces appendiculatus substrate hydrophobicity favours adhesion (Terhune and Hoch, 1993). On host leaves, the presence of epicuticular wax structures appears important for germ tube adhesion by Puccinia sorghi because on ‘waxless’ mutants of maize the germ tubes fail to adhere or orientate and therefore to locate or respond to stomata (Wynn and Staples, 1981). Germ tube ECM has been implicated in adhesion for a number of rust species (Beckett, 1990; Chaubal et al., 1991) and in P. sorghi the ECM consists of glycoproteins rich in acidic amino acids and β-1,3-glucan polymers, although there is likely to be variability in ECM constitution between rust species (Chaubal et al., 1991). When urediniospore germ tubes of most rust fungi encounter a stoma, the tip of the germ tube continues growth until it lies over the stomatal guard cells. Providing the germ tube is adhered, elongation ceases and the cytoplasm accumulates in the tip which differentiates into a swollen appressorium separated from the germ tube by a septal wall. This differentiation is known to be a further contact-mediated response that is shown by many different rust fungi (Allen et al., 1991). It has been best characterised in U. appendiculatus where studies using artificial substrata showed that germ tubes differentiate appressoria in response to encountering a surface ridge. Furthermore, maximal response is obtained following contact with a single ridge of height 0.5 μm; this correlates closely to the mean height of the lip of its host (P. vulgaris) guard cells (Hoch et al., 1987; Terhune et al., 1993). This extraordinary ability to recognise minute topographic details is arguably even more remarkable in certain Puccinia spp. There had been some controversy regarding the signals perceived by P. graminis and other cereal rusts because they respond poorly to single ridges or grooves, leading to suggestions that other signals associated with stomata may be more important stimulants of appressorium formation. However, although a range of chemical and environmental factors can stimulate differentiation (Hoch and Staples, 1991), Read et al. (1997) showed that while germ tubes of P. graminis tritici (wheat stem rust) and P. hordei (barley brown rust) did not respond significantly to single ridges, a very high proportion (83–86%) of germ tubes

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differentiated appressoria after encountering multiple ridges and grooves mimicking the topographic conformation of cereal stomatal complexes. Studies of U. appendiculatus by Hoch and co-workers provide the best insight into signal control of appressorium differentiation by a rust fungus (reviewed by Tucker and Talbot, 2001). Good evidence suggests that membrane stress imposed within the germ tube tip as it encounters a stomatal ridge engages a mechanosensitive ion channel capable of transporting various cations including Ca2+ . The fact that provision of Ca2+ in vitro can induce appressorium formation by U. appendiculatus is consistent with the proposition that Ca2+ flux plays a part in the differentiation process. Within 4 min of signal perception the cytoskeleton and cytoplasmic vesicles become reorganized along the cell wall of the germ tube tip and it seems likely that microtubule organisation is also involved in appressorium differentiation. That protease-treated germlings fail to respond to topographic signals supports the idea that trans-membrane proteins may link extracellular sensing proteins associated with the ECM to cytoplasmic proteins as part of the regulatory process. The involvement of integrins in this system is indicated from findings showing the reversible inhibitory effects of a number of RGD peptides on appressorium differentiation by U. appendiculatus (Corrêa et al., 1996).

12.7

Entry into the host leaf

12.7.1 Direct penetration of the host surface from appressoria Fungi may penetrate their host surface by physical force, by enzymatic degradation or by a combination of the two strategies. M. grisea offers the classic example of forceful entry into the host epidermal cell. This relatively unsubtle strategy can be extremely effective because host cell responses that involve minor modifications to strengthen the underlying plant surface/cell wall structures are unlikely to impede penetration. The sophistication of the strategy lies, however, in the complex processes of appressorium development that allow the generation of sufficient force to effect penetration. Cell collapse assays reveal that turgor within M. grisea appressoria can rise up to 8 MPa immediately preceding emergence of a specialised hypha, the penetration peg, that can breach not only the plant cuticle and outer cell wall to enter the epidermal cell lumen, but also has the capacity to penetrate extremely hard artificial substrata (Howard et al., 1991). To counteract the penetrative force, ECM glues the appressorium tenaciously to the surface. This enormous turgor is due to the accumulation of large quantities of glycerol, attributed to the action of triacylglycerolipase on lipids imported from the spore, which causes rapid water influx and the generation of hydrostatic pressure (reviewed by Tucker and Talbot, 2001). From an architectural perspective, the appressorium is able to contain the pressure by synthesis of a veneer of melanin that lines and strengthens the chitin-rich appressorial wall. This effectively seals the wall and allows turgor pressure to build up in the appressorium. Melanization is seen in appressoria of many other fungi

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including P. ampelicida and Colletotrichum spp., and in these too physical force is likely to be key to penetration. Thus the penetration peg of C. graminicola generates a force of around 16.8 μN, which is certainly sufficient to breach most plant cuticles (Bechinger et al., 1999). Nevertheless, the ability to generate turgor is not dependent on melanization because the hyaline appressoria of B. graminis are also capable of generating 2–4 MPa turgor pressure (Pryce-Jones et al., 1999). In this case, however, the evidence indicates that penetration is achieved by a combination of physical force and the secretion of cutin and cell wall degrading enzymes (Suzuki et al., 1998; Pryce-Jones et al., 1999). As appressoria mature and penetration pegs emerge and attempt to breach the host surface, host cells often respond by strengthening their wall and in many cases by depositing apoplastic wall appositions (papillae) beneath appressoria (Zeyen et al., 2002). These defensive responses clearly depend on passage of information through the plant cell wall and it is obviously important that plant cell walls have evolved to allow this transmission of information. One of the most rapid responses to pathogen attack is an oxidative burst focused in the plant cell wall/cytoplasm directly beneath the fungal appressorium. Thus, a fungal penetration peg entering the cell wall will be confronted with an oxidising environment which may disrupt further fungal development. It seems, however, that pathogens have developed antioxidative systems to cope with this problem. In B. graminis up-regulated expression of a catalase gene coincides with the timing of penetration and the CATB protein is secreted at the host–pathogen interface (Zhang et al., 2004). This is a closely controlled phenomenon that is dependent upon plant characteristics because no equivalent up-regulation of gene expression or focus of secreted protein occurs on an artificial surface. This data taken collectively with assays designed to detect fungal antioxidant activity, provoked speculation that this detoxification may play a role in B. graminis pathogenicity. Alternatively, it could be that antioxidants are needed to strengthen the fungal cell wall as it penetrates. However, neither hypothesis could be confirmed due to the lack of a robust transformation system in B. graminis. The idea was pursued, therefore, using M. grisea, and here a catalase B knockout mutant showed slightly attenuated pathogenicity and a slight reduction in cell wall strength (Henderson 2005). However, in M. grisea a knockout (MMT1) of a metallothionein (with high affinity for zinc) which acts as a powerful antioxidant, produced mutants unable to penetrate intact cuticle (Tucker et al., 2004) but able to cause disease if the cuticle was removed before inoculation. Cuticle removal obviously causes wound responses that are likely to include the rapid accumulation of plant reactive oxygen species. Thus, in M. grisea, MMT1 is more likely associated with differentiation of a fully functional cell wall, since MMT1 mutants are supersensitive to cell wall degrading enzymes, than in detoxifying plant-generated active oxygen species. With successful penetration of the cell wall and any defensive barriers erected in response to attempted entry, plant resistance to many pathogenic fungi probably depends upon factors that lie beyond the influence of the leaf surface. This may not be entirely true of the powdery mildew fungi. For these, the feeding structure

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(haustorium) develops as the tip of the penetration peg swells and differentiates within invaded epidermal cells, but the fungal hyphae are epiphytic. As they grow, these hyphae secrete ECM and hyphal adhesion is presumably as important to continued colony development as germ tube attachment is to the germling. However, to our knowledge, no studies have yet considered the effect that leaf surface features may have on development of established powdery mildew colonies.

12.7.2 Entry via stomata Successful penetration through the stomatal pore takes a fungus beyond the influence of leaf surface features and any defence these may offer. Although the need for an elaborate strategy to support entry through the stomatal pore may seem unnecessary, penetration is often achieved in darkness when stomata are closed. Unlike many fungi that directly penetrate epidermal cells, however, it is questionable whether those that enter via the stomatal pore employ hydrostatic force. Turgor pressure within U. appendiculatus appressoria is only 0.35 MPa, but it is possible that concentration of cytoskeletal elements in the penetration hypha (Mendgen et al., 1996) contributes to physical force which is capable of distorting the lip of artificial guard cells (Terhune et al., 1993). Nevertheless, entry is likely to be facilitated by enzymatic activity. In U. viciae-fabae, differentiation of appressoria and penetration hyphae is accompanied by secretion of a number of proteases and cellulolytic enzymes (Mendgen et al., 1996). Given that penetration is successful, the penetration hyphae swell within the substomatal cavity to form a vesicle from which infection hyphae emerge that grow endophytically until they contact a plant cell where they differentiate a haustorial mother cell and penetrate the host cell to form a haustorium. In the absence of resistance, endophytic hyphae ramify, many additional haustoria are formed, sporulation commences and the pustule ruptures the plant surface to release its spores.

12.8

Conclusions

Recent years have seen rapid progress in studies of interaction between leaf surfaces and certain pathogenic fungi (Gniwotta et al., 2005). In revealing the complexity of these interactions, however, we have exposed how little we truly understand of the basis of even the few favoured ‘model’ systems that have received attention. Coupled with conventional approaches, the advent of stable transformation systems applicable to both hosts and fungi and associated -omics technology, the continuing development of sophisticated techniques for molecular cell biological studies and advancement in methodologies for chemical analyses of small quantities, the coming decades promise deeper understanding. This will inevitably help towards unravelling the basis of host resistance mechanisms operating at the leaf surface to prevent or limit disease development. It is already clear that in compatible relationships, vital host–pathogen interactions commence within seconds of an encounter.

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The challenge is to identify phases of interaction where intervention to prevent or control loss due to disease is possible through, e.g. plant breeding or the development of novel targets for fungicides.

Acknowledgements We thank B.J. Thomas sincerely for providing all of the micrographs of B. graminis and E. pisi presented in this Chapter.

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Talbot, N.J., Kershaw, M.J., Wakley, G.E., deVries, O.M.H., Wessels, J.G.H. and Hamer, J.E. (1996) MPG1 encodes a fungal hydrophobin involved in surface interactions during infection-related development of Magnaporthe grisea, The Plant Cell, 8, 985–999. Terhune B.T. and Hoch H.C. (1993) Substrate hydrophobicity and adhesion of Uromyces urediospores and germlings, Experimental Mycology, 17, 241–252. Terhune B.T., Bojko, R.J. and Hoch H.C. (1993) Deformation of stomatal guard cell lips and microfabricated artificial topographies during appressorium formation by Uromyces, Experimental Mycology, 17, 70–78. Tsuba, M., Katagiri, C., Takeuchi, Y., Takada, Y. and Yamaoka, N. (2002) Chemical factors of the leaf surface involved in the morphogenesis of Blumeria graminis, Physiological and Molecular Plant Pathology, 60, 51–57. Tucker, S.L. and Talbot, N.J. (2001) Surface attachment and pre-penetration stage development by plant pathogenic fungi, Annual Review of Phytopathology, 39, 385–417. Tucker, S.L., Thornton, C.R., Tasker, K. et al. (2004) A fungal metallothionein is required for pathogenicity of Magnaporthe grisea, Plant Cell, 16, 1575–1588. Warwar, V. and Dickman, M.B. (1996) Effects of calcium and calmodulin on spore germination and appressorium development in Colletotrichum trifolii, Applied and Environmental Microbiology, 62, 74–79. Woolacott, B. and Archer, S.A. (1984) The influence of the primary germ tube on infection of barley by Erysiphe graminis f.sp. hordei, Plant Pathology, 33, 225–231. Wright, A.J., Carver, T.L.W., Thomas, B.J., Fenwick, N.I.D., Kunoh, H. and Nicholson, R.L. (2000) The rapid and accurate determination of germ tube emergence site by Blumeria graminis conidia, Physiological and Molecular Plant Pathology, 57, 281–301. Wright, A.J., Thomas, B.J. and Carver, T.L.W. (2002a) Early adhesion of Blumeria graminis to plant and artificial surfaces demonstrated by centrifugation, Physiological and Molecular Plant Pathology, 61, 217–226. Wright, A.J., Thomas, B.J., Kunoh, H., Nicholson, R.L. and Carver, T.L.W. (2002b) Influences of substrata and interface geometry on the release of extracellular material by Blumeria graminis conidia, Physiological and Molecular Plant Pathology, 61, 163–178. Wynn, W.K and Staples, R.C. (1981) Tropisms of fungi in host recognition, in Plant Disease Control: Resistance and Susceptibility (eds R.C. Staples and G.A. Toenissen), Wiley, New York, pp. 45–69. Xu, J.R. and Hamer, J.E. (1996) MAP kinase and cAMP signaling regulate infection structure formation and pathogenic growth in the rice blast fungus Magnaporthe grisea, Genes and Development, 10, 2696–2706. Yang, S.L. and Ellingboe, A.H. (1972) Cuticle layer as a determining factor for the formation of mature appressoria of Erysiphe graminis on wheat and barley, Phytopathology, 62, 708–714. Zeyen, R.J., Carver, T.L.W. and Lyngkjær, M.F. (2002) The papilla response, in The Powdery Mildews: A Comprehensive Treatise (eds R.R. Belanger, A.J. Dik, W.R. Bushnell and T.L.W. Carver), APS Press, Minnesota, pp. 107–125. Zhang, Z. and Gurr, S.J. (2001) Expression and sequence analysis of the Erysiphe graminis mitogenactivated protein kinase genes, mpk1 and mpk2, Gene, 266, 57–65. Zhang, Z., Henderson, C and Gurr, S.J. (2004) Blumeria graminis secretes an extracellular catalase during infection: potential role in suppression of host defence, Molecular Plant Pathology, 5, 537–548. Zhang, Z., Priddey, G. and Gurr, S.J. (2001) The Erysiphe graminis protein kinase C gene pkc1 and pkclike gene are differentially regulated during germling morphogenesis, Molecular Plant Pathology, 2, 327–338.

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

13 Plant–Insect interactions on cuticular surfaces Caroline Müller

13.1

Introduction

Insects explore the surface of a plant for many different purposes: they may search for an ideal food source or a spot to oviposit, for prey present on a plant or for a shelter to hide from predators, parasitoids or unfavourable abiotic conditions. The outermost surface of the primary parts of higher plants, the cuticle, is composed of the cuticular layer and the cuticle proper which is covered by epicuticular waxes. The certain chemical and physical properties of the cuticle (Chapter 4) influence the abilities of insects to move on the substrate and offer information about the characteristics and suitability of a plant (Eigenbrode and Espelie, 1995; Jenks and Ashworth, 1999; Müller and Riederer, 2005). The ability to attach to the surface is relevant on all trophic levels: plants show different cuticle properties that impede attachment of herbivores for tissue protection but also for catching insects in case of carnivorous plants. Herbivorous insects need a surface on which their legs and also their eggs will adhere and to which their mouthparts will have access. For predators and parasitoids the attachment on the plant surface will influence their predation and parasitation efficiency on herbivores. After successful contact with the plant cuticle, the chemical characteristics of the epicuticular waxes affect the acceptance or rejection behaviour the insect will show towards the plant. The chemical recognition of the suitability of a plant is particularly important for monophagous or oligophagous herbivores that can feed and develop only on one or a few plant species within one plant family. But also for generalists, the cuticular surface will offer valuable cues about acceptability of a plant. Classically, scientists have focused on secondary metabolites that are characteristic for certain plant species or families to link plant–insect relationships. However, it is becoming more and more clear that these compounds often will only play a role secondarily once an insect has damaged the plant cuticle and comes into contact with epidermal and mesophyll tissue. Only some secondary metabolites are indeed located in or on the cuticle. This chapter summarises the multifaceted aspects of the role of cuticular surfaces in plant–insect interactions.

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13.2

399

Access to the plant surface

13.2.1 Impeding attachment from the plant perspective Plants can be protected against herbivory by properties that hinder attachment of legs and eggs. The reproductive tissue of several plants is elevated on a greasy pole, a stem densely covered by epicuticular wax crystals, that prevents access of crawling insects (Juniper, 1995). Many buds, fruits and seeds are sticky, glaucous or coated with secreted mucilage which averts attack by insects (Juniper, 1995). Supracellular structures such as trichomes or thorns on the surface of leaves and stems can hinder crawling and walking of insects physically (Kerner, 1879; Jeffree, 1986; Romeis et al., 1999), hamper access of mouthparts to the nutritious tissue and negatively influence larval survival and oviposition behaviour of females (Städler, 2002). In several plant species a higher trichome density is induced by feeding of specific herbivores (Agrawal, 2000; Traw and Dawson, 2002) which increases morphological resistance when needed. The epicuticular waxes can impede locomotion due to physical and chemical characteristics (Müller and Riederer, 2005) and thereby avert access to stems, leaves, flowers and fruits for feeding, oviposition and nymphoposition (Kerner, 1879; Stork, 1980; Eigenbrode and Espelie, 1995; Ni and Quisenberry, 1997; Ni et al., 1998; Powell et al., 1999). Prominent waxblooms as well as bloomless glossiness, dependent on the presence and crystalline properties of the epicuticular wax layer, have been demonstrated to provide resistance to insects by influencing attachment (Stoner, 1990; Bodnaryk, 1992; Eigenbrode and Pillai, 1998; Eigenbrode, 2004). Loose crystalline material on the surface or an often observed epicuticular exfoliation are also discussed to reduce insect adhesion (Juniper, 1995; Eigenbrode, 1996). Carnivorous plants may derive some portion of their nutrients from insect prey. Highly adapted features of the cuticular surface are needed to trap and retain the insects. Downward pointing hairs are but one characteristic. The rim (peristome) of Nepenthes bicalcarata pitcher leaves causes an ‘aquaplaning’ by disrupting the attachment of the soft adhesive pads of visiting insects. This happens when the radial ridges of the smooth overlapping epidermal cells are completely covered by a liquid film from nectar and rainwater. Furthermore the surface topography provokes anisotropic friction for claws of the insect legs (Bohn and Federle, 2004). The slippery zone of Nepenthes alata pitchers is lined by platelet-shaped crystals, consisting mainly of triacontanal. These polymeric crystals are probably rather stable, resisting erosion by insect feet and thereby impeding attachment of approaching insects and of caught prey trying to escape (Riedel et al., 2003). In the bromeliad species Brocchinia reducta and Catopsis berteroniana (Bromeliaceae) the surface waxes consist of thread-shaped crystalloids that form a dense homogenous network. The crystals easily break off from the epidermis, thereby interacting with adhesive fluids secreted by the pads of the legs of visiting flies. The attachment of the fly is thereby hindered and insects will be trapped (Gaume et al., 2004).

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Some myrmecophilous plants are protected against natural enemies by living in symbiosis with ants. The epicuticular wax covers of different species of the genus Macaranga (Euphorbiaceae) form an efficient physical barrier against all ‘foreign’ non-specialised ant species, allowing only the associated ant species to climb and inhabit the stems (Federle et al., 1997).

13.2.2 Attachment from the insect perspective The attachment of herbivorous and carnivorous insects is in general higher on surfaces with reduced wax layers (Eigenbrode and Pillai, 1998; Eigenbrode and Kabalo, 1999; Eigenbrode and Jetter, 2002; Gorb and Gorb, 2002; Eigenbrode, 2004), but the reverse is also commonly seen (Eigenbrode and Espelie, 1995; Brennan et al., 2001; Eigenbrode and Jetter, 2002; Eigenbrode, 2004). Specialised morphological structures or behaviour might allow insects to overcome mechanical impediments of the plant cuticular surface that usually hamper attachment. Chrysoperla carnea (Stephen) (Neuroptera: Chrysopidae) produces mucilaginous secretions from an anal adhesive organ that mediate suction on waxy blooms (Eigenbrode, 1996). The tarsae of the chrysomelid beetle Hemisphaerota cyanea (Say) (Coleoptera: Chrysomelidae) are oversized and collectively bear some 60 000 adhesive bristles, each with two terminal pads. By touching ground with an increased number of these bristles, the beetle can improve a secure hold on the substrate (Eisner and Aneshansley, 2000). Each tarsus of the bug Coreus marginatus L. (Heteroptera: Coreidae) is provided with a pair of smooth flexible pulvilli adapted for attachment to the relatively smooth surface of its host plant Rumex crispus L. (Polygonaceae) (Gorb and Gorb, 2004). Weaver ants [Oecophylla smaragdina (Fab.), Hymenoptera: Formicidae] possess a flexible adhesion pad between the claws – the arolium – that allows, in combination with a wet adhesive secretion, strong attachment forces to smooth surfaces (Federle et al., 2002). The particular proportion between length of tibia and femur may enable the aphid Brevicoryne brassicae (L.) (Homoptera: Aphididae) to apply its tarsae perpendicular to the plant surface, thereby reducing lateral vectors that could hamper adhesion (Southwood, 1986). Walking on the edge of a leaf and thereby opposing the tarsae can have a similar effect, whereby more force is applied perpendicularly to the surface than by gravity (Eigenbrode, 1996). Hessian flies [Mayetiola destructor (Say), Diptera: Cecidomyiidae] prefer adaxial over abaxial leaf sides for oviposition due to more pronounced physical features such as grooves and ridges (Kanno and Harris, 2000a); possibly, the eggs attach better to this side compared to the relatively smooth abaxial side.

13.3

Recognition cues for insects

The cuticular surface of the plant is the primary contact zone for an approaching insect. Therefore, it is highly adaptive for a herbivore to recognise a suitable plant

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by the epicuticular waxes or secondary compounds present on the surface, without wasting energy and time on a non-host or even running the risk of poisoning by biting into unsuitable leaf tissue. Table 13.1 lists studies where effects of plant surfaces on herbivores are investigated. The references provided are those that were published after the thorough review by Eigenbrode and Espelie (1995) ten years ago on the effects of epicuticular lipids on insect herbivores or those that were not cited by these authors. Most studies (more than two-thirds) were done on crop plants, particularly on cabbages and cereals. A quarter of the studies include tree species (Table 13.1). Among the insect species studied in this context, one-third belong to Lepidoptera. Within the beetles, most is known on behaviour of chrysomelids on plant surfaces, within the Hymenoptera mainly ants were studied. From these data it becomes obvious that many further investigations using different plant as well as insect taxa are needed to gain a more general picture on effects of the cuticle on insects. The various behaviours of the insects that have been shown to be evoked by epicuticular waxes or surface extracts of plants are discussed in the following sections.

13.3.1 Deterrent properties By impeding locomotion or attachment through trichomes or various wax bloom properties on the plant surface (see earlier), access of insects to the plants is forestalled (Stoner, 1990; Bodnaryk, 1992; Eigenbrode and Pillai, 1998; Eigenbrode, 2004). However, insects able to walk on the plant cuticle can be hindered by a ‘secondary strategy’ from feeding or ovipositing. Sesquiterpenes are located in trichomes, but are easily released on the plant surface when the cuticular sac of the trichomes matures (Talley et al., 2002). These sesquiterpenes can be toxic to neonate larvae of Leptinotarsa decemlineata Say (Coleoptera: Chrysomelidae) (Carter et al., 1989; see Table 13.1). Feeding by several herbivores can induce amounts of secondary compounds such as alkaloids and flavonoids in trichomes, thereby altering the chemical composition of the plant surface to which these compounds are exuded. Such trichome exudates are known to act toxically or as a deterrent towards many insect species (Roda et al., 2003). Chemical compounds of the cuticle are known to evoke rejection behaviour, as e.g. 1-hexacosanol and short–chain fatty acids (C3 –C13 ) that prevent aphid settling (Phelan and Miller, 1982; Powell et al., 1999) or carboxylic acids and wax esters that have antixenotic properties, deterring feeding (Shepherd et al., 1995). Long-chain alcohols and amyrins reduce biting and increase walking time of neonate caterpillars [Plutella xylostella (L.), Lepidoptera: Plutellidae] on leaves of a cruciferous host (Eigenbrode and Pillai, 1998; see Table 13.1). Insect damage, such as punctures by aphids, has been shown to induce wax production (Bystrom et al., 1968). Deterrent compounds on the leaf are not always produced by the plant itself: destruxins derived from the entomopathogenic fungus Metarhizium anisopliae (Metschnikoff) Sorokin accumulate on the plant surface where they show

Papilio polyxenes F. (Lepidoptera: Papilionidae)

Leptinotarsa decemlineata Say (Coleoptera: Chrysomelidae) Mayetiola destructor (Say) (Diptera: Cecidomyiidae)

Daucus carota L. (Apiaceae)

Lycopersicon hirsutum f. hirsutum (Solanaceae)

Hexane

Leachates

Chrysomela scripta F. (Coleoptera: Chrysomelidae)

Ostrinia nubilalis Hbn. (Lepidoptera: Pyralidae)

Allium ampeloprasum L. (Alliaceae), Zea mays L. (Poaceae)

Long-chain primary alcohols and α-tocopherylquinone in varying concentrations and ratios Free amino acids, soluble carbohydrates (sugars)

Sugars involved in oviposition preference

Different acceptance for oviposition (13 components potential stimulants, 19 potential deterrents for oviposition) Feeding stimulants

Toxic to neonate larvae

Zingiberene (sesquiterpene)

Fifty different wax compounds

Oviposition stimulants

Flavonoid glycosides, chlorogenic acid

Chloroform (10 s) and subsequently near-boiling water (1 s) Compound in trichomes located on the leaf surface Hexane (20 s)

Evoked behaviour

Involved chemistry

Solvents for extraction/wax removal or source of surface compounds

Populus clones (Salicaceae)

Triticum aestivum cultivars (Poaceae) with different surface wax composition

Insect species (Order: Family)

Plant species (Family)

Derridj et al. (1996)

Coyle et al. (2003)

Cervantes et al. (2002)

Carter et al. (1989)

Brooks et al. (1996)

Reference

Table 13.1 Studies on attractive and deterrent effects of plant surface characteristics on herbivores. In general, leaves were tested, except where noted otherwise. Only references that were published after the review by Eigenbrode and Espelie (1995) or that were not cited by these authors are given

402 BIOLOGY OF THE PLANT CUTICLE

Leaf-cutter bees Megachile sidalceae Cockerell (Hymenoptera: Megachilidae) Fifth-instar Helicoverpa armigera Hbn. (Lepidoptera: Noctuidae)

Euphydryas chalcedona (Doubleday) (Lepidoptera: Nymphalidae) Delia floralis Fallen (Diptera: Anthomyiidae)

Chrysophtharta bimaculata Oliver (Coleoptera: Chrysomelidae)

Cercis canadensis var. mexicana (Rose) M. Hopk.) (Fabaceae), glossy and glaucous leaves Cajanus cajan (L.) Millsp. (Fabaceae), pods

Mimulus aurantiacus Curtis (Scrophularia-ceae)

Eucalyptus regnans F. Muell. (Myrtaceae), tip versus centre of leaves

Brassica genotypes (Brassicaceae)

Neonate Plutella xylostella (L.) (Lepidoptera: Plutellidae)

Brassica oleracea varieties (Brassicaceae)

Thirty-six wax components in varying proportions on tip and centre

Glucosinolates and other compounds

Dichloromethane and subsequently methanol (5 s per solvent) Chloroform (10 s)

Methanol

Quercetin-3-methyl ether, 3-hydroxy-4-prenyl5-methoxystilbene2-carboxylic acid (stilbene) Seven resins: ortho dihydroxy resins and methoxylated flavonoids

Methanol

Primary alcohols; a mixture of α- and β-amyrins; C14-alkanoic acid

Other compounds than glucosinolates as oviposition stimulants Oviposition preference for leaf tips probably not correlated with leaf chemistry

No correlation between different levels of resins and feeding

Reduce biting; reduce the time of biting and increase the time of walking; increase the time of palpating More cuttings on leaves with crystalline adaxial surface wax than on glabrous Feeding stimulant

Continued

Howlett and Clarke (2003)

Hopkins et al. (1997)

Hare (2002)

Green et al. (2003)

Eigenbrode et al. (1999)

Eigenbrode and Pillai (1998)

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

403

Populus deltoides x P. nigra (Salicaceae)

Brassica napus L. (Brassicaceae), glossy and waxy strains and herbicide-treated Six grasses (Poaceae)

Chrysomela scripta F. (Coleoptera: Chrysomelidae)

Mayetiola destructor (Say) (Diptera: Cecidomyiidae)

Plutella xylostella (L.) (Lepidoptera: Plutellidae)

Atta cephalotes L. (Hymenoptera: Formicidae) Delia radicum (L.) (Diptera: Anthomyiidae)

Forty-two tropical species

Brassica oleracea cv. botrytis CC Cross (Brassicaceae)

Insect species (Order: Family)

Plant species (Family)

Table 13.1 Continued

Hexane (30 s)

n−Docosanol, n-tetracosanol, n-hexacosanol (C26 ), n-octacosanol (C28 ), n-triacontanol (C30 ), α-tocopherylquinone

Oviposition preferences on surface extracts similar to those on real plants Feeding stimulants

Justus et al. (2000) Oviposition preference for glossy and herbicide-treated leaves

Cool (10◦ C) dichloromethane (50 s)

Hurter et al. (1999)

Oviposition stimulants

1,2-Dihydro-3-thia4,10,10b-triazacyclopenta[.a.]fluorene-1carboxylic acid (CIF)1

Lin et al. (1998)

Kanno and Harris (2000b)

Hubbell et al. (1984)

Repelled fungus-growing ants

Terpenoids, steroids and waxes

Soxhlet extracts in chloroform of whole leaves Dichloromethane and subsequently two times methanol (5 s per solvent)

Reference

Evoked behaviour

Involved chemistry

Solvents for extraction/wax removal or source of surface compounds

404 BIOLOGY OF THE PLANT CUTICLE

Tanacetum vulgare L. (Asteraceae)

Triticum aestivum L. (Poaceae)

Several ant-species (Hymenoptera: Formicidae)

Different Macaranga ant-plants (Euphorbiaceae), stems Eucalyptus maculata Hook. (Myrtaceae)

Atta sexdens rubropilosa Forel (Hymenoptera: Formicidae) Mayetiola destructor (Say) (Diptera: Cecidomyiidae) Cassida stigmatica Suffr. (Coleoptera: Chrysomelidae)

Plutella xylostella L. (Lepidoptera: Plutellidae)

Water or methanol; chloroform or hexane (5 min)

Lobesia botrana Den. & Schiff. (Lepidoptera: Tortricidae) Delia radicum L. (Diptera: Anthomyiidae)

Hexane/cellulose acetate

Arrestants to females

Continued

Müller and Hilker (2001)

Morris et al. (2000)

Oviposition stimulants 1-Octacosanal and 6-methoxy-2benzoxazolinone

Dichloromethane

Markstädter et al. (2000)

Marsaro et al. (2004)

Slippery for ants

Marazzi et al. (2004b)

No bioactivity

Large amounts of triterpenoids form crystalline wax blooms

Oviposition stimulants

Glucosinolates

Marazzi et al. (2004a)

Maher and Thiery (2004)

Oviposition stimulants; not stimulating Oviposition stimulants

Lombarkia and Derridj (2002)

Oviposition stimulants

Glucosinolates and CIF1

Sugars and sugar alcohols (in particular fructose, sorbitol, myo-inositol)

Dichloromethane (3 × 30 s)

Chloroform and subsequently two times in methanol (5 s per solvent) Chloroform and subsequently two times in methanol (5 s per solvent)

Ultra-pure water (spraying)

Cydia pomonella L. (Lepidoptera: Tortricidae)

Brassica napus cv. Express (Brassicaceae)

Brassica napus cv. Express (Brassicaceae)

Malus domestica var. golden delicious (Rosaceae), fruits and leaves Continued Vitis vinifera cv. Chasselas (Vitaceae), grapes

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

405

Phaedon cochleariae (F.) (Coleoptera: Chrysomelidae)

Tupiocoris notatus Distant (Heteroptera: Miridae)

Brassica napus L. var. ‘Martina’, Nasturtium officinale R. Br. (Brassicaceae)

Nicotiana attenutata Torr. Ex Wats. (Solanaceae)

Aphis fabae Scopoli, Sitobion avenae F. (Homoptera: Aphididae)

Diuraphis noxia (Mordvilko) (Homoptera: Aphididae) Diuraphis noxia (Mordvilko) (Homoptera: Aphididae)

Five genotypes of Triticum aestivum L. (Poaceae)

Triticum aestivum L. c.v. ’Arapahoe’ and ’Halt’, Hordeum vulgare L. c.v. ’Morex’, Avena sativa L. c.v. ’Border’ (Poaceae) Vicia faba L. (Fabaceae, host); Avena sativa L: (Poaceae, non-host)

Insect species (Order: Family)

Plant species (Family)

Table 13.1 Continued

1-Hexacosanol predominant on A. sativa

Glandular trichomes, content excreted onto the leaf surface

Flavonols: quercetin and seven methylated derivatives

Powell et al. (1999) Triggered stylet penetration; non-host only accepted after wax removal No stimulatory activity by wax extracts and/or glucosinolates; feeding preference for de-waxed leaves Quercetin as feeding attractant

Roda et al. (2003)

Reifenrath et al. (2005)

Ni et al. (1998)

Ni and Quisenberry (1997)

More or less antixenotic

Differences in density of trichomes and wax flakes Only limited influence of wax removal on probing and nymphoposition preferences

Reference

Evoked behaviour

Involved chemistry

Chloroform : methanol: Minor amounts of glucosinolates in water (2 :1 : 1) solvent extracts (3 × 20 s); gum arabic as adhesive

Chloroform (10 s)/cellulose acetate

Ethyl ether

Solvents for extraction/wax removal or source of surface compounds

406 BIOLOGY OF THE PLANT CUTICLE

Amphorophora idaei Börner (Homoptera: Aphididae)

Plutella xylostella L. (Lepidoptera: Plutellidae)

Brassica oleracea L. (Brassicaceae)

Amphorophora idaei Börner (Homoptera: Aphididae)

Atta sexdens rubropilosa Forel (Hymenoptera: Formicidae) Helicoverpa armigera Hbn. (Lepidoptera: Noctuidae) Helicoverpa armigera Hbn. (Lepidoptera: Noctuidae)

Rubus idaeus L. cultivars (Rosaceae)

Cajanus scarabaeoides (L.) Thouars (Fabaceae) Cajanus cajan (L.) Millsp. and C. platycarpus (Fabaceae) Rubus idaeus L. cultivars (Rosaceae)

Didymopanax vinosum Marchal (Araliaceae)

Spencer (1996), Spencer et al. (1999)

Oviposition stimulants

Artificially mixed

Continued

Shepherd et al. (1999b)

Different resistant properties to aphids

Dichloromethane (10 s)

Shepherd et al. (1999a)

Settling preference on leaves with greater wax coverage and higher levels of the compounds of shorter chain length

Dichloromethane (10 s)

Sterols, particularly cycloartenol, branched alkanes, presence or absence of C-29 ketones and symmetrical C-29 secondary alcohol, and other compounds Higher levels of cycloartenyl esters, α-amyryl esters and other compounds in wax from the resistant cultivar n-Alkane mixture in combination with glucosinolate

Shanower et al. (1997)

Feeding stimulants

Shanower et al. (1997)

Salatino et al. (1998)

Acetone

Deterrent for some colonies

Feeding deterrents

Lupeol, primary n-alcohols

Water

Chloroform (three times for 30 s)

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

407

Leptinotarsa decemlineata Say (Coleoptera: Chrysomelidae)

Solanum berthaultii Hawkes (Solanaceae)

1 CIF, cabbage identification factor.

Pieris brassicae (L.) (Lepidoptera: Pieridae)

Brassica oleracea L. (Brassicaceae)

Dichloromethane and subsequently in methanol (3 s per solvent) Ice-chilled methylene chloride (10 + 10 + 5 s); further fractionated

Pentane (20 s)

n-Alkanes (hexacosane, heptacosane, octacosane, nonacosane, tritriacontane) Glucobrassicin (3-indolyl-methylglucosinolate)

Chloroform (three times for 30 s)

Atta sexdens rubropilosa Forel (Hymenoptera: Formicidae)

Ostrinia nubilalis Hbn. (Lepidoptera: Pyralidae)

Differences in several wax esters between waxy and glossy leaves

Hexane (50 min)

Mnesampela privata (Guenée) (Lepidoptera: Geometridae)

Eucalyptus globulus subsp. pseudoglobulus (Naudin ex Maiden) Kirkpatr. and putative hybrid (Myrtaceae) Nine woody species

Zea mays L. (Poaceae)

Involved chemistry

Solvents for extraction/wax removal or source of surface compounds

Insect species (Order: Family)

Plant species (Family)

Table 13.1 Continued

Steinbauer et al. (2004)

Udayagiri and Mason (1997)

van Loon et al. (1992)

Yencho et al. (1994)

Oviposition preference for waxy leaves and young leaves; reduced performance of offspring on such leaves Different wax extracts caused different responses (no negative influence, slight and strong deterrence) Oviposition stimulants

Oviposition stimulant

Feeding deterrents, mainly in non-volatile fraction

Sugayama and Salatino (1995)

Reference

Evoked behaviour

408 BIOLOGY OF THE PLANT CUTICLE

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

409

antifeedant properties (Amiri et al., 1999). Compounds derived from yeasts, possibly polyamines, can prevent oviposition (Städler, 2002). Generalist herbivores might use surface chemicals as cues in associative learning of aversion. The polyphagous grasshopper Schistocerca americana (Drury) (Orthoptera: Acrididae) first has to bite into leaf tissue to recognise an unpalatable plant but afterwards, palpation of the surface is sufficient to evoke rejection (Chapman and Sword, 1993). Larvae and adults of the tansy leaf beetle Galeruca tanaceti (L.) (Coleoptera: Chrysomelidae) prefer to feed on leaves of Tanacetum vulgare L. (Asteraceae) where waxes are removed with cellulose acetate treatment (Müller and Hilker, 2001) over intact leaves, while they do not discriminate between waxy and de-waxed leaves of Brassica pekinensis (Lour.) Rupr. (Brassicaceae) (Figure 13.1). This generalist has a rather poor performance with long (a)

Larval feeding (%) n.s.

100 80 60 40 20 0

(b)

Tanacetum vulgare

Brassica pekinensis

Adult feeding (%) n.s.

80 60 40 20 0

Tanacetum vulgare

Brassica pekinensis

Figure 13.1 Percentage of individual larvae (a) and adults (b) of Galeruca tanaceti, feeding on intact (black bars) or de-waxed (white bars) leaf parts of the host plants Tanacetum vulgare and Brassica pekinensis. Leaflets (T. vulgare) or leaf discs (B. pekinensis) were dipped half in cellulose acetate dissolved in acetone (5–10% w/v). After evaporation of acetone, a white film that appeared on the surface of the treated side was carefully removed by forceps. Leaves of one species were offered to individual secondinstar larvae or adults (N = 20 per host plant and per developmental stage. ∗ P < 0.05; ∗∗∗ p < 0.001; n.s., not significant; Wilcoxon signed-rank test for paired differences, two-sided). While G. tanaceti avoids waxes of the poor host plant T. vulgare, waxes of the highly suitable host plant B. pekinensis (cultivar without trichomes) do not influence feeding.

410

BIOLOGY OF THE PLANT CUTICLE

developmental times and a low pupal weight when reared on T. vulgare compared to B. pekinensis (Müller, 1999). The surface waxes of T. vulgare presumably imply negative information for this generalist, either chemically or mechanically. The specialist beetle Phaedon cochleariae (F.) (Coleoptera: Chrysomelidae) prefers de-waxed over intact leaf parts of its host plants Nasturtium officinale R. Br. and Brassica napus L. var. Martina (Brassicaceae) (Reifenrath et al., 2005). In this case, epicuticular waxes might cover access to or ‘hide’ feeding stimulants present in leaf cells of the interior. This might be an adaptive trait for the protection of the plant against herbivorous specialists. Leaf-cutting, fungus-growing ants have been shown to be repelled or deterred to different degrees by terpenoids, steroids and waxes extracted from various potential host plants (Hubbell et al., 1984; Sugayama and Salatino, 1995). Certain compounds of epicuticular waxes of Didymopanax vinosum Marchal (Araliaceae) were only deterrent for some but not all colonies of Atta sexdens rubropilosa Forel (Hymenoptera: Formicidae) (Salatino et al., 1998; Table 13.1).

13.3.2 Attractive properties Many studies focus on secondary plant compounds that are characteristic for certain plant species or families and should therefore be used by insect specialists for host recognition (Harborne, 1995). Within the last two decades several speciesspecific wax compounds of the cuticle have been determined (Chapter 4) that could mediate the host-finding process. Waxy compounds such as long-chain alkanes, alcohols, carboxylic acids, as well as secondary metabolites such as quinones and flavonoids located at the plant surface have been shown to be involved in feeding stimulation (Adati and Matsuda, 1993; Lin et al., 1998; Coyle et al., 2003; Green et al., 2003). More studies dealt with oviposition stimulants and revealed various alkanes, aldehydes, flavonoids, chlorogenic acids, glucosinolates, the ‘cabbage identification factor’ (1,2-dihydro-3-thia-4,10,10b-triaza-cyclo-penta[.a.]fluorene1-carboxylic acid) (Brooks et al., 1996; Udayagiri and Mason, 1997; Hurter et al., 1999; Spencer et al., 1999; Marazzi et al., 2004a,b) as well as sugars that leak on the plant surface (among others released from extrafloral nectari es) (Derridj et al., 1996; Lombarkia and Derridj, 2002; Leveau, 2004) as active principles in host recognition (see Table 13.1). In several insect species, epicuticular wax compounds act in synergism with secondary metabolites as oviposition stimulants (Roessingh et al., 1992; Spencer, 1996; Spencer et al., 1999; Morris et al., 2000; Marazzi et al., 2004a). A given herbivorous species, such as for example the Colorado potato beetle (Leptinotarsa decemlineata) or the Cotton bollworm Helicoverpa armigera Hbn. (Lepidoptera: Noctuidae), can distinguish between closely related plant species primarily by infochemicals perceived at or near the leaf surface (Harrison, 1987; Shanower et al., 1997). Waxes of a given host plant, on the other hand, can comprise opposing information for herbivores with different degrees of specialisation. While the polyphagous generalist G. tanaceti is deterred by waxes of leaves of the less

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

(a)

411

Larval feeding (mg) 1.4

n.s.

1.2 1.0 0.8 0.6 0.4 0.2 0.0 Tanacetum vulgare

(b)

Adult feeding (mg) 1.6 n.s. 1.4 1.2 1.0 0.8 0.6 0.4 0.2 0.0 Tanacetum vulgare

Cirsium arvense

n.s.

Cirsium arvense

Figure 13.2 Mean amount (±se) of individual third-instar larvae (a) and adults (b) of Cassida denticollis, feeding on intact (black bars) or de-waxed (white bars) leaf parts of the host plant Tanacetum vulgare and the non-host Cirsium arvense. Leaflets were dipped half in cellulose acetate dissolved in acetone (5–10% w/v). After evaporation of acetone, a white film that appeared on the surface of the treated side was carefully removed by forceps. Leaves were offered to individual third-instar larvae or adults (N = 20 per host plant and per developmental stage. ∗ P < 0.05; n.s., not significant; Wilcoxon signed-rank test for paired differences, two-sided).

suitable host T. vulgare (see earlier and Figure 13.1) the oligophagous specialist Cassida denticollis Suffr. (Coleoptera: Chrysomelidae) that feeds on a few species of Asteraceae only, seems to be not influenced by waxes. Third-instar larvae and adults of C. denticollis do not discriminate between intact or de-waxed leaflets of T. vulgare and accept both in a comparable manner. Leaflets of the non-host Cirsium arvense (L.) Scop. are not accepted for feeding, neither with nor without cuticular waxes, by C. denticollis (Figure 13.2). For the strictly monophagous specialist Cassida stigmatica (Coleoptera: Chrysomelidae), epicuticular waxes of T. vulgare constitute an important recognition cue in oviposition stimulation. Females of C. stigmatica prefer to lay eggs on the abaxial leaf side of the plant. While they do not discriminate between leaves with and without waxes when being offered the adaxial side only,

412

BIOLOGY OF THE PLANT CUTICLE

they clearly prefer to oviposit on intact leaves offered with the abaxial side over de-waxed parts (Müller and Hilker, 2001). As the chemistry of the epicuticular waxes also influences the fine structure of the cuticular surface (Jeffree, 1986; Markstädter et al., 2000), it cannot easily be differentiated if the insect might use mainly the chemical and/or mechanical information for its host plant selection process (Eigenbrode and Espelie, 1995; Kanno and Harris, 2000a,b). Due to the chemical composition and micromorphology, optical characteristics of the cuticular surface are highly affected (Chapter 6). Visual cues, such as UV-reflectance, might be important recognition properties for pollinators, dispersers and non-pollinating herbivores (Juniper, 1995). However, a causal link between visual cues of the cuticle and attractiveness to herbivores has rarely been addressed (Eigenbrode and Espelie, 1995; Justus et al., 2000; Steinbauer et al., 2004). Preferences of insects can be significantly influenced by leaf colour which is a result of wavelength reflectance of different intensities (due to surface properties). This has been shown for example for the leafhopper Empoasca fabae (Harris) (Homoptera: Cicadellidae) (Bullas-Appleton et al., 2004). The colour perception of an insect can be further affected by reflection polarisation, again determined by wax structure (Grant et al., 1993), if it has a polarisation-dependent colour vision system. Papilio butterflies are sensitive to such interferences (Kelber, 1999; Kelber et al., 2001). Perception of polarisation reflection might enable them to discriminate shiny from matt surfaces (Horváth et al., 2002) which are in turn determined by the presence of a waxy layer or other microstructures. It is likely that visual and chemical cues of the surface act in concert in the host recognition process of an insect (Prokopy and Owens, 1983; Degen and Städler, 1997). For predators, an infestation of a plant with potential prey might be recognised by changes in the visual appearance caused by the herbivore feeding. Sawfly damage induces amongst others an increase of surface flavonoids in the leaves of mountain birch (Betula pubescens, Ehrh., Betulaceae) which affects the UV spectral maxima. Predators such as birds could well respond to these cues to detect their prey (Mäntylä et al., 2004). The pupal parasitoid Pimpla turionellae (L.) (Hymenoptera: Ichneumonidae) has been demonstrated to perceive changes in the chromatic and achromatic plant surface appearance that are due to an infestation with its endophytically living host (Fischer et al., 2004).

13.4

Mimicry

Mimicry systems are characterised by an organism (mimic) simulating signal properties of a second organism (model) which are in turn recognised by a third organism (operator). The organism that mimics a model has an advantage due to the fact that the operator falsely identifies it as a model (Vane-Wright, 1980). In several cases mentioned later it is not entirely unambiguous to determine the model or operator organism. Nevertheless, similarities of two organisms in complex traits like hydrocarbon patterns imply an ecological meaning.

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

413

13.4.1 Mimicry from the plant perspective Plants have evolved intriguing mimicry systems to attract insects as pollinators or dispersers by false cues. They might be either mimicking food or brood resources by odours or contact stimuli that visually and/or chemically resemble e.g. fresh or rotten fruit or by odours that actually mimic sex, brood or alarm pheromones of the cheated insect (Dettner and Liepert, 1994). When volatiles are attractive, they need to be emitted from the plant surface, and are either produced there directly by trichomes or in the interior of the plant and subsequently transported to the outer wax layer (Chapter 7). In many of the mimicry systems, hydrocarbons are involved that mediate signals in the short range as contact cues. The plant family of Orchidaceae is probably best known to include a high number of plants that need very specific insect species as pollinators and therefore have evolved diverse strategies to attract them, often without giving them an actual reward (Wiens, 1978). Compounds in orchid species resembling female sex pheromones of the respective pollinators have been identified as C11 –C19 unsaturated and saturated hydrocarbons, primary and secondary alcohols, aliphatic short and long-chain esters and mono- and sesquiterpenes and their esters (Dettner and Liepert, 1994). The relative proportions of the hydrocarbon compounds can however differ between the orchid and the pollinating insect (Schiestl et al., 2000). Also, some unique compounds such as trace amounts of (ω-1)-hydroxy and (ω-1)-oxo acids, especially 9-hydroxydecanoic acid, were identified to be released from an orchid whose flowers seem to be even more attractive to pollinating male wasps than their females (Ayasse et al., 2003). Seeds of several tropic epiphytes are dispersed by ants. In extract of these seeds commonly 6-methyl salicylic acid methyl ester was detected which triggers a dosedependent response in Camponotus femoratus (F.) (Hymenoptera: Formicidae). This compound also occurs in the mandibular glands of Camponotus ants and acts as a brood pheromone (Davidson et al., 1990). To stimulate ants to carry the seeds, 6-methyl salicylic acid methyl ester must be detectable at the surface. The sesquiterpene hydrocarbon (E)-β-farnesene is present in glandular hairs and is continuously released from intact wild potato plants. As this sesquiterpene is also the alarm pheromone of several dozen aphid species it is an efficient way of the plant to keep aphids away (Gibson and Pickett, 1983).

13.4.2 Mimicry from the insect perspective Espelie (Espelie and Bernays, 1989; Espelie and Brown, 1990; Espelie et al., 1991) noted that plant surface chemicals could be transferred to the insect surface either by direct contact or metabolically. These plant chemicals could either be derived from trichomes as for example found in herbivores feeding on Solanaceae, or might be a powder consisting of long-chain alcohols of the plant epicuticular waxes deposited on the insect cuticle. Chemicals mimicking the plant surface might be also dietary (Espelie and Bernays, 1989) or even synthesised de novo by an insect (Espelie and Brown, 1990). It is suggested that such plant chemicals on the insect surface might

414

BIOLOGY OF THE PLANT CUTICLE

help the herbivore to be cryptic to predators and parasitoids (Espelie et al., 1991). A social wasp that nests in trees of Acacia collinsii Saff. (Fabaceae) is probably protected against aggression by an ant species occupying these Acacia trees due to a mimicry system. The hydrocarbon pattern, characterised by certain alkene–alkane pairs, is not only similar between the surface wax of the wasp and the ant, but also n-alkanes of the surface wax of the wasp nest resemble those detected in the surface wax of acacia thorns in which the ants live (Espelie and Hermann, 1988).

13.5

Methods of investigation

To investigate the role of plant epicuticular waxes on the behaviour of visiting insects, various approaches have been employed. On the one hand, mutants or genotypes of plant species differing in wax morphology and chemistry are used to test influences of the surface characteristics on attachment and acceptability by herbivores and carnivores (e.g. Bodnaryk, 1992; Ni et al., 1998; Eigenbrode and Kabalo, 1999; Kanno and Harris, 2000b; Cervantes et al., 2002; Chang et al., 2004). Even within a given plant, the epicuticular waxes of leaves can strongly vary with developmental stage and thus offer a genetically identical substrate for elegant comparative work (Brennan et al., 2001; Steinbauer et al., 2004). On the other hand, or additionally, waxes are extracted from plant material (leaves, stems, fruits) in various ways (see later). These extracts can be applied on neutral substrates and offered in bioassays to test for the activity of wax compounds (e.g. Sugayama and Salatino, 1995; Hurter et al., 1999; Kanno and Harris, 2000b; Cervantes et al., 2002; Lombarkia and Derridj, 2002). Such extracts are also used to test for electrophysiological responses (e.g. Hurter et al., 1999; Justus et al., 2000; Steinbauer et al., 2004). Bioassay-guided fractionation might help to isolate the active principles within the plethora of surface compounds (Malony et al., 1988; Foster and Harris, 1992; Yencho et al., 1994) that can be analysed by chemical analysis (see Table 13.1). Extracts are often gained by dipping plant material shortly in solvents of different polarity (see Table 13.1). However, a drawback of this extraction procedure is the fact that compounds might be derived from locations other than the actual outermost epicuticular leaf surface (Roessingh et al., 1992; Riederer and Markstädter, 1996; Jetter et al., 2000) and mostly not sampled from one specific leaf side only. If leaves are kept in darkness for a few hours before extraction, stomata will be at maximal closure. This lowers the risk of washing out compounds from the leaf interior through treatment with solvents (Reifenrath et al., 2005). Wax load as well as chemical composition can vary on abaxial and adaxial leaf sides (Premachandra et al., 1993; Yang et al., 1993a; Eigenbrode and Pillai, 1998; Eigenbrode et al., 1999; Reifenrath et al., 2005). Accordingly, many insects show oviposition preferences for one leaf side that might be determined by differences in chemistry and structure (Kanno and Harris, 2000a; Müller and Hilker, 2001).To probe leaf sides separately, intact leaves can be placed on a flexible rubber mat, and a glass cylinder gently pressed onto the exposed surface. In these cylinders

PLANT–INSECT INTERACTIONS ON CUTICULAR SURFACES

415

solvents are shortly agitated and then removed for testing (Premachandra et al., 1993; Jetter et al., 2000; Reifenrath et al., 2005). A third way to investigate how insects are influenced by the cuticle is the removal of epicuticular waxes and test of the remaining tissue. Such de-waxed plant material can then be offered against intact material to test for behavioural answers (Powell et al., 1999; Müller and Hilker, 2001; Reifenrath et al., 2005; Figures 13.1 and 13.2). When waxes are removed by dipping plant material in solvents, the chemistry of the surface and the interior can be drastically changed. Therefore it is more suitable to remove waxes mechanically. A (side-)specific mechanical removal of epicuticular waxes without damaging underlying tissue could be achieved by treatment with cellulose acetate (Powell et al., 1999; Müller and Hilker, 2001) or gum arabic (Jetter et al., 2000; Jetter and Schäffer, 2001; Riedel et al., 2003; Reifenrath et al., 2005), respectively. After evaporation of solvents (acetone or water, respectively), both adhesives form a film that can be removed. In case of gum arabic, the polymerous film can be dissolved in a solvent for further chemical analysis (Riedel et al., 2003; Reifenrath et al., 2005). Frozen liquids can be used as a cryo-adhesive (Ensikat et al., 2000; Riedel et al., 2003). For this purpose, cylinders are gently pressed on a leaf surface, filled with glycerol or water and frozen in liquid nitrogen. The epicuticular waxes will stick to the frozen liquid and allow to test the wax film (after evaporation of the liquid) or the de-waxed leaf in a bioassay, and to analyse the chemical composition of the waxes.

13.6

Application in biological pest management

In many crop plants varieties exist that differ in wax cover, wax composition or trichome density and therefore are more or less suitable for a herbivorous pest. For example, glossy plant mutants of Brassica oleracea L. (Brassicaceae) are more prone to water stress. This can cause an accumulation of deterrent compounds which in turn reduces feeding attacks by aphids (Cole and Riggal, 1992). Trichomes have been shown to be efficient physical barriers against herbivorous pests for a number of important crop plant species within the Brassicaceae, Poaceae, Malvaceae (cotton) and other plant families (Peter and Shanower, 2001). However, altering just one of the plant surface features, through traditional breeding or genetic engineering, might not have the desired economic benefit: for herbivorous insects a variety of cues are important in the host acceptance process, detected by chemo- as well as mechanoreceptors and playing in concert together. Therefore, reliable predictions about the feasibility of breeding antixenotic resistance are difficult (Kanno and Harris, 2000b). Furthermore, the selection of a certain genotype that is less suitable for one insect pest species, might impact suitability to other organisms. Other insect pests or fungi could be even more attracted. Wax characteristics also influence visiting predators or parasitoids (McAuslane et al., 2000; Eigenbrode et al., 2000; Eigenbrode, 2004) whose presence would be advantageous for the plant. Therefore, interactions on

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all trophic levels have to be considered in choosing the suitable crop cultivar for planting. Using pesticides can change the surface characteristics drastically, for example by affecting the fine structure of the epicuticular wax layer (Chapter 7) or wettability. This in turn influences the acceptability by herbivores. Treatment with a carbamate herbicide (S-ethyl dipropylthiocarbamate) of canola (Brassica napus L.) has been shown to increase oviposition by the diamond back moth (P. xylostella L., Lepidoptera: Plutellidae) (Justus et al., 2000). Stress hormones such as jasmonic acid and analogues are involved as signal transducers in plant defence against pathogens and herbivores. However, treatment of celery leaves (Apium graveolens cv. secalinum, Apiaceae) with these substances increases the acceptability by the carrot fly Psila rosae (F.) (Diptera: Psilidae). Due to the jasmonic acid treatment, furanocoumarins accumulate in the leaf surface, which in turn stimulate oviposition by the insect pest (Stanjek et al., 1997). Induced changes of the surface chemistry of crop plants can thus have strong impacts on suitability to herbivores. Also, changes in abiotic conditions can influence plant–insect interactions. Caution is advised when insight from studies of plants grown in the greenhouse or growth chambers is transferred to the field. The epicuticular lipid composition can vary considerably between both environmental conditions and impact herbivore responses to various degrees (Woodhead, 1981; Yang et al., 1993b).

13.7

Conclusion

Since the last thorough review by Eigenbrode and Espelie (1995) on effects of plant surface lipids on insects, much progress has been made. Main new findings have been reported in this chapter. Several more studies have elucidated the important role of leaf surface characteristics and even of particular lipid compounds for host acceptance and deterrence of herbivores. Elegant research approaches shed more light on influences of cuticle properties on the third trophic level, mainly by investigations of tarsal attachment, as well as on the underlying biomechanical characteristics. The same is true for knowledge about carnivorous plants. The development of highly selective probing methods for epicuticular waxes now allows the clear separation of surface cues from cues that are actually only active once the insect has damaged the leaves with its mouthparts or by scratching with its legs or ovipositor to get access to inner plant material. Still very little is known about visual effects of waxes on host plant selection by insects, probably due to experimental issues. Also, in the area of changes of surface characteristics due to induction an augmentation of knowledge can be expected, as in general, research on plant responses after induction increased within the last years. It would be desirable to link studies on different levels, plant physiology, molecular biology, biomechanics, behaviour and ecology, in order to get a broader picture of the impact of the plant cuticle in biotic interactions and even the ecosystem. For example, the physiological role of the plant cuticle, primarily to prevent water loss,

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might be linked with the preference behaviour of an insect (Müller and Riederer, 2005). The insect might be able to determine plant quality by leaf toughness, chemistry and transpiration properties acting together with some factors yet to be elucidated.

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Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd

Index

1,2-dihydro-3-thia-4,10,10b-triaza-cyclopenta[.a.]fluorene-1-carboxylic acid, 404, 410 1,5-alkanolide, 83 10,16-dihydroxyhexadecanoic acid, 38, 128, 133, 319 10-hydroxynonacosan-4-one, 154 10-hydroxynonacosan-5-one, 154 10-nonacosanol, 58, 64, 76–7, 79, 83, 94, 96–8, 100, 105, 107 12-hydroxydodecanoic acid, 100 14-hydroxynonacosan-15-one, 154 15-hydroxynonacosan-14-one, 154 15-nonacosanol, 99 18-hydroxy-9,10-epoxyoctadecanoic acid, 319 1-hexacosanol, 377, 401, 406 1-octacosanol, 71, 104 1-octanol/water partition coefficient, 257, 273 2,4-dichlorophenoxyacetic acid, 256, 259, 264, 295 4-hydroxycinnamic acid, 224–5 6-methyl salicylic acid methyl ester, 413 9,10,18-trihydroxyoctadecanoic acid, 319 9-hydroxydecanoic acid, 413 ABC transporter, 121, 199, 202–203, 205–206, 316, 321 Abies balsamea, 30 Abies concolor, 63 Abies sp., 54, 79 abrasion, 57, 84, 105, 138 absorbance, 217–19, 226, 228–9, 233, 237 absorptance, 218–20, 228, 234 absorption, 10, 92, 105, 201, 216–18, 220, 223–9, 231–2, 234, 236–8, 290 Abutilon striatum, 27 Abutilon venosum, 32, 92 Acacia collinsii, 414 Acacia pychantha, 67 Acacia senigal, 27 Acacia sp., 67, 69, 71, 414 Acanthaceae, 32 accelerator, 268–4, 295 Acer campestre, 306–308 Acer mono, 59, 62 Acer platanoides, 230 Acer pseudoplatanus, 41, 62, 125 Acer saccharum, 27 Acer sp., 27, 41, 59, 62, 75, 230, 306–308 Aceraceae, 59, 62

acetal, 160–161 acetate ester, 157 Acinetobacter sp., 345 Actinidia melanandra, 60, 93 activation energy, 264, 269, 271–2, 283–4, 302–303 active ingredient, 250, 269–70 acyl-ACP, 183–4, 186, 197, 201 acyl-CoA binding protein, 201, 204 acyltransferase, 127, 190 adaptation, 2, 339, 358 adenosine phosphate binding cassette, 102 adhesion pad, 373, 400 adhesion, 6, 313, 316, 322–3, 325, 342–4, 370–376, 379–80, 383–4, 388, 391, 399–400 Adiantaceae, 24, 70 Adiantum capillus-veneris, 24 Adiantum raddianum var. raddianum, 24 adjuvant, 251, 271, 295 Aegiceras corniculatum, 60, 75, 103 Aegicerataceae, 60 aerosol, 5, 251, 341 Aesculus hippocastanum, 41, 59 Agathis australis, 63, 81, 97 Agathis sp., 79 Agavaceae, 24, 26, 69 Agave americana, 17, 19, 20, 23–4, 34–6, 39, 41, 45, 47–9, 52–5, 91, 95, 127 Agave lutea, 24 Agropyron repens, 67 Agropyron sp., 195 Agrostemma githago, 322 air/water partition coefficient, 254 Aizoaceae, 61, 70 alarm pheromone, 413 alcohol ethoxylate, 257, 268–9, 271–2 aldehyde, 68, 71–2, 76–7, 86, 93, 98, 104, 128, 146, 148, 150–151, 160–162, 164, 166–7, 169, 170–171, 173, 182, 184, 187, 189–90, 192–4, 314, 319, 381, 386, 410 alga, 40, 46, 105, 107, 127 alkaloid, 159, 401 alkan-2-ol, 158, 182, 195–7 alkane, 36, 38, 68, 71–2, 76, 84, 88, 96, 98–9, 101, 150–152, 157, 162–4, 166–8, 171, 173, 182–4, 187, 190–195, 199–200, 202, 314, 319, 328–9, 377, 407–408, 410, 414

424 alkanediol, 64, 72, 79, 80–81, 83–4, 97, 100, 105, 148, 152, 154–5, 158, 162 alkyl benzoate, 158 alkyl ester, 72, 86, 88, 145, 148, 151, 157–8, 162, 164, 166, 183, 187 allelochemical, 252 Allium ampeloprasum, 402 Allium cepa, 24, 38–9, 65, 68 Allium cepa cv. ‘Ailsa Craig’, 65, 68 Allium porrum, 66, 72, 78, 98, 154, 170–171, 191 Allium sp., 71 Aloe arborescens, 30 Alstromeria aurantiaca, 68 Alstromeriaceae, 68 Alternaria sp., 357 Amaryllidaceae, 25, 27, 61 Amherstia nobilis, 69 amino acid, 183, 193, 198, 224, 229, 289, 319, 321, 326, 388, 402 Ammophila arenaria, 60 Amorphophallus maximus, 65 amorphous, 7, 12–13, 15, 17, 23, 41–4, 47–51, 53–5, 57–9, 70, 72, 75, 88–91, 98–9, 104–105, 107–109, 130, 138, 174–5, 268–9, 295, 382 Amphorophora idaei, 407 amyrin, 75, 156–7, 198, 401, 403 Anabasis articulata, 24, 27, 68 Anacardiaceae, 26, 60, 63–4, 70 Anarthia scabra, 66 Anarthriaceae, 66 Andrachne telephioides, 68 Andromeda polifolia, 65, 83 angiosperm, 38–9, 41, 56, 79, 88, 93, 105, 160 anisotropy, 95 Annonaceae, 61 ant, 75, 400–401, 405, 410, 414 Anthericum liliago, 68 anthocyanidin, 227 anthocyanin, 220, 224–5, 227–30 Anthurium warocqueanum, 236 antibiosis, 355, 358 antifeedant, 409 Antirrhinum majus, 158, 236 aphid, 400–401, 406–407, 413, 415 Aphis fabae, 406 Apiaceae, 27, 32, 59, 61–2, 68 Apium graveolens, 24, 49, 416 Apium graveolens cv.secalinum, 416 Apium sativum, 24 Apium sp., 47, 54, 416 appressorial germ tube, 369, 379–80 appressorium, 368–9, 379–91 Aquifoliaceae, 31 Aquilegia alpinum, 63, 79–80 Aquilegia formosa, 63 Aquilegia sp., 100 Aquilegia vulgaris, 63, 72, 97

INDEX Arabidopsis thaliana, 16, 24, 27, 31–2, 41, 44, 51, 59, 62, 72–3, 84, 102, 148–9, 151–2, 154–5, 160, 163, 171–2, 182–3, 185–90, 193–4, 199–203, 205, 207, 224, 312–28 arabitol, 350 Araceae, 27, 59–60, 65–6 Araliaceae, 25, 27, 60, 238, 407, 410 Ardisia crenata, 24, 27, 36, 94 Arecaceae, 61, 70 Arecacidae, 86 Argyroxiphium sandwicense, 234 Aristolochia duriar, 66 Aristolochia elegans, 66 Aristolochia gigantea, 60, 152, 154, 172 Aristolochia sp., 60, 66, 73, 86, 88, 99, 100–101, 152, 154, 172 Aristolochiaceae, 60, 66 Arrhenius plot, 264, 283–4, 302–304 arthropod, 6–7, 251–2 Arum maculatum, 27 Arundinaria sp., 61, 86, 93, 95 Asclepiadaceae, 70 ascorbate, 192 Asparagaceae, 68 Asparagus officinalis, 154 Asparagus retrofractus, 68 Aspergillus flavus, 375 Asphodelaceae, 68 Aspidiaceae, 28 Aspleniaceae, 29 Asteraceae, 25, 27, 59–60, 65–6, 405, 409, 411 asymmetrical secondary alcohol, 58, 63–4, 77, 79, 83, 96–7 asymmetry, 98, 259 Athanasia parviflora, 41 atmosphere, 2–3, 6, 161, 251, 267, 292, 294, 296, 301, 341, 348 atomic force microscopy, 89–90, 94, 102, 135, 137–9, 141, 348 Atrichum undulatum, 63 Atriplex hymenelytra, 235 Atriplex semibaccata, 31 Atta cephalotes, 404 Atta sexdens rubropilosa, 405, 407–408, 410 attachment, 11, 324, 342–3, 372, 383, 391, 398–401, 414, 416 Aureobasidium pullulans, 337, 346 Avena fatua, 24, 30, 32, 67 Avena sativa, 45, 67, 195, 406 Avena sativa cv. ‘Black supreme’, 67 Avena sp., 64, 67 Bacillus pumilus, 345 bacteria, 4, 6, 126, 206, 251, 289, 334–51, 353–8 Baptisia australis, 67 Barbacenia tubulosa, 66 barrier, 3–4, 7, 11, 36, 38, 50–51, 75, 126, 168, 250, 254–9, 260, 262, 264, 267–8, 270–273, 280–283,

INDEX 292–3, 295, 299–300, 302–303, 306–309, 316, 319, 326, 355, 368, 390, 400, 415 baurenol, 198 Begonia pavonina, 237 behaviour, 6, 96, 135, 137–8, 141, 216–17, 230, 237–8, 252, 339, 351, 374, 398–401, 414, 416–17 Benincasa hispida, 61, 86, 88 Benthamia alyxifolia, 69 benzyl alcohol, 158 Berberidaceae, 63 Berberis sp., 63 Bergenia purpurascens, 27 Berula erecta, 27, 32, 51 Beta vulgaris, 17, 24, 31, 39, 43, 48, 52, 54, 57, 59, 88 betacyanin, 225, 229 betalain, 224–5, 229 betalamic acid, 229 betaxanthin, 229 Betula pubescens, 412 betulin, 156 Bignoniaceae, 30 bioavailability, 353–4 biocontrol, 342, 349, 355–8 biofilm, 46, 105, 107, 342–3, 351 biological pest management, 415 biopolyester, 126 bioreporter, 339–40, 348, 354, 356 biosynthesis, 8, 127, 141, 154–6, 166, 168, 182–3, 185–9, 191–2, 194–6, 198–200, 202, 207, 318–20, 325, 327–8, 339 biosynthetic pathway, 56, 128, 166, 182, 184, 189–90, 193, 196–7 bis-N, O-trimethylsilyltrifluoroacetamide, 148 Blechnaceae, 59 Blechnum capense, 59 bloom, 56, 232, 234, 399–401, 405 Blumeria graminis, 368–71, 373–81, 383–7, 390 Blumeria graminis f. sp. hordei, 375 Bobartia gracilis, 24 Boraginaceae, 59, 65 Brassica napus, 32, 43, 61–2, 68, 70, 99, 151, 154, 187, 190, 319, 322, 404–406, 410, 416 Brassica napus mutant nilla, 68 Brassica napus mutant Rigo, 61 Brassica oleracea, 17, 30, 32, 61–2, 66, 68, 80, 86–8, 93, 99, 101–102, 151–2, 154–5, 164, 167–8, 171–2, 182, 187, 189–91, 193, 199, 403–404, 407–408, 415 Brassica oleracea mutant gl2, 61 Brassica oleracea mutant gl4, 68 Brassica oleracea var. botrytis, 30, 32 Brassica oleracea var. capitata, 62, 66 Brassica oleracea var.caulo-rapa, 66 Brassica oleracea var. gemmifera, 32, 62, 66, 80 Brassica pekinensis, 409–410 Brassica sp., 42, 70–72, 78, 84, 96–8, 105, 158, 232, 325, 403–410, 415–16 Brassicaceae, 24, 27, 30–32, 59, 61–3, 66, 68, 70, 88, 164, 325, 403–410, 415

425 Brevicoryne brassicae, 400 Brocchinia reducta, 399 Bromeliaceae, 234, 399 Bromus interruptus, 59, 67 bryophyte, 40, 54, 60 Bupleurum ranunculoides, 68, 86, 103 Bupleurum salicifolium, 62 Bupleurum sp., 93 Burmannia biflora, 68 Burmanniaceae, 68 Butomaceae, 61 Butomus umbellatus, 61 butryrospermol, 198 Buxaceae, 24, 65, 66 Buxus sempervirens, 24, 65, 72, 83, 84, 98, 152, 155 Buxus sp., 79 Cactaceae, 70, 164 Cajanus cajan, 403, 407 Cajanus platycarpus, 407 Cajanus scarabaeoides, 407 calcium, 16, 109, 283–5, 295, 308, 380, 387 Calibanthus hookeri, 68 Calliandra haematoma, 69, 103 Calligonum comosum, 67 Callitris columnaris, 63 callose, 323–4 Caltha palustris, 30 Camassia cusickii, 68 cAMP, 381–2, 387 Camponotus femoratus, 413 Campylobacter sp., 357 Cannabis sativa, 27, 30, 32 Caprifoliaceae, 33, 65 Capsicum annuum, 30, 256, 297–8 carbohydrate, 224, 280, 289, 295, 383, 402 carbon dioxide, 3–4, 347 carbon source, 320, 351–3 carboxylase, 185, 316, 320 Carex flacca, 61 Carludovica palmata, 66 carnauba, 57, 86 carnivorous plant, 169, 398–9, 416 carotenoid, 224–5, 229–30 Carya illinoensis, 232 Caryophyllaceae, 26–7, 30, 60, 65 Cassida denticollis, 411 Cassida stigmatica, 405, 411 Cassytha pubescens, 23 Catopsis berteroniana, 399 cell wall, 5, 11, 13–19, 22–3, 41, 43–6, 48–54, 57, 91–2, 94–5, 102, 107–109, 127, 202, 206–207, 222, 224–7, 236–8, 251, 254, 266, 280–281, 287, 289, 292, 314–16, 322, 343, 346, 357, 368, 370, 372, 375–6, 379–80, 388–90 cellobiohydrolase, 381 cellulase, 16, 43, 357 cellulose acetate, 173, 405–406, 409, 411, 415 cellulose, 13–14, 16, 43, 48, 280, 295, 343, 381–2, 386

426 Centranthus ruber, 60, 62, 84 Centranthus sp., 88, 98 Cercis canadensis, 403 Cerinthe major purpurascens, 76, 84 Cerinthe major, 47, 65, 98 Cerinthe minor, 65, 72, 83, 98, 158 Ceroxylon andicola, 70 Ceroxylon sp., 57 chain length distribution, 151, 162–3, 171, 174 chain length specificity, 154, 183, 185 chalcone, 224 Chamaecyparis lawsoniana, 63, 81, 97 Chamaecyparis obtusa, 63 Chamaecyparis sp., 79 Chamaegigas intrepidus, 27 Chamerion angustifolium, 59, 67, 76, 86, 95 Cheilanthus sp., 70 Chelidonium majus, 63, 79, 80, 97 Chenopodiaceae, 24, 27, 30–31, 59–60, 67–8 Chenopodium album, 59, 67–8, 103 Chionochloa rigida, 59 Chlorococcus sp., 105 chlorogenic acid, 402, 410 chlorophyll fluorescence, 220 chlorophyll, 147, 220, 229, 231, 313–14, 317, 320, 340 Chlorophyta, 40 Chondrus crispus, 40 Chrysanthemum frutescens, 65 Chrysanthemum morifolium, 59 Chrysanthemum segetum, 65, 84, 98 Chrysomela scripta, 402, 404 Chrysoperla carnea, 400 Chrysophtharta bimaculata, 403 Cicer arietinum, 33 cinnamyl alcohol, 158 Cirsium arvense, 156, 411 Cirsium horridulum, 13, 295 Citrus aurantifolia, 130–134, 137–8, 141 Citrus aurantium, 174, 256, 265, 282–3, 296–7, 299–300 Citrus jambhiri, 325 Citrus limon, 12, 17, 27, 51, 59–60, 72, 86, 98 Citrus limon cv. ‘Adamopolou’, 59, 60 Citrus paradisi, 30, 33, 39, 60–61, 156 Citrus paradisi cv. ‘Marsh seedless’, 60–61 Citrus sinensis, 30 Citrus sp., 16–17, 38, 326 Clarkia elegans, 62, 72, 80, 84, 151, 154 Clarkia sp., 98 Clavibacter michiganensis, 343 Clematis vitalba, 59, 88 Clivia miniata, 17, 21–2, 25, 34, 37–9, 41, 43, 45, 47–9, 54, 91 Clivia nobilis, 25 cloud, 57, 251 Clusiaceae, 66 CO, 192 Cochliobolus, 357 co-crystallisation, 98–101

INDEX Coffea arabica, 30, 59 coil, 79, 83–4, 97–8 Colchicaceae, 69 Coleochaete orbicularis, 40 Colletia cruciata, 61 Colletotrichum coccodes, 384 Colletotrichum dematium, 384 Colletotrichum graminicola, 370, 372, 383–4, 390 Colletotrichum lagenarium, 384 Colletotrichum trifolii, 387 Colletotrichum sp., 368, 379, 384–5, 387, 390 Colochasia esculenta , 66 colourless flavonoid, 219, 224, 226–7 Columellia oblonga, 65 Columelliaceae, 65 Combretaceae, 70 Commelinaceae, 31 Commelinidae, 86 communication, 312, 326, 329, 349, 355 compartmentation, 192, 202 competition, 75, 349, 352, 355 condensation reaction, 185, 191, 197 conductance, 293–4, 304, 306–309 confocal laser scanning microscopy, 221, 340 conifer, 16, 43, 98, 100, 102, 104–105, 107, 223, 226–7, 230, 233, 237 conjugated double bonds, 223, 229 connectivity index, 257 Conocephalum conicum, 60 Convallaria majalis, 68, 103–104 Convallaria sp., 68, 73, 86, 103–104 Convallariaceae, 68 convergence, 1, 6, 7, 88 Convolvulaceae, 25, 27 Convolvulus arvensis, 14 Copernica cerifera, 57, 70, 72, 86, 95 Copernica cowellii, 70, 86 Copernica hospita, 70 Copiapoa cinerea, 70 Coreus marginatus, 400 cork, 127, 204 Cornaceae, 59 Corydalis cava, 65 Cotinus coggygria, 63 Cotyledon orbiculata, 233 coumaric acid, 128, 224 Crambe maritima, 62, 88 Crassulaceae, 65, 164, 233 Crataegus prunifolia, 60 Cretaceous, 38, 44 Crocus chrysanthus, 25, 92 crop, 55, 126, 141, 373, 401, 415–16 cross polarisation-magic angle spinning spectrometry, 130, 138 cross-link, 15, 20, 38, 50, 109, 129–31, 134, 138–9, 141 cryo-adhesive, 89, 169, 173, 415 Cryptococcus sp., 346 Cryptogrammaceae, 70 Cryptomeria japonica, 30, 230

427

INDEX crystal formation, 96, 100, 171 crystal morphology, 58, 72, 84, 88, 96, 98–100, 104–105, 107, 110 crystal, 5, 19, 36, 38, 41, 45–6, 57–8, 71–5, 77–9, 82, 84, 86–91, 93–105, 107, 109–110, 163–5, 168–72, 174–5, 194, 200, 207, 231, 235, 252, 262, 268, 280, 283, 285, 295–6, 300, 315, 348, 370–371, 376–8, 381–2, 386, 399, 403, 405 crystalline domain, 163, 175, 262 crystallinity, 75, 174 crystallisation, 74, 78, 77, 82, 87, 95–101, 104, 169, 171–2 crystallite, 84, 88–9, 95–6, 99, 101–102, 109–110 crystalloid, 100, 399 Cucurbitaceae, 61, 68 Cunninghamia lanceolata, 25 Cunninghamia sp., 54 Cupressaceae, 63 Cuscuta campestris, 25 Cuscuta gronovii, 17, 27, 41 Cuscuta odorata, 27 cutan, 39–40, 47–8, 50, 55–6, 107, 109, 127, 134 cuticle proper, 12–23, 34–45, 47–55, 57, 88–92, 94–5, 104, 107–110, 315–16, 320, 328, 344, 386, 398, 416 cuticle/air partition coefficient, 254 cuticle/PEG partition coefficient, 266 cuticle/water partition coefficient, 254–7, 266, 280, 296–8 cuticula, 1, 13 cuticular layer, 12, 14–15, 18–20, 22–3, 34–7, 39, 41–5, 47–55, 57, 91–2, 94, 108–109, 207, 398 cuticular membrane, 11–16, 18–22, 34–7, 39–42, 44–5, 47–55, 57, 89, 91–5, 101, 104, 108, 126, 135, 137, 146, 173, 256, 259–62, 265, 267, 282, 287, 292–305, 307, 314–15, 319 cuticular pore, 91–3 cuticular transpiration, 292–4, 299–300, 301–303, 306–309 cuticular transport, 3, 250, 258–9, 259–62, 264, 267, 280–282, 284, 287, 294 cuticular wax, 5, 11, 13, 19, 36, 40–41, 45–6, 56–9, 71–4, 76–7, 79, 82, 84, 87–90, 92–6, 100–102, 104–105, 107–110, 141, 145–75, 182–207, 223, 225, 231, 233, 238, 251–2, 254, 257–9, 261–5, 267–70, 273, 280, 283, 295, 299–300, 302–303, 315, 320–321, 328, 343, 370–371, 374, 376, 377–8, 381–2, 386, 388, 398–401, 410–416 cutin oligomer, 129, 131–4, 137 cutin, 1, 5–6, 14, 16, 21, 36–40, 43–5, 47–9, 56, 91, 94, 104, 107–109, 126–7, 129–34, 137–41, 145, 165, 173, 185, 200, 202, 205, 207, 222, 257, 258, 273, 283, 285–6, 289, 295, 303, 307, 316, 320–322, 326, 330, 353, 377, 381, 386, 387, 390 cutinase, 31, 44, 51, 91, 314, 320, 322, 343, 357, 372–3, 375, 377, 379, 381 cutinisation, 109 Cyathodes colensoi, 61 Cyclanthaceae, 66

cyclisation, 161, 196, 198 Cydia pomonella, 405 Cynanchum sarcostemma, 70 Cyperaceae, 59, 61 cystolith, 37, 43, 49, 109 Cytophagales, 345 cytoplasm, 102, 183, 201–202, 204, 313–15, 321, 343, 369, 375–6, 388–90 Dactylis glomerata, 67 Daphne tangutica, 61 Dasylirion sp., 55 Daucus carota, 402 Dawsonia beccarii, 63 Dawsonia sp., 56, 63, 66, 72 Dawsonia superba, 66 decarbonylase, 187, 192–4, 196, 328–9 decarboxylation, 191, 196 degradation, 104–105, 107, 129, 131, 134–6, 141, 322–3, 389 degree of succulence, 307 dehydratase, 183, 186–8, 207 Delarbrea michieana, 238 Delia floralis, 403 Delia radicum, 404–405 deliquescence, 252 Dendrocalamus giganteus, 61 depolymerisation, 37–8 deposit, 15, 47, 109, 235, 251–4, 262, 288, 348, 372, 374 deposition, 1, 43, 47, 109, 173, 199, 207, 251–2, 266, 295, 341, 368, 370, 372–4, 377–8, 380, 384–5, 387 derivatisation, 148 desaturase, 193, 316–17, 319 desorption, 201, 259–63, 272, 280 development, 2, 5, 12, 14–16, 18, 23, 34, 38–9, 41, 43, 45–50, 52–4, 101–102, 107–109, 126, 146, 161, 166–7, 173–4, 182, 185, 188, 199–200, 261, 300, 305, 312, 315, 327, 329, 355, 368, 377, 382, 385, 387–92, 416 Devonian, 1–2 Dianellaceae, 69 Dianthus caryophyllus, 27, 65, 83, 97 Dianthus sp., 195 Didymopanax vinosum, 407, 410 Dieffenbachia maculata, 41, 59 differential scanning calorimetry, 140–141, 174 differentiation, 91, 300, 304, 326–9, 339, 380–391 diffuse reflectance, 230 diffusion coefficient, 258, 261–5, 268, 270, 280, 295–6, 298 diffusion, 3, 36, 91, 135, 174, 201, 231, 250, 258–72, 280–283, 285–7, 289, 295–8, 300–302, 307 β-diketone, 64, 71–2, 74–6, 79, 83–4, 93, 95–8, 100–101, 103, 195–7 diol, 162 Dionaea muscipula, 25, 27–8 Dioncophyllaceae, 66

428 Dioscoreaceae, 60 diphenylboric acid 2-aminoethylester, 223 dissolution-diffusion, 250 diterpenoid, 75, 159 Diuraphis noxia, 406 Doryanthaceae, 69 double bond, 127, 134, 157, 160, 196, 198, 223–4, 229, 270 Dracaenaceae, 68 driving force, 3, 5, 266, 292–4, 301–302, 307 droplet, 5, 13, 92, 105, 169, 235–6, 251–3, 261, 266, 269, 354, 371, 373 Drosera burmanni, 62 Drosera sp., 30, 92 Droseraceae, 25, 27–8, 30, 62 Drosophila melanogaster, 205 Drosophyllum lusitanica, 28, 92 drought avoidance, 292, 306, 308 drought resistance, 304 Dryopteris filix mas, 28 Dudleya brittonii, 170, 233 dynamics of wax composition, 165

Eccremocarpus scaber, 30 eceriferum, 188, 313, 315–16, 321, 324, 386, 388 Echeveria sp., 233 ecology, 7, 54, 145, 163, 165, 199, 206–207, 342, 412 ecotoxicology, 266 Elaeocarpus sp., 238 electron diffraction, 93 electron spin resonance, 268 electron-dense, 15, 17–23, 35, 39, 43–5, 47, 50–51, 53, 91, 94, 107–109, 314, 322, 325–6 electron-lucent, 12, 14, 17, 19–23, 35, 39–40, 42–5, 50–51, 53, 91, 107–109 Elegia verticciaris, 66 Elodea canadensis, 32, 43 Elodea sp., 53 elongase, 183–5, 188, 196, 202–203, 300, 316, 318, 325 elongation, 158, 182–6, 188, 191, 195–6, 201–203, 318–20, 322, 380–381, 385, 388 elongation-decarboxylation, 191 Elytrigia sp., 71 embryogenesis, 322, 325–6 embryophyte, 40 emigration, 344, 347, 357 Empoasca fabae, 412 Encelica californica, 234 Encelia farinosa, 234 Encephalartos sp., 154 endoplasmic reticulum, 15, 17, 22, 35, 43–4, 47–8, 51–2, 54, 91–2, 94, 108, 183, 203, 300 enhancer, 267 Enterobacter agglomerans, 345 environment, 3–5, 11, 23, 54, 99, 102, 104, 107, 126, 137, 141, 145, 160, 165–8, 199, 206–207, 235, 250–252, 256, 264–5, 268, 273, 283, 289, 292,

INDEX 300–302, 304, 306–307, 309, 312, 329, 334, 338, 340, 342, 345–6, 349–51, 354–6, 358, 368, 370, 372, 376–7, 384–5, 388, 390, 416 Eocene, 38 Epacridaceae, 61 Epicoccum nigrum, 350 epicuticle, 6, 103 epidermis, 1, 5–7, 11–13, 16–17, 19, 21, 23, 28, 31, 33–4, 41, 44–6, 49–50, 55, 73, 89, 92–3, 95, 102–104, 107, 127, 183, 185, 188, 190–191, 193–4, 199–206, 216–21, 223, 225–38, 251, 280–281, 287, 289, 292, 305, 312, 320, 326–9, 340, 346, 348, 356, 368–70, 374–6, 378–80, 386, 389, 391, 399 epiphitness, 349–52, 355 epitaxy, 104, 110 epoxidation, 127 Equisetaceae, 66 Equisetum arevense, 66 Eragrostis curvula, 65 Ericaceae, 30, 59, 65 Eriospermaceae, 69 Eriospermum triphyllum, 69 Erwinia ananas, 356 Erwinia herbicola, 339, 348, 352, 354, 356, 358 Erwinia sp., 344–5, 356 Eryngium rostratum, 25, 41 Eryngium sp., 45, 61, 86 Erysimum cheiri, 230 Erysiphe pisi, 368–9, 375, 377–9, 385 erythrodiol, 156 Erythronium rostratum, 63 Erythroxylaceae, 69 Erythroxylum coca, 69, 103 Escherichia coli, 187, 190 Espelatia schulzii, 234 ester cutin, 21, 37–9, 43, 47 esterase, 322, 343, 373 esterification, 38, 136 estolide, 71–2, 93, 100, 159 ether bond, 129 Eucalyptus camaldulensis, 65, 69, 73, 103 Eucalyptus cephalocarpa, 97 Eucalyptus cinerea, 31, 34, 53, 94 Eucalyptus cloeziana, 67 Eucalyptus coccifera, 67 Eucalyptus glaucescens, 65, 97 Eucalyptus globulus, 58, 65, 72, 103, 105, 170, 408 Eucalyptus gunnii, 65 Eucalyptus maculata, 405 Eucalyptus microcarpa, 67 Eucalyptus miniata, 67 Eucalyptus nova-anglica, 65 Eucalyptus ovata, 96 Eucalyptus pauciflora, 67, 69 Eucalyptus papuana, 25, 27, 95 Eucalyptus perriniana, 25, 28, 65 Eucalyptus platypus, 67, 70 Eucalyptus regnans, 403

INDEX Eucalyptus sp., 64, 67, 69, 71, 75, 77, 79, 83, 94, 96, 98, 104, 152, 154, 164, 195, 232–3, 405, 408 Eucalyptus umbrawarrensis, 79 Eucalyptus viminalis, 65 Euglena gracilis, 189–90 Euphorbia cerifera, 59, 70 Euphorbia characias, 68 Euphorbia cyparissias, 156 Euphorbia lathyris, 66, 89, 155 Euphorbia milii, 68 Euphorbia peplus, 66, 68 Euphorbia serrata, 68 Euphorbia sp., 71, 89, 155–6 Euphorbiaceae, 33, 59, 60, 62, 66, 68, 70, 164, 400, 405 Euphydryas chalcedona, 403 evaporation, 96, 99, 101–102, 262, 266, 300, 348–9, 411, 415 evolution, 1–4, 7, 56, 238, 357, 385 Exochorda racemosa, 63, 77, 79–80, 97 Exomotheca bullata, 60 external cuticular layer, 12, 14–15, 20, 22, 37, 45, 47–8, 51–5, 91, 108–109 extracellular material, 371–80, 383, 386, 388–9, 391 extracellular polysaccharide, 238, 343, 351, 354 extrusion, 93, 95, 102, 169

Fabaceae, 24, 26–31, 33, 59–62, 66–71, 76, 164, 228, 403, 406–407, 414 Fagaceae, 28–9, 59, 67–9 Fagus sylvatica, 28, 59, 68, 72, 104, 163, 174, 306–308 farina, 88 fatty acid ester, 157 fatty acid synthase complex, 183–5, 197 fatty acid, 21, 23, 38, 71–2, 76, 88, 126, 131, 148, 150, 157–60, 162–4, 166, 171, 173, 182–3, 185–6, 188–92, 194, 197, 201–202, 207, 270–271, 300, 316–20, 325, 387, 401 fern, 40, 44, 55, 88, 155, 160, 237 fernane, 155 fertilisation, 289, 312, 322, 324, 329 ferulic acid, 128, 225 Festuca arundinacea, 67, 73, 103 Festuca glauca, 65, 97 Festuca sp., 195 fibre optics, 218, 233, 237 Ficus elastica, 14, 17, 25, 27, 48, 54, 296 Ficus lyrata, 28 Ficus sp., 55 filament, 58, 61–2, 75–6, 78–9, 97, 204 First Fick’s law, 258–9, 292 flame ionisation detector, 147 flavon-3,4,-diol, 227 flavonoid, 88, 160, 219, 223–8, 232, 236, 401–403, 410, 412 flavonol, 224–7, 406 flow cytometry, 337 flower, 2, 151, 158, 161, 216, 226–30, 232, 236, 399, 413

429 fluorescence microscopy, 44, 207, 220–221, 227, 340–341, 348 fluorescent in situ hybridization, 337, 341 fluorimetry, 220–221 fog, 251, 348, 353 formulation, 250–252, 266–7, 269, 273 fossil, 11, 38, 44, 56 Fourier transform infrared spectroscopy, 130, 161, 174 Fragaria ananassa, 28, 32, 62 Fragaria ananassa cv. ‘Cambridge Favourite’, 62 Fragaria ovalis, 62 Fragaria sp., 75 freezing method, 336 friedelin, 75, 156 Fritillaria meleagris, 60 Fritillaria pallidiflora, 60, 88 frost, 140, 356, 358 fructose, 339, 353–4, 405 Fumariaceae, 65 Funaria hygrometrica, 19, 25, 56, 91 Funaria sp., 40, 51 Funariales, 25 fungi, 4, 6, 91, 105, 107, 126–7, 129, 251, 289, 314, 320, 322, 334–6, 338, 340, 342–7, 349, 356–8, 368–92, 415 fungicide, 271, 392 furanocoumarin, 416 Galanthus nivalis, 61, 89–90, 102 Galanthus sp., 88 Galeruca tanaceti, 409–410 Galium aparine, 30 Galphimia brasiliensis, 28, 92 Galtonia candicans, 61 gametophyte, 11, 19, 26, 32, 40, 56, 70, 75, 98 gas chromatography, 129, 147–9, 151, 157, 194, 315 Gasteria planifolia, 25, 34 Gasteria verrucosa, 25 Gaultheria depressa, 59 gene exchange, 356 Genista aetnensis, 30 Geraniaceae, 64 Gerbera jamesonii, 59 germanicol, 198 germling, 368, 377, 379–84, 389, 391 Ginkgo biloba, 63, 75, 79, 81, 83, 97, 301 Ginkgoaceae, 63 gland, 3, 11, 33, 92, 159, 223, 348, 354, 406, 413 glass transition temperature, 140 glaucous, 56, 75, 79, 232–4, 315, 318, 399, 403 globule, 22–3, 43, 49, 51, 107–109, 204 glossy, 56, 75, 188–9, 194, 199–200, 314–16, 319, 328, 399, 403–404, 408, 415 glucose, 225, 353 glucosinolate, 403, 405–408, 410 glutinane, 156 glycerol, 129, 159, 350, 389, 415 glycoprotein, 343, 372, 383, 388 glycosylated flavonoid, 226

430 glyphosate, 281, 283 Gnetaceae, 59 Gnetum gnemon, 59 Golgi, 202, 204–205 Gordonia axilaris, 25, 28, 31 Gossypium hirsutum, 13, 24–5, 28, 66, 89, 190 Gossypium hirsutum cv. Green Lint, 28 grain, 55, 58, 322, 325 granule, 49, 58, 60, 75, 103, 205 green fluorescent protein, 201–203, 205, 321, 337, 339–41, 348, 354, 356 Grevillea bipinnatifida, 68 Griselinia littoralis, 59 guard cell, 19, 33, 43–4, 46, 50, 55, 91, 104, 107, 222, 229, 305, 388, 391 gymnosperm, 38–9, 56, 71–2, 79, 93, 100, 127, 151–2, 154, 159, 161, 164, 230, Gypsophila acutifolia, 60, 171

Habropetalum dawei, 66 Hakea leucoptera, 25, 31 Hakea suaveolens, 23, 25, 94 Haloragaceae, 24, 59 Hedera helix, 17, 25, 27, 34, 37, 41, 45, 54, 60, 166, 174, 263–4, 296, 299–301, 303–308 Hedychium flavum, 61 Helianthus annuus, 60 Heliconia collinsiana, 61 Heliconia densiflora, 59 Heliconia sp., 86, 93, 103 Heliconiaceae, 59, 61 Helicoverpa armigera, 403, 407, 410 Hemerocallidaceae, 69 Hemerocallis citrina, 69 hemiacetal, 160 hemi-biotrophic fungi, 368, 376 Hemisphaerota cyanea, 400 hentriacontan-16-one, 88, 98, 170–172 hentriacontan-14,16-dione, 71–2, 79, 155, 170 herbicide, 57, 404, 416 Herpolirion novae-zelandiae, 69 Herreria montevidensis, 69 Herreriaceae, 69 hexacosanal, 72, 381 hexadecanoate, 319–20 hindrance factor, 298 Hippocastanaceae, 59 homoiohydry, 2, 11, 55 homologous series, 96, 100, 150–152, 157–8, 161–3 honeydew, 353 Hordeum chilense, 388 Hordeum sativum, 67 Hordeum vulgare, 32, 65, 67, 74, 84, 97, 105, 155, 158, 160, 164, 168, 170, 182, 195, 226, 406 Hordeum vulgare cv. ‘Proctor’, 67 hornwort, 11, 40 host, 6, 368, 370–391, 400–01, 406, 410–412, 415–16 Hosta sieboldiana, 65

INDEX Hostaceae, 65 human skin, 270 humidity, 66, 137, 138–9, 167, 251, 262, 281–4, 287, 292–4, 299, 301–302, 306–307, 317, 326, 338, 342, 347, 368, 373 Humulus lupulus, 25 Humulus sp., 54 Hyacinthaceae, 61, 68 Hyacinthoides non-scripta, 39 Hyacinthus orientalis, 89 hydathode, 3, 33, 235, 251 Hydrangea hortensis, 59 Hydrangea macrophylla, 17, 28, 51 Hydrangeaceae, 59 hydration, 137–9, 141, 281, 302, 304, 316–18, 322–5, 329, 372–4 hydrocarbon, 36, 38–9, 71, 96, 98, 101–102, 146, 150, 155, 157–8, 160, 174, 412–14 Hydrocharitaceae, 32 Hydrocotyle bonariensis, 59 hydrophilic pathway, 295, 297, 300, 308 hydrophilic, 51, 102, 269, 272, 295, 297–8, 300, 302, 308, 342, 371–2, 374, 377, 379, 381, 383–6 hydrophobic, 13, 21, 23, 38, 40, 146, 204, 207, 280, 302, 315, 342–4, 370–374, 377, 379, 381–5 hydrophobicity, 56, 342–3, 371, 379, 381–2, 384, 386, 388 hydrophobin, 383, 387 hydroxy kaurane, 75 hydroxyaldehyde, 158 hydroxycinnamic acid, 223–6, 228–9 hydroxyphenyl butanol, 158 hydroxyphenyl propanol, 158 hydroxy-β-diketone, 72, 75, 79, 83, 93, 97, 100, 152, 195–6 Hygrophila corymbosa, 32 hygroscopic, 252, 348 Hypericaceae, 69 Hypericum bucklei, 69, 103 Hypericum calycinum, 232 Hypericum hircinum, 66 hypha, 91, 105, 107, 251, 336, 343, 379, 389, 391 ice nucleation, 339, 356, 358 Ilex aquifolium, 31, 306–308 Ilex integra, 14, 31, 48, 51 immigration of phyllosphere colonisers, 341–2, 344, 347 immunogold, 16, 375–6 indole-3-acetic acid, 351, 353–4, 357 infection structure, 368, 387 infochemical, 410 infrared, 129–30, 142, 161 inner sorption compartment, 257, 260–262 inorganic ion, 281–3, 285–7, 289 insect cuticle, 413 insect, 6, 11, 105, 169, 199, 227, 229, 232, 236, 341–2, 347, 398–417 integrating sphere, 217–19, 231

INDEX integument, 1, 3, 6–7, 326 intercellular space, 11, 13, 50, 147 internal cuticular layer, 12, 14–16, 20, 22, 34, 37, 45, 47, 49, 51–5, 108 intracuticular wax, , 36, 57, 89, 147, 159, 165, 168–9, 171, 173–5, 295, 300, 381 iodotrimethylsilane, 132, 134–5 ion permeability, 281, 284 ion, 3, 4, 16, 149, 160, 192, 227, 250, 259, 281–3, 285–7, 289, 354, 380, 389 ionic organic compound, 281 Ipheion uniflorum, 89 Iridaceae, 24–6, 31, 69 iridoplasts, 238 Iris germanica, 17, 20, 25, 39, 52, 54 Iris sp., 34 Isoetales, 41 isolated cuticle, 13, 37, 223–5, 269–72, 281, 283, 289 isomer, 38, 80, 99, 100, 148–52, 154, 155, 157, 193, 198 isopropyl amide, 270 jasmonic acid, 416 Juglans regia, 174, 301, 303–304, 306, 308 Juncus inflexus, 59 juniperic acid, 100 Kalanchoe pumila, 233 Kensitia pillansii, 66 ketoacyl elongation, 182, 195–6 ketoalcohol, 158 ketol, 152, 154–5, 162 Kleinia articulata, 66 KMnO4 , 15, 19, 34–7, 39, 43–5, 56, 94 Krystalloid, 95 Laburnum sp., 67, 69 lactone, 65, 72, 79, 83, 98, 158, 351, 355 Lactuca sativa, 25, 27, 57, 59, 228 lamella, 12, 14–23, 33–40, 43–5, 48–54, 56, 91, 94–5, 107–109 lamellar, 23, 34, 36, 38–40, 49, 94 lamellate, 12, 17–21, 23, 34–5, 37–9, 43–5, 48–9, 51–4, 58, 91, 107–109 lamellation, 48, 50 Lamiaceae, 32, 160 land plant, 2, 11, 40, 54–6, 109 Lauraceae, 60–61 Laurus nobilis, 60 Laurus sp., 88 leachate, 353, 402 leaching, 3, 4, 289, 313–14, 317, 320, 347, 353–4 leaf blight, 356 leaf expansion, 45 leaf printing, 335 leaf washing, 335–7, 344 lectin, 383

431 Lecythidaceae, 66 Lecythis chartacea, 66 Lemna minor, 32, 42, 53 Lemnaceae, 32 Lentibulariaceae, 24, 27, 31, 33 Leptinotarsa decemlineata, 401–402, 408, 410 Leptospermum laevigatum, 62 Leucojum aestivum, 60, 88 Leymus arenarius, 65 Libertia elegans, 14, 25, 47 Libertia ixioides, 25 life-style, 54, 359 light microscope, 56–7, 93, 259, 288, 378 light, 4–5, 23, 56, 79, 167–8, 199, 216, 220, 228–31, 233–7, 259, 283–5, 338–40, 368, 378–9, 385 lignin, 16, 47, 109, 127, 223 Ligustrum ovalifolium, 28 Ligustrum vulgare, 226 Liliaceae, 24–5, 28, 30, 33, 60, 63–6, 68 Lilium candidum, 28 limiting skin, 259–62, 265–7 lipase, 132–3, 135, 389 lipid binding protein, 202, 204 lipid raft, 202, 204 lipid transfer protein, 202, 205 lipophilic non-electrolyte, 250–273, 284 lipophilic organic compound, 4, 295, 297, 307 lipophilic pathway, 295, 297 lipophilicity, 256–7, 265, 271–3 Liriodendron sp., 60, 88 Liriodendron tulipifera, 26, 28, 38, 172, 301, 303 liverwort, 11, 40, 56 Lobelia dortmanna, 28, 43, 51 Lobeliaceae, 28 Lobesia botrana, 405 locomotion, 399, 401 Lolium perenne, 345 Lolium sp., 382 Lonicera japonica, 33 Lonicera korolkovii, 65, 83, 98 Loranthaceae, 69 Lotus effect, 5, 56, 105 Lunularia cruciata, 60 lupeol, 157, 170, 198, 407 Lupinus albus, 66, 70 Lupinus luteus, 66 Lycopersicon esculentum, 5, 11, 17, 31, 52, 59, 127, 130–131, 134, 137–41, 155, 165, 173, 227, 255–6, 296–301, 343, 357, 383 Lycopersicon hirsutum, 402 lycopod, 40 Macaranga hypoleuca, 62 Macaranga lamellata, 62 Macaranga tanarius, 170 Macaranga sp., 75, 156, 400, 405 Macaranga triloba, 60 Magnaporthe grisea, 91, 356, 368–70, 372, 374, 379, 383–5, 387, 389–90

432 Magnolia grandiflora, 31, 38, 59 Magnolia sp., 60, 88 Magnoliaceae, 26, 28, 31, 59–60 Magnoliidae, 88 male sterile, 324 Malephora spp., 70, 86 malonyl-ACP, 183, 185 Malpighiaceae, 28 Malus domestica, 283, 405 Malus hupehensis, 59 Malus pumila, 12–13, 26–28, 31–2, 34, 53, 127, 129, 227–8, 230 Malus sp., 27–8, 33, 60, 72 Malvaceae, 24–5, 27–8, 32, 66 Maranta leuconeura, 61 Marantaceae, 61 Marchantia paleacea, 60 Marchantia polymorpha, 19, 60 Marchantia sp., 32, 40, 56 Marchantiales, 32 mass spectrometer, 47, 132–3, 141, 147–9, 151, 157 mass transfer, 250, 258–60 Mayetiola destructor, 400, 402, 404–405 mechanical property, 5, 135, 137–8 mechanical, 5–6, 36, 57, 79, 89, 92, 100–101, 104–105, 135, 137–8, 141, 169, 173, 217, 232, 400, 410, 412, 415–16 Medicago sativa, 200 Medicago truncatula, 200 Megachile sidalceae, 403 Melaleuca hypericifolia, 69 melanin, 389 Melanthiaceae, 69 membrane thickness, 129, 296, 298 Mentha aquatica, 32, 51 meristem, 312, 327–8 Mesembryanthemaceae, 62, 66, 69 Mesembryanthemum crystallinum, 229 mesomorphic leaf, 307 mesophyll, 11, 13, 21, 40, 216–17, 219, 221, 224, 226, 228, 230–231, 233, 236, 267, 398 mesophyte, 54 Metarhizium anisopliae, 401 methanol, 12, 36–7, 39, 129, 132, 136, 158, 217, 335, 353, 403–406, 408 methyl ether, 157, 403 methyl oleate, 270–271 methylamine, 353 Methylobacterium sp., 344 methylotrophic bacteria, 335–6 microbe, 4–6, 334–59 microbial community, 4, 334–59 microchannel, 92, 94 microfibril, 13, 18, 49–53, 91–2, 94–5, 300 microorganism, 11, 43, 53, 105, 126, 289, 334–59 middle lamella, 16 mimicry, 412–14 Mimulus aurantiacus, 403 minimum transpiration rate, 304, 307

INDEX mitogen activated protein kinase, 382 Mnesampela privata, 409 mobility, 38, 101, 131, 258–65, 267, 269, 271–3, 280–281, 286, 296 molar volume, 262–5, 273, 283, 293, 296–7 molecular size, 261–2, 265, 272, 283 monensin, 204 Monoclea gottschei, 40 monodisperse alcohol ethoxylate, 257, 269, 272 Monodora sp., 61 monoglyceride, 270 monooxygenase, 193 Moraceae, 25, 27–8, 30, 32 morphogenesis, 6, 8, 381, 387 moss, 11, 19, 40, 56, 75, 100, 151 motility, 351 multiflorenol, 198 Musa paradisaica, 61, 72, 86, 93 Musa sp., 103 Musaceae, 61, 86 Muscari atlanticum, 28 Musci, 62 mutant, 5, 44, 61, 68, 86–7, 175, 182–3, 185–6, 188–91, 193–6, 199, 201, 205, 230, 232, 234, 236, 312–29, 343–4, 350–351, 355, 383, 387–8, 390, 414–15 Myosotis arvensis, 57, 59 Myrica germanica, 149, 151–2, 158 Myriophyllum sp., 24, 57, 59 Myriophyllum verticillatum, 24 Myrsinaceae, 24, 27 Myrtaceae, 25, 27–8, 31, 64–5, 67, 69–70, 71, 403, 405, 408 N-acyl homoserine lactones, 351, nanoindentation, 137 naphthenic oil, 270 Narcissus pseudonarcissus, 27, 43 Narcissus sp., 41 Narthecium asiaticum, 69 Nasturtium officinale, 406, 410 Naturstoff reagent, 223 nectar, 92, 399 nectary, 33, 92, 410 negative staining, 79, 84, 97 Nelumbo nucifera, 56, 64, 72, 79, 81, 83, 100, 152, 154, 170 Nelumbonaceae, 64 Nematothallus sp., 56 Nepenthaceae, 69 Nepenthes alata, 69, 161, 170, 399 Nepenthes bicalcarata, 399 Nepenthes rufescens, 69 Nepenthes sp., 71, 86, 93 Nerium oleander, 306–308 niche, 339, 342, 350, 352, 354 Nicolaia elatior, 66 Nicotiana attenuata, 406 Nicotiana glauca, 26, 45, 51, 61, 88

INDEX Nicotiana sp., 159 Nicotiana tabacum, 26, 31, 51, 59 nitrogen, 159, 353, 415 NO2 , 107 nonacosan-10,13-diol, 83 nonacosan-10-one, 154 nonacosan-14,15-diol, 154 nonacosan-14-ol, 99, 148–9, 151, 193 nonacosan-14-one, 88, 154 nonacosan-15-ol, 72, 84, 148–9, 151, 191, 193 nonacosan-15-one, 72, 84, 99, 154, 182, 191, 193 nonacosan-4,10-diol, 72, 83 nonacosane, 72, 83, 99, 154, 182, 191, 193–4, 329, 408 nonacosan-5,10-diol, 83 non-ester cutin, 37, 47, 129 Nothofagaceae, 59 Nothofagus solandri var. Cliffortioides, 59 Nothofagus sp., 59 Notholaena sp., 70 Notothylas orbicularis, 40 nuclear magnetic resonance spectroscopy, 47, 129, 130–131, 133–4, 137–41, 174, 206, 268 nutrient, 228, 289, 335, 342–7, 350, 352–5, 357–8, 368, 399 nymphoposition, 399

obligate biotrophic fungi, 368, 376, 385 octa-9,10-decenoate, 320 octadecanoic acid, 295 Odosicyos sp., 68 Oecophylla smaragdina, 400 Oenothera organensis, 28 Oidium neolycopersici, 383 oil body, 204 oil, 41, 190, 252, 266, 268–71, 354 Olea chrysophylla, 223 Olea europaea, 68, 155, 227–8, 345 Olea sp., 234 Oleaceae, 28–9, 68 oleanane, 155–6 oleanolic acid, 71–2, 89, 156–7 oleosin, 204 Oligotrichum hercynicum, 63 Onagraceae, 28, 59, 62, 67 optical property, 5, 216–38 Opuntia engelmannii, 164 Orchidaceae, 413 organ fusion, 312–15, 318, 320–321, 328–9 Oryza sativa, 55, 67, 91, 161, 190, 194, 204–205, 207, 319, 356–7, 368–9, 374–5, 384 osmiophilic, 13, 17, 21, 23, 40, 51, 204–205 osmotolerance, 351 Osmunda regalis, 61, 88, 152, 154, 158, 172 Osmundaceae, 61 Ostrinia nubilalis, 402, 408 ouricuri, 57, 93 oviposition stimulant, 410 oviposition, 6, 399–400, 409–411, 414, 416

433 Oxalidaceae, 66 Oxalis hedysaroides, 66 oxidoreductase, 316, 322 oxidosqualene, 196, 198 oxygen, 3, 41, 150, 157–8, 160, 192–3, 221, 349, 390 P450 enzyme, 127, 319 Paeonia mlokosewitchii, 61 Paeonia officinalis, 61, 154 Paeonia suffruticosa, 66 Paeonia sp., 61, 88 Paeoniaceae, 61, 66 palmitone, 88, 98, 154 Pandanaceae, 66 Pandanus montanus, 66 Pantoea agglomerans, 343 Papaver rhoeas, 158 Papaver somniferum, 63, 65, 79–81, 83, 97, 170 Papaver sp., 158–9, 164 Papaveraceae, 63, 65, 151–2, 154, 164 Papilio, 402, 412 parasitoid, 398, 412, 414–15 particulate matter, 107, 251 partition coefficient, 254–8, 260, 263, 266, 273, 280, 295–8 partitioning, 159, 205, 250–251, 254–5, 266, 268, 352 passive adhesion, 372–4 pathogen, 4, 126, 137, 141, 199, 312, 336, 342, 350–351, 356–9, 368, 371, 375, 382–7, 390–391, 401, 416 pathogenesis, 379, 385–6 pectin, 14, 16, 18, 22, 33, 43, 45, 48, 107–109, 205–206, 295 pectinase, 16, 357 pellicle, 40 penetration peg, 380, 389–91 penetration, 35–6, 91, 218, 223, 232, 237, 262, 267–72, 280–281, 283–7, 302, 322, 324, 329, 343, 379–80, 382, 384, 389–91 peristomatal transpiration, 305 permeance, 258–60, 263, 265–6, 292–9, 301–304, 306–309 pesticide, 252, 267, 273, 416 petal, 27, 30–31, 41, 43, 95, 152, 158, 216, 226–8, 230, 232, 236 petroleum, 96, 271 Petunia hybrida, 232 Phaedon cochleariae, 406, 410 phase separation, 171, 174 phase transition, 138, 140, 302–303 Phaseolus vulgaris, 12–13, 24, 26, 28, 31, 43, 50, 54, 57, 59, 88, 232, 383, 388 Phenacospermum guianense, 61 phenanthroline, 193 phenolics, 36, 221, 223–4, 226, 231, 237 phenyl propanoid, 158 phenylethyl alcohol, 158 phenylpropanoid metabolism, 224 pheromone, 252, 413

434 Phillyrea latifolia, 92, 227 Philodendron sp., 14 Phoma sp., 357 Phoradendron flavescens, 13 Phormiaceae, 69 Phormium cookianum, 69 Phormium sp., 34, 47, 54 Phormium tenax, 21, 26, 34, 41, 47, 94 phosphate, 222, 271–2, 335, 353 phosphonate, 271 photo-dimerisation, 160 photo-oxidation, 104 photosynthesis, 2, 4–5, 11, 40, 56, 105, 216, 221, 228, 231–3, 236–7, 353 Phragmites australis, 61, 89, 95 Phyllagathis rotundifolia, 237 Phyllirea latifolia, 29 Phyllitis scolopendrium, 29 phylloplane, 334, 346 Phyllosticta ampelicida, 368, 372, 376, 383–5, 387, 390 phylogeny, 55, 70, 77, 344 Phytophthora megasperma f. sp. glycinea, 384 phytosterol, 159 phytotoxin, 354, 357 Picea abies, 15, 17, 23, 29, 41, 43, 47–8, 50–51, 63, 91 Picea glauca, 63 Picea pungens, 63, 81, 97–8, 154, 170, 223, 232–3, 237 Picea pungens var. glauca, 63 Picea rubens, 29 Picea sitchensis, 15, 17, 19, 23, 29, 43, 46, 48, 50–51, 63, 79, 81–2, 94–7, 101–102, 105, 107 Picea sp., 50, 54, 79, 109 Pieris brassicae, 408 pigment, 5, 205, 216–38 pili, 343, 351 Pimpla turionellae, 412 Pinaceae, 25–6, 29–31, 63–4, 70 Pinguicula vulgaris, 223 Pinus sylvestris, 29 Pinus balfouriana, 64 Pinus banksiana, 105, 228 Pinus nigra, 31, 64 Pinus nigra var. maritima, 64 Pinus radiata, 64, 94 Pinus sp., 64, 79, 226, 228 Pinus strobus, 29, 64, 105 Pinus sylvestris, 29, 31, 64, 70, 79, 95, 226 Pistacia palaestina, 20, 26 Pistacia sp., 54 Pistacia vera, 60 Pisum sativum, 24, 62, 67, 72, 77, 80, 98, 103, 151–2, 164, 170, 187, 190–192, 226, 228, 232, 234, 377 Pisum sativum cv. ‘Meteor’, ‘Kelvedon Wonder’, ‘Alaska’, 67 Pisum sp., 71 Pityrogramma sp., 70, 88 Pityrogramma triangularis, 70 Plagiochasma rupestre, 60

INDEX plant protection, 4, 250–273, 289 Plantaginaceae, 26, 29, 31 Plantago major, 17, 26, 29, 31, 36, 50, 93, 94 plasma membrane, 201–206, 321 plasmid, 349, 356–7 Plasmopara viticola, 356 plasticiser, 283, 285–6 plastid, 183–5, 197, 201 Plutella xylostella, 401, 403–405, 407, 416 Poa colensoi, 65, 93 Poa nemoralis, 67, 103 Poa sp., 71 Poaceae, 24, 26, 30–33, 59–62, 64–5, 67–8, 71, 77, 79, 83, 86, 103, 152, 154, 159, 164, 402, 404–406, 408, 415 Pogonatum belangeri, 64, 81 Pogonatum rubenti-viride, 65, 98 Pogonatum urnigerum, 64, 81 point of deliquescence, 252 polar compound, 146, 250, 281, 287, 289 polar domain, 283, 287 polar organic solute, 3 polar pathway, 3, 280–289, 295, 298, 301 polarisation, 130, 204, 230–231 pollen tube, 92, 322 pollen, 316–17, 320, 322–5, pollination, 322–4 pollinator, 227–30, 232, 236, 412–13 pollutant, 3–4, 104–105, 161, 250, 271, 273 polydisperse alcohol ethoxylate, 272 Polygala myrtifolia, 63, 66 Polygalaceae, 63, 66 Polygonaceae, 29, 59, 67, 400 polyketide, 158, 195 polylamellate, 17, 36–7 polymer, 4, 11, 13, 21, 37–40, 45, 47–8, 50, 71–2, 86, 91, 93, 100, 102, 104, 107, 109–110, 126–41, 146, 160, 259, 267–8, 270, 280, 282–3, 295, 297, 299–300, 302–304, 343, 353, 388, 399, 415 polymorphic, 96, 98–9 polyphenols, 221, 223 polysaccharide, 14, 35, 37, 39–41, 43, 47–51, 91–2, 94, 108–109, 223, 238, 295, 300, 302, 343, 351, 354, 372 Polytrichaceae, 63–4, 66 Polytrichales, 40, 56, 63–6 Polytrichum formosum, 64 Polytrichum juniperinum, 64, 66 Polytrichum sp., 54, 56 Populus canescens, 283–4, 286 Populus deltoides, 404 Populus nigra, 404 pore, 3, 19, 40–41, 89, 91–5, 102, 105, 109, 169, 206, 237, 281–2, 288–9, 294–5, 297–9, 301–302, 304, 307, 387, 391 porosity, 99, 298 porphyrin, 192 Posidonia australis, 32 Posidoniaceae, 32

435

INDEX Potamogeton crispus, 17, 29, 33, 43, 57, 59 Potamogeton sp., 33, 53–4 Potamogetonaceae, 29, 33, 59 potassium, 100, 281, 283 potato, 62, 75, 127, 346, 413 Potentilla fruticosa, 62 Potentilla sp., 75 powdery mildew, 336, 356, 368, 372–3, 375–7, 379, 383, 386, 390–391 predator, 398, 412, 414–15 prediction equation, 257, 265, 296, 307 Prenia sladeniana, 69, 103 primary alcohol, 38, 71, 73, 75, 77–8, 83, 88, 96, 98, 103–105, 150, 157–8, 162–4, 166–7, 169, 171, 184, 189–90, 193–4, 200, 272, 329, 377, 402–403 primary cell wall, 14–18, 22–3, 41, 43, 45, 48, 52–3, 92, 94, 107–108, 206, 376 primary germ tube, 369, 377–82, 386 primordium, 17–18, 312, 325, 327–8 Primula sp., 70 Primulaceae, 70, 88, 160 procuticle, 16–19, 21, 33, 40, 91, 107–108 pronase, 322, 372 Prosopis sp., 66, 77 Prosopis tamarugo, 69 Proteaceae, 25, 31, 68 protoderm, 17, 41, 320, 326–9 Prunus avium, 300 Prunus domestica, 58, 64, 80, 155 Prunus domestica cv. ‘d’Agen’, 64 Prunus laurocerasus, 29, 31, 59, 89, 155, 161, 166–7, 169, 173–4, 282, 299, 301 Prunus lusitanica, 38 Prunus persica, 29, 59 Prunus persica cv. ‘Red Haven’, 59 Prunus sp., 58, 64, 72, 79, 80, 89, 155, 161, 166, 282, 300–301 Pseudomonas fluorescens, 343, 353, 356 Pseudomonas syringae, 339, 341–3, 346–7, 349–52, 354–8 Pseudomonas, 343–5, 354, 356 Pseudotsuga menziesii, 26, 64, 97 Pseudotsuga sp., 54, 79 Psila rosae, 416 Psilotaceae, 26 Psilotum nudum, 26, 40 Pteridaceae, 70 pteridophyte, 40 Pteris sp., 70 Puccinia graminis tritici, 388 Puccinia hordei, 388 Puccinia sorghi, 388 Puccinia sp., 368, 388 pycnidiospore, 372 pyrolysis, 47 Pyrus communis, 12, 29, 33–4, 36, 53, 283, 299, 303 Pyrus communis cv. ‘Passe Crassanne’, 29 quantitative property-property relationship, 257

quantitative structure-property relationship, 257 quartz, 98, 216 Quercus petraea, 306, 308 Quercus pubescens, 67, 69, 76, 89 Quercus robur, 67, 69 Quercus sp., 103 Quercus velutina, 13, 17, 29, 48, 55 quorum sensing, 351, 355

radiation, 2, 5, 11, 56, 105, 160–161, 216–38, 347–8, 350–351 rain, 4, 100–101, 105, 235, 251, 341, 343, 347–8, 353, 368, 370, 399 rainforest, 54 Ranunculaceae, 30, 59, 63–4, 164 Ranunculus penicillatus, 345 Raphanus sp., 322 Rayleigh scattering, 230–231, 233 Rebouilia hemisphaerica, 60 reconstituted wax, 257, 261–4, 269, 271–3, 295 red-fluorescent protein, 340 reductase, 183–4, 186–90, 192, 194, 196, 201–203, 316–17, 319, 321 reflectance, 194, 218–19, 228, 230–237, 412 reflection, 5, 104, 217–18, 227, 230, 231, 412 regulation, 140, 166, 168, 194, 199–200, 203, 207, 326–9, 351, 355, 370, 379–80, 385, 388–90 relative water deficit at maximum stomatal closure, 307 remote sensing, 216, 231 reproduction, 322–6 resorcinol, 158 Restionaceae, 66 reticulate, 12, 15, 17–20, 22–3, 37, 43–5, 48–53, 55–6, 92, 94, 109, 301 reticulum, 15, 17, 22, 35, 43–5, 47–8, 51–2, 54, 91–2, 94, 109, 183, 202–203, 300 Rhamnaceae, 61 rheology, 137–8 Rhodococcus fascians, 356 Rhododendron ponticum, 59 Rhododendron sp., 83, 195 Rhodophyta, 40 Rhodotorula sp., 346 rhodoxanthin, 230 Rhus cotinus, 64, 70, 79, 81, 97 Rhus cotinus atropurpurea, 64, 70, 81 ribbon, 47, 58, 62, 72–3, 75–7, 84, 98, 101, 369, 377 Ribes nigrum, 29, 33 rice blast, 91, 356, 368 Ricinus communis, 33, 156, 158, 170 Robinia sp., 67, 69 rod, 58, 61, 72, 75, 79, 83–4, 86, 90, 93, 95, 98, 102, 105 rodlet, 58, 60, 79, 86, 88–9, 93, 95, 101, 103, 171–2 Rosa canina, 58 Rosa cv. ‘Baccara’, 66, 64 Rosa damascena, 152 Rosa x hybrida, 168

436 Rosa sp., 62, 64, 66, 75–6, 79, 152, 168 Rosaceae, 26–9, 31–3, 59–60, 62–4, 66 Rosaceae, 26–9, 31–3, 60, 62–4, 66, 75, 151, 164, 405, 407 roughness, 252, 254 Rubiaceae, 30, 59 Rubus idaeus, 407 Rumex conglomeratus, 29 Rumex crispus, 400 Rumex obtusifolia, 57, 59 Rumex obtusifolius, 59 Rumex sp., 29 rust, 370, 372–3, 388–9 Rutaceae, 27, 30, 33, 60–61

sabinic acid, 100 Saccharomyces cerevisiae, 185, 188, 201–202 Saccharum officinarum, 56, 58, 61, 71–2, 86, 93, 95, 161 Saelania glaucescens, 62, 75 Salicaceae, 70 Salix alba argentea, 70 Salmonella sp., 357 salt, 3, 35, 234, 283, 348 Sapium sepiferum, 70 Sarracenia psittacina, 26 Sarracenia purpurea, 26, 32 Sarracenia sp., 92 Sarraceniaceae, 26, 32 Saxifraga granulata, 33 Saxifragaceae, 27–9, 33 scale, 58, 91, 94, 97 scanning electron microscope, 46, 57–8, 73, 78, 84, 86, 88–90, 93–5, 107, 167, 173, 288, 321, 369, 371, 374, 378, 380, 383 scattering, 5, 56, 79, 217–18, 230–231, 223, 235–7 Sceletium compactum, 61 Schefflera actinophylla, 283 Scopolodendron sp., 29 screening, 5, 194, 217–18, 220, 226–8, 230–231, 233–5, 313–15, 318, 324, 329–30, 339 Scrophulariaceae, 33 Secale cereale, 26, 68, 92, 195, 226 Secale cereale cv. ‘Lovaspatonia’, 68 Secale sp., 164, 226 secondary alcohol, 58, 64, 68, 71, 76–7, 79, 82–4, 87, 96–101, 134, 148, 15, 154–5, 158–9, 161–4, 182, 184, 190–195, 199, 202, 329, 407, 413 secondary metabolite, 4, 206, 250, 273, 354, 398, 410 Sedum telephium, 65 segregation, 89, 100, 173–4 Selaginella sp., 237 Selaginella uncinata, 236–7 Selaginellales, 41 self-assembly, 23, 36, 38, 96, 101, 109, 383 self-diffusion, 174, 298, 301–302 semi-volatile, 4, 252–4 Septoria sp., 357

INDEX sesquiterpene, 401–402, 413 S-ethyl dipropylthiocarbamate, 416 sex pheromone, 413 shade, 55, 229, 236–7, 299 siderophore, 354 sieve effect, 236–7 signal transduction, 326, 381–2, 386–7, 416 signalling, 204, 238, 324, 326, 377–9, 381, 383, 386–7 Silene dioica, 26 Siluarian, 1 silver, 36, 39, 287–8 Silybum marianum, 234 silylation, 148 Simmondsia chinensis, 66, 189–90, 318–19 simulation of foliar uptake, 260, 262, 272 simultaneous bilateral desorption, 259, 260 Sisyrinchium filifolium, 26, 31, 51 Sitobion avenae, 406 sitosterol, 159, 206 size-selectivity, 262–3 skin, 1, 226–8, 259–62, 265–7 SO2 , 161 Solanaceae, 26, 30–31, 38, 51, 59, 61–2, 402, 406, 408, 413 Solanum berthaulii, 408 Solanum tuberosum, 62, 75, 127, 346, 413 solubilisation, 190 solubility, 146, 183, 206, 257, 259, 266, 280–281, 286, 302 soluble cuticular lipids, 23, 36–9, 57, 96, 109 Sorghum bicolor, 61–2, 68 Spartocytisus filipes, 26, 29, 31, 33, 94 Spartocytisus sp., 54 Spathiphyllum wallissii, 60 spectrophotometry, 217, 220 spectroradiometer, 218 specular reflectance, 230–232, 236 Sphagnum fimbriatum, 19, 40 Sphagnum palustre, 40 sphingolipid, 188–9, 201–202, 204–205 Spinacia oleracea, 27, 30, 39, 52, 54, 60, 127, 182 spiral, 47, 65, 72, 75–6, 82–3, 97–8, 335 spore deposition, 368, 370, 374, 377–8, 384–5, 387 spore, 5, 91, 251, 338, 342–3, 357, 368, 370–380, 382–5, 387, 389, 391 Sporobolomyces sp., 346 sporophyte, 11, 19, 40, 56, 72, 91, 100, 151 sporulation, 391 squalene, 196, 198 Stellaria media, 30, 57, 60 Stephanotis floribunda, 283 stigma, 25, 30–31, 33, 92, 322–5 stigmasterol, 159 stoma, 1–4, 11–12, 31, 33, 40, 44, 46, 50, 55–6, 86, 91, 93, 102–105, 107, 147, 221, 229, 237, 253, 281, 284, 287–8, 292, 294, 303–309, 340, 346, 348, 387–9, 391, 414 stomatal closure, 284, 292, 303–304, 307–309 Strelitzia reginae, 61, 86

INDEX Strelitzia sp., 61, 86, 93, 95, 97, 103 Strelitziaceae, 61, 86 structural colour, 237–8 structural type, 17, 24, 49, 52, 54, 71 Stypandra glauca, 69 suberin, 34, 36, 44, 127 sublethal water deficit, 307 substomatal, 13, 55, 147, 356, 391 sucrose, 281, 353–4 sugar, 289, 352–4, 402, 405, 410 sun leaf, 55, 236, 299 superficial wax, 57, 89, 95 surface lipid, 57, 416 surfactant, 257, 266, 268–72, 354 Syagurus coronata, 30, 61, 57, 61, 70, 93 Symplocos hallensis, 30 Symplocos paniculata, 26, 30 Syringa vulgaris, 41 syringomycin, 354, 357

tabtoxin, 357 Tamaricaceae, 27, 31 Tamarix pentandra, 27, 31 Tamus communis, 60 Tanacetum vulgare, 405, 409–411 tapetum, 204 Taraxacum officinale, 17, 27, 43, 52–4, 60 Taraxacum sp., 54 taraxasterol, 198 taraxerane, 156 taraxerol, 75 Targiona hypophylla, 60 taxonomy, 70, 77 Taxus baccata, 158, 170 Taxus sp., 79 Teflon, 372, 384 temperature, 36, 46, 76, 86, 89, 99, 107, 132, 134, 136, 138–41, 146–7, 160, 167, 174, 188, 255, 262–4, 267, 272, 283–6, 292–3, 295, 299, 302–304, 343, 348, 352, 356, 368, 374, 384 Terminalia cf seyrigii, 70 terpene, 270 terpineol, 270–271 tetracosanoic acid, 171, 191 Thalictrum flavum, 64, 81 Theaceae, 25–6, 28, 30–31 Thellungiella halophila, 63 Thellungiella parvula, 63 thickness, 14–19, 21–3, 34, 36–7, 40, 42, 45, 52–5, 92, 94–5, 108, 129, 146–7, 165, 173, 185, 200, 204, 217, 220, 235, 237–8, 259, 261, 268, 273, 296, 298–9, 348 thin-layer chromatography, 148 thioesterase, 183–4, 197 Thymelaceae, 61 Ticoa harrisii, 27 Tilia cordata, 60 Tilia magnifica, 69

437 Tilia tomentosa, 155 Tiliaceae, 60, 69 tocopherol, 159 toluidine blue, 315–16, 318–19, 322, 327 tomato, 5, 11, 17, 31, 52, 59, 127, 130–131, 134, 137–41, 155, 165, 173, 227, 255–6, 296–301, 343, 357, 383 tortuosity, 262–3, 268, 296, 298 Tradescantia virginiana, 31 Tradescantia sp., 41 trafficking of wax constituents, 199, 201, 205 transcription factor, 199–200, 327–9 transfer, 102, 169, 190, 201–202, 204–206, 231, 250, 254–5, 258–60, 326, 356 transmembrane domain, 190, 205–206, 321 transmission electron microscope, 13, 17, 23, 40, 52, 58, 84, 88, 93–4, 204, 321 transmittance, 217–20, 228, 230, 236–7 transpiration, 2–3, 7, 168, 292–4, 299–309, 417 transport property, 8, 260, 262, 265, 267, 269, 273, 294, 300 transporter, 102–103, 199, 202–203, 205–206, 321 triacontanal, 86, 169–70, 194, 381, 399 triacyl glycerides, 325 triacylglycerol, 188 triacylglycerolipase, 389 tri-butyl-phosphate, 271 Trichoderma sp., 358 Trichomanes elegans, 237 trichome, 33, 70, 73, 76, 92, 159, 223, 231, 234–5, 281, 287–8, 318, 320, 346, 348, 351, 354, 370, 399, 401–402, 406, 413, 415 Trifolium pratense, 68, 99 Trifolium repens, 60, 68–9, 72, 78, 93, 103 Trifolium sp., 71, 93, 103 trilinolein, 326 triterpene acetate, 88 triterpene acid, 72 triterpene synthase, 198 triterpene, 75, 198 triterpenoid biosynthesis, 156, 196–9 triterpenoid, 75, 88–9, 148, 155–7, 159, 164, 169–70, 173–4, 196–9, 405 triterpenol, 156–7 Triticum aestivum, 31, 44, 65, 68, 71–2, 91, 97, 155, 160, 195, 222, 402, 405–406 Triticum sp., 71, 164, 405–406 Triticum turgidum, 233 tritriacontan-10-ol, 149 tritriacontan-16,18-dione, 71–2, 79 Tropaeolaceae, 31 Tropaeolum majus, 31, 64, 72, 79, 81, 83, 97, 102 Tropaeolum speciosum, 41, 64 tube, 19, 46, 47, 58, 63–5, 69, 72, 74–7, 79–80, 82–4, 91–8, 100–103, 105, 107, 323, 326, 335, 343, 369, 375, 377–89, 391 Tulipa gesneriana, 30, 33, 58, 64, 81 Tulipa kaufmanniana, 64, 79, 83, 97 Tulipa sp., 219

438 Tupiocoris notatus, 405 turgor, 304, 306, 309, 389–91 Typha angustifolia, 61 Typha elephantina, 61 Typha latifolia, 61 Typha sp., 86 Typhaceae, 61

Ulex europaeus, 60 ultra-hydrophobicity, 56 ultraviolet radiation, 2, 5, 29, 133, 161, 216–18, 220–224, 226–9, 231–4, 237–8, 347–51, 412 Umbelliferae, 24–5 unbranched, 150–151, 155 unilateral desorption from the outer surface, 260–263, 266, 272 urediniospore, 370, 372–3, 388 Uromyces appendiculatus, 383, 388–9, 391 Uromyces sp., 368 ursane, 155–6 ursolic acid, 72, 89, 156–7 Utricularia laterifolia, 18, 24, 27, 41 Utricularia monanthos, 33, 92 Utricularia sandersonii, 17, 21, 31 Utricularia sp., 23, 49, 52 UV excitation, 220 UV fluorimetry, 220 UV screening, 217–18, 220, 226–8, 231, 233–4 uvaol, 156 Uvularia floridiana, 69

Vaccinium ashei, 65, 101 Vaccinium myrtillus, 234 Vaccinium reticulatum, 30, 94 vacuole, 5, 219, 224–9, 236–8, 321 Valerianaceae, 60, 62 Vanilla planifolia, 27 vapour pressure, 252, 271, 293 vapour, 3–4, 11, 36, 250, 252–4, 271, 293, 296, 302–303, 348 vascular plant, 11, 40, 42, 54–6, 127, 151 Velloziaceae, 66 verbascoside, 226 Veronica anagallis-aquatica, 33 very long chain fatty acid, 182–5, 188–9, 201–204, 207, 318–19, 324–5 vesicle, 70, 201, 204–205, 389, 391 viable-but-not-culturable-microbial cells, 337, 350 Vicia faba, 30–31, 33, 60, 127, 129, 158, 226, 283, 285, 287–8, 406 Vicia sativa, 127 Vinca major, 14 Vinca minor, 305–306 viscoelasticity, 137 Vitaceae, 60, 66 Vitis vinifera, 39, 60, 66, 71–2, 155, 226–7, 405

INDEX Vitis vinifera cv. ‘Siebel’, 66 Vitis vinifera cv. ‘Sultana’, ‘Kishmish’, ‘Thompson seedless’, 66 volatile, 3, 4, 93, 101, 252–4, 266–7, 373, 413 volatility, 4, 132, 270–271 Wachskörnchen, 58 Wachsplättchen, 58 Wachsschichten, 58 Wachsstäbchen, 58 Wachsüberzug, 58 wasp, 413–14 water activity, 2, 292–4 water droplet, 5, 13, 235–6 water permeability, 7, 91, 268, 280, 283, 286–7, 292–309, 321 water potential, 292–3, 304–306, 348 water status, 292 Watsonia bulbifera, 69 wax amount, 57, 102, 165–8, 173, 183, 185, 200–299, 300, 320–321 wax composition, 89, 145–75, 198–9, 300, 315, 344, 402, 415 wax extraction, 37, 145–7, 223, 280, 285–6, 299 wax film, 57, 76, 86, 89, 91, 95, 103, 109, 165–6, 168–9, 171, 173, 295, 415 wax plate, 16, 58, 68–9, 71–9, 82–4, 86–8, 90–91, 93, 96–9, 101–105 wax platelet, 75, 88, 103–104, 169, 170–171, 174, 399 wax secretion, 92, 104, 204 wax/water partition coefficient, 257, 266 wet deposition, 251 wettability, 104–105, 348, 416 wheat, 16, 44, 57, 71, 79, 83, 90–91, 104, 195, 207, 221, 375–6, 388 Williamodendron quadrillocellatum, 61 wind, 105, 251–3, 341, 343, 368, 370, 373 Wooleya farinosa, 62, 86 Xanthomonas campestris, 356–7 Xanthomonas sp., 344 xeromorphic leaf, 307–308 x-ray, 93, 95, 174, 206 yeast, 6, 188–9, 201–204, 251, 289, 319, 326, 334, 337–8, 342–7, 349, 351, 355–7, 409 Yucca filamentosa, 69 Yucca sp., 55 Zea mays, 33, 68, 71, 105, 164, 182, 186, 188–90, 193–4, 199, 207, 228, 231, 312, 317, 319, 325–6, 343, 388, 402, 408 Zingiberaceae, 61, 66 Zostera capensis, 33 Zosteraceae, 33

Biology of the Plant Cuticle Edited by Markus Riederer, Caroline Müller Copyright © 2006 by Blackwell Publishing Ltd (a)

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Plate 1 Atomic force microscopy (AFM) images of epicuticular wax regenerating on a living Galanthus nivalis leaf adaxial surface, following removal of the earlier-secreted wax with epoxy resin glue. The images show changes in a 6 × 6 μm area during an 80-min time frame. Flat, lobed plates extend over the whole surface to form first a monolayer (light grey) (a,b) and then bilayered structures (dark grey) (c,d). Independently, rod-like crystals arise directly from the cleaned cuticle proper (CP) surface, extending by growth at their distal ends, not from the base. Diagonal arrows (a,b) and vertical arrows (c,d) mark reference points that remain in fixed positions during rod extension growth. The black arrow in (a) marks a crystal that in (b) has been removed by the AFM tip. (a–d) bars = 1 μm; reproduced with permission from Koch et al. (2004), Journal of Experimental Botany, 55, 711–718. (e,f) SEM images of wheat leaf wax recrystallised from chloroform on glass forming an amorphous film (e) and on freshly cleaved highly ordered pyrolytic graphite (HOPG; PLANO GmbH, Wetzlar, Germany) (f), producing highly ordered wax crystals; reproduced with permission from Kerstin Koch (see also Koch, K., Barthlott, W., Koch, S., Hommes, A., Wandelt, K., Mamdouh, H., De-Feyter, S. and Broekmann, P. Structural analysis of wheat wax (Triticum aestivum, c.v. ‘Naturastar’ L.): from the molecular level to three dimensional crystals, Planta, in press).

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Plate 2 Gas chromatographic (GC) separation and mass spectrometric (MS) identification of plant cuticular wax components. (a) GC–FID trace of the wax mixture extracted from inflorescence stems of Arabidopsis thaliana ecotype Columbia. (b) Mixed mass spectrum of trimethyl silyl (TMSi) derivatives of nonacosan-14-ol and nonacosan-15-ol [corresponding to the GC peak at 23.5 min in (a)] showing characteristic α-fragments at m/z = 285, 299, 313. (c) GC–MS traces of the α-ions, demonstrating that the isomeric 14- and 15-alcohols cannot be separated under the GC conditions used. (d) Mixed mass spectrum of TMSi derivatives of tritriacontan-10-ol, tritriacontan-12-ol and tritriacontan-14-ol (from leaf wax of Myricaria germanica) showing characteristic α-fragments at m/z = 229, 257, 285 and m/z = 369, 397, 425, respectively. (e) GC–MS traces of the α-ions, demonstrating that the isomeric 10-, 12- and 14-alcohols can be partially separated under the GC conditions used.

Plate 3 Green fluorescent protein (GFP) experiments demonstrate the location of the proteins involved in wax production. These images represent a projection of optic sections collected with the laser scanning confocal microscope. The enoyl-CoA reductase (CER10-GFP) component of the elongase was found in the endoplasmic reticulum of pavement cells in the leaf of Arabidposis thaliana. ABC transporter CER5 was localised to the plasma membrane in A. thaliana stems (GFP-CER5). Magnification bar = 10 μm.

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leaf Plate 4 Isolines of the relative concentration of a semi-volatile compound in the unstirred layer in the vicinity of the leaf surface. Relative concentrations are colour-coded from red (highest) to dark blue (lowest). In the finite-element model leading to this picture a regular distribution on the leaf surface of solid deposits of the organic compound (red spots) and stomata was assumed. The concentration distributions at two wind speeds (0 and 1 m s−1 ) are shown. Figure from Riederer et al. (2002) by kind permission by Oxford University Press.

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(c)

(d)

Plate 5 Multispectral autofluorescence microimaging of a 100-μm-thick cross-section from a Triticum aestivum L. leaf in phosphate buffer. Autofluorescence images in panel (a) were excited at 365 nm and detected in the blue range at 470 nm, and in panel (b) were excited at 436 nm and detected in the yellow range at 580 nm. Combination of monochrome images, (a) and (b), with blue and yellow colours assigned to the 470 and 580 emission bands, respectively, is shown in panel C (bar = 50 μm). Panel (d) depicts fluorescence intensity profiles for the 470 and 580 nm bands along the two, x and y, directions indicated by dotted arrows in (c). Panels (c) and (d) clearly reveal inhomogeneous fluorescence characteristics and, thus, quite large variations in spatial localisation of different fluorescing compounds: the UV-induced blue fluorescence emanates from cell walls, while the blue-induced yellow signal appears to be confined to cuticles, guard cells and sclerenchyma bands (G. Agati, Corrado Tani and Z.G. Cerovic, unpublished data).

Plate 6 View of a colonised bean leaf using confocal laser scanning microscopy. Individual bacterial colonisers producing GFP are visible as green fluorescent dots. The centre shows a stoma; the cuticular top of an epidermal plant cell is visible on the left top corner while at the bottom right the picture slices into the leaf’s palisade parenchyma. Chlorophyll is coloured red in this picture.

Plate 7 (a–d) Light (LM) and (e–f) scanning electron microscopic (SEM) investigation of silver (Ag) deposits around stomata and in trichomes of V. faba leaves after treatment with AgNO3 . (a and e) Stomata of untreated leaf surfaces serving as control. (b and f) Stomata of AgNO3 -treated leaf surfaces with characteristic silver deposits surrounding the stomatal pore. (c and g) Trichomes of untreated leaf surfaces serving as control. (f and h) Trichomes of AgNO3 -treated leaf surfaces with characteristic silver deposits in the base and head of the trichome. Data from Schlegel et al. (2005).

m

Recognition of a signal

m

m

m

Co-pollination of wild-type and mutant pollen

(d)

H2O

m

Polen tube grows

Successful fertilization of the mutant pollen

Pollen Pollen germination hydration and degradation of stigmatic cuticle

H2O

Recognition of a Hydration of both signal from wildwild-type and type pollen mutant pollen

m

Adhesion of Recognition of wild-type pollen a signal (lipids?) on stigma surface

(b)

Accumulation of callose in the papillae Pollen hydration

Stigmatic papillae

Pollen

Plate 8 Schematic diagram of successive events in compatible pollination. (a) Diagram of stigma of Arabidopsis thaliana. (b) Sequential events following compatible pollination. (c) Events following pollination by mutant pollen defective in pollen–stigma recognition. (d) Restoration of mutant. Abbreviation: m, pollen fertility by co-pollination with wild-type pollen.

Adhesion of mutant pollen on stigma

(c)

(a)