Plant Solute Transport

Plant Solute Transport Edited by ANTHONY YEO Haywards Heath, West Sussex, UK TIM FLOWERS School of Life Sciences University of Sussex, UK

Plant Solute Transport Edited by ANTHONY YEO Haywards Heath, West Sussex, UK TIM FLOWERS School of Life Sciences University of Sussex, UK

 C

2007 Blackwell Publishing

Blackwell Publishing 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 Pty Ltd, 550 Swanston Street, Carlton, Victoria 3053, Australia Tel: +61 (0)3 8359 1011 The right of the Authors to be identified as the Authors 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 2007 by Blackwell Publishing Ltd ISBN: 978-14051-3995-3 Library of Congress Cataloging-in-Publication Data Plant solute transport/edited by Anthony Yeo and Tim Flowers. p. cm. Includes bibliographical references. ISBN-13: 978-1-4051-3995-3 (hardback : alk. paper) ISBN-10: 1-4051-3995-1 (hardback : alk. paper) 1. Plant translocation. II. Flowers, T. J. (Timothy J.) QK871.P53 571.2–dc22

I. Yeo, A. R.

2007 2006027577

A catalogue record for this title is available from the British Library Set in 10/12 pt Times by TechBooks, New Delhi, India Printed and bound in Singapore by Markono Print Media Pvt Ltd 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 Preface

xiii

Contributors

xvii

1

2

General introduction ANTHONY YEO 1.1 Introduction 1.2 Synopsis 1.3 Concluding remarks Reference Solutes: what are they, where are they and what do they do? TIM FLOWERS 2.1 Solutes: inorganic and organic 2.2 Analysis of inorganic elements 2.2.1 Obtaining material for analysis 2.2.2 Optical methods 2.2.3 Mass spectrometry 2.2.4 X-ray fluorescence 2.2.5 Ion-specific electrodes 2.2.6 Ion chromatography 2.3 Solute concentrations 2.4 Organic compounds 2.5 Range of solutes found in plants 2.6 Localisation 2.6.1 Stereological analysis 2.6.2 Inorganic elements and electron microscopy 2.6.3 Ion-specific microelectrodes 2.6.4 Direct sampling 2.6.5 Use of fluorescent dyes 2.6.6 Flux analysis 2.6.7 Organic compounds 2.7 What do they do? 2.7.1 Vacuoles 2.7.2 Organelles and the cytoplasm 2.7.3 Cell walls 2.7.4 Conclusions References

1 1 3 14 14 15 15 15 15 16 16 17 17 17 17 18 19 19 19 20 21 22 22 23 25 25 25 26 26 26 27

iv 3

4

CONTENTS

The driving forces for water and solute movement TIM FLOWERS and ANTHONY YEO 3.1 Introduction 3.2 Water 3.3 Free energy and the properties of solutions 3.3.1 Free energy and chemical potential 3.3.2 Water potential and water potential gradients 3.3.3 Osmosis and colligative properties 3.4 Cell water relations 3.5 Water movement 3.5.1 Water movement through the soil 3.5.2 Water in cell walls 3.5.3 Water movement across a root (or leaf) 3.5.4 Water movement through the xylem and phloem 3.6 Solute movement 3.6.1 Chemical, electrical and electrochemical potentials and gradients 3.6.2 Diffusion – Fick’s first law 3.6.3 Diffusion potential 3.6.4 Nernst potential 3.6.5 Donnan systems 3.6.6 Goldmann equation 3.7 Coupling of water and solute fluxes References Membrane structure and the study of solute transport across plant membranes MATTHEW GILLIHAM 4.1 Introduction 4.2 Plant membranes 4.2.1 Plant membrane composition 4.2.2 Plant membrane structure 4.3 Studying solute transport across plant membranes 4.4 Transport techniques using intact or semi-intact plant tissue 4.4.1 Plant growth 4.4.1.1 Solution design 4.4.1.2 Using inhibitors 4.4.2 Accumulation and net uptake 4.4.3 Radioactive tracers 4.4.4 Fluorescent solute probes 4.4.5 Electrophysiology 4.4.5.1 Voltage-based measurements (membrane potential and ion concentration) 4.4.5.2 Voltage clamping 4.5 Using isolated membranes for transport studies

29 29 29 31 31 32 33 34 35 38 39 39 40 40 41 41 42 43 43 44 44 45

47 47 47 47 50 51 52 52 52 53 53 54 55 57 58 60 60

CONTENTS

4.5.1 4.5.2

Isolating membranes Assaying transport activities of protoplasts and membrane vesicles 4.6 Using molecular techniques to inform transport studies 4.6.1 Revealing the molecular identity of transporters and testing gene function 4.6.2 Location of transport proteins 4.6.3 Heterologous expression 4.7 Combining techniques (an example of increasing resolution and physiological context) 4.8 Future development 4.9 Conclusions Acknowledgements References

5

Transport across plant membranes FRANS J. MAATHUIS 5.1 Introduction 5.1.1 Plant solutes 5.1.2 Definitions and terminology 5.1.3 Some formalisms 5.2 Passive transport 5.2.1 Diffusion through membranes 5.2.2 Facilitated diffusion through carriers 5.2.3 Transport through ion channels 5.2.3.1 Potassium channels 5.2.3.2 Calcium channels 5.2.3.3 Non-selective ion channels 5.2.3.4 Chloride channels 5.2.4 Transport through water channels 5.3 Primary active transport 5.3.1 Primary proton pumps 5.3.1.1 P-type ATPases 5.3.1.2 V-type ATPases 5.3.1.3 The pyrophosphatase 5.3.2 Primary pumps involved in metal transport 5.3.2.1 P-type Ca2+ pumps 5.3.2.2 Heavy metal ATPases 5.3.3 ABC transporters 5.4 Secondary active transport 5.4.1 Potassium uptake 5.4.2 Nitrate transport 5.4.3 Sodium efflux 5.4.4 Non H+ -coupled secondary transport

v 60 61 63 63 64 65 66 66 67 67 67

75 75 76 76 79 81 81 82 83 84 85 85 85 85 87 87 88 89 90 90 90 91 92 92 93 94 95 95

vi

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CONTENTS

5.5 Concluding remarks References

96 96

Regulation of ion transporters ANNA AMTMANN and MICHAEL R. BLATT 6.1 Introduction 6.2 Physiological situations requiring the regulation of ion transport 6.2.1 Change of cell volume 6.2.2 Nutrient acquisition 6.2.3 Stress responses 6.3 Molecular mechanism of regulation 6.3.1 Transcriptional regulation 6.3.2 Post-translational regulation 6.3.2.1 Autoinhibition 6.3.2.2 14-3-3 proteins 6.3.2.3 Calmodulin 6.3.2.4 Cyclic nucleotides 6.3.2.5 Heteromerisation 6.4 Traffic of ion transporters 6.5 Conclusions and outlook References

99

Intracellular transport: solute transport in chloroplasts, mitochondria, peroxisomes and vacuoles, and between organelles ¨ KATRIN PHILIPPAR and JURGEN SOLL 7.1 Introduction 7.1.1 Research to identify solute transport proteins in plant organelles 7.1.1.1 Benefits of a model plant: Arabidopsis thaliana 7.2 Chloroplasts 7.2.1 The function of plastids 7.2.2 Transport across the outer envelope: general diffusion or regulated channels? 7.2.2.1 A porin in the outer envelope of plastids? 7.2.2.2 OEPs, a family of channels with substrate specificity 7.2.2.3 Outer membrane channels and porins: evolutionary aspects in chloroplasts and mitochondria 7.2.3 Transport across the inner envelope: phosphate translocators, major facilitators and carriers 7.2.3.1 The phosphate translocator family

99 99 99 102 106 107 108 109 109 111 113 114 116 117 120 120

133 133 133 134 136 137 137 138 138

142 142 142

CONTENTS

7.2.3.2 Major-facilitator-mediated transport 7.2.3.3 Carriers in the inner envelope of plastids 7.2.4 Transport across the inner envelope: ABC transporters and ion transport 7.2.4.1 ABC transporters 7.2.4.2 Ion transport 7.2.4.3 Transport of metal ions 7.3 Mitochondria 7.3.1 The function of plant mitochondria 7.3.2 Transport across the outer membrane: the porin VDAC 7.3.3 Transport across the inner membrane: carriers 7.3.3.1 Transporters involved in ATP production 7.3.3.2 Carriers for transport of TCA cycle intermediates 7.3.3.3 Amino acid transport across mitochondrial membranes 7.3.3.4 Carriers involved in β-oxidation of fatty acids 7.3.4 Transport across the inner membrane: ABC transporters and ion channels 7.3.4.1 ABC transporters 7.3.4.2 Ion channels 7.4 Peroxisomes 7.4.1 Function of peroxisomes in plant metabolism 7.4.2 Solute transport across the peroxisomal membrane 7.4.2.1 A porin in the peroxisomal membrane 7.4.2.2 Specific transport proteins in the peroxisomal membrane 7.5 Photorespiration: transport between plastids, mitochondria and peroxisomes 7.6 Vacuoles 7.6.1 Generating a pH gradient across the tonoplast: H+ -ATPase and H+ -pyrophosphatase 7.6.2 Transport of malate and sucrose across the tonoplast 7.6.2.1 Malate 7.6.2.2 Sucrose 7.6.3 Aquaporins and ABC transporter in the tonoplast 7.6.3.1 Aquaporins in the vacuole are tonoplast-intrinsic proteins 7.6.3.2 ABC transporters in the tonoplast 7.6.4 Ion transport 7.6.4.1 Ion channels 7.6.4.2 Calcium, sodium and magnesium uptake involves active transport 7.6.4.3 Transport of transition metals References

vii 144 146 147 147 149 150 153 153 154 156 156 158 159 160 160 160 161 162 163 163 163 165 166 167 168 170 170 171 171 171 172 173 173 175 177 178

viii 8

9

10

CONTENTS

Ion uptake by plant roots ROMOLA J. DAVENPORT 8.1 Introduction 8.2 Soil composition 8.3 Root exploration of the soil 8.4 Physical factors affecting root uptake: depletion zones and Donnan potentials 8.5 Radial transport of solutes across the outer part of the root 8.5.1 The role of apoplastic barriers 8.5.2 Root hairs and cortical cells 8.6 Solute uptake from different root zones 8.7 Transport of solutes to the xylem 8.8 The kinetics of solute uptake into roots 8.8.1 Radioisotopic studies 8.8.2 Other methods 8.8.3 Kinetics of uptake in response to solute availability 8.9 Conclusion References Transport from root to shoot SERGEY SHABALA 9.1 Introduction 9.2 Transport of water 9.2.1 Xylem structure 9.2.2 Physics of water flow and evolutionary aspects of conduit development 9.2.3 Water flow between xylem elements: safety mechanisms 9.2.4 Hydraulics of the sap lift: general overview 9.2.5 Driving force for water movement in the xylem 9.2.6 Controversies and additional mechanisms 9.3 Transport of nutrients 9.3.1 General features of xylem ion loading 9.3.2 Ionic mechanisms of xylem loading 9.3.2.1 Potassium 9.3.2.2 Sodium 9.3.2.3 Anion channels 9.3.2.4 Gating factors 9.3.3 Xylem-sap composition 9.3.4 Factors affecting ion concentration in the xylem 9.3.5 Xylem unloading in leaves References Solute transport in the phloem JEREMY PRITCHARD 10.1 Introduction

193 193 193 194 196 197 197 198 201 203 204 204 207 207 209 209 214 214 214 214 216 217 219 221 222 224 224 225 225 226 227 227 228 229 230 231 235 235

CONTENTS

10.2

Phloem anatomy 10.2.1 Sieve tubes 10.2.1.1 Sieve tubes are anucleate 10.2.1.2 Sieve plate blockage 10.2.2 Plasmodesmata 10.2.2.1 Plasmodesmatal structure 10.2.2.2 Plasmodesmatal selectivity 10.3 Phloem composition 10.3.1 Carbohydrate 10.3.1.1 Sucrose 10.3.1.2 Other carbohydrates 10.3.2 Inorganic ions 10.3.2.1 Variation in sieve element composition 10.3.2.2 K+ /sucrose reciprocity 10.3.3 Nitrogen 10.3.4 mRNA 10.3.4.1 Protein metabolism message 10.3.4.2 Structural genes and cell-wall enzymes 10.3.4.3 Interaction with DNA/RNA 10.3.4.4 Carbohydrate metabolism 10.3.4.5 Redox–oxidative stress 10.3.4.6 Amino acid metabolism 10.3.4.7 Transport 10.3.4.8 Interaction with the environment 10.3.5 Proteins 10.3.5.1 Oxidative stress 10.3.5.2 Defence 10.3.5.3 Calcium and sieve element structure 10.3.5.4 Metabolism 10.3.6 Macromolecular trafficking 10.4 Sieve element water relations 10.4.1 Sieve element water relations 10.4.1.1 Sieve element osmotic pressure 10.4.1.2 Sieve element turgor pressure 10.4.2 Flow in the phloem 10.4.3 Phloem loading 10.4.3.1 Symplastic or apoplastic loading? 10.4.3.2 Transporters facilitating apoplastic loading 10.4.3.3 H+ /ATPase 10.4.4 Phloem unloading 10.4.4.1 Evidence for unloading pathway: root tips 10.4.4.2 Evidence for unloading pathway: developing fruits 10.4.4.3 Evidence for unloading pathway: seed coats

ix 236 236 236 237 238 238 238 240 240 240 240 241 241 242 242 243 244 244 245 245 245 245 245 246 246 246 247 247 247 248 248 249 249 249 250 251 251 254 255 257 257 259 259

x

CONTENTS

10.4.5 Resource partitioning through the phloem Exploitation by other organisms 10.5.1 Micro-organisms and viruses 10.5.2 Sap-feeding insects 10.5.3 Plants 10.5.4 Other organisms 10.6 Conclusions References

260 261 261 261 262 262 262 263

Factors limiting the rate of supply of solutes to the root surface ANTHONY YEO 11.1 Introduction 11.2 Supply of nutrients to the root surface 11.2.1 Absence of the nutrient element in the growth medium in any form 11.2.2 Bioavailability of the element 11.2.3 Movement of nutrients towards roots 11.2.4 Homogeneity or heterogeneity (spatial and temporal) in availability 11.2.5 Losses 11.3 Acquisition and uptake of nutrients by the root 11.3.1 Affinity and capacity of transport processes in the roots 11.3.2 Exploration and exploitation of soil volume by roots 11.4 Acquisition of phosphorus 11.5 Protected cropping systems: hydroponics as an example of ‘ideally’ controlled conditions 11.6 Concluding remarks References

275

10.5

11

12

Mineral deficiency and toxicity ANTHONY YEO 12.1 Introduction 12.1.1 Terminology 12.2 Deficiency and efficiency: iron in alkaline soils 12.2.1 ‘Strategy I’: reduction-dependent iron uptake 12.2.2 ‘Strategy II’: phytosiderophores 12.3 Phosphate uptake in soils that are low in phosphate 12.3.1 Cluster roots and root exudates 12.3.2 Mycorrhizal symbiosis 12.4 Toxicity and tolerance–aluminium in acid soils 12.5 Toxicity and tolerance–essential and non-essential metals 12.5.1 Hyperaccumulation 12.5.2 Ion transport in hyperaccumulators 12.5.3 Phytochelatins 12.5.4 Function of hyperaccumulation

275 276 276 276 278 279 279 280 280 282 284 286 287 287 290 290 291 293 295 296 299 299 300 301 303 304 305 306 308

CONTENTS

13

14

xi

12.6 Concluding remarks References

308 309

Water-limited conditions ANTHONY YEO 13.1 Introduction 13.2 Plant responses to reduced water availability 13.3 Mechanisms to reduce water loss: regulation of stomata and regulation of leaf area 13.3.1 Stomatal regulation 13.3.2 Leaf area regulation 13.3.3 Consequences: interaction with leaf temperature 13.4 Mechanisms to maintain water potential gradients: osmotic adjustment 13.4.1 Water potential of drying soil 13.4.2 Osmotic adjustment 13.4.3 Compatible solutes/osmolytes/osmoprotectants 13.4.4 Water movement from protoplast to apoplast in freezing injury 13.5 Mechanisms to acquire more water: root properties 13.5.1 Constitutive formation of deep roots 13.5.2 Facultative formation of deep roots 13.5.3 Root conductance 13.6 Mechanisms to increase water-use efficiency: C4 and crassulacean acid metabolism (CAM) 13.6.1 C4 photosynthesis 13.6.2 CAM 13.7 Gene regulation 13.8 Concluding remarks References

314

Salinity ANTHONY YEO 14.1 Introduction 14.2 External concentration of salt up to about 50 mM NaCl 14.3 External concentration of salt up to about 100–150 mM NaCl 14.4 External concentration of salt above about 150–200 mM 14.5 ‘Molecular’ tolerance 14.6 Cellular tolerance 14.7 Moving on to a cell in a plant 14.8 Salt glands 14.9 Selectivity at the root 14.9.1 Root selectivity for chloride 14.10 Transport from root to shoot 14.10.1 Transport of chloride to the xylem

314 315 318 318 320 321 322 322 323 324 326 326 326 327 327 328 329 331 334 335 335 340 340 341 343 344 345 346 347 347 348 353 353 356

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CONTENTS

14.11 Transport from shoot to root 14.12 Leaf cells 14.13 Prospects 14.14 Concluding remarks References

356 357 361 364 365

Desiccation tolerance ANTHONY YEO 15.1 Introduction 15.2 Occurrence of desiccation tolerance 15.3 Desiccation tolerance in seeds 15.3.1 Intracellular physical characteristics 15.3.2 Intracellular de-differentiation 15.3.3 ‘Switching-off’ metabolism 15.3.4 Antioxidant systems 15.3.5 Protective molecules 15.3.6 Amphiphilic molecules 15.3.7 Oleosins 15.3.8 Damage repair 15.4 Vegetative tissues 15.4.1 Gene expression 15.4.2 Physical characteristics 15.4.3 Metabolism and antioxidants 15.4.4 Low-molecular-weight carbohydrates 15.4.5 Hydrins or LEA proteins 15.4.6 Signals 15.4.7 Constraints to the development of desiccation tolerance 15.5 Concluding remarks Acknowledgements References

371

Index The colour plate section appears after page 78

371 372 372 374 374 375 375 376 378 379 379 379 382 382 383 383 385 385 386 388 388 388 391

Preface Plants generate oxygen, consume carbon dioxide and convert the energy of the sun into food. Life on earth, as we know it, could not have developed and cannot exist without them. Human physical needs are serviced by plant products, from using their molecules as pharmaceuticals to using their bodies for timber. The extent of interest in flowers, gardens and landscapes indicates the psychological importance of plants to us. The acquisition and transport of solutes is fundamental to plant processes at all levels of organisation, and underlies their ability to colonise the land. The purpose of this book is to examine solute transport as a subject in its own right and consider it from the molecular to the ecological and agricultural contexts. Plant cells are full of a vast array of solutes, some in very large quantities. Plants expend considerable amounts of energy and resources upon acquiring or synthesising these solutes that are necessary for the plant’s existence. The need for such quantities arises from the way plants grow and the environments in which they grow. Plants increase in size principally through cell expansion: the volume of individual cells increases over time. To achieve this, water must move into the cell. The structure of plants and plant parts, as well as their rigidity and shape, depends largely upon a hydrostatic skeleton whereby the solution within the cell is contained under pressure by a viscoelastic cell wall. Even the leaves of trees wilt without this. Accumulating and retaining water against hydrostatic pressure requires an opposing force, and this is provided by the osmotic effect of accumulated solutes. This is one reason why the concentration of solutes inside the plant must be much greater than that in the external medium. Plants also face a continual battle to acquire water at least as fast as they lose it. In order to grow, plants must obtain carbon dioxide from the air. The stomatal pores that allow this automatically permit water vapour to pass in the opposite direction from the moist leaf to the usually much drier air. Plant cells must be able to replace this water from the soil as well as compete with the atmosphere to retain some of this water in the plant. Once again, the osmotic forces provided by the accumulation of solutes are the plants’ main weapons. Plants have also evolved in a marine environment where the concentration of inorganic ions was considerable. To avoid dehydration, early plant cells also needed to have an equivalent concentration of solutes. Vital processes such as protein synthesis developed at this time and have since been conserved rigidly. This can explain the requirement plant cells retain for elevated and specific concentrations of inorganic ions in their cytoplasm.

xiv

PREFACE

The marine environment provides abundant and fairly continual replacement of nutrient requirements, but this is not true for all soils. Since plants colonised the land, they have needed to forage and apply strategies to seek and mobilise nutrients from the limited quantities available in many field situations. In addition to these inorganic ions, plant cells contain a vast array of solutes, from small molecules up to proteins and nucleic acids. These are components of cellular biochemistry; as materials, intermediates, products and co-factors in pathways and cycles. Solutes are important for the storage and mobilisation of reserves. The co-existence of all these different processes is dependent upon compartmentalisation by membranes within and between organelles (e.g. vacuoles, chloroplasts and mitochondria), and for this a multitude of transport processes of varying specificity and capacity are required. Directly or indirectly, most of these processes are energy-driven. In plants, the primary energy currency is the proton motive force: proton gradients set up by conversion from high-energy chemical bonds and by the photosynthetic and respiratory electron transport chains. Plant solute transport today has to meet these requirements for the uptake, synthesis and movement around the plant of sufficient quantity and quality of solutes for all of these needs. This must be achieved across the range of ecological conditions in which plants grow, ranging from the relatively sufficient conditions provided in agriculture to those severely limited by the availability of water, of nutrients, and those affected by non-optimal temperatures and by mineral toxicity. This book sets out to provide a coherent coverage of solute transport in plants. The first section covers the physical concepts behind the solute and water movement and the roles of solutes in the plant. The second section covers the transport of solutes at the molecular, cellular, tissue and whole-plant levels of organisation; from the nanometre distances across a membrane to the 100 or more metres required to traverse a tall tree. This section includes a discussion of the membranes that provide the compartmentalisation central to living processes and that allow different cells to perform different functions and different processes to go on within the same cell. The methods of measuring solute transport at different levels of organisation are also addressed. The movement of solutes by pumps, carriers and ion channels is discussed, covering movement from within an organelle to movement around the plant. The two long-distance transport systems – the xylem and phloem – and the forces that drive movement in the two systems link the tissue and whole-plant levels of organisation. The final section of the book examines how solute transport has been adapted in plants growing in a range of conditions from carefully tended horticulture to those of environmental stress. The conflicting priorities of ecological and agricultural adaptation are highlighted. Plant Solute Transport aims to provide an in-depth coverage of this substantial topic, from the molecular to the ecological scale. There is a gap, which we seek to fill, between the large general textbook covering all of plant physiology (perhaps including growth and development and/or biochemistry and molecular biology) and the highly detailed multi-author volume addressing one specific area (such as membrane transport). This volume is directed particularly at research workers and

PREFACE

xv

graduate students, but has a wide enough coverage to be of use to third-year students in plant sciences. The up-to-date research is grounded in the underlying physics and chemistry and placed in the context of what solute transport must achieve for plants in both ecological and agricultural contexts. Anthony Yeo Tim Flowers

Contributors Dr. Anna Amtmann Plant Sciences Group. Division of Biochemistry and Molecular Biology, IBLS, University of Glasgow, Glasgow G12 8QQ, UK Professor Michael R. Blatt Plant Sciences Group. Division of Biochemistry and Molecular Biology, IBLS, University of Glasgow, Glasgow G12 8QQ, UK Dr. Romola J Davenport Oxford Institute of Ageing, University of Oxford, Manor Rd, Oxford OX1 3UQ, UK Professor Tim Flowers Department of Biology and Environmental Science, School of Life Sciences, John Maynard Smith Building, University of Sussex, Falmer, Brighton, BN1 9QG, UK, and School of Plant Biology, 35 Stirling Highway, Crawley, Western Australia 6009, Australia Dr. Matthew Gilliham School of Agriculture, Food and Wine, University of Adelaide, PMB 1, Glen Osmond, South Australia, 5064, Australia Dr. Frans J. Maathuis Biology Department, Area 9, University of York, York YO10 5DD, UK Dr. Katrin Philippar Department Biologie I, Botanik, Ludwig-MaximiliansUniversit¨at, Menzingerstr. 67, D-80638 M¨unchen, Germany Dr. Jeremy Pritchard School of Biosciences, The University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK Dr. Sergey Shabala School of Agricultural Science, University of Tasmania, Private Bag 54, Hobart, Tasmania, 7001, Australia Professor Jurgen ¨ Soll Department Biologie I, Botanik, Ludwig-MaximiliansUniversit¨at, Menzingerstr. 67, D-80638 M¨unchen, Germany Dr. Anthony Yeo Department of Biology and Environmental Science, School of Life Sciences, John Maynard Smith Building, University of Sussex, Falmer, Brighton, BN1 9QG, UK and School of Plant Biology, 35 Stirling Highway, Crawley, Western Australia 6009, Australia. ARY current address: [email protected]

1 General introduction Anthony Yeo

1.1

Introduction

Plant cells are full of solutes, both dissolved inorganic ions and low-molecular-mass organic molecules. The concentration of solutes inside plant cells is higher than that in the growing medium, and it is much higher for the large majority of terrestrial plants. Plants expend considerable amounts of energy and resources upon acquiring or synthesising these solutes, so perhaps the first question to ask is, ‘why do they do it?’ In part the reasons are historical. The salinity of the early oceans was substantially greater than it is today (Knauth, 1998). The conditions in which life evolved are still debated. It is believed that life might have been evolved in situations where freshwater diluted this salinity; however, the great majority of early life arose in the oceans. For simple physical reasons (water flows across their semipermeable membranes influenced by osmotic forces; see Chapter 3), it was necessary for cells to match the water potential of the seas to remain hydrated; so an equivalent concentration of solutes was needed. Some fundamental living processes of cells were laid down during this arcane period – long before life colonised the land. Observation has shown that these processes have been rigidly conserved; for instance the ionic requirements for protein synthesis (see Chapters 3 and 14). The ghost of the past commands the conditions that plants have to maintain in the cytoplasm of their cells today (a hundred or hundreds of mM of solutes). This is even though the concentration of salts in the growing medium may now be of the order of μM and mM, a tiny fraction of that in the present-day or ancient oceans. In part the reasons are physical. The first challenge of life on land was to remain hydrated. As plants evolved from wetlands to dry land, the availability of water became less. Retaining water against the non-osmotic components of water potential became a priority for the first time. The soil was periodically dry, and the cells of plant roots had to retain water against the water potential of the drying soil. In addition to this, the leaves of plants were in a medium that was hardly ever saturated with water, that is the air. The moist surfaces of cells lost water to the demands of the unsaturated air – because of the vapour pressure difference. This has been the nowin situation of plant life on land. The need to acquire atmospheric carbon dioxide for photosynthetic carbon fixation meant that the cells could not be permanently waterproofed – letting in carbon dioxide meant letting out water. Cells not only had to obtain their water from drying soil, but also had to compete with the voracious demands of transpiration – some 98% of water used (see Chapter 3) – and for this they had to depend upon their own osmotic pressure.

2

PLANT SOLUTE TRANSPORT

In part the reasons are structural. Without enough water, plants and even the leaves of trees wilt. Plants still rely largely on a hydrostatic skeleton maintained by turgor pressure; that is the positive hydrostatic pressure that the cell contents exert upon the surrounding structural cell walls (see Chapter 3). Cells use the osmotic component of water potential (hence the dissolved solutes) to build the turgor pressure. Without this, leaves (or large parts of the plant in the absence of the structural thickening found in woody stems) become flaccid. Such leaves are then unable to fulfil the needs of photosynthesis and may be irreversibly damaged. The large majority of plant growth is by cell expansion. In contrast with animals, mature cells of plants contain a large central vacuole (which may be 90% or more of the volume). This is the principal way in which plants generate size, be it to get up into the light or down into the wet soil, or to expand leaves and ramify roots to capture carbon dioxide, water and nutrients. A continual increase on the quantity of solutes is needed to sustain the concentration within the growing cells, without this the turgor pressure would decrease and there would be no growth. For all these reasons, it is a fundamental requirement for survival that plants fill their cells with solutes, whether this is in the form of inorganic ions concentrated from the growing medium or with organic solutes synthesised from sources (of principally: carbon, nitrogen, phosphorus, oxygen and hydrogen) in the atmosphere and soil. Plants need both the major inorganic ions (for instance, potassium, magnesium and nitrates) and the numerous ions that serve the role of specific ‘micronutrients’. On land these resources had to be found from an environment in which they could become rapidly depleted – in contrast to the sea, where, even at low concentrations, there was normally continual replacement. Nowadays, ‘fertigation’ and nutrient film techniques are common in commercial horticulture to prevent such depletion. In the soil, plants must often forage for the materials they need. Overall the flows of water and solutes are locked together in a dance of physical laws. Evapotranspiration causes a mass flow of water through the soil-plant-atmosphere system and the accumulation of salts drives localised flow of water which brings with it dissolved salts. It takes two to tango. The solutes of plant cells and their roles are diverse. Quantitatively, the largest components are dissolved inorganic ions and low-molecular-mass organic molecules. But the term solute also includes compounds of greater molecular mass as components and products of biosynthetic and catabolic pathways and cycles, up to and including soluble proteins and nucleic acids. Not all soluble species always exist in, or are always transported in, solution. Soluble inorganic ions must often be transported anhydrously across the membrane bilayer by protein carriers. Also, there are species that are not soluble in water but are nonetheless transported throughout the plant; for instance insoluble proteins and viral particles. The transport of solutes occurs over a large range of scale, some 10 orders of magnitude, from the order of 10 nm to cross a cell membrane to the order of 100 metres to ascend the tallest tree. The nature of the events and driving forces that underlie transport over such differences in scale are extremely different for the same solute. A potassium ion carried to the top of a tree in the xylem is in solution in water,

GENERAL INTRODUCTION

3

but a potassium ion being transported across a membrane by a carrier is not in solution but is bound reversibly to a transport protein. Movement up the xylem of a tall tree is by a mass flow of solution driven largely by the evaporation of water at the leaf surface, while accumulation across a membrane is driven either directly or indirectly by energy derived from a biochemical process. Membranes provide the compartmentalisation that is central to living processes; allowing different cells to perform different functions and allowing different processes to go on within the same cell. The concentrations of soluble metabolic intermediates of the citric acid cycle within the mitochondrion can be made relatively independent of the concentrations of the same solutes in the cell as a whole. This allows the same solute to be used for different purposes in different parts of the same cell. Extreme examples are the vacuolar compartmentalisation of malate in CAM plants (see Chapter 13) and of salts in halophytes (see Chapter 14); in both cases permitting the retention of concentrations that would destroy the cytoplasm. More generally, compartmentalisation within membrane-bound compartments provides efficiency, allowing high concentrations to exist in one place without the need for the enormous quantities that would be needed to provide the same concentration throughout the cell. The compartmentalisation of protons is universal in plant cells, with pumping out of the cytoplasm both across the plasma membrane to the outside and across the tonoplast into the vacuole. This not only provides the neutral-toalkaline pH needed in the cytoplasm, but the electrochemical potential gradient of protons. In plant cells, it is this proton motive force (PMF) that is used both to store and couple the energy derived from biochemical processes (ATPases and pyrophosphatases, the photosynthetic and respiratory electron transport chains) with the active transport of other solutes.

1.2

Synopsis

There is a wide range of inorganic and organic solutes in plants. Chapter 2 is an introduction to methods for their extraction and analysis. Inorganic elements can be measured by optical properties (by flame emission and atomic absorption spectroscopy), mass spectroscopy, X-ray fluorescence, with ion-specific electrodes, and by ion chromatography. Analysis of organic solutes is usually achieved by chromatographic separation, often in conjunction with mass spectroscopy and nuclear magnetic resonance. Intracellular localisation can be achieved either via transmission or scanning electron microscopy preceded by precipitation, freezing or freezesubstitution. Ion-specific intracellular electrodes can also be used, as can direct sampling using a modified pressure probe. Individual ions can be monitored in cells loaded with fluorescent probes, and tracer fluxes can be interpreted using analysis of compartmental models. Chapter 2 also introduces the roles of solutes in the vacuole, cytoplasm, organelles and cell walls. Chapter 3 begins by describing the properties of water that are important to its behaviour in biological systems: the hydrogen-bonding that confers structure and order, latent heat, thermal capacity, tensile strength, surface free energy (tension) and

4

PLANT SOLUTE TRANSPORT

incompressibility. The large dielectric constant gives water its solvent properties, its ability to perform charge shielding and provide hydration shells, which link to its roles in maintaining the higher order structure of macromolecules. It is difficult to understand how plants acquire and transport solutes without understanding the physical bases of ion and water movement. What are the driving forces? Which way do ions and water ‘want’ to go? How do plants move and accumulate solutes against physical and chemical gradients? Chapter 3 continues with a consideration of Gibbs free energy and chemical potential, water potential and water potential gradients, osmosis and other colligative properties. It includes the derivation of equations for water movement in cells and in the soil–plant–atmosphere system (resistances and the Ohm’s law analogy), and of how surface tension develops negative hydrostatic pressures in drying soils and cell walls. The chapter then moves on to solute movement; diffusion and Fick’s law, and to permeabilities and fluxes. The contribution of electrical charge is explored in the derivation of the Nernst equation, Donnan systems and the Goldman equation. Finally, irreversible thermodynamics is introduced as it applies to the analysis of coupled flows of solutes and solvents. With this background, the subsequent section of chapters (4 to 10) looks at how solutes are moved at individual membranes and, on an increasingly integrative scale, within and between cells and around the plant, both up in the xylem and down in the phloem. Chapter 4 considers the structure and composition of plant membranes – of which there are about 20 types, all comprised of lipids, proteins and carbohydrates in the approximate ratio of 40:40:20. The amphiphatic nature (both hydrophobic and hydrophilic) of lipids underlies the formation of bilayer membranes. These have little intrinsic solute permeability. This is conferred in biological membranes by embedded transporter proteins mediating either active or passive movement and providing varying degrees of regulation. The overall structure of membranes is currently considered to consist of lipid-ordered microdomains, with rather less freedom of movement in the plane of the membrane that was inherent in the first fluid mosaic model. The transport proteins are often multimeric and distributed in membranes in clusters. Techniques for studying solute transport in membranes are discussed next beginning with those applicable to intact (or semi-intact) tissues and moving on to adaptation of these techniques for use with isolated membranes. There is an emphasis on design and composition of experimental solutions, particularly their osmolarity, and on consideration of unstirred layers, and the difference between the study of net transport and unidirectional fluxes. Methods available include inhibitor studies, radioactive tracers, fluorescent probes and electrophysiology – the last including multi-barrelled electrodes. Individual membrane types can be isolated via protoplasts, and sometimes by direct mechanical means, separated by differential centrifugation and identified by marker analysis. Aqueous polymer two-phase isolation provides information regarding sidedness. Analysis can be performed on vesicles or tiny pieces of a membrane attached to a micro-electrode. Techniques such as fluorescence microscopy and patch-clamping can yield considerable spatial

GENERAL INTRODUCTION

5

and temporal resolution, enabling the detection of the activities of single ion channels. Molecular techniques now allow the in silico characterisation of the possible function of membrane proteins where there is sufficient information on available databases. Forward and reverse genetic screens can be used to endeavour to relate gene to function, as can the use of over-expression and expression in heterologous systems (generally in yeasts or in Xenopus laevis oocytes). The location of proteins within the plant and cell can sometimes be determined by expression using reporter gene constructs. For all techniques of investigations there is a compromise between resolution and invasiveness (or distance from physiological reality). The importance of confirming a result obtained with one technique using a different approach, cannot be overstated. The details of transport across membranes is considered for simple inorganic solutes, anions and cations (see Chapter 5). Any membrane protein involved in cross-membrane movement of substrates is defined as a transporter. Transporters can be classified as to whether the event they mediate is active or passive, and if it is active, whether it is a primary process or a secondary one utilising energy already stored in proton gradients. Transporters may also be classified as pumps, ion channels, or carriers – the last includes the provision of passive transport at higher selectivity but lower capacity than ion channels. A further form of classification is that of uniport, symport and antiport. All these terms will be met in different combinations in the literature. Broadly speaking, primary, ATP-driven, pumps set up proton gradients to drive secondary transport. Primary pumps are also directly involved in the transport of calcium and heavy and transition metals. Secondary transport in plants is generally coupled to proton gradients, and participates in the uptake and movement of hundreds, if not thousands, of different substrates. Finally, it is the ion channels that are almost exclusively responsible for passive transport. They mediate only passive transport and are either open or closed, known as gating, which may be regulated by voltage, ligands, or may be mechano-sensitive (e.g. stretch-activated). In addition, there is a selectivity filter that operates on the basis of physical size and charge properties. Channels may be inward- or outward-rectifying according to whether they are permitting passage into or out of the cell. The transport rate of channels may be millions per second. Water movement across membranes is always passive and directed by the gradient in water potential. Water may cross membranes via their intrinsic permeability to water and also through proteinaceous pores: aquaporins. The selectivity of aquaporins is related to size and they have very high capacity (over 109 per second). Primary pumps use chemical, redox or light energy to move solutes against their electrochemical gradient. ATPases have low capacity (around 100 per second), and consequently large numbers of these proteins are required. There are also primary pumps for calcium and some other metals, such as for copper in chloroplasts. Transport rates of primary pumps are hundreds per second. Secondary active transport pumps solutes against their free energy gradient, but the energy derives from coupling to the proton gradient set up by the primary pump(s) and can be either

6

PLANT SOLUTE TRANSPORT

symport (in the same direction as protons) or antiport (in opposite directions). These are often termed carriers, as they are neither primary pumps nor channels. Such carriers have higher selectivity than channels but lower capacity (hundreds or thousands per second). Major nutrients, such as potassium, are taken up through channels at high external concentrations, and by active processes that are induced upon potassium starvation, at low external concentrations. Plants have many transport processes that need to operate in different ways to address different environmental conditions and developmental stages, as well as differences between different cells and tissues. The processes require regulation (Chapter 6), which occurs at several levels, e.g. gene expression, mRNA degradation, protein turn-over, protein activity and membrane trafficking. Regulation involves both positive and negative feedback, and the transporters themselves are both components and targets of signalling pathways (e.g. calcium, auxin and ABA). Chapter 6 considers examples of the regulation of transporters in adaptive processes, the molecular mechanisms underlying transcriptional and post-transcriptional regulation, and the regulation of transporters by membrane trafficking. Regulation of solute transport is required to effect changes in cell volume, both for sustained growth and for the cyclical changes in volume needed in stomatal guard cells for control of stomatal aperture. The pathways leading to co-ordinated regulation of potassium and chloride channels during stomatal closure are examined. High-affinity uptake of nutrients is often induced by deficiency situations, since there may be less costly pathways of uptake when the same nutrient is in abundant supply. Some transporters are induced by a change from high supply to low supply, and some transporters are induced by a change from nil to low supply. Fine-tuning may be via differential regulation of apparently functionally redundant isoforms. Nutrient transport is regulated not only by availability but by the nutrient status of the plant. Transport is also linked to carbon status, and thus is controlled indirectly by environmental factors that affect photosynthesis. Response to many environmental stresses is dependent upon regulation. For example, ‘unwanted’ entry of sodium into the root cells in saline conditions will lead to membrane depolarisation, which will open depolarisation-activated calcium channels leading to a rise in cytoplasmic calcium activity, which is in turn a signal to enhance the activity of the sodium-proton antiport carrier at the plasma membrane, which pumps sodium out again. The limited information regarding the molecular components of the transcriptional regulation of nutrient transporters are summarised. Post-transcriptional regulation involves auto-inhibitory domains, protein–protein interactions (e.g. with protein kinases, calmodulins and 14-3-3 proteins), and ligand binding (e.g. ion channel gating by cyclic nucleotides). The 14-3-3 proteins are highly conserved and regulate a wide range of targets including a number of ion channels. Calmodulins are small calcium-binding proteins that are able to translate intracellular calcium signals into a variety of cellular responses. Cyclic nucleotides are widely used in signal transduction, and evidence is building that higher plants use cGMP as a secondary messenger. Finally, the role of membrane trafficking is reviewed. SNARE (soluble NSF attachment receptor) proteins have been identified in higher plants; they are a group of membrane proteins that are highly conserved

GENERAL INTRODUCTION

7

in eukaryotes and are at the centre of the molecular machinery involved in vesicle trafficking and membrane fusion. Plant processes involve a complex traffic between organelles, and between organelles and the cytoplasm. Organelles have their own transport systems and these are integrated with cellular metabolism (Chapter 7). Chloroplasts are part of the plastid family that includes storage plastids and amyloplasts. They contain the light-harvesting centre and the photosynthetic electron transport chain. Chloroplasts have distinct outer and inner membranes, plus the thylakoid system. The outer envelope (OE) has a range of proteins (OEPs) which are selective channels for solutes essential to plastid function. The inner membrane contains the phosphate translocator family and members of the major facilitator superfamily. There are transporters for di- and tri-carboxylates and carbohydrates, for ATP/ADP exchange, and for a range of specific ions (including nitrate and sulphate which are reduced in the plastid) and there are also symporters for transition metals. Mitochondria are semi-autonomous organelles with a smooth outer membrane and a much-folded inner membrane, which is the energy-transducing membrane. The compartments are the intermembrane space and the protein-rich core, or matrix. One key role of mitochondria is the synthesis of ATP formed by oxidative phosphorylation – the PMF generated by the respiratory chain drives the ATP synthase complex. The outer membrane contains the VDAC porin which is freely permeable to solutes of up to 4–5 kDa: specific permeability barriers reside with the inner membrane. Carriers on the inner membrane include the phosphate carrier, the ATP/ADP carrier, and carriers for intermediates of the tricarboxylic acid cycle, amino acids, and a carrier for succinate/fumarate (which links β-oxidation in the peroxisomes with the TCA cycle). There are also ion channels for potassium and calcium. Peroxisomes are bounded by a single membrane. They are involved in βoxidation and are part of the photorespiratory cycle; they also generate reactive oxygen species and contain appropriate protective mechanisms. Glyoxisomes convert lipid reserves to sucrose. The peroxisome family has a ‘specific porin’ as well as transporter proteins including the peroxisomal ATP/ADP carrier. The photorespiratory pathway is split between the chloroplast, mitochondrion and peroxisome. Vacuoles are multifunctional and are involved in the storage of different metabolites, quantitatively extreme examples being malate (in CAM) and sucrose (in storage tissue). The vacuole is the largest organelle and usually comprises the major volume fraction: it is bounded by a single membrane, the tonoplast. The tonoplast contains proton ATPases and pyrophosphatases which together generate a PMF. A major facilitator imports malate. Tonoplast intrinsic proteins (TIPs, aquaporins) mediate water flow. There are ABC transporters for the accumulation of secondary metabolites and xenobiotics. There are a range of ion channels and carriers mediating the movement of solutes needed for cell expansion, guard cell movement, and compartmentalisation (such as of sodium). Chapter 8 addresses the main factors affecting and controlling the uptake of charged solutes by plants, from the soil solution to the transpiration stream. It

8

PLANT SOLUTE TRANSPORT

describes root anatomical and physiological responses to the availability of nutrients in the soil and the general processes involved in the transport of solutes into and out of root cells. The Casparian strip blocks apoplastic radial movement of water and solutes when it develops, and in many species this barrier is backed up with an hypodermis. Some leakage may occur, particularly when lateral roots are initiated. There is also a symplastic continuity from cell to cell via plasmodesmata. Root hairs are modified epidermal cells that increase surface area and root radius, and appear to be most important in the acquisition of immobile nutrients. In some instances, epidermal cells are modified as transfer cells. The cells of the cortex may be involved in nutrient uptake depending upon whether the epidermal cells can satisfy the needs of the plant and upon whether they have already depleted the concentration to which the cortex is exposed. The tissue and cell expression pattern of high-affinity transporters varies between different nutrients. Cortical cells may also be involved in re-uptake of nutrients that have been effluxed by cells in outer layers of the root. Uptake varies along the length of the root, being minimal at the apex (which is phloem-supplied). Root hairs are usually concentrated behind the apex. Uptake of mobile nutrients may occur along the root but uptake of immobile nutrients is mainly near the tip. Uptake of calcium occurs in young roots only where an apoplastic radial pathway remains available. Xylem loading varies longitudinally, clearly affected by the stage of xylem development. Xylem loading is independent of initial uptake, at least for some solutes. There is evidence that shoot requirements can dictate root uptake and translocation rates. Net uptake is the sum of influx and efflux, and the latter can be a very high percentage of the former. Analysis of tracer uptake is usually related to a three-compartment model: (1) the cytosol of cells of the outer root, (2) vacuoles and (3) transport to the shoot. After the initial uptake, filling of vacuoles and transport to the shoot are in parallel. The kinetics of tracer uptake have been interpreted as dual isotherms since the 1960s. This is now considered to represent the co-existence of low-capacity-high-affinity systems at low external concentration, and high-capacity-low-affinity systems at higher external concentration. These may be either channels or carriers. The xylem has evolved for long-distance upward transport of water and solutes (Chapter 9). The xylem has a large capacity to carry the replacement of transpirational losses and is a leak-proofed conduit with the mechanical strength to avoid collapse under negative hydrostatic pressure. Xylem comprises vessels and tracheids (collectively, tracheary elements, the conducting pathway), fibres and parenchyma (the only living cells in the xylem). The xylem parenchyma cells are densely cytoplasmic with ER, ribososmes and mitochondria. Vessel elements are 5–500 μm (typically 40–80 μm) in diameter and joined end-to-end via perforation plates into vessels that may be several metres long. Tracheids are 10–25 μm and are interconnected by pit fields at their overlapping, tapered ends. The classic interpretation of water movement in the xylem is the cohesion-tension theory. There have also been additional mechanisms suggested which include: mucopolysaccharides to help maintain water flow, osmotic water lifting (root pressure), ionic control of xylem conductance and an electrical driving force.

GENERAL INTRODUCTION

9

The concentration of major solutes in the xylem is mostly in the mM range though these concentrations are variable between species and may depend upon shoot demand. The osmotic pressure of the xylem is usually not considerable. Sampling of the xylem is difficult because most methods are very invasive, though there has recently been use of xylem-feeding insects. Loading of potassium into the xylem is probably via depolarisation-activated outward-rectifying potassium channels. There are three types of anion channels involved in xylem loading, and this is mostly a passive process. Sodium loading could be via a non-selective channel but probably via sodium-proton antiport. Unloading of solutes from the xylem into the leaf is plausibly under hormonal control, and a complex network of veins exists to reduce damage due to excessive concentration of xylem contents when water is withdrawn. Proton ATPases are probably the driving force behind both active and passive unloading, with co-transport processes important, for example, for sugars. The other long-distance transport system in plants is the phloem (Chapter 10). The transport pathway consists of sieve tubes which are an end-to-end arrangement of sieve elements (each 40–500 μm by 5–50 μm) joined at a sieve plate. The plate is perforated by pores and is the major resistance to flow. The sieve tube contains mitochondria but is anucleate. Sieve tubes may live many years and have protection against oxidative damage. The other main component of the phloem is the companion cells which are connected to the sieve tubes via plasmodesmata, through which all proteins destined for the anucleate sieve element must pass from the companion cell. Analysis of sieve tube contents has been made mostly using phloem-feeding insects or by bark incision. The major carbohydrate in most species is sucrose, at hundreds of mM, though in some species the major transported carbohydrate differs; for example, sorbitol, raffinose or mannitol. Potassium and sucrose are the major osmotica and there is a reciprocity between them, maintaining turgor pressure with varying carbon supply. Phloem transports many other nutrients and, recently, the implications of the transport of mRNA and proteins is complementing and revising the understanding of the phloem. Over 200 proteins have been identified, although sieve elements are unable to synthesise proteins themselves. Osmotic pressure in sieve tubes at ground level is generally 1–2 MPa. Phloem is loaded at sources (sites where there is synthesis, as in photosynthesis, or else breakdown of storage compounds) and solutes are removed at sinks (where contents are diluted, metabolised, or stored elsewhere). Turgor pressure differences between sources and sinks underlie the pressure flow hypothesis of bulk movement in the phloem. Solutes move into the sieve element from the companion cell. Entry of solutes into the companion cell can take one of the two routes, apoplastic or symplastic. Much evidence favours the former. Sucrose is loaded by a proton co-transport carrier, powered by the PMF set up by the proton ATPase. Loading of potassium into the sieve element-companion cell complex is important both in the transport of potassium in the phloem and in the regulation of the loading process itself. Aquaporins are also present, as are transporters for the loading of many other substances. Unloading may be by either symplastic or apoplastic routes; this differs with species, organ, and stage of development.

10

PLANT SOLUTE TRANSPORT

In the third section of the book we set out to put this information in an ecological and agricultural context (Chapters 11–15). We describe the factors, other than the transport processes themselves, which limit the supply of nutrients to plants in field conditions and even when growing in carefully tended artificial environments. Next, we look at deficiency and toxicity; some of the ways in which plants have evolved to cope with the ‘not enough’ and ‘too much’ of elements and minerals in their growth environment. We then go on to look at how the use of solutes, both in quantity and quality, has been adapted to more extreme environments: the demands of hot, dry deserts, freezing mountains and saline marshes. All of these entail dealing (by avoidance or tolerance) with some form of externally imposed dehydration. There is also a crucial stage in the life cycle of most plants, the internally controlled dehydration concomitant with seed formation. This is true desiccation tolerance and, while this is common place during reproduction, it is very rare in the vegetative tissues of vascular plants. Many factors, in addition to the properties of the transport processes themselves, affect the rate of uptake of nutrients by plants (Chapter 11). Plants are able to take up nutrients from concentrations that are very low in comparison with those in the soil solution, certainly in fertile soils; except in the case of phosphorus which is commonly at limiting concentrations. Although present, nutrients are not always available: many processes affect the supply of nutrients from the bulk (soil) solution to uptake sites on the roots of plants. These include bioavailability and mobility (the rate of diffusion is impeded by absorption on, and chemical interaction with, the soil). Mass flow of soil solution provides a large-capacity route of nutrient supply, but the contribution of bulk flow to nutrient supply decreases with the decrease in soil water content. There will always be boundary (unstirred) layers around the root in which movement is principally by diffusion. Whenever the flux density of uptake exceeds the flux density of supply, there will be depletion zones around the root, greater than the unstirred layer, across which nutrients must also diffuse. Since diffusion becomes less effective as the distance increases, such supply is commonly limiting, and in many situations the rate of transfer across boundary and depletion zones limits the rate of uptake by the plant. Distribution of nutrients in the soil is also heterogeneous in both space and time, and interception of nutrients also involves roots exploring and exploiting new volumes of soil. Uptake of nutrients depends on both affinity and capacity (flux density) of transport processes. High affinity transporters may provide enough capacity to avoid deficiency of major nutrients and sufficiency of trace nutrients, but are not able to supply the quantitative needs of the plant to support rapid growth. A spectrum of transport processes exists with lower affinity, higher capacity alternatives providing the uptake at higher external concentrations. Concentrations of major nutrients in the xylem are generally in the mM range, and external concentrations in the same range are generally needed to support maximal growth, even in well-mixed solutions, even though K m values for high-affinity transporters are often in the μM range. Maintaining optimal growth in horticulture increasingly relies upon the controlled supply of nutrient solution to the plant in hydroponics, which has many advantages as well as

GENERAL INTRODUCTION

11

some disadvantages. Phosphorus stands out as the major nutrient that is commonly limiting to plant growth in field situations, mainly due to low bioavailability rather than chemical deficiency. A total of sixteen elements are essential to the growth of all plants, and a further four have been demonstrated to be essential in some species (Chapter 12). Plants have evolved mechanisms to maximise uptake of minerals that are in limiting supply. The two strategies for the acquisition of iron in neutral and alkaline soils, where iron is present in quantity but unavailable, are discussed. Plant responses centre either on reduction of ferric to ferrous iron in the soil or on chelation of ferric iron, uptake and subsequent reduction. Another example concerns the acquisition of phosphate, which also includes alteration of conditions in the soil as well as the development of cluster roots and symbiotic associations. Even ‘essential’ elements may be present at external concentrations that can elicit uptake to toxic internal concentrations. This is most widespread for aluminium and manganese in acidic soil. Aluminium is used as an example of how plants can detoxify metals in the soil and tolerate them in the plant. The process commonly involved is chelation, either pre-emptively in the soil by secreting exudates, or within the plant by using chelates combined with compartmentalisation. Toxicity also arises from non-essential elements, particularly transition and heavy metals. Although locally significant, these are rare events both geologically and anthropogenically. Because of this, there will have been little selection pressure to develop specific metal detoxification systems and such tolerance as exists is thought to have arisen from serendipitous recruitment of existing processes (the phytochelatins). A group of plants known as hyperaccumulators achieve enormous concentrations of metals in their tissues. Although phytochelatins, which may have evolved to provide homeostasis for essential metals, can cope with low-level chronic exposure, the hyperaccumulators function by compartmentalisation of metals in the vacuole. There is evidence that high concentrations of metals in the leaves can deter herbivory and this has been advanced as an evolutionary explanation for their extraordinary metal contents. Water availability (Chapter 13) is a major factor in the zonation and distribution of plants, with nearly half of all land being classified as dry land. Terminology around avoidance and tolerance is confusing and difficult, but a three-stage concept of drought has a clear mechanistic and physiological basis. Essentially, these stages are (I) water status can be maintained even with stomata open, (II) stomatal control of water status as water availability decreases, and (III) inability to control water status even with stomata closed. The model also helps clarify the blurred distinction between the agricultural and ecological agendas when it comes to coping with drought: these agendas are often in opposition. Ecological success is linked with survival to complete the life cycle, even if this means slowing down or shutting down as water availability decreases, including pre-emptive adaptations to reduce water usage. Agricultural success is concerned with using as much water as available, and maintaining photosynthesis under drought, in order to produce maximum yield. Plants respond to water deficit in many ways ranging from rapid regulation of stomatal conductance to constitutive anatomical modifications seen in desert species.

12

PLANT SOLUTE TRANSPORT

Reduction in stomatal conductance and leaf area will not only reduce water loss, and conserve soil water, but also will reduce growth and yield. Transpiration is also central to heat dissipation in hot climates, and there are thermal considerations linked intimately with water conservation. Solute accumulation is most often considered in relation to osmotic adjustment. Although osmotic adjustment is clearly important in ecological survival, its role in improving yield in agricultural contexts has been severely challenged. Solutes may also be important as compatible solutes, in drought as in any situation that leads towards reduced hydration, though demonstration of a physiological role requires that compartmentalisation be sufficient. Solute transport underpins the photosynthetic adaptations of CAM and C 4 photosynthesis through the storage and transport of fixed carbon as malate. Both provide substantial increase in water use efficiency, and C 4 photosynthesis is associated with high productivity. Water deficit has been shown to affect the expression of numerous genes. Significantly, the way in which deficit is applied accounts for most of the differences in expression. This underlines the essentiality of applying treatments that are physiologically relevant. Commonly-affected genes were generally involved in downregulation of growth, again emphasising the difference between agricultural objectives and what plants ‘naturally tend to do’. Salinity (Chapter 14) is unusual amongst stresses in that the adapted native flora is not particularly stressed – salinity stress is mainly an agricultural event. Excess salt uptake damages plants in the long term when salt accumulates in the cytoplasm or cell walls, particularly in leaves that are at the end of the line in the transpiration stream. Even halophytes ‘exclude’ most (perhaps 90%) of the salt in the medium, but are well able to manage the remainder. Species that are more salt-sensitive rely on limiting salt uptake and require near-perfect exclusion, close to 98%; this is both very expensive in terms of active extrusion and leaves many questions about achieving osmotic adjustment unanswered. Exclusion is a viable option only at very low external concentrations, and halophytes optimise the regulation of salt transport to the shoot rather than depending on exclusion. However, variation in salt tolerance in crops is usually associated with reduced salt uptake and so is diametrically opposed to the mechanisms that confer salt tolerance in halophytes. Non-selective cation channels, high affinity potassium transporters, and LCT1 have emerged as the potential pathways for sodium entry. In non-halophytes, this (largely unwanted) sodium entry is opposed almost entirely by active extrusion. Despite the damaging consequences, sodium is, in most scenarios, moved actively into the xylem by proton antiport. This is perhaps demanded by the needs of root ion homeostasis. A range of ‘scavenging’ processes (reabsorption and retranslocation) exist to recover excess salt uptake, but are of limited capacity; large capacity would depend on the ability to actively efflux the recovered sodium in situations where efflux is already unable to limit net uptake. Compartmentalisation of salt in the vacuole, together with synthesis and localisation of a compatible solute in the cytoplasm, is central to the tolerance seen in halophytes. Compartmentalisation depends upon minimising leakage across the tonoplast, rather than continually pumping sodium back in.

GENERAL INTRODUCTION

13

Overexpression of sodium-proton antiporters has been reported to increase the salt tolerance of some species, but the evidence is confusing and equivocal. The ion relations cannot be separated from the response to osmotic shock (an artefact of some experimental designs) and further limitations in analyses and experiments compromise interpretation. Nevertheless, there is, theoretically, potential to enhance the tolerance of the more tolerant species by manipulating their ion transport. In this scenario, halophytes will be a source of expertise on how to coordinate ion transport, rather than a source of cherry-picked genes. There is also some potential for minimising sodium influx pathways at the lower end of the salinity spectrum, where osmotic stress is not an issue. The majority of crop species lie, however, in the middle ground, where exclusion-based tolerance takes them further away from the successful halophytes, and this poses a dilemma in plant breeding. The tolerance of desiccation (Chapter 15) is common in the development of seeds and of pollen, but is rare in the vegetative tissues of vascular plants. Desiccation differs from water deficit in a qualitative manner; it means the absence of cytoplasmic water. Water is no longer present to shield charges and the surfaces of macromolecules, and the hydrophobic effect no longer exists; the physical chemistry of the cell is entirely different. Tolerance of desiccation also implies tolerance of the metabolic disruption entailed during de- and re-hydration. Tolerance in orthodox seeds requires tolerance of mechanical damage (shrinkage), metabolic damage, of the desiccated state itself and of rehydration. It is a slow and progressive process requiring the programmed and pre-emptive shut down of the cellular machinery. This depends in a co-ordinated way upon intracellular physical characteristics, de-differentiation of the cell, switching off metabolism, effective antioxidant systems, development of protective molecules (low molecular weight carbohydrates and LEA proteins that can preserve the cytoplasm in the desiccated state), oleosins to surround lipid bodies, and mechanisms for repairing damage. Protective molecules function in water-replacement or in glass-formation (the vitrified state). The failure of recalcitrant seeds to develop desiccation tolerance can be due to a deficit in any one of these mechanisms, in overall co-ordination, or simply be prevented by anatomy. Vegetative tolerance in vascular plants is exemplified in the resurrection plants. Tolerance to desiccation is developed slowly, as in seeds, and so differs from the desiccation tolerance of bryophytes, which is constitutive, and can be moved in and out of quite rapidly. Vegetative desiccation tolerance depends, as in seed development, on physical characteristics that permit shrinkage, on metabolic shut-down and antioxidants, and on solutes, including LEA proteins, that can act in water replacement and vitrification. Desiccation tolerance in both seeds and vascular plants is slow, coordinated and pre-emptive. Competitive advantage rests with the predictability that shutting down is ‘worthwhile’. Whilst this is clear for orthodox seeds, it may limit the advantage of vegetative desiccation tolerance to rare niches. This may help explain the rarity of desiccation tolerance in vascular plants. It may also be the case that few species have the mechanical ability to shrink. Another limitation is how roots dehydrate without damage when in contact with the soil matrix, and this has been little investigated.

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1.3

PLANT SOLUTE TRANSPORT

Concluding remarks

Throughout the book we attempt to link the increasing knowledge of cellular and molecular bases of solute and water movement with the roles that these fulfil in the whole plant under both ideal and stressful conditions – and show how this is dictated by the physical laws that govern solute and water movement. A great deal of plant physiology, indeed the raison d’ˆetre of much research, is concerned with improving plant performance in situations in which they are, in some way or another, ‘stressed’. There are two major themes that come out of this. The first is that response to stress operates at all levels of organisation, requiring co-ordination at the level of the whole plant; whether this is in relation to mineral deficiency, mineral toxicity, water deficit, salinity, or to the need to tolerate desiccation. The second is that there is often a conflict between the ecological and the agricultural advantages. Competitive advantage can be about survival, and conserving resources can be an advantage if it increases the chance of completing the life cycle, even if it means slowing down. Agricultural advantage tends towards brinkmanship; that is, getting the maximum yield depends on taking water use to the limit. What suits agriculture may not work in the wild, and vice versa. There is, understandably, perhaps inevitably, an ever-increasing focus on details, individual processes, individual genes, and individual proteins. It is, however, vital to keep in sight the much wider stage on which the consequences and manipulations of individual processes, and manipulations of those processes, are played out.

Reference Knauth L. P. (1998) Salinity history of the early oceans, Nature 395, 554.

2 Solutes: what are they, where are they and what do they do? Tim Flowers

2.1

Solutes: inorganic and organic

Plants depend on solutes in solution for most of their biochemistry and to develop the turgor pressure necessary for growth and form. In this chapter, approaches to qualitative and quantitative analysis of solutes present in cells and their subcellular compartments are outlined (further experimental details can be found in Chapter 4). For inorganic ions, quantitative analysis can usually be achieved in a single step, while organic compounds mostly have to be separated before their concentration can be determined.

2.2

Analysis of inorganic elements

The solutes that are found in cells are either accumulated from the environment or created within the plant; generally, organic compounds are synthesised while inorganic solutes are acquired from the soil. There is a variety of methods by which inorganic elements can be detected and quantified in plants – in extracts or in plant material that has been vaporised. Analysis broadly depends on one of the following: (a) the optical properties of elements when burning; (b) the mass of the element or its ions; (c) the emission of X-rays (X-ray fluorescence); (d) the use of ion-specific electrodes or (e) the chromatographic separation of ions (ion chromatography).

2.2.1

Obtaining material for analysis

For some analytical techniques, plant material can be used directly either by vaporising it at high temperature (e.g. see Section 2.2.2) or by freezing it at very low temperature (e.g. see Section 2.6.2). In most cases, however, mineral elements are extracted prior to analysis. This is most simply done by heating a plant or plant part in water or dilute acid (e.g. 100 mM acetic acid) – a couple of hours at 80◦ C will extract virtually all of any Group 1 cations (e.g. sodium or potassium) present in that tissue. In order to ensure complete dissolution of mineral elements, the plant material can either be heated in a mixture of concentrated nitric acid and sulphuric acid and the extract diluted for analysis or heated alone (dry ashed) at high temperature (550◦ C) to remove all the organic components before dissolution of the inorganic matter in dilute nitric acid (Humphries, 1956). Some constraints are imposed by the nature of (and impurities in) the acids used to dissolve the samples or by loss of those elements that are volatile below the dry-ashing temperature.

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2.2.2

PLANT SOLUTE TRANSPORT

Optical methods

When vaporised (atomised) in a flame, atoms emit light, as electrons that have absorbed energy fall back into lower energy shells; the wavelength of the emitted light is characteristic of the element and the number of photons emitted is in proportion to the elemental concentration. These characteristics of the light emitted when elements are burned form the basis of one of the simplest means of determining the mineral composition of plants. An extract is converted into a fine mist (nebulised) and blown into a flame (e.g. acetylene burning in air). Elements do not need to be separated: all that is needed is a flame, a monochromator (an optical device that selects the wavelength, generally a diffraction grating) and a detector (e.g. a photomultiplier) to measure how much light is emitted at that specific wavelength. For example, any potassium in an extract will be atomised and emit light at a wavelength of 766.5 nm; the amount of light will be determined by the photomultiplier and hence the concentration of potassium can be estimated. This is flame emission spectrometry (FES). Not all elements emit sufficient light for analysis using FES, but they can still be analysed using their absorption of light when atomised in a flame. In atomic absorption spectrophotometry (AAS), the monochromator and detector are used to determine how much light of a specific wavelength is absorbed after passing through an atomised element. The light is emitted from a lamp that contains the element to be analysed (e.g. potassium), which is heated by a tungsten filament in an atmosphere of an inert gas such as argon. For potassium, the lamp will emit light at wavelengths of 769.9, 766.5, 407.7 and 404.4 nm. The monochromator is used to select one of these wavelengths and the element in the flame will absorb light in proportion to its concentration – and so can be determined quantitatively. Lamps can be obtained for a wide range of elements and although an air/acetylene flame is not hot enough to atomise all elements (e.g. silicon), hotter flames can be obtained by burning acetylene in nitrous oxide. While conventional FES and AAS use a nebuliser to deliver solution into a flame, it is also possible to atomise elements using a hollow graphite rod that is heated electrically to several thousand degrees. This ‘graphite furnace’ (GF) can be used to atomise a solution or solid material without the need for extraction: elemental analysis is by AAS (termed GF-AAS). A further possibility is to spray solution into a stream of argon that flows into a ‘torch’ where the gas stream is heated to about 10 000◦ C using a radio-frequency generator. At this temperature, a plasma forms where the atoms are present in an ionized state. This is known as an inductively coupled plasma or ICP. Elemental composition can be determined using optical emission spectroscopy.

2.2.3

Mass spectrometry

ICP can also be combined with mass spectrometry, where the analysis depends not on optical emission, but on the determination of the mass of elements ionised in the plasma (ICP-MS). The solute can be introduced to the plasma via a nebuliser or directly, without first making a liquid extract, using a laser (laser ablation ICP-MS).

SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO

17

Unlike AAS or FES, which provides information on a single element for each test, ICP-MS can provide information on many elements in a single analysis.

2.2.4

X-ray fluorescence

Elements can also be estimated by their emission of X-rays. Just as light is emitted as electrons return to the ground state after absorbing energy when raised to high temperature, X-rays are emitted when elements are excited in a beam of X-rays (high-energy photons) or high-energy electrons: the process is known as X-ray fluorescence (XRF). This is a very powerful tool for biologists, especially when used in conjunction with an electron microscope (see Section 2.6.2).

2.2.5

Ion-specific electrodes

The estimation of the concentration of elements using ion-specific electrodes is based on completely different properties of materials than those discussed so far. A membrane is used to separate ions and this leads to a difference in voltage across that membrane. The best known of ion-specific electrodes is the pH electrode. In this electrode, the membrane is made of a glass that is permeable to hydrogen ions, but not to other ions. Hydrogen ions diffuse through the glass and come to equilibrium with the external solution; this leads to a difference of voltage across the membrane, which is measured using a high-impedance voltmeter in conjunction with a standard ‘reference’ electrode: the potential difference is directly proportional to the logarithm of the ionic concentration in the external solution (according to the Nernst equation; see Section 3.6.4). Apart from hydrogen ions, there are electrodes whose potential is responsive to the concentration of NH 4 + , Ba2+ , Ca2+ , Cd2+ , Cu2+ , Pb2+ , Hg2+ , K+ , Na+ , Ag+ , Br− , CO 3 2− , Cl− , CN− , F− , I− , NO 3 − , NO 2 − , ClO 4 − , S− and SCN− – none is as specific as the pH electrode and so care has to be taken in the presence of other ions that can also cross the membrane (see also Section 4.4.5.1).

2.2.6

Ion chromatography

This is a form of liquid chromatography where ions in solution are separated by their interaction with a resin. Generally, the ions are detected by the conductivity of the effluent from the column in which the ions have been separated; ion chromatography can be used to detect and quantify anions as well as cations and each analytical run provides information on all the ions that are separated.

2.3

Solute concentrations

The results of analyses generally provide data on the quantity (e.g. in grams or moles) of a substance extracted from a known mass of fresh or dry weight. From these data it is possible to calculate the content per plant or plant part or the concentration expressed per unit dry mass or fresh mass or mass of water. However, it is not

18

PLANT SOLUTE TRANSPORT

always easy to determine the concentration in situ as this depends on knowing not only how much of the solute is present but also how much water is present. There is also a question of whether that water behaves as free liquid water or is influenced by macromolecules that are present. If water exists in microdomains of rapidly exchanging regions of high and low density (Wiggins, 2001) with different solvent properties and if these domains are influenced by surfaces and solvents, then the effective ion concentrations are likely to be different from those calculated from estimates of total quantities of solutes and water. Although concentrations can be estimated, effective ion concentrations or activities (the parameter that determines the reactivity and movement of an ion in solution) generally remain unknown, except where they have been measured directly using an ion-specific microelectrode (although there are computer programs that allow their estimation; see Section 4.4.1.1). In a dilute solution (say less than 10 mM) containing a single solute, the solute interacts primarily with the solvent (water), but as solutions become more concentrated, there is an increasing solute–solute interaction and this affects the properties of the solution – both the ‘effective’ concentration of the solute and the properties of the solvent. The effective concentration of the solute is described by its activity. The activity (a) of a solute j (the units remain mM) is a function of the concentration (c, mM) such that: a j = λ j cj

(2.1)

where γ j is the activity coefficient of the solute j. In solutions, cations exist with anions and it is the mean activity coefficient (λ ± ) that is estimated. Activity coefficients are generally less than 1, and especially so for charged solutes such as ions. For example, at 25◦ C a 100 mM solution of KCl has a mean molal activity coefficient of 0.77 (Robinson and Stokes, 1959). In a mixed solution of 100 mM monovalent cations and anions and 25 mM divalent cations and anions – the sort of solution that might occur in plant cells – the mean molar activity coefficient for the monovalent ions is just 0.7 (see Chapter 3 in Nobel, 2005).

2.4

Organic compounds

It is generally more complex to separate and quantify organic compounds than inorganic ions, since there are so many organic molecules in so many different classes – for example, sugars, sugar alcohols, amino acids, organic acids and proteins. In general, these compounds have to be separated chromatographically and then, once separated, there is a variety of methods that can be used to detect individual compounds. Separation may be achieved by liquid or gas chromatography, which may be used in conjunction with mass spectrometry or nuclear magnetic resonance (NMR). In recent years, increasing use has been made of NMR to analyse the composition of plant material in vivo or in extracts. NMR occurs when atoms with ‘spin’ (a fundamental property of nature like electrical charge) are exposed to an oscillating magnetic field while held in a powerful stationary magnetic field. For the analysis of plant metabolites, important atoms that exhibit spin are 1 H, 2 D, 13 C, 14 N, 23 Na and

SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO

19

31

P (Krishnan et al., 2005; Mesnard and Ratcliffe, 2005); 23 Na has recently been used to map the distribution of sodium in stem tissues (Rokitta et al., 2004).

2.5

Range of solutes found in plants

Solutes that are found in plant cells can be broadly divided by function (see Section 2.7) as well as whether they are inorganic or organic. For all plants, N, P, S, K, Mg, Ca, Fe, Mn, Zn, Cu, B, Mo, Cl and Ni are recognised as ‘essential’, while there is discussion about Na and Si (see also Reuter and Robinson, 1997; Chapter 12). Amongst the essential elements, some (such as Mo and Ni) are required in low quantities for specific biochemical functions (e.g. nitrate reductase requires Mo and urease Ni); others (such as S, P, N, Mg, Ca and K) are present in much higher quantities (tens of thousands of times the quantities of Ni and Mo; see Marschner, 1995). It is, however, extremely difficult to generalise about the concentrations of inorganic ions in plants, because plant species and habitats vary dramatically. For example, most plants growing in normal soils have very little sodium in their leaves, perhaps tens of micromoles per gram dry weight, but for plants growing in saline soils the concentration of Na can reach several millimoles per gram dry weight (see Chapter 14). As already mentioned, organic solutes are extremely diverse; they make up the major metabolites found in all cells (e.g. the sugars and sugar phosphates of glycolysis, the Krebs cycle and the Calvin cycle) and secondary metabolites (which vary greatly between species and tissues), as well as hormones (e.g. indole acetic acid, abscisic acid, gibberellins and kinins), storage compounds (sugars such as sucrose in many species) and osmoprotectants (such as glycine betaine; see, e.g., Chapter 13). More details of some of these groups will be found in later chapters of this book; this chapter continues with a summary of how the localisation of solutes in cells can be discovered.

2.6

Localisation

Plants are visibly complex when viewed under a microscope, their different cells having clearly different appearances and ultrastructure: the meristematic cells of the growing plants have dense cytoplasmic contents, while the cells in leaves contain chloroplasts and the parenchymatous cells of the root cortex appear dominated by large, apparently empty, vacuoles.

2.6.1

Stereological analysis

Plant cells, all of which are bordered by cell walls, contain separate compartments enclosed by membranes. Knowledge of the volume of these cellular compartments is necessary in order to determine the distribution and concentration of solutes in cells. The relative volume of an individual cellular compartment can be estimated by stereological methods that use two-dimensional images to generate quantitative

20

PLANT SOLUTE TRANSPORT

estimates of relative volume (Weibel et al., 1966; Russ and Dehoff, 2001). The basis of the analysis is the computation of the area of the whole cell and its compartments from the two-dimensional image: estimates of area are generally made by counting intersections of the cell and its organelle(s) with a grid or a random set of points. In mature cells, the vacuole is the dominant compartment, while in photosynthetic cells the chloroplasts form the dominant subcompartment within the cytoplasm. For example, 72.5% of the volume of the mesophyll cells of Suaeda maritima is occupied by the vacuole; the remainder of the volume is made up by the chloroplasts (12.7%), the rest of the cytoplasm (9.8%), the cell wall (4.3%) and the mitochondria (0.6%) (Hajibagheri et al., 1984).

2.6.2

Inorganic elements and electron microscopy

Some elements are electron dense and can be seen directly in an electron microscope, although it is not possible to distinguish between different electron-dense elements without further analysis. However, since electron microscopes produce focused beams of high-energy electrons that cause X-rays to be emitted from the specimens being examined, a microscope fitted with an appropriate X-ray detector (of the energy or wavelength) can be used to identify and quantify the elemental composition of the specimen. X-ray microanalysis can be undertaken with both scanning and transmission microscopes and software has been developed that can be used to map elements within cells viewed with the microscope (Morgan et al., 1999). A vital aspect of all such studies, however, is an ability to fix the element under consideration in its natural site in the cell. It is pointless to analyse the distribution of a solute if the very process of analysis has caused the substance to be redistributed. Solutes can be fixed either by precipitation – that is a chemical reaction to make an insoluble compound – or by freezing. A good example of the former approach is the use of sodium cobaltinitrite (a soluble salt) to precipitate potassium cobaltinitrite (an insoluble salt) in guard cells (Willmer and Fricker, 1996). The insoluble potassium salt can be visualised under a light microscope by conversion to cobalt sulphide; the cobalt in the cobaltinitrite is electron dense and so directly visible with an electron microscope. Using this technique it has been possible to demonstrate the movement of potassium ions during stomatal opening and closing (Willmer and Fricker, 1996; see also Section 6.2.1). Chloride can be localised in plant cells by precipitation with silver, which is also electron dense. As a preparative technique to fix solutes in their natural compartments, rapid freezing is more versatile than chemical precipitation for both scanning and transmission electron microscopy. The aim of cryopreservation is to freeze samples so rapidly that damage is not caused by the formation of ice crystals; necessary rates of freezing are around 10 000◦ C/s. Commonly, material is frozen at about −186◦ C, by direct immersion in melting or liquid nitrogen. However, this allows gaseous nitrogen to form between the coolant (liquid nitrogen) and the sample. The low thermal conductivity of the gas dramatically reduces the rate of freezing. It is better to cool a solvent such as 2-methyl butane (containing 8% methylcyclohexane to lower its freezing point) with liquid nitrogen (Harvey et al., 1976) as this avoids the

SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO

21

formation of gaseous nitrogen around the sample. Other more elaborate methods of freezing are to use a cold metal block, a jet of propane or high pressure (see chapters in Hajibagheri, 1999). High-pressure freezing (sometimes called ‘slam freezing’) uses a pressure of about 200 MPa to minimise the formation of ice crystals during freezing and hence enhance cryopreservation of tissue that is more than a monolayer of cells (see Hohenberg et al., 2003, and articles in the special issue). Frozen material can be used directly in both transmission and scanning microscopes with a cold stage. In the latter, images of the surface are produced and that surface can be obtained by sectioning prior to freezing or by fracturing the sample in the microscope (freeze fracture) or by milling. Milling can be achieved by a beam of gallium ions produced in the microscope and is used to produce a flat surface (focused ion beam or FIB milling; see Drobne et al., 2005a,b) and avoid artefacts of analysis produced by the topography of an unmilled surface. There is also a variety of techniques to remove water from the specimen before it is introduced into the electron beam. For scanning electron microscopy, water can be removed from the sample by freeze drying. Alternatively, samples can be dehydrated in acetone or ethanol before being subjected to ‘critical point drying’ to minimise distortions due to the effects of surface tension (see Hall, 1978); at the ‘critical point’, surface tension is reduced to zero. For analysis of thin sections by transmission electron microscopy, it is vital to minimise solute movement during the embedding process. Before embedding, frozen material is transferred, in the presence of a dehydrating agent, to an organic solvent – ether, ethanol or acetone – that is miscible with the resin to be used. The solvent is maintained below the freezing point of water, which is removed from the tissue to the dehydrating agent and substituted by the organic solvent – hence ‘freeze substitution’. Once substitution is complete, a suitable resin can be added and polymerised by the action of ultraviolet light. Cutting sections is perhaps the most difficult stage of the procedure, because the sections cannot be floated onto water without the loss of water-soluble elements. Consequently, sectioning has to be done dry – in the absence of water (organic solvents that will not dissolve the substances to be analysed do not have an appropriate surface tension to enable the sections to be floated away from the knife).

2.6.3

Ion-specific microelectrodes

Minature specific-ion electrodes can be used to report the activities of certain ions (such as H+ , K+ , Na+ and Ca2+ ; see also Section 2.2.5) within cells. Since the majority of the cell volume is occupied by the vacuole, measuring ion activities in this compartment is relatively simple. However, measuring ion activities within the cytoplasm of a mature plant cell is significantly more difficult. It is hard to locate an electrode, whose tip has a diameter of about 1 μm, in a cytoplasm that is only 1–2 μm wide. Electrophysiologists have, however, developed a way of identifying the position of an electrode tip using measurements of pH (Felle, 1993; Walker et al., 1998). Using pH electrodes, it has been established that the vacuole is considerably more acidic (pH about 5; Felle, 2005) than the cytoplasm (pH of about 7.2). By

22

PLANT SOLUTE TRANSPORT

constructing double- and triple-barrelled electrodes where one of the electrodes reports the activity of hydrogen ions, this allows the location of the electrode to be identified from the pH, and hence the other element(s) to be estimated in cytoplasm and/or vacuole. For example, using triple-barrelled electrodes reporting pH and the activities of sodium or potassium, sodium and potassium activities in barley could be separated into two populations (Carden et al., 2003), one having a mean pH of 5.6 (vacuolar) and the other a mean pH of 7.4 (cytoplasmic pool).

2.6.4

Direct sampling

Since the central vacuole comprises between 75 and 95% of the volume of a mature plant cell, its contents can be estimated simply by the analysis of sap expressed from tissues. Vacuolar sap is easily obtained from plant cells following one or more cycles of freezing and thawing, which makes the membranes surrounding the cell (the plasma membrane) and its vacuole (the tonoplast) leaky (see, e.g., Gorham et al., 1984). Centrifugation or simply pressure from a glass rod will extract vacuolar sap. Once a clear solution has been obtained, it can be subjected to any of the many methods available to determine its solute content. This sap will be contaminated by solutes from other compartments, but the contamination will be trivial in the context of any analysis that is undertaken to evaluate the major solutes that are stored in the vacuole. It has also proved possible to isolate intact vacuoles from plant cells (e.g. Leach et al., 1990) and these can be analysed for their contents, although exchange of solutes across the vacuolar membrane during the isolation cannot be ruled out. For mature cells, it is also possible to sample the vacuolar sap directly, using a microcapillary as a syringe. The procedure (Malone et al., 1989) was developed from a technique used to determine the turgor pressure within cells – the pressure probe. A glass capillary is drawn to an outer tip diameter of about 4 μm and filled with oil and connected to a pressure transducer. When used for microsampling, the turgor pressure in the cell is allowed to force vacuolar contents into the micropipette; the samples obtained can be used for determination of their osmotic pressure or for elemental analysis using XRF (Fricke et al., 1994). As already noted, extraction procedures are subject to contamination, and where a compartment is small in relative volume, contamination is an all-important issue. While chloroplasts can be extracted non-aqueously from plant cells, thus retaining water-soluble solutes, the binding of solutes from other compartments to the chloroplast envelope is an important source of potential contamination. Interestingly, chloride is retained in the chloroplasts even after aqueous preparation (see Flowers, 1988), suggesting that the envelope keeps its ability to retain solutes during preparative procedures developed to maximise biochemical activity. Further information on the solutes of subcellular organelles can be found in Chapter 7.

2.6.5

Use of fluorescent dyes

There are many dyes whose properties change depending on their environment: a simple example is the colour change of a pH indicator as the concentration of hydrogen ions changes. Similar properties can be used in conjunction with light microscopy to identify the location and concentration of solutes within cells. For

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23

example, there are a number of dyes whose fluorescence changes as a function of the concentration of minerals in their environment (see Section 4.4.4). Such dyes, generally, have to be injected into cells, but once present they can be illuminated with a specific wavelength of light which excites their fluorescence. So, fura-2 can report the concentration of calcium within a cell and sodium- and potassium-binding benzofuran isophthalates (SBFI, sodium-binding benzofuran isophthalatecan and PBFI, potassium-binding benzofuran isophthalate) report the concentrations of sodium and potassium. SBFI appears to provide a reasonable estimate of cytosolic Na, but Na interfered with the use of PBFI in root hairs exposed to external sodium chloride (Halperin and Lynch, 2003).

2.6.6

Flux analysis

The three major compartments of plant cells are, from the outside looking in, the cell wall, the cytoplasm and the vacuole. The cytoplasm and the vacuole are delimited by membranes that represent a resistance to the movement of solutes – the plasmalemma and the tonoplast, respectively. Solutes entering or leaving the cytoplasm have to cross only the plasma membrane; solutes entering or leaving the vacuole must cross both membranes. Flux analysis is based on following the movement of a labelled (generally labelled with a radionuclide) solute in a system which is otherwise at equilibrium. Analysis of the movement of the label provides information on the characteristics of the membranes and the compartments they define. In practical terms, tissue is equilibrated with a solution containing the solute under investigation (e.g. KCl) containing a tracer of known specific activity (specific activity is the ratio of labelled to unlabelled species; in the case of KCl, the tracer could be either 42 K or 36 Cl). After a sufficient period of time for the specific activity of the labelled solute to become uniform in the tissue (this may be hard to achieve and remain uncertain for the vacuole), the efflux of the tracer into an unlabelled, but otherwise identical, external solution is followed with time. This can be achieved by transferring the labelled tissue into fresh unlabelled solution at, say, 1, 2, 4, 8, 15, 30, 45, 60, 90, 120, 150 and 180 minutes after the initial removal from the labelled solution. The amount of tracer in the solutions is determined and that remaining in the tissue at the various sampling times calculated. The data can be analysed according to a model that assumes efflux occurs from three compartments (extracellular spaces, ECS; cytoplasm, C and vacuole, V). This three-compartment model assumes efflux of tracer is composed of three first-order rates of loss of activity superimposed on one another. For loss from the ECS, C and V: ln c = −kt + z

(2.2)

where c is the concentration, t is time, k is a rate constant and z is a constant from an integration. Thus the relationship between the logarithms of the concentration of ions in the tissue against t should be linear. The logarithm of the amount of isotope remaining in the tissue at various times is plotted against time and a straight line fitted to the linear portion of the curve: this represents the efflux from the vacuole (Figure 2.1). The total efflux is then corrected for efflux from the vacuole to provide a linear relationship for flux from

24 Logarithm of remaining activity

PLANT SOLUTE TRANSPORT

6.2

5.7

5.2 0

50

100

Logarithm of remaining activity

A

6 5.5 5 4.5 4

0

5

10

15

0.5

1 1.5 Time (minutes)

20

25

Logarithm of remaining activity

B

C

6

5.5

5 0

2

2.5

Figure 2.1 Flux analysis of 86 Rb-labelled potassium in maize roots. Maize roots (of 1 cm lengths) were immersed in a solution containing 1 mM KCl and 0.1 mM CaCl 2 , labelled with 86 Rb for 3 h. The labelled roots were rinsed rapidly to remove surface film and immersed in a series of unlabelled, but otherwise identical, solutions for a total period of 105 min, with solutions changed rapidly (each minute) at first. Radioactivity in the eluants, and remaining in roots at the end of the period, was determined by Cerenkov radiation. The logarithm of the remaining activity was plotted against time (A), showing a final, linear, phase interpreted as corresponding to efflux from across the tonoplast. This was subtracted and the data replotted (B), revealing a second linear phase interpreted as corresponding to efflux across the plasma membrane. Repeating the procedure again (C) revealed a third linear phase, corresponding to exchange of the extracellular spaces. Rate constants for exchange can be calculated from fitted linear regressions and the potassium content of the different compartments estimated.

the cytoplasm and, finally, efflux from the extracellular spaces, which includes the cell wall (see Flowers and Yeo, 1992, and Section 4.4.3). The data can provide information on the rate constants for loss from the compartments and, provided the compartment reaches the specific activity of the external solution, the content of the compartment. The analysis of the movement of radioactive tracers can also be used to determine the characteristics of influx. Recently, short-term experiments have been

SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO

25

used to provide information on the influx of 22 Na into cells. Roots of arabidopsis were exposed to sodium chloride and the unidirectional influx, estimated from measurements taken over a period of 2 min (Essah et al., 2003).

2.6.7

Organic compounds

As with the identification of organic solutes, their localisation in cells is much more difficult than with inorganic solutes. Although the elements C, N and O can be detected and mapped in electron microscopes, the great variety of possible compounds makes the analysis of specific molecules difficult. Consequently, specific reactions must be made to take place in situ. In some cases a stain or an antibody can be reacted with a specific organic compound and visualised either by its colour, its fluorescence when viewed under a fluorescence microscope or, in the case of antibodies, by the attachment of electron-dense gold particles to the antibody (see Hajibagheri, 1999). In most published cases, the technique has been used to localise proteins: recent examples are the localisation of isoprene synthase in poplar (Schnitzler et al., 2005) and proteins associated with the spread of beet necrotic yellow vein virus (Erhardt et al., 2005). The localisation of smaller solutes has rarely been attempted, although it has proven possible to localise glycine betaine to the cytoplasm of the cells of the salttolerant plant Suaeda maritima. By reacting iodoplatinic acid with glycine betaine in situ, an electron-dense deposit was formed that could be shown to contain both iodine and platinum (the reaction was carried out under conditions in which the solute was retained in the tissue by preparation using freeze substitution; Hall et al., 1978). The localisation of organic compounds can also be determined following cell fractionation in aqueous or non-aqueous conditions although the number of compartments that can be defined by this process is limited to nuclei, chloroplasts and mitochondria (e.g. Riens et al., 1991; Farre et al., 2001). As mentioned previously (Section 2.6.4), contamination by binding of substances to the external membranes of organelles is a potential problem, as is leakage through the same membranes.

2.7

What do they do?

The function of a solute in a cell depends, in part, on the particular solute and, in part, on its location. Clearly it is not possible to ascribe here a function to the myriad of solutes that occur in plant cells.

2.7.1

Vacuoles

Vacuoles are the largest compartments in mature plant cells. All cells are derived from meristematic cells and generally undergo a massive expansion of the vacuole during development. Meristematic cells have a volume of about 0.5–8 pL (a picolitre is 10−12 of a litre), and as development proceeds, cell volumes increase and reach values between 50 and 5000 pL. The vacuoles are the sites of storage of most (in terms of quantity – that is volume times concentration) of the solutes present in mature cells.

26

PLANT SOLUTE TRANSPORT

A particularly important aspect is the storage of energy in the form of a reservoir of protons within the vacuole. These protons can be exchanged in the acquisition of sucrose or other mineral elements. A large number of plant extracts have been analysed since the 1880s, primarily to obtain estimates of the osmotic pressure within cells: these range from fractions of an MPa to 5 or more MPa (Steiner, 1939; Flowers et al., 1977). The accumulation of solutes in the vacuole is essential for the generation of turgor pressure in cells (see Section 3.2.3), which in turn provides the shape and form of non-woody species and the driving force for the growth in all plants. Many plant extracts have also been analysed for the nature of the solutes present; the major components of vacuoles are sugars, potassium, calcium, magnesium and nitrate ions. However, since there can be large differences in the composition of sap between plants, depending on the species and the environment in which the plant is growing, the analysis is valuable only in context. For example, in the roots of beet, it is sugars that constitute the major solutes stored for the generation of energy for new growth after winter. Where plants are growing on saline soils, the sap is commonly dominated by sodium and chloride ions whose accumulation adjusts the plant water potential (see Section 3.2.2) to that of the external medium. In plants utilising crassulacean acid metabolism (see Section 13.6.2), malic acid may be present in high concentrations, particularly at the end of the night.

2.7.2

Organelles and the cytoplasm

The cytoplasm and its constituent organelles are the sites of enzymes (proteins), which require relatively high (about 100 mM) concentrations of ions for their stability and activation. Potassium plays a particularly important role in the activation of enzymes (Leigh and Wyn Jones, 1984), while other elements such as zinc and magnesium are important co-factors in many enzymatic reactions (Marschner, 1995). This specificity contrasts with the osmotic role which can often be fulfilled by a variety of solutes. During water deficit, adaptation to low temperature and in plants adapted to growth under saline conditions, specific solutes can be synthesised to act as osmoprotectants and cryoprotectants (cf. Section 13.6); some of these at least (e.g. glycine betaine) are likely to be located largely in the cytoplasm.

2.7.3

Cell walls

Cell walls occupy about 3% of the total cell volume and of that volume about 1% is water-available space (Flowers and Yeo, 1986). This means that small changes in ion concentrations in that compartment can have a disproportionately large effect on the water relations of the cell (Flowers and Yeo, 1986) and are able to alter turgor (Clipson et al., 1985).

2.7.4

Conclusions

Plant cells depend on solutes for their turgor pressure and growth. Growth requires the uptake of water driven by gradients in solute concentration. Solutes are also

SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO

27

required as substrates for reactions that generate the physical substance of cells. Solutes are at the centre of plant life. Advances in chemical analysis have allowed many of the solutes in cells to be identified and their location discovered and led to the description of the ‘proteome’ (the proteins in cells), the ‘metabolome’ (the profile of small metabolites in cells) and the ‘ionome’ (the ions in cells; Salt, 2004). Understanding how plants respond to changes in their ionome is at the heart of this book.

References Carden, D.E., Walker, D.J., Flowers, T.J. and Miller, A.J. (2003) Single-cell measurements of the contributions of cytosolic Na+ and K+ to salt tolerance. Plant Physiology 131, 676–683. Clipson, N.J.W., Tomos, A.P., Flowers, T.J. and Wyn Jones, R.G. (1985) Salt tolerance in the halophyte Suaeda maritima (L.) Dum.: the maintenance of turgor pressure and water potential gradients in plants growing at different salinities. Planta 165, 392–396. Drobne, D., Milani, M., Zrimec, A., Berden Zrimec, M., Tatti, F. and Draslar, K. (2005a) Focused ion beam/scanning electron microscopy studies of Porcellio scaber (Isopoda, Crustacea) digestive gland epithelium cells. Scanning 27, 30–34. Drobne, D., Milani, M., Zrimec, A., Leser, V. and Berden Zrimec, M. (2005b) Electron and ion imaging of gland cells using the FIB/SEM system. Journal of Microscopy-Oxford 219, 29–35. Erhardt, A., Vetter, G., Gilmer, D., et al. (2005) Subcellular localization of the triple gene block movement proteins of beet necrotic yellow vein virus by electron microscopy. Virology 340, 155–166. Essah, P.A., Davenport, R. and Tester, M. (2003) Sodium influx and accumulation in Arabidopsis. Plant Physiology 133, 307–318. Farre, E.M., Tiessen, A., Roessner, U., Geigenberger, P., Trethewey, R.N. and Willmitzer, L. (2001) Analysis of the compartmentation of glycolytic intermediates, nucleotides, sugars, organic acids, amino acids, and sugar alcohols in potato tubers using a nonaqueous fractionation method. Plant Physiology 127, 685–700. Felle, H.H. (1993) Ion-selective microelectrodes – their use and importance in modern plant-cell biology. Botanica Acta 106, 5–12. Felle, H.H. (2005) pH regulation in anoxic plants. Annals of Botany 96, 519–532. Flowers, T.J. (1988) Chloride as nutrient and osmoticum. In: Advances in Plant Nutrition, Vol. 3 (eds L¨auchli, A. and Tinker, B.), pp. 55–78, Praeger, New York. Flowers, T.J., Troke, P.F. and Yeo, A.R. (1977) The mechanism of salt tolerance in halophytes. Annual Review of Plant Physiology 28, 89–121. Flowers, T.J. and Yeo, A.R. (1986) Ion relations of plants under drought and salinity. Australian Journal of Plant Physiology 13, 75–91. Flowers, T.J. and Yeo, A.R. (1992) Solute Transport in Plants. Blackie Academic and Professional, London. Fricke, W., Leigh, R.A. and Tomos, A.D. (1994) Epidermal solute concentrations and osmolality in barley leaves studied at the single-cell level – changes along the leaf blade, during leaf ageing and NaCl stress. Planta 192, 317–323. Gorham, J., McDonnell, E. and Wyn Jones, R.G. (1984) Salt tolerance in the Triticeae: Leymus sabulosus. Journal of Experimental Botany 35, 1200–1209. Hajibagheri, M.A., Hall, J.L. and Flowers, T.J. (1984) Stereological analysis of leaf cells of the halophyte Suaeda maritima (L.) Dum. Journal of Experimental Botany 35, 1547–1557. Hajibagheri, M.A.N.. (ed.) (1999) Electron Microscopy Methods and Protocols. Humana Press, Totowa, NJ. Hall, J.L.. (ed.) (1978) Electron Microscopy and Cytochemistry of Plant Cells. Elsevier/North Holland Biomedical Press, Amsterdam. Hall, J.L., Harvey, D.M.R. and Flowers, T.J. (1978) Evidence for the cytoplasmic localization of betaine in leaf cells of Suaeda maritima. Planta 140, 59–62.

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Halperin, S.J. and Lynch, J.P. (2003) Effects of salinity on cytosolic Na+ and K+ in root hairs of Arabidopsis thaliana: in vivo measurements using the fluorescent dyes SBFI and PBFI. Journal of Experimental Botany 54, 2035–2043. Harvey, D.M.R., Hall, J.L. and Flowers, T.J. (1976) The use of freeze-substitution in the preparation of plant tissues for ion localisation studies. Journal of Microscopy 107, 189–198. Hohenberg, H., Muller-Reichert, T., Schwarz, H. and Zierold, K. (2003) Special issue on high pressure freezing – foreword. Journal of Microscopy-Oxford 212, 1–2. Humphries, E.C. (1956) Mineral components and ash analysis. In: Modern Methods of Plant Analysis, Vol. 1 (eds Paech, K. and Tracey, M.V.), pp. 468–502. Springer, Berlin Krishnan, P., Kruger, N.J. and Ratcliffe, R.G. (2005) Metabolite fingerprinting and profiling in plants using NMR. Journal of Experimental Botany 56, 255–265. Leach, R.P., Wheeler, K.P., Flowers, T.J. and Yeo, A.R. (1990) Molecular markers for ion compartmentation in cells of higher plants. I. Isolation of vacuoles of high purity. Journal of Experimental Botany 41, 1079–1087. Leigh, R.A. and Wyn Jones, R.G. (1984) A hypothesis relating critical potassium concentrations for growth to the distribution and functions of this ion in the plant cell. New Phytologist 97, 1–13. Malone, M., Leigh, R.A. and Tomos, A.D. (1989) Extraction and analysis of sap from individual wheat leaf cells: the effect of sampling speed on the osmotic pressure of extracted sap. Plant, Cell and Environment 12, 916–926. Marschner, H. (1995) Mineral Nutrition of Higher Plants. Academic Press, London. Mesnard, F. and Ratcliffe, R.G. (2005) NMR analysis of plant nitrogen metabolism. Photosynthesis Research 83, 163–180. Morgan, A.J., Winters, C. and Sturzenbaum, S. (1999) X-ray microanalysis techniques. In: Electron Microscopy Methods and Protocols (ed Hajibagheri, M.A.N.), pp. 245–276. Humana Press, Totowa, NJ. Nobel, P. (2005) Physiochemical and Environmental Plant Physiology. Elsevier Academic Press, Amsterdam. Reuter, D. and Robinson, J.B., (eds) (1997) Plant Analysis: An Interpretation Manual. CSIRO Publishing, Collingwood, Victoria, Australia. Riens, B., Lohaus, G., Heineke, D. and Heldt, H.W. (1991) Amino-acid and sucrose content determined in the cytosolic, chloroplastic, and vacuolar compartments and in the phloem sap of spinach leaves. Plant Physiology 97, 227–233. Robinson, R.A. and Stokes, R.H. (1959) Electrolyte Solutions. Butterworths, London. Rokitta, M., Medek, D., Pope, J.M. and Critchley, C. (2004) Na-23 NMR microimaging: a tool for non-invasive monitoring of sodium distribution in living plants. Functional Plant Biology 31, 879–887. Russ, J.C. and Dehoff, R.T. (2001) Practical Stereology. Springer, New York. Salt, D.E. (2004) Update on plant ionomics. Plant Physiology 136, 2451–2456. Schnitzler, J.P., Zimmer, I., Bachl, A., Arend, M., Fromm, J. and Fischbach, R.J. (2005) Biochemical properties of isoprene synthase in poplar (Populus x canescens). Planta 222, 777–786. Steiner, M. (1939) Die Zusammensetzung des Zellsaftes bei hoheren Pflanzen in ihrer okologischen Bedeutung. Ergebnisse der Biologie 17, 152–254. Walker, D.J., Black, C.R. and Miller, A.J. (1998) The role of cytosolic potassium and pH in the growth of barley roots. Plant Physiology 118, 957–964. Weibel, E.R., Kistler, G.S. and Scherle, W.F. (1966) Practical stereological methods for morphometric cytology. Journal of Cell Biology 30, 23–38. Wiggins, P.M. (2001) High and low density intracellular water. Cellular and Molecular Biology 47, 735–744. Willmer, C. and Fricker, M. (1996) Stomata. Chapman & Hall, London.

3 The driving forces for water and solute movement Tim Flowers and Anthony Yeo

3.1

Introduction

This chapter begins by describing some basic properties of water before embarking on an outline of the thermodynamics of solutions: the objective is to provide a background sufficient for the understanding of the forces that cause water and solutes, particularly ions, to move through plants. There are sections dedicated to water movement, ion movement and the linked flows of water and solutes.

3.2

Water

A water molecule consists of two hydrogen atoms covalently bonded to an atom of oxygen. Oxygen is the more electronegative so there is a greater probability of electrons being close to the oxygen than to the hydrogen, leading to partial charge separation and a partial negative charge on the oxygen and partial positive charges on the hydrogen atoms. This makes water a polar molecule and the electrostatic attraction between adjacent molecules constitutes a force known as the hydrogen bond. This lends ‘structure’ or ‘order’ to water, where there are continual transient associations between clusters of molecules. When water freezes, all the molecules are joined by hydrogen bonds: only 15% of these bonds break on melting and at 25◦ C, 80% of the hydrogen bonds are still intact, meaning that water may be termed ‘semicrystalline’ (Nobel, 2005). It is these associations between water molecules, and the free energy change needed to disintegrate them (some 20 kJ mol−1 ), that accounts for water being a liquid at temperatures regarded as ‘normal’: the structurally similar, but nonpolar, hydrogen sulphide is a gas at similar temperatures. The presence of hydrogen bonds in liquid water underlines properties vital for the physiology of plants. Work must be done to separate water molecules into the gas phase (the latent heat of vaporisation) and the heat required to evaporate water (44 kJ mol−1 at 25◦ C) is enough to dissipate most of the heat load of solar radiation falling on a wet surface. Water also has a comparatively large thermal capacity (a relatively large amount of energy is needed to raise its temperature). These two properties of water (the latent heat of vaporisation and the thermal capacity) are crucial to plants, buffering fluctuations in environmental temperature and providing the basis for transpirational cooling.

30

PLANT SOLUTE TRANSPORT

A further consequence of the mutual attraction between water molecules – its tensile strength – is integral to transpiration. It is its tensile strength that gives water its cohesion and allows it to hang in long columns in the xylem (see Section 9.2): the attraction between liquid and solid surfaces is adhesion, and when, according to the material, adhesion is greater than cohesion, this results in capillary rise. Cohesion underlies the phenomenon of surface tension (or surface free energy): the mutual attraction between molecules tends to minimise the surface at a liquid–gas interface. This is why water tends to form droplets and why there is a concave curvature at a water surface. The concave shape is produced because surface tension generates a negative pressure within the water, which is inversely proportional to the radius. The tension is negligible in a glass of water, but enormous at the pore sizes found in clay soils and plant cell walls (see Section 3.5.1). As water is virtually incompressible, it can support both tensions (negative pressures) and positive pressures, the latter allowing it to provide the hydrostatic skeleton of cells (turgor pressure; see Section 3.3.3). The polar nature of water also gives it a large dielectric constant and makes water one of the most general solvents known, particularly for the charged ionic species that are required by plants for their mineral nutrition. Being slightly charged, the polar water molecules are able to associate with, and shield, the electrical charges of other ions, preventing them from interacting with each other and, over certain ranges of activities (see Sections 2.3 and 3.3.3 for a definition of activity), preventing them from precipitating from solution. This is essential for maintaining the mix of ionic species needed by cells. This shielding effect is also what provides the protective hydration shells around macromolecules, by electrostatic attraction to charged or partially charged groups on the surface. This helps make proteins soluble and limits unwanted aggregation, or other interaction between macromolecules, within the cell – a problem that arises when cells are dehydrated during water deficit induced by drought, freezing or salinity. There are two other consequences of the ordering of water molecules at surfaces for the structure and activity of proteins – and hence cellular biochemistry. Thermodynamic considerations underlie the folding pattern of proteins with predominantly hydrophilic (charged or partially charged) groups at the surface and hydrophobic (uncharged groups) in the interior (similar to the lipid bilayer membrane; see Section 4.2.1). Structuring of water at protein surfaces is one of the factors contributing to the tendency of proteins to maintain their tertiary structure. The water in such hydration shells is more ordered than the ‘quasi-crystalline’ water at distance from a surface, and such structured water is less able to fulfil the liquid properties of water, including acting as a solvent. This leads to the concept of solute-available space within cell compartments. In the cluttered cytoplasm, packed with jostling membrane systems and macromolecules, a considerable proportion of the water may be locked away in hydration shells. Extreme views have been that little of the water is truly liquid. This has profound implications for what a ‘concentration’ in a cell compartment is or means (already mentioned in Section 2.3). Solutes not only dissolve in water, they can affect its solvent properties as well, and if they do so are known as cosolvents. They are of two types. A chaotrope is

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

31

a cosolvent that decreases structuring of water (for example urea and other protein denaturants). A kosmotrope is a cosolvent that increases structuring of water (for example glycerol, glycine betaine and other protein stabilisers). This is particularly important in an ecological context where certain solutes have protective roles in dehydration resistance and in dehydration tolerance (see Chapter 15).

3.3

Free energy and the properties of solutions

In biological systems, the largest flow of water is from soil, through plants to the atmosphere – the so-called soil–plant–atmosphere continuum. Under natural conditions, liquid water enters the system following rainfall – it may be distant rainfall with subsequent surface or subsurface flow, but precipitation of liquid water starts the cycle – and water ends up in the gas phase, in the atmosphere. Little (only about 2%; see Munns, 2005) of the water flowing in the system is retained in plants.

3.3.1

Free energy and chemical potential

From a thermodynamic point of view, the soil–plant–atmosphere system can be seen as a system that operates at approximately constant temperature and pressure. Atmospheric pressure does vary, but variations are normally small (about 10% of the average atmospheric pressure at sea level, which is 101.3 kPa): the extremes are about 87 and 109 kPa (http://en.wikipedia.org/wiki/Atmospheric pressure). Temperature also varies (the extremes on the world surface are about −90 to +58◦ C; http://en.wikipedia.org/wiki/Temperature extreme), but it is assumed that temperatures are virtually constant for short periods – where a temperature rises from 10 to 40◦ C over 6 h, then that would be an increase of only about 0.08◦ C min−1 . For a spontaneous change to occur in such a system at constant temperature and pressure, there must be a decrease in free energy. So, if water moves spontaneously from, say, plant to atmosphere, the free energy of the water must decrease. Free energy – formally, Gibbs free energy (G) – decreases for spontaneous processes at constant temperature and pressure. However, free energy is an extensive property; it depends, like mass, on the quantity of a substance: the bigger the system, the more the free energy. Hence it is important to be able to assess the amount of free energy of a substance independent of its quantity. This free energy per mole of substance is termed the chemical potential (μ). For water, its chemical potential (μ w ) is given by the equation:   ∂G (3.1) μw = ∂n w T,P,E,h,n j where n w represents the number of moles of water, T is the temperature, P is the pressure, E is the electrical potential, h is the height in a gravitational field and nj is the number of moles of other substances. Water will flow spontaneously from high to low chemical potential. The bigger the difference in chemical potential, the

32

PLANT SOLUTE TRANSPORT

greater the driving force for water flux (the amount moving per unit area per unit time). Equation 3.1 indicates that the chemical potential of water (in a system such as the soil–plant–atmosphere system) depends on temperature (T), pressure (P; water can be pumped through pipes), interactions between water and solutes (nj ; solutes lower μ w because they lower the activity of water; see Section 3.3.3), height in a gravitational field (h; it takes work to lift water; water runs freely downhill) and electric fields (E). However, the influence of the latter on the chemical potential of water is insignificant, as water does not carry a net positive or negative charge, and so E can be ignored. Consequently, the chemical potential of water at constant temperature depends on its activity (a w ), pressure (P) and height (h) in a gravitational field. This is expressed mathematically as: μw = μ∗w + RT ln aw + V¯w P + m w gh

(3.2)

where R is the gas constant, T is the absolute temperature, a w is the activity of the water, V¯w is the partial molal volume of water (see below), P is the pressure, m w is the mass per mole of water, g is the acceleration due to gravity, h is the height in a gravitational field and μ∗w is an arbitrary chemical potential of water under standard conditions (a constant of integration). The standard state (Eq. 3.2) is defined when a w = 1 (RT ln aw = 0), P = 1 and the height in the gravitational field is zero (m w gh = 0). In practical terms this is pure water at atmospheric pressure and the height (and temperature) of the system under consideration. The partial molal volume (V¯w ) is the rate of change of volume of water with increasing number of moles of water, when the number of moles of other substances, temperature, pressure, electrical potential and height in a gravitational field is kept constant (i.e. V¯w = (∂ V /∂n w )n j ,P,T,E,h ); its value is 1.805 ×10−5 m−3 mol−1 at 20◦ C.

3.3.2

Water potential and water potential gradients

Equation 3.2 is not readily usable, but it is possible to derive a more practical form of the relationship, firstly by defining ‘water potential’ as the difference between the chemical potential of water at any point in a system and that of pure free water in a standard state; that is: W =

μw − μ∗w V¯w

(3.3)

The term V¯w (m3 mol−1 ), the partial molal volume of water, is introduced to convert the units from those of free energy (J mol−1 ) to pressure (Pa – a Pascal is equivalent to a J m−3 ). Next, by substituting Eq. 3.2 into Eq. 3.3, the following relationship is reached: w = P −  + ρw gh (3.4)   where  is the osmotic pressure −(RT/V¯w ) ln aw and ρ w is the density of water (m w/V¯w ).

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

33

Equation 3.4 is commonly written as: w = p + π + g

(3.5)

where  p is the pressure potential (which may be positive, zero or, where water is under tension, negative),  π is the osmotic potential (a consequence of the presence of solutes and is always negative) and  g is the gravitational potential (which is negligible in cells and small plants but may be significant in tall trees: it is important in soils as the driving force for deep drainage and for the uptake of water for deeprooted species; see footnote in Section 3.4). Thus the awkward arbitrary constant of Eq. 3.2 is removed and the units converted from energy per mole to pressure units – those favoured by physiologists since the discovery of osmotic pressure. More details of the derivation of the formulae can be found in Nobel (2005).

3.3.3

Osmosis and colligative properties

Solute molecules interact with water to lower its free energy. The effect of a solute in lowering the free energy of water is easily demonstrated using a simple osmometer (Figure 3.1). Here, a concentrated solution is contained within a thistle funnel by a semipermeable membrane and then immersed in water. Water flows from the high potential in the surrounding water (0 MPa) into the solution until the increase in pressure, represented by the height of the water column, raises the free energy of the solution to that of the pure water surrounding the membrane. At this point there is no longer a gradient in free energy and net water movement ceases. At equilibrium, the effect of the positive pressure developed by the height of the water column (turgor

Osmotic pressure Solution Semipermeable membrane Pure water

Figure 3.1 Osmosis and the generation of osmotic pressure. A solution is separated from pure water by a semipermeable membrane: solute molecules represented by the single sphere are unable to cross the membrane. There is a net flux of water molecules (three connected spheres) across the membrane until the water potential in the solution increases to 0 MPa, due to the increase in pressure – equivalent to the head of water (osmotic pressure).

34

PLANT SOLUTE TRANSPORT

pressure in a cell) is equal and opposite to that of the solute in lowering of free energy. Such a situation arises for a membrane that is impermeable to the solute – that is a perfect semipermeable membrane (see Section 3.5 below). Equation 3.4 includes a term , the osmotic pressure (= −(RT/V¯w ) ln aw = −π ), that depends on the activity of water. If solute and solvent behave ideally, the osmotic pressure can be expressed in terms of the concentration of a solute (or solutes). Under these conditions:  = RTcs

(3.6)

where c s is the osmolality of the solution (a 1 osmolal solution contains 1 mole of osmotically active particles per kilogram of water). Here the concentration is expressed per unit mass of water (the molal scale) rather than the more commonly used basis of a litre of solution (molar scale): molality does not change with temperature and pressure, as mass is independent of these variables. For dilute solutions of low molecular weight solutes, molal and molar concentrations are similar. Above a concentration of about 0.2 M, the two scales diverge and increasingly so, the higher molecular weight of the solute. So far, the membrane has been considered as effecting a perfect separation between solvent (water) and solute. However, where solute passes through the membrane to some degree, the effective osmotic pressure is reduced. It is easy to imagine the two extremes – a perfectly semipermeable membrane, where all the solute molecules are reflected by the membrane, and a completely permeable membrane, where the solutes pass through the membrane. In the latter case, there would be no osmotic pressure. Membranes can vary in the proportion of solute reflected and can be characterised by their ‘reflection coefficient’ for a given solute. This is the ratio of the effective osmotic pressure to the theoretical osmotic pressure given by Eq. 3.6 (see also Section 3.6). The ability of solutes to change the free energy of water means that a number of properties of water change on addition of a solute, for example the vapour pressure, the freezing point and the boiling point as well as the osmotic pressure. These are the so-called colligative properties of solutions, and provided there are no large solute– solute interactions, there is a linear relationship between solute concentration and solution property. For example, the partial pressure of water vapour in equilibrium with a solution is linearly related to the mole fraction of water in the solution (Raoult’s law). The mole fraction is the ratio of number of moles of water divided by the total number of moles of water plus solute in the solution. The relationship holds to higher concentrations when expressed on the molal (that is per kilogram of solvent) rather than on the molar (per litre of solution) basis.

3.4

Cell water relations

In terms of their water relations, cells are complex in that they have two semipermeable membranes (plasma membrane and tonoplast) plus small organelles embedded in their cytoplasm (the chloroplasts, mitochondria and microbodies, all with

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

35

semipermeable membranes) and a cell wall composed of a water permeable matrix of complex carbohydrates and proteins. In spite of this complexity, cells do behave as osmometers and so when in equilibrium with water outside (o ), their internal (i ) water potential will tend to zero and wo = 0 = wi = p + π + g whence: −p = π + g

(3.7)

For a cell, the gravitational potential has little influence on its water relations. G can be calculated from the equation: G =

m w gh V¯w

(3.8)

where m w is mass per mole (18.016 g mol−1 ), g is the acceleration due to gravity (9.8 m s−2 ), h is the vertical height in m and V¯w is the partial molal volume (1.8 × 10−5 m3 mol−1 ). Since the value of G changes by 9.8 kPa m−1 or approxi∗ mately 0.01 MPa m−1 , this is too small a value to influence the water potential on the dimensions of cells (μm). So, at equilibrium in water: –p = π

(3.9)

It is the restraining cell wall that leads to the development of positive turgor pressures within cells. Turgor pressures can be measured directly with a pressure probe (H¨usken et al., 1978), giving values that range from 20 to 800 kPa (see, e.g., Clipson et al., 1985); values are quite substantial, especially when compared with the pressure of familiar items such as car tyres (about 200 kPa). The turgor pressure within cells can be modulated by solutes accumulating within the matrix of the cell walls, altering its water potential (Clipson et al., 1985; James et al., 2006).

3.5

Water movement

The measurement of water movement through the soil–plant–atmosphere continuum is not a trivial matter. If a plant is contained in a pot or a lysimeter (a container buried in soil), then water movement can be measured by changes in weight. For trees or tracts of vegetation weighing is only rarely possible (see, e.g., http://www.ars.usda.gov/Aboutus/docs.htm?docid=8680), but evaporation can be estimated using micrometeorological methods (see, e.g., Jones, 1992). However it is measured, the quantity of water moving through a plant will depend upon the size ∗ Per

metre of vertical height, the gravitational potential changes by 0.018016 × 9.8 × 1 (kg mol−1 ) × (m s−2 ) × (m) or kg m2 s−2 mol−1 or J mol−1 . To convert this to pressure units, divide by the partial molal volume V¯w , which is 1.8 × 10−5 m3 mol−1 , viz. (1.8016 × 9.8 × 10−2 )/(1.8× 10−5 ) = 9.8 × 103 (J mol−1 ) × (m−3 mol) or J m−3 . Since a J is an N m, a J m−3 is the same as an N m−2 or a Pa.

36

PLANT SOLUTE TRANSPORT

Table 3.1 Representative water potentials in the soil–plant–atmosphere system Component

Potential (MPa)

Wet soil Root Shoot Atmoshere 75% rh Atmosphere 50% rh

−0.1 −0.2 −0.5 −38.9a −93.6a

a

The relationship between the water potential of water vapour in the atmosphere and the relative humidity is given by:   rh RT ln w = (3.10) 100 V¯w

where rh is the relative humidity (%), R is the gas constant (8.314 J mol−1 deg−1 ), T is the temperature (K) and V¯w is the partial molal volume of water (1.805 × 10−5 m3 mol−1 at 20◦ C).

of the plant; a large tree will evaporate more water per unit time than a small plant of arabidopsis. Consequently, it is conventional to calculate flow rates or fluxes on the basis of the area of transport – the units are commonly g m−2 s−1 or mol m−2 s−1 or, for a volume flux, m3 m−2 s−1 (note that this is formally equivalent to a velocity). The driving force for this water movement is the difference in free energy between liquid water in the soil and water vapour in the atmosphere. In the soil–plant–atmosphere system, the water potential in the soil ranges from close to zero in wet soil to rather negative values (perhaps as low as −2.0 MPa, beyond the permanent wilting point for most plants) in dry soils (with a moisture content of 10–15%). The water potential of an atmosphere with 50% relative humidity is −93.6 MPa (Table 3.1; Eq. 3.10). So, here is the major driving force for water movement through the system – the difference in water potential between a high value in the soil, of around −0.5 to –1 MPa, to a low value of about −50 MPa or less in the atmosphere. This is the overall driving force, a large difference in free energy between liquid water in the soil and water vapour in the atmosphere. The movement of water through the soil–plant–atmosphere system has been likened to an electrical circuit – with a sequence of driving forces and of resistances (Figure 3.2; van den Honert, 1948). As in Ohm’s law, fluxes (current in the case of electrical circuits) are proportional to the driving force for movement (a difference in voltage in the case of Ohm’s law) divided by the resistance – or multiplied by the conductivity. These resistances are located in the soil, in the roots, the xylem, the leaves and in evaporation to the atmosphere. The relationship between flux and driving force can be expressed in the following way:

ψw Rw = w L w

JVw =

(3.11a)

JVw

(3.11b)

where JVw is the volume flux of water expressed per unit of area over which the flux occurs (in m3 m−2 s−1 ), L w is a water conductivity coefficient and R w is a resistance to water flow. To calculate the total flux, JVw must be multiplied by the area A.

37

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

Ψw

Atmospheric water potential

Ra

Atmospheric resistance

a

Ψw

Leaf water potential

Rx

Xylem resistance

l

Ψw

Root water potential

Rr

Root resistance

r

s

Ψw

Soil water potential

Figure 3.2 Various segments of the pathway of water movement in the soil–plant–atmosphere system, represented by resistances and potentials.

Using this relationship it is possible to calculate the relative resistances (measured in MPa s m−3 ; the reciprocal of conductance) in the various segments of the flux of water from soil to atmosphere. It is assumed that the majority of the water entering through the roots passes out through the leaves (only 2% is ‘consumed’ in growth). Hence, for a plant with a root system of area Aroot , leaf area Aleaf and volume flux J v , the quantity of water entering equals that leaving, or Aroot Jvin = Aleaf Jvout . Since the system can be divided into separate segments, each obeying the same rule that what enters one segment leaves to enter the next segment (Figure 3.2), flux through the whole system must equal that through the any segment (e.g. the xylem). Hence: xylem

JVroot Aroot =

w

− wroot

Rroot

atmosphere

x yl

= JV A x yl =

wroot − wleaf  leaf − w = w Rxylem Ratmosphere

(3.12) Inserting representative figures from Table 3.1 and rearranging to solve for R atmosphere : atmosphere

Ratmosphere = =

wleaf − w wshoot − wleaf

(Rroot + Rxylem )

−0.5 − (−38.9) (Rroot + Rxylem ) = 113(Rroot + Rxylem ) −0.2 − (−0.5)

This simple calculation illustrates a very important aspect of flow through the continuum that the resistance in the gas phase is significantly greater than the sum of the other plant resistances. R atmosphere is a consequence of the low water permeability of the leaf cuticle, and hence the control of water loss by the (much smaller) stomatal pore area. The calculation is, however, a simplification and if we are to understand

38

PLANT SOLUTE TRANSPORT

the system as a whole, we need to know precisely what drives water movement through the different parts of the system and what constitutes a resistance to flow. In understanding the soil–plant–atmosphere system, it is crucial to realise that the driving forces differ in different parts of the system: in soils, water movement depends on height in the soil profile (e.g. gravity) and on the forces that bind water in the small capillaries that exists between soil particles. In plants, water can, again, move through a matrix composed of small capillaries – the cell walls – or in small tubes (the xylem and phloem) where bulk flow of water occurs under pressure gradients. Between cells, however, water movement depends on the properties of the membranes, which are differentially permeable to water and solutes (the membranes are semipermeable).

3.5.1

Water movement through the soil

Soils are composed of particles that can differ greatly in size – from clays whose particles are less than 2 μm in diameter to sands where the particles can be up to 2 mm in diameter. The water potential of water in a soil is influenced primarily by the height of the water in the soil profile and, as the soil dries, by the negative pressures that develop in menisci formed as air enters drying soils. Dissolved solutes do not create a driving force for water in soils, as soils are not osmotic systems – there are no semipermeable membranes. Any influence of solutes is largely through their effect on the soil structure, which alters the permeability (conductivity) of the soil to water. In wet soils (the ‘wet end’ of soil moisture is measured as field capacity, the water content at which downward drainage under gravity materially ceases) the spaces between soil particles are occupied by water molecules, and so the water potential is determined by the gravitational potential. As the soil dries, however, water is withdrawn into capillaries between the soil particles and air–water interfaces develop. The development of these menisci alters the hydrostatic pressure (P) in the water since:   1 1 (3.13) + P = −σ r1 r2 where σ is the surface tension of water and r 1 and r 2 are two principal radii of curvature of the meniscus (see also Figure 9.4). As soils dry, water retreats into ever smaller capillaries, and so the tension in the water increases and the soil water potential becomes increasingly negative. Movement of water through soils depends on the driving force – the gradient of water potential and the conductivity of the soil to water (cf. Eq. 3.11). So the volume of solution flowing per unit area per second (J v ) depends on the gradient of soil water potential with distance, viz.: Jv = −L s

d s dx

(3.14)

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

39

where the soil water potential  s is given by  s = ps + Gs – the sum of the pressure potential in the soil (here a tension; see Eq. 3.13) and the gravitational potential (remember there is no osmotic component as there is no semipermeable membrane). In wet soils,  s is dominated by Gs and in dry soils by ps . The relationship between the flux of water through a soil and the driving force is termed Darcy’s law (after Henri Darcy, who discovered the relationship towards the end of the nineteenth century). Darcy’s law is a little more complex than might first be apparent, as the hydraulic conductivity of the soil varies with the water content – and declines dramatically as the soil dries. As a consequence, dry soil has an extremely low conductivity to water and provides one reason why root growth is so important to a plant for obtaining water. Once water reaches a root, its initial contact is with the cell walls of the epidermal cells: these represent a similar medium to the soil for water movement.

3.5.2

Water in cell walls

Cell walls are composed of a matrix of cellulose molecules impregnated with a variety of other more or less complex polymers. The pores within cell walls (about 5–30 nm) are, however, smaller than those found in soils. As in the soil, water will be held in microcapillaries and large tensions can arise as cell walls dry, due to the effects of surface tension at air–water interfaces (Figure 9.4). Where the space in the cell wall has a diameter of 10 nm, the tension could be as great as −29 MPa at 20◦ C (Nobel, 2005). In practice, however, large tensions do not develop when water is available in neighbouring cells, as water moves from those cells into the microcapillaries so that they are virtually filled – the tensions that do develop depend on the contact angle between water and the cell wall as well as the radius of the pore (Nobel, 2005).

3.5.3

Water movement across a root (or leaf)

Roots provide a number of possible pathways for the radial movement of water as well as for its longitudinal flow. The majority of radial movement occurs from the soil to the central vascular tissue and in regions of the root where root hairs are prevalent. Water can, potentially, move through the cell walls (apoplastic movement) or within cells (a symplastic route where water flows from cell to cell via plasmodesmata) or a combination of the two, whereby water moves from cell to cell across the root (Steudle and Peterson, 1998). The balance of water movement through the various pathways depends on the resistances to flow, which can change with flow rate and the developmental state of the root with respect to barriers to radial movement of water (Steudle and Peterson, 1998). The apoplastic pathway is potentially dangerous as water movement could carry solutes into tissues without any regulation by living cells. In practice, plant roots have developed barriers to the radial movement of solutes. These barriers are comprised of insoluble bands of suberin (Casparian bands) that are present in the walls of specialised cells that make up the exodermis and the endodermis (cf. Schreiber et al.,

40

PLANT SOLUTE TRANSPORT

2005). These barriers not only prevent unwanted ingress of solutes, but perhaps, more importantly, prevent the relatively concentrated solution in the xylem leaking from the root to the soil. The Casparian bands force water (plus solutes) into the symplastic pathway, except where exodermis or endodermis is breached either by the presence of passage cells or by damage caused, presumably, by the growth of lateral roots through the barrier (see Ranathunge et al., 2005). Where this occurs the so-called ‘bypass flow’ can constitute a significant pathway for the movement of ions from the external medium to the shoots, the best investigated example of which is rice (Yeo et al., 1987, and Chapter 14).

3.5.4

Water movement through the xylem and phloem

Apart from radial movement, water also moves longitudinally through the roots of plants, in the phloem and xylem; both are pressure-driven flows that can be described by the Poiseulle equation: Jv = −

r2 δP 8η δx

(3.15)

where J v is a volume flux with units of m3 m−2 s−1 , r is the radius of the tube, η is the viscosity (Pa s) and δP/δx is the pressure gradient under which flow takes place. The negative sign indicates that flow takes place in the direction of decreasing pressure. The xylem is an apoplastic pathway where water flows from relatively higher pressure in the roots to relatively lower pressure in the leaves: the water column is, however, under tension (see Chapter 9). The phloem, on the other hand, is a symplastic pathway; again water flow is pressure driven, but the pressures are positive, with relatively high values in the source regions and relatively lower values at the sinks, wherever they may be (Chapter 10). The xylem (except for root pressure exudation) operates under negative hydrostatic pressure, while the phloem operates under positive hydrostatic pressure.

3.6

Solute movement

The movement of solutes in plants can take place via the bulk flow of solutions, as in the xylem and phloem, or via specific transporters, where a protein is involved in the flux of a specific substance or group of substances. For neutral solutes the same driving forces occur as for water: the solute moves down gradients of its free energy (chemical potential) determined largely by gradients of activity (concentration). The effects of pressure (Nobel, 2005) and gravity are trivial in the context of cellular solute movements. For ions, however, there is an additional force that plays a very important role in their net movement – electric charge. As ions are charged particles, their movement is influenced by the presence of electric fields.

41

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

3.6.1

Chemical, electrical and electrochemical potentials and gradients

Unlike the case with water, differences in electrical potential (E) have a major influence on the movement of ions as ions carry a net charge (z, positive for cations and negative for anions; the charge carried by 1 mole of protons is 9.65 × 104 C or Faraday’s constant). The chemical potential of an ion j is given by: μ j = μ∗j + RT ln a j + z j FE

(3.16)

where z is the valency, F is the Faraday and E is the electrical potential (cf. Eq. 3.2 for water).

3.6.2

Diffusion – Fick’s first law

Although in the phloem and the xylem solutes move in a mass flow of solution, in many situations the movement of solutes depends upon diffusion. Diffusion results from random movements of solute molecules. Where there are differences in concentration between two sites, there is a greater statistical probability of movement from a region of high concentration to a region of low concentration as there are more molecules in the high concentration region than in the region of low concentration. The flux (J) of a solute j (viz. jj , the quantity of j crossing a unit area per unit time e.g. mol m−2 s−1 ) is directly proportional to the concentration gradient of j, viz.: δc j (3.17) δx where δc j / δx is the concentration gradient of j along the distance axis x and Dj is the diffusion coefficient of j (a coefficient rather than a constant as the value varies with temperature and concentration of j). This relationship is commonly known as Fick’s first law of diffusion, after its discoverer. Diffusion coefficients of common solutes in water at 25◦ C have different values, examples of which are 0.52 × 10−9 m2 s−1 for sucrose and 1.9 × 10−9 m2 s−1 for K+ (with Cl− ). It is informative to note that the time taken for a little over a third (36.8%, 1/e) of a population of K+ ions to diffuse across a cell (50 μm) is 0.6 s, while the time taken for the same proportion of these K+ molecules to diffuse over the distance of 1 m would be about 8 years (Nobel, 2005). Diffusion is not a process suited to long-distance transport in biological systems. In cells, there is a bulk movement of the cytoplasm, known as cytoplasmic streaming, which results in mass movement of solution and so reduces the time taken for solutes to move between parts of a cell. Where diffusion takes place across a barrier, such as a membrane or cell wall, the concentration gradient −δc j /δx can be represented by the difference in average concentration across the barrier divided by its effective width, i.e. the difference of concentration between the outside (o ) and the inside (i ) across the distance x, viz. (coj − cij )/ x. The distance over which solutes diffuse is the width of the barrier, plus any unstirred layers on either side of that barrier (layers where, because of friction between the barrier and the bulk solution, the bulk flow of solution is reduced to zero). Unstirred layer can be greater in width than the thickness of the barrier itself. Because the barrier, the membrane or cell wall, is not of the same chemical J j = −D j

42

PLANT SOLUTE TRANSPORT

composition as the bulk solution, the concentration of solute in the barrier depends on the partition between the two phases. This means the effective concentration   difference across the barrier is K j coj − cij , where K j is a partition coefficient, a dimensionless ratio of the concentration of the solute in the barrier and in an aqueous solution. The flux across the membrane is given by:   (3.18) J j = P j coj − cij where the permeability coefficient Pj is: Pj =

Dj K j

x

(3.19)

The permeability coefficient for K+ in a cell membrane is about 10−9 m s−1 , typical of small charged ions. In cell walls, ions will diffuse through aqueous channels, but these are relatively small in proportion to the unit area over which diffusion is occurring. For a cell wall whose thickness is 1 μm, the permeability coefficient would be 1 × 10−3 m s−1 , considerably greater than that of a cell membrane. However, the permeability coefficient of the same K+ ion in an unstirred layer of 30 μm would be 3 × 10−5 m s−1 , lower than that for the cell wall per se. For larger solutes, whose molecular dimensions are similar to the pore size in the walls, the walls can act as a ‘membrane’ with a reflection coefficient (see Section 3.2.2) less than 1 and osmotic water withdrawal can occur across the walls, as has been demonstrated for tissues where the cells have been disrupted by freezing and thawing (Flowers and Dessimoni Pinto, 1970). The limiting size of molecules that cross cell walls is 3.5–5.2 nm according to species (Carpita et al., 1979). Although the values of Dj , K j and x may be uncertain, Pj is a readily measurable quantity. Provided the flux of a substance can be measured and the internal and external concentrations are known, Pj can be calculated. The chief problem in estimating permeability coefficients is in obtaining the values of coj and cij at the membrane surface, since it is only possible to measure these in the solution on either side of the membrane. The boundary layer, in which the concentration varies with distance from the membrane, confounds estimation of the actual concentrations at the membrane surfaces themselves. The boundary layer cannot be entirely removed by rapid stirring of the solution, especially if intact plant cells are used, as this layer will be located within the cell wall where stirring is not possible. Values of Pj for small molecules such as glucose, glycerol and urea lie in the range from 0.01 to 3 × 10−9 m s−1 . It is difficult to estimate absolute permeabilities for ions, but P Na /P K is about 0.2 and P Cl /P K , 0.003 (see Nobel, 2005).

3.6.3

Diffusion potential

If a salt, such as potassium chloride, is added as a solid to a beaker of water, the ions dissolve in the water and a concentration gradient is established within the beaker, which leads to the diffusion of potassium and chloride ions from high to low

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

43

concentration. If the ions are of different sizes, they will have different mobilities in the solution and diffuse at a slightly different speeds so that a difference in charge develops, which is known as a diffusion potential. In a solution, this occurs over microscopic distances. Such diffusion potentials arise where microcapillaries filled with concentrated electrolyte as a conductor are inserted into cells to determine the potential across a membrane (cf. Section 2.6.3). For an electrode filled with 3 M KCl and inserted into a cell, the diffusion potential is about −2 mV.

3.6.4

Nernst potential

If a cell is placed in a dilute solution of a salt and allowed to equilibrate such that the diffusion of ions into the cell balances the diffusion out of the cell, there is no net movement of ions and the electrochemical potential of, say, K+ inside is equal to that outside the cell. In general terms, at equilibrium, μoj = μij , whence, expanding using Eq. 3.16: μ∗j + RT ln a oj + z j FE o = μ∗j + RT ln a ij + z j FE i

(3.20)

The potential at equilibrium E Nj (named after Nernst, who first derived this relationship) is given by E Nj = E i − E o . Rearranging Eq. 3.20, a oj   z j F(E i − E o ) = RT ln a oj − ln a ij = RT ln i aj whence E Nj = E i − E o =

a oj a oj RT RT ln i = 2.303 log i z j F aj zjF aj

At 25◦ C for a monovalent cation, this reduces to: E Nj = 59.2 log

co ci

(3.21)

Equation 3.21 demonstrates the poise between electrical potential and chemical concentration in systems at equilibrium: a difference in concentration of a monovalent cation of tenfold across a membrane is balanced by a difference in potential of 59 mV when the temperature is 25◦ C (at 20◦ C, the balancing potential is about 58 mV).

3.6.5

Donnan systems

Cell walls offer yet another level of complexity to the diffusion of ions in that walls carry a net negative charge. In such a system, at equilibrium, the Nernst equation can be applied, but here the concentration of anions and cations is different. For example, with K+ at a concentration (and activity) of 1 mM and Ca2+ at 0.5 mM outside a cell and with a cell wall with a concentration of fixed anions equivalent to 100 mM, it is possible to calculate the concentration of potassium and calcium ions

44

PLANT SOLUTE TRANSPORT

in the cell wall and the difference in potential between the outside solution and the cell wall (Briggs et al., 1961): Outside

Cell wall

Cytoplasm

Vacuole

K+ (mM)

1

9.5

150

50

Ca2+

0.5

45

–

–

(mM)

EN(K) (mV)

0

EN(K) (mV)

0

EN(K) (mV)

0

−57 −126 −100

In plant cells, it is likely that the fixed negative charges in the cell walls are largely occupied by calcium ions.

3.6.6

Goldmann equation

For a plant membrane, there are many ions that diffuse across it at any given time. However, for the most part, the fluxes are dominated by the movements of K+ , Na+ and Cl− (and H+ , but as we shall see H+ is pumped and therefore does not contribute to the diffusion potential). In this case it is possible to deduce (see, e.g., Nobel, 2005) that the measured potential across the membrane (E M ) is given by:   o i PK cKo + PNa cNa + PCl cCl  E M = RT ln  (3.22) i o PK cKi + PNa cNa + PCl cCl where P represents the permeability coefficients and c the concentrations. This is the Goldman or Goldman–Hodgkin–Katz equation.

3.7

Coupling of water and solute fluxes

Classical thermodynamics, the basis of most of the equations used so far in this chapter, deals mainly with the study of closed systems – systems that exchange energy but no matter through their boundaries. Plants exchange both energy and matter with their environment and are ‘open systems’. The methods of classical thermodynamics have been developed so that they may be applied to open biological systems providing new insights into the linked fluxes of solutes and water in both plants and animals. Irreversible thermodynamics applies the parameters used in classical thermodynamics to non-equilibrium conditions, i.e. to situations where there is a net flux of a substance, although the system must be close to equilibrium (Prigogine, 1961). We have already seen how the flux of a substance such as water can be described by an equation relating the driving force and the conductivity. For example, water

THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT

45

movement through a plant can be described by Eq. 3.11 (JVw = w L). Here, the volume flux of water is determined simply by the gradient of water potential. However, the movement of water is also affected by the movement of solutes and irreversible thermodynamics describes the flux of a substance in terms of all the forces (X, from 1 to n) that act upon that substance. So, the flux J of a substance j is given by: J j = L j1 X 1 + L j2 X 2 + · · · + L jn X n

(3.23)

where L is a coefficient. For fluxes of water (J w ) and solute (J s) , the equations reduce to: Jw = L ww μw + L ws μs

(3.24)

Js = L sw μw + L ss μs

(3.25)

and

These equations involve two fluxes, two driving forces and four coefficients, known as Onsager coefficients (after the Nobel Prize winner of 1968); the number of coefficients is, in practice, 3 as it can be shown that L ws = L sw . The development of these equations to describe fluxes of total volume (water and solute) was described by Kedem and Katchalsky (1958) and leads to the following relationships (Nobel, 2005) between the volume flux (J V ), the differences between the mean velocities of solute and water (J D ) and the differences in hydrostatic pressure ( P) and turgor pressure ( ) across a membrane: JV = L P P + L PD  JD = L DP P + L D 

(3.26) (3.27)

where L P , L PD , L DP and L D are the Onsager coefficients and L P is the hydraulic conductivity of the membrane. These coefficients have been used to define an important parameter, the reflection coefficient σ (=− LLDPP =− LLPDP ). This can be seen as the proportion of solute molecules ‘reflected’ by the membrane: for a perfect semipermeable membrane, σ = 1; for a membrane that is completely permeable to the solute (and so would not generate any osmotic pressure), σ = 0. The application of irreversible thermodynamics has generated insights into transport processes beyond those gained from the use of empirical relationships and classical thermodynamics.

References Briggs, G.E., Hope, A.B. and Robertson, R.N. (1961) Electrolytes and Plant Cells. Blackwell Scientific Publications, Oxford. Carpita, N., Subularse, D., Montezinos, D. and Delmer, D.P. (1979) Determination of the pore size of cell walls of living plant cells. Science 205, 1144–1147. Clipson, N.J.W., Tomos, A.D., Flowers, T.J. and Wyn Jones, R.G. (1985) Salt tolerance in the halophyte Suaeda maritima (L.) Dum. The maintenance of turgor pressure and water potential gradients in plants growing at different salinities. Planta 165, 392–396.

46

PLANT SOLUTE TRANSPORT

Flowers, T.J. and Dessimoni Pinto, C.M. (1970) The effects of water deficits on slices of beetroot and potato tissue. I: Tissue–water relationships. Journal of Experimental Botany 21, 746–753. H¨usken, D., Steudle, E. and Zimmermann, U. (1978) Pressure probe technique for measuring water relations of cells in higher plants. Plant Physiology 61, 158–163. James, J.J., Alder, N.N., Muhling, K.H., et al (2006) High apoplastic solute concentrations in leaves alter water relations of the halophytic shrub, Sarcobatus vermiculatus. Journal of Experimental Botany 57, 139–147. Jones, H.G. (1992) Plants and Microclimate A Quantitative Approach to Environmental Plant Physiology. Cambridge University Press, Cambridge, UK. Kedem, O., Katchalsky, A. (1958) Thermodynamic analysis of the permeability of biological membranes to nonelectrolytes. Biochimica et Biophysica Acta 27, 229–246. Munns, R. (2005) Genes and salt tolerance: bringing them together. New Phytologist 167, 645–663. Nobel, P. (2005) Physiochemical and Environmental Plant Physiology. Elsevier Academic Press, Amsterdam. Prigogine, I. (1961) Introduction to Thermodynamics of Irreversible Processes. Interscience Publishers, New York, p. 119. Ranathunge, K., Steudle, E. and Lafitte, R. (2005) Blockage of apoplastic bypass-flow of water in rice roots by insoluble salt precipitates analogous to a Pfeffer cell. Plant Cell and Environment 28, 121–133. Schreiber, L., Franke, R., Hartmann, K.D., Ranathunge, K. and Steudle, E. (2005) The chemical composition of suberin in apoplastic barriers affects radial hydraulic conductivity differently in the roots of rice (Oryza sativa L. cv. IR64) and corn (Zea mays L. cv. Helix). Journal of Experimental Botany 56, 1427–1436. Steudle, E. and Peterson, C.A. (1998) How does water get through roots? Journal of Experimental Botany 49, 775–788. van den Honert, T.H. (1948) Water transport as a catenary process. Discussions of the Faraday Society 3, 146–153. Yeo, A.R., Yeo, M.E. and Flowers, T.J. (1987) The contribution of an apoplastic pathway to sodium uptake by rice roots in saline conditions. Journal of Experimental Botany 38, 1141–1153.

4 Membrane structure and the study of solute transport across plant membranes Matthew Gilliham

4.1

Introduction

The transport of solutes across membranes is integral to a vast array of biological processes from plant nutrition to cell signalling, from symbiotic- and plant–pathogen interactions to cell polarity and plant development. Therefore, the number of plant scientists with a potential remit for exploring particular aspects of membranes and their transport systems is great. Numerous detailed texts exist concerning the function, structure and properties of plant membranes (see below), the transport proteins within membranes (see Chapters 5 and 6) and the techniques used to study either. This chapter provides a brief overview of available and emerging techniques, what they can reveal, their advantages and limitations and how they can be used in combination to demonstrate definitively certain transport processes within a plant. Readers interested in a particular technique are referred to citations within the text for further details, including in-depth methodologies. As the nature of the techniques available for the study of solute transport across membranes is dependent upon the properties and structure of plant membranes, a summary of their major features is provided below as context.

4.2 4.2.1

Plant membranes Plant membrane composition

All eukaryotic organisms have lipid- and protein-rich bilayers that delineate individual cells and compartmentalise intracellular regions into distinct organelles or other membrane-bound subcompartments. These membranes both form and provide an essential barrier between functional domains and their external environment acting as major sensors for environmental perception and stress response (e.g. cold, Uemura et al., 2006; salt, Zhu et al., 2000). In addition, plant membranes are key regulators of cellular homeostasis, platforms for metabolic-energy transduction (e.g. mitochondria or chloroplast membranes) and a dynamic matrix from which intracellular signals are released (e.g. Wang, 2005) – topics that will not be covered in any great detail here. Plants contain multiple membrane systems (∼20) that have been classified in terms of their form, composition and function (Figure 4.1). The plasma membrane

48

Membrane

PLANT SOLUTE TRANSPORT Distinctive transport proteins

Main roles

Nuclear envelope

RNA/protein transport

Endoplasmic reticulum

Synthesising, sorting and processing proteins

Golgi

Protein, vesicle trafficking

trans-Golgi network

Protein, vesicle trafficking

Partially coated reticulum

Protein, vesicle trafficking

Multivesicular body

Protein, vesicle trafficking

Secretory vesicle

Protein, vesicle trafficking

Plasma membrane

Signalling, cell wall synthesis, homeostasis

Endocytic vesicle

Protein, vesicle trafficking

Transport vesicle

Protein, vesicle trafficking

Tonoplast Storage vacuole Lytic Undefined

Storage of metabolites and toxins, homeostasis, pigmentation Lysis

Mitochondria

ATP synthesis

P-type H+ -ATPase, PIP1

TIPs, V-PPase TIP2 (α-TIP) TIP1 (λ -TIP) TIP3 (δ-TIP) not TIP1/2

Inner

ATP/ADP translocator

Outer

Porins

Chloroplast envelope

Connectivity/ Trafficking

Photosynthesis

Inner

VDAC

Outer

Porin P-type Cu2+ -ATPase

Thylakoid

Symbiotic membrane, e.g. symbiosome

N2 fixation

Perioxisomes

Lipid mobilisation, glycolate pathway, perioxidation

NOD26-like

Figure 4.1 Major plant membranes, their roles, major transport proteins and connectivity. —, connection through each membrane; ---, connection between indicated compartments.

(PM; sometimes referred to as the plasmalemma) forms the boundary around a cell. However, the connectivity of intracellular membrane systems, through plasmodesmata, to adjacent cells (see also Section 8.5.1) has led some to question this classical notion, viewing plants instead as supracellular organisms (Baluska et al., 2004). Regardless of views on this matter, as membrane cannot form de novo, the trafficking and connectivity between membrane systems facilitates growth, maintains and changes their composition and affords plant ‘cells’ a dynamic and responsive

SOLUTE TRANSPORT ACROSS PLANT MEMBRANES

49

network primed for survival. Although membrane systems are heterogeneous in their exact constituents, the mass of the general building blocks – lipids, proteins and carbohydrates – is maintained at a ratio of approximately 40:40:20 (Staehelin and Newcomb, 2000). However, it is the specific properties of these individual components that allow different membranes to perform their specialised functions. Glycerophospholipids (e.g. phosphatidylethanolamine [PE], phosphatidylserine [PS], phosphatidylcholine [PC], phosphatidylinositol [PI], phosphatidylglycerol [PG] and cardiolipin [CL]; see www.lipidlibrary.co.uk) constitute the most common class of lipids in the PM and mitochondria and also form their major structural components. These glycolipids consist of two hydrophobic hydrocarbon (fatty acid) tails (14–24 C), with at least one tail having one or more cis double bonds. The degree to which tails are saturated affects lipid packing within, and consequently the shape of, the membrane. Esterified to the fatty acid tails are charged polar (hydrophilic) head groups. The high percentage of lipids present with anionic head groups (e.g. PE, PI, PC) gives the PM a relatively high negative surface charge compared to other membranes, a charge that can be used to aid its isolation (see Section 4.5.1). Chloroplast membranes, in contrast to mitochondrial and plasmalemmal membranes, contain glycoglycerol lipids [e.g. mono- (MGDG) and di-galactosylglycerides (DGDG)], rare in most non-photosynthetic membranes, as their major structural lipid components. PS and CL are the distinctive lipids of the mitochondria but in phosphatedeprived conditions, presumably as a phosphate-conservation mechanism, DGDG content increases through direct transfer from chloroplasts (Jouhet et al., 2004). It has been hypothesised that it is the asymmetrical arrangement of MGDG and DGDG, on the inner and outer leaflets respectively, of the thylakoid membrane that allows it to become highly folded and tightly packed, maximising photosynthetic efficiency (Murphy, 1982). PG is also present in anomalously high proportions within the thylakoid membranes where it has been shown to be essential for chloroplast differentiation and autotrophic growth (Hagio et al., 2002). Other classes of membrane lipids include sterols (e.g. sitosterol and 24methylcholesterol) and glycosphingolipids (e.g. glycosylceramide). Whereas sterols have been implicated in the regulation of membrane fluidity and glycosphingolipids are thought to have roles in cell signalling (such as abscisic acid [ABA] signalling; Ng et al., 2001), it is likely that the roles of these two lipid classes may be intimately linked (see Sections 4.2.2 and 14.12). Phosphatidic acid, also involved in ABA signalling, is produced by hydrolysis of membrane lipids by phospholipase D and has been implicated in important signalling pathways such as root growth and programmed cell death (Wang, 2005). The amphipathic (amphiphilic) nature of lipid molecules, which concomitantly form continuous bilayers, generates a selectively permeable barrier around anything membrane-bound and facilitates the potential formation of large solute concentration gradients across the bilayer. Only highly lipid-soluble (e.g. ethanol, glycerol), small non-polar (e.g. O 2 , CO 2 ) and some small polar (e.g. H 2 O, urea) molecules are able to traverse the lipid bilayer passively and directly (Chapter 5). Proteins embedded within lipid bilayers can create additional transport pathways for lipid-impermeable substances or augment transport of those that are lipid-permeable (e.g. aquaporins; Luu and Maurel, 2005). Whilst integral membrane proteins are irreversibly bound

50

PLANT SOLUTE TRANSPORT

and their presence controlled by cytotic events, both peripheral (linked by salt bridges to other proteins or lipids) and lipid-linked proteins (e.g. fatty-acid-, prenyl-groupand sterol-linked) can form reversible associations with the membrane. Proteins can form a direct transport corridor across membranes through pumps, channels, carriers (see Section 5.1.2) or plamodesmata (see also Section 8.5) and control vesicle trafficking or regulate such processes indirectly. The protein constituents of the various membranes within plants can also be distinctive (Figure 4.1) and are therefore useful attribute for identifying particular tissue fractions (see Section 4.3.2).

4.2.2

Plant membrane structure

Membrane composition (and also structure) varies depending on species, cell type and plant physiological address (i.e. the plant’s current status as a result of its physiological and developmental history). For instance, the protein complement and transport properties (and functions) of the PM of xylem parenchyma cells differ from that of the guard cell (e.g. Gilliham and Tester, 2005). Moreover, both the protein and lipid composition of a given membrane can alter with changes in physiological conditions. The fluidity of lipid bilayers is naturally temperature-dependent (they will undergo a liquid-crystal to gel-like phase transition as temperature increases). Upon changes in temperature, to keep membranes at an acceptable fluidity for optimal physiological function, the plant can adapt the lipid composition of its membranes. For example, to increase fluidity of membranes upon cold stress, plants can increase the percentage of unsaturated phospholipids and decrease the percentage of sphingolipids (Uemura et al., 2006; see below). The four-dimensional membrane structure is dynamic and influenced by interactions between lipids, proteins, the cytoskeleton and the cell wall (e.g. McMahon and Gallop, 2005). Insights gained through recent technological advances have found the well-documented fluid-mosaic model of a biological membrane, developed by Singer and Nicholson (1972) (see Figure 1.9; Staehelin and Newcomb, 2000), to be a useful but underdeveloped generalisation of a biological membrane (Engelman, 2005). Drawing from biological membrane studies in other organisms, together with those in plants, evidence is emerging that plant PMs, and potentially other endomembranes, resemble a mosaic of microdomains with a particular molecular composition. This is in contrast with the traditional view that membranes are ‘liquid-disordered’, with most molecules being able to freely diffuse within the membrane plane. Interactions between areas of the membrane rich in sterols (in both lipid leaflets), and sphingolipids (solely in the outer leaflet), form ‘liquid-ordered’ microdomains (Martin et al., 2005). Sphingolipids have long acyl chains that form strong and tightly packed associations, thus endowing these domains with high-melting points. As a consequence, an increase in the proportion of these ‘liquid-ordered’ over ‘liquiddisordered’ domains decreases the fluidity of the membrane. Generically referred to as ‘lipid rafts’, these detergent-resistant membrane fractions are often enriched in glycosylphosphatidylinositol-anchored polypeptides (Bhat and Panstruga, 2005). Associations of these and other proteins, promoted by sphingolipids, are believed to

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51

form the lipid raft into a functional unit with specialised biochemical and signalling roles such as the induction of cell polarity (Fischer et al., 2004). As revealed through X-ray crystallography, it has been shown that transport proteins are more-often-than-not multimeric (e.g. T¨ornroth-Horsefield et al., 2006). Through other studies it has been demonstrated that they can in fact form functional heteromers (e.g. Dreyer et al., 1997). Regardless of their potential presence within lipid rafts, transport protein multimers are also often clustered. Furthermore, they may be in close vicinity to, or loosely associated with, other proteins within the bilayer or with those in the apoplast or symplast which may in turn affect or regulate protein-mediated solute transport activities. In addition, proteins that have large ectodomains, numerous transmembrane spanning regions or are anchored by single helices or lipidic anchors will cover significant areas of the bilayer surface and will therefore influence its structural properties. It has been suggested that lipid bilayers will also vary their thickness to accommodate protein structures (McMahon and Gallop, 2005). Specific interactions occur between lipids and proteins, with lipids acting either as co-factors or to ensure correct protein folding to guarantee membrane functionality (Valiyaveetil et al., 2002). It is therefore not surprising that changes in lipid composition and/or membrane fluidity can affect transport-protein function (e.g. the sterol-induced up-regulation of H+ -ATPase; Opekarov´a and Tanner, 2003).

4.3

Studying solute transport across plant membranes

The ‘information revolution’ spawned by the publishing of scientific journals, protocols and databases on the internet, as well as the recently sequenced and annotated genomes of multiple organisms, has provided the tools to mine data effectively and inform functional studies with unrivalled ease (see Section 4.6; Rhee et al., 2006). Rapid advances in molecular techniques have enhanced the power of many established (and some recently developed) transport assays manyfold, giving ‘traditional’ transport-based phenomenological or physiological investigations a new level of control and complexity. It has long been evident that there exists much intra-organ, -tissue, -cell and -organelle specificity in the transport properties of membranes and the molecular determinants of these differences can now start to be unravelled. Contemporary transport studies generally aim to associate a gene and a protein (or multiples of both) with a particular transport process by manipulating the transport process at the level of the gene, protein, cell and/or whole plant. As a result, the functional characterisation of transport proteins and their regulatory pathways is progressing swiftly (see the following chapters). A transport assay, as defined within the confines of this text, is any technique that can be used to elucidate a particular transport mechanism or infer the involvement of a particular transport process in plant function. There is much overlap in how general techniques can be applied, and so the remaining chapter has been divided into three main sections. Section 4.4 gives a background to most of the major transport assays with reference to their use with whole plants or semi-intact tissue; Section 4.5 describes how some of these techniques can be adapted for use with isolated

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membranes and Section 4.6 summarises some of the molecular techniques that are used to identify or directly manipulate a gene or a protein to enhance transport-based assays.

4.4

Transport techniques using intact or semi-intact plant tissue

4.4.1

Plant growth

Prior to initiating experiments, plant growth conditions should be carefully considered to ensure physical and physiological compatibility with a particular transport assay. Most physiological studies are initially carried out on seedlings for ease of handling, experimental control and efficiency, although many techniques can equally be applied to more mature plants if so needed. To name a few; aerated hydroponics, aeroponics, mica-based artificial soils, saturated filter paper and vertically orientated sterile agar/phytogel plates can be used to provide easy access to the roots. These methods can also give fine control over the nutrient content of the growth medium, which should, in any case, be carefully formulated to mimic physiological situations (it should be noted that full strength Murashige and Skoog, Hoaglands and/or high sucrose will actually perturb plant growth; see Gibeaut et al., 1997, for optimised hydroponic culture). At the same time, non-soil-based media may change root architecture and transport properties (Zimmermann et al., 2000).

4.4.1.1

Solution design

It may be important to attempt to calculate (or measure) the exact composition of the growth medium, but it is imperative for that of experimental solutions. The osmotic potential or osmolality (especially if osmotically stressing the plants or dealing with protoplasts or membrane vesicles) should be set, calculated or measured using an osmometer. Electrostatic attractions between solutes will decrease solute activities (a x ; see Section 2.3), especially at high concentrations (a square bracket is commonly used to designate a concentration, viz. the concentration of x is [x]). It is also important to consider the effect of chelators and the concentration of buffering agents within solutions. For example, by increasing the level of Ca2+ buffering in the cytosol using 1,2-bis(o-aminophenoxy) ethane-N, N, N , N  -tetraacetic acid at 25 mM, it is possible to completely prevent a [Ca2+ ] rise, induced by Ca2+ passage across various membranes, and therefore prevent its potential effects upon ion channel activity further downstream (Alexandre and Lassalles, 1992). GEOCHEM (Parker et al., 1987), Maxchelator® (Stanford University, California, USA) and Visual MINTEQ 2.23 (KTH, Stockholm, Sweden) are all available on the internet and can be used to calculate ion activities and buffering, although all will require users to enter additional constants for certain solutes. These calculated ionic activities can then be used to calculate the electrochemical potential in a solution or difference across a membrane, if required. Alternatively, ion/solute-selective electrodes (see Section 2.2.4) can be used to measure the activities of ions in solution (see Section 4.4.5.1).

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Boundary or unstirred layers parallel to the membrane surface, which occur at any air–water–solid interface, are a diffusional barrier between the bulk solution and membrane and may be exaggerated by the presence of the cell wall. Failure to take unstirred layers into consideration may result in significant errors when measuring biophysical parameters of membranes such as solute permeability (P s ) and hydraulic conductivity (L p ) (Tyree et al., 2005). Furthermore, the electrostatic interactions of ions with the outer membrane surface charge (ψ0o ) will affect the concentration of charged species at the membrane surface compared to that of the bulk solution (Kinraide, 2004). ψ0o , which is naturally negatively charged due to surface groups, can enrich the concentration of cations or deplete the concentration of anions at the PM surface by more than tenfold relative to the bulk solution (Barber, 1980). In contrast, high ionic strength, highly charge-dense ions or a low pH will induce slightly positive values (Kinraide, 2001). Most studies ignore the contribution of ψ0o to E m although it has been proposed that it could influence many flux parameters previously thought to be exclusively the result of direct ion interactions with transport proteins such as substrate saturation, rectification, inhibition by non-transported ions and voltage gating (Kinraide, 2001; see Chapter 5). However, as the ψ0o is subsumed within the total electrochemical potential between outside and inside, it is only important to calculate it if it is thought that the transporter (or particular transport phenomenon) is positioned, and so it experiences only part of the total electrochemical gradient. The cell wall Donnan phase (see Section 3.5.5) is not believed to affect membrane surface ion concentration significantly although it is thought to slightly increase the concentration of cations and decrease the concentration of anions (Kinraide, 2004).

4.4.1.2

Using inhibitors

Pharmacology is used in combination with most transport techniques as a diagnostic for the involvement of specific transport pathways or the membrane location of a transport process. For instance, mercury (Hg2+ ) is used as a diagnostic blocker of aquaporins (Niemietz and Tyerman, 2002), tetraethylammonium (TEA+ ) of K+ channels, gadolinium (Gd3+ ) of (stretch-activated) cation channels (Demidchik et al., 2002a) and niflumate of anion channels (Roberts, 2006). However, care should be exercised, as some blockers appear to have limited specificity (e.g. niflumate blocks both anion and K+ channels with similar potency, Garrill et al., 1996; and TEA+ has been reported to block aquaporins, Yool et al., 2002). A sensible approach to take when using inhibitors is to screen many compounds to build up a pharmacological profile of a transport process or to use engineered blockers (such as antibodies or synthesised chemical libraries that specifically inhibit particular transport proteins or phenomenon; e.g. Blackwell and Zhao, 2003).

4.4.2

Accumulation and net uptake

The concentration and net accumulation of solutes within tissues can be used to imply transport across membranes. For instance, digested tissue (or tissue extract) can be screened (see Section 2.2) in a high-throughput manner using flame photometry,

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ion chromotagraphy or ion-coupled mass spectrometry to identify transport mutants with particular ion profiles (Salt, 2004). Infiltration-centrifugation of samples is used to extract apoplastic-enriched solution, which can be similarly screened to differentiate it from symplastic content (e.g. Lohaus et al., 2001). All these techniques measure average content and have limited spatial-temporal resolution. Two established techniques with useful spatio-temporal resolution that are not invasive, but are costly and are useful for a subset of ions (or elemental compositions) include nuclear magnetic resonance (NMR; see Section 2.4) and X-ray microanalysis (XRMA; see Section 2.6.2). NMR can be used on whole plants to image concentrations non-invasively, or track movements of a limited number of solutes (13 C labelled compounds, e.g. glucose, [H+ ] with 19 F, phosphate using 31 P, Na+ with 23 Na) and, very effectively, water (using 1 H) (Kockenberger, 2001). XRMA can identify a greater number of elements but is used on fixed tissue (so lacks any real temporal resolution). However, when combined with electron microscopy, it can be used to construct revealing maps of ion distribution and compartmentation within tissues (e.g. Storey and Leigh, 2004). XRMA is extremely sensitive and can be used to measure ion content of samples on the picolitre scale, and therefore it has become an integral component of single-cell sampling (SiCSA; Section 2.6.4). SiCSA uses a pressure probe, itself a useful technique, for studying water permeability (P f ), P s and L p of native membranes in vivo, which may indicate the presence of protein-based transport pathways within membranes (Tomas and Leigh, 1999). For SiCSA, the pressurised glass microcapillary (with a tip <1 μm) is inserted into a cell and the vacuolar- or cytoplasmic-enriched samples withdrawn, due to turgor pressure. The elemental composition of specific cells accessible to a microcapillary can then be examined (e.g. Fricke et al., 1994).

4.4.3

Radioactive tracers

Accumulation into plant tissue can also be measured (even visualised) with good time resolution using radioactive isotopes (e.g. 109 Cd in leaves of Thlaspi caerulescens; Cosio et al., 2004). Plant tissue can be incubated in a solution containing a proportion of a particular solute as a radioactive isotope, e.g. 43 K, 86 Rb, 22 Na, 45 Ca, 36 Cl, 36 ClO 3 (for nitrate), and 14 C (labelling compounds such as urea and glucose), which acts as an analogue or tracer for the movement of the related solute form. The amount of radioactivity taken into (or effluxed from) the tissue is then measured over a number of time periods (see Section 2.6.2). To obtain the kinetics of unidirectional influx, experiments should be performed over a timescale of minutes (usually <20 min for most ions and tissues). The concentration dependence (and/or saturation) of uptake can also be obtained through fitting rates obtained from varying the concentration of (non-radioactive) solute with Michaelis–Menton parameters (K m and V max ) (Kaiser et al., 2002). Tracer-flux measurements have been used to identify both high-affinity and low-affinity transport systems in algae (e.g. MacRobbie, 1971) and plant roots (Kaiser et al., 2002). When conducting these experiments, the influence of unstirred layers may need to be considered, especially if V max is high and K m is low, as the presence of a significant unstirred layer may

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become rate-limiting and result in an underestimation of transport parameters (see methodology in Walker and Pitman, 1976). Ions are effluxed by plant tissues, often at similar rates to influx; so long tissue incubation times in radioactive tracer (>1 h) are needed to gain a more thorough understanding of the movement, and accumulation or compartmentation, of solute within cells, tissues and whole plants. Compartmental analysis by tracer efflux (CATE; e.g. Walker and Pitman, 1976; Britto et al., 2006) has been instrumental in increasing our understanding of the ion fluxes across the tonoplast and PM and of ion pool sizes within these compartments (e.g. Britto and Kronzucker, 2003) as well as in elucidating, in particular, the ionic control of stomatal movements (e.g. MacRobbie, 2006). CATE assumes that there are three basic compartments within plant tissue at the cellular level (cell wall, cytosol and vacuole) and all can be fitted with separate influx rates with first-order kinetics. The rate of efflux of tracer from the different compartments of loaded tissue can similarly be fitted with separate rates (with first-order kinetics) and subtracted from accumulation data in order to more accurately determine influx rates compared to those of steady-state measurements (see Section 2.6.6). Tracer-flux experiments are particularly effective when carried out on isolated cells, protoplasts or membrane vesicles as they reduce assumptions concerning redistribution of tracer within complex tissue; however, insightful models can be used in these circumstances to understand better the transport processes through whole plants (e.g. K+ and Cl− , Cram and Pitman, 1972; Na+ , Davenport et al., 2005). Despite its power for some ions, the validity of CATE has been questioned for ion pools that turn over rapidly (e.g. Ca2+ , Britto and Kronzucker, 2001). Sampled tissues must be dissolved or solutions combined with a specific cocktail for measurement of radioactivity on a scintillation counter; therefore radioactive tracer experiments are destructive and cannot be used to study movements of solutes in real time. Finally, licences and strict monitoring of radioactive materials is required unless stable isotopes such as 14 N are used.

4.4.4

Fluorescent solute probes

Epifluorescent photometry and microscopy has long been a powerful tool for measuring the average ion concentration of tissues with excellent temporal resolution (see also Section 2.6.5). Recent advances in confocal and multiphoton microscopy and digital camera technology, coupled with dramatic advances in the available solute probes, are now allowing the imaging of transport processes in some cases at the level of individual proteins and on the millisecond scale (e.g. Fricker et al., 2006). Therefore it is now possible to use these techniques to view the subcellular origin of ion fluxes or the interactions between transport proteins and regulators of transport in real time. There is a plethora of probes available for a variety of ions (and membrane voltage) with a huge range of spectral properties (consult www.probes.com) but care should be taken when choosing probes for plant tissue due to natural autofluorescence (e.g. chlorophyll, NADH). The fluorescent properties of probes measure the activity of target solutes through two basic mechanisms following substrate

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binding or dissociation, (1) by shifting emission or excitation spectra (allowing a ratiometric quantification) or (2) by changing emission or excitation intensity at a single wavelength. Single-wavelength probes have proved useful for measuring qualitative changes of ions if no alternative is available (e.g. Lemtiri-Chlieh et al., 2003). However, ratiometric probes (e.g. 1-[6-amino-2-(5-carboxy-2-oxazolyl)5-benzofuranyloxy]-2-(2-amino-5-methylphenoxy) ethane-N, N, N , N  -tetraacetic acid (fura-2) for Ca2+ and 2 ,7 -bis-(carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF) for H+ ) should be preferentially used as they allow (semi-) quantification after in vivo calibration. Ratiometric probes negate any heterogeneity in measurement due to non-uniform dye accumulation, leakage of probe, photobleaching, thickness of specimen or inequalities in fluorescence detection (Roos, 2000). In vivo calibration is also essential as many probes change their spectral properties in planta due to the altered ionic, osmotic or protein content of cellular environments over those commonly chosen for in vitro calibration (Shaw, 2006). It can also be necessary to monitor any potential concentration changes of ‘interfering ions’ (e.g. [H+ ]) during experiments as these may affect probe fluorescence and produce artifactual results (see M¨uhling and L¨auchli, 2002). Apoplastic movement, concentration and accumulation of solutes can be investigated relatively easily, using membrane impermeant chemical-based probes, by feeding the probe into the transpiration stream or through vacuum infiltration. For instance, the extent of the apoplastic continuum between the roots and shoots for water and solutes was evaluated in rice (e.g. Yeo et al., 1987) and ratiometric quantification of apoplastic H+ , Ca2+ , K+ and Na+ under various scenarios have all been made (e.g. M¨uhling et al., 1998; M¨uhling and L¨auchli, 2000, 2002). Loading of probes into cells across the PM and cellular compartments provides a more significant challenge. Membrane-permeant probes can be loaded by vacuum filtration but cuticles or (suberised) cell walls can restrict or bind dyes. Acetoxymethyl (AM)-linked dyes can be loaded into cells at 4◦ C to avoid AM cleavage by cell wall esterases (e.g. Zhang et al., 1998). However, incomplete cleavage in the cytosol often leads to accumulation of dye within organelles and signal contamination from different subcellular compartments. Low pH (∼4.5) can also be used to mask the charged carboxyl groups that usually render most probes impermeant, but both low pH and temperature may affect the desired physiological response. Most loading techniques require little specialised equipment and so membrane-permeant dyes are frequently used, but as the concentration of loaded probe is difficult to control and the presence of excess probe can ‘buffer’ observable changes in ion pool sizes or produce (phyto-) toxic effects, other techniques are preferred in more sophisticated physiological studies (Shaw, 2006). As a general guideline for these kinds of experiments, 5–50 μM of probe is sufficient and the illumination intensity used should be so low that only after acclimation to the dark should probe loading be perceptible to the eye. To control their concentration, probes are often linked to high molecular weight dextrans to prevent leakage to other compartments and are loaded via microinjection (e.g. root hairs, Foreman et al., 2003; pollen tubes, Feij´o et al., 1999). Microinjection has proven one of the most revealing and reliable methods (e.g. it was used to reveal the first calcium oscillations seen in plant cells; McAinsh et al.,

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1995), but is extremely technically demanding and has low throughput. Recent development of biolistic delivery (Bothwell et al., 2006) promises higher throughput with an equal level of control. However, all the techniques mentioned above are physically invasive and suffer from access problems, therefore providing a dilemma if researchers are required to study cells within the tissue profile. The development of new fluorescent protein (FP)-based nanosensors for solutes promises to overcome many of these problems as they are practically non-invasive, can be resolved at unparalleled spatio-temporal definition and appear to be present in low enough concentrations to not affect plant function (see Dixit et al., 2006, for a technical review). FPs can also be targeted at any cellular or apoplastic compartment (e.g. ER, Johnson et al., 2005; vacuole, Fluckiger et al., 2003), but the reduced fluorescent properties of current FPs at low pH (<6.5) make them less suitable as probes for solute dynamics in acidic environments. Single FP probes have been constructed (e.g. green fluorescent protein [GFP] has been modified to produce the ratiometric pH luorin to monitor cytosolic and apoplastic [H+ ]; Gao et al., 2004). Fluorescence resonance energy transfer (FRET) provides the opportunity to engineer FP sensors for a wide range of ions and metabolites by incorporating specific solute-binding moieties (for review see Lalonde et al., 2005). Upon binding of a specific substrate, a donor FP will transfer energy used in its excitation to an acceptor protein, thus decreasing its emission while increasing that of the interacting donor. In plants, Ca2+ (Allen et al., 1999) and Cl− (Lorenzen et al., 2004) dynamics have been studied, but powerful voltage and sugar probes also exist with many other probes being developed. Related to some FPs in origin, aequorin is a chemiluminescent protein that releases photons upon the binding of free Ca2+ . The quantum yield of aequorin upon binding Ca2+ is much lower than GFP but when overexpressed in a particular compartment (whole plant or cell-type specific targeting to the cytosol, to the cell wall or to intracellular compartments) it does facilitate a technically undemanding assay of average tissue [Ca2+ ] (e.g. Kiegle et al., 2000a; Gao et al., 2004). Aequorin assays have, at best, a tissue-type level of spatial resolution but are particularly effective in revealing information about stimulus-invoked [Ca2+ ] signatures, or the origin of [Ca2+ ] signals, when combined with the use of inhibitors and the specific targeting of apoaequorin to compartments. [Ca2+ ] can be imaged fairly non-invasively in whole plants, detached tissues or protoplasts, with relatively poor temporal resolution (>20 s for stimulus-induced transients), or measured photometrically, with better temporal resolution (∼1 s) but with limited spatial resolution. [Ca2+ ] ‘resting’ levels of the relevant compartment can also be measured, over tens of minutes, with sensitive photon-counting devices (Love et al., 2004).

4.4.5

Electrophysiology

Bioelectricity is a consequence of, and driving force behind, a plant’s transport economy (cf. Section 3.6.1). A plant’s electrical circuit consists of a membrane, which separates two areas of charge, and electrogenic transporters (ion channels, co-transporters, pumps). In conventional terms, this corresponds to a capacitor in

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series with resistors. The compound electrochemical potential gradient (as a consequence of ion concentration equalities) across a membrane (membrane potential – E m ) equates to voltage. Many ‘transport non-literate’ researchers find that data from electrophysiological experiments are presented in an intimidating or, at least at first, a non-intuitive way. However, good basic introductions on how to interpret data or conduct experiments are available (e.g. Tester, 1997). Knowledge of Ohm’s and Fick’s laws and the derived Nernst equation (refer to Sections 3.6 and 5.1.3) are essential to understanding many electrophysiological assays. Some additional technological instructions should then be adequate to carry out many procedures. Several standard texts are referred to for a complete grounding in electrophysiological theory and techniques: Odgen (1994), Sakmann and Neher (1995), Hille (2001) and (specifically for plants) Volkov (2006). As the currents across membranes are generally small in magnitude (single ion channels can pass a current of <1 pA), high resistance recording equipment is ∗ needed. Therefore, it is essential that experiments be performed in an environment free of both vibration and electrical interference (e.g. on an air-table in an earthed Faraday cage). All electrical equipment external to the cage (e.g. computers and amplifiers) should be earthed to one point on the cage and share one universal grounding point. Most electrophysiological techniques are invasive but give direct and instant information about transport activity across membranes, unlike most of the techniques already mentioned which rely on ‘time-integrated’ assays. Unfortunately, most direct electrophysiological assays of transport are possible only for electrogenic transporters (those whose activity results in the movement of net charge); non-electrogenic transport such as that of water, or of ions through electroneutral carriers, cannot be studied directly with these techniques (however see Section 4.4.5.1).

4.4.5.1

Voltage-based measurements (membrane potential and ion concentration)

E m (also referred to as ψ) and the concentration of the ion of interest (x) on either side of a membrane ([x] in , internal; [x] ext , external), as well as the ion valency, define the electrochemical potential gradient for ion x. E m can be measured directly through the insertion of a ‘sharp’ (tip <1 μm) borosilicate/quartz microelectrode (backfilled with an electrolyte, usually ∼0.1–3 M K-Cl/K-acetate) into a membrane compartment. To complete the circuit, in series with the microelectrode should be a voltmeter and a reference electrode in the bulk external solution (see Ogden, 1994). An ion selective electrode (ISE) can be constructed by insertion of an ionophore (selectively permeable for ion x over other ions in a linear [Nernstian] fashion) into the microelectrode tip (cf. Section 2.2.5). [x] can be measured after backfilling the ISE with an electrolyte of known [x] and calibrating its voltage response in the ∗Ohms law dictates that if resistance of the measuring circuit in parallel with the plant membrane is above a threshold (∼1 G), ionic currents across membranes can be measured as a change in E m (Ogden, 1994).

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desired concentration range of [x] (e.g. Miller and Wells, 2006). A specific ISE can only be constructed if the desired ionophore is available or can be synthesised, but there are a wide range available for both cations and anions (see www.sigma.com). Similar solid-state microsensors have also been developed for other solutes such as O 2 , NO and auxins (Mancuso and Marras, 2006). Unfortunately, ionophores vary in their selectivity profiles and ISE signals will suffer from contamination from other ions to varying degrees, notably Mg2+ from Ca2+ , Cl− from NO 3 − and organic anions, and K+ from NH 4 + . However, it may be possible, using several ISEs with different selectivities for the ‘interfering’ ions, to estimate [x] and other ion concentrations in these circumstances (Knowles and Shabala, 2004). When an ISE is inserted into a membrane compartment it will measure E m in addition to [x] in . Therefore, both an ISE and E m electrode have to be inserted into the same membrane compartment and E m subtracted from the ISE signal to obtain [x] in (Miller and Wells, 2006). Once both [x] in and [x] ext are determined, the Nernst equation can predict whether at the ‘resting’ E m the [x] in can be accounted for by purely passive processes or whether active transport plays a role at the measured [x] ext . To reduce damage to cells, multibarelled electrodes are constructed instead of impaling the same cell multiple times. Multibarelled electrodes can also be useful to ascertain which membrane compartment the electrode has penetrated. As the vacuole usually inhabits the majority of the cell interior, impalements may penetrate the tonoplast; therefore any E m reading will be a compound measurement of both that of the tonoplast and PM. A combined pH and E m reading will therefore establish whether the impalement is cytoplasmic or vacuolar (as an approximation, vacuolar – pH 5–6, ∼100 mV and cytoplasmic – pH >7, >−120 mV, when measured in an extracellular K+ of 1 mM; cf. Section 2.6.3). E m measurement has also been informative in understanding the electrogenic basis of stimulus-invoked membrane processes (e.g. action potentials in algae (Beilby, 1989) and stomatal guard cell movements (MacRobbie, 1985). Multiple-calibrated ISE can also be used to estimate ion fluxes around biological tissues with high spatial and temporal resolution. [x] ext is measured at points a known distance away from the source over a set period of time, and the parameters integrated using Fick’s law of diffusion (Sections 3.6.2 and 5.1.3) or an equivalent electrochemical law. Three commercial systems are available: MIFE® (microelectrode ion flux estimation) (University of Tasmania, Hobart, Tasmania, Australia; Newman et al., 1987), Seris (self-referencing ion selective) probe (BCRC, Woods Hole, MA, USA; Kuhtreiber and Jaffe, 1990) and SIET (scanning ion-selective electrode technique) (Applicable Electronics Ltd., Maryland, USA; Shipley and Feij´o, 1999). These have been used to gain valuable information about ion fluxes involved in the polar growth of pollen grains and tubes, roots, root hairs and various other tissues (Lew, 1998; Newman, 2001; Kunkle et al., 2006, Messerli et al., 2006; Shabala, 2006). Caution should be practiced when interpreting flux data, as the convention used when presenting data is usually opposite to that used in other electrophysiological assays such as voltage clamping (see Gilliham et al., 2006b). Both intracellular ISE and external ISE-based flux measurement can be used to study the non-electrogenic movement of solutes across membranes if the relevant sensor is available.

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4.4.5.2

Voltage clamping

The symplastic continuum within plants makes it impossible to effectively control the voltage across most cells; however, this can be achieved in cells that are (relatively) electrically isolated (e.g. root hairs, guard cells; Lew, 2006) or that have been isolated by other means (e.g. protoplasts; see Section 4.5). By controlling E m it is possible to observe the passage of charged solutes across membranes as current (I i ) over time (down to the level of a single channel). V m can be fixed at a nominated level and compared to E m across the poles of a feedback amplifier, the difference being due to I i (the flow of ions across the membrane). The feedback circuit injects a current (I f ) equal and opposite to I i and all parameters can be recorded by computer. I i can be similarly clamped to observe the change in voltage. V m (or I i ) clamping can be performed on larger cells using two intracellular microelectrodes (TEVC) or by using one electrode (dSEVC). V m clamping therefore allows the characteristics of transporters (conductance, gating and selectivity) to be identified over the physiological E m range whilst controlling the electrochemical potential across the membrane. TEVC has revealed much information regarding the ionic basis of stomatal movement in vivo (Blatt and Armstrong, 1993), and while presently underutilized in plant sciences, I i clamping could be useful in studying ligand-gated ion channel activity in intact or isolated tissues.

4.5 4.5.1

Using isolated membranes for transport studies Isolating membranes

Specific cell types or native membranes, such as those buried within the plant tissue profile, that are inaccessible to many of the conventional assays mentioned above, can be investigated directly after isolation from whole plants. The cell wall (and its obvious influence in signalling or the creation of unstirred layers) can be removed and the effect of intracellular components (and their regulation of membrane processes) can be diminished by gaining access to the cell interior or by inverting vesicles. These two obvious advantages of working with isolated membranes, or cells, might also be a disadvantage as transporters may not show true physiological activity or may even show no activity. Incubation of tissues in a cocktail of enzymes that breakdown cell walls and middle lamellae (e.g. cellulase, cellulysin, pectolyase, macerase) will release cells or cell fragments (termed protoplasts or spheroplasts) into solution, bound simply by a PM, but overexposure to the cocktail will decrease membrane viability and integrity (e.g. Vogelzang and Prins, 1992). To protect membranes from rupture, an osmoticum (e.g. mannitol) must be included, although vacuoles can be isolated from protoplasts by osmotic shock (e.g. Pantoja and Smith, 2002). To protect against proteases, bovine serum albumin should be included as a competitive substrate and to protect against phenolic oxidases, polyvinyl pyrrolidone, ascorbate and metabisulfite can all be included; pH will also need to be buffered. It has also been reported that increasing the level of Ca2+ improves protoplast yield and viability (Demidchik et al., 2002b). Mechanical techniques are also used to isolate cells, such as laser

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ablation of root hairs (V´ery et al., 2000) or laser capture microdissection of specific cell types (Nelson et al., 2006). Protoplasts can also be selectively released (e.g. from epidermal cells; Foreman et al., 2003) or identified and sorted based on endogenous properties such as size, viability and chlorophyll content (Wegner and Raschke, 1994; Galbraith and Birnbaum, 2006). Fluorescence-activated (cell) sorting can be used to obtain viable protoplasts from specific cell (or organelle) types by their identification through FP expression using an appropriate regulatory sequence (e.g. GAL4-GFP cell-specific enhancer trap lines; Kiegle et al., 2000b; Galbraith and Birnbaum, 2006). Protoplasts can make a convenient starting point for isolation of internal organelles, or membranes can be fractionated directly and there are established protocols for obtaining most membranes with a high purity (∼>95%; Robinson and Hinz, 2001). Typically, tissue is homogenised (using a pestle and mortar, blender or sonication with glass beads, depending on the shearing strength required to break cell walls) followed by purification. Several steps of differential centrifugation are used to separate cell debris and the different membrane fractions in terms of mass and density (most commonly using sucrose or iodinated compound density gradients). Fraction homogeneity is then confirmed using membrane-specific stains, immunological detection (e.g. tonoplastic intrinsic proteins [TIPs] for tonoplast fractions) or enzyme assays (glucan synthase for PM fractions). To reduce the deleterious effects of vacuolar rupture, tissue should be homogenised at 4◦ C and incubated in a protective solution, similar in composition to a protoplast isolation medium but without the cell wall-degrading enzymes. For transport studies, the sidedness of vesicles should be considered as catalytic sites, for many transporters are on the inside (e.g. ATPases). Aqueous polymer two-phase partition centrifugation (using polyethylene glycol and dextran), which preferentially purifies hydrophobic and negatively charged vesicles, is used to select ‘tight’ (non-leaky) right-side-out PM vesicles into the polyethylene glycol upper phase. Brij 58 (a detergent) can be used to increase the percentage of inside-out vesicles (Johansson et al., 1995), whereas several freeze-thaw cycles can increase the proportion of right-side-out vesicles.

4.5.2

Assaying transport activities of protoplasts and membrane vesicles

Membrane vesicles have been used extensively to characterise pumps (ATPases, PPases; e.g. Sze 1985), co-transporters (Qui et al., 2004) and aquaporins (Alleva et al., 2006) in native membrane fractions (e.g. PM, thylakoid, tonoplast) or through the use of purified proteins reconstituted into artificial liposomes. The accumulation of radioactive isotopes (e.g. 45 Ca; Marshall et al., 1994) or changes in light scattering (or fluorescent properties of an entrapped probe) can be used to measure the kinetic parameters of solute or water movement across isolated membrane vesicles (Verkman, 1995) and rate constants for ATP binding, substrate pumping or transport stoichiometry can also be determined. Fluorescent probes can be loaded into vesicles by many of the techniques mentioned in Section 4.4.4 and via electroporation and detergent permeabilisation, although these often make membranes ‘leaky’. Stopped-flow spectrophotometers allow quick solution exchanges (<1 ms) in small

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volumes (20–100 μl) and are capable of measuring rapid spectral changes in vesicle preparations (<100 ms) induced by the movement of solute (or water) into or out of the vesicle across the membrane of interest. For instance, P f or P s of membranes can be determined by fitting a time course to light scattering changes, if initial vesicle size is known and homogeneous, and this has become a useful technique for investigating aquaporin properties of native membranes in vitro (Niemietz and Tyerman, 2002). Light scattering can be sensitive to motion effects during fluid mixing and changes in refractive index although these can be easily controlled for, whereas the use of fluorescent probes negates these problems but has other caveats (see Section 4.4.4). Patch clamp electrophysiology is a technique that allows the voltage (or current) clamping of protoplasts or membrane vesicles (Hamill et al., 1981). A ‘blunt’ and fire-polished microelectrode (of diameter ∼1 μm) is pressed against the ‘naked’ membrane and forms a high resistance seal preventing the leakage of ions across the junction. Suction can be applied to the interior of the microelectrode and the membrane beneath ruptures, giving access to the internal contents. Patch clamping therefore allows the solution composition of either side of the membrane, as well as V m , to be controlled. Solutions are often carefully designed so that the activity of a single type of ion channel or transporter can be observed in isolation or can be defined (e.g. if K+ transport is the object of study, K+ will be the predominant permeable ion in solution, or alternatively the Nernst potentials for other ions will be engineered to be far apart so that clear comparisons can be made with reversal potentials, E rev ). Regulators of transport can also be determined by their addition to either side of the membrane (e.g. phosphatase-dependent ABA activation of Cl− channels in guard cells; Pei et al., 1997) unlike TEVC or dSEVC. However, the dilution of regulators within intracellular milieu could also result in ‘run-down’ or aberrant transporter activity. Patch clamping can be performed at the ‘whole-cell’ level, which measures ‘macroscopic’ current (the net current through all active electrogenic transporters within the membrane, as with TEVC), or, in its mostly highly resolved form, at a ‘patch’ level, which measures ‘microscopic’ currents in a small section of membrane (the net current through single [or few] electrogenic transporters). Patches can be in an either ‘inside-out’ (internal membrane side facing the bathing solution) or ‘outside-out’ (outside facing the bathing solution) configuration. A ‘cell-attached’ configuration, although having the advantage that cellular contents remain undiluted, is infrequently used as V m and electrochemical gradients are hard to define. Precise measurements of conductance, gating and selectivity can be made for individual channels, or the pumping rate or coupling ratio for pumps, or stoichiometries for transporters, can be ascertained using E rev (e.g. Davies et al., 1996; Gilliham and Tester, 2005; however see Section 4.7). Fluorescent dyes can be loaded via a patch pipette and be used for imaging, or photometry, of ion concentrations whilst simultaneously controlling V m . This can reveal whether certain membrane currents on particular membranes are responsible for specific ion increases (e.g. Lemtiri-Chlieh et al., 2003). The capacitance of the membrane can also be monitored during patch clamp experiments and related (using a constant) to membrane area; this can then be used

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to study processes that involve a change in membrane area such as endocytosis and exocytosis (Homann and Tester, 1997). These may be important for solute trafficking events directly (e.g. capacitive Ca2+ flux) or indirectly (e.g. fusion of ion-channel containing vesicles induced by turgor changes; Meckel et al., 2004). Membrane vesicle preparations, or liposomes that have had reconstituted protein added to them, can also be patch clamped. More commonly, these membrane preparations are added to a planar lipid bilayer (PLB) system to survey their electrogenic activity upon fusion, with the PLB painted across the divide (of ∼200 μm) between two solutions. PLBs use the same electronics as the patch clamp system and so can similarly characterise single channels. PLBs have been useful for observing dominant ion channels in the PM, thylakoid and mitochondria (e.g. White and Tester, 1994). Lipid composition of the PLB must be carefully considered, as surface charge or structural effects may affect channel function; it has also been observed that some lipids can act as ionophores and conduct ion currents (M. Tester, unpublished data).

4.6 4.6.1

Using molecular techniques to inform transport studies Revealing the molecular identity of transporters and testing gene function

In silico selection can be useful for identifying putative transporters such as K+ channels (Gaymard et al., 1998) and aquaporins (Sakurai et al., 2005) but has so far been less definitive for other families (e.g. GLR, Gilliham et al., 2006) for which no transport function has yet been identified. Additionally, as functions for >50% of genes within the sequenced plant genomes remain yet to be ascribed, such approaches may not be so straightforward. Regardless of these limitations, invaluable web-based annotated and hot-linked databases exist (e.g. in arabidopsis; for membrane proteins – aramemnon.botanik.uni-koeln.de, www.suba.bcs.uwa.edu.au, genes –www.arabidopsis.org, and toolkits for homology or motif-based searches – www.ncbi.nlm.nih.gov/blast). Transport functions of proteins can be identified by studying plant populations with different genotypes or phenotypes. Forward genetic screens rely on the isolation of mutant plants grown in specific conditions by phenotypic analysis (e.g. root or shoot growth, survival or altered ion accumulation, an example being sos1; Qui et al., 2002 – see Section 4.4.2) followed by a technique used to isolate the allele responsible for a particular trait, such as QTL analysis or subtractive hybridisation of cDNA (Flowers et al., 2000). However, this technique is rarely successful in isolating transport mutants directly. Phenotypes may often be too subtle to detect or gene function may be compensated by physiological plasticity (otherwise known as pathway ‘redundancy’). Once a target gene has been identified, reverse genetic screens using ‘loss-offunction’ (or ‘knockout’) mutants (created by the insertion of T-DNA, insertion elements or transposons through plant transformation) are useful for analysing the function of a gene or pathway (Peiter et al., 2005). A database of available potential

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‘knockout’ lines of arabidopsis can be found at www.arabidopsis.org and a map of the location of the ‘knockout’ insertional element within the genome is available at atensembl.arabidopsis.info. RNAi allows multiple genes from the same family to be silenced and so reduces the chances of pleiotropic up-regulation of related genes that can mask phenotypes. TILLING, which is a high-throughput polymerase chain reaction based screen of heavily mutagenised plant lines, is used to recover a range of mutations, not only knockouts, within a specific gene and therefore can identify critical residues within a protein (Comai and Henikoff, 2006; see tilling.fhcrc.org:9366). To confirm whether a phenotype is a result of the gene of interest, ‘knockout’ mutants should be recomplemented with the specific gene to restore transport function (e.g. Peiter et al., 2005). Overexpression of target genes can be performed in planta or in homologous or heterologous expression systems (see Section 4.6.3) to see if a transport phenotype has been exaggerated. This can be tested through a relevant assay (e.g. solute accumulation – see Section 3.5.2, or increase ‘whole cell’ conductance – see Section 4.5.2). In planta, overexpression can be constitutive, which may increase the chances of observing a phenotype. However, ectopic expression may also mask phenotypes if cell-specific or developmental processes are crucial to revealing a transport phenotype in which case overexpression can also be driven by (inducible) cell- or tissue-specific promoters (e.g. Moore et al., 2006).

4.6.2

Location of transport proteins

Equally as important to ascribing a physiological transport role to a particular protein (in addition to the transport activity it catalyses) is its spatial and temporal expression. Fractionation has revealed valuable information about transport activities of specific membranes (see Sections 4.5.1 and 4.5.2). However, as membrane proteins are highly hydrophobic and often low in abundance they have been difficult to isolate until recently (ATPases being an exception; e.g. Harper et al., 1989). This area has recently been reinvigorated through the use of chloroform–methanol extraction or blue native PAGE separation, and identification of membrane proteins in most organelles is now underway through mass spectrometry-based sequencing in a high-throughput manner (e.g. Heazlewood et al., 2005). Once a particular protein sequence is known, the gene that encodes the protein can be uncovered in silico, as can any related genes and their function tested through recombinant technologies (see Sections 4.6.1 and 4.6.3). However, as membrane preparations are not 100% pure, this can sometimes lead to the misidentification of the membrane on which a particular protein is located (e.g. Wandrey et al., 2004) and it is therefore important to confirm such information with some of the techniques mentioned below. Reporter genes (e.g. β-glucoronidase, GFP or luciferase) can be fused to gene promoters, if known, to reveal tissue localisation. In addition, gene fusions to GFP (or other markers) may reveal membrane location within a plant or a heterologous system (Peiter et al., 2005). However, such gene fusions may also interfere with membrane targeting or protein function or may mark some membranes anomalously

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due to overexpression. Antibodies raised to specific epitopes may be used in situ but can equally be used on membrane fraction preparations (e.g. immunolocalisation, immunoprecipitation or western blotting) to overcome some of these issues, but the successful preparation of such antibodies is not yet guaranteed. Transcriptional analysis, using northern blots, real-time reverse-transcription polymerase chain reaction and macroarray or microarrays of particular genes can be useful to study how its expression can be regulated. Gene families with close homologues (e.g. aquaporins; Jang et al., 2004), or perhaps pleiotropic regulation of these genes following ‘knockout’ or overexpression of the gene of interest, can also be studied using such analysis. However, transcript levels may not always correlate with protein levels, so it may also be wise to use additional proteomic analysis such as antibodies (e.g. Aroca et al., 2005).

4.6.3

Heterologous expression

Heterologous expression systems provide an additional mechanism for validating and functionally characterising the transport activity (or regulation) of plant genes. Whilst the influence of plant endogenous regulators and metabolism is removed (which may or may not be desirable), it is replaced with those of the host, which may, in turn, modify the in planta transport properties of the protein (see Dreyer et al., 1999). Interaction studies (e.g. the split-ubiquitin two-hybrid yeast system (Obrdlik et al., 2004), which is favoured for membrane proteins, have been used to characterise potential regulators. And it seems likely that, in time, the regulation of transporter function in any system will be better understood by identifying and testing the candidate regulators pulled out in these screens. It should also be noted that protein–protein interactions can be (and should be) verified in planta with techniques such as FRET (or similar). Heterologous expression systems are generally high throughput, once the relevant cDNA is inserted into a suitable vector, and can express the gene of interest within a few days. Escherichia coli is generally not used for characterising eukaryotic membrane proteins, as incorrect folding or toxicity effects following overexpression are common, but has been used to identify and characterise several plant proteins (e.g. Uozumi, 2001). The most utilised system has been yeast (Saccharomyces cerivisiae, Schizosaccharomyces pombe or Pichia pastoris). Functional complementation of yeast strains deficient in certain transport pathways with plant cDNA libraries (or specific genes) is a proven system for identifying the encoded transport proteins from plants through simple growth assays (e.g. AKT1, Sentenac et al., 1992; H+ -ATPase, Palmgren and Christensen, 1993). Yeast can also be used to purify large amounts of protein ready for: reconstitution into liposomes (Lanfermeijer et al., 1997), the study of crystal structure (e.g. T¨ornroth-Horsefield et al., 2006) or the analysis of specific binding residues (i.e. phosphorylation sites; Maudoux et al., 2000). Site-directed mutagenesis of putative critical residues within proteins, followed by functional screens, has also revealed many insights into transport regulation (e.g. Johansson et al., 2006).

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Immature eggs (or oocytes) isolated from ovaries of the South African clawed toad (Xenopus laevis) provide another popular system for characterising transporters. X. laevis oocytes can synthesise proteins from injected cDNA (into the nucleus) or cRNA (cytoplasm) and target plant proteins to the PM (subcellular targeting peptides may have to be removed or added – e.g. Chen et al., 1999; see review by Miller and Zhou, 2000). Both yeast and X. laevis oocytes can be used for assays of; water permeability of aquaporins (by shadow imaging of X. laevis oocytes (Tournaire-Roux et al., 2003) or by spectrophotometer for yeast (Sakurai et al., 2005)), and electrogenic activity of co-transporters or ion channels and pumps by TEVC or patch clamping (e.g. Bertl et al., 1998; Gaymard et al., 1998; Liu et al., 2003). Electroneutral transport can also be studied by radioactive flux (see Section 4.4.3). Cell-cultured insect cells (Sf9/21) or mammalian cell lines (COS) provide other systems used for patch clamp-based characterisation (Dreyer et al., 1999). Plant heterologous and homologous overexpression systems (e.g. transient overexpression in tobacco mesophyll cells, or arabidopsis suspension cultured cells) have also been developed so that transport proteins can be properly processed and patch clamp electrophysiological characterisation can be easily applied (e.g. Bei and Luan, 1998).

4.7

Combining techniques (an example of increasing resolution and physiological context)

Many transport techniques load the dice in favour of viewing a certain transport activity of a protein (e.g. in ion channel selectivity experiments the ion mixtures used do not often relate to those seen in planta). This may result in the incorrect characterisation of the transport properties of the protein. To define properly the stoichiometry or selectivity of transporters and channels, it is mandatory that ion fluxes and ion currents across membranes be measured simultaneously. This can be achieved by combining voltage clamping with ISE-based flux measurement in both heterologous and plant systems (Kang et al. 2003; Gilliham et al., 2006b). Additionally, by combining these techniques it is then possible to conduct selectivity experiments in complex and physiologically relevant solutions.

4.8

Future development

High-throughput functional characterisation of proteins and their regulation is becoming a reality through automation (e.g. multichannel patch clamp (Wang and Li, 2003) or TEVC for X. laevis oocytes (Schnizler et al., 2003)). The use of complex computer-based algorithms has already been used for multiple transport-associated processes (e.g. simulating the molecular dynamics of channel gating; T¨ornrothHorsefield et al., 2006). However, it is becoming apparent that the signalling networks that regulate many transport processes, when integrated into the physiological framework of a functional cell (or plant), are too complex to comprehend without an

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iterative exchange between computer modelling and experimentation. It is therefore likely that computer-based modelling will be combined with experimental data with increasing frequency in the near future.

4.9

Conclusions

As is the norm for experimental science, for every interpretation of an investigational outcome there is often an alternative explanation due to some experimental caveat, and transport studies are certainly no exception. Intrinsic to every experiment is a compromise between the degree of invasiveness or removal from physiological reality and the extent of the biochemical, spatial and temporal resolution/definition achieved. Ultimately, it is the question that decides which technique should be used, but often the most reductionist and highly resolved techniques are those with the least physiological applicability (Tester, 1997). It is fortunate, therefore, that there is a whole host of techniques and resources at the experimenters’ disposal that, when combined, are adequate to address many unanswered questions in plant physiology. The importance of confirming a result reported using one assay with another technique to ascertain a physiological role cannot be overstated. Unfortunately, the choice of transport assay available to a researcher is still limited by expense, expertise and equipment (or its accessibility). All of these may be solved through collaboration (although boundaries and expectations should be clearly defined before entering into any such relationship). It would have been impossible to describe all techniques with sufficient detail for readers to fully understand any one technique in the confines of this chapter. Instead a snapshot has been presented that has hopefully provided some guidance to students and researchers alike, for ways in which transport assays can be used to better understand plant function.

Acknowledgements I am grateful to Bob Barrett, Romola Davenport, Mark Tester and Steve Tyerman for comments on the manuscript, Julia Davies and Tony Miller for guidance and members of the ‘transport group’ Department of Plant Sciences, University of Cambridge for being a source of knowledge and inspiration over the past 8 years.

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5 Transport across plant membranes F. J. Maathuis

5.1

Introduction

In the early nineteenth century, osmosis was discovered – a phenomenon that clearly pointed to a selective barrier between the outside and inside of a cell. Yet it was not until 1877 that the botanist Pfeffer proposed the membrane theory of cell physiology, which assumed the presence of a bag-like lipid structure with semipermeable properties (see Sections 3.3.2 and 4.2.1). This description gave a rudimentary basis for the selective influx and efflux of solutes at a cellular level. However, the suggestion that cell membranes (mainly) consist of lipids had to be revised in the light of the observed high permeability to charged particles such as K+ ions, and the concept was developed that membranes contain ‘pores’ dedicated to the movement of particular solutes. Thus, the notion of specific transport proteins was born and has been extensively refined over the ensuing decades, resulting in a generally accepted concept for cellular membranes: a fluid mosaic of lipids arranged in a bilayer interspersed with a plethora of proteins that function in transport, signalling and cell structure (see Section 4.2.2). Clearly then, it is a membrane, or more precisely the plasma membrane, that forms the barrier between ‘inside’ and ‘outside’ of the cell and, by extension, between ‘life’ and ‘death’. The selectivity and transport functions of membranes ensure that integrity is maintained not only at the cellular level, but also inside cells, where specific functions need to be compartmentalised. For a typical plant cell, this results in a complex system of membranes that includes the aforementioned plasma membrane at the cell periphery, a vacuolar membrane or tonoplast, mitochondrial (see Section 7.3), chloroplast (see Section 7.2) and peroxisomal (see Section 7.4) membranes and internal membranous structures such as the Golgi and ER systems. All these membranes, by their very nature of functioning as barriers, are heavily involved with the movement of solutes, but their transport mechanisms are tailored to specific inter- and intra-cellular requirements. It is obvious that, in many respects, chloroplastic thylakoid membranes involved in light reactions will contain different transport mechanisms from guard cell plasma membranes. The former will need redox-driven proton pumps to convert light energy into chemical energy, whereas the latter will need robust K+ and Cl− transport systems to generate large turgor changes. It is also apparent that plasma membranes in root cells, where soil nutrients are retrieved, will predominantly contain uptake systems for such minerals. On

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the other hand, xylem parenchyma cells that transfer minerals to the xylem for subsequent translocation to the shoot, are more likely to mediate a net efflux function. In addition, the energetics of transport will vary greatly. The flux of some solutes may already ‘want to go’ in the desired direction, for example due to a large diffusion gradient. The influx or extrusion of others may be against a gradient and therefore requires expenditure of energy. Thus, transport system composition, directionality and energetics all vary as a function of the solute itself, local cellular and intracellular requirements and environmental conditions. At a higher level, specific adaptations in plant species may also include transport systems that are found in one species but not another. In spite of the differences discussed above, there will also be many parallels: many basic cellular functions, including transport across membranes, are essentially similar irrespective of the cell type, species or even the kingdoms of life. For example, the conversion of trans-membrane H+ gradients into chemical energy in the form of ATP is mediated by F 1 F 0 H+ -ATP synthases that are essentially the same whether derived from bacteria, plants or humans.

5.1.1

Plant solutes

Plants move or transport many substances ranging from the simple hydrogen ion (H+ ) to very complicated organic compounds such as sugars and amino acids (see Chapter 2), and a broad division can be made between organic and inorganic solutes. This chapter will mainly focus on simple inorganic solutes which often constitute minerals that are essential to plants, but the principles discussed are equally valid for organic solutes. The primary inorganic solutes include both cations (e.g. K+ , Na+ , NH 4 + , Ca2+ and Mg2+ ) and anions (e.g. NO 3 − , Cl− , PO 4 3− and SO 4 2− ). Transport of other inorganic solutes such as trace metals and organic solutes such glycerol, sugars, amino acids and xenobiotics is also of crucial importance to plant physiology but these will only be mentioned briefly.

5.1.2

Definitions and terminology

For mechanisms of membrane transport, several systems of classification exist. Often these have historical origins and, although they persist, they are not always still appropriate. In addition, none of the classifications is unambiguous, which can lead to confusion. An example is the division between ‘transporters’ and ‘ion channels’ based on outdated assumptions that carriers and pumps (see below) bind a substrate and subsequently ‘transport’ it to the opposite side of the membrane, whereas ion channels simply provide a diffusive pore. It is now known that channels also bind their substrate and that the F 0 part of the F 1 F 0 H+ -ATPase can function as a H+ channel. Indeed, many formalisms that describe the kinetic behaviour of channels can equally be used to describe carriers and pumps and vice versa. The convention adopted here will be that any membrane protein involved in cross-membrane movement of substrates is a transporter.

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The first useful classification of transporters refers to active transport and passive transport (Colour plate 5.1a), a distinction that is often made to refer to the direction of movement of a solute with respect to its electrochemical gradient. Left to its own device, any solute will move from a higher to a lower electrochemical potential, or ‘down hill’ in energetic terms. When such movement occurs across membranes it is referred to as passive transport. In contrast, to move a solute from lower to higher electrochemical potential, energy input is required and such ‘up hill’ transport is therefore termed active. There are two potential sources of confusion with these definitions of active and passive transport that can generate misconceptions: frequently ‘passive’ and ‘active’ are believed to mean ‘no energy necessary (i.e. ‘free transport’) and ‘energy necessary’. Although the latter is true, passive transport, as we shall see below, often dissipates trans-membrane gradients that have to be maintained at the expense of energy. In other words, although the transport of a particular solute may in itself be passive, it often indirectly requires energy expenditure by a cell. Secondly, the terms passive and active, which strictly refer to thermodynamic properties of solute fluxes, are regularly associated with specific transport mechanisms. Clearly, passive transporters can never mediate active transport, this would violate the second law of thermodynamics, but active transporters can, and probably will in many conditions, function as purely dissipative and therefore as passive systems. Indeed, transporters have been described that can switch between active and passive modes such as the mitochondrial triose phosphate/nucleotide sugar transporter. The second classification refers to primary and secondary transport (Colour plate 5.1b). Primary transport moves solutes against their electrochemical gradient at the expense of ‘metabolic’ energy. The latter can be redox energy, for example in the form of pyruvate in the mitochondrial respiratory chain, which is converted into an electrochemical proton (H+ ) potential by redox-energy-driven pumps, such as cytochrome oxidase. It can also be in the form of solar energy, used to generate H+ gradients during photosynthesis. In its most familiar form, primary transport uses chemical energy derived from compounds such as ATP. In mammalian cells, the primary K+ -Na+ -ATPase uses around 25% of all generated ATP to maintain transmembrane K+ and Na+ gradients. In plants, ATP-fuelled primary transport across plasma and vacuolar membranes generates transmembrane electrochemical H+ gradients (Colour plate 5.1b). The energy stored in this ‘H+ battery’ can then be used to transport further solutes via specific carriers, which couple down hill reflux of H+ to the movement of other solutes. Such mechanisms, where dissipation of a primary pump-maintained gradient is coupled to the movement of another solute, are referred to as secondary transport (Colour plate 5.1b). The latter can be active, when the substrate moves up its electrochemical gradient, or passive, for example when ions move down their electrochemical gradients through ion channels. In plants, primary transport in the form of ATPase pumps typically establishes H+ gradients and, unsurprisingly, most coupled secondary transport uses these H+ gradients to move other solutes. In animals, the predominant primary pump extrudes Na+ from the cytoplasm and, consequently, most secondary transport in animals is run on ‘sodium batteries’.

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The third classification system is based on the transport mechanism of transporter proteins themselves and includes three main types: pumps, ion channels and carriers (Colour plate 5.1c). Pumps normally are active systems that use chemical, light or redox energy to move substrates against their electrochemical gradient, typically at a rate of a few hundred per second. They fulfil many functions in cells such as generating gradients for secondary transport, extruding cytoplasmic Ca2+ and maintaining ionic homeostasis. Ion channels are purely dissipative and therefore genuinely passive transport systems. They can move substrates (ions) at astonishing rates of several million per second and are ideally suited to function in fast processes such as Ca2+ signalling, propagation of impulses along nerve cells and relaying sensory input. The large transport capacity of ion channels means they are also ideally suited to processes that need movement of large quantities of ions, such as the uptake of important plant nutrients, maintaining turgor and in turgor driven cellular movement. The third group, carriers, forms a rather non-descript collection of transport mechanisms best defined as ‘not being a pump or a channel’ and includes secondary active transporters. The carrier concept stems from classical enzyme kinetics describing rate constants and affinities of substrate binding and release sites on either side of a membrane. Carriers can mediate passive or active transport and a valid question is therefore why cells would need two forms of passive transport system: both ion channels and passive carriers. The answer may have to do with the way different types of transporter confer selectivity for their substrate. Rapid progress has recently been made in determining crystal structures of various membrane transport proteins and this has shed much light on the mechanisms underlying ion selectivity (Gouaux and MacKinnon, 2005). Broadly generalising, ion channels contain large water filled pores as part of their architecture, which allows free travel for (hydrated) ions well into the membrane bilayer. The selectivity filter, which only spans a fraction of the bilayer thickness, is the only restriction to free diffusion and is the only site in passing through the channel at which ions are dehydrated. The combination of electrostatic interactions of the dehydrated ion with charged side chain groups and the filter size determines ion selectivity in ion channels. In contrast, pumps and carriers often lack solvent-filled cavities. In these cases, dehydrated ions are forced to follow a predominantly proteinaceous pathway where conformational changes impact the affinity of binding sites in the carrier. This arrangement leads to considerably slower transport when compared to ion channels but precludes any chance of opening ion pathways to both sides of the membrane at the same time (Gouaux and MacKinnon, 2005). Thus, although channels can be remarkably selective for specific ions, selectivity is nevertheless largely based on the simple concept of size exclusion. Carriers and pumps have ‘proper’ binding sites based on three-dimensional geometry of enzyme and substrate. The latter can therefore have much higher substrate affinities and be far more specific. For example, carriers can discriminate between different sugar molecules in spite of similar sizes or they can distinguish between l and d stereoisomers of amino acids. This high degree of selectivity comes at a cost

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of lower turnover rates, hundreds or thousands per second, compared to millions per second for ion channels. The last classification, which often occurs, refers to another aspect of transport mechanisms namely the question whether only one or more than one substrate is transported (Colour plate 5.1d). In a uniport system, only one type of substrate is moved from one side of the membrane to the other. Uniport mechanisms are therefore by definition passive systems and include all ion channels and a number of carrier systems. In a symport mechanism, the transport of two or more substrates is coupled to each other and all substrates are moved in the same direction. When there is counterflow of substrates a system is referred to as antiport. An example of a uniport system is an ion channel: one type of substrate moving in a file through the channel pore in one direction (in fact there are many ion channels with multi-ion pores that transfer different types of ion at the same time but there is no strict coupling between the two types of ion). Symporters are normally active transport systems that use downhill movement of, for instance H+ , to drive transport of other substrates. Good examples are the plant sucrose:H+ symporters that move sugars from one compartment to another. Many antiport systems have been described in plants; one of the best characterised is NHX1, a system that exchanges one H+ for one Na+ ion, and functions in the vacuolar sequestration of Na+ .

5.1.3

Some formalisms

To understand the distinctions made by various classification systems and how cells run their membrane transport economy, it is necessary to examine some of the fundamental physical concepts that deal with diffusion, energetics and electrochemistry. The action of ATP driven primary H+ pumps in plant cells – and K+ -Na+ pumps in animal cells – has as one corollary that cell interiors are negatively charged (but some other factors also contribute to this phenomenon) with respect to the extracellular compartment. Therefore, cells maintain a voltage difference across the plasma membrane, which in plant cells is typically between −100 and −200 mV. Smaller transmembrane voltages exist across the tonoplast, many plastid membranes and probably endomembranes of the ER and Golgi, although the latter have not been determined. The existence of a trans-membrane voltage (E m ) has large implications for transport of charged solutes across these membranes. The driving force for transport of ions across membranes is determined by two parameters: the chemical gradient, i.e. the difference between the external and cytoplasmic concentration, and the electrical gradient, i.e. the cell membrane voltage. Combined, these two forces are expressed as an electrochemical ion potential (μ). If μ is lower outside the cell than inside, uptake of the ion will require energy and thus has to be active. If μ is higher outside, its uptake can proceed passively (but it does not necessarily do so). At equilibrium, conditions are such that the electrochemical potential is zero, which implies that the chemical and electrical forces are opposite and cancel each other out.

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Now, a short digression into the physics of diffusion. Because of the kinetic energy, molecules are constantly moving (unless the temperature is at absolute zero). This inevitably means that any collection of molecules will distribute itself randomly across all possible positions in space. So if it were physically possible to introduce a number of gas molecules in one corner of an enclosed container, before long the collection of molecules would have spread throughout the entire space and, this being the equilibrium state, would remain so. Thus, diffusion can be formalised as a tendency of any substance to equalise its distribution. This in turn means that a concentration difference of any substance will dissipate if there is free movement of particles. Diffusion through a barrier such as a biological membrane will depend on two parameters: (i) the magnitude of the concentration difference and (ii) the permeability of the membrane for that particular substance. Permeability itself is governed by the resistance of the membrane and its thickness. Clearly a very ‘tightly’ constructed membrane will have a larger resistance than a porous membrane. Similarly, a very thick membrane impedes flow of particles more than a very thin membrane. These properties are incorporated into the diffusion coefficient (D, in m2 sec−1 ), which is specific to any particle and the medium the particle diffuses through. Overall, these concepts are formalised in Fick’s (first) law (see also Chapter 3): Js = −Ds

dCs , dx

(5.1)

which intuitively states that the flux Js of a solute s depends on its diffusion coefficient Ds and its concentration gradient (the variation of Cs per change in space x). Secondly, since ions are solutes with a charge their movement is influenced not only by the diffusion properties described in Eq. 5.1 but also by any electrical field. This force–flux relationship is analogous to that mentioned above for diffusion, only the driving force is the electrical field rather than a concentration gradient and the proportionality constant is slightly different. Js = −μs Cs

dE , dx

(5.2)

where μs is a constant analogous to Ds and dE/dx is the electrical gradient or electrical field. Nernst and Planck incorporated this term, describing flux as a function of the electrical field that acts on a charged particle, into Fick’s law resulting in the Nernst-Planck ‘electrodiffusion’ relationship: Js = −Ds

dCs dE − μs C s . dx dx

(5.3)

In other words, (electro)diffusion of a charged particle depends on its chemical gradient and on the electrical field (voltage difference). When ion transport is studied, it is often the current (flux of charge per unit time) rather than the flux that is the point of interest, since it is often current that is the experimental parameter under investigation and Eq. 5.3 can be multiplied by Faraday’s constant F (see Section 3.5.1) and ionic valence of S, z s , included to yield

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a relationship that expresses ionic current as a function of concentration and voltage:   dCs F z s Cs d E Is = −z s FDs + , (5.4) dx RT d x where R is the gas constant and T is the absolute temperature. At equilibrium, Is = 0 and after rearranging and integration, Eq. 5.4 reduces to the Nernst equation (Section 3.5.4):  + S RT E= ln + out , (5.5) zs F [S ]in which shows that at equilibrium the concentration ([S+ ] out /[S+ ] in ) and electrical force (E) are equal and opposite thus cancelling each other out. At equilibrium, i.e. the condition where the net flux across the membrane is zero, E (the ‘Nernst potential’) is exactly the voltage where the current reverses. This is often referred to as the Nernst potential of a specific ion or simply as E K for potassium or E Na for sodium. We can rearrange Eq. 5.5 and substitute room temperature and z = 1 to get:  + I (5.6) E = 58 log + out , [I ]in for a monovalent cation. We can now see that a 10 times concentration gradient is equivalent to around 60 mV electrical force or, vice versa, that 120 mV electrical force equates to a 100 times chemical gradient. For example, if we assume a typical value for the membrane potential E m of –120 mV, we can immediately derive that passive K+ accumulation (or of any other monovalent cation) is limited to 100 times [K+ ] ext . K+ accumulation to levels higher than 100 times [K+ ] ext therefore requires alternative modes of energisation and relies on active transport. Equation 5.4 can also be used to derive another important formalism that is frequently applied to assess selectivity of membranes or of specific transport systems (Chapter 3). The so called Goldman Hodgkin Katz voltage equation gives the membrane potential (E rev ) at which no net current flows. This parameter is experimentally relatively easy to obtain and, in combination with known ionic concentrations of the permeant ions, for example K+ , Na+ and Cl− , can be used to resolve the relative permeabilities P K , P Na and P Cl according to:   PK [K]o + PNa [Na]o + PCl [Cl]i RT ln E rev = . (5.7) zF PK [K]i + PNa [Na]i + PCl [Cl]o

5.2 5.2.1

Passive transport Diffusion through membranes

Passive transport mediates a solute flux in the direction of a lower (electro)chemical potential. Diffusion is therefore by definition a passive process. Nevertheless, it is a crucially important phenomenon for all life. For example, diffusion of inorganic

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minerals in the soil solution is often a limiting factor for plant nutrition, whereas many smaller cells (e.g. bacteria) rely on diffusion for the cytoplasmic distribution of solutes. Substances can cross membranes by diffusion if they can dissolve in the oily interior of the membrane. Such lipophilic (or hydrophobic) substances include important compounds like O 2 , CO 2 , H 2 O 2 and NH 3 . For example O 2 –CO 2 gas exchange in lungs and in photosynthesising plant tissues operates by this process (Colour plate 5.2a). Another example is the plant hormone ethylene, which plays important role in stress response and in fruit ripening. These processes rely on the physical properties of lipid membranes and on the chemical and physical properties of the diffusing molecule. Such processes do not involve specific proteins (Colour plate 5.2a) and are therefore not submitted to any significant level of regulation.

5.2.2

Facilitated diffusion through carriers

An obvious disadvantage of relying on simple diffusion is the total lack of control which cells can exert on the flux of substances that move by this process. Organisms have therefore developed specific transport systems that allow diffusion across membranes to occur through dedicated proteins. Clearly, such systems can function only when the overall membrane permeability for the particular compound is relatively low, which is certainly the case for charged particles (ions) and also for many important substances such as sugars. Plant cells use facilitated diffusion through carrier-type transporters, through ion channels and through water channels. Ion and water channels will be dealt with in more detail below and the focus here is on some plant carrier systems that mediate facilitated diffusion. Nitrogen is required by plants in vast quantities and many groups have studied the mechanism and identity of nitrogenous compounds that are taken up by plant roots. Most plants acquire N in either of two forms: as nitrate (NO 3 − ) or ammonium (NH 4 + ) ions, or a mixture of the two. The translocation of NH 4 + is mediated by proteins of several transport families each containing multiple isoforms. Recently, one isoform of the tomato AMT (ammonium transporter) family (LeAMT1;1) was heterologously expressed in oocytes of Xenopus laevis (Ludewig et al., 2002), a convenient system to perform voltage clamp experiments (see Section 4.4.5.2). Inward current could be observed when micromolar amounts of NH 4 + were added to the external medium showing (i) net positive charge was entering the oocyte interior and (ii) the transporter has a high affinity. Subsequently, the authors showed that the inward current was not sensitive to any other ion which might act as a coupled driving force. The conclusion, therefore, was that NH 4 + is transported into the cell via a carrier-type uniport mechanism (Colour plate 5.2b), an excellent example of facilitated diffusion. A second example of a putative carrier that mediates facilitated diffusion is HKT1. HKT1 is probably expressed in the plasma membrane and contains seven to nine transmembrane domains (TMD). In arabidopsis, there is only one HKT isoform, but in other species such as rice, the HKT family extends to seven or eight members. Initially, HKT1 was characterised as a K+ :Na+ symporter (Rubio

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et al., 1995), but it has become increasingly evident that at least in arabidopsis, HKT1 only transports Na+ (M¨aser et al., 2002; Berthomieu et al., 2003). Thus, although there may be conditions where HKT1 can function as a K+ :Na+ symport mechanism, it appears that in vivo, HKT1 binds a Na+ ion to both binding sites which makes it kinetically equivalent to a uniport mechanism that mediates facilitated Na+ diffusion. The physiological relevance of this system is still under debate. Antisense expression of this transporter in wheat led to a decrease in unidirectional Na+ influx (Laurie et al., 2003) providing direct evidence that it functions as a root Na+ uptake mechanism. In arabidopsis, there is the suggestion that AtHKT1 may function in Na+ translocation from root to shoot and vice versa (M¨aser et al., 2002; Berthomieu et al., 2003).

5.2.3

Transport through ion channels

Ion channels (Colour plate 5.3) are integral membrane proteins that allow ions to pass when the protein is in the open state. Channels function as regulated pores and, simplistically put, all they do is be open (conduct) or closed. So, in contrast to many carriers and all pumps, ion channels are purely passive transporters. Opening and closing, or gating, of ion channels can depend on many factors but roughly speaking there are voltage-gated and ligand-gated channels and a third group is formed by mechanosensitive channels. Voltage-gated channels (Colour plate 5.3a-c) ‘sense’ the membrane potential and increase/decrease their open probability (Po) as a function of the membrane potential, whereas ligand-gated channels (Colour plate 5.3d) only open after binding an effector molecule, the ligand, which alters the protein conformation and thus leads to channel opening. Mechanosensitive channels play important roles in mammalian functions such as touch and hearing, whereas in plants they are envisaged to sense processes such as turgor-related changes in cell volume. In all ion channels, the pore domain constitutes the transmembrane aperture through which ions are conducted. The pore is usually water-filled, which makes it an attractive environment where ions can diffuse across the membrane. One region in the pore forms the selectivity filter (Colour plate 5.3a), an area that determines which ions are allowed through the pore. There are three main characteristics that define an ion channel: channel conductance, channel selectivity and channel gating (Hille 2001). Conductance G is the reciprocal of resistance R defined in Ohm’s law as: V = IR

or

V = I /G

(5.8)

with V the voltage expressed in volts (V) and I the current expressed in amperes (A). Conductance is expressed in siemens (S) and for ion channels is typically in the range of pS. The larger the conductance, the bigger the flux of ions that can pass through the channel. Often ion channel nomenclature refers to the most permeant ion, or the most permeant and physiologically relevant ion. Which ion permeates will depend on channel selectivity, which is typically defined by the selectivity filter, an area that provides a physical barrier and filters on the basis of ion size but also on the basis

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of electrostatic interactions between the permeating ion and charged residues of the channel protein. Thus, cation channels typically show negative residues lining the channel pore, whereas anion channels contain positive charges in this region. In voltage-gated channels, opening of the pore strictly depends on changes in membrane potential E m . A decrease in membrane polarisation, depolarisation, stimulates opening of depolarisation-activated channels (Colour plate 5.3b), whereas increased membrane polarisation, hyperpolarisation, leads to opening of hyperpolarisation-activated channels. The link that connects changes in E m to channel gating is provided by a voltage sensor domain (Colour plate 5.3a), typically a transmembrane alpha helix with a large number of charged residues. Under the influence of changes in E m , movement of this domain translates into conformational changes in the channel protein leading to opening or closing of the pore. Although opening of voltage-gated channels requires a change in E m , there may be many factors other than membrane voltage that impact on the channel open probability, such as the presence of second messengers, the redox state or the phosphorylation state of the channel protein. The ligand-gated ion channels constitute another main class of ion channel. In this type of transporter, gating occurs only after binding of specific compounds to the channel protein (Colour plate 5.3d). These compounds are called ligands, i.e. substances that bind to another compound to form a complex. The term agonist (literally meaning a ‘competitor’) is also frequently used since often different ligands can have similar actions and compete for the same binding site. In contrast, antagonists bind at the same site but have an opposite action to ligands. Ligand binding to the channel protein causes a conformational change that switches the channel from the closed to the open state or vice versa. Ligand binding is a minimum requirement for channel gating but, as for voltage-gated channels, overall gating properties of ligand-gated channels are often modulated by additional parameters which can include membrane polarisation (Hille, 2001). In the following section, some of the major classes of ion channel found in plants are briefly reviewed.

5.2.3.1

Potassium channels

Potassium (K+ ) channels were the first class of ion channel described in plant cells and it was found that these consisted of two major categories, inward and outward rectifying (Maathuis et al., 1997; see also Colour plate 5.4a). The inward K+ channels are activated upon membrane hyperpolarisation and form a major pathway for K+ uptake, whereas the open probability of outward channels increases when the membrane depolarises, leading to loss of K+ from the cell. Examples of major inward and outward rectifying voltage dependent channels have been cloned (see Very and Sentenac, 2001 for a review) and their overall structure resembles that of mammalian Shaker-type channels with six TMD, a voltage sensing domain in the S4 region and a pore with selectivity filter in the S5–S6 part of the channel (Colour plate 5.4b). The selectivity filter contains the conserved GYGD motif, a sequence found in virtually all ion channels that are selective for K+ , and the S4 region ensures that these K+ channels require a change in E m to transit from closed to open state. Functional channels are made up of four subunits either as homomers

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or as heteromers. Apart from Shaker-like channels, the major families of plant K+ channels also contain two or four TMD-structured channels and voltage-independent K+ channels.

5.2.3.2

Calcium channels

Ca2+ channels have also been intensely studied in plants. In contrast to mammalian Ca2+ channels, plant Ca2+ channels are usually not very selective for Ca2+ and will also conduct many other cations such as K+ (White, 2000). Plant Ca2+ channels have mainly been characterised at the tonoplast where they are believed to play a role in Ca2+ signalling. These vacuolar Ca2+ channels consist mainly of ligand-gated channels that require IP 3 , NAADP or cADPR to be gated (White, 2000). However, one putative Ca2+ channel (TPC1) is voltage gated and has a 12 TMD structure (Peiter et al., 2005).

5.2.3.3

Non-selective ion channels

Non-selective ion channels allow passage of various ions and are often permeable to mono- and divalent ions. Plant non-selective cation channels have been characterised to some extent and generally found to be voltage independent (Demidchik et al., 2002). Although some non-selective cation channels have been shown to be regulated by cyclic nucleotides (Leng et al., 2001; Maathuis and Sanders, 2001; Balague et al., 2003), it is often not clear how their gating is controlled. These channels are believed to be important in signalling, in turgor regulation and in cation nutrition. They are of special relevance regarding plant stress since they may form a major conduit for the entry of toxic ions such as Na+ (Demidchik et al., 2002). Two gene families, cyclic nucleotide gated channels (CNGCs; Talke et al., 2003) and glutamate receptors (GLRs; Davenport, 2002), are believed to encode non-selective channels and are found in many plant species.

5.2.3.4

Chloride channels

Only one family of anion channels has been described in plants (Hechenberger et al., 1996). Members of the ChLoride Channel (CLC) family show large homology to their mammalian counterparts that are mainly involved in cellular volume regulation. Plant CLCs contain 10–12 TMD and are strongly sensitive to E m with a P o vs. V relationship that is bell shaped. Little is known about the function of plant CLCs, but they are probably involved in early events during cell signalling which frequently involve a Cl− efflux, and possibly in nitrogen nutrition since some CLCs are also capable of transporting NO 3 − (Geelen et al., 2000). A further function may be in turgor regulation and turgor-driven movement when large amounts of both cations and anions are moved across tonoplast and plasmamembrane.

5.2.4

Transport through water channels

In all forms of life, water is the solvent for cellular solutes. In addition, water in plants is necessary to generate turgor and to provide a medium for mass flow of solutes (Chapters 9 and 10). Indeed, movement of water is intricately linked to that

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of ions and, as is the case for most ions, its transport is regulated and controlled. However, in contrast to ions, there are no known active transport mechanisms for water and thus water movement is always passive and directed towards a lower water potential. Although water permeability of phospholipid bilayers is substantial, it is now clear that biological membranes contain specific protein-based pathways for the movement of water. These water channels or aquaporins constitute a parallel and regulated pathway for water flux, both at the intracellular and the whole plant level (Chrispeels et al., 2001). Structurally, aquaporins have a protein topology that is very similar to Shakertype ion channels, with each subunit having six TMD. However, rather than one, aquaporin subunits have two pore-forming loops: one in the first half of the protein between TMD two and three and the other between TMD five and six (Colour plate 5.5). The second loop contains a cysteine residue where mercury can bind, leading to channel blockage. The aquaporin-specific signature motif NPA (asparagine, proline, alanine) is involved in the stabilisation of the pore-forming loops. Similar to Shaker ion channels, functional aquaporins consist of tetramers in which the eight pore loops combine to form transmembrane water-conducting paths. The water-conducting pore is around 0.30 nm wide at its narrowest point, a very close fit to the 0.28 nm size of a water molecule, and thus an efficient barrier to other substances. Apart from size exclusion, pore residues interact with the permeating water molecules mainly through H+ bonds. These structural properties ensure that aquaporins are highly selective for water molecules, which move through the protein in a single file, and that water channels are virtually impermeable to any charged species. The latter include protons, a remarkable property since protons can usually be transferred readily through water molecules. Despite this high level of substrate specificity, aquaporins can sustain very high transport rates of ∼3 × 109 water molecules per second. Although aquaporins more or less completely exclude charged species and are highly selective for water, several have been shown to be capable of transporting small non-charged solutes such as glycerol and urea. Sometimes a distinction is made between water-selective aquaporins and those that can conduct water and small solutes, the aquaglyceroporins. Comparative studies using an Escherichia coli isoform of each group showed that although both proteins were structurally and genetically highly similar, minor changes in the pore of the aquaglyceroporin result in a slightly wider selectivity filter and hence glycerol permeability (Wang et al., 2005). In plants, aquaporins are predominantly expressed in vacuolar and plasma membranes and fall into four major classes, of which some are related to their membrane location. PIPs (plasma membrane intrinsic proteins) are expressed predominantly in the plasma membrane and are subdivided into the PIP1 and PIP2 subfamilies (Johanson et al., 2001). In the tonoplast, TIPs (tonoplast intrinsic proteins) are present, with some isoforms specifically targeted to storage vacuoles, and others to lytic vacuoles. NIPs (NOD26-like intrinsic proteins) have largely unknown membrane locations apart from NOD26 itself, which is expressed in root nodules. A fourth class comprises the SIPs (small basic intrinsic proteins), whose membrane location is also largely unknown (Johanson et al., 2001). These four classes tend to be members of

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big gene families with around 35 members in the model species arabidopsis. Such large gene families might suggest that functional redundancy is common for water channels but could also point to a need for isoform-specific expression patterns of aquaporins with finely tuned functional adaptations. There is now good evidence that both transcriptional and post-transcriptional regulation of aquaporin activity takes place in plant membranes (Luu and Maurel, 2005). For example, root hydraulic conductivity can be seen to correspond closely to PIP1 mRNA levels in Lotus japonicus (Johanson et al., 2001) and the diurnal action of motor cells that move Samanea saman leaves closely mirrors levels in PIP2 (Johanson et al., 2001). In response to salt stress, root PIPs and TIPs are rapidly down-regulated but with a time difference between PIPs and TIPs (Maathuis et al., 2003). Post-translationally, both aquaporin glycosylation and phosphorylation appear to be major mechanisms for the regulation of hydraulic conductance. While glycosylation may be involved in the recruitment of protein to the relevant membrane (Vera-Estrella et al., 2004), phosphorylation directly enhances channel activity (Chrispeels et al., 2001; Johanson et al., 2001). Other factors that impact on aquaporin activity are H+ and Ca2+ with both these ions having a blocking effect on water transport through aquaporins.

5.3

Primary active transport

In many conditions, cells require solutes in quantities that cannot be achieved through passive transport. Similarly, many waste products need to be removed or compartmentalised often against their electrochemical potential. This transport can, by definition, not occur via passive transport and requires active systems that directly use energy. A further fundamental requirement of all living cells is the capacity to convert chemical energy into electrochemical energy and vice versa. Primary active transport mechanisms convert metabolic energy to move substrates against a gradient. At the same time, many primary systems, through their activity, energise membranes by establishing electrochemical potential differences. The latter can subsequently be used to energise secondary active transport (Section 5.4).

5.3.1

Primary proton pumps

The majority of chemical energy that is generated from processes such as respiration and photosynthesis is deposited in the phospho-ester bonds of compounds like adenosine triphosphate (ATP), guanyl triphosphate (GTP) and pyrophosphate (PPi). Hydrolysis of the phosphate-ester bond is highly exergonic; for example the conversion of ATP to adenosine diphosphate (ADP) has a G◦ of –32 kJ mol−1 . This energy can be converted into other forms such as an electrochemical gradient. In plant cells, a large amount of ATP is used for transmembrane H+ movement. Extrusion of H+ from the cytoplasm has physiological rationales; for example, for cellular pH regulation, the acidification of cell walls and rhizosphere, or to lower

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the pH in lytic compartments. Nevertheless, the principal function of plant H+ ATPases, or H+ pumps, is in the generation of transmembrane H+ gradients. The pH and electrical potential differences of the H+ gradient can both be used in subsequent transport processes. Plant proton ATPases, as most other pumps, have very low turnover rates of ∼100 H+ per second. This and their crucial role in cellular physiology mean proton ATPases are very prolific enzymes that can make up several percent of total protein. They are predominantly found in the plasma membrane and tonoplast and also in other endomembrane systems such as ER and Golgi complex (see also Chapter 7). Often the level of expression of these pumps is related to physiological conditions and function of tissues and cell types.

5.3.1.1

P-type ATPases

The proton pump located in the plasma membrane is a P-type ATPase (see Geisler et al. (1999), Sze et al. (1999) and Axelsen and Palmgren (2001) for reviews) so called because during its catalytic cycle, phosphorylation of a specific site at the enzyme is essential (Colour plate 5.6). In plants this enzyme generates a proton motive force (PMF) across the plasma membrane, typically in the region of 250–300 mV comprising a membrane potential of around –150 mV (negative inside the cell) and a pH of around 2 units (acidic outside the cell), which is equivalent to –120 mV. Both components of the PMF are used to energise movement of many important nutrients and metabolites through active H+ -coupled transport. In addition, the activity of P-type ATPases is a dominant contributor to membrane polarisation and therefore impacts greatly on the driving force for passive transport. The G◦ for ATP hydrolysis is around 32 kJ mol−1 . This would be equivalent to around –330 mV if it is assumed that 1 ATP is hydrolysed per H+ pumped, since G◦ = −nF E◦. with the stoichiometric coupling ratio n equal to unity, a conversion factor between mV and kJ mol−1 , the Faraday constant F, of 96.5 kJ V−1 mol−1 and E◦ the electropotential in V. However, in physiological conditions, the G for ATP hydrolysis is considerably higher and in the region of 50 kJ mol−1 , theoretically sufficient to generate a PMF of over –500 mV if n were 1. Experiments where H+ fluxes were compared with ATP hydrolysis rates indeed showed that 1 H+ is pumped for every ATP hydrolysed. P-type proton pumps are encoded by multiple-gene families and the enzyme functions in the membrane as a single polypeptide of around 100 kDa. Structurally, P-type pumps contain 10 TMD with the TMD4–TMD5 cytoplasmic loop containing the ATP binding site. The same loop also has a conserved aspartyl residue that becomes phosphorylated and dephosphorylated during every catalytic cycle. Toxins such as vanadate and arsenate mimic phosphate and inhibit the enzyme by binding to the phosphorylation site. The C-terminal residues of the protein function as an autoinhibitory domain. Cleavage of this ∼100 amino acid tail by trypsin, or expression of the shortened mutant protein, leads to activation of the enzyme. The same domain also interacts with 14-3-3 proteins that can bind to it and, through unknown mechanisms, activates the ATPase (Colour plate 5.6). The fungal toxin fusicoccin

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is believed to stabilise the interaction between ATPase and 14-3-3 and thus activate the protein.

5.3.1.2

V-type ATPases

At the tonoplast and various endomembranes, vacuolar or V-type ATPases extrude H+ from the cytoplasm. Recently, V-type ATPases have also been localised to the plasma membrane. In contrast to the P-type, the V-type ATPase protein is remarkably complex requiring the assembly of 14 different subunits (Colour plate 5.7) and has an overall weight of ∼700 kDa. The holocomplex consists of a large head portion, V 1 , where ATP is bound and split. The basal part, V 0 , of the complex protrudes through the membrane and forms the H+ channel. In F 1 –F 0 ATP synthases, the gamma subunit is part of the stalk which transduces potential energy, from H+ travelling through the H+ channel, into ATP synthesis. How the link between the chemical (V 1 ) and electrochemical (V 0 ) energy is precisely structured in V-ATPases is unknown but it is believed that V-type ATPases are very similar to F 1 –F 0 ATP synthases, the enzymes found in mitochondria and chloroplasts that convert redox energy into chemical energy in the form of ATP (Kluge et al., 2003). V-type ATPases are believed to function as rotary motors, similar to their F 1 –F 0 ATP synthase counterparts, with the catalytic A and B subunits driving rotation of a central shaft (subunits D and F), which causes the membrane-bound c ring to rotate. Protons are picked up by the spinning c ring, pumped through conductance channels in the membrane bilayer, and released into the vacuolar lumen. The exact stoichiometry of this process is not known and may vary according to conditions but is believed to be 2 to 3 H+ per ATP hydrolysed. The PMF generated by V-ATPases is particularly important for the maintenance of passive and secondary transport across the tonoplast. The vacuole, with its large volume fraction, constitutes a crucial compartment for storage of ionic and nonionic nutrients and metabolites. In addition, it is the compartment where potentially harmful ions and substances are deposited to safeguard the cytoplasm. All these functions require tightly regulated, and often energised, transport mechanisms at the tonoplast. The trans-tonoplast PMF consists almost entirely of a pH: of the total PMF of around 200 mV, E m is typically no more than 20–30 mV (the vacuolar lumen being positive with respect to the cytoplasm) and the remainder in the form of pH (the lumen being acidic with respect to the cytoplasm). As is the case for the P-type ATPase, V-type ATPases will be expressed in tissueand cell-type-dependent patterns and both transcriptional and post-transcriptional regulation occurs in response to endogenous and external conditions. For the V-type ATPase, this is particularly well documented when plants are exposed to salinity and other abiotic stress (Dietz et al., 2001). Tonoplast-enriched vesicle preparations showed considerably higher ATPase and H+ pumping capacity when isolated from salt-stressed plants rather than plants growing in non-saline conditions (Blumwald and Poole, 1985; Blumwald and Poole, 1987; Staal et al., 1991). Later it was also shown that mRNA levels of many V-ATPase subunits are upregulated after salt stress (e.g. Maathuis et al., 2003). These observations are interpreted as an extra demand

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for tonoplast energisation to drive the substantial H+ -coupled sequestration of Na+ ions in the vacuole (see also Section 14.12).

5.3.1.3

The pyrophosphatase

Plants are one of the few organisms that contain a second type of proton pump at the tonoplast, the pyrophosphatase or PPase. This enzyme breaks down pyrophosphate (PPi) and employs the released energy to move H+ into the vacuolar lumen. This enzyme therefore works in parallel to the V-type ATPase and it remains a question why cells would require such an arrangement. The main purpose of the PPase may be to extract the considerable amount of energy that is available in the form of PPi, a by-product of cellular metabolism that is particularly ubiquitous in young developing tissue. On the basis of sequence homology, plant PPases are divided into type I or type II. The strong dependence of type I PPase activity and reversal potential on cytoplasmic K+ has led to the suggestion that type I PPases pump K+ ions at the same time as H+ (Davies et al., 1994) although this concept is still debated. Type II PPases show no requirement for cytoplasmic K+ . PPases consist of a single catalytic subunit with 13–16 TMD and a MW of around 80 kDa. They may function as homodimers in vivo (Maeshima, 2000). They are generally encoded by small gene families and PPase regulation has been studied in several plants and conditions to gain better insights into the role of this enzyme. For example, PPase expression is induced by conditions such as anoxia, and overexpression of PPase has led to plants showing increased salt and drought tolerance (Gaxiola et al., 2002). In physiological conditions, the free energy release of PPi hydrolysis is around 27 kJ mol−1 though MgPPi rather than PPi itself is the actual substrate. It is generally assumed that the PPi hydrolysis to H+ pumping ratio is one and this stoichiometry could therefore generate a PMF of around 300 mV.

5.3.2

Primary pumps involved in metal transport

Plants need to acquire many metals that play structural roles, are important cofactors to many enzymes, and participate in cellular signalling. Often such metals are required only in small quantities, but their low abundance in the environment makes the presence of high-affinity transport mechanisms a necessity. On the other hand, many conditions, such as polluted soils, may present a danger of metal toxicity and plants therefore also need adequate metal efflux mechanisms.

5.3.2.1

P-type Ca2+ pumps

Extracellular Ca2+ concentrations usually exceed 1 mM. In contrast, cytoplasmic free Ca2+ levels need to remain extremely low to prevent precipitation of Ca2+ phosphates. Cytoplasmic Ca2+ is therefore typically kept in the nanomolar range. The extremely low level of cytoplasmic Ca2+ may have led to the adaptation of this divalent ion as an important signalling intermediate. Indeed, a plethora of internal and external stimuli is known to evoke Ca2+ signals in plants, stimuli such as touch, pathogen attack or drought stress. During Ca2+ signalling, cytoplasmic Ca2+

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concentrations may temporarily rise to around 1 μM. Both the maintenance of low cytoplasmic Ca2+ and a rapid restoration of resting levels after a signalling event require rigorous systems to remove cytoplasmic Ca2+ and deposit it in the apoplast or in intracellular stores (see also Chapter 7) such as the vacuole and the ER. One of the main components to maintain Ca2+ homeostasis is the P-type Ca2+ ATPase. These primary Ca2+ pumps are predominantly found at the plasma membrane and also at the ER, mitochondrial and plastidic membranes. As with other primary pumps, Ca2+ pumps have low turnover rates, but their affinity is extremely high with nanomolar K m values. The general catalytic mechanism of Ca2+ pumps is similar to that described for the P-type H+ ATPase, including a phosphorylated intermediate. There are two main classes (Axelsen and Palmgren, 2001), type IIA and IIB, of plant Ca2+ pumps, both functioning as a single polypeptide with an approximate MW of ∼110 kDa. In contrast to type IIA, type IIB pumps contain a calmodulin binding domain in the N-terminal region which is involved in a Ca2+ based feedback mechanism: When cytoplasmic Ca2+ levels increase, Ca2+ -activated calmodulin binds to the Ca2+ pump and stimulates its activity.

5.3.2.2

Heavy metal ATPases

This group of enzymes catalyses a diverse set of functions in the homeostasis of essential heavy metals such as Cu2+ , Zn2+ , Mn2+ , Fe2+ , Ni2+ and Co2+ (Hussain et al., 2004; Williams and Mills, 2005). Heavy metal ATPases (HMAs) are members of the P-type ATPase superfamily and are also named CPx-ATPases due to a signature domain consisting of a cysteine-proline-x motif, where x is a cysteine, a histidine or a serine. The CPx motif resides in the sixth TMD and is thought to play a role in binding of the transported metal. A good example where CPxATPases are known to mediate essential metal transport processes is the delivery of Cu2+ to the chloroplast where it is needed in the lumen for the copper-containing redox protein plastocyanin. In the chloroplast stroma, copper is a cofactor for the chloroplastic superoxide dismutase (SOD) enzyme, which is crucial for scavenging photosynthesis-generated superoxide radicals. At the chloroplast, two Cu2+ ATPases (PAA1 and PAA2) have been identified (Shikanai et al., 2003; Abdel-Ghaney et al., 2005). On the basis of GFP reporter studies, it was deduced that PAA1 is expressed in the chloroplast envelope whereas PAA2 is most likely targeted to the thylakoid membrane (see also Section 7.2.4.3). Through loss-of-function mutants, it was found that both PAA1 and PAA2 are necessary to deliver plastocyanin Cu2+ to the thylakoid lumen for photosynthetic activity, whereas only PAA1 is required to ensure adequate SOD activity in the stroma. Despite its physiological relevance, Cu2+ and other heavy metals can also be extremely toxic to plants even at micromolar concentrations (see also Chapter 12). After uptake from the soil solution such metals are readily chelated by metal-binding chaperones to avoid toxicity and detoxification also entails vacuolar sequestration and extrusion into the apoplast, functions that are often mediated by heavy metal ATPases.

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5.3.3

PLANT SOLUTE TRANSPORT

ABC transporters

The last class of primary transporters that will be discussed is made up of the ATP binding cassette (ABC) transporters (Sanchez-Fernandez et al., 2001; Martinoia et al., 2002). ABC transporters are encoded by large gene families (∼150 genes in arabidopsis) and fulfil an amazing number of transport functions. In mammals, many ABC transporters are involved in the efflux of xenobiotics including medicinal drugs and are therefore responsible for the ‘multi drug resistance’ phenomenon. In plants, only a few ABC transporters have been characterised but it is clear that these proteins function in the detoxification of xenobiotics such as herbicides and heavy metals, the transport of hormones, and the movement of an exceedingly large range of metabolites. ABC transporters use MgATP as their energy source but the ATP can be replaced by GTP. Like P-type ATPases, ABC transporters are readily inhibited by vanadate, pointing to phosphorylation as an important step during the catalytic cycle. Structurally, they generally consist of two copies of a transmembrane domain (TMD) and an ATP binding domain (ABD). The latter couples ATP hydrolysis to transport of the substrate through the TMD. In fully functioning ABC transporters, the arrangement can either be TMD-ABD-TMD-ABD or ABD-TMD-ABD-TMD. The ABD contains a conserved region with characteristic ‘Walker motifs’ (GXXXXGKT, where X is any residue) and ABC transporter signature motif. As in animals, one of the main functions of plant ABC transporters is the removal of toxic compounds from the cytoplasm. This typically involves vacuolar sequestration of the hazardous compound after conjugation with derivatives such as glutathione, glycosyl or acetyl groups. A large group of ABC transporters from the multi-drug-resistance like protein (MRP) subfamily moves glutathionylated (GS) substances from the cytosol to the vacuolar lumen. For example, the arabidopsis MRP1 ABC transporter was shown to be capable of transporting conjugates of the herbicide metolachlor. The same transporter also illustrates one of the big conundrums regarding ABC transporters, the seemingly contradictory properties of high affinity and low specificity for substrates. For example, AtMRP1 was also shown to transport dinitrophenyl-GS and anthocyanin-GS with high affinity. As is the case for their animal counterparts (e.g. the cystic fibrosis transmembrane conductance regulator) plant ABC transporters may also be involved in the regulation of ion transport (e.g. Leonhardt et al., 1999).

5.4

Secondary active transport

In many cases, metabolites and nutrients have to be moved from lower to higher (electro)chemical potentials and therefore require active transport. In contrast to ATP-driven pumps that typically have high affinities and low turnover rates, active transport through carrier-type mechanisms achieves larger transport rates. Energisation of such mechanisms is realised through coupling of substrate movement to that of other ions that move down their electrochemical potential. In animal cells, primary transport in the form of the K+ -Na+ ATPase establishes a transmembrane

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Na+ gradient and hence secondary transport is generally Na+ coupled. Plants, however, use H+ ATPases to generate a PMF and consequently secondary transport in plants is mostly H+ coupled. Notwithstanding this general concept, exceptions to this rule have been found (see Section 5.4.4). Proton-coupled active transport can consist of two mechanisms: antiport or symport. The former indicates a counterflow of substrate(s) and H+ , whereas in the latter case substrate(s) and H+ move in the same direction across the membrane. The list of both antiport and symport systems observed and anticipated in plants is long and so only some of the main examples are discussed below.

5.4.1

Potassium uptake

+

K is an essential macronutrient for plants and crucial in many metabolic reactions because of its role in enzyme activation. Protein synthesis, photosynthesis and cytoplasmic H+ homeostasis all require high cytoplasmic K+ concentrations. Furthermore, K+ plays a key role in turgor generation. As is the case for many nutrients, K+ uptake into plant roots has low- and high-affinity components with the latter being induced when plants become K+ starved. Electrophysiological studies have shown that, at relatively high external concentrations, passive transport of K+ through ion channels occurs. However, when external concentrations of K+ drop to low (micromolar) levels, its uptake needs to proceed through an active mechanism. This becomes clear by examining the equilibrium condition for K+ distribution across the membrane which is described by: 

K+ out E m = 60 log  + . K in

(5.9)

A membrane potential of –180 mV, a realistic value, can theoretically drive a 1000-fold accumulation of K+ and this could proceed through ion channels. But with [K+ ] in typically being in the region of 100 mM, such passive uptake can no longer proceed whenever the external [K+ ] drops below 100 μM. In those conditions, active transport is required. Electrophysiological experiments showed that this occurs via H+ coupling in a K+ :H+ symport mechanism (Maathuis and Sanders, 1994) that has a high affinity for K+ of around 20 μM. For a symport where K+ and H+ fluxes are coupled, the overall change in free energy is a function of E m , the respective ion gradients, and the coupling ratio n:



n   H+ out K+ out .  n  H+ in K+ in

G = (n + 1)FE m + RT ln

(5.10)

At equilibrium G = 0 and  Em =

RT (n + 1)F

n   H+ out K+ out .  n  H+ in K+ in



 ln

(5.11)

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For the K+ :H+ symport, it was shown that this high-affinity uptake mechanism has an apparent stoichiometry of 1 K+ per H+ (Maathuis et al., 1997). This coupling ratio of unity ensures that theoretically the system can sustain an incredible 108 -fold accumulation of K+ if we assume an E m of –180 mV and a pH of 2. The molecular identity of high-affinity K+ transporters has been established to some extent. In Arabidopsis, members of the KUP/HAK gene family have been found to mediate high-affinity uptake in heterologous expression systems and expression of one isoform, AtHAK5, is induced by K+ starvation (Gierth et al., 2005). HAK5 is expressed in cortical and stelar root cells and a loss-of-function mutant in this gene showed absence of 86 Rb+ uptake, specifically in the high-affinity range. Thus, HAK5 is the most likely candidate for the plant H+ coupled high affinity K+ uptake system, but only a thorough electrophysiological examination of the energetics of the HAK5 encoded transporter will give a definitive answer.

5.4.2

Nitrate transport

In terms of mass, nitrogen is one of the largest constituents of plant biomass: it is acquired in large quantities from the soil. Generally, the forms of nitrogen taken up by plants are NH 4 + and NO 3 − of which NO 3 − is usually preferred. As is the case for K+ , the uptake of nitrate by roots is characterised by distinct high- and low-affinity kinetic phases (Forde, 2000; Orsel et al., 2002). Although external concentrations can be in the millimolar range, particularly in fertilised areas, the negative charge of NO 3 − means it has to overcome a considerable energetic barrier to enter the plant symplast and even in this low-affinity range, nitrate uptake has, therefore, to rely on active transport. It is widely assumed that such transport depends on coupling to H+ and Eq. 5.10 can be used to assess the possibility of various coupling ratios between H+ and NO 3 − . If unity is substituted for n then the overall charge becomes zero and NO 3 − uptake has to rely entirely on the pH as the driving force. With a pH of 2 and typical cytoplasmic NO 3 − concentration of 5 mM, this would restrict uptake to external concentration of 50 μM or more. However, uptake has been shown to proceed at lower external NO 3 − concentrations. In addition, measurements of root cell membrane potentials and voltage clamp experiments with nitrate-transporterexpressing oocytes have shown that the membrane depolarises in response to an increase in external NO 3 − . These observations strongly indicate that entry of nitrate into the symplast carries positive charge and it is therefore assumed that 2 H+ accompany each NO 3 − taken up through a NO 3 − :2H+ symporter (Meharg and Blatt, 1995). Two main classes of genes have been identified that are involved in various forms of NO 3 − uptake. These are the NRT1 and NRT2 gene families, both encoding proteins with a six plus six TMD structure. As a coarse generalisation, the NRT1 gene products are involved in low-affinity NO 3 − uptake, whereas NRT2 genes encode proteins that mediate both constitutive and inducible high-affinity uptake capacity (Orsel et al., 2002).

TRANSPORT ACROSS PLANT MEMBRANES

5.4.3

95

Sodium efflux

Apart from a number of C4 plants, sodium (Na+ ) is not required by plants as a nutrient. On the contrary, Na+ -based salinisation of many regions worldwide is an increasing problem for agriculture, especially since many crop plants are salt sensitive (cf Chapter 14). The negative membrane potential and high external concentrations of Na+ ions mean that the passive Na+ flux into the plant symplast is potentially large. Principally, there are only two ways to avoid death by salt: significantly limit Na+ influx and remove excess Na+ from the cytoplasm where its toxicity is most prevalent. Removal of Na+ is against its electrochemical gradient and therefore coupled to the H+ gradient. Both at the tonoplast and the plasmamembrane H+ :Na+ antiport mechanisms have been characterised that participate in exporting Na+ to the vacuole and apoplast, respectively. Most of this characterisation was carried out using membrane vesicles and measuring Na+ -induced changes in acridine or quinacrine fluorescence (see also Section 4.4.4). The latter compounds report changes in pH which should occur across the membrane when H+ -coupled Na+ fluxes occur. Such experiments revealed that plant H+ :Na+ antiporters are electroneutral, i.e. the coupling ratio is unity, and therefore completely rely on the presence of a pH. Thus, with a vacuolar pH of 5 and a cytoplasmic pH of 7.5, an electroneutral H+ :Na+ antiporter could drive an approximate 300 times accumulation of Na+ in the vacuolar lumen. H+ :Na+ antiporters from the NHX gene family (Yokoi et al., 2002) have been cloned from a range of plant species and the expression level of several tonoplast and plasma membrane H+ :Na+ antiporters greatly impacts on plant salt tolerance (Zhu, 2001; Vera-Estrella et al., 2005). Nevertheless, NHX isoforms may also be capable of transporting other monovalent cations. NHX antiporters typically contain 12 TMD (Colour plate 5.8), are around 120–130 kDa in weight and often have a conserved motif in the third TMD, where amiloride binds. Amiloride, a diuretic used to treat high blood pressure, inhibits many animal and plant H+ :Na+ antiporters. Recent work, however, challenges the generalised NHX topology and provides evidence that the C-terminus is actually in the luminal compartment where it plays an important role in calmodulin-dependent regulation of ion selectivity (Yamaguchi et al., 2005).

5.4.4

Non H + -coupled secondary transport

In contrast to animal cells, Na+ -coupled secondary transport is rare in plants. However, there is convincing evidence that it occurs for some substrates such as urea, NO 3 − and K+ . Na+ :urea symport has only been observed in charophytes and never in higher plants (Walker and Sanders, 1991), whereas Na+ -coupled NO 3 − uptake has been reported only for the halophytic seagrass Zostera marina. A more extensive survey was made into the occurrence of Na+ -coupled K+ uptake (Maathuis et al., 1996) and this showed that apart from charophytic genera, aquatic angiosperms such Egeria, Elodea and Vallisneria contain a second type of high-affinity K+ uptake

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system that is driven through Na+ cotransport. The latter generally has a lower K m than the H+ coupled high affinity system. Since no molecular or mechanistic data are available for the Na+ -coupled system, it is hard to establish its exact physiological role in these aquatic species but it has been suggested to act as a K+ scavenging mechanism when ambient K+ levels are extremely low.

5.5

Concluding remarks

To maintain life, it is essential that cells have the capacity to optimise and control local environments. This will critically depend on two factors: the presence of barriers between compartments in the form of membranes and the mechanisms to control the influx and efflux of compounds. The latter function has developed to a high level of sophistication in higher plants to move solutes efficiently. Broadly speaking, primary, ATP-driven pumps set up H+ gradients to drive secondary transport. Primary pumps are also directly involved in Ca2+ and heavy metal transport. Secondary transport in plants is generally coupled to H+ gradients and participates in the uptake and movement of hundreds, if not thousands of different substrates. Finally, it is the ion channels that are almost exclusively responsible for passive transport. In spite of completing genome sequencing for a number of plant species, many of the transport mechanisms have yet to be identified at the gene level and an even greater number of transport proteins requires characterisation regarding substrate selectivity, kinetics and patterns of expression. Nevertheless, the current data suggest that all these mechanisms are under close control through many processes such as transcription, post-translational modification, membrane voltage modulation and ligand binding. It is this highly regulated nature and the specifically tailored functional aspects of solute transporters that endow plants with the capacity to thrive in almost any global environment.

References Abdel-Ghany, S.E., Muller-Moule, P., Niyogi, K.K., Pilon, M. and Shikanai, T. (2005) Two P-type ATPases are required for copper delivery in Arabidopsis thaliana chloroplasts. Plant Cell 17, 1233–1251. Axelsen, K.B. and Palmgren, M.G. (2001) Inventory of the superfamily of P-type ion pumps in Arabidopsis. Plant Physiology 126, 696–706. Balague, C., Lin, B.Q., Alcon, C., et al. (2003) HLM1, an essential signaling component in the hypersensitive response, is a member of the cyclic nucleotide-gated channel ion channel family. Plant Cell 15, 365–379. Berthomieu, P., Conejero, G., Nublat, A., et al. (2003) Functional analysis of AtHKT1 in Arabidopsis shows that Na+ recirculation by the phloem is crucial for salt tolerance. Embo Journal 22, 2004– 2014. Blumwald, E. and Poole, R.J. (1985) Na+ /H+ antiport in isolated tonoplast vesicles from storage tissue of Beta vulgaris. Plant Physiology 78, 163–167. Blumwald, E. and Poole, R.J. (1987) Salt tolerance in suspension-cultures of sugar-beet – induction of Na+ /H+ antiport activity at the tonoplast by growth in salt. Plant Physiology 83, 884–887.

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Chrispeels, M.J., Morillon, R., Maurel, C., Gerbeau, P., Kjellbom, P. and Johansson, I. (2001) Aquaporins of plants: structure, function, regulation, and role in plant water relations. Aquaporins 51, 277–334. Davenport, R. (2002) Glutamate receptors in plants. Annals Botany 90, 549–557. Davies, J.M., Hunt, I. and Sanders, D. (1994) Vacuolar H+ -pumping ATPase variable transport coupling ratio controlled by pH. Proceeding National Academy Science USA 91, 8547–8551. Demidchik, V., Davenport, R.J. and Tester, M. (2002) Nonselective cation channels in plants. Annual Review Plant Biology 53, 67–107. Dietz, K.J., Tavakoli, N., Kluge, C., et al. (2001) Significance of the V-type ATPase for the adaptation to stressful growth conditions and its regulation on the molecular and biochemical level. Journal Experimental Botany 52, 1969–1980. Forde, B.G. (2000) Nitrate transporters in plants: structure, function and regulation. Biochimica Biophysica Acta-Biomembranes 1465, 219–235. Gaxiola, RA., Fink, G.R. and Hirschi, K.D. (2002) Genetic manipulation of vacuolar proton pumps and transporters. Plant Physiology 129, 967–973. Geelen, D., Lurin, C., Bouchez, D., et al. (2000) Disruption of putative anion channel gene AtCLC-a in Arabidopsis suggests a role in the regulation of nitrate content. Plant Journal 21, 259–267. Geisler, M., Axelsen, K.B., Harper, J.F. and Palmgren, M.G. (2000) Molecular aspects of higher plant P-type Ca2+ -ATPases. Biochimica Biophysica Acta-Biomembranes 1465, 52–78. Gierth, M., Maser, P. and Schroeder, J.I. (2005) The potassium transporter AtHAK5 functions in K+ deprivation- induced high-affinity K+ uptake and AKT1 K+ channel contribution to K+ uptake kinetics in Arabidopsis roots. Plant Physiology 137, 1105–1114. Gouaux, E. and MacKinnon, R. (2005) Principles of selective ion transport in channels and pumps. Science 310, 1461–1465. Hechenberger, M., Schwapach, B., Fischer, W.N., Frommer, W.B., Jentsch, T.J. and Steinmeyer, K. (1996) A family of putative chloride channels from Arabidopsis and functional complementation of a yeast strain with a CLC gene disruption. Journal Biochemistry 271, 33632–33638. Hille, B. (2001) Ion Channels of Excitable Membranes. Sinauer Associates, Inc. Sunderland, MA. Hussain, D., Haydon, M.J., Wang, Y., et al. (2004) P-type ATPase heavy metal transporters with roles in essential zinc homeostasis in Arabidopsis. Plant Cell 16, 1327–1339. Johanson, U., Karlsson, M., Johansson, I, et al. (2001) The complete set of genes encoding major intrinsic proteins in Arabidopsis provides a framework for a new nomenclature for major intrinsic proteins in plants. Plant Physiology 126, 1358–1369. Kluge, C., Lahr, J., Hanitzsch, M., Bolte, S., Golldack, D. and Dietz, K.J. (2003) New insight into the structure and regulation of the plant vacuolar H+ -ATPase. Journal Bioenergetics Biomembranes 35, 377–388. Laurie, S., Feeney, K.A., Maathuis, F.J.M., Heard, P.J., Brown, S.J. and Leigh, R.A. (2002) A role for HKT1 in sodium uptake by wheat roots. Plant Journal 32, 139–149. Leng, Q., Mercier, R.W., Hua, B.G., Fromm, H. and Berkowitz, G.A. (2002) Electrophysiological analysis of cloned cyclic nucleotide-gated ion channels. Plant Physiology, 128, 400–410. Leonhardt, N., Vavasseur, A. and Forestier, C. (1999) ATP binding cassette modulators control abscisic acid-regulated slow anion channels in guard cells. Plant Cell 11, 1141–1151. Ludewig, U., von Wiren, N. and Frommer, W.B. (2002) Uniport of NH4+ by the root hair plasma membrane ammonium transporter LeAMT1;1. Journal Biological Chemistry 277, 13548–13555. Luu, D.T. and Maurel, C. (2005) Aquaporins in a challenging environment: molecular gears for adjusting plant water status. Plant Cell Environment 28, 85–96. Maathuis, F.J.M. and Sanders, D. (1994) Mechanism of high affinity potassium uptake in roots of Arabidopsis thaliana. Proceeding National Academy Science USA 91, 9272–9276. Maathuis, F.J.M., Verlin, D., Smith, F.A., Sanders, D., Fernandez, J.A. and Walker, N.A. (1996) The physiological relevance of Na+ -coupled K+ -transport. Plant Physiology 112, 1609–1616. Maathuis, F.J.M., Sanders, D. and Gradmann, D. (1997) Kinetics of high affinity K+ uptake in plants, derived from K+ - induced changes in current-voltage relationships. Planta 203, 229–236. Maathuis, F.J.M. and Sanders, D. (2001) Sodium uptake in Arabidopsis thaliana roots is regulated by cyclic nucleotides. Plant Physiology 127, 1617–1625.

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Maathuis, F.J.M., Filatov, V, Herzyk, P., et al. (2003) Transcriptome analysis of root transporters reveals participation of multiple gene families in the response to cation stress. Plant Journal 35, 675–692. Maeshima, M. (2000) Vacuolar H+ -pyrophosphatase. Biochimica Biophysica Acta-Biomembranes 1465, 37–51. Martinoia, E., Klein, M., Geisler, M., et al. (2002) Multifunctionality of plant ABC transporters – more than just detoxifiers. Planta 214, 345–355. M¨aser, P., Eckelman, B., Vaidyanathan, R., et al. (2002) Altered shoot/root Na+ distribution and bifurcating salt sensitivity in Arabidopsis by genetic disruption of the Na+ transporter AtHKTI1. FEBS letters 531, 157–161. Meharg, A.A. and Blatt, M.R. (1995) NO 3 - transport accross the plasma membrane of Arabidopsis thaliana root hairs: kinetic control by pH and membrane voltage. Journal Membrane Biology 145, 49–66. Orsel, M., Filleur, S., Fraisier, V. and Daniel-Vedele, F. (2002) Nitrate transport in plants: which gene and which control? Journal Experimental Botany 53, 825–833. Peiter, E., Maathuis, F.J.M., Mills, L.N., et al. (2005) The vacuolar Ca2+ -activated channel TPC1 regulates germination and stomatal movement. Nature 434, 404–408. Rubio, F., Gassmann, W. and Schroeder, J.I. (1995) Sodium-driven potassium uptake by the plant potassium transporter HKT1 and mutations conferring salt tolerance. Science, 270, 1660–1663. Sanchez-Fernandez, R., Davies, T.G.E., Coleman, J.O.D. and Rea, P.A. (2001) The Arabidopsis thaliana ABC protein superfamily, a complete inventory. Journal Biological Chemistry 276, 30231–30244. Shikanai, T., M¨uller-Moul´e, P., Munekage, Y., Niyogi, K.K. and Pilon M. (2003) PAA1, a P-Type ATPase of Arabidopsis, functions in copper transport in chloroplasts. Plant Cell 15, 1333–1346. Staal, M., Maathuis, F.J.M., Overbeek, J.H.M., Elzenga, J.T.M. and Prins, H.B.A. (1991) Na+ /H+ antiport activity in tonoplast vesicles from roots of the salt-tolerant Plantago maritima L. and the salt-sensitive Plantago media l. Physiologia Plantarum 82, 179–184. Sze, H., Li, X.H. and Palmgren, M.G. (1999) Energization of plant cell membranes by H+ -pumping ATPases: regulation and biosynthesis. Plant Cell 11, 677–689. Talke, I.N., Blaudez, D., Maathuis, F.J.M. and Sanders, D. (2003) CNGCs: prime targets of plant cyclic nucleotide signalling? Trends Plant Science 8, 286–293. Vera-Estrella, R., Barkla, B.J., Garcia-Ramirez, L. and Pantoja, O. (2005) Salt stress in Thellungiella halophila activates Na+ transport mechanisms required for salinity tolerance. Plant Physiology 139, 1507–1517. Very, A.A. and Sentenac, H. (2003) Molecular mechanisms and regulation of K+ transport in higher plants. Annual Review Plant Biology 54, 575–603. Walker, N.A. and Sanders, D. (1991) Sodium-coupled solute transport in charophyte algae: a general mechanism for transport energization in plant cells? Planta 185, 443–445. Wang Y., Schulten K. and Tajkhorshid E. (2005) What makes an aquaporin a glycerol channel: a comparative study of AqpZ and GlpF. Structure 13, 1107–1118. White, P.J. (2000) Calcium channels in higher plants. Biochimica Biophysica Acta-Biomembranes, 1465, 171–189. Williams, L.E. and Mills, R.F. (2005) P-1B-ATPases – an ancient family of transition metal pumps with diverse functions in plants. Trends Plant Sciences 10, 491–502. Yamaguchi, T., Aharon, G.S., Sottosanto, J.B. and Blumwald, E. (2005) Vacuolar Na+ /H+ antiporter cation selectivity is regulated by calmodulin from within the vacuole in a Ca2+ - and pH-dependent manner. Proceedings National Academy Sciences USA 102, 16107–16112. Yokoi, S., Quintero, F.J., Cubero, B., et al. (2002) Differential expression and function of Arabidopsis thaliana NHX Na+ /H+ antiporters in the salt stress response. Plant Journal 30, 529–539. Zhu, J.K. (2001) Plant salt tolerance. Trend Plant Sciences 6, 66–71.

6 Regulation of ion transporters Anna Amtmann and Michael R. Blatt 6.1

Introduction

Transport proteins have multiple and diverse functions in plant development and physiology. They mediate the transport of many solutes including inorganic nutrients, primary assimilates and secondary metabolites. They reside in various tissues and membranes where they facilitate the uptake or the release of solutes into/from cells and intracellular compartments. Transport proteins differ in affinity, transport rate and direction of transport (Chapter 5). Such functional diversity is required to ensure that nutrients and metabolites can be optimally used under any given condition. According to developmental stage, metabolic state, composition of the soil and environmental condition, the plant has to integrate uptake, allocation and redistribution of solutes to maximise their usage for growth and reproduction. Such adaptation requires flexible and robust mechanisms to regulate solute transport. Developmental, metabolic and environmental clues have to be recognised and translated into signals that are perceived by the targeted transporters and the resulting fluxes have to be adjusted at the whole-plant level. The regulation of ion transporters occurs at several levels (e.g. gene expression, mRNA degradation, protein turnover, protein activity and membrane trafficking), and involves both negative and positive feedback loops (e.g. inhibition of nitrate transporters by N assimilates and Ca2+ -induced Ca2+ release). Furthermore, membrane transporters are both integral components and targets of signalling pathways (e.g. cytoplasmic Ca2+ oscillations and gradients, auxin transport, abscisic acid [ABA] signalling and salt-overly-sensitive [SOS] pathway). Transcriptional and post-translational regulation of transporters has been the subject of active research for many years, and a huge amount of knowledge about the signals and regulatory mechanisms involved has been accumulated. In this chapter only the surface of this field can be scratched, by listing several examples for adaptive processes involving the regulation of transporters (Section 6.2) and molecular mechanisms underlying transcriptional and post-translational regulation of ion transporters (Section 6.3). In the last section, recent ideas on how ion transporters could be regulated by membrane trafficking are summarised.

6.2 6.2.1

Physiological situations requiring the regulation of ion transport Change of cell volume

The most elemental of situations requiring control of solute transport in plants relate to changes in cell volume. Plant cells utilise inorganic ions – notably K+ and to lesser

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extents Cl− and organic anions – as the primary osmotically active solutes to maintain turgor and to drive irreversible cell expansion as well as reversible changes in cell volume. Clearly, a bias of inorganic ion uptake must be maintained to accommodate increases in cell volume during growth and, on a cell-by-cell basis, a coordinated balance of influx and efflux must be achieved once growth terminates (HoldawayClarke and Hepler, 2003; Very and Sentenac, 2003; Amtmann et al., 2004). A similar coordination of ion transport, albeit for solute loss, underpins reversible changes in cell volume that are characteristic of stomatal guard cells (Willmer and Fricker, 1996), pulvini and similar ‘motor’ cells in plants (Hill and Findlay, 1981). Major targets for transport control in all these cases include the dominant H+ -ATPases that energise the various cell membranes, as well as K+ and Cl− (anion) channels. While less is known that bears on transport control during irreversible growth, much detail of several regulatory pathways have come to light from a few cell models, especially stomatal guard cells, and some recent developments are briefly touched on here. The very large ion fluxes associated with stomatal movements offer a particularly useful handle for analysis of transport control in stomatal guard cells. Between open and closed states, guard cells of Vicia faba, for example, take up or release 2–4 pmol of KCl. On a cell volume basis, these changes correspond to 200–300 mOsm in solute content. Since mature guard cells lack plasmodesmata (Wille and Lucas, 1984), all of this solute flux must pass across the plasma membrane. Electrophysiological measurements under voltage clamp of the H+ -ATPase and dominant ion channel currents show that these are more than sufficient to account for solute uptake during stomatal opening and for K+ and Cl− loss during stomatal closure (Blatt, 1987; Lohse and Hedrich, 1992; Thiel and Wolf, 1997; Schroeder et al., 2001). Nonetheless, attention over the years has centred primarily on the effects of ABA in facilitating solute loss and stomatal closure under water stress, and especially its action in regulating K+ and Cl− channel activities at the plasma membrane. The voltage sensitivities of these ion channels contribute significantly to their regulation under free-running (that is, non-voltage-clamped) conditions (Gradmann et al., 1993; Blatt, 2000). Nonetheless, control of the Cl− channel current (I Cl ) and the two dominant K+ channels – identified with the inward-rectifying (I K,in ) and outward-rectifying (I K,out ) K+ currents and, in Arabidopsis thaliana, principally with the KAT1 and GORK gene products, respectively (Nakamura et al., 1995; Hosy et al., 2003) – is also tightly linked through voltage-independent pathways. Coordinated regulation of I Cl and I K,in is achieved in part through signalling pathways that lead to a rise in cytosolic-free [Ca2+ ] ([Ca2+ ] i ). The coupling of changes in [Ca2+ ] i to ABA is well established (Davies and Jones, 1991; McAinsh et al., 1997; Blatt, 2000; Webb et al., 2001). However, the mechanisms leading to a rise in [Ca2+ ] i and its downstream targets continue to yield new insights. It is clear now that ABA influences [Ca2+ ] i through at least two complementary processes. At the plasma membrane, ABA affects the voltage threshold for activation of Ca2+ channels mediating Ca2+ entry (Hamilton et al., 2000) and thereby affects the interaction between [Ca2+ ] i and voltage evident as oscillations in [Ca2+ ] i (Allen et al., 2000, 2001; Blatt, 2000), possibly through a NADPH-oxidase-dependent process (Kwak et al., 2003). Within the guard cell, ABA promotes Ca2+ release

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101

induced by Ca2+ entry across the plasma membrane via at least one well-defined mechanism that draws on nitric oxide stimulating cyclic GMP- and cyclic ADPribose-activated Ca2+ channels within one or more endomembranes (Neill et al., 2002; Garcia-Mata et al., 2003). Other internal, Ca2+ -associated pathways include inositol-1,4,5-trisphosphate release and [Ca2+ ] i elevation through the actions of phospholipase C (Blatt et al., 1990; Gilroy et al., 1990; Hunt et al., 2003), as well as the actions of inositol-hexakisphosphate (Lemtiri-Chlieh et al., 2000), sphingosine and other membrane lipid metabolites (Lee et al., 1996; Ng et al., 2001; Coursol et al., 2005). How these [Ca2+ ] i signals are translated to alterations in I K,in and I Cl activities is still uncertain. Plausibly, Ca2+ may couple to the channel proteins through a protein kinase cascade similar to that of SOS2/3 signalling pathway that regulates the SOS1 Na+ /H+ antiporter (see Section 6.2.3). Certainly, both the K+ and Cl− channel currents are sensitive to protein kinase and phosphatase antagonists (Luan et al., 1993; Thiel and Blatt, 1994; Grabov et al., 1997) and subject to modulation by the arabidopsis ABI1 (abscisic acid insensitive 1) 2C-type protein phosphatase (Armstrong et al., 1995; Pei et al., 1997). Protein phosphorylation also influences the efficacy of [Ca2+ ] i signalling and the responses of I K,in and I Cl mediated by nitric oxide (Sokolovski et al., 2005). Thus, it seems likely that (de)phosphorylation of a target closely associated with these channels contributes to Ca2+ signal transmission. While the Ca2+ signal predominates in many aspects of regulation for I K,in and I Cl , it is curiously absent in control of the outward-rectifying K+ channels in guard cells, including the GORK K+ channel of arabidopsis (Hosy et al., 2003) and its phloem homologue SKOR (Gaymard et al., 1998). Instead, current through these channels is strongly dependent on cytosolic pH (pH i ) and on extracellular [K+ ] ([K+ ] o ). Increasing pH i augments I K,out in a scalar fashion, consistent with a synergistic binding of two H+ (Grabov and Blatt, 1997) and the rise in pH i evoked by ABA (Irving et al., 1992; Blatt and Armstrong, 1993). Unlike the situation for Ca2+ , virtually nothing is known of the mechanism(s) behind this rise in pH i nor of its site of action, although its kinetics (and that of changes in I K,out ) are sufficiently slow (Blatt and Armstrong, 1993) to be accommodated by cation exchange and charge-balancing events during solute efflux from the vacuole (MacRobbie, 2000). The control of gating by [K+ ] o exhibited by I K,out is unique to plant K+ channels, but is not exclusive to guard cells. Indeed, for many plant Kv channels, the voltage dependence of gating is modulated by the availability of K+ outside (Blatt, 1991). Like the K+ channels of animal cells, these channels open on membrane depolarisation and facilitate K+ flux out of the cell; however, unlike their animal counterparts, the plant K+ channels do so only at membrane voltages positive of the K+ equilibrium potential. In other words, the voltage dependence of gating for these K+ channels in plants shifts with external [K+ ]. This ability to respond to the prevailing [K+ ] o makes good physiological ‘sense’; it guarantees that the channels open only when the driving force for net K+ flux is directed outward, and, in stomatal guard cells it ensures the K+ efflux needed to drive stomatal closure and control gas exchange, even when extracellular K+ varies over concentrations from 10 nM to 100 mM (Blatt and Gradmann, 1997).

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Remarkably, the ability to sense [K+ ] o is integral to the K+ channel protein itself and, therefore, represents one of the very few examples in which the mechanism for ‘nutrient sensing’ is known. It also poses a number of intriguing questions that are fundamental to understanding the mechanisms of voltage-dependent K+ channel gating both in plants and in animals. From a molecular standpoint, gating by K+ implies that cation binding with the channel protein must stabilise or otherwise favour a closed conformation of the channel pore. Recent studies demonstrated that SKOR gating is consistent with K+ binding to a site with pore-like characteristics and identified a critical role for a site deep within the S6 transmembrane helix of the channel protein, adjacent the so-called pore helix, that is essential for this K+ sensitivity (Johansson et al., 2006). The finding is significant, because analogous interactions between the pore helix and the S6 helix are known to affect gating of mammalian K+ channels (cf. Alagem et al., 2003; Seebohm et al., 2003, 2006). In animals, these interactions are favoured by cation occupation of the pore. Thus, in effect, cations in the pore of many animal K+ channels can be thought to ‘push out’ on the pore and, indirectly, to stabilise the open state, a mechanism originally identified by Clay Armstrong who proposed the ‘foot-in-the-door’ hypothesis (see Hille, 2001). However, in SKOR (Johansson et al., 2006) these interactions have precisely the opposite effect: they stabilise the closed state of the K+ channel. Furthermore, site mutations in this domain of SKOR also affect the sensitivity to [K+ ]. How K+ binding translates to these seemingly counterintuitive effects on gating is not clear at present, but will undoubtedly yield insights into K+ channel gating in general. For the moment, however, it can be concluded that there is a tight interplay in the mechanics of gating by K+ and voltage that accounts for the response of the channel to the K+ environment.

6.2.2

Nutrient acquisition

Many transporters are involved in the acquisition, storage and redistribution of macronutrients such as potassium, calcium, magnesium, nitrogen, sulphur and phosphorus as well as micronutrients including iron, zinc, manganese and copper (see also Chapter 12). Depending on supply and demand of mineral nutrients, plants have to adjust the expression levels and activity of individual transporters to achieve a combination of specific transport properties (affinity, substrate specificity and mode of energisation) that is best suited to fulfil the plant’s needs under the given conditions. Regulation of nutrient transport occurs primarily at the level of gene expression; for example, many genes encoding transporters involved in nutrient acquisition are induced or repressed when the concentration of their substrate in the soil changes. In the following description of these events the emphasis is on transporters of macronutrients. Nutrient availability also affects the transcription of transporters such as proton pumps or anion channels that are not directly involved with the transport of the respective minerals but have a function in increasing the driving force for nutrient uptake or in providing charge balance (Maathuis et al., 2003). Several genes encoding high-affinity sulphate (e.g. AtSULTR1;2 and AtSULRT1;2; Buchner et al., 2004) and phosphate transporters (e.g. AtPT1 and

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AtPT2; Al-ghazi et al., 2003) are induced by removal of S or P from the growth medium. By contrast, high-affinity nitrate transporters of the NRT2 family are induced by adding small amounts of nitrate (10–50 μM) to an N-depleted medium (Krapp et al., 1998; Filleur and Daniel-Vedele, 1999). Expression of ammonium transporters from arabidopsis (e.g. AtAMT1;1 and AtAMT1;3) increases during N deficiency, whereas expression of AMT isoforms from tomato and rice are induced by N supply (Loque and von Wiren, 2004). The observation that some transporters are induced by a change from high to low supply and others by a change from nil to low supply might indicate a fundamental difference in the underlying regulatory mechanisms but is of little relevance for the physiological effect. Both responses result in high expression levels of these transporters under conditions that require their function as high-affinity systems in nutrient uptake. Fine-tuning of this response seems to occur through differential regulation of apparently functionally redundant isoforms. For example, SULTR1;1 and SULRT1;2 are both high-affinity transporters induced by low S (10 μM), and they co-localise in the plasma membrane of root cells (Takahashi et al., 2000; Shibagaki et al., 2002); nevertheless, they differ with respect to sulphur sensitivity of their transcriptional response. SULTR1;1 shows sharp induction between 100 and 1500 μM external S, whereas SULTR1;2 transcripts increase more gradually with a maximum at 10 μM external S (Maruyama-Nakashita et al., 2004c). Similarly, members of the arabidopsis AMT family of ammonium transporters vary in their sensitivity to N supply; transfer of plants to N-free medium induces the expression of AtAMT1;1 and AtAMT1;3 within 3 days, whereas induction of AtAMT1;2 and AtAMT2;4 requires more extended periods of N deficiency (Gazzarrini et al., 1999; Sohlenkampa et al., 2000). Such differential regulation of transporter isoforms allows a flexible usage of these systems in a wide range of conditions. Astonishingly few transporters involved in K+ transport respond to varying + K supply with transcriptional changes (Maathuis et al., 2003). Out of some 50 genes expected to have K+ transport capacity (M¨aser et al., 2001), only HAK5, a putative high-affinity K+ uptake system, is consistently reported to be induced by K+ starvation (Armengaud et al., 2004; Shin and Schachtman, 2004; Gierth et al., 2005). Genes for K+ channels although responding to several environmental and hormonal stimuli (Pilot et al., 2003a) do not respond to changes in external K+ supply. It appears that the relative contribution of individual K+ channels to K+ nutrition is primarily determined at the protein level; thus, channel gating is modulated by external K+ as well as several second messengers potentially involved in nutrient signalling (e.g. Ca2+ , pH, nitric oxide, reactive oxygen species [ROS] and cyclic nucleotides [CNs]; see Section 6.2.1 and Amtmann et al., 2006). Nutrient transport is adjusted to not only the availability of nutrients in the environment but also the nutritional status and requirement of the plant (Figure 6.1). Negative feedback control of nutrient uptake by primary assimilates has been shown in several cases. For example, a rise of plant amino acid levels decreases NRT2.1 expression. In tobacco, glutamine is the most effective inhibitor (Krapp et al., 1998), whereas in arabidopsis and barley, arginine or asparagine is more effective than glutamine (Zhuo et al., 1999; Vidmar et al., 2000). Glutamine seems to be involved in

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S

/+N

P

-K Ckn /CRE1 OAS

Suc, Fru, Glu Sac3

Gln, Asn, Arg

GSH, Cys

ROS

PHR SULTR

PT P1BS

H2PO4

AMT

NRT

HAK

SURE

SO42

NH4+

NO3

K+

Figure 6.1 Transcriptional regulation of nutrient transport. The figure gives an overview of stimuli and pathways regulating genes encoding phosphate transporters (PT), sulphate transporters (SULTR), ammonium transporters (AMT), nitrate transporters (NRT) and potassium transporters (HAK). Transporters are induced by depletion (or resupply) of the respective nutrient in the soil. Feedback control is exerted by primary assimilates such as glutathione (GSH) or amino acids (Cys, cysteine; Gln, glutamine; Asn, asparagine; Arg, arginine). Nutrient uptake is linked to the photosynthetic rate (light) through soluble sugars (Suc, sucrose; Fru, fructose; Glu, glucose). In some cases there is evidence for the involvement of specific hormones (Ckn, cytokinin), kinases (CRE1, Sac3) and transcription factors (PHR) in signaling. P1BS and SURE are P- and S-responsive promoter cis-elements respectively. For further details and references, see text.

feedback control of ammonium uptake. Thus, induction of the arabidopsis ammonium transporter AMT1;1 in response to resupply of ammonium nitrate occurs only if assimilation of ammonium into glutamine is inhibited by methionine sulphoximine (Rawat et al., 1999). By contrast, in rice, glutamine induces the expression of OsAMT1;1 (Sonoda et al., 2003) indicating that glutamine can act as a metabolic trigger for both down- and up-regulation of AMT genes depending on the individual gene and the plant species (Loque and von Wiren, 2004). Metabolites of the sulphur assimilation pathway inhibit transcription of sulphate transporters. Barley HvST1 and arabidopsis SULTR1;1 and SULTR2;1 are repressed by glutathione (GSH) and cysteine (Smith et al., 1997; Maruyama-Nakashita et al., 2004c). O-acetyl-l-serine (OAS, a precursor of cysteine synthesis) overrides the negative feedback regulation of HvST1 by GSH (Smith et al., 1997). Both arabidopsis genes are also up-regulated by OAS albeit again with different sensitivity (Maruyama-Nakashita et al., 2004c). Nutrient uptake is tightly linked to carbon status, and thus indirectly controlled by environmental factors that affect the photosynthetic rate. Lejay et al. (2003) tested a number of root ion transporters for regulation by photosynthesis. The arabidopsis genes Amt1.1, Amt1.2 and Amt1.3, NRT1.1, NRT1.2, Hst1, AtPT2 and AtKUP2 were all repressed during the dark. This repression was prevented by continuous light,

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or by adding sucrose at the beginning of the dark period, indicating a link to the photosynthetic rate rather than the circadian rhythm. The authors found a strong correlation between the stimulating effects of light and sucrose and measured an increase in the concentration of soluble sugars in the root tissue during the light period. Neither 2-oxoglutarate nor malate mimicked the effect of sucrose, making it unlikely that carboxylic acids, providers of carbon skeletons for amino acids, play a role in gene repression during the dark period (Lejay et al., 2003). Lejay and colleagues went on to investigate a possible role of known sugar signalling pathways in the regulation of these transporters. Sucrose does not appear to be the signal per se, since glucose and fructose, which do not increase during sucrose application, had even stronger effects than sucrose. Hexokinase (HXK) has been postulated to be a major sugar sensor and a regulatory element for cross talk between C and N metabolism (Sheen et al., 1999; Moore et al., 2003). Experiments using sugar analogues 2-deoxyglucose (2-DOG) and mannose, which are phosphorylated by HXK but are poorly metabolised by glycolysis, and 3-O-methylglucose (3-OMG), which is not metabolised at all, suggested that HXK is required and sufficient for the creation of the sugar signal independent of its function in sugar metabolism (Jang and Sheen, 1994). However, this is not the case for light regulation of nutrient transporters (Lejay et al., 2004). Expression of NRT2.1 was repressed rather than stimulated by 2-DOG or mannose, and glucosamine, an inhibitor of HXK, decreased mRNA levels of NRT2.1 even when sucrose was applied. Thus HXK activity is required but not sufficient to maintain expression of NRT2.1. Similar results were obtained for NRT1.1, AMT1.2 and AMT1.3 and HST1 (Lejay et al., 2003). Further support for the notion that HXK signalling is not involved in light regulation of NRT2.1 came from the observation that the transcriptional response of NRT2.1 to sucrose and light is maintained in sugar-sensing mutants rsr1, sun6 and gin1-1 as well as hxk mutants with altered signalling. Induction of NRT2.1 by sucrose and glucose is, however, abolished in HXK antisense plants, suggesting that catalytic activity and carbon metabolism downstream of HKX are required for sugar regulation of this transporter. A link to carbon metabolism was also established for sulphate uptake. Adenylyl sulphate (APS) reductase, a key enzyme in sulphate reduction, is stimulated by sucrose and glucose (Kopriva et al., 1999), and addition of glucose and sucrose enhances the response of SULTR1;1 and SULTR1;2 to S starvation (MaruyamaNakashita et al., 2004c). Conversely, depletion of carbon sources from the media attenuates induction of SULTR1;1 and SULTR1;2 in sulphur-free medium. Uptake of individual nutrients is linked not only to carbon status but also to the availability of other nutrients. Thus expression of SULTR1;1 and SULTR1;2 responds to nitrogen supply. Low N attenuates the induction of these genes in response to S starvation. The connection point between N and S may lie in the OAS pool (Maruyama-Nakashita et al., 2004c). OAS production involves an amino transfer reaction and therefore depends on the supply of N. P supply does not influence the expression of sulphate transporters but affects the expression of AMT1.1, NRT1.1 and NRT 2.1 (Wang et al., 2002; Wu et al., 2003). It has been suggested that this involves a systemic sucrose signal (Liu et al., 2005). NRT2.1 is also down-regulated during K+ starvation (Armengaud et al., 2004). In this case, amino acids rather than

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sugars appear to act as signals, and it has therefore been hypothesised that enzymes involved in N assimilation are directly dependent on K+ (Amtmann et al., 2006).

6.2.3

Stress responses

The regulation of transport processes is essential for plant responses to biotic and abiotic stresses. For example, acclimation to drought stress (see Chapter 15) requires enhanced water and solute uptake, acclimation to salt stress requires export and compartmentalisation of toxic Na+ , and defence against pathogens requires the release of secondary metabolites and nutrient recycling from necrotic tissue. One example of transporter regulation during drought stress is the regulation of stomatal K+ channels during drought stress, which occurs downstream of an ABA signal as described above (Section 6.2.1). Transporters not only are signalling targets of environmental stress but are also required for signalling as they carry signalling molecules and their precursors between different cellular compartments, cells and tissues. One of the best described responses to environmental stress involving membrane transporters both as stress receptors and as stress targets is acclimation to salt stress. Salt stress caused by high levels of external NaCl leads to the build-up of toxic Na+ in the cytoplasm unless counteracted by the action of transport systems that either export Na+ back into the apoplast or compartmentalise it in the vacuoles (see Chapter 14 for discussion of the effects of salinity on plants). The first measurable signal of salt stress is a rise in cytoplasmic Ca2+ (Knight et al., 1997). It is likely that this rise in cytoplasmic Ca2+ is caused by Ca2+ influx through depolarisation-activated Ca2+ channels (Miedema et al., 2001), as Na+ entry into the cell through nonselective channels (Demidchik et al., 2002) and other Na+ -permeable transporters (e.g. HKT1; Rus et al., 2001) will cause membrane depolarisation (Amtmann et al., 2004; Volkov and Amtmann, 2006). Crucial components of the signalling pathway leading from the initial Ca2+ signal to the enhanced activity of Na+ export systems were discovered by Zhu and colleagues. In a root-bending assay of an ethane methyl sulphonate mutated population of A. thaliana, they identified three loci that resulted in sos mutants (Wu et al., 1996). Positional cloning identified SOS1 as a plasmamembrane-located Na+ /H+ antiporter (Shi et al., 2000), SOS2 as a serine/threonine protein kinase (Liu et al., 2000) and SOS3 as a calcineurin B-like protein containing three EF hands for Ca2+ binding (Liu and Zhu, 1998). Unlike calcineurin, SOS3 is not a phosphatase but interacts physically with the SOS2 kinase (Halfter et al., 2000). Mutations of the SOS2 protein in the N-terminal kinase activation loop and the Cterminal autoinhibitory domain result in constitutive kinase activity. The mutated kinase increases plasma membrane Na+ /H+ exchange activity in wild type as well as in sos2 and sos3 knockout mutants but not in sos1 mutants, indicating that SOS2 acts upstream of SOS1 but downstream of SOS3. Previously it had been shown that interaction between SOS3 and SOS2 leads to activation of substrate phosphorylation by SOS2. It is therefore likely that the SOS2/SOS3 complex activates SOS1 by phosphorylation. However, activation of Na+ /H+ antiport activity by constitutively active SOS2 was weaker in sos2/sos3 mutants than in wild type and differed between sos2 and sos3 genotypes, indicating that other elements required for a fully responsive SOS3 protein are missing or altered in these mutants (Qiu et al., 2002).

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Recent experiments showed that the Na+ /H+ antiporter NHX1 and the Ca2+ /H+ antiporter CAX1, both residing in the tonoplast, are also targets of SOS2 regulation, but in this case SOS2 acts independently of SOS3 (Cheng et al., 2004; Qiu et al., 2004). Both SOS3 and SOS2 are members of large gene families, and it is likely that many other transporters are regulated by SOS2/SOS3 proteins. This raises the exciting possibility that the SOS signalling pathway represents a general link between ionic conditions in the environment and cellular ion homeostasis. Each SOS regulatory module would be formed by a specific signalling triplet consisting of a Ca2+ -binding protein, a kinase and a target transporter (Cheng et al., 2004). Microarray experiments have revealed that transcriptional regulation of ion transporters occurs in response to many abiotic stresses (see for example Fowler and Thomashow, 2002; Kreps et al., 2002; Seki et al., 2002; Maathuis et al., 2003), but in most cases the role of the individual transporters and their regulation in stress acclimation remains to be characterised. Interesting clues can be obtained by comparing expression patterns of ion transporters between plants that are either sensitive or tolerant to the applied stress. For example, AKT1-type K+ channels required for K+ uptake into root cells are differentially regulated by salt stress in salt-tolerant and salt-sensitive rice varieties (Golldack et al., 2003). The vacuolar V-type H+ -ATPase is essential for creating the driving force for Na+ accumulation in the vacuole during salt stress and is generally up-regulated during salt stress. However, the individual subunits targeted by salt-induced transcriptional regulation appear to differ between halophytes and glycophytes (Dietz et al., 2001; Kluge et al., 2003; Maathuis et al., 2003). Furthermore, comparative microarray analysis of A. thaliana and its salt-tolerant relative Thellungiella halophila (Taji et al., 2004; Volkov et al., 2004; Gong et al., 2005; Wong et al., 2006) indicated that many genes that are up-regulated by salt stress in arabidopsis exhibit already high expression levels in T. halophila under low-salt condition. Hence, T. halophila is evolutionarily ‘prepared’ for high salinity and does therefore no longer require salt-induced regulation of certain ion transporters. Examples include the Na+ /H+ antiporter SOS1, the H+ -ATPase AHA2 (Taji et al., 2004) and a large number of V-type ATPase subunits (B. Wang and A. Amtmann, unpublished results). Differential responsiveness of ion transporters in sensitive and resistant species to abiotic and biotic stress should remind us that observation of regulatory processes under stress conditions does not necessarily indicate successful stress acclimation but in many cases merely shows that the plant has pressed a ‘panic button’. Separation of acclimation from panic responses, especially with respect to the underlying signalling pathways, is a major challenge for the interpretation of experimental data in this field.

6.3

Molecular mechanism of regulation

The previous sections have introduced several physiological situations that involve the regulation of ion transporters. A number of molecular signals including intracellular Ca2+ and pH, protein kinases, transcription factors, metabolites and hormones have been confirmed to be essential components of the signalling pathways linking

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environmental stimuli to changes in ion transport. However, in only a few cases is it known how these pathways finally modulate the targeted transporter. In the remainder of the chapter some of the molecular mechanisms that can regulate transporter genes or proteins are described.

6.3.1

Transcriptional regulation

The molecular components of the transcriptional regulation of nutrient transporters remain largely unknown. Regulation of AtSULTR1;1 requires an upstream region of −3031 bp, whereas −1944 bp are sufficient for tissue-specific expression (Maruyama-Nakashita et al., 2004a). Analysis of deletion mutants in this region identified a 16-bp sulphur-responsive element (SURE) between −2777 and −2762 that is sufficient and necessary for enhanced expression of SULTR1;1 in response to S starvation (Maruyama-Nakashita et al., 2005). Regulation of phosphate transporters and other P-responsive genes is under the control of transcription factors of the MYB-CC family such as PHR1 and PHR2, which act as positive regulators (Rubio et al., 2001; Todd et al., 2004). AtPHR1 recognises a GnATATnC motif, the P1BS element. However, although the P1BS motif is present in the promoters of many P-regulated genes, it is not overrepresented in P-regulated genes (Hammond et al., 2003). This could indicate that P regulates PHR genes at the post-transcriptional level, or that other promoter elements are required for P-specific responses (Amtmann et al., 2006). Very little is known to date about upstream events linking changes in soil nutrient availability to gene expression of the respective transporters, but there is evidence that they involve the production of ROS, cytoplasmic Ca2+ signalling and kinase/phosphatase activity. Production of ROS occurs in roots in response to K+ starvation and is necessary for the induction of downstream responses including derepression of HAK5 (Shin and Schachtman, 2004; Shin et al., 2005; Figure 6.1). A rise in intracellular Ca2+ signal, although not yet reported, is likely to occur in response to K+ starvation, since microarrays revealed K+ -induced changes in transcript levels of several Ca2+ -binding proteins and Ca2+ -dependent kinases (CDPKs) (Armengaud et al., 2004). Transcriptional regulation of sulphate transporters appears to involve phosphorylation/dephosphorylation events; in arabidopsis, application of phosphatase blockers (okadaic acid, calyculin A) abolishes green fluorescent protein (GFP) expression under the control of the SULTR1;1 promoter in response to sulphur starvation (Marayuma-Nakashita et al., 2004a), and in Chlamydomonas the regulation of sulphate uptake requires an Snf1-like Ser/Thr kinase (Sac3, Figure 6.1; Davies et al., 1999). Phosphorylation is also likely to be involved in the regulation of nitrate transporters, since all NRT2 sequences contain a target sequence for protein kinase C in their C-terminus (Forde, 2000). Finally, several studies indicate a role for plant hormones in mediating nutritional signals. K+ starvation enhances the expression of enzymes involved in the biosynthesis of ethylene (Shin and Schachtman, 2004) and jasmonic acid (Armengaud et al., 2004), and levels of the two hormones increase in roots and shoots of K+ -starved plants respectively. However, the exact position of ethylene and jasmonate signals within the K+ starvation response remains

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to be elucidated. Expression of sulphate (SULTR1;1 and SULTR1;2) and phosphate (PT1) transporters is repressed by cytokinin, and application of cytokinin suppresses their induction by S or P starvation (Martin et al., 2000; Maruyama-Nakashita et al., 2004c; Hou et al., 2005; Figure 6.1). Signal transduction of cytokinin-dependent responses involves the histidine kinase CRE1 (Inoue et al., 2001), and cre1 mutants no longer show a response of SULTR1;1 and SULTR1;2 to cytokinin (MaruyamaNakashita et al., 2004b). The observation that both cytokinin and CRE1 transcript levels decrease during P starvation (Franco-Zorrilla et al., 2005) further supports the notion of CRE1/cytokinin signalling pathway in nutrient responses.

6.3.2

Post-translational regulation

In addition to changes in gene expression, many transporter proteins respond to environmental clues with changes in protein activity. Post-translational modifications of ion transporters are manifold; the best characterised examples involve: r intramolecular interaction, for example with autoinhibitory domains, r protein–protein interaction, for example with 14-3-3 proteins, calmodulins or protein kinasesand phosphatases, r ligand binding, for example ion channel gating by CNs and r interaction between different subunits in heteromeric proteins, for examples betweenα-subunits of Shaker K+ channels. Each of these mechanisms will be described here in some detail using H+ andCa+ -ATPases, Na+ /H+ antiporters and ion channels as examples (Figure6.2).

6.3.2.1

Autoinhibition

Plant P-type H+ -ATPases (cf. Section 5.3.1.1) have a fundamental role in creating the electrochemical proton gradient that drives nutrient uptake and in controlling currents through voltage-dependent ion channels via the membrane potential. Plant Ca2+ -ATPases in plasma- and endomembranes have a similarly important role in the maintenance and recovery of a low base level of cytoplasmic free Ca2+ , which is a prerequisite for Ca2+ signalling. The capacity to regulate both types of pumps is therefore essential. Both proton and Ca2+ pumps contain well-defined regulatory domains in their C- and N-termini respectively (Geisler et al., 2000a; Morsomme and Boutry, 2000; Baekgaard et al., 2005; Figure 6.2). These serve as autoinhibitors and are thought to interact both intramolecularly with other parts of the pump and extramolecularly with activating proteins including 14-3-3 proteins, protein kinases and calmodulin. The first evidence for autoinhibition of plant H+ -ATPases was provided by biochemical experiments; trypsin treatment of plant plasma membranes resulted in H+ -ATPase activation concomitant with cleavage of a C-terminal fragment (DeWitt and Sussman, 1995). Furthermore, complementation of a yeast strain lacking endogenous proton pump activity with plant H+ -ATPases was unsuccessful unless the C-terminus was removed (Palmgren and Christensen, 1993; Baunsgaard et al., 1996;

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AHA/PMA2

ACA2/8 out

out/ER

cyt

cyt

Ca2+

P Ca2+

KCO1

CNGC1/2

NHX1

out

out

vac

cyt

cyt

cyt

P Ca2+

AID

14-3-3 BD

14-3-3

FC

CaMBD

CaM

CNBD

SelRD

Figure 6.2 Post-translational regulation of ion transport. Regulatory domains for autoinhibition, protein–protein interaction and ligand binding are shown for plasma membrane H+ -ATPases (AHA2 and PMA2), plasma membrane and endomembrane Ca2+ -ATPases (ACA2, ACA8), a putative vacuolar K+ channel (KCO1), putative cyclic-nucleotide-gated channels (CNGC1, CNGC2) and the vacuolar Na+ /H+ antiporter NHX1. Transporter topology is shown as suggested by Baekgaard et al. (2005; H+ - and Ca2+ pumps), V´ery and Sentenac (2002; KCO and CNGC) and Yamaguchi et al. (2003; NHX1). Transmembrane spanning domains as well as extracellular (out) vacuolar (vac), ER-luminal (ER) and cytoplasmic (cyt) loops of the proteins are shown as grey lines. Regulatory domains are represented by different symbols as shown at the bottom (AID, autoinhibitory domain; 14-3-3BD, 143-3 protein-binding domain; FC, fusicoccin; CaM, calmodulin; CaMBD, calmodulin-binding domain; CNBD, cyclic nucleotide binding domain; SelRD, regulatory domain controlling ion selectivity). Individual residues involved in AID interaction are shown as small black boxes. For details and references, see text.

Morsomme et al., 1998). Both trypsin treatment and genetic deletion increased maximal pump activity (V max ), decreased the apparent K m and shifted the pH optimum to more alkaline pH values (Palmgren et al., 1990, 1991; Palmgren and Christensen, 1993). Progressive deletion of AHA2 from the C-terminus showed that deletion of 38 residues is sufficient to decreases the K m and achieve the observed shift in pH optimum but deletion of at least 51 and 61 residues is required to achieve full activity and functional expression, respectively, in yeast (Regenberg et al., 1995). A number of point mutations were identified in the tobacco H+ -ATPase PMA2 that release autoinhibition and improve the coupling ratio between proton pumping and ATP hydrolysis (Morsomme et al., 1996, 1998). The majority of these mutations

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are concentrated in two close regions within the first half of the C-terminal domain. Systematic alanine scanning of the C-terminus of arabidopsis AHA2 also revealed two regions (RI and RII) as being important for autoinhibition (Axelsen et al., 1999). These are located between Lys-863 and Leu-885 and between Ser-904 and Leu-919. Plant Ca2+ pumps of the IIB type also contain an autoinhibitory R domain but, in contrast to their animal counterparts, it is located in the N-terminus. It appears, however, that the position of this domain in the C- or N-terminus is not important for its autoinhibitory function. Thus, relocation of the C-terminal R domain in the animal PMCA4b to the N-terminus had only minor effects on autoinhibition (Adamo and Grimaldi, 1998). As in the case of H+ -ATPases, yeast complementation with plant Ca2+ -ATPases is possible only if the autoinhibitory domain, in this case the N-terminus, is deleted (Harper et al., 1998; Chung et al., 2000; Geisler et al., 2000b; Bonza et al., 2004; Schiott et al., 2004; Schiott and Palmgren, 2005). Point mutations in certain single amino acid residues in the N-terminus have the same effect (Curran et al., 2000; Baekgaard et al., 2006). In the arabidopsis Ca2+ pump ACA8 these include Trp-43 and Phe-60, which are directly involved in autoinhibition. In addition to the R domains, several amino acid residues outside the autoinhibitory domains are important for autoinhibition of H+ and Ca2+ pumps (Morsomme et al., 1996, 1998; Curran et al., 2000; Figure 6.2) suggesting that they could be important for intramolecular recognition of the autoinhibitory domain. In the arabidopsis endoplasmic reticulum Ca2+ pump ACA2, several residues that connect the catalytic domain with transmembrane domains are required for autoinhibition, and it was suggested that they interact with the N-terminal autoinhibitory domain (Curran et al., 2000). In the tobacco plasma membrane proton pump PMA2, point mutations leading to enzyme activation were discovered in the N-terminus, the first and fourth transmembrane spanning domain and the small and large cytoplasmic loops (Morsomme et al., 1996, 1998). Partial tryptic digestion showed that these mutations cause a conformational change that makes the C-terminus more accessible to trypsin. Hence, it is likely that these modifications result in the displacement of the C-terminal inhibitory domain from its interaction site.

6.3.2.2

14-3-3 proteins

14-3-3 proteins are highly conserved proteins that regulate activities of a wide range of targets via direct protein–protein interaction. Short amino acid motifs of the target protein containing phosphoserine or phosphothreonine are bound in a conserved amphipathic region present in each monomer of a dimeric 14-3-3 protein. Hence, interactions depend on the phosphorylation status of the targets and can involve either one or two targets at the same time (Roberts, 2003). In their function as activators/ repressors, adapters or chaperones, they are involved in many cellular processes (Aitken, 1996). In plants, targets of 14-3-3 proteins include metabolic enzymes (e.g. nitrate reductase; Huber et al., 2002; Comparot et al., 2003), transcription factors (e.g. VP1; Schultz et al., 1998), protein kinases (e.g. CDPK2; Camoni et al., 1998) and ion transporters. Regulation of plant P-type H+ -ATPases by 14-3-3 proteins was discovered through the action of fusicoccin (FC), a fungal toxin that provokes

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membrane hyperpolarisation and acidification of the external medium (De Boer, 1997). The features of H+ -ATPases isolated from FC-treated plants resembled those obtained by C-terminal deletion (i.e. higher V max , lower K m , shift of pH optimum; see above) suggesting that FC modifies the C-terminus (Johansson et al., 1993; Lanfermeijer and Prins, 1994). The mystery of how FC regulates plant proton pumps was solved when a 30-kDa protein doublet present in FC receptor preparations was cloned and identified as a member of the 14-3-3 protein family (Korthout and de Boer, 1994; Marra et al., 1994; Oecking et al., 1994). It was subsequently shown that 14-3-3 proteins bind directly to the C-terminus of plant H+ -ATPases and that this interaction is stabilised by FC (Jahn et al., 1997; Oecking et al., 1997; Fullone et al., 1998). 14-3-3 protein binding involves a unique region in the extreme end of the Cterminus which partly overlaps with the RII autoinhibitory domain (Jelich-Ottmann et al., 2001; Fuglsang et al., 2003; Figure 6.2). The 14-3-3 binding domain includes two phosphorylation sites. Phosphorylation of the penultimate threonine residue of the C-terminus (Thr-947 in AHA2) is required to stabilise 14-3-3 binding (Fuglsang et al., 1999; Svennelid et al., 1999; Maudoux et al., 2000). Phosphorylation of this site in vivo has been shown in guard cells in response to blue light (Kinoshita and Shimazaki, 1999) and in root cells in response to aluminium stress (Shen et al., 2005) but the kinase that phosphorylates the threonine residue has not yet been identified. Instead, a kinase of the CDPK family, PKS5, has been shown to phosphorylate a serine residue situated further upstream in the C-terminus (Baekgaard et al., 2005). Phosphorylation of this site (Ser-931 in AHA2) leads to a decrease of 14-3-3 binding and thus to decreased activity of the pump. In vitro experiments identified several phosphatases including alkaline phosphatase (Fuglsang et al., 1999) and PP2A (Camoni et al., 2000) that can disrupt the interaction between the H+ pump and 14-3-3 proteins. First evidence that 14-3-3 proteins regulate ion channels came from the observation that overexpression of a 14-3-3 gene in tobacco (Saalbach et al., 1997) and addition of recombinant 14-3-3 proteins to tomato suspension cells (Booij et al., 1999) resulted in increased K+ outward currents. Since then, modulation of several types of vacuolar (‘slow vacuolar’, SV, and ‘fast vacuolar’ FV) and plasma membrane (inward and outward) K+ currents by 14-3-3 proteins has been reported (De Boer, 2001; Van den Wijngaard et al., 2005), including both activation and inhibition, but linking physiological results with molecular studies proved difficult sometimes. Using surface spectron resonance Sinnige et al. (2005) showed that the barley two-pore K+ channel HvKCO1 (Figure 6.2), previously characterised as a component of the SV current (Schonknecht et al., 2002), interacts with three barley 14-3-3 proteins out of which 14-3-3A exhibits the highest affinity. By contrast, SV currents were strongly reduced by 14-3-3B and 14-3-3C but not by 14-3-3A. A possible explanation for this discrepancy was provided by the recent discovery that SV currents are in fact created by TPC1, another two-pore channel that differs from KCO1 in its topology (Peiter et al., 2005). Hence, the identification of KCO1mediated currents in planta and the role of KCO1 regulation by 14-3-3 proteins for cellular K+ transport now require reassessment. A physiological role of 14-3-3 regulation of K+ channels in seed germination was recently identified in barley

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(Van den Wijngaard et al., 2005). The second phase of germination, apparent in embryonic root (radicle) emergence, is inhibited by ABA and stimulated by FC. ABA and FC also have opposite effects on K+ uptake into the radicle (inhibited by ABA and stimulated by FC) and on K+ currents across the plasma membrane of protoplasts derived from the radicle (ABA reduces K in , FC reduces I K,out and stimulates I K,in ). Inclusion of recombinant barley 14-3-3B protein in the patch pipette solution (cytoplasmic side) resulted in a fast decline of K out currents by 60%, whereas K in currents were increased. Furthermore, addition of a 14-3-3 binding protein to the pipette solution completely abolished I K,in , showing that residual K in in the previous experiment was due to bound endogenous 14-3-3 proteins. Hence, 14-3-3 proteins emerge as an important component of the signalling pathway leading to ABA-dependent seed dormancy. One possible scenario is that ABA activates protein serine/threonine phosphatases ABI1 and ABI2 (Armstrong et al., 1995) causing channel dephosphorylation, dissociation of 14-3-3 protein and channel inhibition (Van den Wijngaard et al., 2005).

6.3.2.3

Calmodulin

Calmodulins (CaM) are small Ca2+ -binding proteins that can translate intracellular Ca2+ signals into a variety of cellular responses. In accordance with this function, CaM are involved in plant responses to a large number of environmental stimuli (Snedden and Fromm, 2001). Targets of this large gene family (approximately 28 members in arabidopsis) include membrane transporters such as Ca2+ -ATPases, putative cyclic-nucleotide-gated channels (CNGCs) and the vacuolar Na+ /H+ antiporter NHX1 (Figure 6.2). The interaction between Ca2+ , CaM and Ca2+ -ATPases creates a negative feedback loop that instigates removal of Ca2+ from the cytoplasm as soon as cytoplasmic Ca2+ levels start to rise. The relative kinetics of the influx of Ca2+ and its removal by Ca2+ pumps shape the Ca2+ signal and determine its frequency, thereby endowing it with some specificity (Plieth, 1999). Upon binding of Ca2+ , calmodulin interacts with the N-terminus of IIB-type Ca2+ pumps and leads to increased activity. There is no consensus CaM-binding site but they are usually 15–30 amino acids long and form an α-helix containing two bulky hydrophobic residues that function as anchors for CaM (Crivici and Ikura, 1995; Yap et al., 2000). N-terminal CaM domains (CaMBD) have been identified in the cauliflower Ca2+ -ATPase BCA1 (Malmstr¨om et al., 1997) and in the arabidopsis Ca2+ pumps ACA8 and ACA9 (Bonza et al., 2000; Schiott et al., 2004). The CaMBD in ACA8 spans from Arg-43 to Lys-68, and the conserved Trp-47 and Phe-60 act as hydrophobic anchor residues for CaM binding. Eleven more hydrophobic or basic residues were found to be important for the stability of the CaM complex. CaM binding interferes with autoinhibition involving six residues in the CaMBD. These include the hydrophobic anchor residues Trp-43 and Phe-60 suggesting that these residues have a dual function in CaM recognition and in autoinhibition (Baekgaard et al., 2006). Kinases are again important modulators of this regulation, but in contrast to their stabilising effect on the C-terminal 14-3-3 protein complex in proton pumps, they seem to inhibit CaM action in Ca2+ pumps. For example, it was shown that the endomembrane Ca2+ -ATPase ACA2 is

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phosphorylated by a CDPK at Ser-45 near the CaMBD, and that this phosphorylation results in inhibition of CaM stimulation and of basal activity (Hwang et al., 2000). The physiological meaning of this apparently inverse effect of intracellular Ca2+ on ACA2 via CaM and CDPK remains to be studied. Recent results from the Blumwald laboratory (Yamaguchi et al., 2003, 2005) have raised the surprising possibility that CaM acts not only in the cytoplasm but also within the vacuole. In a yeast two-hybrid screen they identified a CaM-like protein AtCaM15 as an interacting partner of the vacuolar H+ /Na+ antiporter AtNHX1. AtCaM15 belongs to a subgroup of CaM that somewhat diverges from the more conserved members of the same gene family (CaM1–CaM7). A further yeast two-hybrid assay confirmed CaM15 as a CaM, since it interacted with a known CaM-binding protein, and also showed that CaM15 interacts with the C-terminus of NHX1. Progressive deletions of the C-terminus mapped the binding site to a region between Val-498 and Gly-518. This region indeed has the potential to form a positively charged amphiphilic helix, a characteristic feature of CaM-binding domains. The interaction between AtNHX1 and CaM15 appears to be specific, as it did not occur with a CaM from petunia (CaM81), which has a conserved CaM motif and is identical to AtCaM7. Pull-down assays with FLAG-tagged AtCaM15 showed that CaM binding to NHX1 is dependent on the presence of Ca2+ (1–10 mM) and increases with acidic pH (pH 7.5–5.5). In a previous study, the same group had shown that the C-terminus of AtNHX is located in the vacuolar lumen (Yamaguchi et al., 2003). Treating isolated vacuoles of yeast expressing FLAG-tagged AtCaM15 with proteinase K in either the presence or the absence of Triton X-100 showed that CaM15 could be detected only in the absence of the detergent, indicating that it is indeed localised inside the vacuolar lumen. The vacuolar localisation was subsequently confirmed in planta by fluorescence microscopical analysis of leaf protoplasts transformed with EGFP (enhanced green fluorescent protein) tagged AtCaM15. What could be the physiological relevance of CaM signalling in the vacuole? Regulation of NHX1 by CaM seems to target the cation selectivity of this transporter. The study of Yamaguchi et al. (2005) showed that CaM binding to NHX1 decreased the V max of Na+ /H+ antiport, whereas the V max for K+ /H+ antiport activity was unchanged. Previously they had shown that C-terminal deletion of NHX1 increased the Na+ /K+ selectivity of the antiport activity (Yamaguchi et al., 2003). On the basis of the pH dependence of CaM binding to NHX1 the authors suggest that transient increase in vacuolar pH would release CaM from the C-terminus, thereby increasing the potential of the vacuole to accumulate Na+ .

6.3.2.4

Cyclic nucleotides

Cyclic nucleotides, cGMP and cAMP, are used for signal transduction by many organisms including animals, fungi, bacteria and algae, and there is increasing evidence that higher plants use cGMP as a secondary messenger, for example in chloroplast development and pathogen defence (Bowler et al., 1994; Durner et al., 1998). A rise of intracellular cGMP levels has recently been measured after the onset of drought and salt stress (Donaldson et al., 2004). Treatment of arabidopsis plants with membrane-permeable cGMP leads to transcriptional changes in a large

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number of genes (Maathuis, 2006) among which transporters for monovalent cations are overrepresented. Potential direct targets of CN signalling include protein kinases as well ion channels (Maathuis, 2006). CNGCs in animals are non-selective among cations and play an important role in the transduction of visual and olfactory signals (Finn et al., 1996). Animal CNGCs are activated by binding of cAMP or cGMP to a regulatory domain in the C-terminus. Additional control of gating is exerted by a CaM-binding site in the N-terminus, which interacts with the CN-binding site thereby lowering the affinity for cAMP and cGMP (Finn et al., 1996). In plants, Shaker K+ channels are regulated by CNs (Hoshi, 1995; Gaymard et al., 1996) but the response is slow and requires high concentrations of CN. Hence, although these channels contain putative CN-binding sites, a direct interaction is unlikely. CN action could be exerted through phosphorylation events; for example in V. faba mesophyll cells, a PKA-like kinase is involved in the stimulatory effect of cAMP on the K+ outward rectifier (Li et al., 1994). The first plant CNGC was identified from barley on the basis of its binding to CaM (Schuurink et al., 1998). Homologues of HvCBT1 and animal CNGCs have since been identified in many plant genomes including tobacco, rice and arabidopsis (Talke et al., 2003, for review). Unfortunately, plant CNGCs have proven to be rather recalcitrant to expression in heterologous systems, probably due to the requirement for heterotetramerisation between and αand β-subunits (Maathuis, 2004). Nevertheless, heterologous expression has been achieved for AtCNGC1, CNGC2, AtCNGC4 and NtCBP4. Interestingly, these experiments revealed some features of plant CNGCs that are unknown in their animal counterparts, such as K+ selectivity and inward rectification, but they agreed in the observation that currents were activated by CN (Leng et al., 1999, 2002; Balague et al., 2003; Hua et al., 2003). By contrast, patch-clamp experiments showed that the activity of voltage-independent non-selective cation channels in arabidopsis root cells decreases after addition of CN (Maathuis and Sanders, 2001). This channel type is likely to play a role in Na+ uptake during salt stress (Demidchik et al., 2002), and indeed arabidopsis seedlings treated with membrane-permeable CNs display increased salt tolerance (Maathuis and Sanders, 2001). In plant CNGCs, the CN-binding domain is not located in the N-terminus as in their animal homologues but is located in the C-terminus where it overlaps with the CaM-binding domain (Arazi et al., 2000; Figure 6.2). Yeast two-hybrid assays confirmed the interaction between the C-termini of CNGC isoforms (CNGC1 and CNGC2) and members of the CaM gene family (CaM2 and CaM4; K¨ohler et al., 1999). It has therefore been suggested that the binding of CaM at the C-terminus might interfere with CN binding. Indeed, whole-cell patch-clamp experiments showed that inclusion of AtCaM4 in the (cytoplasmic) pipette solution resulted in a decrease of cAMP-induced currents in HEK cells expressing AtCNGC2 (Hua et al., 2003). Interaction of CaM and CN binding within the C-terminus implies a different regulatory mechanism from the one operating in animal CNGCs. Interestingly, differential regulation of individual CNGC isoforms is indicated by the observation that CaM-binding domains of AtCNGC1 and AtCNGC2 differ in their affinity for CaM (K¨ohler and Neuhaus, 2000). However, there is still a long way to go to understand the physiological role and regulation of CNGCs in plants.

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Heteromerisation

Membrane transporters are often assembled from several subunits. Heterooligomerisation involving several transporter isoforms is likely to be a common phenomenon but has only recently attracted interest from the point of view of transporter regulation. For example, observation of interaction between members of the AMT1 family of ammonium transporters has led to the hypothesis that this type of protein–protein interaction is a regulatory element in AMT protein activity (Ludewig et al., 2003; Loque and von Wiren, 2004). To date, the best studied example of heteromeric protein assemblies is K+ channels of the Shaker family. Functional channels consist of four α-subunits, which in the simplest case form homotetramers. Biochemical experiments and yeast two-hybrid studies have revealed that in plant Shaker channels, interaction between the individual subunits involves three domains in the C-terminal region. The KHA domain at the extreme C-terminus cross-interacts with a region just downstream of the hydrophobic core, and the putative CN-binding domain interacts with itself (Daram et al., 1997). There is now increasing evidence that functional channels can also be provided by heterotetramers. First indication for heterotetramerisation came from the observation that co-expression of different plant Shaker channel transcripts in Xenopus laevis oocytes produced currents that could not be explained by simply adding homotetrameric channel currents (Dreyer et al., 1997). Out of the five subgroups of Shaker channel genes (Pilot et al., 2003b), interaction occurs, at least in heterologous systems, between group I (AKT1-type) and group II (KAT1-type) α-subunits, group III (AKT2-type) and group IV (AtKC-type) α-subunits, and also between group II and group III α-subunits (Dreyer et al., 1997; Baizabal-Aguirre et al., 1999; Pilot et al., 2001, 2003a; Zimmermann et al., 2001). The question remains whether heterotetramerisation occurs in planta and whether it provides a physiological means for regulating K+ currents. The analysis of expression patterns shows that many tissues express at least two types of Shaker channels, thus providing the opportunity for the formation of hybrid channels (Cherel, 2004). However, in most cases it has not been analysed whether co-expressed channels are co-localised in the same membrane. KAT1 and KAT2 are both expressed in guard cells and have been shown to interact in heterologous expression systems (Pilot et al., 2001). Evidence for KAT1/KAT2 interaction in guard cells comes from the apparently contradictive observations that overexpression of mutant KAT1 channels affects stomatal function (Kwak et al., 2001) while KAT1 knockout does not (Szyroki et al., 2001). More intriguingly even, AtKC1, which does not form functional homotetrameric channels in heterologous expression systems, might act as a modulator of AKT1 currents. Both genes are expressed in root hairs but knockout of AKT1 alone is sufficient to completely abolish K+ inward current, thus confirming AtKC as a ‘silent channel’. Nevertheless, disruption of the AtKC gene results in qualitative changes of the inward current affecting its Ca2+ and pH sensitivity (Reintanz et al., 2002) suggesting that AKT1/AtKC heteromers underlie physiological K+ currents in root hairs. Hence, heterotetramerisation of a limited number of co-expressed but differentially regulated subunits has the potential to produce a plethora of K+ currents within a single cell. This would explain why in many cases electrophysiological studies on

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plant cells reveal a complex picture of ion currents that is difficult to reconcile with individual channel features detected in heterologous expression systems.

6.4

Traffic of ion transporters

Eukaryotic cells maintain a traffic of vesicles to shuttle membrane material, proteins and soluble cargo between endomembrane compartments, the plasma membrane and the extracellular space. Vesicles are formed by budding and constriction at the formative membrane surface, and their delivery is achieved by fusion and intercalation with the lipid bilayer of the target membrane (Pratelli et al., 2004; Surpin and Raikhel, 2004). These processes sustain membrane turnover and must be integrated so as to populate cellular membranes with ion transport proteins and to maintain their homeostatic functions. For vesicle fusion in plants there is a growing body of kinetic and physiological data that bear on traffic control, at least at the plasma membrane where physical access is possible in vivo. Exocytotic and endocytotic events at the plasma membrane have been identified with stepwise changes in capacitance that accompany the increase or decrease of membrane surface area during vesicle membrane fusion and removal, respectively (Thiel and Battey, 1998; Blatt and Thiel, 2003). These changes in capacitance are consistent in size with the predicted vesicle dimensions derived from ultrastructural studies (Picton and Steer, 1983; Phillips et al., 1988) and from imaging studies using fluorescent styryl dyes to label internalised membrane (Meckel et al., 2004). Factors shown to affect vesicle traffic in plants include cytosol-free Ca2+ concentration ([Ca2+ ] i ), guanosine nucleotides (Homann and Tester, 1997; Carroll et al., 1998) and osmotic changes (Kubitscheck et al., 2000). Furthermore, evidence for [Ca2+ ] i -dependent and -independent exocytotic pathways underscores the complexity of secretory processing that must occur in parallel within individual cells (Homann and Tester, 1997; Sutter et al., 2000). By contrast, information remains scarce that can speak to the partitioning and delivery of specific ion transport proteins to various target membranes within the cell. Vesicle traffic has been implicated in the spatial distribution of the auxin efflux carrier Pin1 and its sensitivity to the ARF-GEF inhibitor Brefeldin A (Steinmann et al., 1999; Geldner et al., 2001) that disrupts Golgi structure and trafficking (Nebenfuhr et al., 2002). Traffic of the H+ -ATPase has also been suggested to underpin auxinstimulated H+ extrusion and parallel increases in H+ -ATPase protein that take place over a similar timescale (≥10 min) at the plasma membrane (Hager et al., 1991) and may be related to a concerted targeting to the plasma membrane (Lefebvre et al., 2004). Nonetheless, only recently has attention turned to vesicle traffic as a means to controlling specific transporter activities, notably of selected ion channel proteins, and its relation to changes in cell volume. Two developments have placed membrane traffic squarely at the forefront of research in this respect. First, the biophysical studies of Homann, Thiel and colleagues have demonstrated that a reversible exchange of K+ channels occurs in guard cell protoplasts during osmotically driven changes in cell volume. Homann (1998) and Kubitscheck et al. (2000) measured events of exo- and endocytosis from the plasma membrane of

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V. faba guard cell protoplasts using capacitance recording and styryl dyes. Capacitance recording makes use of sine wave retardation to determine membrane surface area (Penner and Neher, 1989; Angleson and Betz, 1997; Thiel and Battey, 1998) and yields kinetic information about vesicle traffic when combined with fluorescence analysis of styryl dye distributions. These studies demonstrated a close coordination of vesicle traffic with cell volume. Their subsequent work indicated that traffic of the predominant K+ channels was integrated with these surface area changes in such a way that the balance of channel population densities was maintained (Homann and Thiel, 2002). Finally, Hurst et al. (2004) and Meckel et al. (2004) have confirmed that these cells respond to osmotic challenge with endo- and exocytosis of a GFP-tagged KAT1 K+ channel after transient biolistic transfections. Details of the molecular mechanism(s) are lacking, but the kinetics are sufficiently rapid to suggest that ion channel traffic – and changes in channel population – may contribute significantly to channel control in some circumstances. Nonetheless, questions still hang over the interpretation of these findings. Most importantly, osmotically driven vesicle traffic appears non-selective, and therefore differs fundamentally from the physiological regulation of channel activities, for example during stomatal movements (Hurst et al., 2004). Thus, how widespread are these phenomena and the circumstances in which they prevail will need to be established. The second development has come from the identification of two SNARE (soluble NSF [N-ethylmaleimide-sensitive factor] attachment protein receptors) proteins associated with ABA signalling in tobacco and arabidopsis (Leyman et al., 1999). SNAREs comprise a group of membrane proteins that are conserved among all eukaryotes and form the core of the molecular machinery for vesicle trafficking and membrane fusion (Jahn et al., 2003; Pratelli et al., 2004; Sutter et al., 2006). Complementary SNAREs, identified by their core residues (either arginine [R] or glutamine [Q]), are localised to different target membrane compartments and vesicles, and interact to form a tetrameric bundle of coiled helices that draws the membrane surfaces together and facilitates fusion. Furthermore, specificity in SNARE interactions is thought to contribute to membrane recognition and vesicle targeting (Mcnew et al., 2000; Paumet et al., 2004). In this context, it is not surprising that the plasma membrane Q-SNAREs NtSyp121 (NtSyr1) from tobacco and AtSyp121 (AtSyr1) from arabidopsis are associated with ABA. Drought stress and ABA have profound effects on cellular compartmentation and cell volume, especially in guard cells. What is noteworthy, however, is that both SNAREs were initially identified in a screen for ABA receptors using an expression-cloning strategy. Furthermore, NtSyp121 was cleaved by the Clostridium botulinum neurotoxin BotN/C and, intriguingly, both neurotoxin treatments and the (so-called Sp2) domain corresponding to the soluble cleavage fragment blocked K+ and Cl− channel responses to ABA when loaded directly into guard cells (Leyman et al., 1999). How might a membrane-trafficking protein account for such rapid and selective channel control? One clue has come from studies of KAT1 K+ channel trafficking and localisation in the presence of the Sp2 domain of these SNAREs. Sutter et al. (2006) made use of a dual labelling strategy, incorporating a pair of haemagglutinin epitopes in the extracellular loops of KAT1 and a photo-activatable GFP (Patterson

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and Lippincott-Schwartz, 2002) at its C-terminus. They found that KAT1 traffic to the plasma membrane was suppressed by the Sp2 domain of both NtSyp121 and AtSyp121 such that roughly 60% of the label was retained within the endoplasmic reticulum, and the effect was selective for the K+ channel when compared with the PMA2 H+ -ATPase. By contrast, both the K+ channel and H+ -ATPase were retained within the endoplasmic reticulum when co-expressed with a dominantnegative Rab1b mutant that blocks export to the Golgi apparatus (Batoko et al., 2000). This shift to endosomal accumulation implies a backlog of synthesised protein that built up on restricting traffic to the plasma membrane, and offers some underpinning for possible actions of ABA. Notably, the specificity of Sp2 for the K+ channel demonstrates that the trafficking pathways of these two integral membrane proteins diverge at a stage post-Golgi, late in transit to the plasma membrane. Thus both the K+ channel and H+ -ATPase must pass through the Golgi apparatus en route to the plasma membrane. At present, however, the basis for KAT1 selectivity is unresolved. For example, it is possible that the KAT1 K+ channel uniquely partitions in association with NtSyp121 and its homologue AtSyp121 late in transfer to the plasma membrane. This interpretation accords with recent evidence in nerve and epithelia for differential trafficking and distributions, even among subsets of Kv- and Kir-type K+ channels (Ma et al., 2001, 2002; Leung et al., 2003; Rivera et al., 2003; Misonou and Trimmer, 2004). Alternatively, however, the traffic and targeting observed for KAT1 may be a general characteristic of a larger subset of plasma membrane proteins, and the H+ -ATPase represents an exception to this rule. Lefebvre et al. (2004) reported that traffic of the PMA2 H+ -ATPases depends on presumably novel and, as yet, unidentified domains within the large, cytosolic loop internal to the protein sequences. Furthermore, Geelen et al. (2002) noted previously that transit of secretory cargo (a soluble GFP) is also sensitive to the Sp2 domain of NtSyp121, suggesting that the K+ channel follows the same export pathway. These are only two illustrative explanations, and they serve to highlight our substantial ignorance about traffic at the plasma membrane. Still more intriguing, localisation of the K+ channel at the plasma membrane was strongly altered in the presence of the Sp2 domains. Whereas KAT1 normally was found anchored within surface microdomains of approximately 0.5-μm diameter, when co-expressed with the Sp2 domains distribution of the K+ channel was diffuse and the protein was mobile within the plane of the membrane. Again, these observations do not speak directly to channel control by ABA, but they do bear witness to actions of the Sp2 domains that are separate from their effects on traffic to the plasma membrane and, thus, indicate roles for the SNAREs that may link directly to cell signalling. SNAREs are important to the targeting and, hence, to different spatial distributions of K+ channels in nerve cells (Ma et al., 2001, 2002; Leung et al., 2003; Rivera et al., 2003; Misonou and Trimmer, 2004) and between apical and basal membranes of epithelia (Bravo-Zehnder et al., 2000; Le Maout et al., 2001). However, an impact on channel mobility and anchoring within the plasma membrane is entirely new. It is not difficult to imagine that anchoring and clustering of the channel proteins are important for effective signal transmission and control of channel activities. Disrupting this organisation and, presumably, local associations

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with upstream signalling elements could have profound effects on this coupling and thereby interfere with ABA signalling, as was first observed for these SNAREs (Leyman et al., 1999). Clearly, it will be of special interest now to identify the immediate protein partners of these SNAREs and to explore the functional impact of channel clustering at the plant plasma membrane.

6.5

Conclusions and outlook

Ion transporters mediate transmembrane solute fluxes that underlie cell volume changes, nutrient acquisition and maintenance of water potential. Regulation of ion transporters is essential to adjust these parameters to plant development and to environmental challenges such as nutrient shortage, drought and salinity. The combination of electrophysiological, biochemical, molecular and genetic approaches has created a wealth of information on responses of ion transporters to environmental cues and the signalling pathways leading to these responses. The signalling pathways involve a number of ubiquitous signalling molecules such as Ca2+ , CNs, ROS and nitric oxide, well-known regulatory proteins such as kinases, phosphatases, calmodulins and 14-3-3 proteins, as well as hormones such as ABA, auxin, ethylene and cytokinin. Nevertheless, there are still substantial gaps in our understanding of the molecular mechanisms that link signalling pathways to transporter abundance and activity. Furthermore, primary receptors perceiving the environmental stimuli still remain to be identified. Active research in the area of ion transport regulation will continue to fill these gaps, and it can be expected that this effort will be supported by continuous improvements of experimental techniques. In particular, phosphoproteomics of membrane proteins (N¨uhse et al., 2004) is likely to reveal much sought information on in vivo (de)phosphorylation events involved in the regulation of ion transporters under various environmental and nutritional conditions.

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7 Intracellular transport: solute transport in chloroplasts, mitochondria, peroxisomes and vacuoles, and between organelles Katrin Philippar and J¨urgen Soll

7.1

Introduction

Compartmentalisation of numerous metabolic pathways, biochemical reactions or substances is central and vital for every eukaryotic cell. Cell compartments are represented by several different, membrane-delimited organelles such as mitochondria and peroxisomes. In plant cells this assignment of tasks is even more pronounced than in the cells of animals, since plants also harbour chloroplasts and vacuoles. The consequence of cell compartmentalisation is an extensive intracellular network, involving the exchange of metabolites and solutes between the different organelles and the cytosol. In turn, solute transporters, mediating and regulating metabolic traffic across organellar membranes, are the bottleneck for the proper function of cellular metabolism. After a short introduction on strategies to identify organellar transporters in plants, this chapter will describe the integration of chloroplasts, mitochondria, peroxisomes and vacuoles into the cellular metabolic network. In following sections, known transport capacities and proteins are summarised and put into their respective physiological contexts. Nomenclature, structure, topology and functional characteristics of common membrane transport proteins are described elsewhere in this book (e.g. Chapter 6); therefore, this chapter will focus on the function of transport proteins that have been identified at a molecular level within specific organelles and their integration into the intracellular metabolic network.

7.1.1

Research to identify solute transport proteins in plant organelles

Beginning in the 1970s, intracellular solute transport in plants was studied by following metabolic fluxes into and out of the respective isolated, intact organelles. Transport capacity was first resolved either by uptake/release of radioactive labelled compounds or by simple swelling/shrinking assays of organelles. Later, isolation and purification of organellar membranes allowed a more detailed study of transport assigned to a certain membrane system. Electrophysiological techniques, like patch-clamp on isolated membranes or organelles as well as the study of membrane systems reconstituted in artificial lipid bilayers, were applied. This research lead to the description of ionic and metabolite fluxes with defined features such as conductance, gating, voltage dependence, and selectivity. The biggest task thereafter was to

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link the different currents and fluxes with the action of a given polypeptide. However, in many cases this is still a central question of modern research. In another approach, biochemical purification and isolation of organellar membrane proteomes identified proteins, which in turn were described as solute channels or carriers. Again, the reconstitution of the corresponding purified recombinant proteins in artificial lipid bilayers or the heterologous expression in appropriate systems (e.g. oocytes of the frog Xenopus laevis) provided in vitro transport function. In addition, current knowledge about the action of plant membrane transporters was gained using functional complementation strategies in yeast mutants, exhibiting a phenotype attributable to the absence of a carrier of known function. Immunological techniques (Western blotting and immunogold labelling) and fusion of the respective peptide to a fluorescent reporter protein (e.g. the ‘green fluorescent protein’ GFP) are commonly used to define the subcellular localisation of membrane transporters.

7.1.1.1

Benefits of a model plant: Arabidopsis thaliana

Because of particular attributes of their physiology and/or organelle isolation techniques, plant organellar transporters have often been isolated from quite diverse species – e.g. pea and spinach chloroplasts allow good separation of outer and inner envelope membranes. However, the model plant Arabidopsis thaliana (thale cress; arabidopsis) nowadays provides a fully sequenced and annotated genome as well as an enormous pool of mutant plants, therefore allowing the study of entire transporter families and their functions in planta. Thus, research focuses more and more on the characterisation of arabidopsis transporters, orthologous to the proteins originally isolated from other plants. In particular, the comparison of arabidopsis genes with transport proteins already identified in mammalian, yeast or prokaryotic model organisms helps to identify new plant membrane transporters. Transmembrane topology and localisation of proteins to subcellular organelles can be predicted in silico by analysis of hydrophobicity and certain targeting sequences. The plant membrane protein database ARAMEMNON (see Table 7.1) offers a comprehensive view of consensus transmembrane topologies, a collection of subcellular targeting predictions and a display of paralogues and orthologues in the arabidopsis, rice and cyanobacterial proteomes. Thus, bioinformatics represent a powerful tool to identify solute transport proteins in plant organelles (see also Section 4.6.1). In spite of the growing amount of information on sequences and functional characterisation of individual transporter proteins, little is known about their physiological roles in planta. This is a pivotal question in current research on plant membrane transport. To assign a biological function to a particular gene, reverse genetics represents a powerful tool. There is a constantly growing pool of arabidopsis mutant libraries in which genes have been randomly knocked out. Furthermore, cloned cDNA sequences as well as expression data (e.g. Affymetrix GeneChip) for nearly all arabidopsis genes are available (see Table 7.1 for the most common public databases). In contrast to functional genomics, proteomics provide a direct way to show the presence of proteins in certain organelles or tissues. Recent studies on the proteome of organelles and organellar membranes have greatly improved the allocation of known membrane transporter families as well as the identification

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Public databases for plant research

The most common public databases for research on the model plant Arabidopsis are listed Databasesa ARAMEMNON: http://aramemnon.botanik.uni-koeln.de – protein, cDNA and genomic sequences, topology consensus and subcellular targeting predictions, paralogous and orthologous genes SIGnAL (SALK Institute Genomics Analysis Laboratory): http://signal.salk.edu – summary and overview of current mutants and cDNA libraries, links to seed stocks and cDNA clones NASC (Nottingham Arabidopsis Stock Centre): http://arabidopsis.info – collection of mutant libraries, seed stocks, transcriptomics, genomics, proteomics TAIR (The Arabidopsis Information Resource): http://arabidopsis.org – maintains a database of all genetic and molecular biology data for Arabidopsis MATB (MIPS Arabidopsis thaliana database): http://mips.gsf.de/proj/thal/db/index.html – contains all Arabidopsis sequences and annotation produced by the Arabidopsis Genome Initiative (AGI), plus mitochondrial and chloroplast genomes NASCArrays (Microarray database NASC): http://affymetrix.arabidopsis.info/narrays/ experimentbrowse.pl – gene expression (data from the Affymetrix AG and ATH1 GeneChip arrays) Genevestigator: https://www.genevestigator.ethz.ch – gene expression (data from the Affymetrix AG and ATH1 GeneChip arrays) Summary of databases and analyses of the organellar proteome from plastids, mitochondria, peroxisomes and vacuoles in plants Proteomesb Plastids – Plprot: http://www.plprot.ethz.ch (Kleffmann et al., 2006) Plprot provides a useful overview and links to all published plastid proteome analyses and databases, including different plastid types (chloroplast, etioplast and proplastid) and plant species (Arabidopsis, rice, and tobacco BY-2). Mitochondria – http://www.plantenergy.uwa.edu.au/applications/ampdb/index.html (Heazlewood and Millar, 2005; Millar et al., 2005) Millar and coworkers present data from analyses of the Arabidopsis mitochondrial proteome Peroxisomes – Araperox: http://www.araperox.uni-goettingen.de (Reumann et al., 2004) The peroxisomal proteome in Araperox is based on computational prediction of targeting signals only Vacuoles – proteome data are given as supplements of the respective publications Tonoplast proteins from Arabidopsis have been isolated and published by different groups: Carter et al. (2004); Sazuka et al. (2004); Shimaoka et al. (2004); Szponarski et al. (2004). Endler et al. (2006) studied the tonoplast proteome on barley mesophyll vacuoles a

The May, July and August 2005 issue of the journal Plant Physiology focused on Biological Databases for Plant Research, giving a detailed overview of recent research and results. The April 2006 issue of the Journal of Experimental Botany is dedicated to Plant Proteomics, providing current knowledge about efforts on the proteomic field. b

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of new proteins and functions. Databases and references for organellar proteomes are listed in Table 7.1. Although several solute transport proteins have been identified in plant organelles, there are still many transport functions to be linked to specific proteins, many questions to be answered as well as many unidentified or yet unexpected transport capacities awaiting discovery and characterisation. As an example, the photorespiratory cycle (see Section 7.5), which involves the metabolite exchange between chloroplasts, peroxisomes and mitochondria, has been unravelled biochemically, while the molecular identity of the transporters involved remains enigmatic. Knowledge of the integration of known transport proteins into metabolic pathways mostly remains patchy. Thus, research on solute transporters in plant organelles is a constantly growing field, which has to focus on explanations of certain metabolic pathways and the in planta function of transport proteins. In the near future, improvement and integration of all the techniques mentioned above will provide a deeper insight into intracellular solute transport in plants.

7.2

Chloroplasts

Chloroplasts, which originated from free-living cyanobacteria, are the site of photosynthesis and thus build up the basis for all life dependent on atmospheric oxygen and carbohydrate supply. Chloroplasts are kidney-shaped and self-replicating organelles enclosed by outer and inner envelope membranes. Both envelope membranes are distinguishable by their structure, function and biochemical properties, but also cooperate in the synthesis of lipids or in protein translocation (Douce and Joyard, 1990; Joyard et al., 1998). Enclosed between the outer and inner envelope is the intermembrane space, while the stroma represents the chloroplast matrix. Inside the stroma, chloroplasts harbour a third membrane system, the thylakoids, which represent the site of light-harvesting complexes and the photosynthetic electron transport chain. Chloroplasts, around 5 μm in size, are unique and essential to plants and green algae. However, chloroplasts represent only one type of the plastid organelle family in higher plants. Proplastids in meristematic tissue and etioplasts in darkgrown plantlets develop during biogenesis into the mature, autotrophic chloroplast of the green leaf. In contrast, storage plastids are heterotrophic organelles that convert photosynthates derived from source tissue into storage compounds, which can be mobilised during plant development (Bowsher and Tobin, 2001; Lopez-Juez and Pyke, 2005). Amyloplasts (starch) in the endosperm of seeds, in cotyledons, tubers or fruits, elaioplasts (oil) in seeds of oilseed plants and chromoplasts (carotenoids) in flowers and fruits represent the major storage-type plastids. Root plastids, with major functions in the oxidative pentose phosphate pathway and nitrogen assimilation, also are members of the so-called non-green plastids (for a review on plastid differentiation see Waters and Pyke, 2005). The various plastid types are dynamically interconvertable; hence, the name plastid originates from the Greek word ‘plastikos’, meaning ‘plastic or shapeable’.

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7.2.1

137

The function of plastids

The plastid organelle family conducts vital biosynthetic functions in every plant cell (for an overview see Tetlow et al., 2005; Weber et al., 2005). First and foremost, chloroplasts carry out photosynthesis and release O 2 , thereby providing the basis for most of the life on earth. Via the reductive pentose phosphate cycle (Calvin–Benson cycle) in the stroma, atmospheric CO 2 is converted into carbohydrates that are directly used for starch and sucrose biosynthesis inside the chloroplast and cytosol, respectively. Furthermore, glyceraldehyde-3-phosphate can enter the chloroplastidic isoprenoid biosynthesis, leading to the production of isoprene, prenylquinones, carotenoids, chlorophyll side chains and several hormones. The Calvin cycle shares intermediates with the oxidative pentose phosphate pathway, leading to synthesis of nucleic acids as well as aromatic amino acids, polyphenols and lignins (shikimate pathway). Photorespiration, which involves transport of metabolites between the organelles (chloroplasts, mitochondria and peroxisomes), is described in Section 7.5. Nitrite produced in the cytosol during nitrogen assimilation is transported into the plastid and reduced to ammonia by plastidic nitrite reductase. All subsequent reactions of primary assimilation and amino acid synthesis take place within plastids of photosynthetic and non-photosynthetic tissue. In plants, de novo synthesis of fatty acids is carried out by a multisubunit synthetase complex in plastids. Furthermore, plastids are the site for sulphate reduction, and evidence for a plastidic glycolysis is accumulating. Although mitochondria in plants harbour an Fe–S cluster biogenesis system involving the ATP-binding cassette (ABC) transporter STA1/ATM3 (see Section 7.3.4.1), Fe–S cluster biogenesis also occurs in plastids (Balk and Lobreaux, 2005). In chloroplasts, Fe–S clusters for example are required for a functional cytochrome b 6 -f complex, ferredoxin and photosystem I, ensuring electron flow in the thylakoids. These manifold biosynthetic functions of plastids require the existence of different selective transport mechanisms across the envelope membranes to provide the cell with carbohydrates, organic nitrogen and sulphur compounds. On the other hand plastids take up inorganic cations (K+ , Na+ , Mg2+ , Ca2+ , Fe2+ , Cu2+ , Mn2+ and Zn2+ ), anions (NO 2 − , SO 4 2− and PO 4 3− ), and a variety of organic biosynthetic pathway intermediates such as phosphoenolpyruvate, dicarboxylic acids, acetate, amino acids, and ATP to fulfill their biosynthetic tasks. A comprehensive overview of the established and predicted solute transporters in the plastid envelope is given by Weber et al. (2005). In the following we will summarise and discuss the transport capacities that have been experimentally assigned to defined proteins.

7.2.2

Transport across the outer envelope: general diffusion or regulated channels?

Flux measurements and electrophysiological characterisation of plastid envelope membranes led to the postulation that transport across the outer envelope is a diffusion process through a general porin. But, as in peroxisomes (Section 7.4.2.1), this

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porin has not been identified at the molecular level. In contrast, several proteins with transport capacity can be assigned to the outer envelope (outer envelope proteins [OEPs; see below]). However, while the inner envelope carriers (Section 7.2.3) show a distinct substrate specificity, to what extent transport through channels of the outer membrane is regulated in vivo remains a matter of debate.

7.2.2.1

A porin in the outer envelope of plastids?

The solute permeability of isolated chloroplasts was first examined by electron microscopy and distribution of radioactive labelled compounds (Heldt and Sauer, 1971). These early studies were followed by electrophysiological approaches after reconstitution of outer envelope membranes into artificial lipid bilayers (Fl¨ugge and Benz, 1984) and by applying the patch-clamp technique on isolated, intact organelles (Pottosin, 1992, 1993). In summary, the results obtained suggested that the plastidic outer envelope is equipped with a general diffusion pore, related to the voltagedependent anion channel (VDAC), a porin from mitochondria (see Section 7.3.2). Therefore, it has long been assumed that the outer envelope of plastids is freely permeable for small-molecular-weight solutes up to 10 kDa. Correspondingly, it was believed that the osmotic barrier against the cytosol is formed exclusively by the inner envelope. Subsequently, Fischer et al. (1994) localised the mitochondrial VDAC to non-green plastids of pea roots but not to chloroplasts of green leaves. However, recent studies on the subcellular distribution of the VDAC protein between plastids and mitochondria in green and non-green tissue show that VDAC is present in mitochondria only (Clausen et al., 2004; cf. Section 7.3.2). Thus, the molecular identity of a VDAC-like porin in the outer envelope of chloroplasts is equivocal.

7.2.2.2

OEPs, a family of channels with substrate specificity

Transport across the outer envelope is mediated by a set of channel proteins, which are unique to plastids, have distinct substrate specificities and are called outer envelope proteins (see Figure 7.1). OEP16, OEP21, OEP24 and OEP37 – classified according to their molecular weight – were all isolated as protein bands from fractionated outer envelope membranes from pea chloroplasts. Thus, they represent abundant proteins in this membrane system. In addition to the preprotein-conducting channel Toc75 (Hinnah et al., 1997), these OEPs have been functionally characterised in vitro by electrophysiological measurements in artificial lipid bilayers. They all represent high-conductance solute channels with the highest open probability at 0 mV. Their distinct substrate specificities indicate separate roles in different metabolic processes, challenging the notion that they are general diffusion pores. OEP16: a channel for amino acids and amines. In artificial lipid bilayer membranes, OEP16 forms a cation-selective, high-conductance channel (1.2 nS in 1 M KCl) with a remarkable permeability to amino acids and amines (Pohlmeyer et al., 1997; Steinkamp et al., 2000). The OEP16 pore is impermeable to 3phosphoglycerate (3-PGA) or to sugars (glucose, fructose, sucrose and sorbitol) although the predicted pore size (about 1-nm width) is large enough to allow transport of these solutes. OEP16 shows sequence similarities to components of the protein translocase of the inner mitochondrial membrane, and to a lesser extent to LivH,

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INTRACELLULAR SOLUTE TRANSPORT Pollen development

Seed germination Embryo development

Solutes

Export of photoassimilates: TP TP, 3-PGA, Pi

Fatty acids, Amino acids?

OEP37 OE P1 6

OEP24

P OE 21

Cold acclimation Amino acids, amines

ATP TP

THY

Cl-

CLC

Stroma OE

IMS ATP synthesis

IE

Shikimate pathway: aromatic compounds

Pi Xyl-5-P

PEP

Export of photoassimilates: TP

Pi

GPT

1

PPT1

Gluc-6-P

Pi

T XP

TP T

Pi TP

Pi Xyl-5-P Exchange of intermediates Pentose phosphate pathway

Male + female gametophytes: import of carbon skeletons

Figure 7.1 Solute transport across the chloroplast envelopes: OEPs and phosphate translocators. The chloroplast of higher plants posseses three membrane systems: outer envelope (OE), inner envelope (IE), and the thylakoids (THY), defining three compartments: intermembrane space (IMS), stroma, and thylakoid lumen. Possible transport capacities and functions of the OEP proteins in the OE as well as the phosphate translocators in the IE are depicted. Upper half: In the outer envelope, OEP16 (four α-helices) constitutes a channel specific for amino acids and amines. A possible function for OEP16 might be the supply of plastid-derived amino acids during cold acclimation of plants. The β-barrel pore OEP21 is regulated by ATP and triose phosphates (TP) from the intermembrane space. High TP levels, generated by export from the stroma via TPT (see below), can thus induce a net efflux of TP, 3-phosphoglycerate (3-PGA) or inorganic phosphate (Pi). The rather unspecific channel OEP24 (12 β-sheets) has a defined function in solute transport during pollen development, maybe in concert with GPT1 in the inner envelope (see below). The metabolites transported by OEP37 still have to be identified; most likely, this β-barrel pore functions in transport of fatty acids and/or amino acids during seed germination and embryo development. The chloride channel CLC (12 α-helical domains), identified in outer envelopes, could mediate transport of Cl− ions. Lower half: The triose phosphate/phosphate translocator TPT in the inner envelope exports assimilated triose phosphates in exchange for phosphate, necessary for ATP synthesis. Function of the glucose-6-phosphate/phosphate translocator GPT1 is required for proper development of male and female gametophytes. In these non-photosynthetic tissues plastids have to import carbon skeletons in the form of glucose-6-phosphate (Gluc-6-P). The phosphoenolpyruvate/phosphate translocator PPT1 in C3 plants imports phosphoenolpyruvate (PEP) as precursor for synthesis of aromatic compounds in the stroma (shikimate pathway). Since XPT is able to transport Xyl-5-P in exchange for phosphate, it can shuttle pentose phosphate pathway intermediates between stroma and cytosol.

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an amino acid transporter in Escherichia coli (Rassow et al., 1999). Interestingly, OEP16 does not form a β-barrel pore, but is made of four α-helical transmembrane domains (Linke et al., 2004). This is very unusual, since porin-type channel proteins in outer membranes of Gram-negative bacteria – the evolutionary progenitors of plastids – in general are β-barrels. Thus, OEP16 somewhat resembles transporters or ion channels of the bacterial periplasmic membrane. Contradictionary to a function in amino acid transport, recent research proposes that OEP16 from barley acts as a precursor translocase for protochlorophyllide oxidoreductase (PorA) import into chloroplasts (Reinbothe et al., 2004, 2005). However, characterisation of mutants of all arabidopsis OEP16 isoforms does not support a function in PorA protein import (Philippar et al., in press). Furthermore, expression of OEP16 is induced by light and cold, while PorA transcripts are down-regulated upon illumination (Su et al., 2001). Studies on the cold induction of OEP16 in barley (Baldi et al., 1999) again suggest a function for OEP16 in amino acid transport during cold acclimation of cereals, because cold stress causes the accumulation of free, de novo synthesised amino acids, most likely produced in plastids. OEP21: a channel for phosphorylated carbohydrates, regulated from the intramembrane space. OEP21 has been shown to form a rectifying, anion-selective channel, which is regulated by ATP and triose phosphates (TPs) from the intramembrane space (B¨olter et al., 1999). The channel has a lower conductivity (0.95 nS in 1 M KCl) than OEP16 and is permeable to inorganic phosphate (Pi) and phosphorylated carbohydrates, central intermediates in the solute fluxes between chloroplasts and cytosol (e.g. TPs, 3-PGA, hexose phosphates). Only very recently, a refined analysis of the OEP21 channel properties revealed a β-barrel channel formed by eight β-strands with a wider pore vestibule (2.4 nm) at the intermembrane site and a narrower filter pore of ∼1 nm (Hemmler et al., 2006). In the open conformation, a higher conductance at one side of the channel corresponds to a current rectification in the direction from the intermembrane space to the cytosol. OEP21 contains two binding sites for ATP, one high-affinity site at the centre of the pore, and a second, low-affinity site at the vestibule. The latter is represented by a C-terminal, putative Fx4K motif, facing the intermembrane space. Binding of ATP to the inner site blocks the channel, while binding to both sites decreases anion selectivity. TPs can bind to both sites with the same affinity and thus compete with ATP. In consequence, increasing TP–ATP ratios at the intermembrane space releases the current block and increases anion selectivity, resulting in a net efflux of TPs (see Figure 7.1). Interestingly, one of the two arabidopsis OEP21 proteins, OEP21.2, is equipped with the Fx4K motif, while the second isoform, OEP21.1, is lacking this regulatory site. Thus, future experiments with the two OEP21 isoforms in the model plant should pinpoint the in planta regulation and function of OEP21. OEP24: a channel with specified function in a defined cell type. The channel properties of the slightly cation-selective OEP24 closely resemble those described for general diffusion pores (Nikaido, 2003). The 2.5-nm pore of the highconductance (2.1 nS in 1 M KCl) channel OEP24 allows the passage of TPs, ATP, inorganic pyrophosphate (PPi), dicarboxylate and positively or negatively charged amino acids (Pohlmeyer et al., 1998). Furthermore, OEP24 can functionally replace

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the mitochondrial VDAC (see Section 7.3.2) in VDAC-deficient yeast mutants (R¨ohl et al., 1999) and thus might represent a rather non-selective channel in the outer envelope. Secondary structure predictions suggest that OEP24 has a β-barrel-like structure containing 12 membrane-spanning β-strands (Schleiff et al., 2003). In arabidopsis, two isoforms of OEP24 are present. Interestingly, mutation of OEP24.1 leads to a defect in pollen germination (A. Timper, J. Soll and K. Philippar, unpublished). In early pollen development, fatty acid biosynthesis in plastids supplies the growing pollen grain with energy and lipid material, needed for pollen tube emergence and growth (Yamamoto et al., 2003). Thus, the rather unspecific pore OEP24 seems to have a very specific and defined function, triggered by expression in plastids during early pollen development (cf. GPT1; Section 7.2.3.1). To conclude, these findings prove the concept that metabolic fluxes across the outer envelope in vivo are mediated by OEP proteins, which are not redundant porins, but represent specific channels essential for plastidic function. OEP37: a new member of the chloroplast outer membrane channels. Only very recently, OEP37 was described as a new member of the outer envelope channels (Goetze et al., 2006). The reconstituted recombinant protein OEP37 from pea forms a cation-selective, rectifying high-conductance channel with a voltage-dependent open probability being maximal at 0 mV. The channel pore reveals an hourglassshaped form with different diameters for the vestibule (3.0 nm) and restriction zone (1.5 nm). OEP37 displayed affinity for the precursor of the chloroplast inner membrane protein Tic32 and for a synthetic peptide. Most likely, the channel pore of OEP37 is formed by 12 β-strands (Schleiff et al., 2003). In arabidopsis, transcripts of AtOEP37 are ubiquitously expressed throughout plant development and accumulate in early germinating seedlings as well as during late embryogenesis. The plastid intrinsic protein could be detected in isolated chloroplasts of cotyledons and rosette leaves. However, the knockout mutant oep37–1 shows that the proper function of this single-copy gene is not essential for development of a mature plant. In conclusion, OEP37 may constitute a novel peptide-sensitive channel in the outer envelope of plastids with function during embryogenesis and germination (Figure 7.1). Although, the pore characteristics of OEP37 are described in detail, substrate specificity and putative metabolite transport capacities await further characterisation. However, because of its expression in storage-type plastids of green cotyledons in seeds it is tempting to speculate that OEP37 might transport stored fatty acid compounds and/or amino acids. CLC: a chloride channel present in outer envelopes. A putative chloride channel CLC-f, which shares similarity with cyanobacterial CLC proteins, was localised to the outer membrane of spinach chloroplasts (Teardo et al., 2005; Figure 7.1). The protein with 12 α-helical transmembrane domains is expressed in etioplasts and chloroplasts but not in root plastids. In addition to the channel pores described above, distinct anion-selective channels have been recorded by the patch-clamp technique and by incorporating envelope vesicles into lipid bilayers (Pottosin, 1992; Heiber et al., 1995). Thus, the authors argue that CLC-f might be responsible for these currents. However, more experimental data have to be gathered, to link CLC-f function to a role in the outer envelope of chloroplasts.

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7.2.2.3

PLANT SOLUTE TRANSPORT

Outer membrane channels and porins: evolutionary aspects in chloroplasts and mitochondria

Gram-negative bacteria, representing the evolutionary progenitors of plastids as well as mitochondria, contain three different classes of porins in their outer membrane: (i) general porins, which do not bind solutes and act as size-selective diffusion pores; (ii) solute-selective specific porins, binding solutes with low affinity; and (iii) ligand-gated and energy-dependent porins with high affinity for solutes (Klebba and Newton, 1998; Nikaido, 2003). In conclusion, it seems likely that the outer membrane of chloroplasts and mitochondria are equipped differently, depending on the metabolic needs of the organelle. While mitochondria with VDAC kept a general diffusion pore that can be regulated by metabolites, substrates and nucleotides (see Section 7.3.2), the situation in plastids is more complex. They are equipped with low-affinity, highly specific channels (OEP16), the substrate- and ATP-gated OEP21 and the specifically expressed, but rather general, porin OEP24, reflecting the entire spectrum of bacterial porins (for reviews and discussion see Fl¨ugge, 2000; Soll et al., 2000; B¨olter and Soll, 2001). Although Gram-negative bacteria in general are considerd as the progenitors of both organelles, chloroplasts (from ancient cyanobacteria) and mitochondria (from proteobacterial ancestors) have most probably arisen from two independent endosymbiotic events. An unambiguous identification of a porin on the molecular level was possible only in mitochondrial outer membranes (VDAC; Section 7.3.2), while the existence of a porin in the outer envelope of chloroplasts as well as in peroxisomal membranes (Section 7.4.2) relies only on the characterisation of metabolite fluxes. Recent research in the field, however, suggests that an enormous number of yet uncharacterised membrane transporters exist in plant organelles. Thus, the view of a general diffusion pore may prove to be an oversimplification. In the following two sections we will review the solute transporters of the inner envelope membrane in chloroplasts. In general, these transporters are built of α-helical transmembrane domains. However, several proteins have been localised from a mixed preparation only (outer and inner envelope) or from the chloroplast envelopes in general (e.g. by fusion to GFP). Thus, it is possible that proteins assigned to the inner envelope in this review are integral to the outer envelope instead. With the α-helical CLC and OEP16 (see above), it is evident that membrane proteins in the outer envelope of chloroplasts are not exclusively of β-barrel topology. Future research is thus required to discriminate unequivocally between outer and inner envelope localisation of transporters.

7.2.3 7.2.3.1

Transport across the inner envelope: phosphate translocators, major facilitators and carriers The phosphate translocator family

Members of this family are phosphate antiporters that exchange various phosphorylated carbon compounds for Pi between plastids and the cytosol. The four phosphate translocators identified so far (see Figure 7.1) belong to a family of drug/metabolite transporters, which most likely function as homodimers. A membrane topology with eight α-helical domains is suggested for the entire phosphate translocator family in

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143

plants (Weber et al., 2005). For a more detailed overview of this family we refer to recent reviews by Knappe et al. (2003a), Weber (2004) and Weber et al. (2005). TPT, the triose phosphate/phosphate translocator. TPs are generated at the expense of photosynthetic energy and reducing equivalents (ATP, NADPH) by the Calvin–Benson cycle in the chloroplast stroma. Subsequently, TPT exports the fixed carbon in the form of TPs, which are then integrated into sucrose in the cytosol. In turn, released Pi in the cytosol is transported back into the chloroplast, where it is required for ATP synthesis (Figure 7.1). TP can also be used to produce transistory starch. TPT was first isolated and localised to the inner envelope in spinach chloroplasts (Fl¨ugge et al., 1989); subsequently, substrate specificity and channel characteristics were shown using purified recombinant proteins (Loddenk¨otter et al., 1993; Schwarz et al., 1994). Although TPT is a single-copy gene in arabidopsis, mutants defective in TPT function alone do not show a severe phenotype, but an alteration in carbon metabolism (Schneider et al., 2002). To compensate for the loss of TPT, photoassimilates are allocated to the cytosol in a different way, including starch breakdown and formation and export of free sugars (mostly maltose by MEX1, see below). Simultaneous knockout of both starch synthesis and TPT, however, prevented this compensation and lead to impaired growth and photosynthesis. GPT, the glucose-6-phosphate/phosphate translocator. The glucose-6phosphate/phosphate antiporter was isolated from heterotrophic pea root plastids (Kammerer et al., 1998). Since plastids of non-green tissue are not photosynthetically active, they have to fuel their biosynthetic pathways (e.g. fatty acid, amino acid and starch biosynthesis) by import of glucose-6-phosphate via GPT. In arabidopsis, two GPT proteins are present, which represent functional proteins when reconstituted into liposomes (transport of Pi, glucose-6-phosphate, TPs and phosphoenolpyruvate) as well as in planta (Niewiadomski et al., 2005). While mutation of GPT2 had no obvious impact on plant growth and development, loss of GPT1 revealed a pleiotropic effect on male and female gametophyte development. Mutant pollen development, for example, was associated with reduced lipid-body formation and disintegration of membranes, linking GPT1 function with plastidic fatty acid biosynthesis occurring in early pollen development (cf. OEP24; Section 7.2.2). PPT, the phosphoenolpyruvate/phosphate translocator. PPT was originally found and proven to function in phosphoenolpyruvate transport in maize endosperm, maize roots, cauliflower buds, tobacco leaves and arabidopsis leaves (Fischer et al., 1997). Plastids of C4 (e.g. maize) and crassulacean acid metabolism (CAM) plants have to import pyruvate generated by malic enzyme. In turn, pyruvate is converted into phosphoenolpyruvate, which then leaves the plastid via PPT to be metabolised in primary carbon fixation and biosynthesis of storage carbohydrates, respectively. In C3 plants like arabidopsis and non-photosynthetic tissue of C4 plants, PPT imports phosphoenolpyruvate into the plastid rather than export it (Fischer et al., 1997). In these cases, phosphoenolpyruvate is required as a precursor for the plastidlocalised shikimate pathway (Figure 7.1). The arabidopsis PPT1-mutant cue1 is lacking plastid-derived aromatic compounds such as anthocyanins, thus underlining the import role for PPT1 (Streatfield et al., 1999). However, this role was challenged by the isolation of a second PPT gene, PPT2, from arabidopsis and the finding that PPT1

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is expressed in the vasculature (leaves and roots) while PPT2 is found ubiquitously in leaves but not in roots (Knappe et al., 2003b). For discussion on the role of PPTs in plant development and metabolism, see Weber (2004) and Weber et al. (2005). XPT, a xylulose-5-phosphate/phosphate translocator. The most recent addition to the plastidic phosphate translocator family is XPT from arabidopsis, found to catalyse the exchange of xylulose-5-phosphate (Xyl-5-P) with Pi or TPs (Eicks et al., 2002). This transporter allows the exchange of pentose phosphate pathway intermediates between the cytosol and plastids, and thereby connects the pentose phosphate pathways in both compartments. In addition, it permits the generation of reducing equivalents from Xyl-5-P by the oxidative pentose phosphate pathway.

7.2.3.2

Major-facilitator-mediated transport

With exception of the maltose transporter MEX1, dicarboxylates, carbohydrates and adenylates are transported across the inner envelope by transporters of the major facilitator superfamily. The major facilitator superfamily (MFS) is also called the ‘uniporter–symporter–antiporter family’ (Pao et al., 1998). MFS transporters are single-polypeptide, secondary carriers with 12 α-helical transmembrane domains, capable of transporting solutes in response to chemiosmotic gradients. The MFS transporters characterised in the inner envelope generally function as antiporters (see Figure 7.2). Dicarboxylate transport in plastids. Transport of dicarboxylates such as malate, 2-oxoglutarate, glutamate and oxaloacetate is important during carbon and nitrogen assimilation. Ammonia, generated by nitrite reduction in the cytosol and/or photorespiration (see Section 7.5), is directly assimilated into glutamate by the action of the glutamine synthetase/glutamate synthase (GS–GOGAT) cycle. Therefore 2-oxoglutarate, the carbon skeleton for ammonia assimilation, must be imported from the cytosol and the product glutamate has to be exported. This transport is mediated by a malate-coupled two-transporter system (Figure 7.2) involving a 2-oxoglutarate/malate (DiT1/OMT1) and a glutamate/malate antiporter (DiT2/DCT1). In addition, a high-affinity oxaloacetate transporter as part of the malate/oxaloacetate shuttle, balancing the stromal ATP/NADPH ratio, is proposed (Taniguchi et al., 2002, and references therein). DiT1/OMT1, a 2-oxoglutarate/malate antiporter. The recombinant protein SoDiT1, originally derived from spinach chloroplasts (Menzlaff and Fl¨ugge, 1993), was shown to have the same substrate specificities as the authentic chloroplast protein (Weber et al., 1995). Taniguchi et al. (2002) identified the orthologue, AtOMT1, in arabidopsis. Here the recombinant OMT1 transported malate, 2-oxoglutarate and oxaloacetate, but not glutamate, when reconstituted into proteoliposomes. Gene expression was induced by light and by nitrate, and the authors suggest that AtOMT1 should be able to transport 2-oxoglutarate/malate as well as functioning as an oxaloacetate transporter. However, Renne et al. (2003) argue that under in vivo conditions DiT1/OMT1 should rather function as 2-oxoglutarate/malate translocator. DiT2/DCT1, a glutamate/malate transporter. In the early 1980s, it was shown that the classical photorespiratory mutant dct in arabidopsis, which needs high CO 2 to survive, is defective in chloroplast dicarboxylate transport (Somerville and

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INTRACELLULAR SOLUTE TRANSPORT Photorespiration

Glutamate

DiT1

DiT2

Malate

ADP

Malate 2-Oxoglutarate

G

Carbon + nitrogen assimilation

Starch

NT T

ATP

IE

Amino acids, fatty acids, starch

Biosynthesis + Breakdown

T lc

Glucose

MEX1

Maltose

Monocots: starch THF ADP-glucose

AMP

FOL1

BT1

Folate

Figure 7.2 Major facilitators and carriers in the inner envelope of plastids. The function of the major facilitator and carrier proteins in the inner envelope (IE) applies to the biosynthetical and catabolic capacity in the plastid stroma. According to Renne et al. (2003), the two major facilitator proteins DiT1 and DiT2 constitute a malate-coupled transport system for glutamate export and 2-oxoglutarate import. They function during carbon and nitrogen assimilation, e.g. in the photorespiratory cycle (cf. Figure 7.8). Starch breakdown in the stroma at night leads to the export of the products maltose via MEX1 and glucose via GlcT. While MEX1 is a transmembrane protein with nine α-helical domains, GlcT belongs to the major facilitators. The biosynthetic capacity of plastids requires the import of energy in the form of ATP, which is exchanged for ADP across the inner envelope by the nucleotide transporter NTT (major facilitator). In monocotyledonous plants, starch synthesis in storage plastids is dependent on import of ADP-glucose by the carrier Brittle1 (BT1). Another carrier with six α-helical domains in the inner envelope is FOL1, involved in folate import during biosynthesis of tetrahydrofolate (THF).

Ogren, 1983). The corresponding transporter protein in arabidopsis is DiT2/DCT1 (Taniguchi et al., 2002; Renne et al., 2003). The recombinant protein transported glutamate, aspartate and to a lesser extent 2-oxoglutarate in exchange for malate, and the cDNA was able to complement the mutant phenotype of dct. In conclusion it is suggested that DiT2/DCT1 exports malate in exchange for glutamate across the inner envelope of chloroplasts. Carbohydrate transport across the inner envelope. At night, transistory starch in the chloroplast stroma is predominantly broken down to glucose and maltose, which in turn represent the major carbon export from chloroplasts in the dark (for review, see Smith et al., 2005). In isolated chloroplasts fluxes of glucose and maltose could be demonstrated (Sch¨afer et al., 1977; Herold et al., 1981; Rost et al., 1996). MEX1, the maltose transporter. A mutant that is defective in maltose export from plastids (maltose excess1, mex1) provided the material for the isolation of

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MEX1, a transmembrane protein with nine α-helical domains (Figure 7.2; Niittyl¨a et al., 2004). Mutant (mex1) plants are small and pale green and have leaf maltose concentrations about 40 times higher than in wild-type plants. MEX1 is a novel maltose transporter that is unrelated to other sugar transporters of the major facilitator family. GlcT, a putative glucose transporter. Reexamination of the kinetics and labelling of glucose uptake into spinach chloroplasts lead to the identification of a protein, which represents a putative glucose transporter in the inner envelope of chloroplasts (Weber et al., 2000). GlcT belongs to the major facilitators and is closely related to the mammalian glucose transporter family. In conclusion, a possible function during starch breakdown and glucose export from chloroplasts at night is suggested. However, expression of GlcT also occurs in tissues that do not contain starch (Butowt et al., 2003), pointing to a possible function in carbon import as well (see Weber, 2004, for discussion). NTT, the ATP/ADP transporter in plastids. Amino acid, fatty acid and starch biosynthesis, as well as protein import, in plastids require energy in the form of ATP. The plastidic nucleotide transporter NTT, which catalyses the uptake of ATP in exchange for organellar ADP (Figure 7.2), was first identified in arabidopsis and functionally characterised in E. coli cells (Neuhaus et al., 1997; M¨ohlmann et al., 1998; Tjaden et al., 1998b). In planta function during starch biosynthesis was demonstrated in transgenic potato tubers that exhibited increased or decreased amounts of the NTT protein (Tjaden et al., 1998a). By analysing expression pattern and mutants of the two NTT proteins (major facilitator family) present in arabidopsis, Reiser et al. (2004) found that plastidic ATP/ADP transport activity is not required to pass through the complete plant life cycle. However, NTT was necessary for an undisturbed development of young tissues as well as a controlled cellular metabolism in mature leaves.

7.2.3.3

Carriers in the inner envelope of plastids

Two proteins belonging to the mitochondrial carrier family (MCF) were identified in the inner envelope of plastids. MCF proteins, first characterised in the inner membrane of mitochondria (see Section 7.3.3) are made of six α-helices and act as dimers. BT1, an ADP-glucose transporter. In storage plastids, starch synthesis is dependent on the uptake of cytosolic precursors. In amyloplasts of dicotyledons, carbon compounds are imported as glucose-6-phosphate via GPT (Section 7.2.3.1). The necessary energy for conversion into ADP-glucose and subsequently into starch is provided by the ATP/ADP antiporter NTT (see above). Plastids in cereal endosperm, however, have to import ADP-glucose, since they cannot convert glucose1-phosphate and ATP into ADP-glucose inside the stroma (M¨ohlmann et al., 1997). The corresponding transporter for ADP-glucose was first isolated from the inner membrane of maize endosperm plastids in the low-starch mutant brittle1 (bt1; Shannon et al., 1998), and an orthologous transporter is also found to be defective in a low-starch mutant of barley (Patron et al., 2004). BT1, which most likely exchanges ADP-glucose with AMP, is an MCF protein. Interestingly BT1 orthologues

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147

were recently discovered in dicotyledonous plants such as potato and arabidopsis (Leroch et al., 2005). The protein StBT1 from potato was shown to catalyse adenine nucleotide uniport with similar affinities for AMP, ADP and ATP. In contrast to ADP-glucose import in monocots, StBT1 is suggested to provide the cytosol with adenine nucleotides that have been synthesised in plastids. FOL1, a plastid-localised folate carrier. Biosynthesis of tetrahydrofolate (THF) in plants involves a complex intracellular traffic of THF and its precursors between cytosol, mitochondria and plastids (see Bedhomme et al., 2005, and references therein). However, the recently identified arabidopsis protein AtFOL1, which is similar to mammalian mitochondrial folate transporters (Bedhomme et al., 2005), was the first plant protein discovered that is able to transport folate. FOL1 is targeted to the chloroplast envelope and can complement folate uptake to a heterologous cell line. Since knockout mutants of AtFOL1 are not impaired in folate uptake into chloroplasts, the presence of a second plastidic folate transporter is likely.

7.2.4 7.2.4.1

Transport across the inner envelope: ABC transporters and ion transport ABC transporters

A total number of 19 ABC transporters represent the largest group of transporters, predicted to be present in chloroplast envelopes (Weber et al., 2005). Moreover, vacuolar ABC transporters (Section 7.6.3.2) have been shown to transport chlorophyll catabolites (Lu et al., 1998). However, none of the classical eukaryotic full or half ABC transporters could be assigned with a function in plastids. Nonintrinsic ABC proteins (NAPs) contain the ABC, but lack the membranespanning domains of a classical eukaryotic ABC transporter (for a description of ABC transporter structure see Chapter 5.3.3). They resemble prokaryotic ABC transport systems, in which a soluble ATP-binding domain interacts with the transmembrane permease subunit to generate a functional ABC transporter (for overview of plant and prokaryotic ABC transporters see Higgins, 2001; Sanchez-Fernandez et al., 2001; Garcia et al., 2004). The plastid-localised protein ABC1/NAP1 was isolated from a mutant impaired in phytochrome-A-mediated response to far red light (laf6, ‘long after FR’; M¨oller et al., 2001). ABC1, which is most similar to proteins from cyanobacteria, belongs to the subfamily of small soluble, non intrinsic ABC proteins. Since ABC1 localises to the periphery of chloroplasts it might interact with a transmembrane permease subunit to form a functional ABC transporter. Loss of ABC1 in chloroplasts leads to deficiency in chlorophyll and accumulation of the chlorophyll precursor protoporphyrin IX in the cytosol. Thus, the authors conclude that ABC1 is involved in light signalling as well as in transport of protoporphyrin IX across the inner envelope (Figure 7.3). NAP7 is another plastid-localised ABC/ATPase, involved in the biogenesis of plastidic Fe–S clusters (Xu and M¨oller, 2004). No evidence, however, that NAP7 is involved in transport, exists.

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PLANT SOLUTE TRANSPORT

NO2SO42-

Lipid biosynthesis

Proto IX

NO2-

AB C1

Lipids

Nitrogen, sulphate reduction

Proto IX

SO42PO43-

Chlorophyll

IE

H+

Ca2+ signalling: formation of root nodules or mycorrhizal symbiosis

Cl-, Pi, Glutamate

ANTR2

PO L

K+ ?

CA S

pH control

PHT 2;1

H+

Na+/K+ 3 X2 CH

IMS

?

?

AB C1

?

OE

?

TGD1 TGD1

Far-red light

H+

Figure 7.3 ABC transporters and ion transport in the inner envelope of plastids. The soluble ATPase subunit ABC1 is proposed to form a functional ABC transporter with a yet unidentified permease subunit in the inner envelope (IE). In turn, ABC1 by ATP hydrolysis fuels the import of the chlorophyll precursor protoporphyrin IX (proto IX) from the intermembrane space (IMS). In addition to chlorophyll biosynthesis, proto IX might be involved in far-red light signalling. The permease TGD1 most likely transports lipids across the inner envelope. Thereby TGD1 is involved in the biosynthesis of thylakoid lipids, which occurs at the IE and OE membrane systems. For a functional ABC transporter two halfmolecules of TGD1 are necessary, and the interacting ATPase subunits are unknown. For nitrogen assimilation and sulphate reduction, chloroplasts have to import nitrite and sulphate from the cytosol. However, pathways and involved proteins are unidentified. In addition to the phosphate translocators (Figure 7.1), the major facilitator PHT2;1 allows the accumulation of phosphate in the stroma. By exchanging protons for sodium and/or potassium, the major facilitator CHX23 regulates stromal pH and drives proton-coupled uptake mechanisms. The major facilitator ANTR2 is capable to mediate the required uptake of Cl− ions, and thus can contribute to pH control as well. CASTOR (CAS) and POLLUX (POL) have four α-helical transmembrane domains and most likely transport K+ , which in turn would be required for Ca2+ signalling and endosymbiotic events in the root.

In contrast, the arabidopsis protein TGD1 (Xu et al., 2003) shows similarities to ABC-domain-lacking, membrane-intrinsic permease half-molecules of bacterial ABC transporter complexes (Higgins, 2001). TGD1 was identified in a highthroughput screen for mutants with altered lipid metabolism. Recently, TGD1, which appears to be a component of a lipid transporter (Figure 7.3), was properly localised to the inner envelope membrane (Xu et al., 2005). Mutants of TGD1 show embryo abortion and accumulate triacylglycerols, oligogalactolipids and phosphatidate, whereas chloroplast lipids are altered in their fatty acid composition. Thus, transport by TGD1 is involved in galactolipid biosynthesis, which is linked to the chloroplasts envelope.

INTRACELLULAR SOLUTE TRANSPORT

7.2.4.2

149

Ion transport

Sulphate, nitrite, phosphate. Sulphate and nitrite reduction occur in plastids; therefore, the SO 4 2− and NO 2 − ions have to pass the envelope membranes (Figure 7.3). Although action of the phosphate translocators (Section 7.2.3.1) already provides a transport system for phosphate, PO 4 3− is transported in an alternative way. Douce and Joyard (1990) propose a carrier system, based on exchange of SO 4 2− with Pi for sulphate uptake into chloroplasts. In the model unicellular green alga, Chlamydomonas reinhardtii, a putative, envelope-localised ABC-type transporter functions in sulphate uptake (reviewed in Melis and Chen, 2005). However, homologues of these genes have not been retained in vascular plants, so the pathway for sulphate import into chloroplasts of plants still remains enigmatic. Although nitrite uptake into chloroplasts is crucial for ammonia assimilation (Section 7.2.1), the transport process across the envelope is not well understood and is discussed as a passive diffusion versus regulated transport (see Douce and Joyard, 1990; Galvan et al., 2002, and references therein). Identification of the NAR1 (nitrate assimilation related) family in Chlamydomonas reinhardtii, containing transmembrane proteins with six α-helical domains, similar to bacterial formate/nitrite transporters, favours a controlled transport process (Galvan et al., 2002). However, since orthologous proteins in higher plants could not be identified unequivocally, the question whether nitrite uptake is by diffusion or via a carrier remains open (Figure 7.3). Characterisation of the phosphate transporter PHT2;1 of arabidopsis, which shares similarity to H+ /Pi symporters of bacterial origin, showed that a Pi transport alternative to the phosphate translocators exists (Versaw and Harrison, 2002). PHT2;1, predicted to contain 12 α-helical transmembrane domains, localises to chloroplasts and mediates phosphate uptake when expressed heterologously in yeast cells. Analysis of a null mutant reveals that PHT2;1 activity affects Pi allocation within the plant and modulates Pi starvation responses. Since it is most likely that PHT2;1 is an H+ /Pi symporter, the pH difference that is maintained across the inner envelope membrane could be used to energise Pi import into the stroma (Figure 7.3). It is suggested that in addition to the phosphate translocators, which exchange metabolites with Pi in a 1:1 stoichiometry (Section 7.2.3.1; Figure 7.1), the chloroplast possesses with PHT2;1 an alternative mechanism for Pi import allowing the concentration of Pi in the stroma. Potassium, chloride and protons modulate pH. During photosynthesis, lightdriven H+ gradients are generated between the cytosol (pH ≈ 7.0), the chloroplast stroma (pH ≈ 8.0) and the thylakoid lumen (pH ≈ 5.0). In the stroma, concentrations of the physiologically important ions are 150 mM K+ , 50 mM Cl− and 5 mM Mg2+ . The steady-state membrane potential across the inner envelope is in the order of –100 mV (negative in the stroma) and across the thylakoid membrane about 10 mV (positive in the lumen). Ion channels in the inner envelope and the thylakoid membrane appear to be involved in generation and regulation of these proton gradients and membrane potentials (Heiber et al., 1995). Because of the pH optimum, of around 8.0, of the key enzymes of the Calvin–Benson cycle, the photosynthetic capacity of chloroplasts is regulated by stromal pH. Furthermore, the

150

PLANT SOLUTE TRANSPORT

presence of Cl− ions is required for proper function of the oxygen-evolving complex (Ferreira et al., 2004). Flux studies and electrophysiological analyses lead to a model, integrating proton and potassium transport across the inner envelope (Heiber et al., 1995, and references therein). The stromal pH is regulated by K+ /H+ exchange across the chloroplast envelope, involving the action of a H+ -ATPase (see Douce and Joyard, 1990) and a K+ channel (Berkowitz and Peters, 1993; Wang et al., 1993; Mi et al., 1994; Heiber et al., 1995; Mi and Berkowitz, 1995). A low conductance chloride channel has also been identified in envelopes. Thus, it has been proposed that potassium and chloride channels together with the H+ -ATPase are important for the regulation of stromal pH. Unfortunately, up to now, the molecular identities of these proteins remain unknown. However, a putative Na+ (K+ )/H+ exchange protein, CHX23, has been localised to the plastid envelope in arabidopsis (Song et al., 2004; Figure 7.3). Leaves of chx23 mutants displayed a high sensitivity to NaCl and a higher cytosolic pH than did wildtype leaves. Furthermore, thylakoid biogenesis was impaired in mutant chloroplasts. CHX23 has 12 predicted α-helical transmembrane domains and belongs to the family of plant Na+ (K+ )/H+ antiporters (major facilitator). Most likely, CHX23 is an ion antiporter that functions in an appropriate adjustment of pH in the chloroplast stroma and the cytosol and is necessary for chloroplast biogenesis. The protein ANTR2 (for anion transporter) was shown to be localised to the plastid inner envelope of arabidopsis and spinach (Roth et al., 2004; Figure 7.3). The major facilitator ANTR2 contains 12 putative transmembrane domains and belongs to the animal NaPi-1 family of proteins, which are involved in the transport of Pi, chloride and glutamate. Thus, it is concluded that ANTR2 can mediate transport of phosphate, chloride and glutamate across the inner envelope. Only very recently CASTOR and POLLUX, two proteins from Lotus japonicus, which localise to root plastids, have been shown to be crucial for the development of endosymbiotic fungal and bacterial relationships with root cells (Imaizumi-Anraku et al., 2005). Mutant plants in CASTOR or POLLUX genes are not able to form root nodules or arbuscular mycorrhizal symbiosis and lack the Nod-factor-induced calcium spiking in these cells. Since both proteins show significant structural similarity to calcium-gated potassium channels, the authors speculate that CASTOR and POLLUX mediate ion fluxes between plastids and cytosol, which are a prerequisite for calcium spiking and hence signal transduction leading to endosymbiosis.

7.2.4.3

Transport of metal ions

Calcium. One of the most important signals that relates to a plant’s response to its environment is an increase of cytosolic Ca2+ elicited by light/dark stimuli. Isolated chloroplasts take up Ca2+ upon illumination, a process that probably is mediated by Ca2+ transport across the inner envelope membrane (Sai and Johnson, 2002, and references therein). Thus, Ca2+ fluxes across the chloroplast envelopes contribute to Ca2+ signalling by regulating cytosolic Ca2+ levels and controlling processes in the chloroplast. In the stroma, Ca2+ has an impact on enzymes for photosynthetic CO 2 fixation, while in the thylakoid lumen Ca2+ is required for the function of PSII.

INTRACELLULAR SOLUTE TRANSPORT

151

1 PPF

Ca2+ ?

MRS2-1 1

Light-dependent Mg2+ uptake

Cytoplasmic Ca2+ Ca2+ signalling

Mg2+ Chlorophyll

IMS

Photosynthetic electron transport

IE

Cu2+ SOD: detoxification H+

PAA2

OE

Fe2+, Zn2+, Cu2+, Mn2+ A1 HM

PAA1

Me tal

Cu2+

ADP ATP

ADP

ATP

Figure 7.4 Transport of metal ions in chloroplasts. PPF1 might be a candidate protein for Ca2+ transport across the chloroplast envelope. In turn, PPF1 would contribute to the regulation of cytoplasmic Ca2+ and corresponding signalling events. Localisation to the inner envelope and Ca2+ transport capacity of PPF1, however, are not clarified yet. MRS2–11 (two to three α-helical domains) can mediate lightdependent uptake of Mg2+ into chloroplasts. Magnesium in turn is incorporated into chlorophyll. Uptake of the transition metals iron, zinc, copper and manganese most likely is coupled to proton symport and mediated by a yet unknown protein in the inner envelope. The heavy-metal ATPases PAA1 and HMA1 transport Cu2+ across the inner envelope, while PAA2 mediates copper uptake into the thylakoid lumen. Transition metals in the chloroplast stroma function as cofactors for superoxide dismutases (SOD) during detoxifcation of oxygen radicals. Import of Fe, Cu and Mn is crucial for photosynthetic electron transport.

Using the Ca2+ -selective photoprotein aequorin, targeted to chloroplasts in transgenic arabidopsis, Johnson et al. (1995) were able to monitor the increase in stromal intrinsic Ca2+ following light-to-dark transition. However, the corresponding Ca2+ transport protein has not yet been identified. Interestingly, the pea protein, PPF1, orthologous to ALB3 in arabidopsis, represents a putative calcium ion carrier (Wang et al., 2003; Figure 7.4). Overexpression of PPF1 delays flowering and increases chloroplastic Ca2+ levels, while reduction of PPF1 leads to decreased Ca2+ in chloroplasts. Further, it is suggested that PPF1 controls programmed cell death (PCD) in apical meristems of flowering plants by regulating cytoplasmic calcium content (Li et al., 2004). However, ALB3 in arabidopsis is localised to thylakoid membranes and required for the membrane insertion of members of the light-harvesting chlorophyll-binding protein (LHCP) family (Sundberg et al., 1997; Woolhead et al., 2001; Spence et al., 2004), questioning its function as a Ca2+ channel.

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PLANT SOLUTE TRANSPORT

Magnesium. In chloroplasts, stromal magnesium, which increases upon illumination, is involved in the regulation of key enzymes of carbon fixation and photosynthesis and represents the central cation of the chlorophyll molecule (Berkowitz and Wu, 1993; Shaul, 2002; Ishijima et al., 2003). AtMRS2–11, a protein that belongs to the MRS2 subfamily in the CorA superfamily of magnesium transporters (Knoop et al., 2005), localises to the chloroplast envelope (Drummond et al., 2006). Furthermore, MRS2–11, which contains two to three, C-terminal transmembrane domains, confers magnesium uptake to a yeast mutant and is regulated by light in a diurnal manner. Thus the evidence is that MRS2–11 is involved in light-dependent magnesium uptake into chloroplasts (Figure 7.4). Transport of transition metals. Because of their redox potentials, the transition metals Mn, Fe and Cu play a vital role in photosynthetic electron transport (Raven et al., 1999). While the photosynthetic apparatus represents one of the most ironenriched systems (PSII, PSI, cytochrome b 6 -f complex and ferredoxin) in plants, copper ions catalyse electron transfer via plastocyanin and a cluster of Mn atoms is required as the catalytic centre in the oxygen-evolving complex. Furthermore, stromal-localised Fe- or Cu/Zn superoxide dismutases scavenge reactive oxygen species. In addition, Zn is known to function as cofactor (RNA polymerase, zincfinger domains) in plastid transcription. Despite these essential functions for metal ions in chloroplasts, very little is known about metal transport proteins in the plastid envelopes. Iron. During germination and development and during iron stress, ferritin clusters in plastids serve as an iron store (Briat et al., 1999; Connolly and Guerinot, 2002); iron is absolutely required for photosynthetic electron transport, and Fe–S cluster biogenesis inside plastids involves the import of Fe2+ ions (Balk and Lobreaux, 2005; see also Section 12.2). Thus, plastids represent one of the most ironenriched systems in the plant cell. However, the iron transporter in the plastid envelope is not yet identified. Direct measurements of iron transport on isolated pea chloroplasts have shown that iron is transported in the form of ferrous ions across the inner envelope (Shingles et al., 2001, 2002). Fe2+ uptake into chloroplasts is most likely energised by a proton gradient and can be inhibited by Zn2+ , Cu2+ and Mn2+ . The putative metal uptake protein (Figure 7.4) would thus mediate an Fe2+ /H+ uniport and be able to transport Zn2+ , Cu2+ and Mn2+ as well. Copper. The two copper ATPases, PAA1/HMA6 and HMA1, in the inner envelope, as well as PAA2/HMA8 in the thylakoid membrane system (Figure 7.4; cf. 5.3.2.2), represent the only metal ion transport systems in chloroplasts that have so far been identified at the molecular level (Shikanai et al., 2003; Abdel-Ghany et al., 2005; Seigneurin-Berny et al., 2006). Both proteins belong to the group of heavy metal ATPases within the superfamily of P-type ATPases (for overview see Williams and Mills, 2005). While PAA1 and PAA2 were identified in screens for mutants with high chlorophyll fluorescence, HMA1 was first found in a proteomic analysis of chloroplast envelopes. Mutants of PAA1 and HMA1 both affect Cu content and Cu/Zn superoxide dismutase activity in chloroplasts; thus, both ATPases represent the Cu uptake system across the inner envelope.

INTRACELLULAR SOLUTE TRANSPORT

7.3

153

Mitochondria

Mitochondria are highly dynamic and complex semi-autonomous organelles, composed of a smooth outer membrane surrounding an extensively folded inner membrane of significantly larger surface area than the outer membrane, generating two aqueous compartments, the intermembrane space and the matrix, a protein-rich core. Mitochondria are typically long and oval shaped, ranging in size from 0.5 to 1 μm. The energy-transducing membrane (ATP synthesis) is the inner mitochondrial membrane, which has a highly pleomorphic structure due to numerous membrane invaginations, forming cristae stacks (see Bowsher and Tobin, 2001; Logan, 2006, for mitochondrial structure and compartmentalisation). So-called translocation contact sites between the outer and inner membrane have enabled the co-isolation of a protein import translocase complex (Schleyer and Neupert, 1985; Dekker et al., 1997; Schulke et al., 1999). The matrix contains the enzymes of the pyruvate dehydrogenase complex, the tricarboxylic acid cycle, and for the oxidative decarboxylation of glycine (during photorespiration), as well as pools of metabolites including NAD, NADH, ATP and ADP. It is suggested that the enzymes of the respective metabolic pathways in the matrix are arranged in multi-enzyme complexes, leading to an aqueous space in between, through which solutes can easily diffuse (Partikian et al., 1998). The most fundamental role of mitochondria is the synthesis of ATP formed by oxidative phosphorylation (Saraste, 1999). ATP production is coupled to the controlled dissipation of a proton electrochemical gradient across the inner membrane. This proton gradient is generated by the respiratory chain and in turn drives the ATP synthase complex to synthesise ATP from ADP and Pi.

7.3.1

The function of plant mitochondria

Many basic features of mitochondrial structure and function, developed at an early stage of evolution, have been highly conserved between animals and plants. Besides respiration and cellular energy supply, these include numerous transport systems for anions in the inner membrane and a general diffusion pore in the outer membrane. In contrast to animals, the bulk of fatty acid oxidation in plants is confined to peroxisomes (cf. Douce and Neuburger, 1989). However, mitochondria are involved in numerous other metabolic processes including the biosynthesis of amino acids, vitamin cofactors, and iron–sulphur clusters (for reviews see Mackenzie and McIntosh, 1999; Bowsher and Tobin, 2001). Furthermore, the plant mitochondrion is one of the three cell compartments involved in photorespiration (see Section 7.5; Douce and Neuburger, 1999), and is essential to several other plant-specific metabolic pathways including photosynthesis (Raghavendra and Padmasree, 2003) and the use of carbon and nitrogen storage compounds during seed germination (Picault et al., 2004). Mitochondria are implicated in Ca2+ -mediated signalling (Vandecasteele et al., 2001; Logan and Knight, 2003) and have been shown to be involved in PCD (Jones, 2000; Youle and Karbowski, 2005). In plants, PCD is regulated via the mitochondrial alternative oxidase, an enzyme that is unique to plants

154

PLANT SOLUTE TRANSPORT

and prevents oxidative stress (see Marechal and Baldan, 2002). In arabidopsis, the complete glycolytic pathway could be associated with mitochondria (Giege et al., 2003). Thus, there is a large-scale movement of metabolites, nucleotides and cofactors into and out of the mitochondria, linking the organelle with cellular metabolism.

7.3.2

Transport across the outer membrane: the porin VDAC

The outer membrane of mitochondria is freely permeable to solutes up to a size of 4–5 kDa (Figure 7.5; Benz, 1994), perhaps a legacy of the presence of general diffusion pores in analogy to the pore-forming proteins in the outer membrane of Gram-negative bacteria. In mitochondria, this β-barrel pore, with a molecular weight around 30–36 kDa, is called VDAC (voltage-dependent anion channel). Although primary amino acid sequences may vary, secondary structural elements, including an N-terminal α-helical domain and 16 antiparallel amphiphatic β-sheets are highly conserved among eukaryotic VDAC porins. While the pore is formed by the βsheets, the α-helical domain is involved in voltage-sensing and gating (Mannella, 1998). In lipid bilayer membranes in vitro, VDAC forms an aqueous channel with low substrate selectivity and thus functions more or less as a size-exclusion filter. However, evidence has accumulated that gating of VDAC is regulated by several factors such as NADH or proteins localised in the intermembrane space, indicating a complex mechanism of metabolite exchange in vivo (Colombini et al., 1987; Holden and Colombini, 1993; Zizi et al., 1994; Rostovtseva and Colombini, 1997; Vander Heiden et al., 2000). In yeast, it has been shown by mutant analysis that VDAC in the outer mitochondrial membrane is essential for mitochondrial respiration (Dihanich et al., 1987; Lee et al., 1998). Overexpression of a VDAC isoform from rice induces apoptosis in a mammalian cell line; therefore, it was suggested that VDAC in plants, as in animals, acts as a conserved element of PCD by participating in the release of intermembrane space proteins (Godbole et al., 2003). The first plant VDAC was identified electrophysiologically in purified outer membranes of pea mitochondria (Schmid et al., 1992). Since then, mitochondrial porins have been isolated and characterised from several plant species (Aljamal et al., 1993; Blumenthal et al., 1993; Abrecht et al., 2000a,b; Godbole et al., 2003; Wandrey et al., 2004). The VDAC protein from pea roots, originally described in non-green plastids (Fischer et al., 1994; cf. Section 7.2.2), was recently shown to be localised to mitochondria alone (Clausen et al., 2004). Immunoblot analysis, in vitro import experiments and fusion to GFP gave signals solely in mitochondria. In arabidopsis and L. japonicus, five different isoforms of VDAC are present and expressed constitutively throughout the plant (Clausen et al., 2004; Wandrey et al., 2004). Interestingly, the plant kingdom is thus equipped with more VDAC proteins than humans (3) or yeast (2), giving the opportunity to diversify physiological functions. However, all expressed VDAC isoforms (one VDAC is a possible pseudogene) in arabidopsis are detected within the mitochondrial proteome (Heazlewood et al., 2004; Millar et al., 2005). In the legumes L. japonicus and soybean, VDAC was immunodetected by in situ hybridisation in mitochondria and unknown vesicular structures close to the plasma membrane, but neither in peroxisomes nor in plastids

155

INTRACELLULAR SOLUTE TRANSPORT Solute fluxes across the OM ≤ 5 kDa

OM

IMS

IM

matrix H+

PiC

Arg breakdown

ATP synthesis Pi

+

Arg

TCA cycle

ADP

BA C1

OH-/ Pi

VDA C

AAC ATP

Heat H+

Mobilisation seed storage proteins

C SF

DTC

UCP Cold acclimation

Orn

Suc

ATP export

Di-, tricarboxylates

Ammonia assimilation

Fum Lipid mobilisation

Figure 7.5 Transport across the mitochondrial membranes: VDAC and carriers. Mitochondria are delimited by two membrane systems: the outer membrane (OM) and the inner membrane (IM), which is extensively folded and represents the energy-transducing membrane, harbouring the electron transport chain and ATP synthase complex (not shown). In turn, two compartments, the intermembrane space (IMS) and the matrix, are built. Upper half: Because of the β-barrel pore VDAC (16 β-sheets), the outer membrane of plant mitochondria is freely permeable to solutes with molecular weight up to 5 kDa. Thus, the voltage-gated VDAC is essential for mitochondrial function and central for transport of numerous solutes. Lower half: The inner membrane is equipped with several proteins of the mitochondrial carrier family (six α-helical domains). The phosphate carrier PiC and the ATP/ADP carrier AAC are required for ATP synthesis. PiC imports the inorganic phosphate (Pi) used for phosphorylation of ADP, which is imported by AAC. While PiC transport can be driven by proton symport or OH− /Pi antiport, AAC exchanges ADP with the synthesised ATP and thus supplies the plant cell with energy. In contrast, the uncoupling protein UCP is a carrier that bypasses ATP synthesis by importing protons. The resulting energy is dissipated as heat and thus UCP most likely functions in cold acclimation of plants. DTC can exchange di- and tricarboxylates across the inner membrane and thus feeds the TCA cycle in the matrix. A function of DTC during transport of ammonia from mitochondria to plastids (ammonia assimilation) is possible (cf. Figure 7.8). The amino acid carrier BAC1 is capable to mediate arginine (Arg) uptake, most likely in exchange for ornithine (Orn) during mobilisation of seed storage proteins in early seedling development. SFC as well functions in seed germination by importing succinate (Suc) in exchange for fumarate (Fum). Suc is produced by β-oxidation of fatty acids during lipid mobilisation.

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PLANT SOLUTE TRANSPORT

(Wandrey et al., 2004). In summary, by improvement and usage of miscellaneous assays for subcellular localisation of membrane proteins, evidence is accumulating that the predominant site for plant VDAC function is the outer membrane of mitochondria. Localisation of this porin in plastids or peroxisomes (cf. Sections 7.2.2 and 7.4.2.1) is most likely due to contamination of membrane fractions or cross-reactivity of antibodies. In contrast to the outer membrane, the inner mitochondrial membrane builds up the permeability barrier for solutes. Because ATP synthesis relies on the electrochemical proton gradient across the inner membrane, it is generally impermeable to charged or polar molecules. Therefore, numerous transport activities and proteins have been characterised in this membrane, including carriers, ABC transporters and ion channels.

7.3.3

Transport across the inner membrane: carriers

A mitochondrial carrier family of related proteins that span the inner membrane and mediate the selective transport of solutes has been shown to operate in yeast, animals and plants (reviewed in Laloi, 1999; Picault et al., 2004). All MCF proteins share a common secondary structure with six α-helical transmembrane domains, are nucleus encoded, operate as homodimers and have a molecular mass around 32 kDa. By genomic and proteomic analysis, up to 58 putative mitochondrial carriers have been reported in arabidopsis (Millar and Heazlewood, 2003; Picault et al., 2004). However, in databases and publications, several MCF members have annotated functions based on sequence similarities only. On the other hand, extensive measurements of metabolic fluxes across isolated mitochondrial membranes in the past 40 years have led to the postulation of numerous carrier functions without identifying the corresponding proteins (for an overview see Laloi, 1999; Picault et al., 2004). Thus, in the following sections we will principally focus on those transport capacities with molecular-assigned carrier proteins (Figure 7.5).

7.3.3.1

Transporters involved in ATP production

Oxidative phosphorylation, which leads to the formation of ATP in the matrix of mitochondria, is dependent on import of phosphate and ADP. Phosphate is taken up via a phosphate carrier (PiC) and ADP is exchanged with ATP by the ATP/ADP carrier (AAC). Furthermore, the uncoupling protein (UCP) is involved in the regulation of oxidative phosphorylation by decreasing the proton electrochemical potential difference across the inner membrane (see below). PiC, the phosphate carrier. PiC is responsible for a fast uptake of phosphate, which serves as substrate for phosphorylation of ADP. It can catalyse the phosphate (H 2 PO 4 − )/proton symport or phosphate/hydroxyl ion antiport as well as the exchange of matrix and cytosolic phosphate (Figure 7.5). This electroneutral transport is driven by the pH difference maintained across the membrane by the mitochondrial electron transport chain (for details see Laloi, 1999). The existence of a phosphate translocator in plant mitochondria has been suggested by swelling assays of isolated mitochondria in ammonium phosphate solution. Phosphate transport activity could

INTRACELLULAR SOLUTE TRANSPORT

157

be assigned to a membrane fraction from solubilised pea mitochondria (McIntosh and Oliver, 1994) and cDNA similarity screening finally led to the isolation of PiC from soybean, maize, rice and arabidopsis (Takabatake et al., 1999). The mitochondrial PiC in plants seems to be highly expressed in developing organs where tissues contain dividing cells, requiring a high energy level (overview by Laloi, 1999). This expression indicates that, together with the adenine nucleotide translocator (see below), the PiC plays an important physiological role in the energy supply for plant cells. AAC, the ATP/ADP carrier. AAC is the most abundant carrier protein in the mitochondrial inner membrane; it catalyses the exchange of ATP, synthesised by oxidative phosphorylation in the matrix, with cytosolic ADP (Figure 7.5). Pharmacological studies have demonstrated that ATP/ADP transport in plant mitochondria involves an AAC similar to that of mammals (see Laloi, 1999, for details). Subsequently, AAC was purified from maize mitochondria and shown to catalyse ATP/ATP and ATP/ADP exchange when reconstituted into liposomes (Genchi et al., 1996). Haferkamp et al. (2002) showed the same for the three mitochondrial AACs present in arabidopsis (Saint-Guily et al., 1992; Schuster et al., 1993). Expression of AAC is high and ubiquitous, the extent depending on the developmental state and regulation by external stresses. The main function for AAC is clearly in oxidative phosphorylation for import of ADP and export of ATP. However, roles in male sterility, uncoupling of mitochondria (see below) and apoptosis have been discussed (see Laloi, 1999). In an effort to compare mitochondrial and plastidic ATP/ADP transport, envelope membranes from pea root plastids, spinach chloroplasts and pea leaf mitochondria were reconstituted into liposomes (Sch¨unemann et al., 1993). On the basis of the determined transport characteristics, the authors conclude that plastid and mitochondrial AACs have derived from different ancestors. This could be confirmed by the identification of plastidic AACs from arabidopsis that belong to the major facilitator family and so are structurally different from the mitochondrial carriers (NTT; see Section 7.2.3.2). In summary, mitochondria evolved peculiar AACs that efficiently export ATP, whereas plastids acquired a different type of nucleotide transporter that seems to be specialised in ATP uptake (cf. Haferkamp et al., 2002). UCP/PUMP, uncoupling proteins in plant mitochondria. Mitochondrial UCPs in animals are carriers that transport protons present in the intermembrane space back to the matrix, thereby bypassing ATP synthase and thus dissipating the proton electrochemical potential difference (Figure 7.5). This process, which is mediated by free fatty acids, results in an increase in mitochondrial respiration, and the energy liberated by the oxidation of different substrates is dissipated as heat. In plants, the first evidence for the existence of a UCP-like protein called PUMP (plant uncoupling mitochondrial protein) was provided by analysis of potato mitochondrial respiration (for review see Laloi, 1999). Correspondingly, a cDNA encoding a peptide with high similarity to mammalian UCPs, StUCP, was identified (Laloi et al., 1997). In arabidopsis, two genes for UCP/PUMP have been isolated (Maia et al., 1998; Watanabe et al., 1999). Reconstituted into lipid bilayers, AtPUMP1 as well as the maize orthologue ZmPUMP catalyse linoleic-acid-induced proton fluxes (Borecky et al., 2001; Favaro et al., 2006). Plant UCPs are ubiquitously expressed, peak in

158

PLANT SOLUTE TRANSPORT

developing organs and, interestingly, some members of this family are induced by cold treatment. Thus, the latter might indicate that these proteins could be involved in heat production. Thermogenesis in plants has been previously described in different developmental processes such as fruit ripening and flowering, or after exposure to chilling temperature, which might contribute to cold acclimation or resistance to chill (see Laloi, 1999). However, there is still much ongoing work on the role of UCPs in plant metabolism, which might also be involved in protection against free oxygen radicals, since gene induction has also been reported in response to oxidative stress (Hourton-Cabassa et al., 2002, 2004; Brandalise et al., 2003a); overproduction of AtPUMP1 led to an increase in tolerance to oxidative stress (Brandalise et al., 2003b). To sum up, plant mitochondria contain two energy-dissipating systems: the alternative oxidase, which may prevent the build-up of a transmembrane potential, and the UCPs, which decrease this potential.

7.3.3.2

Carriers for transport of TCA cycle intermediates

The TCA (tricarboxylic acid) or Krebs cycle plays an important role not only in the breakdown of respiratory substrates but also in many biosynthetic pathways by supplying diverse intermediates. Both functions of the TCA cycle involve the activity of mitochondrial carriers either for the import of respiratory substrates such as pyruvate, malate and oxaloacetate, or for the constant export of intermediates. Previous and extensive biochemical characterisation of carrier function on mitochondria from diverse plants suggested the presence of monocarboxylate and dicarboxylate carriers, a citrate (tricarboxylate) transporter, an oxaloacetate transport system and the oxoglutarate/malate translocator to shuttle all these solutes (for overview see Laloi, 1999). However, only partial cDNA sequences for these transport proteins could be isolated. Thus, it appears most likely that except for the monocarboxylate pyruvate, most metabolites involved in the TCA cycle can be transported via the recently identified dicarboxylate–tricarboxylate carrier (DTC; see below). On the other hand, the MCF sequences in arabidopsis include three putative dicarboxylate carrier proteins (Picault et al., 2004), awaiting functional characterisation. Further, physiological and biochemical evidence for the export of reducing equivalents in the form of malate via malate/oxaloacetate or lactate/malate shuttles is still accumulating (Pastore et al., 2003; de Bari et al., 2005). Thus ‘the jury is still out’ on the in planta roles for dicarboxylate and tricarboxylate carriers in mitochondria. DTC, a dicarboxylate–tricarboxylate carrier. The transport of specific dicarboxylates and tricarboxylates (intermediates of the TCA cycle) across the inner mitochondrial membrane is required in several metabolic processes such as amino acid synthesis (nitrate/ammonium assimilation), export of reducing equivalents (for photorespiration), fatty acid metabolism (lipid mobilisation and fatty acid elongation), gluconeogenesis and isoprenoid biosynthesis. By overexpression in E. coli and reconstitution into phospholipid vesicles, it has been demonstrated that DTC proteins from arabidopsis and tobacco are capable of transporting both dicarboxylates (such as malate, oxaloacetate, oxoglutarate and maleate) and tricarboxylates (such as citrate, isocitrate, cis-aconitate and trans-aconitate) by a counterexchange mechanism (Figure 7.5; Picault et al., 2002). Furthermore, nitrate supply to

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nitrogen-starved tobacco plants leads to an increase in DTC mRNA in roots and leaves. DTC, which is ubiquitously expressed and widely found in the plant kingdom, differs from mammalian carriers with regard to its very broad substrate spectrum. Thus, it is concluded that the presence of DTC resolves previous inconsistencies concerning putative malate, citrate and oxaloacetate carriers in plants (see above; Laloi, 1999; Picault et al., 2002, 2004). Since DTC shows a high degree of similarity to oxoglutarate/malate carriers of animal mitochondria, Picault et al. (2002) propose that the oxoglutarate/malate carrier and DTC originated from a common ancestor, which in animals evolved into the distinct oxoglutarate/malate and tricarboxylate carrier, whereas in plants into DTC. As for AAC (see above), the MCF protein DTC with six α-helical membrane domains is structurally different from the dicarboxylate transporters in plastids, which belong to the major facilitator family (see Section 7.2.3.2). A role for DTC in nitrogen assimilation is proposed, because export of citrate or oxoglutarate from mitochondria has been suggested to be involved in shuffling ammonia from mitochondria to plastids (Lancien et al., 1999; Hodges, 2002; compare also photorespiration, Figure 7.8). Uptake experiments on plant mitochondria indicate that pyruvate transport has biochemical features similar to electroneutral pyruvate uptake into mammalian mitochondria, occurring in exchange for OH− (for an overview see Laloi, 1999). In pea mitochondria, a 19-kDa protein was assigned by a specific antibody to a protein fraction, capable to exchange pyruvate/pyruvate (Vivekananda and Oliver, 1990). It was proposed that PTP (pyruvate transport protein) acts in a multiple subunit protein, which in consequence would not belong to the MCF family. Since until now, no genes have been identified in any organism, the molecular nature of PTP remains enigmatic.

7.3.3.3

Amino acid transport across mitochondrial membranes

In plant mitochondria, uptake of glycine and export of serine during the photorespiratory cycle (see Section 7.5; Figure 7.8) requires an amino acid transporter. Although it has become more and more evident that glycine uptake involves a carrier (Laloi, 1999), neither the molecular identity nor whether serine can be transported by the same protein is known. Abiotic stress, such as high salinity or drought, causes proline accumulation in plants, involving proline transport into mitochondria where proline catabolism occurs (Di Martino et al., 2006, and references therein). Recently, transport activity by a putative carrier for proline and a postulated proline/glutamate shuttle was measured in wheat mitochondria (Di Martino et al., 2006). BAC, basic amino acid carriers. Two arabidopsis basic amino acid carriers, related to the yeast ornithine carrier, complemented the respective yeast mutants and were designated AtmBAC1 and AtmBAC2, respectively (Catoni et al., 2003a; Hoyos et al., 2003). The recombinant purified BAC1 was reconstituted into phospholipid vesicles and transported the basic amino acids arginine, lysine, ornithine and histidine (in order of decreasing affinity). High expression of BAC1 in seedlings is consistent with arginine uptake into mitochondria during arginine breakdown in early seedling development, when this amino acid serves as a nitrogen storage form (Figure 7.5; Hoyos et al., 2003). In contrast, the highest levels of BAC2 transcripts

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were found in flowers (i.e. pollen), in the vasculature of siliques and in aborted seeds, pointing to a different function of this protein (Catoni et al., 2003a).

7.3.3.4

Carriers involved in β-oxidation of fatty acids

The β-oxidation of fatty acids (occurring in peroxisomes, see Section 7.4) produces acetyl-CoA, which is converted via the glyoxylate cycle into succinate. This dicarboxylate must be transported into the mitochondria to be metabolised within the TCA cycle by succinate dehydrogenase, which is accessible to its substrate only from the mitochondrial matrix. Succinate is exchanged across the inner membrane for fumarate or malate via the succinate–fumarate carrier (SFC, see below). AcetylCoA generated by the β-oxidation of fatty acids can alternatively be taken up by mitochondria as acetyl-carnitine via the carnitine carrier (CAC) shuttle system, a yet unidentified protein (Lawand et al., 2002). SFC, a succinate–fumarate carrier. Complementation of a yeast mutant carrying a deletion of the SFC gene enabled functional identification of a mitochondrial succinate translocator, AtmSFC1, from arabidopsis (Figure 7.5; Catoni et al., 2003b). Expression of SFC1 in etiolated seedlings points to a role in the export of fumarate during lipid mobilisation at early seed germination, while in mature plants expression in developing and germinating pollen suggests a role in ethanolic fermentation.

7.3.4 7.3.4.1

Transport across the inner membrane: ABC transporters and ion channels ABC transporters

The ABC family is one of the largest protein families in living organisms (see also Sections 5.3.3, 7.2.4.1, and 7.6.3.2). These transporter proteins have various substrates, including ions, carbohydrates, lipids, xenobiotics, antibiotics, drugs and heavy metals (Martinoia et al., 2002). Arabidopsis contains approximately 130 ABC proteins, but the precise functions and substrate specificities of most of these transporters still remain obscure (Sanchez-Fernandez et al., 2001; Garcia et al., 2004). Three putative ABC transporters of the mitochondria subfamily of arabidopsis (ATM) are known and group into the ‘half-transporters’ with one transmembrane and one ABC domain. AtATM3 (alias STA1), whose deficiency causes dwarfism and chlorosis, most likely exports Fe–S clusters from mitochondria (Kushnir et al., 2001). The authors suggest that plant mitochondria possess an evolutionarily conserved Fe–S cluster biosynthesis pathway, which is linked to the intracellular iron homeostasis by the function of ABC transporters (Figure 7.6). Subsequent studies showed that AtATM3 is important for Cd(II) and Pb(II) resistance, possibly functioning as a transporter of glutathione-conjugated metals and Fe–S clusters across the inner mitochondrial membrane (Kim et al., 2006). Assembly of cytochrome c in plant mitochondria follows a pathway distinct from that of yeast and animals and more similar to that described for α- and γ proteobacteria. Faivre-Nitschke et al. (2001) gathered evidence that a potential ABC transporter in wheat mitochondria is involved in cytochrome c biogenesis in plants.

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161

Iron homeostasis Fe–S cluster

Kin Sur

ATM3 ATM3

IM

Energy dissipation

Matrix

K+ K+

?

Fe–S cluster biogenesis

H+

Cold acclimation, volume regulation, ... C Cl 1 Cl Nt

? Ca2+ signalling

Ca2+ Volume regulation, control membrane potential

Figure 7.6 ABC transporters and ion channels in the inner mitochondrial membrane. Two subunits of the ABC ‘half-transporter’ ATM3 mediate export of Fe–S clusters from the mitochondrial matrix. Thus, ATM3 links cellular iron homeostasis with mitochondrial Fe–S cluster biogenesis. The activity of ion channels in the inner mitochondrial membrane has been demonstrated, but the nature of the corresponding proteins is unknown. Import of K+ ions is most likely mediated by an inward-rectifying K+ channel (K in ), coupled to a sulphonylurea receptor (Sur). Together with a potential K+ /H+ exchanger this potassium transport system might function in energy dissipation analogous to UCP (see Figure 7.5). Ca2+ most likely is transported by a uniporter, analogous to animal mitochondria. The chloride channel CLC-Nt1 (12 α-helical transmembrane domains) was localised to tobacco mitochondria. A putative role of Cl− transport is volume regulation (together with K+ ) and control of the potential across the inner membrane.

7.3.4.2

Ion channels

Ion transport across the inner mitochondrial membrane has been reported for potassium (energy dissipation), calcium (cell signalling) and anions/chloride. However, the molecular identities of the corresponding transport proteins still remain unknown. Potassium. In recent years, a new energy-dissipative mechanism was described in plant mitochondria involving K+ import into the mitochondrial matrix and K+ /H+ exchange (Pastore et al., 1999; Petrussa et al., 2001, 2004; Chiandussi et al., 2002). In this manner, K+ , the most abundant cation in the cytosol, regulates coupling between respiration and ATP synthesis in plant mitochondria. K+ import appears to be inhibited by ATP, suggesting the channel is similar to mammalian mitochondrial, ATP-sensitive K+ channels. The discovery of such a channel brought new implications to the physiology of this organelle, because the existence of a K+ import

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channel acting together with a potent K+ /H+ exchanger (Diolez and Moreau, 1985) would allow regulation of the proton potential through a K+ cycle analogous to that found in animals, which possess a K+ transporter most likely composed of a sulphonylurea receptor and an inward-rectifying K+ channel (Figure 7.6; Garlid, 1996; Mironova et al., 2004). Hypothetical roles for this channel would be volume regulation (as proposed for the two other energy-dissipating processes [UCP, AOX, see Section 7.3.3.1]), thermogenesis, apoptosis (via cytochrome c release) and/or prevention of oxidative stress. Flux experiments on isolated mitochondria from soybean suspension cultures indicate the involvement of a K+ -ATP channel during the manifestation of PCD induced by H 2 O 2 or NO (Casolo et al., 2005). However, ATP sensitivity of K+ transport into plant mitochondria is still a matter of debate, since Ruy et al. (2004) report the existence of a highly active ATP-insensitive K+ import pathway in plant mitochondria. Calcium. During cell activation, animal mitochondria play an important role in Ca2+ homeostasis because of the presence of a fast and specific Ca2+ channel in the inner membrane, the mitochondrial Ca2+ uniporter (see Montero et al., 2004, and references therein). The role of mitochondrial calcium in plant cell signalling has received little attention, although Logan and Leaver (2000) showed using mitochondrial targeted aequorin that internal Ca2+ is modulated by physiological and environmental stimuli. It is speculated that as in animals, the main targets of mitochondrial Ca2+ are the dehydrogenases of the TCA cycle. By activating these enzymes, calcium could stimulate respiration, and in turn increase ATP production. Anions/chloride. It has long been established that the inner membrane of plant mitochondria is permeable to chloride (Beavis and Vercesi, 1992). As a result of classical mitochondrial swelling assays it was proposed that anion uniport in plant mitochondria is mediated via a pH-regulated channel related to the ‘inner membrane anion channel’ (IMAC) of animals. Since Lurin et al. (2000) localised the tobacco chloride channel, CLC-Nt1, to mitochondria (Figure 7.6), they speculate that this protein might be the IMAC-like plant orthologue. Functions of IMAC in plant mitochondria would be volume regulation and control of the potential across the inner membrane.

7.4

Peroxisomes

In contrast to plastids and mitochondria, peroxisomes are enclosed by a single membrane. Initially, the term microbody was introduced by mammalian microscopists to describe a membrane-surrounded particle of unknown function. Since then, subclassifications, such as peroxisomes and glyoxysomes, have been used for microbodies with different metabolic pathways and tissue distribution (Tolbert, 1971). The synonymously used term peroxisome refers to the primary function in compartmentalisation and thus protection from oxidases, which produce hydrogen peroxide and reactive oxygen species. Because of their function in oxidative metabolism, peroxisomes are essential, ubiquitous organelles, present in all eukaryotes.

INTRACELLULAR SOLUTE TRANSPORT

7.4.1

163

Function of peroxisomes in plant metabolism

In plants, peroxisomes are small organelles (0.2–0.5-μm diameter) and have been differentiated into at least three different classes, namely glyoxysomes, leaf peroxisomes and unspecialised peroxisomes (Beevers, 1979). They generate reactive oxygen species and contain defence mechanisms in the form of catalase (detoxification of H 2 O 2 ), superoxide dismutase and the membrane-bound ascorbate peroxidase. In addition, a central function for most plant peroxisomes is the β-oxidation of fatty acids. However, the situation is seen currently as being more complex, since an exceptionally large number of morphological and metabolically specialised peroxisomes have been discovered in plant cells (for reviews see Reumann, 2000; Mano and Nishimura, 2005; Theodoulou et al., 2006). Leaf peroxisomes are present in photosynthetic tissue and besides β-oxidation of fatty acids are responsible for photorespiration, even though the entire process is distributed between chloroplasts, leaf peroxisomes and mitochondria (Reumann, 2000). In contrast, glyoxysomes reside in cells of storage tissue, such as endosperm, and in cotyledons during germination of oilseed plants. They contain enzymes for fatty acid oxidation and for the glyoxylate cycle and play a pivotal role in the conversion of lipid reserves into sucrose. In senescing tissue, glyoxysomes might additionally be involved in degradation of amino acids. The functions of glyoxysomes and peroxisomes are known to interconvert during cellular processes (Hayashi et al., 2000). In germinating seedlings, for example, glyoxysomes in cotyledons are functionally transformed into leaf peroxisomes upon illumination. The reverse process has been observed when leaves undergo senescence. Apart from the primary functions in oxidative metabolism, βoxidation of fatty acids, photorespiration (leaf peroxisomes) and lipid mobilisation (glyoxysomes), plant peroxisomes also play a significant role in nitrogen assimilation (root nodule cells of leguminosae; Verma, 2002), degradation of branched amino acids and biosynthesis of plant hormones including jasmonic acid and auxin (Stintzi and Browse, 2000; Zolman et al., 2001; Feussner and Wasternack, 2002).

7.4.2

Solute transport across the peroxisomal membrane

The functioning of peroxisomes in their diverse physiological processes requires a tightly regulated transport of solutes and metabolites across their membrane. However, to date, the plant peroxisomal membrane is one of the least well characterised, mainly because of the difficulties in isolating pure membranes free of contamination by other organelles. According to yeast nomenclature, plant PMPs – peroxisomal membrane proteins – are generally classified according to their molecular mass in kilodaltons.

7.4.2.1

A porin in the peroxisomal membrane?

Membrane isolation of spinach leaf peroxisomes and glyoxysomes of castor bean endosperm led to the electrophysiological characterisation of an anion-selective, specific porin (Reumann et al., 1995, 1997, 1998). This peroxisomal porin was

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distinctively different from VDAC in mitochondria (cf. Section 7.3.2). For example, the peroxisomal porin showed a substantially lower single-channel conductance and a stronger anion selectivity than the mitochondrial VDAC, and thus formed smaller, more specific channel pores, reminiscent of ‘specific porins’ in Gram-negative bacteria. It was proposed that metabolism of peroxisomes is in general not compartmentalised by the boundary membrane, but by the strictly organised arrangement of matrix enzymes in multienzyme complexes (Heupel and Heldt, 1994). In consequence, the porin-like channel mediates the diffusion of a broad range of negatively charged metabolites (Figure 7.7; cf. Reumann, 2000). During photorespiration in

Negatively charged metabolites?

Porin?

Phytohormone biosynthesis FA-CoA

ß-oxidation

AMP + PPi FA

S CT PMP 38

Hormone precursors (FA-CoA?), FA Lipid mobilisation in glyoxysomes

ATP CoASH

ADP

? PM P2 ? 2

? CoASH

Figure 7.7 Solute transport across the peroxisomal membrane. A ‘specific porin’ in the peroxisomal membrane would mediate the diffusion of a broad range of negatively charged metabolites. However, the molecular identity of this porin is still unclear. PMP38, the peroxisomal ATP/ADP carrier is similar to AAC, the ATP/ADP carrier in mitochondria (Figure 7.5) and imports ATP, required for synthesis of fatty acyl coenzyme A (FA-CoA), in exchange for ADP. Regulation of PMP38 expression points to a function in lipid mobilisation during seedling development. Theodoulou et al. (2006) propose that CTS, an ABC transporter, imports free fatty acids (FA), which are esterified to coenzyme A (CoA), and then are catabolised by β-oxidation. Again CTS action is required during lipid mobilisation. In turn, the cofactor CoASH has to be transported via a still unknown protein. Uptake of already activated fatty acids (FA-CoA) by CTS, however, cannot be excluded. In addition CTS might transport auxin precursors and/or jasmonic acid, showing a broad substrate range, reminiscent of the action of multidrug resistance ABC transporters. Substrates and function for the transmembrane protein PMP22 have not been identified yet.

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leaf peroxisomes, the porin would be able to transport the intermediates glycolate, glycerate, glutamate and α-ketoglutarate while in glyoxysomes it would mediate the flux of citrate, isocitrate, succinate and malate (glyoxylate cycle). Further, the porin should provide redox equivalents via a shuttle function for malate/oxaloacetate (peroxisomes) or malate/aspartate (β-oxidation in glyoxysomes). In this model, only the transport of neutral amino acids, like Ser or Gly, or of fatty acids for β-oxidation requires the activity of other transport proteins. However, up to now the molecular identity of this ‘multi-tasking’, specific, peroxisomal porin could not be assigned. An attempt by Corpas et al. (2000) led to the isolation of a porin-like peptide from glyoxysomal membranes of cucumber. This peptide belongs to a 36-kDa protein band and shows strong similarity to VDAC from pea, arabidopsis and other oilseed plants. Whether one of the five VDAC isoforms present in arabidopsis (cf. Section 7.3.2) localises to peroxisomes remains an open question. Because of the difficulties in isolating adequate amounts of pure peroxisomal membranes, proteome analysis has been performed only in silico by analysis of peroxisomal targeting sequences in the entire arabidopsis proteome (Reumann et al., 2004). Disappointingly, among the 282 putative peroxisomal proteins only two transporters, namely the ABC transporter CTS and the ATP/ADP carrier PMP38 (see below), could be identified. Thus, this attempt helped to identify the nature of neither the “specific porin” nor membrane transporters with new functions.

7.4.2.2

Specific transport proteins in the peroxisomal membrane

The question whether metabolite transport across the peroxisomal membrane is mediated by a porin-like channel or, in contrast, needs specific transport proteins is still a matter of debate. In yeast and mammalian peroxisomal membranes, the presence of transport ATPase activity, a pH gradient and the requirement of specific shuttles for NADH and acyl-CoA argue against the porin model (see Mullen and Trelease, 1996; Theodoulou et al., 2006, for discussion). Furthermore, genetic manipulation of transport systems revealed that import and export of several ions, metabolites and cofactors are tightly controlled and protein mediated. However, in plants only two peroxisomal membrane transporters have been identified at the molecular level, so far. PMP38, the peroxisomal ATP/ADP carrier. Preparation of glyoxysomal membrane fractions from pumpkin led to the discovery of the peroxisomal AAC (Fukao et al., 2001). The arabidopsis orthologue PMP38 is an integral membrane protein, similar to the mitochondrial AAC (Section 7.3.3.1), contains six predicted α-helical transmembrane domains and therewith belongs to the mitochondrial carrier family MCF. The peroxisomal localisation of AtPMP38 was verified by immunoblot analysis on fractionated organellar membranes and immunogold labelling. Since expression of AtPMP38 is decreased when glyoxysomes transform into peroxisomes during illumination of seedlings, it is suggested that PMP38 imports ATP, required for synthesis of fatty acyl-CoA in the glyoxysomal matrix. Thus, PMP38 would function in concert with the fatty acid β-oxidation cycle (Figure 7.7). CTS, an ABC transporter in plant peroxisomes. The necessary import of fatty acids for β-oxidation is most likely performed by the full-size ABC transporter CTS

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(‘COMATOSE’, for review see Theodoulou et al., 2006). Peroxisomal ABC transporters in all eukaryotes belong to subfamily D and, with the exception of the plant CTS, all ABCD proteins are ‘half-size’ (for ABC transporters see Sections 7.2.4.1, 7.3.4.1 and 7.6.3.2). Originally, CTS (Footitt et al., 2002) was identified in three independent forward genetic screens and is also known as PXA1 (Zolman et al., 2001) or PED3 (Hayashi et al., 2002). By immunoblot analysis on fractionated organellar membranes and immunogold labelling, CTS was localised to peroxisomes. Isolation and characterisation of the respective mutant alleles provided considerable insight into the function of ABC transporters in plant peroxisomes. PXA1 and PED3 mutants are blocked in β-oxidation of precursors of auxin as well as of fatty acids. CTS mutant alleles are impaired in germination, and cotyledons show a pronounced inability to break down lipid bodies. A model was suggested in which CTS mediates transport of fatty acids, fatty acid acyl-CoA or cofactors for β-oxidation into peroxisomes (Figure 7.7; Theodoulou et al., 2006). Moreover, CTS might transport precursors of auxin as well as the lipid-derived jasmonic acid (Theodoulou et al., 2005) and thus be involved in the peroxisomal biosynthesis pathways of these phytohormones. Another potential membrane transporter from plant peroxisomes is PMP22 from arabidopsis, which was isolated by database screening (Tugal et al., 1999). PMP22 is an integral membrane protein (four to five transmembrane domains), similar to mammalian and yeast peroxisomal proteins, but up to now no function has been assigned to this protein. Although not a transmembrane protein, another interesting peptide is represented by the arabidopsis orthologue to the mammalian sterol carrier protein 2 (SCP-2), an intracellular, small basic protein that enhances the transfer of lipids between membranes (Edqvist et al., 2004). AtSCP-2 localises to peroxisomes and can catalyse the in vitro lipid transfer between membranes. It is speculated that SCP-2 might play a role in β-oxidation of fatty acids during chlorophyll catabolism.

7.5

Photorespiration: transport between plastids, mitochondria and peroxisomes

Because of the oxygenase function of ribulose-1,5-bisphosphate carboxlyase/ oxygenase (Rubisco) in the light, leaves of C3 plants evolve CO 2 and consume O 2 , leading to a complex multi-compartment pathway called photorespiration. The photorespiratory pathway involves cross talk and solute transport between plastids, mitochondria and peroxisomes (Figure 7.8; Douce and Neuburger, 1999). Photorespiration starts in the chloroplast stroma with the oxygenation of ribulose-1,5bisphosphate by Rubisco to produce one molecule of 3-PGA, which is fed into the Calvin cycle, and one molecule of 2-phosphoglycolate (2-Pglt), which is dephosphorylated. The resulting product glycolate (Glt) is shuttled to peroxisomes [see Figure 7.8 for main reactions and transport pathways (i) to (vi)]. The molecular identity of glycolate transporters in the chloroplast envelopes as well as in the peroxisomal membrane is unknown, although a peroxisomal porin would be able to mediate

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INTRACELLULAR SOLUTE TRANSPORT Plastid Peroxisome O2 Rubisco

(i)

2-Pglt

Glt

?

Mitochondrion

Glt

(iv)

3-PGA Glc

Calvin cycle

?

?

Gox ?

(ii)

Glc

Gly ?

?

Gly Gly

Glu GOGAT

?

Ser

Glu

?

?

Ser

CO2 NH4+

Mal Mal 2-OG

Gln

DiT2

(iii)

DiT1

Ru 1,5-BP

? 2-OG

(v)

Glu NH3 GS

Orn or Glu ? ?

(vi) Cit or Gln ?

? DTC

Figure 7.8 Solute transport during photorespiration. Photorespiration involves solute transport between plastids, peroxisomes and mitochondria. Reactions and transport processes are described in the text and depicted according to Douce and Neuburger (1999), Reumann (2000) and Linka and Weber (2005). For the sake of simplicity, biochemical reactions, enzymes and cofactors are not pictured in detail. Furthermore, the outer membrane of plastids and mitochondria is omitted. While in mitochondria, VDAC can mediate transport of the respective solutes across the outer membrane (see Section 7.3.2), the pathway through the outer envelope of plastids is unclear (see Section 7.2.2). The numbers (i)–(vi) represent the respective transport steps as described in the text. Transport proteins with unknown molecular nature are represented by ‘?’. The dicarboxylate transporters DiT1 (import of 2oxoglutarate [2-OG]) and DiT2 (export of glutamate [Glu]) in the inner envelope of plastids function via the exchange of malate (Mal, cf. Figure 7.2). They are involved in transamination processes (see Section 7.2.3.2). The released CO 2 in mitochondria is assumed to diffuse back to chloroplasts (dotted arrow), while for NH 4 + export (dashed arrows) two different pathways are proposed (Linka and Weber, 2005, for details). A function of the dicarboxylate/tricarboxylate transporter DTC during ammonia export from mitochondria is possible (cf. Section 7.3.3.2). Back in the chloroplast, ammonia is assimilated into glutamate by the action of glutamine synthetase/glutamate synthase (GS–GOGAT) cycle. Please note that the cytotoxic NH 4 + /NH 3 have to be transported and transiently stored in the form of amino acids.

this transport activity (see Section 7.4.2.1; Reumann, 2000). Next, glycolate is oxidised to glyoxylate (Gox), which is subsequently transaminated to glycine (Gly) in the peroxisomes. Glycine has then to be transported to mitochondria for further reactions (see below). While glycine can pass the outer membrane of mitochondria via VDAC, the export of this amino acid from peroxisomes is definitely not mediated by a porin (Reumann, 2000). Thus, transporters for glycine in the peroxisome and inner mitochondrial membrane have not yet been identified. In the mitochondria, two molecules of glycine are converted to one molecule each of serine (Ser), ammonia and carbon dioxide; serine is then transported back to the peroxisome.

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As for glycine, the relevant transport proteins in both organelles are unknown (cf. Sections 7.3.3.3 and 7.4.2). In peroxisomes, serine is converted to glycerate (Glc) and shuttled back to the chloroplast, again by unknown transport systems. However, a carrier for both glycolate and glycerate in the chloroplast inner envelope has been suggested (Douce and Neuburger, 1999, and references therein). Back in the chloroplast, glycerate is phosphorylated to 3-PGA and fed into the Calvin cycle. In summary, the reaction cycle converts two molecules of 2-Pglt into one molecule of 3-PGA, one molecule CO 2 and one molecule of ammonia. Ammonia assimilation in chloroplasts (see below) and transamination in peroxisomes (Gox to Gly) involves the action of a malate(Mal) coupled two-transporter system most likely represented by the dicarboxylate transporters DiT1/OMT1 and DiT2/DCT1 in the inner envelope of plastids (see Section 7.2.3.2). Again, if not mediated by a porin, transporters in the peroxisomal membrane are unknown. In mitochondria, photorespiration generates massive amounts of the cytotoxic metabolite ammonia, which in turn is assimilated in plastids by the GS–GOGAT system (see Section 7.2.3.2). Thus, a shuttle system between mitochondria and plastids is required for ammonia, transiently storing this toxic metabolite in the form of amino acids (Linka and Weber, 2005). However, it is still unclear whether this pathway involves the function of an ornithine–citrulline (Orn–Cit) or a glutamate–glutamine (Glu–Gln) shuttle (see Douce and Neuburger, 1999; Linka and Weber, 2005, for discussion). To sum up, the cross talk and intracellular transport between plastids, peroxisomes and mitochondria during photorespiration involve at least six independent transport steps across the membranes of three organelles. While the biochemistry of the photorespiratory cycle is an established textbook material, the identity of the metabolite transporters involved still has to be unravelled.

7.6

Vacuoles

Like plastids, vacuoles are plant-specific organelles and represent the major compartment of the plant cell, irrespective of the cell type. In mature plant cells, the central vacuole, delimited by a single membrane called the tonoplast, occupies 80– 90% of the cell volume. Plants have only a limited capacity to excrete potentially toxic compounds. Thus, the best known function of vacuoles is the compartmentalisation and detoxification of xenobiotics (i.e. synthetic chemicals present in the plant’s environment). Based on this function it has been suggested that the distance between life and death is 7.5 nm, the thickness of the tonoplast (Matile, 1984). However, plant vacuoles are morphologically and functionally diverse organelles, with many additional roles, including recycling of cell components, regulation of turgor pressure, cytoplasmic pH and cytoplasmic calcium, as well as storage of primary (amino acids, sugars and malate) or secondary metabolites (Barkla and Pantoja, 1996; Maeshima, 2001; Yazaki, 2005). Excess metabolites are transported into the vacuole, which serves as a transient storage pool, and released to the cytoplasm when required for metabolism. Furthermore, the space-filling function of the vacuole is essential for cell growth, because cell enlargement is accompanied by

INTRACELLULAR SOLUTE TRANSPORT

169

expansion of the vacuole rather than of the cytoplasm. The vacuole plays a role in tolerance to environmental stress, including exposure to chemicals, heavy metals, plant pathogens and salt stress (see Yamaguchi et al., 2002, and references therein; Chapter 14). In general, the vacuole, as the biggest organelle, is necessary for plant cell homeostasis. Thus, vacuoles contain a large number of hydrolytic and biosynthetic enzymes, inorganic ions, soluble carbohydrates, organic acids, amino acids, secondary compounds and modified xenobiotics (Maeshima, 2001). All these are compounds that have to be transported across the tonoplast. Some vacuoles function primarily as storage organelles, others as lytic compartments. In consequence, more than one kind of vacuole has been observed in cells undergoing differentiation, maturation and autophagy, as well as in fully differentiated cells (see Bethke and Jones, 2000, and references therein).

7.6.1

Generating a pH gradient across the tonoplast: H + -ATPase and H + -pyrophosphatase

Transport activity across the tonoplast is dominated by two primary active transport mechanisms requiring high-energy metabolites for their operation: the vacuolar H+ -ATPase (V-ATPase; see also Section 5.3.1.2) and the H+ pyrophosphatase (V-PPase; see also Section 5.3.1.3). Thus, plant vacuoles are unique among eukaryotic organelles in having two proton pumps (Figure 7.9). Both proteins are among the most abundant peptides in the vacuolar membrane and have been extensively characterised at the functional and molecular level (see Barkla and Pantoja, 1996; Maeshima, 2001, for a detailed description). By using energy in the form of ATP, the V-ATPase is pumping protons from the cytosol into the vacuole, thereby generating a pH gradient across the tonoplast. This proton motive force in turn provides the driving force for a wide range of secondary active and passive transport processes. Furthermore, the V-ATPase controls cellular pH homeostasis. The V-ATPase is a multisubunit enzyme complex, structurally related to F 0 F 1 -ATPases. A possible function for the V-ATPase is salt tolerance of plants (Maeshima, 2001) as well as seedling growth, since mutants of DET3, the C-subunit of the V-ATPase, show deficiencies in hypocotyl cell expansion (Schumacher et al., 1999). Probably, a decrease in V-ATPase limits the accumulation of vacuolar solutes and hence the osmotic driving force for growth. An alternative vacuolar proton pump is the V-PPase, for which PPi is used instead of ATP as the energy donor. The V-PPase is essential for maintaining the acidity of the large central vacuole. The H+ /PPi stoichiometry has been determined to be 1.0, and the steady-state pH gradient generated across the tonoplast against the cytoplasmic pH (≈7.0) is approximately 3 pH units. In contrast to the V-ATPase, the V-PPase consists of a single polypeptide and acts as a dimer. Overexpression of V-PPase in Arabidopsis leads to salt- and drought-tolerant plants (Gaxiola et al., 2001), showing that V-PPase activity energises the accumulation of toxic cations inside the vacuole by the Na+ /H+ antiporter (see Section 7.6.4.2 and 14.12 for discussion of the physiology).

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pH homeostasis Cell turgor (growth) Photosynthesis

ATP

tDT

Pi e Pas V-P e Pas V-P

Salt tolerance Cell growth

H+

Mal

Suc

SU T4

PPi

Osmoregulation Cell volume

ADP V-ATPase

H2O H+

pH gradient

Storage

TIP TIP

Urea Glycerol

Storage of xenobiotics, secondary metabolites

Detoxification

IDI 7 IDI 7

MR P

GS (Glutathione) -X X-Glucose X-Glucuronic acid

Phytosiderophores? Iron acquisition?

Figure 7.9 Solute transport across the tonoplast. Two proton pumps in the tonoplast acidify the plant vacuole. While the V-ATPase is a multisubunit complex (F 1 F 0 -ATPase), the V-PPase consists of a single polypeptide and acts as a dimer. Both proteins are linked to a function in salt tolerance and plant growth, but first and foremost they establish a pH gradient across the tonoplast, which drives secondary transport of metabolites, e.g. calcium, sodium and magnesium uptake (see Figure 7.10). The major facilitator tDT (12 α-helical transmembrane domains) imports malate (Mal) into the vacuole, thereby controlling cell turgor and growth as well as pH homeostasis. Malate and photosynthetically derived sucrose (Suc) are stored in the vacuole. The latter as well is transported by a major facilitator protein: SUT4 in Arabidopsis. A tetramer of TIP subunits mediates water flow across the tonoplast and thus is crucial for osmoregulation and cell expansion/shrinking. Further TIPs can facilitate transport of small solutes like urea or glycerol. ‘Full size’ ABC transporters of the MRP subfamily transport xenobiotics and secondary metabolites (X) into the vacuole and thus are crucial for detoxification processes. In general these compounds are conjugated to either glutathione (GS) or glucose or glucuronic acid. The ‘half-type’ ABC transporter IDI7 as well localises to the tonoplast. It can only be speculated that IDI7 transports phytosiderophores for iron acquisition in grass roots.

7.6.2 7.6.2.1

Transport of malate and sucrose across the tonoplast Malate

In plant cells, excess malate is stored within the large, central vacuole. Malate serves as a storage form of fixed CO 2 , as charge balance and as an osmolyte for maintenance of cell turgor. The transport of malate into the vacuole is crucial for

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the regulation of cytoplasmic pH and the control of metabolism (e.g. in CAM; see also Section 13.6). The existence of a vacuolar malate/citrate transporter has been described at the functional level, using flux analysis, membrane-potential- and pHdependent fluorescence probes, as well as electrophysiological analysis on isolated vacuoles or tonoplast membranes (Emmerlich et al., 2003; Hurth et al., 2005, and references therein). The vacuolar malate transporter tDT (tonoplast dicarboxylate transporter) was subsequently isolated from arabidopsis (Emmerlich et al., 2003). AttDT contains 12 α-helical transmembrane domains (major facilitator family) and exhibits high sequence similarity to the human sodium/dicarboxylate cotransporter (Figure 7.9). In deletion mutants, cellular as well as vacuolar malate and fumarate contents were strongly reduced while citrate levels were increased (Emmerlich et al., 2003; Hurth et al., 2005). Furthermore, the residual malate import into vacuoles of mutant plants was not inhibited by citrate. Knockout plants also exhibited an increased respiratory coefficient, indicating a shift of the respired substrates from carbohydrates in wild type to mainly organic acids in the mutant. In conclusion, mutant analysis showed that arabidopsis vacuoles contain, besides tDT, another transport capacity for malate as well as a channel for dicarboxylates and citrate. The function of AttDT, however, was critical for regulation of pH homeostasis.

7.6.2.2

Sucrose

In leaves, a large proportion of the sucrose produced by photosynthesis is transported into the vacuole during the light period for storage. At night, sucrose is released and loaded into the phloem for transport to the sink tissues (see Endler et al., 2006, and references therein). Thus, vacuolar sucrose transport is crucial for cytoplasmic sucrose concentrations and, in turn, the function of photosynthesis. Furthermore, many monocotyledonous plants, such as barley and wheat, synthesise fructans for carbohydrate storage or as cold-, drought- and salt-stress protectants. Fructan synthesis occurs within the vacuole and requires sucrose. Facilitated diffusion of sucrose across isolated vacuoles has been demonstrated in barley (Kaiser and Heber, 1984; Martinoia et al., 1987). Only very recently, the corresponding vacuolar sucrose transporter was isolated using a proteomic approach analysing the tonoplast fraction of purified mesophyll vacuoles from barley (Endler et al., 2006). The identified protein, HvSUT2, and its arabidopsis orthologue, AtSUT4, belong to the major facilitator family (see Section 7.2.3.2) and localise to tonoplast membranes (Figure 7.9). The authors conclude that both proteins are involved in the transport and vacuolar storage of photosynthetically derived sucrose. Furthermore, this proteomic analysis provides additional protein sequences, e.g. a potential vacuolar hexose transporter (Endler et al., 2006).

7.6.3 7.6.3.1

Aquaporins and ABC transporter in the tonoplast Aquaporins in the vacuole are tonoplast-intrinsic proteins

The so-called tonoplast-intrinsic proteins (TIPs; see also Section 5.2.4) represent the most abundant proteins in the vacuolar membrane. In vitro, when expressed in oocytes of X. laevis, many of the TIPs function as water channels (aquaporins; Bethke

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and Jones, 2000; Maeshima, 2001, and references therein). Aquaporins facilitate water transport across biomembranes in an osmotic pressure-dependent manner. TIPs as well as PIPs (plasma membrane intrinsic proteins) belong to the ubiquitous family of membrane-intrinsic proteins. These proteins have a common structure of six membrane-spanning α-helices and two short α-helices each including an NPA motif and function as homotetramers (Figure 7.9). The TIP endomembrane-type aquaporins are unique to plants, and arabidopsis contains ten genes coding for TIP isoforms. They mediate water exchange between the cytosolic and vacuolar compartments and play a central role in cell osmoregulation under osmotic stress, drought and salinity, as suggested by several detailed analyses of TIP gene expression (see Maeshima, 2001; Luu and Maurel, 2005). TIP-facilitated water flow may be important in protecting the cell against plasmolysis, since water stress affects the abundance of TIP mRNAs and proteins (Barrieu et al., 1999; Sarda et al., 1999). Furthermore, the high abundance of γ -TIP in the motor cell of mimosa plants points to a function in rapid cell expansion and/or shrinking during movement of mimosa leaves, triggered by mechanical touch (Fleurat-Lessard et al., 1997). Because different TIP isoforms express in different types of vacuole, TIPs have been proposed to be markers of vacuolar function and development (Jauh et al., 1999; Moriyasu et al., 2003). In addition to facilitating water flow, TIPa from tobacco was shown to transport urea and glycerol (Gerbeau et al., 1999). Thus, plant TIPs may function in both water and solute transport and participate in long-term regulation of cytosolic and vacuolar volumes. In Mesembryanthemum crystallinum, immunodetection of a TIP isoform in membrane fractions revealed that this aquaporin can be redistributed from tonoplast to other endomembrane fractions after hyperosmotic treatment with mannitol (Vera-Estrella et al., 2004). This redistribution provides insight into a novel mechanism for regulation of aquaporins in response to osmotic stress and may point to a TIP function in vesicle-sorting mechanisms.

7.6.3.2

ABC transporters in the tonoplast

Full-size ABC proteins of the MRP subfamily (multidrug-resistance-related protein) are localised in the vacuolar membrane and facilitate the accumulation of secondary metabolites and xenobiotics (for an overview of plant ABC transporters, see Section 5.3.3; Higgins, 2001; Sanchez-Fernandez et al., 2001; Garcia et al., 2004). In general, secondary metabolites and xenobiotics are conjugated to glutathione, glucuronic acid or glucose before storage in the vacuole (Figure 7.9). Thus MRP proteins sequester glutathionylated compounds, glucuronides, glucosides as well as malonylated chlorophyll catabolites (see Martinoia et al., 2002; Yazaki, 2005). The ABC transporters MRP3 and MRP4 in maize vacuoles are required for uptake of anthocyanin glucosides into the vacuole (Goodman et al., 2004). Mutants of MRP3 have a distinct pigmentation phenotype in the adult plant that results from a mislocalisation of the pigment as well as significant reduction in anthocyanin content. However, in arabidopsis, vacuolar transport of anthocyanin most likely is mediated by an H+ antiport mechanism via a multidrug and toxic compound extrusion protein (MATE) (Yazaki, 2005, and references therein). The MRP subfamily in arabidopsis contains 15 members and represents the best characterised plant ABC

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transporters. However, it is still not clear how many kinds of ABC transporters are in the tonoplast, what substrates they transport and how they are regulated in vivo. AtMRP2 has been localised to the arabidopsis tonoplast (Liu et al., 2001) and shows glutathione-conjugate transport into yeast vacuoles. Thus, MRP2 most likely functions in detoxification of xenobiotics, which are conjugated to glutathione. However, MRP2 is also able to transport chlorophyll catabolites during leaf senescence (Lu et al., 1998; Tommasini et al., 1998) and has been described as transporting glucuronides (Liu et al., 2001). An MRP-like protein from rye mediates uptake of flavone glucuronides into tonoplast vesicles. This transport is dependent on the membrane potential, pointing to a possible regulation by vacuolar proton pumps (Klein et al., 2000). Transport mechanisms for the major barley flavonoid saponarin, however, are different in monocots and dicots. Uptake into barley vacuoles occurs via a proton antiport, while the transport into vacuoles from arabidopsis, which does not synthesise flavone glucosides, displays typical characteristics of ABC transporters (Frangne et al., 2002). For a more detailed description of the multifunctionality of vacuolar ABC transporters we refer to Martinoia et al. (2002) and Yazaki (2005). IDI7 is a half-type ABC transporter, localised to vacuoles of barley roots, which is induced by iron deficiency (Yamaguchi et al., 2002). IDI7 groups into the TAP subfamily and shows similarity to ATM3/STA1, which most likely transports metal conjugates or Fe–S clusters across the mitochondrial membrane (see Section 7.3.4.1; Figure 7.6). Thus, it is speculated that IDI7 transport may be involved in secretion of phytosiderophores from iron-deficient barley roots.

7.6.4 7.6.4.1

Ion transport Ion channels

The first ion channels described in the tonoplast of plant cells were the slowactivating (SV) and fast-activating (FV) channels (Hedrich et al., 1986; Hedrich and Neher, 1987; see also Section 6.3.2.2). The SV channel is ubiquitous in plants, is the predominant contributor to tonoplast conductance, transports mono- as well as divalent cations and is activated by cytosolic calcium. Thus, the SV channel is proposed to function in Ca2+ -induced Ca2+ release from the vacuole. The FV channel, in contrast, has a selectivity reminiscent of potassium channels (selective for monovalent cations and blocked by divalent cations) and may function in the release and uptake of K+ during cellular osmoregulation. In addition, in guard cells, a K+ -selective channel which is activated by Ca2+ has been described (Sch¨onknecht et al., 2002, and references therein). This vacuolar K+ channel (VK) most likely functions during opening and closing of stomata. The ion currents so far described as well as other conductances of the tonoplast have either been characterised electrophysiologically by patch-clamping isolated vacuoles or been characterised by fluxes of radiolabelled solutes (for overview see Barkla and Pantoja, 1996). However, the molecular nature of most of these ion channels is still unclear. TPK1/KCO1, a vacuolar potassium channel. The ‘two-pore K+ channel’ TPK1, formerly known as KCO1, in insect cells represents an outward-rectifying K+ -selective channel, which is Ca2+ dependent (Czempinski et al., 1997).

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Subsequently, TPK1 was localised to the vacuolar membrane (Czempinski et al., 2002; Sch¨onknecht et al., 2002). A decrease in SV-type currents of mesophyll cell vacuoles in a TPK1-knockout mutant suggests that TPK1 might be involved in formation of the SV conductance (Sch¨onknecht et al., 2002). However, recent studies of TPK1 conductance in yeast vacuoles (Bihler et al., 2005) and the characterisation of TPC1 (see below) indicate that TPK1 does not mediate SV-type currents but rather represents the vacuolar K+ channel described in guard cells (Figure 7.10). Ca2+ signalling Ca2+ signalling

ADP

ATP

Ca2+ H+

CA X

BCA1

C1 TP C1 TP

Ca2+

Ca2+

Salt tolerance

Ca2+

K+

TP TP K1 K1

Na+ K+

Storage

Volume regulation of vacuole + cell

X NH

H+

pH homeostasis

Mg2+, Zn2+

MHX H+

Storage

Zn2+ P MT 3 1+

Fe mobilisation under iron starvation

NR AM 3+ P 4

Fe2+, (Zn2+, Mn2+)

Zn, metal tolerance homeostasis

Figure 7.10 Ion transport across the tonoplast. Most likely the vacuolar K+ channel in guard cells is represented by a dimer of the Ca2+ -dependent, K+ -selective channel TPK1. In yeast vacuoles TPK1 shows no rectification, thus allowing K+ uptake and efflux. A possible function for TPK1 is vacuole and cell-volume regulation during opening and closing of stomata. Further TPK1 might be involved in regulation of the slow vacuolar (SV) channel TPC1. The Ca2+ -dependent Ca2+ release of the (SV) channel is mediated by TPC1. Since TPC1 (12 α-helical domains) comprises two subunits of the Shaker-like K+ channels which function as tetramers, TPC1 most likely acts as a dimer. TPC1 function is closely linked to processes regulated by cellular Ca2+ signalling. Ca2+ uptake and storage in contrast is mediated by a Ca2+ -ATPase (BCA1 in cauliflower) and by CAX, a Ca2+ /H+ exchanger (11 α-helical domains). Activity of both proteins is crucial for Ca2+ signalling. Alike for CAX, ion uptake through the exchanger NHX (Na+ ) and MHX (Mg2+ , Zn2+ ) is driven by the proton gradient across the tonoplast, which is established and maintained by the V-ATPase and V-PPase (see Figure 7.9). NRAMP proteins are capable to transport several metal cations, including Fe2+ , Zn2+ and Mn2+ . The family members NRAMP3 and NRAMP4 localise to the tonoplast and are proposed to function in iron mobilisation under iron-limiting conditions. The cation diffusion facilitators (six α-helical domains) MTP1 and MTP3 are responsible for Zn uptake into the vacuole and act during metal tolerance and homeostasis.

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Nevertheless, the K+ channel TPK1 might be involved in regulation of the SVchannel conductance in vacuoles. TPC1 represents the slow vacuolar channel. Recently, the Ca2+ -dependent Ca2+ release channel, which is known from numerous electrophysiological studies as the slow vacuolar channel (SV), has been identified and functionally characterised in arabidopsis (Peiter et al., 2005). The corresponding protein TPC1 (two-pore channel 1; Figure 7.10) comprises two subunits of the Shaker-like potassium channels (each with six α-helical domains; see Section 5.2.3.1; Very and Sentenac, 2003) and a cytosolic linker with two EF hands for Ca2+ binding. The protein is localised to the tonoplast, and deletion mutants lack a functional SV channel activity in vacuoles. Thus, TPC1 constitutes an essential component of the SV channel. Further, tpc1 mutants are defective in abscisic-acid-induced repression of germination and in the response of stomata to extracellular calcium, resembling the mutant phenotype of det3, lacking a component of the V-ATPase (see Section 7.6.1). These results demonstrate the function of vacuolar solute transport during cellular Ca2+ signalling.

7.6.4.2

Calcium, sodium and magnesium uptake involves active transport

Uptake of Ca2+ , Na+ and Mg2+ (Zn2+ ) into the vacuole is an active process (Hirschi, 2001; Maeshima, 2001), either energised by ATP or by the proton gradient established by the vacuolar proton pumps (see Section 7.6.1). The latter transport is mediated by cation/proton exchange proteins in the tonoplast membrane (Figure 7.10; Section 5.4). Calcium. The storage-type vacuole serves as a primary pool for free calcium ions in the plant cell. Ca2+ transporters thus regulate cytoplasmic Ca2+ levels and control intracellular Ca2+ signalling (see TPC1 above). Uptake of Ca2+ into the vacuole is mediated by two proteins: a Ca2+ -ATPase and a Ca2+ /H+ antiporter. High Ca2+ -ATPase activity has been reported for purified tonoplast vesicles from several plant species (Sze et al., 2000). Although several genes for Ca2+ -ATPases exist in arabidopsis, two Ca2+ -ATPases, LCA and BCA1, were isolated from vacuoles of tomato and cauliflower, respectively (Ferrol and Bennett, 1996; Malmstr¨om et al., 1997). Structure–function studies on BCA1 (Malmstr¨om et al., 2000) revealed that the N-terminus of this calmodulin-stimulated Ca2+ -ATPase provides regulatory functions, e.g. calmodulin binding, phosphorylation and autoinhibition. Together with the Ca2+ -ATPase, the Ca2+ /H+ antiporter CAX is crucial for 2+ Ca accumulation in the vacuole. Again, Ca2+ /H+ exchange across the vacuolar membrane has been measured in diverse plant species and the corresponding proteins have been isolated from arabidopsis (AtCAX1–3; Hirschi et al., 1996, 2000; Cheng et al., 2005) and mung bean (see Maeshima, 2001). CAX transporters are predicted to have 11 α-helical transmembrane domains, interruted by a central hydrophilic motif (Hirschi, 2001). CAX function was shown by heterologous expression in yeast mutants. Overexpression of AtCAX1 in tobacco and tomato increased vacuolar Ca2+ and led to phenotypes that were reminiscent of calcium deficiency in the cytosol (Hirschi, 1999; Park et al., 2005). Thus, CAX1, whose expression is induced by exogenous Ca2+ , plays a key role in Ca2+ homeostasis and/or Ca2+ signalling. Since

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CAX transporters have a low affinity but high capacity for Ca2+ , it is suggested that they lower cytosolic Ca2+ when concentrations are high (e.g. directly after external or internal stimuli). Subsequently, the Ca2+ -ATPase (high affinity, low capacity) can fine-tune cytosolic Ca2+ concentration (see Hirschi, 2001; Maeshima, 2001). This interplay was confirmed by the characterisation of cax1 and cax1/cax3 double mutants in arabidopsis which showed reduced activity of tonoplast Ca2+ /H+ exchange and V-type H+ -ATPase but increased pumping of the Ca2+ -ATPase and expression of other putative vacuolar CAX genes (Catala et al., 2003; Cheng et al., 2003, 2005). The mutant phenotypes suggest involvement of CAX function, and linkage to Ca2+ signalling, in many different processes, such as plant development, hormonal responses or cold acclimation (Catala et al., 2003; Cheng et al., 2003, 2005). CAX1 in leaves and CAX3 in roots are suggested to function synergistically in plant growth and nutrient acquisition. Interestingly, AtCAX2 shows transport capacity for Mn2+ and Cd2+ as well, and thus may function in the tolerance of plants to heavy metal ions (see Section 7.6.4.3; Hirschi et al., 2000; Hirschi, 2001). Recent research is focusing on regulation (e.g. by pH) and upon structure–function analysis of arabidopsis CAX proteins (Shigaki et al., 2003, 2005; Pittman et al., 2004, 2005). Sodium. Accumulation of excess Na+ into the vacuole is crucial during salt stress or salt tolerance of plants (see also Chapter 14). By transporting Na+ away from the cytosol, the plant cell can avert ion toxicity and also utilise Na+ as osmoticum to maintain turgor. The Na+ /H+ group of antiporters (NHX) has long attracted attention in relation to salt tolerance in plants. An arabidopsis Na+ /H+ antiporter (AtNHX1) was isolated as an orthologue to the yeast antiporter NHX1 (Apse et al., 1999; Gaxiola et al., 1999). The protein has nine transmembrane domains, three membrane-associated hydrophobic regions and a hydrophilic C-terminal domain facing the vacuolar lumen (Yamaguchi et al., 2003). Thus, the structure of NHX is different from the chloroplastidic Na+ (K+ )/H+ exchange protein CHX23 (see Section 7.2.4.2) and any other known Na+ /H+ antiporter. The localisation and function of AtNHX1 in the tonoplast have been determined by immunological methods and functional complementation in yeast. Interestingly, transgenic plants overexpressing AtNHX1 grew in the presence of 200 mM NaCl, and NHX1 mRNA is increased by high salt conditions. These observations support a role for the vacuolar Na+ /H+ antiporter in salt tolerance (but see also Chapter 14 for a critique). Studies on a T-DNA insertional mutant of AtNHX1, however, reveal that the protein also has K+ /H+ exchange activity and its contribution to ion homeostasis is crucial not only for salt tolerance but also for leaf and seedling development (Apse et al., 2003). In the flowers of Japanese morning glory, function of NHX triggers a colour change from reddish purple (buds) to blue (open flowers) by increasing the vacuolar pH (Fukada-Tanaka et al., 2000; Yamaguchi et al., 2001; Ohnishi et al., 2005). Thus, function of NHX is also crucial for vacuolar pH homeostasis. Magnesium (and zinc). Mg2+ , the most abundant divalent cation in the cytosol, is essential for the function of many enzymes (e.g. phosphatases and ATPases) and constitutes the central ion of the chlorophyll molecule. Zinc (see also Section 7.6.4.3) is also crucial for enzyme function and is involved in the regulation of gene

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expression (zinc-finger proteins). AtMHX is an arabidopsis tonoplast transporter that can exchange protons with Mg2+ and Zn2+ ions (Shaul et al., 1999). Similar to CAX, 11 α-helical transmembrane domains are predicted for MHX (Figure 7.10). When expressed in vacuoles of tobacco BY-2 cell culture, AtMHX displayed proton-activated currents of Mg2+ , Zn2+ and Fe2+ . In planta, when overexpressed in transgenic tobacco, AtMHX rendered sensitivity to excess magnesium and zinc in the growth medium. Since MHX transcripts are mainly associated with xylem elements throughout the arabidopsis plant, it is suggested that MHX functions in partitioning of Mg2+ and Zn2+ between various plant organs, by mediating transient storage in the vacuole. Recent studies are focussing on the regulation of AtMHX gene expression (David-Assael et al., 2005).

7.6.4.3

Transport of transition metals

Transition metals such as manganese (Mn), copper (Cu) or iron (Fe) fulfill crucial functions in plant cells, by acting as cofactors for proteases or superoxide dismutases and catalysing electron transfer in mitochondria and chloroplasts. Zinc (Zn) is an essential plant micronutrient and functions as cofactor for many enzymes. However, excessive accumulation of metals in the cytosol is potentially toxic, leading to inhibition of root growth, decreased photosynthesis and chlorosis. Thus, excess metal ions are often transported into the vacuole for storage. In consequence metal cation transporters are critical in maintaining cellular metal homeostasis. NRAMPs (natural resistance associated macrophage proteins) in mammals and yeast have been shown to function as transporters for multiple classes of metal cations, including Fe, Mn and Cd. Arabidopsis has six NRAMP genes, and the isoforms NRAMP3 and NRAMP4 have been localised to the vacuole (Figure 7.10; Thomine et al., 2003; Lanquar et al., 2005). Plant NRAMP proteins contain 12 αhelical transmembrane domains and are capable of transporting Fe when expressed in yeast. Expression of NRAMP3 and NRAMP4 is induced by iron deficiency, and disruption of AtNRAMP3 upon Fe starvation led to increased Mn and Zn accumulation in roots (Thomine et al., 2003). It was proposed that NRAMP3 influences metal accumulation by mobilising vacuolar metal pools to the cytosol, in particular during iron deficiency. However, single mutants did not present with a clear phenotype, because of complementation of NRAMP3 function by NRAMP4. The nramp3/nramp4 double mutant exhibited a strong defect during seed germination under low iron supply (Lanquar et al., 2005). These results point to a function in iron mobilisation from vacuolar metal stores during seed germination. The tonoplast Zn transporter (MTP/ZAT) was first identified in arabidopsis (van der Zaal et al., 1999). The ZAT protein (zinc transporter of Arabidopsis thaliana; Figure 7.10) has six putative transmembrane domains, belongs to the cation diffusion facilitator family and is similar to mammalian Zn transporters. Arabidopsis transgenic lines overexpressing ZAT exhibited enhanced accumulation of Zn in the root and showed a marked increase in resistance to high Zn concentrations in the growth media. Later on, ZAT was renamed MTP1, for metal tolerance protein 1. Kobae et al. (2004) localised AtMTP1 to the vacuolar membrane and showed that mtp1 mutant lines display enhanced sensitivity to high Zn concentrations. Orthologues of

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AtMTP1 have been implicated in the metal tolerance of hyperaccumulator plants (Desbrosses-Fonrouge et al., 2005, and references therein). All plant MTP1-like proteins complement zinc hypersensitivity of a yeast mutant, and AtMTP1 transports Zn2+ when heterologously expressed in yeast and X. laevis oocytes (DesbrossesFonrouge et al., 2005). Thus, MTP1 sequesters excess Zn into vacuoles. Only very recently, the MTP3 protein in arabidopsis was localised to the tonoplast and shown to also exhibit Zn transport capacity (Arrivault et al., 2006). Overexpression of MTP3 causes Zn accumulation and enhanced Zn tolerance in arabidopsis plants. Interestingly, iron deficiency induced MTP3 expression in roots, while a knockdown of MTP3 enhanced Zn accumulation in shoots. Thus, it is concluded that MTP3 is required for cellular metal homeostasis and excludes Zn from the shoot under Zn oversupply and iron deficiency, the latter protecting the cell from Zn2+ displacing a limited supply of Fe2+ from its binding sites.

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Van der Zaal, B.J., Neuteboom, L.W., Pinas, J.E., et al. (1999) Overexpression of a novel Arabidopsis gene related to putative zinc-transporter genes from animals can lead to enhanced zinc resistance and accumulation. Plant Physiology 119, 1047–1056. Vandecasteele, G., Szabadkai, G. and Rizzuto, R. (2001) Mitochondrial calcium homeostasis: mechanisms and molecules. IUBMB Life 52, 213–219. Vander Heiden, M.G., Chandel, N.S., Li, X.X., Schumacker, P.T., Colombini, M. and Thompson, C.B. (2000) Outer mitochondrial membrane permeability can regulate coupled respiration and cell survival. Proceedings of the National Academy of Sciences of the United States of America 97, 4666–4671. Vera-Estrella, R., Barkla, B.J., Bohnert, H.J. and Pantoja, O. (2004) Novel regulation of aquaporins during osmotic stress. Plant Physiology 135, 2318–2329. Verma, D.P.S. (2002) Peroxisome biogenesis in root nodules and assimilation of symbiotically-reduced nitrogen in tropical legumes. In: Plant Peroxisomes: Biochemistry Cell Biology and Biotechnological Applications, 1st edn (eds Baker, A. and Graham, I.A.), pp. 191–220. Kluwer Academic, Norwell, MA. Versaw, W.K. and Harrison, M.J. (2002) A chloroplast phosphate transporter, PHT2;1, influences allocation of phosphate within the plant and phosphate-starvation responses. The Plant Cell 14, 1751–1766. Very, A.A. and Sentenac, H. (2003) Molecular mechanisms and regulation of K+ transport in higher plants. Annual Review of Plant Biology 54, 575–603. Vivekananda, J. and Oliver, D.J. (1990) Detection of the monocarboxylate transporter from pea mitochondria by means of a specific monoclonal antibody. FEBS Letters 260, 220–224. Wandrey, M., Trevaskis, B., Brewin, N. and Udvardi, M.K. (2004) Molecular and cell biology of a family of voltage-dependent anion channel porins in Lotus japonicus. Plant Physiology 134, 182–193. Wang, D., Xu, Y., Li, Q., et al. (2003) Transgenic expression of a putative calcium transporter affects the time of Arabidopsis flowering. The Plant Journal 33, 285–292. Wang, X., Berkowitz, G.A. and Peters, J.S. (1993) K+ -conducting ion channel of the chloroplast inner envelope: functional reconstitution into liposomes. Proceedings of the National Academy of Sciences of the United States of America 90, 4981–4985. Watanabe, A., Nakazono, M., Tsutsumi, N. and Hirai, A. (1999) AtUCP2: a novel isoform of the mitochondrial uncoupling protein of Arabidopsis thaliana. Plant & Cell Physiology 40, 1160– 1166. Waters, M. and Pyke, K. (2005) Plastid development and differentiation. In: Plastids, Annual Plant Reviews, Vol. 13 (ed M¨oller, S.G.), pp. 30–59. Blackwell Publishing, Oxford. Weber, A., Menzlaff, E., Arbinger, B., Gutensohn, M., Eckerskorn, C. and Flugge, U.I. (1995) The 2-oxoglutarate/malate translocator of chloroplast envelope membranes: molecular cloning of a transporter containing a 12-helix motif and expression of the functional protein in yeast cells. Biochemistry 34, 2621–2627. Weber, A., Servaites, J.C., Geiger, D.R., et al. (2000) Identification, purification, and molecular cloning of a putative plastidic glucose translocator. The Plant Cell 12, 787–802. Weber, A.P. (2004) Solute transporters as connecting elements between cytosol and plastid stroma. Current Opinion in Plant Biology 7, 247–253. Weber, A.P.M., Schwacke, R. and Fl¨ugge, U.I. (2005) Solute transporters of the plastid envelope membrane. Annual Review of Plant Biology 56, 133–164. Williams, L.E. and Mills, R.F. (2005) P1B-ATPases – an ancient family of transition metal pumps with diverse functions in plants. Trends in Plant Science 10, 491–502. Woolhead, C.A., Thompson, S.J., Moore, M., et al. (2001) Distinct Albino3-dependent and -independent pathways for thylakoid membrane protein insertion. The Journal of Biological Chemistry 276, 40841–40846. Xu, C., Fan, J., Froehlich, J.E., Awai, K. and Benning, C. (2005) Mutation of the TGD1 chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. The Plant Cell 17, 3094– 3110.

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8 Ion uptake by plant roots Romola J. Davenport

8.1

Introduction

This chapter addresses the main factors affecting and controlling the uptake of charged solutes by plants, from the soil solution to the transpiration stream. It describes root anatomical and physiological responses to the availability of nutrients in the soil and the general processes involved in transport of solutes into and out of root cells. The transport proteins involved in uptake of specific solutes are not described (for these, see Chapters 5–7), but examples of uptake of various solutes are given throughout.

8.2

Soil composition

Plant roots are exposed to a heterogeneous soil environment. Soils show varying degrees of horizontal and lateral heterogeneity depending on their composition. Additionally, plants significantly modify the local soil environment, producing a rhizosphere of root influence that may extend for millimetres around the roots. The extent of plant modifications to the rhizosphere depends on the effect under consideration and on soil characteristics (Tinker and Nye, 2000; Jones et al., 2004). Plants exude sugars, amino acids and organic acids that support local microbe populations and consequently influence solute levels through consumption and decomposition. Plant secretion of acids and bases changes the rhizosphere pH and affects the solubility of, especially, phosphorus and heavy metals. Plants also influence the availability of solutes and water through uptake and efflux and alter local solute composition through selective uptake of solutes. In addition, plants release specific enzymes and chelating agents that increase uptake or reduce toxicity of solutes. The availability of nutrients to plant roots depends both on local concentrations and the solubility and mobility of solutes. The distribution of many nutrients is patchy due to the processes of soil formation, the distribution of decaying organic matter and the activities of plants and other soil organisms. These processes affect the distribution of largely insoluble nutrients such as P and Fe to a much greater extent than more soluble compounds such as monovalent cations and anions that are redistributed between patches by diffusive and bulk flow processes. The mobility of nutrients depends not only on their solubility but also on interactions with other soil components. Soils contain many fixed anionic residues (in clay matrices and the carboxyl and hydroxyl groups on organic particles) that interact with and reduce

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mobility of even very soluble cations such as K+ . By contrast, NO 3 − interacts little with the soil matrix and diffuses and leaches very easily, and despite the patchy distribution of organic sources of N, the products of nitrification are rapidly dissolved and distributed within a soil. Soil pH has dramatic effects on the solubility and mobility of many nutrients. For instance, in acidic soils P is relatively soluble and so leached readily, leading to lower total P than in alkaline soils, where P is mainly insoluble but can be available to those organisms capable of solubilising it. Soil pH also affects microbial activity and hence affects the rate of release of nutrients derived from organic decomposition.

8.3

Root exploration of the soil

Roots can respond to soil conditions at a physiological level by changing the transport activities of particular cell types (for example up-regulating nitrate transporters on detection of nitrate), and at a morphological level by altering growth patterns in order to detect and mine soil nutrients (Hutchings and John, 2004). These strategies are not exclusive. While morphological changes are usually considered more costly than changing the activity of a transporter, the returns on investment in root biomass will depend on which nutrients are growth limiting and other constraints on root development, such as water availability and the supply of photosynthates for root growth. For poorly mobile nutrients with patchy distribution, roots (or their fungal symbionts) need to explore the soil to detect and exploit localised sources. For instance, upon detection of P-rich areas, roots often show intense branching and ramification and/or extensive root hair development (Figure 8.1; Drew, 1975). Ramification may be crucial in enhancing uptake of insoluble P and Fe compounds where the root cannot rely on diffusion for delivery but must secrete solubilising agents at very close range (e.g. the cluster roots of the Proteacea; see also Section 12.3). In this case the production of fine and branching roots serves to increase surface area for uptake and reduce diffusion distance. This is also why mycorrhizal symbioses are particularly important for uptake of P and also relatively insoluble micronutrient metals. However, roots also show a similar growth response to NO 3 − , which in soil would be released over time by decomposition and be distributed relatively rapidly to nearby roots (Figure 8.1). In this case the ramification of roots into these regions may serve more to secure the pocket against competition than to mine the area per se (Robinson, 1996; Hodge, 2004). There is further discussion of nutrient acquisition and mineral deficiency in Chapters 11 and 12. Fungal associations are key in increasing root exploration of soil in the majority of plant species, although importantly not in the family Brassicaceae to which the model plant Arabidopsis thaliana belongs. Fungal hyphae are single celled in thickness and so can ramify through soil on a finer scale and at a fraction of the biomass cost of thicker, multi-cellular roots. Fungi also possess highly effective mechanisms for solubilisation of soil minerals (via high levels of secretion of enzymes and chelating and solubilising agents; Landeweert et al., 2001), and many decompose organic matter directly. They may also compete effectively with other

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Figure 8.1 Root responses to localised supply of nutrients. Barley plants were grown in sand and supplied with a full-nutrient solution (regions marked ‘H’) or with a solution lacking the specified nutrient (regions marked ‘L’). Reproduced from Drew (1975), Figure 4, with permission of Oxford University Press.

soil microbes for nutrients that are then exchanged with the plant host for photosynthates. Mycorrhizae have been demonstrated to enhance plant acquisition of P, micronutrients and N (Hodge, 2005). Plants may also foster local fixation of atmospheric N through highly specialised symbioses (the most common being the legume–Rhizobia complex) and through looser mutualisms involving exudation of photosynthates that encourage N fixation by free-living soil bacteria. In some cases, roots may be modified to provide microhabitats for free-living N-fixing bacteria (Baldani et al., 1997). Plant exudates can also increase local detritivorous microbial activity that results in faster release of inorganic nutrients via decomposition of organic materials (James, 2000). Much of the research on plant nutrition is conducted in young plants grown in nutrient cultures in the absence or strict control of soil microbes (McCully, 1999). Such conditions make it possible to tease out the environmental determinants and

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genetic factors involved in, particularly, the physiological responses of plant roots to abiotic conditions, which are most amenable to reductionist investigation. There is, however, a relative deficit of understanding the complexities of root development and interactions with soil microbes under realistic soil conditions, due to the difficulties of isolating causal factors amidst complexity and of accessing complex systems without perturbing them. An understanding of root behaviour in real soils is crucial when attempting to transfer plant traits observed or engineered in laboratory conditions to the field, where the observed trait may be irrelevant or expressed very differently.

8.4

Physical factors affecting root uptake: depletion zones and Donnan potentials

Nutrients are delivered to roots via diffusive and mass flow processes. Solutes diffuse towards the roots down concentration gradients established by root uptake. Uptake can result in zones of depletion around roots, where diffusion is not rapid enough to replace absorbed solutes. In this case, root cells experience lower solute concentrations than measured in the bulk solution and the kinetics of uptake are affected. Solutes are also delivered to root cells via mass flow of water drawn into the roots by the transpiration stream. The extent to which this occurs depends on the rate of transpiration and the dryness of the soil, and will vary diurnally. Where mass flow occurs, nutrients will be supplied to the root regardless of uptake rates of specific solutes and depletion zones are less likely to develop. In the case of simple diffusion, it is possible that uptake of scarce solutes is restricted to epidermal cells (especially root hairs), where the diffusive pathway from the bulk soil solution is shortest. In this case cortical cells will be exposed to boundary layers depleted by epidermal cell activity and may contribute relatively little to uptake (although they are likely to participate in re-uptake of effluxed solutes; see Section 8.5.2). However if mass flow is significant, then solutes will be carried into the cortical apoplast and cortical and endodermal cells may participate in uptake of even scarce nutrients, often greatly increasing the surface area available for nutrient absorption. Since the absorption of water and solutes occurs independently, it is possible for mass flow to cause the accumulation of solutes at exo- or endodermal barriers to concentrations exceeding those of the bulk solution. In practice, this is unlikely to occur except in cases such as relatively dry saline soils where mass flow may be significant and root NaCl or Na 2 SO 4 uptake rates are low compared to the concentration of salt in the soil solution (Tinker and Nye, 2000). Preferential uptake of particular nutrients and the development of depletion zones can cause the solute composition of the root apoplast to differ significantly from the bulk solution. Additionally, the electrochemical properties of plant cell walls may affect the ionic composition of the solution to which root cells are exposed. Cellwall components contain a high density of fixed, mainly anionic charges. Cell-wall residues preferentially bind particular ions (especially H+ and Ca2+ ). In addition,

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the net negative charge of cell walls creates a Donnan potential (especially in dilute conditions) that attracts cations and repels anions in a valency-dependent manner. The activities and ratios of ions are therefore quite different in cell walls compared with the bulk solution (see Section 3.6.5). Cell membranes also carry a net negative charge due to phosphate groups on phospholipids, but it is not known to what extent the Donnan potentials generated by cell walls and membranes affect the ionic composition of the solution to which membrane transporters are exposed. It is possible that measures of transport rates and ion selectivity may reflect the responses of transport proteins to very different ionic conditions to those assumed on the basis of the composition of the bulk solution (Miedema, 2002; Kinraide, 2004).

8.5 8.5.1

Radial transport of solutes across the outer part of the root The role of apoplastic barriers

The Casparian strip of the endodermis forms a suberised barrier to apoplastic movement of water and solutes (see below for exceptions). Thus while solutes (and water) can move either symplastically or apoplastically up to the endodermal barrier, they can usually only cross that barrier in the symplast (Figure 8.2). Therefore solutes must enter the symplast via cells in the outer part of the root, across cell membranes outside the Casparian strip of the endodermis. In most roots this uptake zone comprises the epidermis, cortex and outer face of endodermal cells (where this is unsuberised – see Section 8.6). In addition, many species have a hypodermis, usually immediately under the epidermal layer, which may provide an initial barrier to movement of at least some solutes (Hose et al., 2001). Since most cells are connected by plasmodesmata, the symplast forms a continuum linking the cortical and

apoplastic pathway transcellular pathway symplastic pathway

epidermis

cortex

endodermis

stele

xylem

Casparian strip

Figure 8.2 Pathways of solute and water movement from the soil to the root stele. Charged solutes can move by diffusion or bulk flow through the apoplast of the outer part of the root but are prevented from entering the stele by the hydrophobic barrier of the Casparian strip (or more extensive suberisation of the endodermis in older parts of the root). To enter the stele, solutes must enter the symplast at some point outside the Casparian strip. Solutes may move from cell to cell via plasmodesmata (the symplastic pathway), or move into and out of cells via the apoplast (the trans-cellular pathway).

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stelar parts of the root, and therefore solute uptake may require only a single uptake step at any point in the outer part of the root. In some species it appears that radial transport across the root occurs via both symplastic and apoplastic pathways, due to an incomplete endodermal barrier. In rice, the uptake of Na+ from high concentrations is proportional to the uptake of the dye PTS, which is membrane impermeant and moves apoplastically, suggesting that Na+ uptake occurs mainly via an apoplastic route (Yeo et al., 1987; Yadav et al., 1996). The authors propose that the endodermal leak is a small proportion of the total volume flow. Consequently, at the low concentrations at which most solutes are present in the soil the symplastic pathway will dominate. However for solutes present at high concentrations, as in the case of NaCl in saline conditions, the low permeability of the endodermis will nevertheless permit a significant apoplastic leak of NaCl to the shoot, with toxic consequences. Peterson and Moon (1993) have proposed that exo- and possibly endodermal leaks may occur when lateral roots initiate. Lateral roots develop from the pericycle cell layer to the inside of the endodermis, and the developing lateral root forces its way through the endodermis and outer cell layers to emerge. This transient rupture of the endodermis may permit apoplastic leakage of solutes to the shoot. Recently, White and colleagues have presented controversial evidence for apoplastic transport of divalent cations to the shoot in arabidopsis and in the Zn hyperaccumulator Thlaspi caerulescens (White, 2001; Ernst et al., 2002; White et al., 2002). It is possible that endodermal leaks occur but only contribute significantly to shoot uptake in the case of ions that are present at high soil concentrations or have very low symplastic permeability.

8.5.2

Root hairs and cortical cells

Root hairs are specialised for nutrient acquisition. They are modified epidermal cells that provide a high surface area for uptake and increase root exploration of the soil by penetrating the soil surrounding the root, reducing the diffusive distance from soil to root. Where solutes are present in limiting amounts, it is possible that uptake by root hairs depletes the soil solution moving into the root to the extent that cortical cells are unable to extract any remaining solutes and therefore play no role in uptake. Evidence for the importance of root hairs in absorption of particular solutes includes increases in root hair length and/or density under conditions of particular nutrient deprivation, and differences in nutrient acquisition rates or growth on low nutrient levels in genotypes differing in root hair length or abundance. Root hairs seem to be particularly important in accessing immobile nutrients and show especially extensive proliferation and elongation under conditions of P deficiency (Forde and Lorenzo, 2001). Efficiency of P uptake has been shown to correlate with root hair length and/or density within species (e.g. Gahoonia et al., 1997; Narang et al., 2000), although there is also evidence for a variety of strategies that may substitute for root hair responses – for instance increased root length and biomass and increased mycorrhizal colonisation (e.g. Wissuwa and Ae, 2001; Smith et al., 2003). The role of root hairs in uptake of more mobile nutrients is probably less critical than for less mobile elements and shows more interspecific variation. For instance, low

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NO3 − supply stimulates root hair growth in a number of species, but had no effect in tomato and some grasses (Forde and Lorenzo, 2001). Limiting levels of Fe may stimulate root hair elongation and proliferation in species that secrete solubilising and reducing agents (so-called ‘strategy I’), but not in the Poaceae that secrete phytosiderophores to complex Fe (‘strategy II’) (Section 12.2.2; Schmidt, 1999). The role of root hairs in nutrient uptake has also been demonstrated using mutants in which root hair development is aberrant or absent. However, evidence for poor growth or low uptake of nutrients in these mutants should be treated with caution because in some cases the mutations affecting root hair development may also affect processes important in nutrient sensing or uptake per se, with concomitant effects on root hairs. For example, the arabidopsis root-hairless rhd2 mutant is defective in expression of an NADPH oxidase involved in a signal transduction pathway that increases Ca2+ uptake to trigger root hair cell expansion under nutrient limitation, and may also affect the activity or expression of other solute transporters (Foreman et al., 2003; Shin et al., 2005). Other types of evidence for the importance of root hairs include experiments measuring uptake under conditions where it should be confined to epidermal cells (including very brief exposure to labelled solute or plasmolysis); these suggest that epidermal uptake accounts for the bulk of K+ uptake (Clarkson, 1996). The role of cortical cells in initial uptake of nutrients may depend on the extent to which root-hair uptake either satisfies plant needs or depletes the apoplastic concentrations to which cortical cells are exposed. If plant requirements are low compared to the local availability of a given nutrient, then cortical cells may contribute significantly to uptake, particularly in species where the cortex comprises many cell layers and so provides a high surface area relative to that of epidermal cells. Conversely, cortical cells may contribute little to initial uptake at low external concentrations of solutes. However, if uptake is confined mainly to root hairs under conditions of nutrient limitation, then it might be expected that the expression of high-affinity uptake mechanisms would occur specifically in root hairs. For some solutes, at least, this does not appear to be the case. For example, expression of high-affinity K+ (Ashley et al., 2006) and sulphur (Maruyama-Nakashita et al., 2004) transporters is responsive to low nutrient availability and not confined to root hairs, but includes cortical cells as well. In contrast, Fe transport, which requires secretion of solubilising or chelating agents, may be confined mainly to the root periphery (Bauer and Bereczky, 2003). High-affinity P influx mechanisms may also be confined to the epidermis (in the absence of mycorrhizal infection) (Daram et al., 1998; Chiou et al., 2001), whereas there is evidence for both root-hair-specific and cortical cell expression of high-affinity transporters for NH 4 + and NO 3 − (Miller and Cramer, 2004). A key caveat to these expression studies is that a definite role for the expressed proteins in high-affinity nutrient uptake under low nutrient-availability conditions has not been demonstrated in most cases, so expression patterns could reflect other roles in nutrient transport apart from uptake. If cortical cells participate in high-affinity uptake from the apoplast under conditions of nutrient limitation, then this suggests that depletion-zone effects do not confine uptake to root hair cells. Alternatively, it may indicate that solutes that enter

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the symplast initially via root hairs do not move wholly symplastically to the stele but exit and re-enter the symplast multiple times before crossing the endodermis. This so-called ‘transcellular’ pathway (Figure 8.2) has generally been considered too energetically expensive to be plausible, especially in the case of anions (Clarkson, 1993). However, it is nevertheless possible that many solutes reach the stele via some combination of the symplastic pathway through plasmodesmata and symplastapoplast-symplast transfers. Roots show high rates of efflux of many solutes (see Section 8.8.1), indicating that membrane transport of most solutes is bidirectional (except in the case of solutes that change chemical form within the cytosol; for instance any Al3+ entering the cytosol is likely to precipitate). Depending on the solute, this efflux may be specific or may occur due to the imperfect selectivity of membrane transporters and the inherent bidirectionality of most ion channels. Nonselective cation channels that transport both monovalent and divalent cations are ubiquitous in plant plasma membranes, and the plant anion channels characterised so far are permeable to a range of anionic solutes. Thus solutes may be leaked from the symplast and then ‘mopped up’ from the apoplast by cortical cells. Additionally, there is evidence in arabidopsis that mature root hairs are only weakly symplastically connected to the rest of the root symplast, and solutes taken into root hairs may require unloading and re-uptake by cortical cells for onward transport to the shoot. The evidence for symplastic isolation of mature root hairs is derived from several sources. Studies of dye movement indicate the progressive symplastic isolation of epidermal cells as they mature, although the dye molecules are large and may therefore report a narrowing rather than complete closure of plasmodesmata (Duckett et al., 1994). However, electrophysiological studies of the electrical continuity between root hairs and adjacent cells found low electrical conductivity between mature root hairs and adjacent epidermal cells, indicating that the movement of even small ions from root hairs to other epidermal cells (cortical cells were not tested) was restricted (Lew, 1994; Meharg et al., 1994). There is also evidence that, in some species at least, plasmodesmatal connections between epidermal and cortical cells are few relative to, for instance, connections between cortical and endodermal cells (Ma and Peterson, 2001). These data are rather surprising, since an apoplastic step between root hairs and uptake into the rest of the root symplast seems wasteful particularly in the case of limiting nutrients actively acquired. However, it is possible that this additional complexity gives the plant greater opportunity to control solute acquisition, and this is worth the additional metabolic expense and loss of solute involved. In addition to the simple division of cell types already described in the outer part of the root, some species have cell types specialised for nutrient uptake. In particular, Fe deficiency elicits in many species the differentiation of epidermal cells into transfer cells that are highly invaginated and provide a high surface area for solute exchange (Schmidt, 1999). Passage cells, in which only the Casparian strip develops, may punctuate the barriers of the exo- and endodermis once the majority of cells in these layers become fully suberised (Section 8.6). These passage cells may play a variety of roles, including membrane uptake of solutes across suberised exoand endodermal barriers (especially for uptake of Ca, or solute absorption across

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the exodermis when the epidermis has died) and may constitute a low-resistance pathway for water movement (Peterson and Enstone, 1996).

8.6

Solute uptake from different root zones

In addition to differences in cell function across the radius of the root, there is variation in uptake along the length of the root. The root apex is specialised for growth and differentiation and shows only low uptake rates, deriving most of its nutrition from the phloem. The apex is thought to be relatively impermeable to apoplastic movement of solutes (Enstone and Peterson, 1992), and the xylem is not functional and therefore delivery to the shoot is minimal from this zone. Behind the elongation zone is a zone of epidermal root hair development. Root hairs may persist in older parts of the root in some species, but are usually concentrated in a zone behind the apex. As the root grows from the tip, new root hairs develop to maximise uptake from un-mined regions of soil. The mature parts of the root are often considered not to be involved in solute uptake, because they exist in regions of already mined soil. Indeed, the outer part of the root may be sloughed off in some species in older parts of the roots (Wenzel and McCully, 1991). However, the extent to which older parts of the roots participate in solute uptake depends on the solute and the species, and on the pattern of root development in the growth conditions used. P and Fe are relatively immobile solutes and therefore rapidly and semi-permanently depleted from the surrounding soil, whereas mobile solutes may be replaced by diffusion and therefore remain available to older parts of the root. Uptake of Fe may be confined mainly to the root apex in species that secrete siderophores, and to the root hair zone in species that secrete solubilising agents (Clarkson, 1996; Bauer and Bereczky, 2003), whereas uptake of K+ , Na+ and NO 3 − is more evenly distributed along the root, at least at non-limiting concentrations (Lazof et al., 1992; Clarkson, 1996). Most of the evidence for P uptake indicates that uptake occurs along the length of the root, despite the problem of P depletion of the rhizosphere of older roots (Clarkson, 1996; Rubio et al., 2004). These findings may reflect growth conditions used. For instance, when roots are P-starved older roots may be employed to scavenge P remaining in already depleted soil when the young roots are unable to supply adequate P. Similarly, plants grown in uniform hydroponic conditions (where depletion zones will not develop) are less likely to restrict uptake to younger root zones. Ca2+ , although often available in millimolar amounts and relatively soluble, is taken up in younger parts of the root only (Clarkson, 1996). The most plausible explanation for this was proposed by Clarkson (1984), who considered that the toxicity and consequently very low activity of Ca2+ in the cytosol made the symplast an unlikely route for uptake of Ca2+ in the millimolar amounts required for its role as a macronutrient in cell walls. He proposed that Ca2+ travelled from the soil to the shoot in the apoplast, with a single symplastic step through the endodermal cells themselves. Thus the outer face of the endodermal cell membrane would be specialised for Ca2+ influx, and the inner face for Ca2+ efflux. However in many species,

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the Casparian strip exists as a single band of suberisation only in the younger part of the root and expands in mature roots to cover the endodermal cell membrane completely, preventing membrane transport by endodermal cells and restricting uptake of solutes to cells outside the endodermis. The uptake of Ca2+ across endodermal cells would therefore be restricted to those parts of the root where only the Casparian strip was present. The longitudinal pattern of uptake of solutes for translocation to the shoot may also depend on the developmental state of xylem cells. There is some controversy over the state of xylem development in young roots. In some species grown under field conditions, the late metaxylem vessels, which constitute the major highconductance pathway for water uptake when dead, have been shown to remain alive (and therefore high-resistance) throughout the root-hair zone and up to 30–50 m from the root apex (Wenzel et al., 1989; McCully, 1999). It is possible that the relatively low conductivity of these regions of the root could limit nutrient transfer to the shoot, and it has been suggested that solutes absorbed in these regions are stored until xylem maturation allows efficient translocation, creating a delay between nutrient absorption and translocation. Immature metaxylem vessels have been shown to contain very high levels of K+ in particular, and McCully and Canny (1988) estimated that perhaps 10% of K+ transported to the shoot derived from the release of this stored K+ during the programmed cell death of these xylem vessels in maize. However, where early metaxylem vessels (with lower conductivity than late metaxylem) are dead and open (which may occur from 4–10 cm from the root apex; McCully, 1999), there is little evidence for inadequate water conductivity and therefore it is likely that these sections of the young root could supply relatively high volumes of solutes despite the high resistance of the late metaxylem elements (Steudle and Peterson, 1998). The contribution of different sections of the root to transport to the shoot is usually measured using a compartmental labelling system, where specific sections of the root are exposed to radioactively labelled nutrient solution and the appearance of radioactivity in the shoot is measured and expressed per unit of exposed root. The plants used for these experiments are often grown hydroponically, and it is possible that xylem maturation differs under these conditions from that of plants grown in soil and exposed to fluctuating water potentials and thus these experiments may not reflect the longitudinal pattern of solute uptake in field-grown roots. Although roots are often modelled as single linear units in physiological descriptions of transport processes, roots are highly branched and may also consist of different root types with different transport properties. In seedlings, the basal roots will contribute extensively to solute uptake and it is possible to distinguish easily between young and mature root regions. However, as lateral roots and new basal and adventitious roots develop it becomes necessary, and also much more technically difficult, to determine the contributions of the different root types. In clonal plants as well as some tree species, different roots supply different parts of the shoot (a phenomenon known as ‘sectoriality’) and thus patchiness in soil nutrients can result in different nutrient status of different parts of the shoot (e.g. Orians et al., 2002) and may also promote differences in uptake patterns between roots depending on differences in shoot signals.

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8.7

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Transport of solutes to the xylem

Long-distance transport of solutes to the shoot occurs via the transpiration stream in mature (dead) xylem (see also Section 9.3). To reach the shoot, solutes must cross the endodermal barrier via the symplast, in species where the endodermis is intact. In root zones where the xylem vessels and tracheids are mature (i.e. dead), solutes must be unloaded from the symplast into the apoplast surrounding and including the xylem. This unloading step has been shown to be independent of initial uptake into the root for at least some solutes (Pitman et al., 1977; Clarkson, 1993) and represents a point of control of solute transfer to the shoot. The most obvious point for unloading of solutes from the symplast is from the xylem parenchyma cells immediately adjacent to the xylem vessels, and these cells often show characteristics of transfer cells, being invaginated and highly metabolically active (De Boer and Volkov, 2003). However, in some species the plasmodesmatal connections between xylem parenchyma cells and the rest of the symplast are relatively few, suggesting that there may be substantial unloading from the symplast before the xylem parenchyma (Ma and Peterson, 2001). In this case, unloading may occur from pericycle cells, which are highly connected to the adjacent endodermis and often show high transport capacity (Vakhimistrov et al., 1972; Ma and Peterson, 2001). When there is bulk flow of water through the stele, unloaded solutes would be carried into the xylem fluid from any part of the stele. The mechanism of unloading of solutes into the xylem differs between solutes and may also differ for a given solute along the length of the root and according to transpiration rates. For instance, whether xylem loading of a solute requires active transport will depend on the chemical or electrochemical gradient for its release. This gradient may vary along the length of the root, as release of solutes along the root increases the xylem concentration relative to the cytosolic concentration. Moreover, when transpiration rates are high, xylem contents may be relatively dilute compared to low transpiration conditions, favouring passive release. Measures of xylem osmolarity and composition are usually made from cut stems and hence provide little information on variations in xylem content along the length of the root. For ions, the situation is further complicated by the possibility of variations in the potential difference between the xylem and the cytosol of xylem parenchyma cells. Measurements of the potential difference between the xylem and the bulk solution indicate that the ‘trans-root potential’ is in the range of +30 mV to −90 mV, and sensitive to changes in exposure of the shoot to light (Wegner and Zimmermann, 2002; De Boer and Volkov, 2003). Stelar cells may also regulate their membrane voltage according to physiological conditions, including drought (Roberts and Snowman, 2000), possibly giving the plant flexibility with respect to the mechanisms deployed for xylem loading. There is some controversy over whether plants rely on transpiration for solute delivery to the shoot (Smith, 1991; Tanner and Beevers, 2001). There is evidence that the solute content of the xylem varies inversely with the transpiration rate, suggesting either that solute loading rates into the xylem remain the same, and solutes are progressively diluted by increasing water flow at higher transpiration rates, or the roots adjust xylem loading to maintain constant rates of delivery to

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the shoot (Smith, 1991). Split-root experiments, where the shoot solute content is manipulated independently of root content, indicate that shoot nutritional requirements can dictate root uptake and translocation rates. The signals that regulate xylem loading remain mysterious but must include signals from both the root and shoot. K+ translocation is probably best characterised, and will be used as an example. In plants grown under a range of K+ concentrations, root K+ uptake is inversely proportional to root K+ content (e.g. Siddiqi and Glass, 1987). However, split root experiments indicate that K+ -replete roots do not show the normal reduction in K+ uptake but continue to absorb and translocate high levels of K+ when the shoot is deficient (Marschner, 1995). Unloading of K+ into the stele is usually presumed to occur via stelar parenchyma cells, and electrophysiological analyses indicate a high frequency of outward-rectifying K+ channels (KOR) that would permit K+ efflux (De Boer and Volkov, 2003). The shoot signals that transmit information to the root are unknown but may be the phloem concentration or rate of delivery of K+ , although a correlation between shoot tissue K+ and phloem K+ concentration has only sometimes been demonstrated (White, 1997). Root signals also contribute to control of K+ translocation. When maize roots detect drought signals (ABA), K+ uptake into roots is maintained but translocation is inhibited (Roberts, 1998), presumably allowing the root to reduce osmotic potential and maintain water uptake from drying soils. The inhibition of translocation is reflected in the decline of KOR activity in protoplasts from stelar parenchyma cells in maize and in down regulation of transcript levels of the stelar K+ -selective outward-rectifying channel, SKOR, in arabidopsis (Gaymard et al., 1998; see also Sections 6.2.1 and 9.3.2.1). There is also evidence, for some solutes, of re-uptake into stelar cells lining the xylem in the older parts of the root. This re-uptake could be for storage or for redistribution of solutes from the actively growing regions of the root to older parts. In the case of toxic solutes, it may serve as a mechanism to enhance root extrusion and reduce transfer to the shoot (Tester and Davenport, 2003). As a consequence of the bidirectional exchange of solutes between the xylem fluid and xylem parenchyma cells, caution must be exercised in determining whether transporter expression or transport activity, measured in xylem parenchyma cells, represent xylem loading mechanisms.

8.8 8.8.1

The kinetics of solute uptake into roots Radioisotopic studies

The net uptake of most solutes into the plant body is not a unidirectional process but the sum of bidirectional movements of solutes across root plasma membranes. A significant proportion of initial influx may be effluxed – up to 95% of Na+ (e.g. Davenport et al., 2005), 75% of NH 4 + (Britto et al., 2001) and 20–50% of NO 3 − for example (Aslam et al., 1996; Scheurwater et al., 1999). Therefore, measurements of changes in the content of plant tissues over time (between successive harvests) can

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only give estimates of net uptake and produce little information about the transport processes involved in initial uptake. To measure unidirectional uptake, it is necessary to measure uptake over short intervals using a marker that can be distinguished from the solutes already in the plant. Radioisotopes of the solute of interest are the most useful markers to use, because they are chemically almost identical to the solute under study but can be readily distinguished from it. Plants can be grown under steady-state conditions, then exposed to a nutrient solution radioactively labelled but otherwise identical to the normal growth medium, and uptake followed over time in intact, transpiring plants or in excised segments (see also Section 4.4.3). The interpretation of the time course of isotope uptake depends on the assumptions made about the structure of the root. Classical models of isotope uptake assume a three-compartment system, where initial uptake into the first compartment (the cytosol of cells in the outer part of the root) is the fastest process and indicated by an initial rapid linear phase of isotope accumulation (Figure 8.3; Walker and Pitman, 1976). The accumulation is linear initially because although there is bidirectional transport of the solute across the plasma membrane, the proportion of the solute A root total

10

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Figure 8.3 Solute uptake in a three-compartment root system model (root cytosol, root vacuole and shoot). The root total and shoot isotope content can be measured, and the root cytosolic and vacuolar contents estimated from the model. Root uptake shows an initial rapid phase of accumulation attributable to labelling of the root cytosol (A). As the cytosol fills, the rate of uptake slows (as efflux of isotope begins to occur) and then the cytosol becomes fully labelled and root uptake becomes dominated by the slower influx to the vacuole. Once the vacuole is fully labelled, the root total ceases to change (B). Shoot uptake occurs from the cytosol, and the appearance of isotope in the shoot occurs with a lag corresponding to the time taken for isotope to accumulate in the root cytosol. Shoot uptake continues without saturation (B), unless there is recirculation of isotope from the shoot via the phloem.

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in the cytosol that is radioactively labelled is low and therefore there is negligible efflux of the isotope. As levels of isotope build up in the cytosol, efflux of the isotope becomes significant and the rate of accumulation begins to slow and assume a second linear rate. This represents the rate of influx of the isotope into the vacuole. This rate is usually assumed to be slower than initial influx – if it is the same or faster, then it cannot be distinguished from initial influx. In this case, uptake will be described by a single linear time course until the root vacuoles become labelled to the extent that there is significant efflux from the vacuole. The third compartment represents the shoot. Labelled solutes generally appear in the shoot with a lag that corresponds to the time required for isotope to accumulate in the compartment from which xylem loading occurs and to accumulate sufficiently to reach a point where a measurable proportion of the solute flux to the shoot is radioactive. Once the root isotope content has equilibrated with the external medium, the solute uptake to the shoot should become linear (although in practice, this may not occur given diurnal fluctuations in transpiration and xylem loading). This rate to estimate the absolute rate of solute translocation (in this case, the net sum of xylem loading and re-uptake). This model assumes that filling of the vacuole and shoot occurs in parallel; decisions about the number of compartments and their arrangement will depend on the complexity of the pattern of observed uptake. Time courses of solute uptake frequently approximate the kinetics predicted by a three-compartment model and can be used to derive measures of unidirectional influx, vacuolar influx and net translocation to the shoot, as well as indirect estimates of efflux rates (Walker and Pitman, 1976; Davenport et al., 2005). However even where the kinetics of uptake appear simple, it is likely that they represent the sum of a much larger heterogeneous set of compartments and transport rates, given the morphological complexity of most roots. Nevertheless simple estimates of influx and other transport rates provide a basis for the comparison of effects of growth conditions, solute concentrations and other factors affecting root transport. Radioisotopic methods can also be used to estimate the rates of efflux from tissues, the contributions of different root sections to uptake, storage and translocation, and to separate the processes of xylem loading and unloading. Efflux rates can be measured by loading tissues with radioactively labelled solute and then washing the tissues in successive rinses of unlabelled solution and counting the radioactivity in the wash (or in the tissue if some portion is harvested at each rinse). The loss of isotope from the tissue must be modelled making assumptions about the number and arrangement of compartments releasing isotope to give rates of efflux from each compartment (MacRobbie, 1981; see Section 2.6.6). Split-root experiments allow the solute content of the root system and shoot to be manipulated independently to separate feedback (of local root content upon root influx) from shoot signals affecting root uptake (e.g. Drew and Saker, 1984). The technique of exposing only sections of the root to isotope (e.g. Cholewa and Peterson, 2004; see also Section 8.6) can be used to dissect the contribution of different sections of the root system to solute translocation to the shoot. This method can also be used to estimate reuptake into mature parts of the root by measuring labelled solute appearing in the upper parts of the root not exposed to labelled solution. While some of the label in

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the mature part of the root will represent exchange with binding sites in the xylem apoplast, comparison of the amount of label in mature root sections under different conditions of nutrient supply, or between genotypes, can indicate factors affecting withdrawal of solutes from the xylem into stelar cells. One problem with using radioisotopes to measure unidirectional fluxes across membranes is the presence of the cell wall, which has a high binding capacity particularly for multivalent cations. For anions and monovalent cations, a brief rinse of the tissue in a non-radioactive solution usually suffices to displace apoplastically bound isotope. However, for multivalent cations much longer rinses are required and even then the displacement of bound isotope is usually incomplete. Rates of uptake of multivalent cations into root cells are usually low and therefore cannot necessarily be distinguished from residual cell-wall binding. Significant efflux of isotope may also occur during prolonged rinsing, leading to an underestimate of cellular uptake. For these reasons it is not possible to measure directly the influx of multivalent cations into plant roots, and only transfer to the shoot can be reliably estimated (Reid and Smith, 1992).

8.8.2

Other methods

Net root uptake and translocation processes can be studied using methods that measure net flows of solutes. Ion-selective microelectrodes (Section 2.6.3) can be used externally to measure net fluxes between roots and the external medium (Section 4.4.5.1). Net fluxes across cell membranes can be measured using voltage-recording intracellular microelectrodes to evaluate changes in voltage or conductance, and changes in internal activities of solutes can be measured using intracellular ionselective microelectrodes, ion-sensitive dyes or proteins and X-ray microanalysis (see Chapter 2). Net fluxes to the root xylem can be estimated by changes in xylemfluid composition measured using X-ray microanalysis, ion-sensitive microelectrodes, xylem feeding insects and extrusion from pressurised cut roots (see Section 9.3.3). The transport capacities of different cell types can be estimated from electrophysiological studies of isolated protoplasts, although these must then be contextualised with measures of membrane potentials in situ and estimates of the solute concentrations to which the cells are normally exposed. The roles of particular transport proteins or cellular features can be investigated by use of specific inhibitors and manipulation of gene expression, with the caveat that gene misexpression may have pleiotropic effects (see Chapters 2–4 for detail descriptions of these techniques).

8.8.3

Kinetics of uptake in response to solute availability

In addition to morphological responses to availability of nutrients, roots also respond by changing the expression and activity of transport mechanisms for solute uptake. These physiological responses have been studied mainly in the absence of complex soil environments and hence do not necessarily reflect the pattern of transport activity in the presence, for instance, of mycorrhizal infections. Additionally, only a relatively small number of species have been studied, and it may be inappropriate to extrapolate from these to other species.

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The kinetics of root uptake have been most intensively studied in the case of K+ . Initial characterisation of K+ uptake in barley at different K+ concentrations (using 86 Rb+ , an isotope with a longer half-life than 42 K+ ) indicated a ‘dual isotherm’ pattern composed of two saturating Michaelis–Menton type curves (Epstein et al., 1963). Uptake increased rapidly with increasing K+ and saturated around 0.2 mM K+ , with a second saturating isotherm of uptake between 0.5 and 20 mM K+ . A similar pattern was found for Na+ uptake in barley (Rains and Epstein, 1967a). The two components of uptake were attributed to separate high- and low-affinity mechanisms of uptake. Closer characterisation of the low-affinity component of K+ uptake indicated that it did not comprise a single curve and the authors suggested that this might reflect the contributions of multiple carrier sites of differing affinity (Rains and Epstein, 1967b), while other researchers suggested that dual isotherms and more complex kinetics would be predicted given the complexity of plant roots and changes in diffusion rates and membrane potential at different K+ concentrations (Cheeseman, 1982; Kochian and Lucas, 1982). Later measurements of K+ (Rb+ ) uptake in maize produced a different kinetic pattern, with saturating high-affinity uptake at low K+ and a linear phase of uptake in the range 1–50 mM K+ (Kochian and Lucas, 1982). This pattern was attributed to the activity of two uptake systems: a high-affinity, low-capacity transporter responsible for the bulk of uptake at low K+ concentrations and a low-affinity, high-capacity transporter, assumed to be an ion channel, which dominated uptake in the higher concentration range (Kochian et al., 1985). Similar patterns of uptake have been observed for a range of solutes, including Na+ in wheat (Davenport, 1998), NH 4 + in rice (Wang et al., 1993a,b; 1994) and NO 3 − in barley (Siddiqi et al., 1990). Dual isotherms of saturating influx have been observed in other cases, including K+ in K+ -starved arabidopsis (Gierth et al., 2005) and PO 4 − in maize (Sentenac and Grignon, 1985). An important point when considering kinetic studies in different species is whether the uptake measured was unidirectional influx or net uptake – only the former gives information about the transport mechanism for uptake. Despite the apparent simplicity of the kinetics of root uptake, characterisation of solute transporters and mutants lacking expression of these transporters has indicated that uptake of K+ , NH 4 + and NO 3 − at least is effected by a range of different transporters of differing affinities and induction patterns. In arabidopsis and wheat, it appears that both ion channels and H+ -coupled symporters contribute to highaffinity uptake even from low micromolar concentrations of K+ (Hirsch et al., 1998). Some K+ transporters are capable of both passive and active transport and may switch from high-affinity transport to low-affinity, high-capacity transport at higher K+ concentrations (so-called ‘dual-affinity’ transporters; Fu and Luan, 1998). Similarly, NH 4 + uptake is active and H+ -coupled at low external concentrations but may be mediated by ion channels, including K+ -selective and cation non-selective channels, at higher NH 4 + levels (Kronzucker et al., 2001). Uptake of anions is usually energetically more expensive than uptake of cations, where the electrical gradient across the plasma membrane favours uptake, and uptake of NO 3 − appears to be active and H+ -coupled even at 20 mM external NO 3 − (Glass et al., 1992). Nevertheless both high- and low-affinity NO 3 − transporters have been identified,

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as well at least one showing ‘dual affinity’ behaviour (Crawford and Glass, 1998; Liu et al., 1999). Phosphate transport is also assumed to be active and H+ -coupled at all concentration ranges, and some phosphate transporters appear to contribute significantly to P uptake at both high and low P concentrations (Shin et al., 2004). Plants are usually grown at a single nutrient level and then tested for their ‘instantaneous’ influx or net uptake rates at different solute concentrations. Depending on the solute and species, pre-treatment at very low concentrations of a given solute may increase or decrease high-affinity transport rates and may increase or reduce low-affinity transport capacity. For instance, K+ starvation induces high-affinity K+ transport, whereas high-affinity NO 3 − transport is induced by detection of extracellular NO 3 − . High-affinity uptake is reduced in K+ and NO 3 − -replete roots, in the case of NO 3 − by accumulation of products of NO 3 − assimilation rather than by NO 3 − itself (Forde, 2002). Low-affinity K+ influx may be unaffected (e.g. maize; Kochian and Lucas, 1982), reduced (e.g. wheat; Davenport, 1998) or enhanced (e.g. arabidopsis; Shin and Schachtman, 2004) by K+ starvation. In some cases the transcript or expression levels of particular transporter genes have been elegantly correlated with the time course of reduction or increase in influx rates (Okamoto et al., 2003). While there is plentiful evidence of inhibition of root uptake by high root concentrations of the solute or its metabolic products, it is also clear that shoot signals indicating above-ground nutrient status can override local root signals, as demonstrated by split-root experiments where the shoot solute content is manipulated independently of the roots (see Section 8.7).

8.9

Conclusion

Faced with patchy and fluctuating solute availability, plants utilise a variety of mechanisms to extract solutes from the soil. This chapter has focused on morphological and physiological responses to nutrient availability in conditions where plant roots must absorb nutrients directly from the growth solution without the complex effects of mycorrhizal and other symbiotic associations. This necessarily gives an incomplete picture of the complexity of plant responses to the rhizosphere in field conditions. Another area of neglect is the effect of toxic solutes present in the soil, which the plant must either exclude (and in some cases such as aluminium, chelate) or compartmentalise to avoid damage; these aspects are covered elsewhere in the book (see for example, Chapters 12 and 14).

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Narang, R.A., Bruene, A. and Altmann, T. (2000) Analysis of phosphate acquisition efficiency in different Arabidopsis accessions. Plant Physiology. 124, 1786–1799. Okamoto, M., Vidmar, J.J. and Glass, A.D.M. (2003) Regulation of NRT1 and NRT2 gene families of Arabidopsis thaliana: responses to nitrate provision. Plant and Cell Physiology 44, 304–317. Orians, C.M., Ard´on, M. and Mohammad, B.A. (2002) Vascular architecture and patchy nutrient availability generate within-plant heterogeneity in plant traits important to herbivores. American Journal of Botany 89, 270–278. Peterson, C.A. and Enstone, D.E. (1996) Functions of passage cells in the endodermis and exodermis of roots. Physiologia Plantarum 97, 592–598. Peterson, C.A. and Moon, G.J. (1993) The effect of lateral root outgrowth on the structure and permeability of the onion root exodermis. Botanica Acta 106, 411–418. Pitman, M.G., Wildes, R.A., Schaefer, N. and Wellfare, D. (1977) Effect of azetidine 2-carboxylic acid on ion uptake and ion release to the xylem of excised barley roots. Plant Physiology 60, 240–246. Rains, D.W. and Epstein, E. (1967a) Sodium absorption by barley roots: role of the dual mechanisms of alkali cation transport. Plant Physiology 42, 314–318. Rains, D.W. and Epstein, E. (1967b) Sodium absorption by barley roots: its mediation by mechanism 2 of alkali cation transport. Plant Physiology 42, 319–323. Reid, R.J. and Smith, F.A. (1992) Measurement of calcium fluxes in plants using 45 Ca. Planta 186, 558–566. Roberts, S.K. (1998) Regulation of K+ channels in maize roots by water stress and abscisic acid. Plant Physiology 116, 145–153. Roberts, S.K. and Snowman, B.N. (2000) The effects of ABA on channel-mediated K+ transport across higher plant roots. Journal of Experimental Botany 51, 1585–1594. Robinson, D. (1996) Resource capture by localized root proliferation: why do plants bother? Annals of Botany 77, 179–185. Rubio, G., Sorgon`a, A. and Lynch, J.P. (2004) Spatial mapping of phosphorus influx in bean root systems using digital autoradiography. Journal of Experimental Botany 55, 2269–2280. Scheurwater, I., Clarkson, D.T., Purves, J.V., et al. (1999) Relatively large nitrate efflux can account for the high specific respiratory costs for nitrate transport in slow-growing grass species. Plant and Soil 215, 123–134. Schmidt, W. (1999) Mechanisms and regulation of reduction-based iron uptake in plants. The New Phytologist 141, 1–26. Sentenac, H. and Grignon, C. (1985) Effect of pH on orthophosphate uptake by corn roots. Plant Physiology. 77, 136–141. Shin, H., Shin, H-S., Dewbre, G.R. and Harrison, M.J. (2004) Phosphate transport in Arabidopsis: Pht1;1 and Pht1;4 play a major role in phosphate acquisition from both low- and high-phosphate environments. The Plant Journal 39, 629–642. Shin, R., Berg, R.H. and Schachtman, D.P. (2005) Reactive oxygen species and root hairs in Arabidopsis root response to nitrogen, phosphorus and potassium deficiency. Plant and Cell Physiology 46, 1350–1357. Shin, R. and Schachtman, D.P. (2004) Hydrogen peroxide mediates plant root cell response to nutrient deprivation. Proceedings of the National Academy of Science of the United States of America 101, 8827–8832. Siddiqi, M.Y. and Glass, A.D.M. (1987) Regulation of K+ influx in barley: evidence for a direct control of influx by K+ concentration of root cells. Journal of Experimental Botany 38, 935–947. Siddiqi, M.Y., Glass, A.D.M., Ruth, T.J. and Rufty, T.W. (1990) Studies on the uptake of nitrate in barley. Plant Physiology 93, 1426–1432. Smith, J.A.C. (1991) Ion transport and the transpiration stream. Botanica Acta 104, 416–421. Smith, S.E., Smith, F.A. and Jakobsen, I. (2003) Mycorrhizal fungi can dominate phosphate supply to plants irrespective of growth responses. Plant Physiology. 133, 16–20. Steudle, E. and Peterson, C.A. (1998) How does water get through roots? Journal of Experimental Botany 49, 775–788.

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Tanner, W. and Beevers, H. (2001) Transpiration, a prerequisite for long-distance transport of minerals in plants? Proceedings of the National Academy of Science of the United States of America 98, 9443–9447. Tester, M. and Davenport, R. (2003) Na+ tolerance and Na+ transport in higher plants. Annals of Botany 91, 503–527. Tinker, P.B. and Nye, P.H. (2000) Solute Movement in the Rhizosphere. Oxford University Press, NY. Vahkimistrov, D.B., Kurkova, E.B. and Soloviev, V.A. (1972) Some characteristics of plasmodesmae and lomasome-like formations in barley roots and their relationship to transport of substances. Fiziologiia Rastenii 19, 951–960. Walker, N.A. and Pitman, M.G. (1976) Measurements of fluxes across membranes. In: Encyclopedia of Plant Physiology, Vol. 2(A) (eds Luttge, E. and Pitman, M.), pp. 93–126. Springer-Verlag, Berlin. Wang, M.Y., Siddiqi, M.Y., Ruth, T.J. and Glass, A.D.M. (1993a) Ammonium uptake by rice roots. I. Fluxes and subcellular distribution of 13 NH 4 + . Plant Physiology 103, 1249–1258. Wang, M.Y., Siddiqi, M.Y., Ruth, T.J. and Glass, A.D.M. (1993b) Ammonium uptake by rice roots. II. Kinetics of 13 NH 4 + influx across the plasmalemma. Plant Physiology 103, 1259–1267. Wang, M.Y., Siddiqi, M.Y., Ruth, T.J. and Glass, A.D.M. (1994) Ammonium uptake by rice roots. III. Electrophysiology. Plant Physiology 104, 899–906. Wegner, L.H. and Zimmermann, U. (2002) On-line measurements of K+ activity in the tensile water of the xylem conduit of higher plants. The Plant Journal 32, 409–417. Wenzel, C.L. and McCully, M.E. (1991) Early senescence of cortical cells in the roots of cereals: how good is the evidence? American Journal of Botany 78, 1528–1541. Wenzel, C.L., McCully, M.E. and Canny, M.J. (1989) Development of water conducting capacity in the root systems of young plants of corn and some other C4 grasses. The Plant Physiology 89, 1094–1101. White, P.J. (1997) The regulation of K+ influx into roots of rye (Secale cereale L.) seedlings by negative feedback via the K+ flux from shoot to root in the phloem. Journal of Experimental Botany 48, 2063–2073. White, P.J. (2001) The pathways of calcium movement to the xylem. Journal of Experimental Botany 52, 891–899. White, P.J., Whiting, S.N., Baker, A.J.M. and Broadley, M.R. (2002) Does zinc move apoplastically to the xylem in roots of Thlaspi caerulescens? The New Phytologist 153, 201–207. Wissuwa, M. and Ae, N. (2001) Genotypic differences in the presence of hairs on roots and gynophores of peanuts (Arachis hypogaea L.) and their significance for phosphorus uptake. Journal of Experimental Botany 52, 1703–1710. Yadav, R., Flowers, T.J. and Yeo, A.R. (1996) The involvement of the transpirational bypass flow in sodium uptake by high- and low-sodium-transporting lines of rice developed through intervarietal selection. Plant, Cell and Environment 19, 329–336. Yeo, A.R., Yeo, M.E. and Flowers, T.J. (1987) The contribution of an apoplastic pathway to sodium uptake by rice roots in saline conditions. Journal of Experimental Botany 38, 1141–1153.

9 Transport from root to shoot Sergey Shabala

9.1

Introduction

Photosynthetic processes in leaves are strongly dependent on a constant supply of water and mineral nutrients, as well as various low molecular weight organic compounds and hormones. Most of these are either synthesized in (e.g. cytokinins and gibberellins) or taken up by (e.g. water and mineral nutrients) plant roots and thus have to be transported to the shoot to enable normal leaf functioning. In its turn, a constant influx of assimilates from leaves to roots is needed for normal root performance. Accordingly, plants have a sophisticated system for the long-distance transport of solutes between roots and shoots. Such long-distance transport of water and solutes takes place in the vascular system of xylem and phloem (see Chapter 10 for a discussion on transport in the phloem). The focus of this Chapter is on solute transport in the xylem.

9.2 9.2.1

Transport of water Xylem structure

Xylem is a plant tissue specialized in conducting solutes from root to shoot and can be considered as a continuous system of inter-connected tubes with a relatively low resistance to the flow of water. Accordingly, its anatomical structure has two prominent features: (i) efficient leak prevention and (ii) remarkable mechanical strength, enabling it to withstand the substantial negative pressure required to drive water from soil to leaves. Xylem consists of four different cell types: vessels, tracheids, fibres and parenchyma. Of these, vessels and tracheids (often called tracheary elements) are responsible for the transport of water and solutes. Only the parenchyma cells are living; all others are dead, secondary lignified cells (Figure 9.1A). These occur most abundantly in the rays of woody species, but may be also scattered throughout the xylem. As a rule, gymnosperms have only tracheids, while nearly all angiosperms have both vessel elements and tracheids. Both vessel elements and tracheids are elongated cells. Their geometry is, however, rather different. Vessels are usually of a larger diameter, typically between 40 and 80 μm (ranging from 5 μm to 0.5 mm), and are formed from a series of cells connected end-to-end via perforation plates. Such plates have openings at which the secondary wall fails to form, and the primary wall and middle lamella are dissolved,

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(b) a

(c)

Rpit In

Pn

i=0

In

∫Rlumen(I)d/

In−1 Pn−1

P2

i=2

Rpit

In−1

Rpit I2

I2

I1 I1

i=1

I0

I1

∫Rlumen(I)d/

Rpit

P0

a

∫Rlumen(I)d/

Rpit

P1

(A)

{P2}

{P1} a

I0

a

(B)

Figure 9.1 Xylem structure and water flow. (A) Xylem structure in woody species (Populus grandidentata). Xylem vessel elements and inter-vessel pits are clearly seen (from Zimmermann, 1983; with permission of Springer-Verlag). (B) Idealized water-flow path through the xylem and its equivalent n-layer electrical diagram depicting lumen (R lumen ) and pit (R pit ) resistance to water flow. Reproduced from Comstock and Sperry, 2000, with permission of New Phytologist Trust.

enabling unrestrained passage of water. To enable mechanical strength, vessel elements are typically strengthened by various thickenings (rings or spirals). Vessel elements are so aligned that they form long tubes called vessels (many cells). Each vessel may extend from a few millimetres to several meters (Zimmermann, 1983) so that in some modern angiosperms, a few individual vessels can span the entire height of a tree. Tracheids are single, elongated, lignified cells of much smaller diameter (10–25 μm) inter-connected by pits (areas lacking the secondary wall) in all directions, but not formed into longitudinal pipes. Instead, tracheid cells have tapered ends that overlap. The so-called pit membrane, a modified primary cell wall consisting a dense network of hydrophilic cellulose polymers, is water permeable (Holbrook and Zwieniecki, 1999) and thus enables relatively unobstructed sap flow. Pits in the tapered region of the cell allow water to move upward from one tracheid into the next, while numerous pits across the sides also allow passage of water between adjacent cells. The development of tracheary elements occurs from the procambium and vascular cambium following programmed cell death and a consequent loss of their protoplasts (de Boer and Volkov, 2003). One specific feature of tracheids and vessel elements is the presence of a secondary wall, consisting largely of cellulose, lignin and hemicellulose. This secondary wall covers most of the primary wall and confers considerable compression strength on the cell, preventing it from collapsing under extreme tension. A key determinant

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of the mechanical properties of the cell wall is lignin, a polyphenolic polymer that influences both wall rigidity and compressive strength. It is reported that the observed difference in lignification corresponds well with variations in ion-mediated change in xylem hydraulic properties among different taxa (Boyce et al., 2004). In Ginkgo biloba and conifer tracheids, the primary cell wall and middle lamella are more heavily lignified than the secondary wall. More developed angiosperms have non-lignified, compound primary walls between vessel elements. Such weak lignification of compound primary cell walls should enhance ion-mediated changes in xylem flow rate, but will also diminish cell adhesion among tracheids and vessels, reducing the effective capacity of conducting cells to withstand mechanical stresses. Therefore, an apparent functional conflict between efficiency and safety is obvious.

9.2.2

Physics of water flow and evolutionary aspects of conduit development

The xylem conduits form an extensive network of capillaries. These conduits are not of the same length and, in most plants, the ends of each individual conduit are randomly distributed longitudinally than lined up in transverse plane (Comstock and Sperry, 2000). Nonetheless, a generalized equivalent circuit diagram can be drawn to quantify parameters of water flow through the xylem (Figure 9.1B). In this simplified model, the flow path is divided into n discrete tiers of conduits. Most of the time, water flows in the large, unobstructed lumen of the capillaries. However, it must also cross through inter-conduit pits at a frequency determined by the length of individual conduits relative to the overall pathway. Therefore, two major types of resistance can be identified: pit (R pit ) and lumen (R lumen ) resistance. The pit resistance is defined as the total wall-crossing resistance, associated with all water in the pathway moving from one conduit tier to another. The lumen resistance is dependent on both the length of all conduits in an appropriate tier as well as on their diameter (Comstock and Sperry, 2000). The quantitative relation between the flow rate in the conduit and xylem geometry is determined by the Hagen-Poiseuille’s law (Zimmermann, 1983): V =−

πr 4 ∂ P 8η ∂ x

(9.1)

where V is the volume flowing per tube, η is the dynamic viscosity of water and r the radius of the conduit; the term ∂ P/∂ x represents the gradient in pressure and π has its usual meaning. Consequently, the resistance in the lumen is: Rlumen =

πρ

8η

r4

(9.2)

where ρ is the water density. This equation suggests that the flow rate (volume per unit time per tube rather than volume per unit time per unit area) through capillaries is proportional to the fourth power of the radius of the capillary. Therefore, a twofold increase in the conduit diameter leads to a 16-fold increase in the flow rate. Not surprisingly,

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the flow through a large vessel is much more rapid than through tracheids and small vessels. As such, peak velocities for xylem flow are 13 mm s–1 in trees with large vessels (Taiz and Zeiger, 1991), while in wheat plants the mean speed of water is 0.8 mm s–1 (Passioura, 1988). The difference is in the radius of their xylem vessels. Also there appears to be a strong evolutionary trend in conduit development. The first evidence for xylem conduits in the form of tracheids is found in the upper Silurian (Edwards and Davies, 1976). As discussed above, to avoid implosion, the walls of tracheids include secondary layers for thickening and are impregnated with the stiffening agent lignin. Such lignified and thickened walls presumably evolved in response to the collapsing forces of water under tension in the early conduits and are found in a wide range of tissues in modern plants. Increased mechanical strength and a selective pressure for a more efficient water transport from roots to shoots led to a progressive increase in tracheid diameter over time (Comstock and Sperry, 2000). Associated with this was an increase in conduit length. However, at large conduit diameters, the pit resistance became increasingly limiting (Figure 9.1B), contributing 50% or more of the total hydraulic resistance of the pathway (Calkin et al., 1986). A selective pressure to reduce pit resistance eventually resulted in the evolution of longer xylem conduits to reduce the number of crossings in the flow pathway (Figure 9.1B; Comstock and Sperry, 2000). However, this affects water transport in two opposing ways: (i) it reduces the number of wall crossings (thus, the hydraulic resistance to flow within the xylem) but (ii) it increases the risk of a greater loss of conductance due to cavitation (discussed in details in the next section). A sensible compromise can be achieved if conduits are not of equal length in all portions of the pathway, but instead, the frequency of the end wall is proportional to the magnitude of water potential gradient at each point – when conduits are longest in the basal portion (roots) and progressively shortened higher up the stem towards the foliage (Comstock and Sperry, 2000). Also, driven by above trends, a new type of conduit emerged – the vessel. Vessel members evolved from tracheids by the progressive enlargement of inter-conduit pits and the degradation of pit membranes to form open perforation plates. The individual membranes at the top and bottom of the vessel retain their end walls and define the total length of the vessel. Importantly, vessels do overlap for a large fraction of their length, providing the surface area for inter-conduit pits thus enabling an efficient water flow from one vessel to another (Zimmermann, 1983).

9.2.3

Water flow between xylem elements: safety mechanisms

Rapid flow through large vessels comes at the price of safety. Significant cavitation (the vapourizing of liquid xylem sap that is under negative pressure) can occur in plants under natural water stress conditions, leading to embolism (formation of the vapour-filled cells). In most cases, impregnation of the cell-wall matrix with lignin prevents embolism. There are cases, however, when the conduit becomes damaged. The most obvious instance is cavitation and the following embolism caused by freezing stress (Figure 9.2C). It is known that conduits with smaller radii are more resistant to cavitation by freezing stress than those with larger radii (Davis et al.,

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(B)

(A)

(C)

(D)

Figure 9.2 Embolism and safety features of sap ascent. The flow of water between tracheids is mediated by numerous pits along the pathway. Under normal conditions, all tracheids are filled with water (B), and upstream water flow is unobstructed (A). Significant cavitation can occur in plants under natural water stress conditions (such as freezing), leading to embolism (formation of the vapour-filled cells; C). As a result of the negative pressure from neighbouring cells, pits in embolized tracheid (shaded area in D) are closed, and water flow bypasses the embolized element. Panels B and C are reproduced from McCully et al., 2000, with permission of Blackwell Publishing. Bar = 50 μm.

1999), and a strong trend does exist for decreasing conduit radius with latitude (Comstock and Sperry, 2000). Other cases may include the rupture of protoxylem during organ expansion, by abscission of leaves and fine roots, by herbivory or pathogen damage or by mechanical damage. The ruptured conduits allow air to leak into the xylem. As the water is withdrawn from the damaged conduit, the conduit becomes embolized (filled with air). Thus, conducting systems composed of very long conduits are extremely susceptible to failure from mechanical point injuries and pathogen attack anywhere along the pathway (Zimmermann, 1983). How can the embolism be prevented from spreading along the xylem? As the lignified secondary walls are not permeable to water, the water flow between adjacent conduits occurs through the thin and porous pits (round, thin places where the cells are separated only by the primary walls) in the common wall (Figure 9.3). The pits consist of the porous pit membrane derived from the compound middle lamella of the adjacent cells. This divides opposing pit ‘chambers’ formed by the openings in the secondary walls (Comstock and Sperry, 2000). In plain language, pits constitute the gates between vessels and from vessels to adjacent parenchyma

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cells. As most pits are of a very small diameter, they are capable of excluding air entry by capillary forces (Pickard, 1981). In other words, surface tension acts as an interfacial water–air stopper, preventing air from being sucked into tiny pores present in all plant cell walls. Because of this feature, pits are said to act as non-return valves. The total area of pit fields, their shape and pattern of lignification vary dramatically between species and may constitute up to 15% of the total vessel area (de Boer and Volkov, 2003). Pit fields can be either simple, un-reinforced structures (simple pits) or more elaborate bordered pits in which secondary walls mechanically support the pit membrane (due to its ‘arc-type’ structure). Such reinforcement allows the pit membrane to be as large as possible and thereby maximizes water exchange between vessels. Bordered pits are characterized by overarching walls that form a bowl-shaped chamber. The secondary walls extend over the centre of the pit and the primary walls are swollen in the centre of the pit to form a torus (thus, torus-margo structure). As the primary wall around the torus is porous, the torus can act as a valve, closing when pressure on one side is greater than that on the other (Figure 9.3). Such a system will act as a check valve anchoring the air–water meniscus by capillary forces. This allows negative pressure to persist in the undamaged conduit system (Zimmermann, 1983) and is thus vital to protect against cavitation and a subsequent embolism. Pits are also instrumental in repairing the ‘broken pipes’ (Holbrook and Zwieniecki, 1999), i.e. removing air bubbles. It is believed that the latter occurs through gas dissolution in the surrounding water (de Boer and Volkov, 2003), although specific details of this process remain unclear.

9.2.4

Hydraulics of the sap lift: general overview

The overall driving force for water lifting in plants is ultimately the chemical potential difference of water between the soil and the atmosphere (see Section 3.5). However, the mechanisms by which water rises against gravity are still controversial, despite extensive research over more than 150 years (Pickard, 1981; Zimmermann, 1983; Sperry, 1995; Tyree, 1997; Zimmermann et al., 2002). The tallest tree ever measured is believed to be Eucalyptus amygdaliana at Mount Baw Baw, Victoria, Australia (143 m in 1885; Salisbury and Ross, 1992). At the same time, even the best vacuum pumps can lift water no more than 10 m. As a result, until the end of the nineteenth century the mechanism by which trees are able to draw water to heights of more than 100 m remained a mystery. It was only in 1894 when H. Dixon and J. Joly formulated the hypothesis of sap ascent by cohesion and adhesion (or Cohesion–Tension theory as it becomes known after 1914; Tyree, 1997; Zimmermann, 1983). Three basic elements of this theory are the driving force (gradient in water potential between the soil and atmosphere), hydration/adhesion (an attractive force between unlike molecules) and the cohesion of water (an attraction between like molecules). The essence of this theory is that water ascends plants in a metastable state under tension, i.e. with xylem pressure more negative than that of vapour pressure of water. The driving force is generated by surface tension at the evaporating surfaces of the

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Embolised cell

2 3 1 4 (A)

(B)

(C

(D) Figure 9.3 Structure and function of bordered pits in xylem conduits. (A and B) The secondary walls (2) extend over the centre of the pit and the primary walls (1) are swollen in the centre of the pit to form a torus (3). With both neighbouring elements filled with water, the pressure is equal on both sides, and the water flows essentially unobstructed through the porous pit membrane (4). (C) When one of the conduit elements becomes embolized, pressure imbalance pushes the torus so it plugs the hole, acting as a non-return valve. (D) SEM illustrating bordered pits connecting two vessel elements in Eucalyptus. Reproduced from Atwell et al., 1999, with permission from Australian Society of Plant Scientists. An ‘arc-type’ structure reinforces the pit and allows it to withstand high pressure required to draw water flow in the xylem.

leaf (Figure 9.4). Loss of water by transpiration sets up a tension that is transmitted to the xylem, and from there, down continuous columns of water to the roots. A central part of this model is that water molecules cling to each other through the hydrogen bonds they form (so-called cohesion; see also Section 3.2). When water molecules are close together, their positive and negative regions are attracted to

TRANSPORT FROM ROOT TO SHOOT

221

R1

R2

R3 Figure 9.4 The surface tension as a driving force for hydraulics water lift in plants. Surface tension forces lower xylem pressure in the liquid directly behind the menisci (the air–water interfaces; shown as dashed lines for three different positions). The xylem pressure is inversely proportional to the radius (R) of curvature of a meniscus. As R decreases in result of evaporation, the tension increases pulling more water into the xylem (see also Eq. 3.13).

the oppositely charged regions of nearby molecules. Such a force of attraction is called a hydrogen bond and gives the water column remarkable mechanical strength of solid wire (high enough to withstand tensions of several MPa, typically found within xylem). The water potential gradient causes water to move out of xylem, through the leaf parenchyma to evaporate at the leaf surface. Due to cohesion, as one water molecule is removed, others are drawn up along the continuous column. Thus, through the evaporative power of the atmosphere, a continuous ‘chain’ of water can be drawn up to the leaf canopy. The tension created in this way could even suck water from the surrounding soil. One critical condition is the absence of microscopic gas bubbles disrupting hydrogen bonds and causing the breakage of the water column (the main reason why vacuum pumps fail to lift water for more than 10 m). The small vessel diameter and adhesion (attraction of water to vessel wall) are therefore important features enabling the hydraulic lift in a plant’s xylem.

9.2.5

Driving force for water movement in the xylem

As discussed above, the major driving force for water lifting in plants is a gradient in water potential ( w ) between the soil and atmosphere. Water potential increases (becomes less negative) in the following sequence: atmosphere leaf cells > xylem sap > root cells > soil solution (see Table 3.1). With between 450 and 600 molecules of water being evaporated from the leaf surface to assimilate one CO 2 molecule, the transpiration stream is a major component forcing xylem transport during the daylight and is responsible for the negative pressure in xylem vessels (Zimmermann et al., 2002). There is another component however, which might play an important role under environmental conditions of low transpiration

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(e.g. high humidity, darkness). This is a gradient in hydrostatic pressure (or root pressure), created by the secretion of ions into the xylem and a consequent water flow through surrounding cells by osmosis (Marschner, 1995). A nice illustration of the existence of root pressure is a phenomenon known as guttation, when plants exude fluid through the edges or tips of their leaves. Such guttation is often found under conditions of good water supply and poor transpiration (e.g. high humidity). However, it appears that not every species develops root pressure (Fisher et al., 1997). Therefore its overall contribution to hydraulic water lift in plants is not very significant, at least for majority of ‘physiological’ conditions (see Section 9.2.6). Contrary to general believe, air does not have to be very dry to establish a steep water potential gradient between the soil and the atmosphere. Simple calculations (Eq. 3.10) show that at 20◦ C and a relative humidity of 98%, the atmospheric water potential  atm is –2.72 MPa, i.e. enough to move a column of water to a height of 277 m. When the relative humidity of the air is 90%,  atm = –14.2 MPa. As the soil-water potential is usually within the range of 0–1.5 MPa, this is more than enough not only to lift water to a substantial height within the plant but also to extract water from the deeper soil horizons. In addition to the holding power of negative water potentials in living cells, there is much greater holding power of hydration within the cell wall of the apoplast (adhesion). As the amount of water diminishes in the cell walls or leaf xylem elements following the transpiration stream, curved menisci begins to form between the cell-wall polysaccharides and in the intracellular spaces (Figure 9.4). Surface tension forces lower the xylem pressure in the liquid directly behind the menisci (the air–water interface). This occurs because the free energy of the system is lowest with a minimum surface area. Quantitatively, the xylem pressure is inversely proportional to the radius of curvature of a meniscus according to Eq. 3.13 (P = − 2σr , where P is pressure, σ the surface tension of water (0.072 Pa m) and r is the radius of meniscus m. During evaporation, r decreases (Figure 9.4), so the tension (–P) increases. When the radius of curvature is as little as a few μm, the magnitude of the resulting tension becomes physiologically significant (for example, P = –1.44 MPa for a meniscus of 10 nm radius). This creates a low water potential in adjacent regions, including adjoining cell walls and protoplasts (Tyree, 1997). The above tension is transmitted from the cell-wall surface to the xylem so that the solution in the xylem is under tension. The hydrogen bonds provide the tensile strength of water, so even under a tension of several MPa a water column does not break (the empirical value for the tensile strength of pure water in capillaries is 30 MPa; Nobel, 1983). The latter is reduced dramatically by impurities, however, emphasizing the importance of avoiding a cavitation.

9.2.6

Controversies and additional mechanisms

The Cohesion–Tension theory implies that the tension gradient between the root and leaf xylem is the only driving force for hydraulic water lift in the xylem. Indeed, probing of vessels of a large number of tree and herbaceous species demonstrated

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the existence of both negative pressure and axial gradients in transpiring plants (Zimmermann et al., 2002). Also, as early as 1965, Scholander and co-authors (1965) showed that the balancing gas pressure required to force sap through the protruding cut end of an excised leaf in a pressure chamber provides a good estimate of the original xylem tension of the intact plant. Also consistent with the theory, increased light intensity and temperature result in a decrease of the xylem pressure towards more negative values, and a good correlation between xylem pressure and flow velocity was established (Wistuba et al., 2000). There are several controversies, however, that cannot be explained by the classical Cohesion–Tension theory. For example, tensions of at least 3 MPa (even more if the requirement of water lift to the soil surface is taken into account) are required to lift water to foliage of a 100-m-high tree. Such tensions are not measured experimentally. Xylem tension exceeding 0.6 MPa is rarely observed within the xylem pressure probe, the most direct method of measuring xylem water potential (Zimmermann et al., 2002). A maximum tension of only 0.2 MPa was measured in the midrib of well-hydrated leaves of the 35-m-tall tropical tree Anacardium excelsum (Zimmermann et al., 1994); this is hardly enough to lift water by only cohesion force. Also, diurnal gradients in xylem tension measured in liana were sometimes opposite to those required by the Cohesion–Tension theory (Benkert et al., 1995). Finally, hydrophobicity of the xylem wall is totally ignored by the Cohesion–Tension theory even though it reduces the cavitation threshold considerably. Taken together, these findings suggest the possibility of some other mechanisms contributing to sap ascent in the xylem. According to Zimmermann et al. (2002), tall plants have apparently developed additional strategies for lifting water against gravity. Several possible mechanisms have been suggested. Contribution from gel-like compounds. Various acid mucopolysaccharides are known to be attached to the walls (Plumb and Bridgman, 1972). Such compounds may be important for ‘conditioning’ the xylem elements in order to keep continuous water films up to the foliage even when vessels are partially filled with air and/or vapour (Zimmermann et al., 2002). Also, these substances may act as cryoprotectants, preventing xylem sap from freezing. Osmotic water lifting. Osmotic water lifting is also theoretically possible and has been advocated by Zimmermann et al. (2002). However, most papers report the xylem sap osmolality being around 40 mOsmol, questioning the role of osmosis in water movement in the xylem. Nonetheless, an osmotic pressure of ∼0.6 MPa was measured in the xylem sap in trunks and branches of birch and maple trees during bud burst (Wistuba et al., 2000; Zimmermann et al., 2002). This is high enough to drive water up to large heights even in the absence of transpiration. Ionic control of xylem conductance. It was shown that the presence of an osmotically insignificant (as low as 10 μM KCl) concentration cations substantially influenced xylem conductance (van Ieperen et al., 2000). It is suggested that such a change in the hydraulic resistance in response to changes in the ionic

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composition of the xylem sap is due to the pit membranes connecting one vessel to another (Zwieniecki et al., 2001). In these membranes, water flows through microchannels made up by cellulose microfibrils, hemicellulose and pectins (mainly D-galacturonic acid; Zwieniecki et al., 2001). The size of these microchannels increases when the pectin matrix shrinks in response to the binding of ions to the negative charged matrix. Involvement of electrical driving force in water rise (Zimmermann et al., 2002). A thin film of ions on the negatively charged inner surface of the vessels may be considered as an electric double-layer capacitor from the point of view of non-equilibrium thermodynamics. This may substantially contribute to water lifting (Amin, 1982). However, no hard experimental evidence supporting this idea has yet been gained. Nonetheless, a strong correlation between oscillations in trans-root potential and xylem pressure was shown (Wegner et al., 1999), calling for further investigations of the causal link between these two phenomena.

9.3

Transport of nutrients

Not only water but also a wide range of inorganic and organic nutrients is transported from the root to the shoot via the xylem. There is compelling evidence suggesting that nutrient translocation can be independent of the sap flow and might be regulated at the level of xylem loading in xylem parenchyma (e.g. Siebrecht et al., 2003). Specific features of xylem parenchyma cells are a rather dense cytosol, well-developed endoplasmic reticulum and numerous ribosomes, mitochondria and peroxisomes (de Boer and Volkov, 2003). All these features indicate a high level of metabolic activity. Importantly, xylem parenchyma cells have fewer cortical microtubules in their external cytoplasmic layer at the point of contact pits. The latter is important both to prevent the deposition of a thick secondary wall and regulate the activity of ion and water channels. Today, the key role of xylem parenchyma in both nutrient transport and long-distance signalling in plants is undisputable (Malone, 1996; de Boer and Volkov, 2003). In this section, electrophysiological and molecular evidence for mechanisms underlying ion exchange at the parenchyma/ xylem boundary are summarized, and factors controlling xylem ion loading are discussed.

9.3.1

General features of xylem ion loading

Like many other issues related to xylem physiology, mechanisms for loading of nutrients into the xylem are still under debate (see also Section 8.7), with both passive and active models being advocated. It was hypothesized in the early days that xylem loading in the root is a passive process, due to a lack of oxygen in the core of root tissue (a so-called leakage hypothesis postulated by Crafts and Broyer (1938)). This idea has been further discussed by others, and the modern view is that we are dealing with much more sophisticated mechanisms (de Boer and Volkov, 2003).

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Electrophysiological techniques have demonstrated the presence of both cation and anion channels in xylem parenchyma (Wegner and Raschke, 1994; Wegner and de Boer, 1997; K¨ohler and Raschke, 2000; K¨ohler et al., 2002), suggesting that these channels are responsible for the loading of solutes into the xylem (Tester and Leigh, 2001). In addition, active models assuming the involvement of the H+ pumps at the symplasm–xylem interface have been advocated (de Boer and Volkov, 2003). This view is supported by both findings of high plasma-membrane ATPase activity in xylem parenchyma cells (reviewed by de Boer and Volkov, 2003) and by the fact that the potential difference between xylem vessel and parenchyma cell is strongly regulated by auxin and fusicoccin, two known activators of H+ -ATPase (de Boer et al., 1985; de Boer, 1997). Irrespective of the different views on the mechanism, there is a general agreement that xylem loading is regulated separately from the ion uptake in cortical cells by the nutritional status of the plant, and with the amount of nutrient recycled via the phloem being a feedback signal to control radial nutrient uptake by roots (Drew et al., 1990).

9.3.2 9.3.2.1

Ionic mechanisms of xylem loading Potassium

Potassium is a key nutrient involved in a plethora of physiological and metabolic processes in plant cells. With K+ concentration being as high as 6% of the leaf dry weight, it is not surprising that K+ loading and transport in the xylem is critical to plant performance under variable environmental conditions. Both molecular and electrophysiological data suggests that efflux of K+ from xylem parenchyma cells into the xylem is carried out by depolarization-activated outwardly rectifying K+ channels (Wegner and Raschke, 1994; Roberts and Tester, 1995; Wegner and de Boer, 1997; Gaymard et al., 1998; de Boer and Volkov, 2003). This is further illustrated in Figure 9.5. These channels, named SKOR (for shakertype K+ outward rectifying) were first detected in root parenchyma of barley (Wegner and Raschke, 1994; Wegner and de Boer, 1997) and maize (Roberts and Tester, 1995; Roberts, 1998). They were reported to activate in a time-dependent manner at membrane potentials slightly positive of the reversal potential of K+ (E K ). Barley SKOR channels were highly selective between K+ and Na+ and also facilitated the passage of Ca2+ (Wegner and de Boer, 1997; de Boer and Volkov, 2003). Recent work on poplar suggested that different types of SKORs may be present in the same tissue (Langer et al., 2002; Arend et al., 2005) with different activation kinetics and voltage-gating properties. While one of these channels (termed PTORK) mediates K+ release upon depolarization, another channel (PTK2) is essentially voltage independent. PTK2 carries inward K+ flux at hyperpolarized potential, and K+ release upon depolarization, when expressed in heterologous systems (Langer et al., 2002). Both these channels are found predominantly in the xylem parenchyma surrounding the vessels, and in the phloem (Arend et al., 2005), only in actively growing poplar plants. Also, a strong correlation between poplar SKORs activity and seasonal wood production was found (Langer et al., 2002).

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K+ Na+

HKT1

H+

H+ ATP H+ Sug K+

KAT2

K+

AKT2/3

K+, Na+

SKOR

K+, Ca2+

NORC

K+, Ca2+ Na+, A-

Root

KE

Leaf

AA-

GluR2

K+ H+ H+

ATP

SOS1

H+ Na+

IRAC

NO3-

QUAC SLAC

ClMal2-

Figure 9.5 Carrier and channel transport systems involved in ion loading into and unloading from the xylem in ‘idealised’ plant species.

9.3.2.2

Sodium

Control of Na+ loading into the xylem appears to be a key feature of plant salt tolerance (Maathuis and Amtmann, 1999; Tester and Davenport, 2003; see also Chapter 14). Different Na+ transport mechanisms are present at the symplast/xylem boundary in the root (Figure 9.5), with one of the likely candidates being a nonselective outward-rectifying (NORC) channel (Wegner and Raschke, 1994; Wegner and de Boer, 1997). This channel is permeable to both cations (P Na ≈ P K ) and anions and shows strong voltage dependence (open at zero potential). The NORC channel is also controlled by external Ca2+ and thus may potentially be involved in xylem loading and signalling. However, it remains to be shown whether this channel is actually involved in Na+ loading into xylem (de Boer and Volkov, 2003). Another

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prominent candidate for Na+ transport across the symplast/xylem boundary is the Na+ /H+ exchanger SOS1 (Ding and Zhu, 1997; Section 14.10). As its activity is facilitated by a large pH gradient between xylem sap and parenchyma cell cytosol, it appears that Na+ loading into xylem might be in fact an active process! Another possibility for Na+ loading into the xylem is a glutamate receptor gene, AtGluR2. This gene has strong homology to non-selective cation channels in mammalian tissues (Lam et al., 1998) and was reported to be expressed in cells adjacent to the conducting vessels in arabidopsis leaves (Kim et al., 2001).

9.3.2.3

Anion channels

9.3.2.4

Gating factors

Three major types of anion channels facilitate anion (Cl– , NO 3 – , malate2– ) release into xylem (K¨ohler and Raschke, 2000; White and Broadley, 2001): (i) an inwardly rectifying anion channel (X-IRAC), characterized by activation at hyperpolarization and open times up to several seconds; (ii) a quickly activating anion channel (XQUAC), important for anion efflux at voltages between –50 mV and the equilibrium potential for the prevailing ion; and (iii) a slowly activating anion channel (X-SLAC), activating above –100 mV. Two of these channels (QUAC and IRAC) were found to be sensitive to cytosolic Ca2+ . Importantly, it was shown that NO 3 – in the xylem exerted positive feedback on QUAC conductance through the change in its voltage dependency (K¨ohler et al., 2002). Taken together, these results suggest that xylem loading with anions is likely to be a passive process.

There appears to be a large number of factors controlling the process of xylem loading, including apoplastic ion concentrations, cytosolic Ca2+ , pH, auxin and abscisic acid (ABA) (de Boer et al., 1985; de Boer and Volkov, 2003). Both external and internal acidification led to a decrease in the steady-state currents through SKOR and guard-cell outward rectifying (GORK) channels (Ache et al., 2000; Lacombe et al., 2000). Changes in pH of the xylem sap can influence xylem loading via two avenues: (i) being a driving force for antiport and symport or (ii) as a regulator of ion transporters or signalling molecules. Elevated cytosolic Ca2+ (Wegner and de Boer, 1997) and ABA (Gaymard et al., 1998; Roberts, 1998) also significantly affects SKOR activity. The latter may have rather significant adaptive consequences, as ABA translates the status of water availability of cells and tissues into metabolic and developmental adaptation of plants and thus can serve as an endogenous signal in the regulation of ion loading of the xylem vessels. Last but not least, apoplastic (xylem) ion concentration per se is a rather potent regulator of xylem loading. Expression of SKOR channels was induced by K+ and repressed by ABA and by conditions of K+ depletion (Gaymard et al., 1998). At the same time, the high-affinity K+ uptake transporter AtKUP1 was enhanced by K+ depletion (Fu and Luan, 1998). As a significant amount of K+ transported to the shoot is re-translocated to the root via the phloem (up to 40% in barley; Wolf and Jeschke, 1987), changes in the apoplastic K+ will result in a shift in the voltage–current relations (Wegner and de Boer, 1997) and thus will provide a required feedback for the xylem loading based on the shoot’s request.

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PLANT SOLUTE TRANSPORT Table 9.1 Concentration of major solutes in the xylem sap of several plant species Concentration range (Mm) Solute/species

Poplara

Banksiab

Sunflowerc

NO3 − H2 PO4 − SO4 2− Cl− K+ Ca2+ Mg2+

1–3 0.5–1.5 0.2–2 0.2–0.8 2–6 0.5–1.5 0.4–1.2

0.01 0.11 0.25 2.92 2.39 0.48 0.55

5–15 0.2–0.7 nm nm 2–8 0.3–1.2 0.2–0.8

nm – not measured. Siebrecht et al., 2003. b Pate and Jeschke, 1993. c Schurr and Gollan, 1990.

a

9.3.3

Xylem-sap composition

Composition of the xylem sap is highly variable and modified according to the requirements of the shoot tissues. Toxic ions can be removed from the sap and essential nutrients recycled intensively. Apart from mineral nutrients, organic molecules may be also present in substantial concentrations. In general, the osmotic composition of xylem sap is rather low (∼40 mOsmol l–1 ), indicating relatively low overall nutrient concentrations. However, xylem-sap composition is highly variable (Table 9.1) and can be rapidly modified by the shoot’s demand. Transfer from xylem to phloem also provides a mean of diverting essential elements from the main transpiring surfaces (older leaves) to growing tissues where they are required. The latter occurs via highly specialized transfer cells and requires active transport. There are also some serious methodological issues relating to quantification of the ionic sap composition. Cutting the shoot can cause contamination of the exudate with components of destroyed cells. Also, an interruption of phloem flow may affect the concentration of nutrients that cycle between roots and shoots. In fact, mineral nutrient dynamics have hardly ever been measured in the xylem of intact, transpiring plants (Watson et al., 2001). Since a direct extraction of xylem sap is confounded by the presence of large negative pressures (Tyree, 1997), most methods use facilitated exudation by means of the pressure bomb (Schurr, 1998). Cryo-analytical methods (X-ray microanalysis) are another popular tool. However, these methods have a very poor detection limit (∼10 mM) and are thus not suitable for measuring the majority of ions except may be K+ (Enns et al., 1998). Recently, a new method using xylemfeeding insects (such as Philaenus spumarius) applicable to intact transpiring plants have been introduced (Watson et al., 2001). With this method, insects’ excreta are collected at regular intervals and xylem nutrient composition is determined by atomic absorption spectroscopy (AAS) or flame photometry. Non-invasive techniques such as NMR imaging or spectroscopy are also a viable alternative (de Boer and Volkov, 2003).

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As evident from Table 9.1, most solutes in xylem sap are inorganic ions, with average concentrations for most of these being in the millimolar range. Quite often, however, nitrate and ammonium are assimilated into organic forms in the root to be transported to the shoot. Also complexed are metal ions such as Zn, Cu and Fe; these are almost exclusively chelated to amino acids. Organic molecules can also be present in substantial concentrations. Sugar concentrations of ∼5 mM were reported for maize xylem sap (Canny and McCully, 1988), and in some temperate deciduous trees such as maple the amount of carbohydrates in the xylem sap can reach much higher values (between 1% and 5%; Larochelle et al., 1998). Phytohormones are also found in xylem sap but often at concentrations several orders of magnitude lower than those required to elicit a physiological response (e.g. ABA concentration between 1 and 50 nM; Wilkinson and Davies, 2002). Xylem sap pH also shows a great deal of variability. In perennial species, the pH of the xylem sap is acidic (pH 5.5) at the beginning of spring and close to neutrality in winter (Fromard et al., 1995). Sap acidity is usually attributed to the activity of H+ -pumping ATPases, which are much more concentrated in the plasma membranes of xylem parenchyma cells than in other plant cell types. Xylem sap pH can increase in response to soil drying (Wilkinson and Davies, 2002) and is reduced under iron stress conditions (Lopez-Millan et al., 2000). These changes in the sap pH will affect both the electrical and chemical component of the proton electrochemical potential ( μ H ) and thus modify the driving force required for active xylem loading. Rather significant spatial heterogeneity also exists, with a gradient of apoplastic pH of ∼0.5–1 units reported between the centre of the xylem veins as compared with surrounding cells in the leaf apoplast (M¨uhling and L¨auchli, 2000).

9.3.4

Factors affecting ion concentration in the xylem

Both external and internal factors contribute to xylem loading. Among internal factors, root respiration and the carbohydrate status of the roots are the most important (Marschner, 1995). Feedback mechanisms (e.g. K+ cycling) have already been discussed above. It is reported that in wheat and rice ∼60% amino-N and ∼30% of xylem K+ originates from the phloem (Grignon and Sentenac, 1991). Among external factors, effects of light intensity on ion uptake and release are well documented (e.g. Casadesus et al., 1995). It appears that these effects are not only limited exclusively to the supply of photosynthates but also modulated by other factors related to the shoot’s demand for growth. Root zone temperature is another significant contributor and may selectively affect both radial transport and xylem loading. Closely related to light and temperature are diurnal variations in xylem sap concentration (Macduff and Bakken, 2003; Siebrecht et al., 2003). These fluctuations arise from perturbation in nutrient supply and demand and can be responsible for up to 70% reduction in the translocation rate from root to shoot (Siebrecht et al., 2003). Nutrient availability is another key factor. The ‘rule of thumb’ is that the concentration of any ion in the sap solution is usually proportional to its availability in the soil solution (White and Broadley, 2001). However, significant interactions between nutrients may take place, both at uptake sites and during translocation in the xylem.

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Xylem ion concentration tends to change dramatically under stress conditions. Under drought conditions, K+ loading into the xylem is reduced, presumably as a result of increased ABA level in the xylem sap (Roberts and Snowman, 2000). This may be required to sensitize stomata to ABA in the leaf by reducing K+ availability as a guard-cell osmoticum (Wilkinson and Davies, 2002). Water stress also reduces H+ -ATPase activity in parenchyma cells (Hartung and Radin, 1989), affecting loading of other ions into the xylem. Salinity effects on Na+ and K+ loading into xylem are well documented (e.g. Maathuis and Amtmann, 1999; Tester and Davenport, 2003), with a sharp decline in xylem sap K+ /Na+ ratio reported.

9.3.5

Xylem unloading in leaves

As discussed above, most of the solutes and water taken up by the roots are transported into the leaves via xylem vessels. Here nutrients must be delivered to a myriad of individual cells composing the leaf. The question is how these fluxes are diverted from the place with highest transpiration (mature leaves) to major sinks (e.g. developing leaves or fruits) where they are most needed. A plausible explanation is that this process is under a strict hormonal control and thus may be regulated by both endogenous cell hormones and variety of signalling substances transported by the transpiration stream (e.g. ABA, auxin, cytokinin, systemin; Wegner and de Boer, 1997). Due to water loss (transpiration), nutrient concentration in the xylem of leaves can dramatically increase. Unless some of this excessive solute accumulation at the terminal sites of the transpiration stream is not removed, necrosis on the tips of margins of the leaves may occur (Marschner, 1995). To prevent this problem, plants have developed sophisticated mechanisms for xylem-ion unloading in leaves (Figure 9.5). A complex network of major and minor veins crossing the leaf blade is used. This network is highly branched (all together, six branch orders are found in the leaves of dicotyledonous plants). Via this network, ions are able to be delivered to the proximity of the sink, unloaded by shoot/leaf xylem parenchyma cells and transported to the sink cell via symplastic diffusion. Specific mechanisms of this process remain largely unknown (de Boer and Volkov, 2003), but both passive and active mechanisms are likely to be involved (Figure 9.5). It has been suggested by various authors that K+ re-absorption by parenchyma cells occurs via potassium inward-rectifying (KIR) channels (de Boer and Volkov, 2003), energized by the activity of the H+ pump. Two types of KIR channels in the barley xylem parenchyma cells were reported by Wegner and de Boer (1997) and suggested to be involved in xylem unloading of K+ . Both these channels were regulated by G-protein and closed at potentials more positive than E K . Another (third) hyperpolarization-activated K+ channel was found in inside-out patches only. Two different types of KIR channels in leaf xylem protoplasts were found in maize (Keunecke et al., 1997), although the molecular identity of these channels remains to be elucidated. There is also explicit evidence that xylem unloading may be mediated by secondary active transport mechanisms. H+ /K+ exchangers have been suggested to operate at the parenchyma symplast–xylem interface (de Boer et al., 1985), the best

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characterized of which is the arabidopsis AtHKT1 transporter. This transporter is located in xylem parenchyma cells (Sunarpi et al., 2005) and, being Na+ selective (Uozumi et al., 2000), can mediate unloading of Na+ from xylem vessels, thus protecting leaves from limited quantities of Na+ (see Section 14.10). A high-affinity K+ uptake transporter (PtKUP) was reported to be present in poplar parenchyma tissue (Langer et al., 2002); its transcript level was highly up regulated under K+ deficiency conditions. The driving force for both passive and active mechanisms of xylem unloading appears to be the H+ -ATPase pump (Maathuis and Sanders, 1994; M¨uhling and L¨auchli, 2000). Increased abundance of plasma membrane H+ -ATPase was reported under conditions of low K+ supply in poplar (Arend et al., 2004). Immunolabelling experiments showed that this increase was restricted to vessel-associated cells of the wood ray parenchyma suggesting a key role for the plasma membrane H+ -ATPase in unloading K+ from the xylem stream (Arend et al., 2004). It should be mentioned that light-induced stimulation of H+ -ATPase activity (Kinoshita and Shimazaki, 1999) and associated acidification of the apoplast (Shabala and Newman, 1999) are widely reported. These may hyperpolarize the plasma membrane in leaf parenchyma cells and enable K+ unloading from the xylem via hyperpolarization-activated KIR channels. Enhanced H+ pumping into the xylem also creates a sharp pH gradient, used to transport various nutrients via a variety of H+ -cotransport mechanisms (Figure 9.5). The latter is especially critical for unloading of sugars and anions from the xylem flow.

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Comstock, J.P. and Sperry, J.S. (2000) Theoretical considerations of optimal conduit length for water transport in vascular plants. New Phytologist 148, 195–218. Crafts, A.S. and Broyer, T.C. (1938) Migration of salts and water into xylem of roots of higher plants. American Journal of Botany 24, 415–431. Davis, S.D., Sperry, J.S. and Hacke, U.G. (1999) The relationship between xylem conduit diameter and cavitation caused by freezing. American Journal of Botany 86, 1367–1372. de Boer, A.H. (1997) Fusicoccin – a key to multiple 14-3-3 locks? Trends in Plant Science 2, 60–66. de Boer, A.H., Katou, K., Mizuno, A., Kojima, H. and Okamoto, H. (1985) The role of electrogenic xylem pumps in K+ absorption from the xylem of Vigna unguiculata – the effects of auxin and fusicoccin. Plant Cell and Environment 8, 579–586. de Boer, A.H. and Volkov, V. (2003) Logistics of water and salt transport through the plant: structure and functioning of the xylem. Plant Cell and Environment 26, 87–101. Ding, L. and Zhu, J.K. (1997) Reduced Na+ uptake in the NaCl-hypersensitive SOS1 mutant of Arabidopsis thaliana. Plant Physiology 113, 795–799. Drew, M.C., Webb, J. and Saker, L.R. (1990) Regulation of K+ uptake and transport to the xylem in barley roots – K+ distribution determined by electron-probe X-ray-microanalysis of frozenhydrated cells. Journal of Experimental Botany 41, 815–825. Edwards, D. and Davies, E.C.W. (1976) Oldest recorded in situ tracheids. Nature 263, 494–495. Enns, L.C., McCully, M.E. and Canny, M.J. (1998) Solute concentrations in xylem sap along vessels of maize primary roots at high root pressure. Journal of Experimental Botany 49, 1539–1544. Fisher, J.B., Angeles, G., Ewers, F.W. and LopezPortillo, J. (1997) Survey of root pressure in tropical vines and woody species. International Journal of Plant Sciences 158, 44–50. Fromard, L., Babin, V., Fleuratlessard, P., Fromont, J.C., Serrano, R. and Bonnemain, J.L. (1995) Control of vascular sap ph by the vessel-associated cells in woody species – physiological and immunological studies. Plant Physiology 108, 913–918. Fu, H.H. and Luan, S. (1998) AtKUP1 – a dual-affinity K+ transporter from Arabidopsis. Plant Cell 10, 63–73. Gaymard, F., Pilot, G., Lacombe, B., et al. (1998) Identification and disruption of a plant shaker-like outward channel involved in K+ release into the xylem sap. Cell 94, 647–655. Grignon, C. and Sentenac, H. (1991) pH and ionic concentrations in the apoplast. Annual Review of Plant Physiology and Plant Molecular Biology 42, 103–128. Hartung, W. and Radin, J.W. (1989) Abscisic acid in the mesophyll apoplast and in the root xylem sap of water-stressed plants: the significance of pH gradients. Current Topics in Plant Biochemistry and Physiology 8, 110–124. Holbrook, N.M. and Zwieniecki, M.A. (1999) Embolism repair and xylem tension: do we need a miracle? Plant Physiology 120, 7–10. Keunecke, M., Sutter, J.U., Sattelmacher, B. and Hansen, U.P. (1997) Isolation and patch clamp measurements of xylem contact cells for the study of their role in the exchange between apoplast and symplast of leaves. Plant and Soil 196, 239–244. Kim, S.A., Kwak, J.M., Jae, S.K., Wang, M.H. and Nam, H.G. (2001) Overexpression of the AtGluR2 gene encoding an Arabidopsis homolog of mammalian glutamate receptors impairs calcium utilization and sensitivity to ionic stress in transgenic plants. Plant and Cell Physiology 42, 74–84. Kinoshita, T. and Shimazaki, K. (1999) Blue light activates the plasma membrane H+ -ATPase by phosphorylation of the C-terminus in stomatal guard cells. The EMBO Journal 18, 5548–5558. K¨ohler, B. and Raschke, K. (2000) The delivery of salts to the xylem. Three types of anion conductance in the plasmalemma of the xylem parenchyma of roots of barley. Plant Physiology 122, 243– 254. K¨ohler, B., Wegner, L.H., Osipov, V. and Raschke, K. (2002) Loading of nitrate into the xylem: apoplastic nitrate controls the voltage dependence of X-QUAC, the main anion conductance in xylem-parenchyma cells of barley roots. Plant Journal 30, 133–142. Lacombe, B., Pilot, G., Gaymard, F., Sentenac, H. and Thibaud, J.B. (2000) pH control of the plant outwardly-rectifying potassium channel SKOR. FEBS Letters 466, 351–354.

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10 Solute transport in the phloem Jeremy Pritchard

10.1

Introduction

The colonisation of the land by plants was a fundamental point in the evolutionary history of the earth. The land was a very different environment from that of the sea where the water provided a medium for supporting plants and transport of substances as diverse as nutrients and gametes. The separation of the sources of different nutrients, broadly into water and inorganic ions in the soil and carbon in the atmosphere, drove the diversification of plant form, leading to the evolution of organs such as roots and leaves. The separation of nutritional acquisition now required transport systems, with xylem moving water and minerals from root to leaf while the carbohydrate requirements of autotrophic roots were supplied by the phloem translocation system. Competition for light and other resources increased the spatial separation of leaves and roots such that phloem must often transport sugars over many tens of meters. Since all life on earth is dependent on the carbon fixed by plants, it is worth remembering that nearly all this carbon is transported through the phloem at some time. In natural situations, plants competing with each other or adapting to non-ideal environments allocate their limited carbon resources to different process through the translocation system; the efficiency of its regulation is a major component of their fitness. In addition, increases in crop yield over the last 10 000 years that underpin many developments in the world economy are largely driven by manipulation of the efficiency and direction of the allocation of photosynthate around plants. Thus an understanding of the mechanism of phloem transport and its regulation is essential to inform developments both in conservation and agriculture. In addition, advances in molecular techniques have given phloem the additional function of an information conduit around the plant, implicated in regulation in many developmental processes including the elusive trigger for flowering. This Chapter outlines the broad structure of the phloem system, describing the sieve element, the companion cells and the plasmodesmata that connect them. The contents of the sieve tube are examined, from the more conventional carbohydrates and minerals through to the messenger RNA and proteins that are now complementing and revising our understanding of the phloem. Transport in the phloem system is driven by a bulk flow through the sieve elements, driven by difference in turgor pressure between sources (where solutes are synthesised, e.g. photosynthesis, or produced from breakdown of stored products) and sinks (where transported substances are diluted, metabolised or stored). The processes by which transported

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substances are loaded into and out of the phloem are discussed and the various membrane transporters that are thought to be involved are outlined. Finally, the various organisms that exploit the phloem are briefly considered. Throughout, phloem refers to the translocation system in general, where workers have not discriminated between its different component parts. Where the sieve tube has been examined specifically, it is referred to as sieve tube or sieve element.

10.2

Phloem anatomy

The phloem consists of the main transport pathway, the sieve tube, closely connected to the companion cell by plasmodesmata. The anatomy of the sieve tube reflects its major function – that of facilitating bulk flow of solution. There is a long literature describing this anatomy (e.g. Esau, 1977; Fahn, 1990; summarised in Sheedy et al., 1995). Mature sieve elements consist of elongated cells with elliptical cross section; the length of tubes ranges from 40–500 μm and radius from 5–50 μm depending on species and stage of development. As sieve tubes mature, the end walls develop into sieve plates, which can be transverse or oblique. Each sieve plate contains pores of around 0.5–1.5 μm in length and 1–15 μm in diameter; the numbers of pores range from tens to hundreds and can therefore cover 20–80% of the area of the sieve plate. The sieve plate forms the major resistance to flow of solution along the sieve tube (Nobel, 1983).

10.2.1

Sieve tubes

Sieve tubes do not have nuclei; they contain mitochondria (Wu and Zheng, 2003), although there may be fewer than in normal cells (van Bel et al., 2002), and plastids (Behnke, 1995). Golgi are also lost during transformation from protophloem to mature sieve element (Eleftheriou, 1996). Endoplasmic reticulum (ER) occurs in the sieve element and bean (Vicia faba). ER is attached to the sieve element membrane by ‘clamps’ (Ehlers et al., 2000). Mature sieve element contains many proteins. In legumes, crystalline proteins can be seen in plastids of up to 100 μm in size, while smaller sieve element plastids are seen in most species (Will and van Bel, 2006). No protein synthesis is thought to occur in mature sieve element so that these proteins are likely to have been formed early in sieve element development or to have passed from the companion cell via plasmodesmata. These proteins may have a role in phloem sealing and defence as discussed later but are not present in all sieve elements; for example they were not observed in the sieve elements of soybean (Glycine max) seed pods (Yaklich et al., 2001).

10.2.1.1

Sieve tubes are anucleate

During development, the sieve element looses its nucleus by a process of programmed cell death. The absence of a nucleus in the sieve element is central to an understanding of the operation and regulation of the phloem translocation system. In contrast to the other major transport pathway of the xylem (Chapter 9), the sieve

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element is very much alive and operates as a metabolically active symplastic compartment. While sieve tubes may live for only a matter of weeks in a short-lived annual such as the model plant Arabidopsis thaliana, in longer-lived plants such as palms, (monocotyledonous trees that lack secondary thickening), the sieve tubes probably remain functional for many years (Zimmermann, 1974). During its life, a sieve tube is exposed to a range of damage including chemical, mechanical and radiation (Raven, 1991). Chemical damage is often manifested as oxidative stress – the presence of reactive oxygen. The phloem has a number of adaptations to reduce oxidative damage including the low numbers of reducing sugars, lack of photosynthesis and absence of gas-filled spaces (Raven, 1991). Oxygen tensions are reported to be low in castor bean (Ricinus communis) phloem (van Dongen et al., 2003). The sieve element also contains systems for detoxification of reactive oxygen species (Hancock et al., 2003).

10.2.1.2

Sieve plate blockage

The major axial resistance of a sieve element is the sieve plate. Mechanical damage can cause these plates to become blocked by proteinaceous or membranous material (e.g. Ehlers et al., 2000). Blockage may be accompanied by deposition of callose (Nakashima et al., 2003). Such sieve element blockage can also be a defence response; for example callose deposition was induced in sieve plates of wheat by Russian wheat aphid (Diuraphis noxia), which subsequently restricted flow in the sieve element (Botha and Matsiliza, 2004). Blockage of sieve element can also be adaptive: during dormancy sieve elements are often blocked by callose plugs on the sieve plates of, for example, vines, and removal of callose plugs in the spring is accompanied by resumption of translocation (Aloni et al., 1991). However, not all sieve element blockage is due to callose occlusion of the sieve plate since cessation of translocation induced by cooling, vibration or electric shock was not affected by inhibitors of callose synthesis (Pickard and Minchin, 1992). Recently, a novel protein-based defence system has been identified in legumes. Sieve element proteins can reversibly change confirmation from a condensed to dispersed state. In vivo, this change is induced by membrane damage or permeabalisation and by turgor changes in the sieve element (Knoblauch et al., 2001). The magnitude of the response is reduced in the presence of chelators, leading to the suggestion that the change is induced by Ca2+ in vivo. This was confirmed in vitro where rapid (<1 s) and reversible change in protein conformation could be induced by application and removal of Ca2+ . This hypothesis is consistent with the low concentrations of calcium observed in the sieve element (Will and van Bel, 2006). Influx of calcium into the sieve element could be mediated by the extensive calcium channels in the sieve element membrane (Gayle and Franceschi, 2000). When in the dispersed conformation, the protein is thought to be swept along in the translocation stream and blocks the sieve plate. Such blockage can act as defence against phloem feeding herbivores such as aphids, but provides a more widely applicable mechanism by which loss of phloem material and maintenance of pressure in the translocation system can be maintained following a variety of physical damage (Will and van Bel, 2006). Currently, this elegant defence system has been identified in legumes,

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but while identical mechanisms are not present in other plants (e.g. monocots), they presumably possess a homologous sealing mechanism.

10.2.2

Plasmodesmata

Since the mature sieve elements lack a nucleus, the companion cell provides many of the function of the sieve element. Companion cells contain a full complement of cellular machinery, including nucleus, functional ribosomes, mitochondria and ER (van Bel et al., 2002). Some companion cells have significant vacuoles (Kempers et al., 1998). Crucially to phloem function, sieve elements are connected to companion cells by numerous plasmodesmata. A large proportion of the sieve element solutes, and perhaps all of its mRNA and protein complement, must enter the sieve element through plasmodesmata.

10.2.2.1

Plasmodesmatal structure

The presence and properties of the plasmodesmata are central to the function of the translocation system. Since the mature sieve element lacks a nucleus, all the water and solutes, in addition to necessary proteins and other macromolecules, must pass from the companion cell to the sieve element through the plasmodesmata. Plasmodesmata are pores between cells that allow cytoplasmic continuity between adjacent cells. It is essential for phloem function that water and solutes can move from cell to cell by bulk flow, without crossing a membrane. However, while plasmodesmata are plasma membrane-lined pores, they have a more complex structure. A rod of smooth ER, the desmotubule, runs through the pore, with ER extending into the cytoplasm of adjacent cells. Proteins are thought to extend radially from the desmotubule to the plasma membrane lining the pore, further obstructing the pore (Schulz, 2005). Recent models suggest that the cytoskeleton, represented by actin microfilaments, are closely associated with the desmotubule, with myosin acting as the molecular motor driving large molecules through the pore (Oparka, 2004). An increasing number of proteins are associated with the plasmodesmatal pore and some are implicated in targeting macromolecules to the plasmodesmata (Oparka, 2004). Such targeting mechanisms can be exploited by plant viruses to aid their infection of plants (Lucas, 2006).

10.2.2.2

Plasmodesmatal selectivity

Water and small solutes are able to move through open plasmodesmata with minimal constraint, while larger molecules can be restricted. Use of membrane impermeant dyes indicated a size exclusion limit of 1–7 kDa, depending on species and stage of development (Overall and Blackmann, 1996). Protein trafficking. Since sieve elements lack a nucleus, all their proteins must come through plasmodesmata from companion cells. The well-studied SUT1 sucrose symporter provides a good example. Generally its mRNA is located in the companion cell and associated with the plasmodesmata, while the protein can move

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through plasmodesmata into the sieve element in potato, tobacco, tomato (Kuhn et al., 1997) and wheat (Aoki et al., 2004). Some proteins are able to move relatively unrestricted through plasmodesmata; a number of proteins expressed in the companion cell of rice could transfer through the sieve element (Fukuda et al., 2005). However it is now clear that many proteins and mRNA have restricted movement through the plasmodesmata. In a pumpkin–cucumber (Cucurbita maxima, Cucumis sativus) graft system, phloem serpin proteins and phloem filament protein PP1 both accumulated in source sieve element, but only the larger PP1 could exit into sink sieve element, demonstrating that plasmodesmata selectivity is not simply a function of size (Petersen et al., 2005). Movement proteins can chaperone RNA from cell so cell; for example, the cucumber protein CmPP16 binds to and mediates the movement of RNA through plasmodesmata (Xonoostly-Cazares et al., 1999). Viruses use movement proteins to mediate their own RNA movement from cell to cell (Lucas, 2006). Such chaperones presumably operate by unfolding the RNA or protein and feeding it through the maze of molecules that make up the plasmodesmata. Some proteins larger than the size of exclusion limit can mediate their own transport thought plasmodesmata, for example the phloem thioredoxin from rice (Ishiwatari et al., 1995). Other mechanisms, apart from molecular chaperones, may facilitate movement of large molecules through plasmodesmata, including a role for nuclear proteins, myosin-mediated transport via the cytoskeleton or vesicle trafficking (Oparka, 2004). The trafficking and subsequent transport of RNA into the sieve element may be facilitated by chaperones that form ribonucleoprotein complexes; these chaperones may have motifs conserved across plant species (Gomez et al., 2005). While it is unlikely that there is a single mechanism facilitating movement through plasmodesmata, such movement is fundamental to the operation of the phloem system: understanding movement through plasmodesmata will contribute to the control of virus infection of plants and macromolecular trafficking of signal molecules such as mRNA. Regulation of plasmodesmatal aperture. In addition to the influence of chaperones, plasmodesmata resistance can be altered in other ways. In tobacco trichomes, a difference in pressure between adjacent cells of around 0.2 MPa closed plasmodesmata and prevented the movement of membrane impermeant dye between adjacent cells (Oparka et al., 1992). Plasmodesmatal conductivity is under metabolic control since aperture is altered both by anoxia (Cleland et al., 1994) and metabolic inhibitors (Wright and Oparka, 1997). Following drought, increased transport of solutes to pea root tips was associated with an increase in the cross-sectional area of plasmodesmata in the root tip (Schulz, 1994). Deposition of callose has also been implicated in the closure of plasmodesmata (Radford et al., 1998). Cotton fibre development was regulated by opening and closing of plasmodesmata, a process that correlated with callose deposition and degradation at the plasmodesmata collar (Ruan et al., 2004). A coarser level of control can be exercised by the simple presence or absence of plasmodesmata; for example as observed in symplastic and apoplastic loading (van Bel and Gamalei, 1992) and in the reduction in plasmodesmata density during extension growth (Seagull, 1983).

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Phloem composition

The sieve element is a single cell and regulates its composition like any other. However, the sieve element needs to transport specific solutes at the same time as accumulating sufficient solutes to generate the turgor needed to drive flow; together with its close relationship with the companion cell, this has made the sieve element subject to more stringent selection pressures than many other cell types. Accordingly, its composition varies with species, environment and development. Phloem sap consists of water, carbohydrates, amino acids, inorganic ions, proteins, hormones, lipids and various secondary compounds. In addition to the transport of solutes, increased sophistication in analytical techniques is revealing another role as an important route for transduction of developmental information around the plant.

10.3.1 10.3.1.1

Carbohydrate Sucrose

Much of what we know about sieve element composition is influenced by the sampling methods used to extract the sap. The use of phloem feeding aphids to tap relatively pure phloem sap has been used extensively (Kennedy and Mittler, 1953; Fisher and Frame, 1984). The stylet of the large willow aphid, Tuberolachnus salignus, can be cut with a razor blade, and it produces around 1 μl of sap per hour (Mittler, 1957). Willow phloem sap was dominated by sucrose with concentrations between 50–150 mM (Mittler, 1958; Weatherly et al., 1959). Sap produced through aphid stylectomy on the American basswood tree Tilia Americana had sucrose concentrations varying between 150–750 mM depending on the stage of development (Hill, 1962). Sap extracted from incisions in the bark of the white ash (Fraxinus americana) indicated that the major carbohydrate in phloem sap was sucrose (Zimmermann, 1957). In maize, stylectomy using brown plant hoppers allowed measurement of phloem sucrose concentrations of 900 mM, and it was the only sugar detected (Oshima et al., 1990). Sap collected by stylectomy from the sieve element of arabidopsis was dominated by sucrose (Deeken et al., 2002). Similarly, in rice, phloem carbohydrate was dominated by sucrose (Fukumorita and Chino, 1982), but raffinose, mannitol stachyose and some sugar alcohols were detected in phloem exudate from incisions in 16 tree species, with sucrose present in exudates from all species tested (Zimmermann, 1957, 1958). Due to the ease of collecting phloem sap from incisions in its bark, much work has been done on the phloem composition of R. communis; its phloem sap carbohydrate is also dominated by sucrose (Hall and Baker, 1972; Mengel and Haeder, 1977; Smith and Milburn, 1980a). Similarly, sucrose formed 95% of the total sugars in phloem exudates of tree tobacco (Nicotiana glauca) (Hocking, 1980). Facilitated exudation in legumes can provide samples enriched in phloem sap and these studies also confirm sucrose as the major phloem carbohydrate in this species (Pate et al., 1974).

10.3.1.2

Other carbohydrates

While sucrose is the main carbohydrate in most species examined, this is not always the case. Reducing sugars are not detected in sieve element sap, probably associated

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with need to minimise oxidative damage (Raven, 1991). In peach (Prunus persicae), sieve element sap collected from aphid stylets contained sucrose at 140 mM but total carbohydrate was dominated by sorbitol with a concentration of 560 mM (Moing et al., 1997). In F. americana, trace amounts of raffinose were detected (Zimmermann, 1957). In both olive (Liakopoulos et al., 2005) and celery (Noiraud et al., 2001), mannitol is the major translocated carbohydrate.

10.3.2

Inorganic ions

The major cation in phloem sap is potassium in species as diverse as willow (Hoad and Peel, 1965), castor bean (Hall and Baker, 1972; Smith and Milburn, 1980a), G. max (Fellows et al., 1978), Lupinus albus (Pate et al., 1979), N. glauca (Hocking, 1980), rice (Oshima et al., 1990), barley (Gould et al., 2004b) and arabidopsis (Deeken et al., 2002; Hall et al., 2006). It is a general observation that Na+ is present in phloem sap at much lower concentrations than K+ (Hoad and Peel, 1965; Hall and Baker, 1972; Pate et al., 1979; Smith and Milburn, 1980a). A low level of sodium in the phloem has been suggested to be an adaptation to protect young growing tissues from Na+ accumulation, and may be a charactertistic of more salt tolerant plants (Munns, 2005). Despite the low levels of Na+ in the phloem, it has been suggested that Na+ re-circulation out the leaf in the sieve element is a mechanism to reduce sodium levels in the leaf based on the observation that ethylene diamine tetraacetic acid (EDTA) facilitated phloem exudate had increased Na+ levels during salt stress in arabidopsis (Berthomieu et al., 2003). Similarly, sodium concentrations were elevated in castor bean phloem following salt stress (Zhong et al., 1998). However other data contradicts this view; in arabidopsis, sieve element Na+ concentrations were unchanged following exposure of plants to 50 mM NaCl (Hall et al., 2006). In castor bean (Hall and Baker, 1972; Smith and Milburn, 1980a) and Lupinus species (Pate et al., 1974), the Mg2+ /Ca2+ ratio was high. Calcium is at low concentrations in phloem sap and usually considered to be relatively immobile in the phloem (Jeschke and Pate, 1991). The relative phloem immobility of calcium can lead to build up of the ion; for example, in barley, calcium built up in older parts of the leaves, whereas potassium remained low as it was exported in the phloem (Fricke et al., 1994). Calcium deficiency or accumulation can lead to physiological problems such as blossom end rot (Guichard et al., 2001).

10.3.2.1

Variation in sieve element composition

Solute composition of phloem is not constant and varies between different plant species, during development and following environmental perturbation (e.g. Smith and Milburn, 1980a; Merritt, 1996; Gould et al., 2004b;). In maize, variation in root extension over the day-night cycle was interpreted as resulting from variation in translocation of carbohydrate through the phloem to the root (Muller et al., 1998). However, diurnal variation in sieve element composition is not common to all plants, since absence of photoassimilate during the night can be buffered by carbohydrate uptake from storage tissues (Weatherly et al., 1959). Low light reduced sieve element sucrose in Sinapis alba from over 300 mM to below 30 mM (Mittler, 1958). Large

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changes in sieve element sucrose occurred in T. americana around bud break (Hill, 1962). Like many solutes, nitrogen concentration in the sieve element is not fixed and varies with position, development, environment and genotype. Such differences have important implications for our understanding of the mechanisms underlying regulation of phloem composition. The high levels of phloem glutamine in pre-tuber potatoes declined during filling (Karley et al., 2002). In pea, there were significant differences between sieve elements on the upper and lower side of the petiole, but no difference in the concentration of amino acids between primary, secondary and tertiary veins was observed (Sandstr¨om and Pettersen, 1994). In castor bean, total sieve element organic nitrogen was unaffected by the type of nitrogen supplied, but there was an effect on glutamine, serine and aspartamine (Allen and Smith, 1986). In oats and barley, total sieve element amino acids varied during development (Weibull, 1987); a similar pattern was seen in rice (Fukumorita and Chino, 1982) with high concentrations in young plants, lower in more mature plants and increased concentrations again as plants matured. In willow, sieve element amino acid concentrations decreased during bud burst from around 20 mM to 3 mM, but increased again in older plants (Mittler, 1958). It is a general observation that amino acid levels are elevated in the sieve element following drought (Ponder et al., 2000; Hale et al., 2003). It has even been reported that atmospheric pollution can affect phloem amino acid concentrations (Bolsinger and Fluckiger, 1989).

10.3.2.2

K + /sucrose reciprocity

While studies have focused on the absolute concentration of solute in the phloem, it is clear that this is not fixed and varies with environmental conditions. Early studies on willow sieve element composition identified K+ and sucrose as the major osmotica in sieve element sap and also noted that sucrose and K+ could partially replace each other so that under conditions of low sieve-element sucrose, K+ increased and vice versa (Hoad and Peel, 1965). This reciprocity was subsequently confirmed in castor bean (Smith and Milburn, 1980a) where a dark-induced decrease in sucrose was partially compensated by an increase in K+ concentration. Such a relationship is important when carbon supply is limited, as it allows maintenance of turgor and therefore continued sieve element transport. Conversely, there was no change in sucrose composition of castor bean sieve element when sieve element K+ was increased (Mengle et al., 1977). These observations led to a greater understanding of the role of potassium in the mechanism of sucrose loading into the sieve element (Philippar et al., 2003).

10.3.3

Nitrogen

Transport of amino acids appears to be one of the major functions of the translcoatiosn system. Like many solutes, the concentration and type of amino acids varies with species, environment and development. Sieve element amino acid concentrations have been reported from 49 mM in arabidopsis (Zhu et al., 2005) to over 600 mM in rice (Fukumorita and Chino, 1982). Total amino acid concentration of

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alfalfa sieve element sap was increased by drought (Girousse et al., 1996); this was a regulated change since asparagine concentration unaltered despite changes in some other amino acids. Nitrate is usually present in lower concentration in the sieve element, for example, in castor bean at 3 mM (Smith and Milburn, 1980a). Sieve element nitrate can vary according to whether NH 4 + or NO 3 − was supplied to the roots (Allen and Smith, 1986), and in N. glauca it varied over the diurnal cycle (Hocking, 1980). Particular attention has been paid to sieve element amino acid composition, since it is important in affecting the performance of sap-feeding herbivores (Douglas, 2003) and regulation of seed quality (Sugimoto et al., 1998). There was no difference in the sieve element amino acid concentration of barley and oats: both were dominated by aspartic acids, glutamic acids, glutamine and serine but sieve element sap was low in methionine, glycine, histidine and tryptophan (Weibull et al., 1986; Weibull, 1987; Ponder et al., 2000). A broadly similar pattern was seen in the amino acid composition of a number of brassica species (Weibull and Melin, 1990), S. alba, (Corbesier et al., 2001), Dactylis glomerata, Arrhenatherum elatius (Hale et al., 2003) and arabidopsis (Zhu et al., 2005).

10.3.4

mRNA

Despite the observation that mature sieve elements lack a nucleus, RNA is detected in phloem (e.g. Sasaki et al., 1998; Xoconostle-Cazares et al., 1999; Asano et al., 2002; Nakazono et al., 2003; Vilaine et al., 2003). Analysis of sieve element cDNA provides information about gene expression in the companion cell. There are two reasons why mRNA might be present in the sieve element. First, it may be dragged by bulk flow of solution from companion cell to sieve element and therefore passively reflect gene expression in the companion cell, for example the thioredoxin and actin mRNA detected in the sieve element of rice (Sasaki et al., 1998). Second, some mRNA may have a role in long-distance transfer of information, termed macromolecular trafficking (Chen and Kim, 2006). The localisation of micro RNA to phloem tissues suggests that these novel molecules may also regulate plant development through the translocation system (Yoo et al., 2004). Immunolocalisation and in situ hybridisation can provide useful information on mRNA location in phloem tissue (e.g. Ruiz-Medrano et al., 1999; XoconostleCazares et al., 1999); however, these approaches preclude the identification of genes or proteins not predicted by existing models. An alternative approach is to identify all genes expressed in phloem tissue. Such studies are limited by the relatively small proportion of phloem tissue in any plant organ and the difficulty in obtaining pure or enriched samples. Insufficient sap could be obtained from cut aphid stylets (Pritchard, 1996) to make cDNA libraries, in part because sieve element sap has a very low mRNA concentration (Doering-Saad et al., 2002). The expression of phloem-specific genes – as compared to genes that are expressed in other tissues as well – can be usefully studied at the intracellular, intercellular (e.g. companion cell to sieve element) and long-distance (e.g. source and sink) levels (Vilaine et al., 2003). Genes can be put into different functional classes, depending on current

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annotations, although the process is limited since some genes will be in more than one functional class and annotations are currently incomplete.

10.3.4.1

Protein metabolism message

In maize, laser micro-dissection was used to sample vascular tissues; transcript for protein from a ribosomal subunit was detected (Asano et al., 2002). The presence of this message does not necessarily reflect the presence of the corresponding protein; no ribosomal proteins have been detected in sieve element sap from a range of species including C. sativus (Walz et al., 2004), Triticum aestivum (Hayashi et al., 2000) or R. communis (Barnes et al., 2004). Ubiquitin proteins are frequently found in phloem tissues or sieve element sap (e.g. Hayashi et al., 2000; Barnes et al., 2004; Walz et al., 2004). Ubiquitin and proteasome-dependent proteolysis is required for the regulation of protein turnover (Ingvardsen and Veierskov, 2001). In the phloem, these systems have been hypothesised to be involved in protein stability (Schobert et al., 1995) or potentially in signalling (Vilaine et al., 2003). Most studies examining phloem gene expression have noted the presence of gene transcripts implicated in protein turnover and degradation (Nakazono et al., 2003; Vilaine et al., 2003). Transcripts sharing homology to an ubiquitin-conjugating protein from Capsicum annuum, a 20-S proteasome beta subunit and an ubiquitin-conjugating enzyme from A. thaliana were found in sieve element sap from R. communis (Doering-Saad et al., 2006).

10.3.4.2

Structural genes and cell-wall enzymes

Phloem transcripts often have significant homology with genes associated with cell structure. Broadly, these can be divided into genes coding for extracellular wall components and those involved in cytoskeletal formation. Profilin binds actin, and its transcript was present at 15-fold greater concentrations than actin in castor bean sieve element sap (Schobert et al., 1998; Barnes et al., 2004). Profilin has been hypothesised to prevent actin polymerisation and therefore to be responsible for the absence of microtubules in sieve element cells (Toth et al., 1994). A potential actin-binding protein, myosin, was also found in the phloem sap (Doering-Saad et al., 2006). Myosin proteins have previously been located in the sieve element and have been hypothesised to have a role in altering the size exclusion limit of plasmodesmata (Baluska et al., 2001; Oparka, 2004). Message for a synaptobrevinlike protein was identified in castor bean (Doering-Saad et al., 2006) and could have a role in vesicle trafficking through plasmodesmata. A number of transcripts encoding cell-wall proteins were found in castor bean and celery phloem (Vilaine et al., 2003; Doering-Saad et al., 2006). These included an extensin mRNA, previously reported to be constitutively expressed in the phloem (Shirsat et al., 2003), and UDP glucose dehydrogenase mRNA, also localised to the vascular tissue of Phaseolus vulgaris (Robertson et al., 1996). A message annotated as a precursor for a cellwall invertase was also identified; cell-wall invertases have been previously localised to potato phloem (Hedley et al., 2000). A number of these structural genes have been reported to be induced by stress, illustrating the difficultly of assigning specific, individual functions to genes. For example, chitinase precursors involved in cell-wall

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remodelling are widely implicated in interactions with various pathogens (Kasprzewska, 2003).

10.3.4.3

Interaction with DNA/RNA

Since the sieve element is strongly implicated in long-distance transport of RNA message, it is not surprising to find messages with annotations related to DNA or RNA binding (Nakazono et al., 2003; Vilaine et al., 2003). Transcripts for RNAbinding proteins may be chaperones in long-distance transport of other macromolecules (Lucas and Lee, 2004). Others possess annotations implying involvement in regulation of transcription, DNA binding and RNA processing which might be expected to occur in companion cells and not in the anucleate sieve elements. However, if mRNA moves through the phloem sap as information, then RNA binding and chaperoning systems will be important in sieve element function.

10.3.4.4

Carbohydrate metabolism

The phloem pathway is central to carbohydrate transport and cDNAs from a castor bean library-encoded protein with involvement in carbohydrate metabolism (Doering-Saad et al., 2006). These included a message for sucrose synthase, a protein previously located to companion cells in maize (Nolte and Koch, 1993) and may be involved in processing phloem-translocated sucrose (Komatsu et al., 2002; Konishi et al., 2004).

10.3.4.5

Redox – oxidative stress

In a long-lived, anucleate yet highly active cell type, such as the sieve element, oxidative stress is a potential problem (Raven, 1991; Walz et al., 2002). In celery, many sequences were found related to oxidative stress (Vilaine et al., 2003), while in castor bean there were transcripts which had functions related to oxidative stress, including ascorbate peroxidase and thioredoxin (Doering-Saad et al., 2006). Thioredoxin is one of the dominant proteins found in the sieve element and may also be involved in chaperoning proteins through plasmodesmata (Ishiwatari et al., 1995).

10.3.4.6

Amino acid metabolism

cDNA libraries encode proteins with roles in amino acid metabolism, including a methionine synthase, a glutamine synthase and an ornithine carbamoyl transferase (Hoffmann-Benning et al., 2002; Doering-Saad et al., 2006). Transcripts for glutamine synthase were up regulated by infestation by phloem-feeding aphids in both tobacco leaves (Voelckel et al., 2004) and celery (Divol et al., 2005).

10.3.4.7

Transport

Aquaporins, a sulphate transporter, a chloride channel and a proton ATPase were founds in a phloem library from maize (Nakazono et al., 2003). The presence of mRNA for a sucrose transporter, an aquaporin and a proton ATPase was found using aphid stylectomy to sample the sieve element, followed by RT PCR (DoeringSaad et al., 2002). Both message and protein were localised to spinach phloem (Fraysse et al., 2005). However, aquaporins were not selectively expressed in celery

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phloem tissue (Vilaine et al., 2003). A castor bean phloem library contained an ATP-binding cassette (ABC) and a nitrate transporter cDNA, along with a cyclic nucleotide-gated ion channel, perhaps transporting K+ (Doering-Saad et al., 2006). This transporter has been noted to interact with the Ca2+ calmodulin-binding system previously observed in phloem (Hayashi et al., 2000; Barnes et al., 2004; Nakamura et al., 2006). Two transcripts from a castor bean phloem library had homology with components of a membrane ATPase, which operates in the sieve element–companion cell complex (Doering-Saad et al., 2002).

10.3.4.8

Interaction with the environment

The phloem may be the site of interaction with many pathogens and is also a potential site of signal transduction during environmental perturbations. A cysteine-proteinase inhibitor homologue was up regulated following aphid infestation in sorghum (ZhuSalzman et al., 2004). A transcript for a homologue to an arabidopsis diseaseresistance protein occurred in castor bean and rice (Asano et al., 2002; Zhu-Salzman et al., 2004; Doering-Saad et al., 2006). Message for similar proteins were up regulated by aphids in sorghum (Zhu-Salzman et al., 2004). Proteins for disease resistance have been found in phloem (Hoffmann-Benning et al., 2002), including an ultraviolet-B repressible protein from Pisum sativum. A number of stress-related transcripts, for example glutathinone transferase, often implicated in xenobiotic metabolism, was seen in celery (Vilaine et al., 2003) and rice phloem (Nakazono et al., 2003).

10.3.5

Proteins

Since the mature sieve element has no nucleus it might be expected to contain few proteins. However, there are well over 200 proteins in the sieve element, and increasing numbers of these are being identified and are conserved across very different plant species (Walz et al., 2004), reflecting the central importance of the sieve element to plants. There is emerging information on the phloem protein complement (Hayashi et al., 2000; Hoffmann-Benning et al., 2002;Vilaine et al., 2003; Barnes et al., 2004; Walz et al., 2004; Giavalisco et al., 2006).

10.3.5.1

Oxidative stress

The long life-time of the sieve element in the absence of a nucleus makes it important to maintain its function in the face of abiotic and biotic stress. For example in palm trees, sieve elements may remain functional for over 100 years (Zimmermann, 1974). Plants with such longevity have a number of potential strategies to reduce the damaging effects of reactive oxygen species. It has been suggested that low oxygen tension is not simply a consequence of high metabolism and tissue locations, but also a positive adaptation to reduce the effect of reactive oxygen species (Raven, 1991). Less passive strategies to alleviate the damage of oxidative stress include thioredoxin and ascorbate systems, the proteins of which are frequently found in the sieve element – for example the full machinery for synthesis of ascorbic acid

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has been demonstrated in the phloem sieve element of Cucurbita pepo and Apium graveolens (Hancock et al., 2003).

10.3.5.2

Defence

As well as damage originating from within, the sieve element is often the target for many organisms and possesses many proteins which may be involved in defence. These include the filamentous proteins PP1 and PP2 that contribute to sieve element sealing at the sieve plate in some species. In addition there are proteins associated with limiting pest damage, such as lectins (Dinant et al., 2003), and components of the myrosinase/glucosinolate defence mechanism (Kehr, 2006). A cysteine–proteinase inhibitor was found in a protein library from castor bean (Barnes et al., 2004), and a homologue was up regulated following aphid infestation in sorghum (Zhu-Salzman et al., 2004) and brown plant hopper infestation of rice (Yuan et al., 2005). Proteins annotated as disease-resistant proteins have been found in phloem by other researchers (Hoffmann-Benning et al., 2002).

10.3.5.3

Calcium and sieve element structure

The importance of calcium to many sieve element processes including calloseinduced sealing (Gayle and Franceschi 2000) and forisome function is consistent with calcium-binding proteins such as calmodulin (Nakamura et al., 2006). Profilin is also often found in the sieve element and thought to determine the polymerisation status of the cytoskeleton, including actin, within the phloem continuum (Schobert et al., 2000). It is increasingly apparent that the sieve element is an important and highly regulated conduit for information in the form of proteins and mRNA, so it is not surprising to find a range of proteins in the sieve element that bind and potentially chaperone other macromolecules; these include RNA-binding proteins, metallothionins, cyclophilins and ubiquitins (Schobert et al., 1995, 1997; Barnes et al., 2004).

10.3.5.4

Metabolism

The focus so far on the sieve element as a conduit for water and dissolved solutes and its absence of a local guiding nucleus has perhaps overlooked its potential role in supporting significant metabolism. One reason for this might be that high rates of bulk flow might not be expected to facilitate a stable enzyme complement in the sieve element. However, it is not unreasonable to expect that such proteins are tethered by bindings analogous to those observed for forisomes by van Bel and co-workers (Ehlers et al., 2000). While a sieve element lacks a nucleus and protein synthesis, it still has ability to perform a wide range of metabolism (Giavalisco et al., 2006) and possesses enzymes including those for ascorbate synthesis (Hancock et al., 2003) and alkaloid biosynthesis (Bird et al., 2003). The full complement of enzymes for glycolysis have been located to the phloem and speculated to have a role in callose synthesis (Geigenberger et al., 1993). Several proteins consistent with a role in the metabolism of amino acids have been located to the phloem including methionine synthase, glutamine synthase and ornithine carbamoyltransferase, which were found in sieve element sap (Hoffmann-Benning et al., 2002). Sucrose synthase protein has

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been located to companion cells in maize (Nolte and Koch, 1993) and Phosphoenolpyruvate carboxykinase was localised to cucumber sieve elements (Chen et al., 2004). In future, a more subtle role for the sieve element in regulation of plant growth and development may become apparent as advantages of having both transport and synthetic capabilities are revealed.

10.3.6

Macromolecular trafficking

Despite the anucleate nature of the sieve element, it still requires proteins to remain functional; the function of mRNA in sieve element sap has been less easy to explain. As ideas about the complexity and the selectivity of the plasmodesmata have developed, the suggestion that RNA transport through the phloem has a functional significance has gained experimental support. Far from being passively swept out of a companion cell in bulk flow, the RNA in the sieve element may have significant regulatory effects in the sinks to which it is transported (Lough and Lucas 2006). In this regard, it is interesting that RNAase activity was low in sieve element sap, consistent with the stability of the message necessary for it to move long distances intact (Doering-Saad et al., 2002). mRNA can indeed move considerable distances in the phloem and have important effects on both leaf development and flowering. In situ hybridisation demonstrated the long-distance movements of mRNA through grafted tissue and subsequent alteration of leaf morphology (Ruiz-Medrano et al., 1999). In transgenic tomato, mutant RNA could move through a graft of wild-type tissue and generate the mutant phenotype on new growth (Kim et al., 2001). Similarly, gibberellin-insensitive RNA selectively transported into the sieve element were able to move through graft tissue to generate an altered leaf phenotype (Haywood et al., 2005). A major goal of plant physiology has been to link perception of day length with the signal for flowering – florigen (Zeevart, 1976). Phloem-borne RNA may prove to be this signal. RNA for the transcription factor Constans was driven by a companion-cell-specific construct and was able to move through a graft and bring forward the time of flowering (Ayre and Turgeon, 2004). Similarly, light treatment of a single arabidopsis leaf could initiate flowering and correlate with the trafficking of the mRNA of the flowering locus gene to the shoot apex (Huang et al., 2005).

10.4

Sieve element water relations

Experiments investigating sieve element/phloem composition have often exploited exudates obtained from cut aphid stylets or incisions (Fisher and Frame, 1984; Pritchard, 1996). Implicit in these techniques are the observation that the sieve element is under positive pressure. Despite a clear consensus that pressure differences were responsible for driving sap flow through sieve element as outlined in the M¨unch hypothesis (Luttge and Higginbotham, 1979), direct evidence has only recently become available. Direct measurement of turgor pressure in sieve element from roots of barley was consistent with bulk flow driven by turgor pressure (Gould et al., 2004a).

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Real-time observation of movement of sieve element sap confirms bulk flow of solution in sieve tubes (see www.uni-giessen.de/∼gf1114/rgvbel/longmov.html).

10.4.1 10.4.1.1

Sieve element water relations Sieve element osmotic pressure

While it is difficult to compare data in the literature because of differences in experimental conditions or collection methods, sieve element osmotic pressure of unstressed or control plants at ground level is relatively constant between 1.0 and 2.0 MPa, for example 1.3–1.5 MPa in rice (Fukumorita and Chino, 1982), 1.15 MPa in N. glauca (Hocking, 1980), 1.25–1.45 MPa in castor bean (Mengel and Haeder, 1977; Smith and Milburn, 1980a), 1.4–1.9 MPa in barley (Ponder et al., 2000; Gould et al., 2004b,), 1.1 MPa and 1.3 MPa in D. glomulata and A. elatus, respectively (Hale et al., 2003), and 1.26 MPa in arabidopsis (Hall et al., 2006). The constancy of osmotic pressure, despite variation in the underlying solute composition, reflects the importance in maintaining sieve element turgor and underlines its central role in translocation. The observation that sieve element sap has a higher osmotic pressure than most other cells in the plant, as illustrated by the observation that sieve tube exudate could plasmolyse leaf parenchyma cells (Curtis and Asai, 1939), is important in the pressure flow hypothesis. Exudation rate decreased in castor bean following excision of photosynthesising leaves, a treatment that reduced the osmotic pressure of the sap (Smith and Milburn, 1980b). In bean, calculation using osmotic pressure values indicated a turgor difference of 0.2 MPa between source sieve element and sink, values consistent with the observed velocity of translocation (Housley and Fisher, 1977). The role of turgor pressure in driving translocation is elegantly demonstrated by following the rate of sap exudation from the stylets of aphids feeding on willow. The rate of sap exudation could be increased by pressurising the xylem or abolished by manipulating xylem (apoplast) water potential with mannitol solutions (Peel and Weatherley, 1963). These observations additionally demonstrate that the sieve element cannot be considered in isolation and it operates in a whole plant context. The linkage between xylem and phloem water relations is clearly demonstrated by the observation that diurnal variation in phloem turgor (as deduced from changes in exudation rate) was abolished if transpiration was prevented (Peel and Weatherely, 1962). A 0.5–1.0 MPa osmotic gradient was measured between source and sink in castor bean and was abolished in the dark (Milburn, 1974). In N. glauca there was an osmotic gradient of 1 MPa over a height of 4.5 m (Hocking 1980).

10.4.1.2

Sieve element turgor pressure

Indirect methods. Despite its central role in driving phloem transport, few studies have been able to directly measure sieve tube turgor. Indirect methods, such as plasmolysis or calculation from osmotic pressure, are not reliable due to the assumptions that must be made. An alterative strategy has been to use the Hagen–Poiseulle relationship describing the forces driving a fluid along a tube, calculating sieve element pressure from the rate of exudation of sieve element sap from cut aphid stylets.

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Generally, these methods provide values that appear feasible but are confounded by assumptions about the minimum stylet diameter. Exudation from stylets on willow gave pressure values between 2 and 4 MPa (Mittler, 1957) while in wheat, calculated sieve element pressures ranged from 0.2 to 4 MPa (Pritchard, 1996). Direct methods. A few studies have successfully overcome the technical difficulties in accessing the sieve element and been able to measure directly the turgor pressure of the sieve tube. These have provided the information predicted by the M¨unch hypothesis, demonstrating that sieve elements are indeed under substantial positive pressure and there can be a significant gradient of pressure along the phloem. Phloem pressure was directly measured in mature oak trees, revealing a high pressure of 1.4 MPa at 1.5 m above the ground (Hammel, 1968); this pressure was significantly higher (1.55 MPa) at 6.3 meters demonstrating a small pressure gradient to drive flow of around 0.03 MPa m−1 . This gradient is consistent with a low resistance of the sieve element in the mature, conducting sieve element that was examined. In Ecballium elatius, the squirting cucumber, turgor pressures ranging from 0.03 to 0.8 MPa were measured (Sheikholeslam and Currier, 1977a). Gradients in turgor in favour of flow between source leaf and sink of around 0.43 MPa were measured, and following a drought stress of over 1.8 MPa a positive turgor remained in the sieve tube and the gradient-driving flow from source to sink was reduced to 0.18 MPa, consistent with a reduction in assimilate transport (Sheikholeslam and Currier, 1977b). In willow, turgor pressure in the sieve tube was measured by attaching a manometer to cut aphid stylets (Wright and Fisher, 1980). A turgor of between 0.51 and 0.93 MPa was measured, but these values did not show a good agreement with values calculated from osmotic pressure and water potential. However, osmotic pressure was calculated from sucrose concentrations alone, an assumption that could have been distorted by the potential for replacement of sucrose by K+ and so an underestimation of sieve element osmotic pressure. Direct measurement of turgor in the sieve element of the pedicel and grain of wheat revealed a high turgor pressure in the pedicle of 2.4 MPa decreasing into the grain to 1.2 MPa (Fisher and Cash-Clark, 2000). The high driving force over a short distance represents a high resistance of the pathway in this unloading step in comparison to the low resistance of the pathway inferred in oak (Hammel, 1968). Similarly, in barley roots, there was a turgor difference of about 0.7 MPa driving solute from sieve element to sink, again high in comparison to the lower resistance seen in pathway sieve element (Gould et al., 2005). In the barley system, sieve tube turgor was 1.6 MPa and was reduced when K+ was in short supply, emphasising the influence of the environment of sieve tube turgor (Gould et al., 2004b).

10.4.2

Flow in the phloem

It is established that transport of water and solutes in sieve elements occurs through a pressure-driven bulk flow (van Bel et al., 2002; Gould et al., 2004b). The properties of the sieve element ‘pipe’, the solution within it and the relative pressure differences of connected sources and sinks determine both the rate and direction of transport.

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Many models have built upon the suggestion of M¨unch and are providing increasing complex ways in which the phloem works. The basic assumptions of these models are of flow through a tube as described in the Hagen–Poiseuille relationship (Eq. 9.1; see Section 9.2.2). This relationship states that the flow is a function of the pressure difference at each end of the tube, the viscosity of the fluid in the tube, the tube elasticity and solution osmotic pressure and resistance which is a function of the length of the tube and, importantly, the fourth power of tube radius. In practice, the resistance of a sieve element is probably dominated by the resistance of the sieve plate (Nobel, 1983; Thompson and Holbrook, 2003) and not by the open lumen of the tube. Essentially, flow along the phloem is a function of a driving force and a resistance. The driving force is the osmotically generated difference in turgor pressure. Resistances to both water and solutes operate at a number of levels, ranging from resistance of membrane barriers (e.g. carriers and channels), plasmodesmatal density and conductance, the diameter, number and openness of the sieve plates, and finally the number of functional sieve elements. The anatomy of the phloem system has been successfully used to predict its transport properties using the Hagen–Poiseuille relationship (Sheehy et al., 1995). Bulk flow of solution in the phloem can now be imaged in intact plants using NMR and reveals rates similar to the calculated flow rates (Kockenberger et al., 1997).

10.4.3

Phloem loading

The rate of exudation of phloem sap often remains constant despite manipulations that affect the number of sinks or the availability of solutes to generate turgor. Despite the increase in the availability of photoassimilate during the day, the exudation rate through cut stylets was unchanged over the diurnal cycle in willow (Weatherley et al., 1959). Collection of sap either by cutting or stylectomy imposes a new sink on the plant; despite this increased demand, exudation rate and osmotic pressure of sap from incisions, for example on castor bean (Smith and Milburn, 1980b) or cut stylets on barley (Pritchard, 1996), can remain constant for long periods of time. Taken together, these observations suggest that phloem can alter the loading of solutes and water to keep pace with the altered demands. It is therefore important to understand the mechanism by which solutes are loaded into the sieve element.

10.4.3.1

Symplastic or apoplastic loading?

Solutes move into the sieve element from the companion cell. Entry of solutes into the companion cell can take one of two routes, apoplastic or symplastic (10.1). Apoplastic loading requires that solutes are taken up across a membrane from the apoplast (Figure 10.1A). In symplastic loading, solutes, usually photosynthates, move from their site of production in the mesophyll cells into the sieve element through open plasmodesmata without leaving the symplast (Figure 10.1B). The absence of an open symplastic route anywhere in the pathway means that the route is apoplastic and membranes must be crossed (van Bel and Gamalei, 1992). Apoplastic loading will be the only route if the sieve element-companion cell complex becomes isolated from mesophyll cells (van Bel and Van Rijen, 1994). As a leaf

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ic

Sieve tube pd

pr

Companion cell

mRNA

aq +

H -ATP

suc

Parenchyma cell

Mesophyll cell

n

ch

A

Sieve tube

Companion cell

Parenchyma cell

Mesophyll cell B

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develops, it undergoes transition from sink to source – import stops when plasmodesmata close or are lost in the main veins, and export begins when minor veins begin to load sugars into the phloem (Turgeon, 2006). Further evidence that loading is apoplastic includes the absence of plasmodesmata, making transport across membranes essential. Mapping of the frequency of plasmodesmatal connections, or plasmodesmograms, reveals which route of loading is possible (van Bel and Gamalei, 1992). Examples of both routes can be found. Plasmodesmatal continuity between minor veins and bundle sheath is probably the ancestral state and there has probably been a general, but not exclusive, trend to the apoplastic route (Turgeon et al., 2001). However, an open symplastic route (i.e. presence of large numbers of plasmodesmata in the bundle sheath/mesophyll boundary) does not necessarily indicate a symplastic mode of loading. There was a symplastic plasmodesmatal configuration in the mask flower, Alonsoa meridionalis, but the higher turgor of the sieve element compared to the mesophyll was consistent with apoplastic loading, emphasising the need to combine analysis of both driving forces and resistances (Voitsekhovskaja et al., 2006). Inhibition of apoplastic sucrose loading with p-chloromercuribenzene-sulfonate (PCMBS) reduced phloem loading in apoplastic loaders and sucrose-accumulating species that had a high frequency of plasmodesmata, but had no effect on those species with symplastic configurations (Turgeon and Medville, 2004). Consistent with the different modes of loading, sucrose concentrations are low and relatively constant in the apoplast of symplastic loaders, whereas apoplastic loaders had higher levels of apoplastic sucrose that was increased if export was inhibited (Voitsekhovskaja et al., 2000). Models of loading have been considered in an ecophysiological context. The extra control afforded by the apoplastic route has been suggested to be an adaptation to deal with the need to move large amounts of photosynthate in a short growing season, as found in temperate plants. The symplastic route is interpreted ← Figure 10.1 (A) Schematic representation of apoplastic phloem loading. Sucrose is synthsesised in the mesophyll cell chloroplasts (ch) and moves into the cell wall. Sucrose diffuses through the cell walls of parenchyma cells to the wall of the companion cell. Sucrose is actively loaded into the companion cell through a sucrose proton symporter (suc), energised by outwardly directed proton ATPases (H+ ATP). Once in the companion cell, sucrose can diffuse through open plasmodesmata (pd) into the sieve element. Accumulation of sucrose into the sieve element–companion cell complex lowers its water potential; water entry is facilitated by aquaporins (aq), generating the high turgor that drives translocation. Electrical balance and loading of other ions is mediated by a range on ion channels (ic). Companion cells contain a nucleus (n) while the sieve tube does not. mRNA and proteins (pr) synthesised in the sieve element can move selectively through open plasmodesmata into the sieve element where they are transported in the bulk flow of sap. (B) Schematic representation of symplastic phloem loading. Sucrose is synthesised in the mesophyll cell chloroplasts (ch) and diffuses down its concentration gradient through open plasmodesmata, through the companion cell into the sieve element. A concentration gradient can be maintained if sucrose is polymerised into longer sugars that cannot diffuse back into the companion cell. Accumulation of solutes in the companion cell complex lowers its water potential; water entry is facilitated by aquaporins (aq), generating the high turgor that drives translocation. Electrical balance and loading of other ions is mediated by a range on ion channels (ic) and H+ -ATPases.

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as an adaptation to transporting photoassimilate at a lower, constant rate throughout the season. Consistent with this, increases in light levels were accompanied by an increase in assimilate export from leaves in apoplastic-loading species, whereas symplastic loaders had a more limited carbon export capacity (Amiard et al., 2005). The global distribution of apoplastic and symplastic loading mechanisms is seen as evidence in favour of this hypothesis. Apoplastic loading allows more control over solute uptake, and under low temperature and drought stress some plants are able to switch between symplastic and apoplastic loading (van Bel and Gamalei, 1992). The fact that maize plants subjected to reduced temperatures had an increased number of plasmodesmata at the bundle sheath mesophyll boundary was seen as an adaptation to facilitate assimilate export at low temperatures (Sowinski et al., 2003). However, other studies have found no difference in the response of leaf export to lowered temperature between symplastic and apoplastic loaders (Schrier et al., 2000). A sucrose concentration higher in the sieve element than that in the mesophyll cells is consistent with apoplastic loading since in the presence of an open symplastic pathway, sucrose would tend to diffuse back out of the sieve element. The polymer trap hypothesis, in which small sugars diffuse into the sieve element and are then trapped inside following polymerisation, can explain how high concentrations of sugars can be accumulated within the sieve element against such a concentration gradient (Turgeon, 1996). Elevation of invertase activity in the apoplast increased potato yield, supporting an apoplastic unloading route (Sonnewald et al., 1997). Loading of solutes was reduced by anaerobic treatments of source leaves resulting in reduced sieve element sugars, osmotic pressure and turgor pressure, therefore demonstrating the role of active transport in loading the phloem (Gould et al., 2005).

10.4.3.2

Transporters facilitating apoplastic loading

Sucrose. In order to enter the sieve element–companion cell complex, solute and water must cross the plasma membrane. Such movement is usually through carriers or channels. Transporters for the major classes of phloem solutes have all been localised to phloem tissues. Since sucrose dominates sieve element composition, it is not surprising that sucrose carriers have received much attention. Early observation on the uptake of sucrose by leaf disks showed optimal uptake at acid external pH (Giaquinta 1977) and inhibition of sucrose uptake by PCMBS (Delrot et al., 1980), leading to a model of sucrose uptake through proton – sucrose transporters. In the context of uptake of sucrose from the apoplast, this scheme is consistent with the low pH of the cell wall and the pH of the sieve element of over 7.5. The sucrose transporter is SUT1, identified in many plant species (Kuhn et al., 1999). Complementation studies in yeast have demonstrated that the SUT1 transporter can take up 14 C-labelled sucrose and such uptake is pH dependent and reduced by protonophores such as CCCP (Riesmeier et al., 1992). Down regulation of the SUT1 genes in potato and tobacco led to accumulation of sugars and starch in source leaves (Burkle et al., 1998). Carbohydrate accumulation was more pronounced under high light when, presumably, more sugar is produced in the source leaf and unable to be exported

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(Kuhn et al., 1996). The homologue of SUT1 in arabidopsis, SUC2, was localised to both export and transport phloem using beta-glucuronidase (GUS) localisation (Truernit and Sauer, 1995), while in situ hybridisation localised SUT1 mRNA to source phloem (Reismeier et al., 1993; Kuhn et al., 1997). Arabidopsis plants expressing green fluorescent protein (GFP) driven by a SUC2 promoter showed that AtSUC2 expression correlated with 14 C-sucrose uptake and transition of developing leaves from sink to source (Wright et al., 2003). In wheat, SUT1 protein was localised to the plasmamembrane of loading and transport sieve elements, suggesting a role in both apoplastic loading and retrieval of sucrose (Aoki et al., 2004). While SUT1, or its homologue SUC2, has received much attention, this transporter is a member of a gene family currently containing nine members (SUC1– SUC9); of these, seven appear to be functional in the plant. SUT4 is involved in sucrose loading in minor veins (Weise et al., 2000). SUC3 is expressed in mesophyll cells and may represent an early step in the apoplastic pathway (Meyer et al., 2004). SUC1 and SUC5 are expressed in floral tissues and may have a role in seed development and dehiscence (Stadler et al., 1999; Baud et al., 2005). However, in maize, the homologue of SUC1 – SUT2 – had increased expression in the phloem tissue (Vilaine et al., 2003). In some species, sucrose is not the only primary product of photosynthesis (see Section 10.3.1.2); for example in celery, mannitol is produced and a mannitol/proton symporter was localised to the mature leaves and phloem sugars. The Rosacea accumulates sorbitol in the sieve element as the translocated carbohydrate and sorbitol transporters were localised to the sieve element of source leaves in apple (Watari et al., 2004). The apoplastic loader Plantago major translocates sucrose and sorbitol and has both sucrose (Barth et al., 2003) and polyol (Ramsperger-Gleixner et al., 2004) transporters.

10.4.3.3

H + /ATPase

The sucrose/proton symporters central to phloem loading are energised by the proton motive force (PMF) generated by proton ATPases pumping protons out of the cytosol. The ubiquitous use of PMF in plants is reflected in a wide distribution of these transporters in most tissues. In arabidopsis, 83 primary ATPases are currently recognised in 12 families. The plasma membrane ATPase family contains 12 members; AHA3 has been localised to phloem tissues in a number of species. AHA3 was located to the plasma membrane of the companion cell; lower amounts of this transporter protein were detected on the sieve element membrane (DeWitt and Sussmann, 1995). Laser capture microdissection of vascular tissues of maize indicated elevated expression of a proton ATPase (Nakazono et al., 2003). In castor bean, one ATPase was located to the sieve element while another to the companion cell (Langhans et al., 2001). Expression of PM H+ /ATPases remained high in castor bean phloem during development (Williams and Gregory, 2004). A role in energising sucrose co-transport is supported by co-localisation with the SUC2 transporter in arabidopsis vascular tissue (Truernit and Sauer, 1995). The mRNA for the proton

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ATPase PPA1 was found in the sieve element of barley seedlings, consistent with a companion cell origin for this gene (Doering-Saad et al., 2002). K + transporter localisation. Potassium is perhaps the most important inorganic ion in plants, being required for osmoregulation, as a co-factor, various tropisms and stomatal opening. It is also required as part of the phloem-loading process. There are a large number of potassium channels and carriers in plants reflecting the diverse roles of potassium (Maser et al., 2001; see also Sections 5.2.3, 5.4 and 14.9). A number of these have been localised to phloem tissue. Transcript for a shaker-like K+ channel VFK1 was localised to sieve element in bean (V. faba; Ache et al., 2001). VFK1 mRNA increased under elevated light when photosynthesis was high, and a role in regulation of sugar loading via modulation of membrane potential was suggested. In maize, the K+ channel Kzm1 localised with SUT1 expression, again consistent with a role in regulation of membrane potential (Philippar et al., 2003). In arabidopsis, AKT2/3 GUS expression localised the K+ transporter to the phloem (Deeken et al., 2000), again transcript levels correlated positively with the rate of photosynthesis, being reduced by darkness or lowered CO 2 , which is consistent with the K+ channel having a role in loading of sucrose into the sieve element. This was confirmed in a knockout mutant of AKT2/3 where absence of K+ transporter activity was accompanied by reduced loading of 14 C sucrose and a lowered sieve element sucrose concentration (Deeken et al., 2002). Thus potassium loading into the sieve element–companion cell complex is important both in the transport of potassium in the phloem and in the regulation of the loading process itself. Aquaporins. The lowering of sieve element–companion cell complex water potential by solute accumulation leads to the uptake of water from the apoplast. Aquaporin water channels facilitate the movement of water across membranes and are therefore expected to be present in phloem tissues. Currently in arabidopsis, 35 aquaporins (including nodulin-like genes, tonoplast intrinsic proteins (TIPs) and plasma membrane intrinsic proteins (PIPs)) have been identified. The 13 PIPs can be divided on the basis of their homology into PIP1 and PIP2; PIP2 was detected in older leaves and was believed to facilitate water uptake by the vascular tissues. In spinach, PIP1 was localised to the sieve elements of leaves (Fraysse et al., 2005), consistent with a role in facilitating water uptake into the phloem. Similarly, the GUS promoter fusion of a PIP from the ice plant, Mesembryanthemum crystallinum, was associated with vascular tissue in leaves. Transcripts for PIP1 and PIP2 were located by laser-capture microdissection (LCM) to the vascular tissues of maize (Nakazono et al., 2003). While PIPs have a role in phloem loading, it is not clear how TIPs contribute to water uptake into the sieve element since mature sieve elements lack vacuoles. Transcript for a tonoplast aquaporin was detected in the sieve element sap of barley (Doering-Saad et al., 2002), while TIP mRNA occurred in a phloem-enriched cDNA library in castor bean (Doering-Saad et al., 2006). In oil seed rape, Brassica napus, a TIP was absent from the phloem tissue but expressed in bundle sheath cells suggesting that TIPs can facilitate water flow towards phloem (Frangne et al., 2001). This emphasises that apoplastic loading can occur away from the sieve element–companion cell complex.

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Amino acids. Amino acids are often the form in which nitrogen is transported around the plant and the site of nitrate reduction can be remote from where the amino acids are metabolised. Maize leaf tissue had lower amino acid concentration, in particular asparagine and glutamate, than sieve element sap, implicating an active uptake process (Weiner et al., 1991). Currently about 50 amino acid transporters are known. These have diverse spatial and temporal distribution (Su et al., 2004) and a number are implicated in phloem loading. An amino acid permease (AAP) transcript was detected in phloem sap of rice (Asano et al., 2002), while AAP2 was expressed in the stem of arabidopsis and suggested to have a role in loading amino acids from the xylem to the phloem (Kwart et al., 1993). A role for AAP1 in loading is strongly suggested by its co-localisation with expression of the sucrose symporter SUT1 during source-to-sink transition in potato leaves (Koch et al., 2003). AAP4 was expressed in root phloem and major leaf veins suggesting a role in phloem loading of amino acids. However, loss-of-function knockouts of this gene did not result in any obvious phenotype (Okumoto et al., 2004). This may reflect the general specificity and overlap in function of the amino acid transporters. However a knockout of the neutral amino acid transporter ANT1 produced a significant increase, in sieve element amino acid concentration (Hunt, 2005). However, despite this change in phloem concentration, there were no large differences in whole-plant phenotype. Other transporters are involved in the loading of organic nitrogen; for example, the compatible solute transporter PROT1 was expressed in the phloem tissue of arabidopsis but other closely related members of the family PROT2 and PROT3 were not (Grallath et al., 2005), indicating the functional diversication that can occur in gene families.

10.4.4

Phloem unloading

Once solutes and water are delivered to a sink by bulk flow, unloading occurs. Regulation of the unloading process is central to maintenance of the pressure differences that drives phloem transport and also determines the relative partitioning of phloem contents to competing sinks. Sinks include a wide range of plant structures including developing leaves, filling storage tissues, seeds, fruit tissues and growing roots. Similar to loading, pathways can be apoplastic or symplastic (Figure 10.2). Some tissues, such as the well-studied seed system, are exclusively apoplastic, while other sinks, such as expanding roots or developing fruits, can show developmental or spatial changes in the pathway of unloading (reviewed in Patrick, 1997).

10.4.4.1

Evidence for unloading pathway: root tips

Early and convincing evidence was presented for direct symplastic unloading of sucrose in root tips by the observation that asymmetrically 14 C-labelled sucrose moving to the root tip in the phloem remained intact (Giaquinta et al., 1983). The labelling of recently fixed carbon (RFC) with 11 C showed that its movement into root tips was unaffected by metabolic inhibitors (Farrar and Minchin, 1991), while 11 C-labelling showed that RFC was rapidly accumulated in the root tip region with high plasmodesmatal connections (Pritchard et al., 2004) consistent with absence of

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suc

Sieve tube

H+-ATP

suc

pd

met

Inv sto

Figure 10.2 Schematic representation of phloem unloading. In sinks such as root tips or developing fruits, sucrose in the sieve tube can be unloaded symplastically from the sieve tube through open plasmodesmata (pd). To lower osmotic pressure, thus maintaining a low-sink turgor pressure to facilitate transport, sugars can be consumed in metabolism (met) or polymerised in storage (sto). If plasmodesmata are closed or absent, loading of sugars into sink cells is apoplastic using H+ -ATPase energised sucrose transporters (suc). Alternatively, or additionally, sucrose unloaded into the cell wall may be cleaved for example by invertases (inv) and then taken up actively by other sugar transporters.

membrane barriers in this region. Photoassimilate accumulates in root tips despite the presence of PCMBS, which blocks sucrose uptake from the cell wall (Dick and ap Rees, 1975). Transport of membrane-impermeant dyes from the phloem into the root tip directly demonstrates symplastic movement between sieve element and growing root cells (Duckett et al., 1994). Both osmotic pressure differences (Warmbrodt, 1987) and direct measurement of turgor pressure in sieve elements of growing root cells (Gould et al., 2004a,b) indicate a positive pressure available to drive solution through open symplastic pathways. Cells in root tips are connected by plasmodesmata (Gunning, 1978; Warmbrodt, 1985; Ma and Peterson, 2001) and the density of these are decreased by longitudinal cell expansion (Seagull, 1983). The symplastic domain can change under different solute availability regimes (Patrick and Offler, 1996), a process facilitated by gating behaviour of plasmodesmata (Schulz, 1994; Baluska et al., 2001). Changes in symplastic unloading can be induced; for example the cyst nematode Heterodonta schachtii induces the development of a feeding complex in arabidopsis roots. This complex is symplastically connected to existing phloem, as demonstrated by the movement of GFP of companion cell origin into the new sink (Hoth et al., 2005). Similarly, in tobacco, the haustorium of the holoparasite dodder (Cuscuta sp.) developed into a symplastic sink for sucrose and the phloem tracer dye carboxyfluorocein (Birschwilks et al., 2006).

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There is also an apoplastic unloading route within the growing root tip. In arabidopsis roots, mature, but expanding, cells lose symplastic continuity with the sieve element (Duckett et al., 1994); the transition from symplastically connected to apoplastically isolated cells occurred within the growing region (Freixes et al., 2002). A similar pattern of movement of symplastic tracer was demonstrated in arabidopsis using phloem-localised GFP (Imlau et al., 1999). Symplastic isolation began around 3 mm from the maize root tip whereas maximal growth was at 5 mm and the region of decreasing relative elemental growth rate extended a further 5–10 mm from the root tip (Hukin et al., 2002). The switch in the pathway of transport was accompanied by an increase in the expression of aquaporin facilitating water transport across membranes in the apoplastic region. In storage roots such as potato, there was a switch from mainly apoplastic unloading in extending stolons to symplastic unloading as tuberisation began (Viola et al., 2001). Transcriptomic analysis of the different regions of the root tip can reveal qualitative differences in the complement of organic and inorganic transporters (Pritchard et al., 2005). Recently, the sucrose symporter protein SUC3 was localised to the sieve element of both transport phloem and a range of apoplastic sinks, and a role in regulating the unloading of sucrose into the cell wall was suggested (Meyer et al., 2004).

10.4.4.2

Evidence for unloading pathway: developing fruits

Expanding cells in young developing fruits start out receiving their solutes symplastically from the sieve element through open plasmodesmata. For example, in apple the sieve element–companion cell complex became isolated and in parallel a monosaccharide transporter increased in abundance, consistent with a role in facilitating apoplastic uptake of assimilate. In grape, changes in plasmodesmatal frequency were consistent with a shift from symplastic-to-apoplastic unloading as fruits matured (Xia and Zhang, 2000). Similarly, the high concentrations of sorbitol in the mature fruit and the presence of sorbitol transporters are cited as evidence of apoplastic uptake in apple (Gao et al., 2003). An apoplastic unloading pathway requires that the appropriate enzymes and transporters are present (Sturm and Tang, 1999). While sucrose may be taken up directly from the apoplast, the often-noted presence of invertase in the cell walls of such sinks suggests that sugars are transported as hexoses (Zhou et al., 2003). Sucrose synthase, which reversibly catalyses the formation of sucrose from glucose and fructose, is involved in the unloading of sucrose from the phloem since inhibition of the enzyme in tomato reduced sucrose import (D’Aoust et al., 1999) while its over-expression gave increased loading of sucrose into fruit tissue (Nyugen-Quoc et al., 1999).

10.4.4.3

Evidence for unloading pathway: seed coats

Nutrients reach developing seeds through the phloem. At the maternal/filial interface, solutes are transferred to the apoplast. It is not currently known how active the unloading process is; it could consist of a change in the proportion of leakage to scavenging/reloading or alternatively or additionally it could be due to the operation of specific unloading channels or carriers – but so far no active unloading processes

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have been identified (Patrick and Offler, 2001). The seed-coat apoplast has a high concentration of solutes, which are taken up from the apoplast by a range of proton ATPase-energised transporters (Patrick and Offler, 2001). This system has proved useful for studying apoplastic unloading. For example, despite the membrane step, manipulation of cell-wall osmotic pressure suggested turgor regulation of solute transport from phloem in the seed coat to the apoplast and uptake into the embryo (Patrick, 1993, 1994).

10.4.5

Resource partitioning through the phloem

Differences in turgor and sieve element anatomy can explain flow along a simple one source–one sink system. Developments of this model can also elegantly explain the differential partitioning of phloem solutes between competing sinks. In general, such models consider the degree of regulation of translocation by source or sink and the partitioning of solutes and water between different competing sinks. In one of the simplest models, the translocation system is considered as a single source, connected by open sieve elements to two competing sinks (Minchin et al., 1993). In an analogous manner to electrical models, flow is a function of the pressure-driving force and the resistance of the pathway. These models provided testable hypotheses that allow the location of the regulation to be dissected. A simple split root system in barley provided a physiological version of the model in which recently-fixed carbon (RFC) from the leaf was labelled with 11 C and delivered to both root tips sinks. Import of RFC was reduced when root extension was inhibited (Farrar et al., 1995). In contrast, application of galactose to roots increased the import of RFC to the whole root while extension growth was decreased (Farrar et al., 1994); thus growth is not the only determinant of sink strength. While carbon import was increased to whole roots by galactose treatment, in the root tips import was decreased concomitant with an increase in turgor suggesting a simple biophysical explanation for the change in local (root tip) import (Pritchard et al., 2004). In addition, these models of phloem transport have been used to explain complex changes in morphology, such as branching patterns in roots where the growth of lateral roots could be explained by the physical properties of competing translocation pathways (Bidel et al., 2000). Models are becoming more sophisticated and sometimes predict non-intuitive behaviour; for example, their dual function of the phloem in both longand short-distance delivery of solutes predicts maximum efficiency when pressure differences along the translocation system are small (Thompson, 2006). Abiotic and biotic factors can alter carbon partitioning; for example it is a common observation that the root/shoot ratio increases under drought (e.g. Egilla et al., 2001), an adaptive response caused by a change in the partitioning of carbon through the translocation system. Defoliation of maize resulted in a greater flux of carbon to the roots (Holland et al., 1996), while phloem-feeding aphids altered the carbon and nitrogen partitioning in alfalfa (Girousse et al., 2005). These changes in carbon partitioning are presumably regulated by alteration in the driving forces and resistances within the translocation system, and the control seems to reside in both source and sink (Minchin et al., 1994).

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10.5

261

Exploitation by other organisms

As the phloem transports a wide range of nutrients around the plant, it is not surprising that a range of organisms exploit the conduit as their feeding site. However, the phloem system is far from defenceless and a number of strategies have been outlined in previous sections. In addition, the sieve element is the site for secondary defence compounds as well as being a major route for their transport. For example in poppy, alkaloid synthesis was localised to the sieve element–companion cell complex (Bird et al., 2003). The toxic compounds rutin and cyanide are found in the phloem of cassava (Calatayud et al., 1994). Glucosinolates can be detected in the phloem of arabidopsis (Chen et al., 2001). Despite these defences, a number of organisms exploit and thrive on the sieve element. The adaptations they have facilitate this exploitation and complement our knowledge of phloem function.

10.5.1

Micro-organisms and viruses

A range of viruses exploit the phloem to transport them around the plant (Lucas, 2006). The ability of plasmodesmata to close prevents the systemic spread of virus particles. Thus, the ability of the virus movement proteins to increase effectively the size exclusion limit of the plasmodesmata and thus allow viral RNA to move from cell to cell is central either to the success of the virus or to the ability of the plant to prevent their spread. A fuller understanding of the mechanism of action of this protein will allow a more detailed understanding of plasmodesmatal function and better strategies to be developed for viral control (Scholthof, 2005). In addition, understanding of this process opens up the possibility of engineering plants with altered phloem resistances to certain organisms at particular stages of development, in order to increase crop yields. While viruses can successfully move around a plant in the sieve element, they have the problem that they must be transmitted from the sieve element to another plant. To do this, they often use other organisms that exploit the phloem, including aphids and white flies. Phytoplasmas are bacteria lacking a cell wall that inhabit the sieve element of many plants. They can cause significant damage and loss of yield as they proliferate within the sieve element leading to its blockage (Christensen et al., 2005). They have a minimal genome and, for example, lack sugar transporters and are therefore adapted to life in the high-carbohydrate environment of the phloem (Christensen et al., 2005).

10.5.2

Sap-feeding insects

Homopteran insects such as aphids and white-fly feed directly on the contents of the sieve element. The major problems they face are locating the sieve element, overcoming any defences in the sieve element and developing and reproducing on sap which is not of ideal nutrient composition, being high in carbohydrate and unbalanced in the essential amino acids. There is as yet no consensus as to what cues aphids use to locate a sieve element but sucrose and certain amino acids may

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play a role (Powell et al., 2006). The egestion of saliva into the sieve element sap during feeding provides a mechanism for overcoming plant defences (Tjallingii, 2006; Will and van Bel, 2006). A symbiotic association with the bacteria Buchnera (Douglas, 2006), coupled with a range of behavioural changes, can allow aphids to reduce the impact of the non-ideal diet.

10.5.3

Plants

A number of plants parasitise the sieve element, including dodder (Cuscuta sp.). Dodder is able to access the phloem of its host directly; this symplastic continuity was demonstrated by transfer of sieve-element-specific tracers from host to parasite (Haupt et al., 2001; Birschwilks et al., 2006). Orobanche species also obtain the majority of their carbon from the host phloem (Hibberd et al., 1999), and microscopical examination suggests symplastic continuity between host and parasite (Dorr and Kollmann et al., 1995).

10.5.4

Other organisms

One vertebrate is known to feed directly on phloem sap – the yellow bellied sap sucker (Sphyrapicus varius) is a woodpecker-like bird from North America. It is able to make incisions in the bark of trees and lick the exuding sieve element sap (Blendinger 1999). Presumably, the natural sealing mechanisms are overcome by its saliva in a manner analogous to aphid feeding; it would be interesting to compare the biochemical properties of the saliva of the bird with that of the aphid. Nematode worms form feeding cysts which can constitute significant new sinks on a plant, to which solutes are delivered symplastically; however, there is no direct contact with the sieve element sap and solutes and water are transferred across the haustorial membranes (Hoth et al., 2005).

10.6

Conclusions

In the last few years our understanding of phloem function has been consolidated, and there is now a relatively clear view of the anatomy of the phloem system and the mechanism of pressure-driven bulk flow, driven by cellular water relations. The biochemical machinery that underpins the movement of solutes and water is beginning to be better understood, both in the function of individual transporters and also their location and expression. In addition, there have been recent discoveries concerning the defence mechanisms, by which sieve elements can protect themselves against the many pests and pathogens that target the phloem; a better understanding of these mechanisms and their specificity will inform the development of novel pest control strategies. It is clear that the sieve element is highly active, and although we have a number of specific examples, we have yet to achieve a general understanding of how sieve element composition responds to changes in the environment; continued analysis

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will reveal which of the many responses are species specific and which are fundamental. In this regard, we know little about how the inorganic ion composition of the phloem changes in response to the environment. While simple physical models can explain some of the complexity of delivery of resources such as carbon around the plant, it is increasingly apparent that subtle changes in resistance also have a role. The development of our understanding of the structure and function of the plasmodesmatal pore is central to this, changing the resistance to flow through phloem domains but also by its selectivity, regulating sieve element composition at the molecular level. Such molecular regulation leads to the emerging dogma of the phloem not only as the circulation system of the plant but also as its nervous system. It is expected that over the next few years, the increasing power of molecular techniques will continue to integrate with our existing (and developing) physiological approaches to establish the phloem system as a central point of the regulation of plant growth, development and response to the environment.

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11 Factors limiting the rate of supply of solutes to the root surface Anthony Yeo

11.1

Introduction

Chapters 11 to 15 of the book consider solute relations under different environmental conditions. Before considering solute transport in conditions of mineral deficiency and toxicity (Chapter 12), water-limited conditions (Chapter 13), salinity (Chapter 14) and desiccation (Chapter 15), some of the general factors that limit the rate of solute supply to the root surface are assessed in this chapter. These factors include the bioavailability and mobility of nutrients in the soil, their heterogeneity and the effects of boundary (unstirred) layers and depletion zones. There has been considerable emphasis on the characteristics of transport processes since the interest in their analysis in terms of Michaelis–Menten kinetics in the middle of the last century (e.g. Epstein and Rains, 1965; Torii and Laties, 1966). It is clear from experimental studies that the affinity of plant solute transport systems can be very great. Plants are able to take up nutrients at substantial rates from concentrations that are extremely low in comparison with those supplied by typical nutrient solutions, in soil-less horticulture and those resulting from the ‘one-off’ application of rapid-release fertilisers. Plants have a spectrum of transport processes of differing affinity: those of high affinity, and others providing greater flux density, and/or lower cost uptake, in the presence of abundant nutrient supply. High affinities have sometimes been related to field conditions, particularly for phosphate, where it has been demonstrated that limiting concentrations in the soil solution are consistent with K m values for high-affinity transport processes. With the exception of phosphate, plants are generally able to acquire nutrients from concentrations that are much lower than those found in ‘average’ nutrient-sufficient soils. The implications are that factors other than the amount of nutrient in the bulk soil or bulk solution limit the amount of nutrient taken up by the roots of plants frequently enough for plants to have evolved many high-affinity transport processes. Many factors are concerned in the process of getting nutrients from the bulk soil or (soil) solution into the roots of plants. These include physical, chemical and biochemical processes in the medium as well as the process of interception – both of nutrients moving towards roots and of roots moving towards nutrients (see also Section 8.3). Nutrient uptake by plants results from several layers of processes that broadly comprise the following.

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A: Supply of nutrients to the root surface: r the presence or absence of the nutrient element in the growth medium in any form; r the bioavailability of that element and the processes, both inorganic and organic, concerned with making it available (particularly the mineralisation of nitrogen and phosphorus); r the mobility of that element in the growth medium and its movement towards roots in terms of its diffusion through, absorption on and chemical interaction with the medium; r the movement of nutrients towards roots including – the effects of soil moisture content on mass flow of soil solution; – the effect of boundary or unstirred layers, and of depletion zones (where the flux density of uptake exceeds the flux density of supply); – the homogeneity or heterogeneity (spatial and temporal) in the availability of that element to the plant; – losses, i.e. nutrients or fertilisers that are lost to the system before they can be intercepted by the plant roots. B: Acquisition and uptake of nutrients by the root: r the affinity (K m ) and capacity (the Vmax or flux density) of transport processes in the roots, including the ability of the plant to provide the energy resources to facilitate the accumulation of nutrients against considerable gradients in electrochemical potential; r the exploration and exploitation of soil volume by roots including – constitutive and plastic factors; – competition (interspecific and intraspecific). Separation of the influence of different factors is complex because they all interact. Further, plants do not commonly grow in isolation except in experimental conditions – competition both intraspecific and interspecific as well as competition with microorganisms are ever-present. Finally, measuring what roots do in the soil, in a competitive situation, is very difficult, and this constrains our knowledge about ‘real’ situations.

11.2 11.2.1

Supply of nutrients to the root surface Absence of the nutrient element in the growth medium in any form

The substantial or complete absence of an essential nutrient will dramatically limit or exclude the presence of plants (see Chapter 12).

11.2.2

Bioavailability of the element

Nutrients may exist in forms that are of limited availability to plants. This includes limited solubility, strong absorption onto charged surfaces, or their presence in complex organic form. Sparing solubility or absorption, such as onto clay minerals, may

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provide a low available concentration in relation to the quantity present and include some buffering equivalent to slow release. In solution culture, the use of chelated species (such as ferric EDTA) performs a similar function of providing a buffered low concentration from a relatively large ‘unavailable’ source. The same concept applies to using ‘sequestered’ iron to counter the chlorosis often encountered when growing acid-loving ornamentals, such as rhododendron, on chalky soils (see Chapter 12). Organic sources of nutrients in the soil (particularly of nitrogen and phosphorus) need mineralisation to become available to plants. The rate of mineralisation of nitrogen from organic matter is related to available soil water (Gregory et al., 1997). Mycorrhizal association may allow exploitation of additional types of resources. The nitrogen source in the soil consists of nitrate and ammonium ions (the forms usually taken up by plants) as well as reserves from which nitrogen is mobilised. Organic nitrogen is made available through mineralisation and nitrification, and the rate of mineralisation is dependent upon the concentration of organic matter in the soil. Some species have preferences for nitrate over ammonium or vice versa, and this relationship is interlaced with other factors. For instance, the uptake of ammonium and nitrate appears to be differentially sensitive to elevation of CO 2 , and there is also evidence that soil temperature differentially affects the uptake of the two different sources of nitrogen (Bassirirad, 2000). Consequently, it is possible to observe a response to either nitrate–nitrogen or ammonium–nitrogen fertilisation even in the presence of ‘adequate’ total nitrogen. For phosphorus, it has been possible to relate yield to extractable soil phosphorus, but such relationships are specific to the method of extraction and to different soils, particularly because of the differing capacities for phosphate absorption between sand and clay soils (Smethurst, 2000). For example, the phosphorus sorption capacity of a clay soil was some 40-fold greater than that of a sandy soil, and this was reflected proportionately in the alkaline-extractable phosphorus that was needed to produce equivalent growth in the different soils in experimental conditions (Bolland et al., 1994). Phosphate is adsorbed onto positively charged minerals such as oxides of iron and aluminium and forms precipitates with common metals such as calcium, aluminium and iron. A collection of absorption/desorption and precipitation/dissolution equilibria controls the concentration of phosphorus in the soil solution and through this its chemical mobility and bioavailability (Hinsinger, 2001). Factors having a major role in determining these equilibria are pH, the presence of ions that compete with phosphorus for ligand-binding sites and the concentration of metals that can form insoluble phosphate precipitates. The conditions in the rhizosphere can be very different from those in the bulk soil because of the presence both of plant roots and of microflora. Plant roots can modify conditions in the soil by extrusion of protons and bicarbonate ions, and by gas exchange, both of which affect soil pH. Plants can also modify soil conditions by exudation of ligand-forming organic solutes. The rate of diffusion of inorganic nutrients, however they become available, differs greatly. The nitrate anion is freely soluble in water and has little interaction with soil structure such that its diffusion coefficient is close to that in solution, at about 10−10 m2 s−1 (Hodge, 2004). At the other end of the scale, phosphate ions are

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strongly absorbed on the soil structure and their mobility is three to five orders of magnitude lower. Phosphate uptake is considered in more detail later in this chapter, as well as in the Chapter 12. The availability of phosphate is unusual in that there are generally low effective concentrations even in ‘fertile’ soils, and phosphate is commonly limiting even in ‘fertile’ soils.

11.2.3

Movement of nutrients towards roots

For uptake, nutrients must reach the sites of transport on the plasma membrane of root cells. This is a matter both of nutrients moving towards roots and of roots moving towards nutrients. There is a mass flow of solution towards the roots due to transpiration (see also Section 8.4). This is most effective in moist soils and for nutrients that are highly soluble and present at reasonably high concentrations (such as nitrate). However, mass flow depends upon the hydraulic conductance of the soil (Eq. 3.14), and there may be little mass flow in dry soils. This is because there is a reduced cross-sectional area available to bulk flow, restricted to regions where there is a film of solution around soil particles, with the path increasingly interrupted by air spaces. Even when there is a substantial mass flow of (soil) solution towards the roots in response to transpiration, this may not be enough to support the rate of transport at the root surface. Potential uptake at the root is characterised by its flux density (or V max per unit surface area measured in μmol m−2 s−1 ). The flux density of supply is the concentration in the soil solution multiplied by the mass flow per unit of root surface area. There will be local nutrient depletion wherever the flux density of transport exceeds the flux density of supply. Nutrient uptake is then liable to be limited by the movement (principally diffusion) of nutrient across the depletion zone or the rate of growth of new roots into unexploited soil. It is possible for solutes at very high concentrations to be accumulated, rather than depleted, in the root zone if the rate of supply by mass flow exceeds the rate of transport. This may in particular occur in saline soils where 90% or more of the dissolved salt is excluded at the membrane surface and left behind in the soil (Chapter 14). Whether or not there is a depletion zone, roots will always be surrounded by a boundary or unstirred layer of solution, whether in soil or in solution (see also Section 8.4). The apoplast also constitutes an unstirred layer immediately surrounding the plasma membrane. The unstirred layer around the root is analogous to the boundary layer of air that surrounds leaves, which is thickest in still air and thinnest in high wind. The boundary layer around roots is thinnest in rapidly stirred or moving solutions and thicker in poorly stirred or unstirred solutions and in the soil. The unstirred layer has no defined boundary and in a complex solution has a different thickness for different solutes. Molecules with large diffusion coefficients encounter a thicker boundary layer than do molecules with smaller diffusion coefficients, the thickness being in proportion to the third root of their diffusion coefficients (Pohl et al., 1998). However, molecules with a larger diffusion coefficient will move across

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the boundary layer faster for any given concentration difference. Thus, the effective boundary layer has dimensions made up of both distance and transit velocity. Within the unstirred layer, ion movement may be partitioned between bulk water flow and diffusion; however, the contribution of bulk flow to ion movement in the boundary layer has been calculated to be small or negligible (Henriksen et al., 1992). The limitations imposed by the rate of diffusion can result in steep concentration profiles within the boundary layer, to the extent that analysis of the (soil) solution may give a poor idea of the effective concentration at the uptake site. Using microelectrodes, a logarithmic profile of nitrate concentration was measured across the unstirred layer around barley roots in solution (Henriksen et al., 1992). Roots can also intercept nutrients (see below) and this is usually of most importance with relatively immobile nutrients. As is also discussed in Chapter 12 (and, below, for phosphate), root exudates can increase acquisition both by chelation and by modifying conditions in the soil, and mycorrhizal associations can also facilitate uptake. Roots also explore and exploit new volumes of soil, and again this is increasingly important for relatively immobile nutrients and in drier soils. Mass flow is more effective at supplying nutrients to the root in moist soils and with high rates of transpiration. It is less important in drying soils where more and more of the pores are filled with air.

11.2.4

Homogeneity or heterogeneity (spatial and temporal) in availability

Natural environments are heterogeneous in space and time and over different scales. Heterogeneity of nutrient availability can be on the scale of micrometres when the concentration gradients in the unstirred layers around roots are considered, or of a scale such as within and between fields, up to those that affect plant zonation. The interaction of plants with heterogeneity, or ‘nutrient patches’, on a scale smaller than a single plant root system, is widely investigated (see Section 11.3.2). Temporal variation can be on the scale of the development of a depletion zone where uptake exceeds supply, through changes after sudden rainfall or fertilisation, to changes due to ecosystem development from colonisation to climax. Even for such a mobile ion as nitrate, heterogeneity is considerable – up to fivefold at a 20-cm spatial resolution in deciduous woodland (Farley and Fitter, 1999).

11.2.5

Losses

Owing to the solvent properties of water, loss by runoff and deep drainage carries nutrients out of the root zone. For example, worldwide nitrogen-use efficiency (NUE) in agriculture is approximately 33% (Raun and Johnson, 1999) and some 50 million tonnes were used for cereal production in 1996 (FAO cited in Raun and Johnson, 1999). Since NUE is based on nitrogen in the grain, it is dependent upon harvest index as well as ‘losses’. Losses of fertiliser application result from a combination of surface run-off, leaching, soil denitrification, volatilisation and plant gaseous emissions. There are large elements of economic and environmental cost involved.

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Acquisition and uptake of nutrients by the root Affinity and capacity of transport processes in the roots

The constants of Michaelis–Menton enzyme analysis are commonly used as descriptors of transport processes. Some confusion can arise because all transport processes, even if showing some form of saturation kinetics, are not necessarily pumps or carriers. And even if a process is a pump or carrier (Section 5.1.2) the rate is not necessarily limited by the transport process, but by access to the transport site, almost certainly involving some aspect of the boundary layer. In addition to the unstirred layer around the root, there is also the relatively tortuous and unstirred pathway through the apoplast to the uptake sites on the plasma membrane, particularly for sites other than the outer face of the epidermis. An increase in external concentration (in conjunction with the membrane potential) increases the driving force for passive movement of a cation across the plasma membrane. Although the number of ions that can pass through a channel in unit time is greater by orders of magnitude than the number that can be moved by a pump or carrier, that number is finite. And the channels are also gated. Since there is a ceiling, passive fluxes through channels may also show nonlinearity with respect to external concentration. Where there are depletion zones, the rate-limiting step does not reside with the transport process, rather with the process of diffusion from the bulk (soil) solution to the uptake site. Diffusion is proportional to the concentration gradient, which bears a complex and ever-changing relationship to the external concentration. The ‘gradient’ between the concentration applied in the bulk solution and that known or supposed as the starting concentration inside the membrane is certainly not fixed. Unless the solute is removed from the cytoplasm at exactly the same rate as the rate of uptake, the concentration difference between the bulk solution and cytoplasm will decrease over time, and generally will be expected to decrease more rapidly at higher external concentrations. This will tend to lead to some form of nonlinear kinetics with respect to the external concentration. The same consideration applies to the boundary (unstirred) layer. In many situations, nutrients will have to diffuse in series across both a depletion zone and an unstirred layer to reach the root surface. Because of the boundary layer (and depletion zone, if present), the concentration difference between bulk solution and cytoplasm (known or estimated) is only a pointer towards the true concentration difference across the membrane or concentration at the site of uptake. The dynamics of the establishment and change in the gradient profile, and so the real concentration at, or concentration difference across, the membrane, may be poorly known. In many, perhaps very many, cases of solute uptake by roots, it is diffusion across the boundary layer and/or the depletion zone that may be rate limiting for uptake. All this makes it intensely difficult to relate what happens at the molecular level to what is happening in a root in the soil. A practical measurement of affinity, which has been made in soils, is the critical concentration in the soil solution. This is defined as the concentration of a nutrient that provides 75% or more of the maximal yield. This analysis is applicable where there is an asymptotic relationship between the concentration in the soil solution and

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the relative yield of the crop (Smethurst, 2000). Such a relationship was found for Trifolium subterraneum over a range of sites in Australia, where 50% of maximal yield was achieved at about 2 μM phosphorus in the soil solution, and a concentration of 5 μM phosphorus in the soil solution elicited 95% of maximal yield (Dear et al., 1992; Figure 1 in Smethurst, 2000). However, depending on other physical and chemical considerations, the quantity of phosphorus in the soil needed to generate such steady-state concentrations in the soil solution can be considerable. In many cases, the rate of mineralisation of phosphorus and the rate of exploration by the roots of plants are liable to be limiting for growth. Overall, the existence of an asymptotic relationship implies that supply of phosphorus is commonly a key limiting factor. An asymptotic relationship between yield and the availability of a nutrient is not at all universal, and the equivalent relationship for nitrogen shows a different pattern – the data points falling on or above an imaginary asymptotic curve (Tisdale et al., 1993; Figure 2 in Smethurst, 2000). Since the majority of data points are above the limiting envelope this implies that nitrogen was generally not the limiting factor in yield. Sands and Smethurst (1995) applied Michaelis–Menten kinetics to analyse earlier experiments conducted using the method of ‘nutrient flux density’. Rather than providing a constant solution concentration, this method consists of pulsed additions of nutrients, increasing with time as the plants grow, such that both a stable relative growth rate and a stable internal concentration of a limiting nutrient are maintained. The results (Sands and Smethurst, 1995) estimated that the equivalent concentration in circulating nutrient solution for 50% maximum RGR would be about 10 μM nitrogen and for 95% maximum RGR about 50 μM nitrogen. Direct continuous measurements of the concentration of nutrients in the xylem sap of transpiring plants have been made by sampling based on the xylem-feeding spittle-bug Philieanus spumarius (Malone et al., 2002). Both earlier analyses of xylem sap (Table 6.1 in Flowers and Yeo, 1992) and direct xylem sampling (Malone et al., 2002) report that xylem concentrations of macronutrients were all in the hundreds of micromolar to millimolar range (Table 11.1; see also Table 9.1). Concentrations of potassium and nitrate were closer to 10 mM. These values are in a

Table 11.1 Approximate range in concentrations of various macronutrients in the xylem of mature pepper plants growing in rockwool in a greenhouse Nutrient

Concentration (mM)

Potassium Calcium Ammonium Nitrate Phosphate Sulphate

6–10 1–2.5 2–2.5 10–17 about 1 about 1

Data were obtained by xylem sap sampling at intervals over a period of 96 h using Philaenus spumarius. Anions and cations were determined by ion chromatography. Estimated from Figure 5 (Malone et al., 2002).

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similar range to the concentrations of major nutrients in typical ‘nutrient solutions’ but they are orders of magnitude greater than K m values for high-affinity transporters. In these conditions, the flux density of transport could never be supplied in soil by advection alone at a concentration remotely near these K m values, since depletion zones would arise almost instantaneously. Even in solution culture, there would need to be enough turbulence to eliminate the depletion zone and the unstirred layer. Presumably, high-affinity transporters developed for conditions of nutrient deficiency, and they have a place alongside lower affinity systems in the spectrum of transporters present in the roots of plants. A spectrum of transport processes, including passive uptake via channels, are usually operational. Except possibly for phosphate, it is very unlikely that any plant making near-maximal growth is relying upon high-affinity transporters for uptake of major nutrients. A crucial advantage of high-affinity transporters is their ability to deplete the concentration of a nutrient at the root surface and thus generate a substantial concentration gradient to drive diffusion across the unstirred layer (and depletion zone). In the cases of phosphorus (and other nutrients with low solubility products for the ionic species required by the plant) the ability of high-affinity transporters is necessary to scavenge the nutrient ion to a low-enough concentration to maintain a favourable equilibrium.

11.3.2

Exploration and exploitation of soil volume by roots

As a plant grows, the depletion of nutrients from the surrounding soil intensifies. The effectiveness of diffusion as a means of supply becomes less as the distance over which diffusion has to occur increases (see Fick’s Law in Chapter 3). Continual growth of the root system is therefore necessary to explore and exploit new volumes of soil. Locally, exploitation is increased by the development of root hairs, though exploration over longer distances is also needed. Since the environment is commonly heterogeneous, the way that plant roots are distributed in the soil commonly reflects and adapts to this. Root hairs (see also Section 8.5.2) develop as protuberances from the epidermis. They increase both the radius of the root and its surface area. The increase in radius (increased volume of explored soil) is probably more important in terms of nutrient acquisition than is the increase in surface area. The development of depletion zones around roots implies that the transport capacity of the root is not usually limiting; it is more the question of supply. Root hairs extend the root surface out into the boundary layer. Mycorrhiza also extend the radius of the root, though by considerably greater distance than do root hairs. A dramatic demonstration of root proliferation in regions of high nutrient concentrations was made using barley in a solution-based system where a central section of the root system was exposed to a higher concentration than was the rest of the root system (Drew, 1975; see Figure 8.1). Root proliferation in the high-nutrient zone has been demonstrated for phosphate, nitrate and for ammonium, but was not seen for potassium. The proteoid roots of the Proteaceae (see Section 12.3.1) can increase the volume of soil explored by a factor of 300 in comparison with the same length of equivalent non-proteoid root (Lamont, 2003).

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In general, roots often proliferate when they encounter a nutrient-rich patch and can also show enhanced uptake capacities in comparison with roots outside these patches. This plasticity had been regarded as a major mechanism by which plants cope with soil heterogeneity; however, it is far from easy to demonstrate that root proliferation benefits the plant with regard to capturing nutrients that have good mobility (Hodge, 2004). Theoretical and experimental support suggesting the beneficial effect of root proliferation in increasing the capture of immobile phosphate exists, but experiments have often failed to show that root proliferation increases the capture of nitrogen from nitrogen-rich patches. It appears that root proliferation is important in nitrogen capture only in certain circumstances. These are where there is interspecific competition for the resource and where the nitrogen patch is of a ‘slow-release’ form (an organic source that needs to be decomposed over time into inorganic ions). In simpler situations, for individual plants and monocultures, and for inorganic sources of nitrogen, positive value of root proliferation may not be demonstrated (Hodge, 2004). That plants do express the proliferation response constitutively in simple experimental conditions underlines the difficulties in drawing the right conclusions. It cannot be assumed that because a response is observed experimentally, it will necessarily be of any benefit to a plant in the field. Root proliferation involves the investment of resources in generating new roots and requires shifts in resource allocation within the root system, or an increase in resource allocation to the root system. There has been only limited support for a general assumption that faster growing species, with more resources, would be capable of more plasticity than slower growing species with less resources to allocate (Hodge, 2004). However, the analysis is complicated by a need to consider relative as well as absolute investment, and the fact that benefits are sometimes clear only if there is competition for the same resource. A distinction was drawn between ‘scale’ effects (where one plant/species has a greater quantity of roots in a nutrientrich patch than does another) and ‘precision’ (where one plant/species produces a greater proportion of its new root growth in a nutrient-rich patch than does another; Campbell et al., 1991). Precision is itself variable within the same species according to the pattern of nutrient distribution (Weijesinghe et al., 2001) and also over time (Hutchings and John, 2004). In the latter case, root positioning into one nutrientrich and three nutrient-poor quadrants was monitored over time and distribution regressed upon total root mass for three grass species. Initially, there was large variation in precision (as measured by total root mass) with some high percentages in the nutrient-rich quadrant, but as plants grew there was convergence towards equal distribution into nutrient-rich and nutrient-poor quadrants (Hutchings and John, 2004). A considerable range of responses are reported in the literature in which nutrient addition sometimes, but not always, increases resource allocation to roots. Root: shoot ratio was increased in about half of the cases reviewed (Robinson, 1994), being unaffected in 45% and reduced in 5% of cases. Differences in response between the Graminae and other plants are also reported (Robinson and van Vuuren, 1998). It is apparent from the simple comparison of the availability of phosphorus and nitrogen that limiting responses vary according to the characteristics of individual

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nutrient availability and mobility in the soil. For nutrients of low mobility (such as phosphorus) there will be enormous heterogeneity over very small scales (i.e. within the rhizosphere of an individual root) and plant response to nutrient status is different from that to the mobile nitrate anion. Rates of mobilisation from nutrient reserves also differ and it is important whether, for example, the nitrogen source is organic or an inorganic fertiliser. Species also differ in their own effects upon mobilisation of nutrients, particularly those such as phosphorus and iron, for which major differences in method apply between graminaceous and other species. Any response observed in the plant may be to the limiting nutrient, and which nutrient is limiting may not be readily obvious. It is, however, clear that although the acquisition of nutrient depends in one sense upon the characteristics of the transport systems, it is overlaid by many other levels of complexity. Distribution of nutrients in the soil is non-uniform in space and time at scales that operate at and below that of the plant and, certainly for immobile nutrients, at levels in the millimetre and micrometre ranges. Plants do not usually grow in isolation, and competition exists both between and within species – and may be most intense in monocultures. Some characteristics of nutrient acquisition are detectable only where there is competition. Heterogeneity varies in intensity, scale and duration, and hence the balance between exploitation and continued foraging may be anticipated to have different consequences in different situations. There is much to learn as to how plants detect and coordinate their responses to heterogeneous environments, but the underlying message is that there is more to nutrient acquisition than the K m and V max of the relevant transporters. The caveat is that these other factors become less and less important, the more the environment of the plant can be controlled, which is the general aim of agricultural and horticultural husbandry. Minimising these ‘extraneous’ effects is the goal of soil-less cultivation systems.

11.4

Acquisition of phosphorus

Mineral deficiency is the main topic of Chapter 12, but some consideration is also given here to phosphorus because plant growth is limited by supplies of inorganic phosphate in most natural ecosystems. This has led to the evolution of adaptive strategies to increase the acquisition of phosphate, including proliferation of cluster roots and the release of organic acids and phosphatases (Smith et al., 2003). In the same way as for organic nitrogen, it is necessary for organic phosphorus to be mineralised (to inorganic phosphate) before it can be taken up by plant transport processes. Complex equilibrium reactions and sorption isotherms result in the concentration of phosphate in solution ranging from below 1 μM in weathered, sandy and alkaline soils (Reisenauer, 1966) to rarely exceeding 8 μM (Smith et al., 2003). Consequently, phosphorus has a unique place amongst macronutrients in that even in fertile soils its effective concentration is usually below 10 μM. The concentration of phosphate in the cytoplasm is maintained within a narrow range between about 5 and 17 mM (Mimura et al., 1996; Mimura, 1999). This represents an accumulation ratio of three or four orders of magnitude with respect to

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the external concentration of phosphate. A further order of magnitude accumulation into the vacuole (with concentrations up to 120 mM; Mimura et al., 1990) provides a substantial store that can be drawn upon in times of external deficiency. High-affinity phosphate transporters in plants belong to the Pht1 family. They function in secondary active transport as H+ /H 2 PO 4 − co-transporters and are predicted to have 12 membrane-spanning domains giving them a ‘doughnut-shaped’ structure with an inner pore through which the ions pass (Smith, 2002). There was early difficulty in establishing the affinity of PHT1 in heterologous systems, but the Arabidopsis thaliana AtPHT1, expressed in tobacco cells, indicated a K m of 3.1 μM for phosphate (Mitsukawa et al., 1977) consistent with it being the high-affinity phosphate transporter. The Pht1 transporters, concerned with high-affinity initial uptake, differ from the Pht2 family. The Pht2 family is concerned with inter- and intracellular movement of phosphate; these are processes which all involve much higher concentrations of phosphate, and thus high affinity is not needed. The rate of phosphate uptake from the external medium is determined by the number of transporter molecules present in the membrane and by their activity. In tomato, it has been demonstrated that in phosphate-deprived plants, enhanced levels of mRNA were translated into increased levels of high-affinity transporter protein and thus an increased number of phosphate transporter protein molecules (Muchhal and Ragothama, 1999; Smith, 2002). Phosphate transporters were also found to exhibit rapid turnover, enabling rapid modulation of phosphate transport by direct transcriptional control. The Pht1 family generally contains highly conserved sites capable of phosphorylation or glycosylation, enabling post-translational modification, with allosteric regulation of activity also possible (Smith, 2002). The expression of genes for phosphate transporters is regulated by the overall phosphorus status of the plant. This is a systemic (whole plant) and not a localised (around the root) response, and under phosphate stress, genes encoding for the high-affinity phosphate transporter are up-regulated. However, deficiency of another nutrient overrides this response, thus restricting the uptake of phosphate that cannot be utilised effectively (Smith et al., 1999). This may reflect that the accumulation of phosphate by proton co-transport is an expensive process, and so there is an advantage of preventing unnecessary uptake. This has implications for the methods to be adopted in breeding plants that are less susceptible to phosphate deficiency. Attempting to increase phosphate uptake by overexpressing genes encoding high-affinity phosphate transporters is applicable to situations where an adequate concentration of phosphate can be maintained at the plasma membrane surface (Smith, 2002). As discussed above, the low solubility and low mobility of phosphate in the soil, combined with its macronutrient requirement by plants, means that depletion zones around the root are the norm. This means that there will be many situations in which the capacity of phosphate transport is not limiting the uptake, but the supply of phosphate to the root surface is. In these circumstances, constitutive overexpression of Pht1 transporters may not only be ineffective, but constitute a costly disadvantage, which the plant’s own regulatory systems would, if left to their own devices, have been able to avoid. In soils with low available phosphate, emphasis is on strategies to improve phosphate movement to the root and possibly on manipulating internal mobilisation of phosphate (Smith, 2002).

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Protected cropping systems: hydroponics as an example of ‘ideally’ controlled conditions

The use of hydroponics in protected cropping has become increasingly widespread, particularly linked to computerised automation, not only of the irrigation system but of the whole greenhouse environment. Hydroponics presents both advantages and disadvantages (Marr, 1994; Savvas, 2003). Hydroponics means growth without soil. Except for inept laboratory experiments, hydroponics does not imply a plant with its roots sitting in a tub of unstirred and unaerated solution. A nutrient solution (fertigation – combining both irrigation and fertilisation) is supplied, usually by drip-feed (now often computer controlled and ‘needs-led’ by sensors monitoring the crop), to plants whose root systems grow in an inert medium. Various inert supports are used: rockwool, perlite, pumice, expanded clay, volcanic minerals, polyurethane foam and so on (Savvas, 2003). A balance of pore sizes is needed to supply both water and oxygen. The only commercial exception is nutrient film (NFT) where plant roots are allowed to develop in enclosed channels set on a gentle incline through which solution flows continually under gravity. In hydroponic culture, the concentrations of major nutrients (N, P, K and Ca) are usually in the (several) millimolar range (Table 1 in Marr, 1994). This again indicates that in situations demanding maximal rates of growth, high-capacity/low-affinity systems predominate. Hydroponics avoids the decline in soil structure and fertility that results from continual cultivation with the same or a similar crop. In greenhouse cultivation (where, for example, tomato may be grown all year, every year) there may be limited potential either for crop rotation or for allowing the land to lie fallow. Drainage problems and soil heterogeneity may also be avoided. Hydroponics also avoids soilborne pathogens and removes the need to sterilise large quantities of soil, which can reduce the quantities of toxic chemicals used in the growing season. Hydroponics using closed, recirculating systems, recycling excess nutrients, also reduces or eliminates seepage down to the water table and environmental contamination of surface- and groundwaters. Some disadvantages of hydroponics are that solution techniques and inert growing support offer much less buffering capacity than there would be in the soil. There is no controlled release of water or nutrients if the system stops, nor is there so much buffering against changes in pH or build-up of toxins. Hydroponics places a ‘high-risk’ reliance on human and technical intervention. Although it is relatively easy to start the season with the system free of pathogens, hydroponics is easier to infect than soil, because pathogenic organisms, should they arrive, will not be in competition with the host of non-pathogenic organisms that would be present in a soil. Probably the greatest limitation in the use of hydroponics is the economic disadvantage in ‘low-tech’ situations. With the investment in large scale (often hectares) of controlled greenhouses for the climate of northern Europe, the additional cost of hydroponics is relatively minor. The situation is not the same in (for example) the Mediterranean region where greenhouses are of smaller scale, intended only to

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modify what are basically favourable natural conditions, and where the proportional cost of adding hydroponics may be excessive (Savvas, 2003). Control is becoming increasingly sophisticated and integrated, as befits a situation in which a single tomato crop may occupy the greenhouse for 11 months of the year and crop for the majority of that time. Control of climate in protected systems has historically used monitoring of climate and the physical side – such as the nutrient solution. There is more recent interest in monitoring not the environment but the state of the crop itself (Ehret et al., 2001).

11.6

Concluding remarks

Many factors, including the bioavailability, mobility and heterogeneity in distribution of nutrients, boundary layers and depletion zones, and soil moisture content, all limit the rate of supply of nutrient to the uptake sites at the surface of the root. Consequently, the affinity and flux density of transport processes are not necessarily the rate-limiting steps in nutrient supply to the growing plant. Maximal rates of growth, aimed for in agriculture, require flux densities of uptake that are usually supported by concentrations of nutrients that are orders of magnitude greater than the K m of high-affinity transporters. Plants have a spectrum of transporters of differing affinity that can support greater flux density and/or more cost-effective modes of uptake in conditions of high nutrient supply. The situation may be very different for plants growing in generally nutrient-poor environments, in which high-affinity transport may predominate, but plants growing in these conditions are unlikely to achieve substantial rates of growth. The ‘exception’ for productive ecosystems is phosphate, a macronutrient that is generally available only in concentrations consistent with high-affinity transport. Additional anatomical and biochemical adaptations are available to confront this limitation. The tight regulation of high-affinity phosphate transport activity by plants, ensuring that it is rapidly up-regulated only when needed and rapidly turned down when phosphate is no longer the limiting factor, has implications. Although it is always rash to generalise, this does suggest that high-affinity transport, which may be at the cost of one molecule of ATP per ion transported, is such an expensive process that it cannot be used indiscriminately. This would have generic bearing on the choice of strategy considered to deal with nutrient deficiency (or toxicity– see Chapter 14 on salinity) by any constitutive modification of solute transport properties.

References Bassirirad, H. (2000) Kinetics of nutrient uptake by roots: responses to global change. New Phytologist 147, 155–169. Bolland, M.D.A., Wilson, I.R. and Allen, D.G. (1994) Effect of P buffer capacity and P retention index of soils on soil test P, soil test P calibrations and yield response curvature. Australian Journal of Soil Research 32, 503–517.

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Campbell, B.D., Grime, J.P. and Mackey, J.M.L. (1991) A trade-off between scale and precision in resource foraging. Oecologia 87, 532–538. Dear, B.S., Helyar, K.R., Muller, W.J. and Loveland, B. (1992) The P fertiliser requirements of subterranean clover, and the soil P status, sorption and buffering capacities from two P analyses. Australian Journal of Soil Research 30, 24–44. Drew, M.C. (1975) Comparison of the effects of a localised supply of phosphate, nitrate, ammonium and potassium on the growth of the seminal root system, and the shoot, in barley. New Phytologist 75, 479–490. Ehret, D.L., Lau, A., Bittman, S., Lin, W. and Shelford, T. (2001) Automated monitoring of greenhouse crops. Agronomie 21, 403–414. Epstein, E. and Rains, D.W. (1965) Carrier-mediated cation transport in barley roots: kinetic evidence for a spectrum of active sites. Proceedings of the National Academy of Sciences of the United States of America 53(6), 1320–1324. Farley, R.A. and Fitter, A.H. (1999) Temporal and spatial variation in soil resources in a deciduous woodland. Journal of Ecology 87, 688–696. Flowers, T.J. and Yeo, A.R. (1992) Solute Transport in Plants. Blackie Academic and Professional, London. Gregory, P.J., Simmonds, L.P. and Warren, G.P. (1997) Interactions between plant nutrients, water and carbon dioxide as factors limiting crop yields. Philosophical Transactions of the Royal Society of London, Series B 352, 987–996. Henriksen, G.H., Raman, D.R., Walker, L.P. and Spanswick, R.M. (1992) Measurement of net fluxes of ammonium and nitrate at the surface of barley roots using ion-selective microelectrodes. II: Patterns of uptake along the root axis and evaluation of the microelectrode flux estimation technique. Plant Physiology 99, 734–747. Hinsinger, P. (2001) Bioavailability of soil inorganic P in the rhizosphere as affected by root-induced chemical changes: a review. Plant and Soil 237, 173–195. Hodge, A. (2004) The plastic plant: root responses to heterogeneous supplies of nutrients. New Phytologist 162, 9–24. Hutchings, M.J. and John, E.A. (2004) The effects of environmental heterogeneity on root growth and root/shoot ratio partitioning. Annals of Botany 94, 1–8. Lamont, B.B. (2003) Structure, ecology and physiology of root clusters – a review. Plant and Soil 248, 1–19. Malone, M., Herron, M. and Morales, M.A. (2002) Continuous measurements of macronutrient ions in the transpiration stream of intact plants using the meadow spittlebug coupled with ion chromatography. Plant Physiology 130, 1436–1442. Marr, C.W. (1994) Greenhouse Vegetable Production: Hydroponic Systems. Kansas State University Agricultural Experiment Station and Co-operative Extension Service, Manhattan, Kansas. Mimura, T. (1999) Regulation of phosphate transport and homeostasis in plant cells. International Review of Cytology 191, 149–200. Mimura, T., Dietz, K.J., Kaiser, W., Schramm, M.J., Kaiser, G. and Heber, U. (1990) Phosphate transport across biomembranes and cytosolic phosphate homeostasis in barley leaves. Planta 180, 139–146. Mimura, T., Sakano, K. and Shimmen, T. (1996) Studies on distribution, re-translocation and homeostasis of inorganic phosphate in barley leaves. Plant, Cell and Environment 19, 311–320. Mitsukawa, N., Okumura, S., Shirano, Y., et al. (1977) Overexpression of an Arabidopsis thaliana high-affinity phosphate transporter gene in tobacco cultured cells enhances cell growth under phosphate-limited conditions. Proceedings of the National Academy of Sciences 94, 7098–7102. Muchhal, U.S. and Ragothama, K.G. (1999) Transcriptional regulation of plant phosphate transporters. Proceedings of the National Academy of Sciences 96, 5868–5872. Pohl, P., Saparov, S.M. and Antonenko, Y.N. (1998) The size of the unstirred layer as a function of the solute diffusion coefficient. Biophysical Journal 75, 1403–1409. Raun, W.R. and Johnson, G.V. (1999) Improving nitrogen use efficiency for cereal production. Agronomy Journal 91, 357–363.

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Reisenauer, H.M. (1966) Mineral nutrients in soil solution. In: Environmental Biology (eds Altman, P.L. and Dittmer, D.S.), pp. 507–508. Federation of American Societies of Experimental Biology, Bethesda, MA. Robinson, D. (1994) The responses of plants to non-uniform supplies of nutrients. New Phytologist 127, 635–674. Robinson, D. and van Vuuren, M.M.I. (1998) Responses of wild plants to nutrient patches in relation to growth rate and life-form. In: Inherent Variation in Plant Growth: Physiological Mechanisms and Ecological Consequences (eds Lambers, H., Poorter, H. and van Vuuren, M.M.I.), pp. 237–257. Backhuys Publishers, Leiden. Sands, P.J. and Smethurst, P.J. (1995) Modelling nitrogen uptake in Ingestad Units using Michaelis– Menten kinetics. Australian Journal of Plant Physiology 22, 823–831. Savvas, D. (2003) Hydroponics: a modern technology supporting the application of integrated crop management in greenhouse. Food, Agriculture and Environment 1, 80–86. Smethurst, P.J. (2000) Soil solution and other soil analyses as indicators of nutrient supply: a review. Forest Ecology and Management 138, 397–411. Smith, F.W. (2002) The phosphate uptake mechanism. Plant and Soil 245, 105–114. Smith, F.W., Cybinski, D. and Rae, A.L. (1999) Regulation of expression of genes encoding phosphate transporters in barley roots. In: Plant Nutrition – Molecular Biology and Genetics (eds GisselNielson, G. and Jensen, A.). Kluwer Academic, Dordrecht, The Netherlands. Smith, F.W., Mudge, S.R., Rae, A.L. and Glassop, D. (2003) Phosphate transport in plants. Plant and Soil 248, 71–83. Tisdale, S.L., Nelson, W.L., Beaton, J.D. and Havlin, J.L. (1993) Soil Fertility and Fertilisers, 5th edn. MacMillan, New York. Torii, K. and Laties, G.G. (1966) Dual mechanisms of ion uptake in relation to vacuolation in corn roots. Plant Physiology 41, 863–870. Wallace, J.S. and Batchelor, C.H. (1997) Managing water resources for crop production. Philosophical Transactions of the Royal Society of London, Series B 353, 937–947. Weijesinghe, D.K., John, E.A., Beurskens, S. and Hutchings, M.J. (2001) Root system size and precision in nutrient foraging: responses to spatial pattern of nutrient supply in six herbaceous species. Journal of Ecology 89, 972–983.

12 Mineral deficiency and toxicity Anthony Yeo

12.1

Introduction

The original definition of an essential element (Arnon and Stout, 1939) comprised the criteria that: r its deficiency prevented the plant from completing its life cycle; r the deficiency is element specific and can be corrected or prevented only by supplying that element; r the element has a nutritional role – as an essential metabolite, involved in enzyme activation, etc. On this basis, 16 elements are recognised as essential in all plants, with a further 4 established in some, but not all, species (Taiz and Zeiger, 1991; Mengel and Kirkby, 2001). In approximately descending order of typical abundance in plant dry matter (Taiz and Zeiger, 1991), the 16 essential elements are hydrogen, carbon, oxygen, nitrogen, potassium, calcium, magnesium, phosphorus, sulphur, chlorine, boron, iron, manganese, zinc, copper and molybdenum. There is a loose separation into ‘macronutrients’ (ranging from tens, to tens of thousands, of micromole per gram of dry mass and ‘micronutrients’ (ranging from a few to a fraction of a micromole per gram of dry mass), with the division falling between sulphur and chlorine. The four other elements are cobalt, silicon, nickel and sodium (Mengel and Kirkby, 2001). Chapters 8 and 11 considered the concentration dependence of nutrient uptake by plants. The overall picture is that the transport systems that plants possess are well adapted to accumulating the nutrients from very low external concentrations. Provided that there is continual replacement, concentrations that are ‘low’ in relation to laboratory culture solutions can support some plant growth – though higher concentrations, exploited by a spectrum of transporters of differing affinity, are needed for optimal growth. However, there is not always continual replacement, and available concentrations can fall below those at which they are limiting to even the high-affinity transporters. There are also conditions in which concentrations of both essential and non-essential minerals rise above those at which they become phytotoxic. The first part of this chapter deals with the efficiency of uptake and utilisation where these minerals are present, but largely unavailable, or of limited mobility in the soil. If the minerals are simply not present there is nothing the plant can do about it. There are other circumstances in which the minerals may be ‘present-but-unavailable’ and this situation is widespread in alkaline soils for

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minerals that, however abundant, are soluble only under acid conditions. In addition to possessing uptake systems of adequate specificity and affinity, plants also possess mechanisms to help mobilise nutrients that are not in a form accessible to those uptake systems. These mechanisms include the acidification of the rhizosphere by the secretion of protons, the reduction of ferric iron and ferric iron chelates at the root surface and the secretion of specific chelating agents known as phytosiderophores. Chelating agents are also used in the detoxification of metals and in the movement and storage of metals about the plant. Of major nutrients, it is phosphate that is most commonly limiting because of its low solubility and low mobility in the soil (see Chapter 11). The second part of this chapter deals with the consequences of concentrations of minerals in the environment that are toxic to many or most plant species. Some of these situations arise from rare occurrences: soils that contain unusually large amounts of transition and/or heavy metals. Natural rare occurrences are metalliferous soils formed on top of unusually metal-rich geological features. Anthropogenic rare occurrences are soils that are contaminated with the byproducts of industrial activity; these are locally problematic but affect relatively small areas (McNeilly, 1994). There are also the widespread occurrences. Naturally, these are the consequences of soil decomposition under acid conditions leading to phytotoxic concentrations of aluminium, manganese and iron, affecting up to 40% of arable farmed land area. The same result comes from anthropogenic sources via the nitrification of ammonium ions applied in the course of agricultural fertilisation (McNeilly, 1994). A complex of availability and toxicity underlies the ‘calcicole/calcifuge problem’ (the reasons that many plants that grow in acid soils cannot grow well in alkaline soils, and vice versa). Plants have developed mechanisms that allow them to acquire nutrients when they are present but not available in forms that can be taken up by the cation transporters. Plants have also developed mechanisms that allow them to cope with otherwise phytotoxic concentrations of metals by both external detoxification and internal tolerance. At the extreme, plants are able to tolerate some metals in amounts that allow them to be used for defence against herbivory. This chapter focuses principally upon four examples that illustrate these problems and their solutions. These are: 1. the two ‘strategies’ for the acquisition of iron from alkaline/neutral soils in which iron is abundant but generally unavailable to plants (Section 12.2); 2. ‘strategies’ for mobilising soil phosphate (Section 12.3); 3. the phytotoxicity of aluminium on acidic soils and methods of aluminium detoxification and tolerance (Section 12.4); 4. detoxification of and tolerance to transition and heavy metals, and in particular, hyperaccumulation (Section 12.5).

12.1.1

Terminology

It will be useful to begin by defining some terms.

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Mineral deficiency – specific processes or growth in general is limited because of supply of a particular nutrient. This may be because it is not present, is present but not in available form or, if available, its uptake is compromised, or it is rendered immobile (possibly after uptake) by other elements or ions. Mineral efficiency – any attribute that enhances the acquisition of nutrients that are in short supply. Mineral use efficiency – the growth per unit of mineral absorbed – by processes such as compartmentalisation and mobilisation within the plant; some species are able to make more effective use of limited supplies than are other species. Mineral toxicity – the concentration of a mineral in the environment leads directly to toxic concentrations within the plant and/or directly causes deficiencies through competition with, or complex with, other essential nutrients. Mineral tolerance – (a) the ability to detoxify or immobilise potentially phytotoxic minerals in the rhizosphere and so prevent uptake; or (b) by using compartmentalisation and/or inactivation, the ability to grow with internal concentrations of minerals that are commonly toxic to other species. Metallophyte – Plants colonising, or endemic to, unusually metal-rich soils. Hyperaccumulator – Plants that accumulate, without obvious detriment, quantities of (usually) heavy metals orders of magnitude in excess of any metabolic needs. Deficiencies arise both from inadequate concentration in the soil solution per se, and from inadequacies in the rate of replacement. Deficiencies can arise in soils for one or more of the following reasons: 1. There is not enough of the nutrient present in the soil in any form. This will prevent or drastically limit the presence of a plant species in that natural environment. In many, but not all, cases, this may be alleviated by application of fertilisers to the soil. 2. The nutrient is in a form in which it cannot be taken up. The nutrient may be in the form of an insoluble precipitate (whether in the natural state of its source in the soil or as a result of co-precipitation with another mineral). Phosphate is sorbed onto soil minerals and colloids and complex equilibrium reactions both define and limit the concentration of inorganic phosphate that is available to plants in the soil solution. Where precipitates are sparingly soluble at soil pH, the replenishment of the soluble phase may be the rateliming factor. In soils, much or most of some metals will be complexed with naturally occurring organic ligands. In some soils the nutrient may be in the ‘wrong’ oxidation state to comply with the specificity of the relevant transporter (e.g. iron oxidised to FeIII rather than reduced to FeII ). 3. The nutrient has low mobility and/or is patchily distributed (see Chapter 11). This is not in itself a problem of solute transport but a question of foraging by roots to continuously explore and then efficiently exploit new volumes

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of soil. This requires investment in the root system and plasticity of the root system to perceive and respond to nutrient patches. 4. Although present and available, uptake of the nutrient is inhibited by competition from chemically similar ions. In this chapter the focus is predominantly on point 2 – the methods that have evolved in plants in order to access ‘unavailable’ nutrients. Point 4, which is concerned with direct competitive inhibition, is discussed mostly with reference to salinity, in Chapter 14.

12.2

Deficiency and efficiency: iron in alkaline soils

Iron is essential for plants, particularly in redox reactions including nitrogen fixation, and as a constituent of electron transport chains. It is also toxic at higher concentrations. In the Fenton reaction, iron acts as a catalyst and produces reactive oxygen species that may damage macromolecules (Connolly and Guerinot, 2002). Iron deficiency is most marked by chlorosis in photosynthetic organisms because of the large use of iron in the photosynthetic apparatus (12 atoms of iron per PS1 [photosystem 1] complex). Studies in eukaryotic systems have shown that the onset of chlorosis is preceded by the disconnection of the LHCI (light-harvesting centre) antennae from PS1 (Moseley et al., 2002), and that cyanobacteria decrease the PSI–PSII ratio when iron is limiting. Adaptation to iron deficiency in cyanobacteria required a replacement of some proteins with new ones, coupled with a decrease in the stoichiometry of electron transfer processes, characteristic of a balance between optimising photosynthesis and minimising photo-oxidative damage (Moseley et al., 2002). This reflects that plants need to cope with both the ‘essential’ and the ‘toxicity’ faces of iron, even when the quantity of iron in the plant is in the ‘normal’ range. Systems with a large demand for iron are sacrificed when iron is limiting, but iron in the plant must be utilised or scavenged when its availability increases, in order to avoid damage by processes such as photo-oxidation. The optimal range for the concentration of iron in the plant is fairly narrow. For rice this is 100–150 μg g−1 dry mass: deficiency is experienced if the concentration is below 50–70 μg g−1 dry mass and toxicity is experienced if the concentration is greater than 300–500 μg g−1 dry mass (International Rice Research Institute [IRRI], 2003). Iron is one of the most abundant elements in the earth’s crust (Guerinot, 2001) so any limitation to availability is not quantity, but quality. Iron is soluble in acid soils but is almost insoluble in aerobic alkaline soils. Thus, of the nutrients that most commonly limit plant growth, iron deficiency cannot usually be alleviated simply by soil fertilisation (foliar feeding will help, as would soil fertilisation with chelated iron if that were economic). Plant transport systems require iron to be in the ferrous FeII form, and not the ferric FeIII form. Plants that occur on alkaline soils adopt one of two methods to increase the availability of iron: which method has evolved is characterised by phylogenetic distribution. The two pathways, Strategy I and Strategy II, are summarised in Figure 12.1.

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Figure 12.1 The two strategies for iron acquisition from sources of ferric (FeIII ) iron. Strategy I (dicotyledonous) plants utilise FRO1/FRO2, the ferric chelate reductase, to reduce ferric chelates to ferrous (FeII ) iron, which is taken up by IRT1, the ferrous iron transporter. Strategy II plants (of the Poacea) secrete phytosiderophores, which complex FeIII and the resulting complex is taken up by the FeIII –phytosiderophore transporter to be reduced to FeII within the plant.

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12.2.1

295

‘Strategy I’: reduction-dependent iron uptake

One method is found in dicotyledonous plants and another in the grasses (Poacea). The system in the dicotyledons is termed ‘Strategy I’. This iron-uptake system involves three steps and is inducible by iron deficiency. Briefly, protons are released into the soil via the action of the plasma membrane H+ -ATPase to produce a lowering of the pH in the rhizosphere. A reductase (Robinson et al., 1999) at the root surface reduces ferric iron chelate to ferrous iron, which is taken up by a transporter (IRT1) (Eide et al., 1996; Vert et al., 2002). The root plasma membrane H+ -ATPase pumps protons outside the cell, and this both contributes to enhanced solubility of iron oxides and generates the driving force for transporter-mediated uptake. Extrusion of protons lowers the external pH and increases the ability of plants to acquire iron from the sparingly soluble sources in the soil (Marshner and R¨omheld, 1994). Most of the ferric iron that is solubilised will be complexed as an organic chelate in the soil solution. Proton efflux also maintains an acidic pH in the apoplast, which is required for activity of the plasma membrane iron chelate reductase. An increase in the steady-state level of root plasma membrane H+ -ATPase was, at least in part, responsible for the increased proton extrusion in iron-deficient cucumber, suggesting transcriptional regulation of the proton pump (Dell’Orto et al., 2000). The acidification process can increase the availability of iron, but this is as ferric iron in aerobic soils. Plants are able to transport free iron only as the ferrous ion. The reduction of ferric chelates to ferrous iron at the root surface is a necessary requirement for iron acquisition by Strategy I plants. The reduction to ferrous iron is achieved by an FeIII -chelate reductase (FRO1 or FRO2). The FRO2 gene was found to be expressed in roots of iron-deficient Arabidopsis thaliana, and encodes a flavocytochrome with intramembranous iron-binding sites and cytoplasmic binding sites facilitating the donation and transfer of electrons (Robinson et al., 1999). FRO2 was demonstrated to be allelic to the frd1 series of nonsense and missense mutations that impair ferric chelate reductase activity, and to be able to complement the mutant phenotype (Robinson et al., 1999). The FRO1 gene from pea also encodes an FeIII -chelate reductase and FRO1 mRNA levels correlated with FeIII chelate reductase activity. FRO1 mRNA was most abundant in the outer cortical cells of the root and was regulated differentially in roots and shoots (Waters et al., 2002). In tomato (Lycopersicon esculentum), LeFRO1 encodes a protein that, when isolated and expressed in yeast, increases FeIII -chelate reductase activity. RT-RCR data from tomato showed that LeFRO1 was expressed widely throughout the plant, and constitutively in leaves, suggesting that it was required for iron homeostasis (Li et al., 2004). The LeFRO1 protein was targeted to the plasma membrane, and in roots the transcription intensity was dependent upon iron status. The respective iron-dependent and constitutive modes of expression indicated that different control mechanisms applied in roots and shoots. The expression of LeFRO1 was disrupted in iron-inefficient mutants (Li et al., 2004). The high-affinity divalent cation transporter, IRT1, the first member of the ZIP family to be identified, is induced in arabidopsis by iron deficiency, being expressed

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specifically in the outer cell layers of the root in response to iron starvation (Vert et al., 2002). IRT2, a close homologue, has a similar function. A mutation containing an IRT1 knock-out (irt1-1) is lethally chlorotic when homozygous and when iron is supplied only in inorganic form in the growth medium. The mutant can be rescued if fed with sequestered (chelated) iron at very high concentrations in the culture medium, indicating that some low-affinity transport occurred by other pathway(s) in the absence of IRT1. The homozygous irt1-1 mutant also lost all or most of the ability to accumulate other divalent cations (zinc, manganese, cobalt and cadmium) under conditions of iron starvation (Vert et al., 2002). This was taken to suggest that while other (ZIP and NRAMP) proteins may be involved in the uptake of these metals, the accumulation of these metals in response to iron deficiency is mediated by IRT1. Split-root experiments (where different parts of the root system are presented with different conditions) indicated that the expression of FRO2 and IRT1 is controlled both locally, by the root iron pool, and remotely, by signals originating in the shoot, and that iron modulates the responses to its deficiency at both mRNA and protein levels. Furthermore, iron acquisition in Strategy I plants may be under diurnal regulation (Vert et al., 2003). Split-root experiments with iron overaccumulating mutants also supported the concept that FeIII -chelate reductase activity is regulated by a signal molecule communicating the iron status of the shoot, although anatomical developments (transfer cells and increased numbers of root hairs) were regulated by the iron concentration of the growth medium (Schikora and Schmidt, 2001). Transition metal transporters have been reviewed recently (Hall and Williams, 2003). FRO2 and IRT1 were induced together in arabidopsis following iron starvation; were coordinately regulated at both transcriptional and post-transcriptional levels; and were coordinately suppressed following resupply (Connolly et al., 2003). The reduction of ferric iron to ferrous iron may be the rate-limiting step in iron uptake by Strategy I plants (Connolly et al., 2003). Strategy I plants cannot uptake iron chelates directly. The reduction-based Strategy 1 is prone to inhibition at high pH and elevated bicarbonate concentrations in calcareous soils. This is because of the pH dependence of both the chemical solubilisation of iron and the enzymatic reduction by FeIII -chelate reductase (Schaaf et al., 2004).

12.2.2

‘Strategy II’: phytosiderophores

Plants of the Poaceae (Gramineae, grasses) use the reduction-dependent pathway only in the minority of cases and adopt a second method in which low molecular mass ligand-forming compounds known as phytosiderophores are released by the root. These phytosiderophores are aminocarboxylate-type hexadentate (having six coordination sites) metal chelators with a high affinity for FeIII (Schaaf et al., 2004). Ferric iron mobilises from sparingly soluble precipitates, assisted by proton release. The constant ‘scavenging’ of ferric iron by chelation maintains some replenishment even if the solubility product of ferric iron is low. Phytosiderophores complex ferric iron, and the resulting complex is taken up via a specific Fe–phytosiderophore transporter. Fe–phytosiderophore transport is dependent upon proton co-transport

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3(S-adenosyl-L-methionine) [nicotianmine synthase EC 2.5.1.43] nicotianamine + 3(S-methyl-5′-thioadenosine)

nicotianamine + 2-oxoglutarate [nicotianamine aminotransferase EC 2.6.1.80] 3′′-deamino-3′′ oxonicotianamine + L-glutamate

3′′-deamino-3′′ oxonicotianamine + NAD(P) H + H+ [3′′-deamino-3′′ oxonicotianamine reductase EC 1.1.1.285] 2′-deoxymugineic acid + NAD(P)+

2′-deoxymugineic acid + 2-oxoglutarate + oxygen [2′-deoxymugineic acid-2′-dioxygenase EC 1.14.11.24] mugineic acid + succinate + carbon dioxide Figure 12.2

The pathway of biosynthesis of mugeneic acid from S-adenosyl-l-methionine.

and is also dependent upon the membrane potential (Schaaf et al., 2004). Naturally occurring iron chelates are not taken up by either plant group: Strategy I plants lack a chelate transporter and Strategy II plants generate their own chelating agent. The mugineic acid family of phytosiderophores are unique to graminaceous plants (Tagaki, 1976). Mugineic acid is closely related to a number of other compounds that also have been identified as phytosiderophores in graminaceous plants: 3-hydroxymugineic acid, 2 -deoxymugineic acid, avenic acid and distichonic acid. The binding coordinating groups are two amine-N, two carboxylate-O and one hydrocarboxylate site (two O), a total of 2 N’s and 4 O’s, in which the central FeIII atom resides. In the synthesis of mugineic acid from S-adenosyl-l-methionine (IUBMB, 2004), condensation of three S-adenosyl methionine molecules eventually (Figure 12.2) yields deoxymugineic acid (Takahashi et al., 1999). Mugineic acid and other members of the family are synthesised from 2 -deoxymugineic acid or from mugineic acid. The secretion of these phytosiderophores increases under iron

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deficiency, and this is correlated with the ability of the plant to overcome this stress. The intermediate, nicotianamine, is itself known to act as an intracellular metal chelator in all higher plants (von Wir´en et al., 1999; Schaaf et al., 2004). Nicotianamine synthase and nicotianamine aminotransferase are both strongly inducible by iron deficiency and their induction correlates with the amount of phytosiderophore secreted. There is a multigene family of nicotianamine synthase genes in the barley genome, and Northern analysis indicated that their expression was root specific and was induced by iron deficiency (Higuchi et al., 1999). The large quantities of phytosiderophores biosynthesised in relation to the extremely low steady-state concentration of the l-methionine precursor led to the proposal that there was a methionine-recycling system in operation (Ma et al., 1995). The phytosiderophore binds to FeIII and the resulting Fe–phytosiderophore is taken up by a specific uptake system. In maize roots, the high-affinity Fe– phytosiderophore transporter is necessary for healthy growth. The maize yellowstripe mutant ys1 (Beadle, 1929), a Mendelian recessive, results from iron deficiency caused by failure to take up iron (since foliar application alleviates the symptoms). The ys1 mutant cultivar releases the phytosiderophore, 2 -deoxymugineic acid, in similar quantities to that released in iron-efficient maize cultivars, but tracer studies showed that uptake of iron was 20-fold lower in ys1 than in the other cultivars. It was concluded, therefore, that the iron inefficiency of the ys1 mutant was due to a defective Fe–phytosiderophore uptake system (von Wir´en et al., 1994). The transport system (encoded by maize ZmYS1) also transported complexes of nickel, ferrous and ferric iron with nicotianamine, and it was concluded that the gene product was a proton-coupled broad-range metal–phytosiderophore transporter (Schaaf et al., 2004). ZmYS1 was also shown capable of transporting Fe–nicotianamine, as well as Cu–phytosiderophore, though the latter was thought to be of limited physiological significance (Roberts et al., 2004). Chelators are also important within the plant because of the potential toxicity of iron above ‘essential’ concentrations. Different chelators will be of differential importance at different pH values, 2 -deoxymugeneic acid being dominant at acidic pH and nicotianamine at alkaline pH, with implications for long-distance transport of metals in the xylem and phloem (von Wir´en et al., 1999). In view of its potential toxicity, iron is stored within cells using ferritin, an iron-storage protein. Ferritin subunits are joined to constitute a hollow sphere capable of storing thousands of atoms of iron (Connolly and Guerinot, 2002). There is evidence for a common ancestral ferritin gene in plants and a family of four genes is known to be present and expressed in the arabidopsis genome (Connolly and Guerinot, 2002). Plant ferritins have transit peptides that target delivery to the plastids (unlike animal ferritins that are cytosolic; Proudhon et al., 1996). Plant ferritins are regulated by iron both transcriptionally and post-transcriptionally, but plant ferritins have not yet been found to contain the iron-responsive elements that are partly responsible for translational control in animal systems (Connolly and Guerinot, 2002). Within the rhizosphere there are siderophores of microbial as well as plant origin. Rhizoferrin, produced by the fungus Rhizopus arrhizus, was as effective a source of iron as were synthetic Fe chelates for Strategy I plants (Yehuda et al., 1996).

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The ferric chelate reductase appears to function with a wide range of chelates. Fe– rhizoferrin was also an efficient source for Strategy II plants, but does not appear to be taken up directly. Comparison of 59 Fe uptake from Fe–rhizoferrin, Fe–EDTA and Fe–phytosiderophore indicated that uptake and translocation of iron from all sources paralleled the diurnal pattern of phytosiderophore release (Yehuda et al., 1996): it was concluded that uptake of iron by plants from microbial siderophores was indirect and mediated by ligand exchange between ferrated microbial siderophores and phytosiderophores. This seems to indicate that the Fe–phytosiderophore transporter cannot transport ferrated microbial siderophores. The graminaceous Fe– phytosiderophore transporter recognises the backbone structure of mugineic-acidderived phytosiderophores (Schaaf et al., 2004).

12.3

Phosphate uptake in soils that are low in phosphate

Plant growth is limited by availability of inorganic phosphate in most natural ecosystems, and as a consequence a range of adaptive strategies have evolved to enable plants to increase the acquisition of phosphate (Smith et al., 2003). Plants are not able to directly utilise any organic phosphorus in the soil until it has been mineralised into phosphate. The resulting phosphate is involved in complex equilibrium reactions, which means that the steady-state concentration of phosphate in the soil solution is below 10 μM even in fertile soils and below 1 μM in many cases (Smith et al., 2003). The implications of this for high-affinity phosphate transport are considered more fully in Chapter 11. The phosphate transport system can cope with very low concentrations, provided that the replacement rate is adequate. The concern in this Section is over how plants deal with situations where the replacement rate is limiting and phosphate deficiency results. Phosphate movement in the soil is mostly by diffusion (Smith et al., 2003), and the slow rate of this diffusion (see Section 3.6.2) results in a depletion zone around the roots, particularly in soils with a low concentration of phosphate (Smith, 2002). Because the concentrations are so low, the driving forces (concentration differences) are also small.

12.3.1

Cluster roots and root exudates

A range of anatomical adaptations have been reported that increase the density of root apices and root hairs per unit volume of soil, commensurate with exploiting a volume of soil in which the distance that a nutrient diffuses is very small. More specialised adaptations include the localised development of lateral roots in areas of relatively phosphate-rich patches (Drew and Saker, 1978; Jackson et al., 1990) and the development of cluster, or proteoid, roots (Dinkelaker et al., 1995; Lambers and Poot, 2003). Cluster roots are characteristic of species of the Proteaceae that are native to phosphate-deficient soils in Australia. These soils are ancient and leached, and their composition binds phosphate tightly, so there has been long-term opportunity for the evolution of adaptive mechanisms. Clusters are rows of rootlets that are determinate in growth pattern: that they do not go on growing sets a boundary

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to what could be a demanding overinvestment in root production. Furthermore, their development is suppressed by increased phosphate supply (De Vos et al., 2001). There are also parallels with the mobilisation of iron. Acidification of the soil around the root as a consequence of proton extrusion (Neumann and R¨omheld, 1999) increases the availability of phosphate. The release of exudates can also help free phosphate from complexes in the soil. These exudates include (but are not restricted to) the carboxylates citrate and malate, and can vary according to the chemical nature of the phosphate supplied (Lambers et al., 2001; Roelofs et al., 2001). Root clusters are ephemeral, and in addition carboxylates are released only in ‘exudative bursts’ lasting a few days (Shane and Lambers, 2005). Release of carboxylic acids is associated with cluster roots but has also been associated with increasing phosphate availability in species that do not exhibit pronounced morphological adaptation. Exudation of acid phosphatases can release phosphate from organic phosphorus complexes in the soil. Keerthisinghe et al. (1998) found no evidence that the K m for high-affinity phosphate transporters differed between cluster roots as opposed to normal lateral roots, but the V max for phosphate transport was found to be greater, suggesting that there may be a greater number of transporter proteins in the cluster roots. Cluster roots showing anatomical, physiological and biochemical adaptation to exploit a volume of soil to the maximum therefore represent a coordinated response requiring the regulation of very diverse characteristics. Some species of the Proteacea store phosphorus in stem tissues seasonally, and can develop symptoms of phosphorus toxicity at relatively low external levels of supply when the storage capacity is exceeded. Also, the capacity to down-regulate phosphorus uptake at supra-optimal phosphorus supply is needed to prevent phosphorus toxicity (Shane and Lambers, 2005).

12.3.2

Mycorrhizal symbiosis

The hyphae of mycorrhizal fungi extend considerable distances from the roots. This provides an alternative method of increasing the volume of soil that the root can access. Hyphae can respond to localised sources of nutrient more rapidly than can the roots of plants because, being narrow, they elongate more rapidly and with less resources than a root. Fungal associations also increase the range of forms of phosphorus that are available to the plant. Most plants are able to form such symbiotic associations but this is unusual amongst plants that form cluster roots, seemingly establishing these as an alternative (Smith et al., 2003). The benefit of mycorrhizal association to the plant is that the investment in resources as part of the symbiotic association is less than that required to exploit the same volume of soil through developing more roots. Plants with coarse, unbranched roots and few root hairs will benefit more from mycorrhizal association than will plants with fine, much-branched roots and plentiful root hairs. The development of mycorrhizal associations is dependent upon the supply of phosphate, and the rate of infection is suppressed above about 140 ppm bicarbonatesoluble phosphate (Amijee et al., 1989). Fertilisation with phosphate decreases both the rate and percentage of infection. If the availability of phosphate is greater than

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that needed for plant growth then the arbuscules (the sites of transfer between plant and fungus) are no longer formed (Abbott and Robson, 1979). Plants may commit to the investment of resources in fungal symbiosis only when ‘needed’. So far the phosphate transporters of fungal symbionts have been found to be similar to those of the plant Pht1 family (Smith et al., 2003).

12.4

Toxicity and tolerance – aluminium in acid soils

Aluminium is the single most common constituent of the earth’s crust and is present in most soils in considerable quantities. However, it is only soluble aluminium that is taken up by plants. The solution properties of aluminium are complex. It is present as Al3+ , (AlOH)2+ and Al(OH 2 )+ cations. Of these ions, the Al3+ cation is the most phytotoxic and dominates at acidic pH values below about 5. Above pH 7.5, aluminium again becomes soluble as the aluminate anion (Al(OH) 4 − ; McBride, 1994). It is because of the almost universal presence of aluminium in parent rocks that any weathering process under acidic conditions, or any other process resulting in soil acidification, will lead to the solubilisation of aluminium as Al3+ . Soils characterised by aluminium toxicity often contain toxic concentrations of manganese and comprise some 40% of the land area used for arable farming (McNeilly, 1994). Manganese is present in soils in lesser quantities than is aluminium and is needed as a micronutrient by plants, but acidic conditions can lead to phytotoxic concentrations of soluble ionic manganese. Liming to increase the pH is the major practice in crop husbandry, but the utility of this is limited by the sheer scale of the areas affected, by the time needed to adjust the whole depth of the root zone and by access to sufficient and economical supplies of lime, particularly for subsistence farmers. The ‘calcicole/calcifuge problem’ was ascendant in the ecological literature of the 1960s (Rorison, 1960; Clymo, 1962; Grime and Hodgson, 1969), and much research pointed to lack of tolerance to aluminium as the crucial factor excluding lime-loving (calcicole) species from acid soils. The implication is that in these species, the genetic information required for the tolerance to aluminium is absent. The effects of aluminium inside and outside the cell are manifold, but the primary effect of aluminium is to stunt and distort root development. Early research (Clarkson, 1965) showed that following exposure to aluminium (or other trivalent metals), cessation of root elongation was closely correlated with the disappearance of mitotic figures, and it was concluded that cell division is highly sensitive to, and is damaged permanently by, short exposures to aluminium. Aluminium interacts with the cytoskeleton of roots and has been shown to affect tubulin polymerisation in vitro (MacDonald et al., 1987). Interference with the cytoskeleton has been held responsible for inducing the swelling that has been observed in root apices and tips of root hairs. Within 1 h, treatment with aluminium resulted in a depletion of microtubules in the distal part of the transition zone of the maize root apex, particularly in the outermost file of cortical cells. Over longer periods of time, treatment with aluminium led to the development of increased

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lesions to the microtubule cytoskeleton in the epidermal and inner cortical files of cells (Sivaguru et al., 1999). There was concomitant, or previous, aluminiuminduced depolarisation of the plasma membrane. Aluminium has also been found to disrupt mitochondrial activity in cultured cells of tobacco and in roots of pea. Aluminium rapidly repressed mitochondrial activity leading, in 12 h, to the production of reactive oxygen species, which were suggested to be the cause of subsequent depletion of ATP and correlated inhibition of root elongation (Yamamoto et al., 2002). Yamamoto et al. (2002) considered that the production of reactive oxygen species was probably the critical event in aluminium inhibition of cell growth. Aluminium-enhanced peroxidation of lipids has been observed, mediated by ferrous ions, in soybean roots (Cakmak and Horst, 1991). Aluminium has also been widely reported to interfere with phosphate uptake and metabolism and with calcium uptake. It is the Al3+ cation that is the principle toxic species of aluminium, and in chelated form aluminium is not phytotoxic. The chelating ability of organic acids now appears to have a key role in tolerance to aluminium through the pre-emptive secretion of organic acids as ligands, the effect of which is the chelation of aluminium in the soil before it enters the root. Developed possibly from the serendipitous benefit of leakage of organic acids by the root, a mechanism comprising organic acid secretion via an aluminiumstimulated channel has been elucidated over recent years. The release of aluminiumchelating ligands, activated by the presence of aluminium, may have a role in the aluminium tolerance of a number of plant species (Ma et al., 2001). In roots of an aluminium-tolerant maize cultivar, the addition of aluminium activated, almost instantaneously, a concentration-dependent release of citrate: half the V max for citrate release was induced by the addition of only 20 μM Al3+ (Pineros et al., 2002). Patch-clamp studies confirmed that there was an aluminium-induced anion channel located in the plasma membrane of both cortical and stelar protoplasts. There was also a separate, aluminium-induced, increase in the citrate content of the root, operating over a longer timescale (Pineros et al., 2002). The authors interpreted these observations as indicating that there was a second mechanism of tolerance in operation, in addition to external chelation. This was based upon internal tolerance to aluminium by chelation and compartmentalisation and hence the increase in bulk citrate as well as the secretion of citrate. An ameliorative effect of silicon upon aluminium toxicity in maize was attributed to entrapment of aluminium in the cell wall, in the form of a hydroxyaluminosilicate complex, thereby reducing the apoplastic mobility of aluminium (Wang et al., 2004). This organic acid defensive mechanism has two requirements – the synthesis of extra organic acids and the mechanism to put them effectively into the root environment. The transport of molecules to the environment appeared to be the rate-limiting step in plants that simply produce, or overproduce, organic acids (Ryan et al., 2001). Since then, the details of the aluminium-activated efflux of organic anions from roots have become well established (Delhaize et al., 2004) and are known as the ‘malate hypothesis’. An aluminium-activated malate-permeable channel was identified in the plasma membrane of root apices (Ryan et al., 1997). The responsible gene (ALMT1) has subsequently been cloned and when expressed in other organisms

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confers an aluminium-activated efflux of malate. When expressed in Hordeum vulgare, ALMT1 conferred aluminium-activated malate efflux and was associated with an increase in the tolerance to aluminium, both in hydroponics and in an acidic soil (Delhaize et al., 2004).

12.5

Toxicity and tolerance – essential and non-essential metals

Some metals, such as copper and zinc, are essential elements, but, together with non-essential metals all can be phytotoxic at elevated concentrations. Non-essential metals whose toxicity is often apparent are, for example, cadmium, chromium, lead and mercury. The response of plants to the semi-metal, arsenic, is usually considered in this group, because there are strong similarities with metals in the modes of uptake and detoxification by plants. Arsenic (and the anions arsenate and arsenite) is found ubiquitously in the environment and there is fallout from smelting and burning of fossil fuels; arsenic is also known to pollute groundwaters (Schm¨oger et al., 2000). Iron and manganese are usually considered separately, since the areas of land affected by their potential toxicity are so very much greater than for those listed above. Toxicity due to non-essential metals generally has limited impact on agricultural production because the areas affected by such metals are relatively small and isolated, although there will be an impact on local communities. There is interest in metal tolerance for the use of plants for reclamation of sites made barren as a result of toxicity of anthropogenic origin, for stabilisation and landscaping and possibly for phytoremediation (the ‘detoxification’ of contaminated sites by planting and harvesting tolerant plants). It has also been proposed to use metal-tolerant plants in biological ‘mining’. The rapid development of tolerant ecotypes of certain species in response to recent anthropogenic pollution has presented a valuable model for studies of natural selection in plant communities in the wild. The occurrence of soils containing phytotoxic levels of non-essential metals in the natural environment is considered too rare for there to have been sufficient selection pressure for the evolution of metal detoxification systems (O’Leary, 1994). By implication, any processes that are used by plants to tolerate such metals are by virtue of chance recruitment. That few species possess the genetic variation to develop metal tolerance is suggested by the very small proportion of the adjacent flora that have succeeded in colonising metalliferous rocks such as the Great Dyke in Zimbabwe (Wild and Bradshaw, 1977), even though these outcrops predate the evolution of the angiosperms (McNeilly, 1994). Since a number of metals (copper, iron and zinc) serve essential roles in small quantities, some of the protective systems (chelation and compartmentalisation) that are found in plants probably have their origin in controlling excessive uptake of these essential elements and providing homeostasis about the plant. There are three different mechanisms of metal toxicity (Sch¨utzend¨ubel and Polle, 2002): 1. Production of reactive oxygen species by auto-oxidation and by the Fenton reaction. Typical of iron and other transition metals.

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2. Blocking of essential functional groups in biomolecules. Typical of nonredox-reactive heavy metals such as cadmium and mercury, which can also cause oxidative stress, lipid peroxidation and accumulation of dihydrogen peroxide. 3. Displacement of essential metal ions from biomolecules. Typical of a range of metals. In the Fenton reaction, ferrous iron is oxidised to ferric iron by interaction with dihydrogen peroxide and generates the hydroxyl radical, possibly via an oxoiron(IV) intermediate (IUPAC, 2003). The hydroxyl radical production cannot be controlled by antioxidants (Sch¨utzend¨ubel and Polle, 2002). There is also a range of mechanisms potentially available for detoxification of and tolerance to metals. These include (Hall, 2002): r r r r r r r r

mycorrhizae; binding to the cell wall; chelation by extracellular exudates; reduced uptake; efflux pumping at the root plasma membrane; chelation in the cytosol by phytochelatins and other ligands; repair of stress-damaged proteins; compartmentalisation of metal chelates within the vacuole via tonoplastlocated ATPases; r compartmentalisation of metal ions in the vacuole via tonoplast proton antiporters. Uptake of non-essential metals is generally assumed to be via ‘mistaken identity’, through competition for sites on inadequately specific transporters ‘intended’ for other ionic species. Potential mechanisms that might allow detoxification of and thus tolerance to metals are concerned with avoiding the achievement of toxic concentrations at critical sites rather than resisting the effects of metals (Hall, 2002). The root plasma membrane may be the first target to be affected by the above interactions with membrane lipids and key transport proteins; however, there is little direct evidence for efflux transporters in plants (Hall, 2002). The information on plant response to metals is scattered over a number of metals of the group, as necessarily are the examples used below. The theme appears, as of now, to be that the generic defences are through chelation and compartmentalisation and that this is again a serendipitous extension of systems that may originally have functioned to maintain the free concentrations of essential metals within fairly tight limits. Much information comes from a group of plants that show an extreme response and accumulate metals in large amounts – the hyperaccumulators.

12.5.1

Hyperaccumulation

A small group of species are notable not only for colonising metalliferous soils but for being ‘hyperaccumulators’ of otherwise toxic metals: one of them in particular,

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Thlaspi caeruleans, has become something of a model species and will thus often be noted. The term hyperaccumulator was originally used to describe plants that were able to hyperaccumulate nickel (Brooks et al., 1977). The term is used for a number of metals and the definition of ‘hyper’ varies with the element involved, but is taken to mean the ability to accumulate 100- to 1000-fold greater concentrations in the shoot than in ‘normal’ non-accumulator counterparts (McGrath et al., 2002). Some 400 taxa of terrestrial plants are categorised as hyperaccumulators of various metals including 300 nickel accumulators (Baker and Brooks, 1989; Brooks, 1998; Salt et al., 1998). Hyperaccumulators of zinc are less common; 16 species are able to do this (Brooks, 1998). T. caeruleans is able to accumulate zinc to at least 25 000 μg g−1 dry mass in hydroponics without evidence of toxicity or reduction in growth (Brown et al., 1995; Shen et al., 1997). In non-accumulator plants, zinc concentrations of 100 μg g−1 are sufficient and 300–500 μg g−1 are toxic (Mengel and Kirkby, 1987). Pteris vittata, the first identified arsenate hyperaccumulator, was able to contain 27 000 μg g−1 of arsenate in its fronds, though signs of phytotoxicity appeared above 10 000 μg g−1 (Wang et al., 2002). The hyperaccumulators contrast with other species adapted to metal-rich soils which function by exclusion of metals from the shoots (Baker and Brooks, 1989). Arsenate resistance is generally achieved through suppression of the high-affinity phosphate uptake system, by which arsenic is taken up through non-specificity (Meharg and Hartley-Whittaker, 2002). More than a curiosity, the hyperaccumulators have received interest as phytoremediators of toxic soils, such as mine waste (McGrath et al., 1993). The practical utility of many species, including the well-researched T. caeruleans, for this purpose is limited by their small biomass production (Ebbs et al., 1997), which has led to the interest in looking for genes to transfer into more vigorous species (Pence et al., 2000).

12.5.2

Ion transport in hyperaccumulators

Comparisons of zinc uptake by roots showed that the K m values were similar but that V max was five times greater in T. caeruleans than in the non-hyperaccumulator Thalspi arvense, indicating that there were more zinc transport sites in the hyperaccumulator (Lasat et al., 1996). A metal transporter cDNA (ZNT1) cloned from T. caeruleans was demonstrated to mediate high-affinity Zn2+ and low-affinity Cd2+ uptake. The transporter was expressed at very high levels in both roots and shoots. The zinc status of the plant regulated the expression of the ZNT1 gene and resulted in overexpression of the ZNT1 transporter and increased influx of zinc in the roots (Pence et al., 2000). T. caeruleans is the only known hyperaccumulator of cadmium (Baker et al., 2000). Many scattered examples suggest that chelation and compartmentalisation (as chelate or as free ions) are generic methods of metal tolerance. In the hyperaccumulator Arabidopsis halleri most of the zinc in the aerial parts was octahedrally coordinated and complexed to malate, and a minor proportion was tetrahedrally coordinated and complexed to carboxyl and/or hydroxyl groups (Sarret et al., 2002). In plants taken from contaminated soil, the zinc in the roots was found as zinc malate,

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zinc citrate and zinc phosphate, though in hydroponics only the phosphate form was found, and presumed to have arisen through precipitation (Sarret et al., 2002): zinc phosphate was present in the aerial parts of the related non-accumulator Arabidopsis lyrata subsp. petraea. In P. vittata arsenate was reduce to arsenite for transport from root to shoot and was stored as such in the fronds: it was concluded to be sequestered free within the vacuole, there being no detection of As–phytochelatin complexes, neither was there sufficient sulphur to provide coordination at the expected (3:1) stoichiometry (Wang et al., 2002). The quantities of arsenic accumulated by P. vittata are exceptional and vastly exceed the capacity of phytochelatins to bind them, making compartmentalisation of inorganic arsenic the only available method. In T. caeruleans, nickel was found to be associated with the cell wall and localised in the vacuole as a Ni–organic acid complex (Kr¨amer et al., 2000). Also in this hyperaccumulator, zinc was found to be sequestered in soluble form in the vacuoles of, predominantly, epidermal cells (K¨upper et al., 1999). Preferential accumulation of zinc in epidermal cells was observed even when the growth medium did not contained elevated concentrations of zinc. Single-cell sap sampling showed that the concentration of zinc in the vacuolar sap of epidermal cells was five or more times that in mesophyll cells and achieved a maximum concentration of 385 mM in plants whose shoots had 2% zinc on a dry mass basis (K¨upper et al., 1999). Zinc tolerance in Silene vulgaris was not related to reduced uptake, nor was it related to phytochelatin synthesis (Chardonnens et al., 1999). Split-root experiments indicated that tolerance resided in root cells (Harmens et al., 1993). Tonoplast vesicles of a tolerant ecotype were able to accumulate more zinc than could vesicles isolated from a sensitive ecotype (Verkleij et al., 1998). Zinc uptake at the tonoplast in S. vulgaris was concluded to be via more than one parallel pathway, the proportions of which differed between ecotypes, actively transporting zinc (and since the accompanying citrate was not accumulated) probably as a free ion (Chardonnens et al., 1999). Nickel hyperaccumulation in Thlaspi goesingense appears in part due to compartmentalisation of nickel within leaf vacuoles. A putative vacuolar metal transport protein from T. goesingense had the features of the cation-efflux family, complemented metal sensitivity in yeast strains lacking the orthologue and was expressed highly in T. goesingense compared with expression in metal-sensitive close relatives (Persans et al., 2001). There are some observations indicating some immobilisation in cell walls, but subcellular localisation is generally vacuolar. Compartmentalisation within the vacuole may be as a free ion.

12.5.3

Phytochelatins

Enzymatically synthesised peptides that formed ligands to cadmium were first described in plants in 1985 and termed phytochelatins (Grill et al., 1985). Metallothioneins are phytochelatins found in many types of organisms and are gene-encoded polypeptides that bind certain metals. These molecules are not primary gene products but have been categorised as Class III metallothioneins (Robinson et al., 1993) to separate them from the more typical metal-binding proteins. Phytochelatins are typically produced in response to acute rather than chronic metal stress and (based

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again upon the lack-of-selection-pressure argument, see above) are considered to be a serendipitous recruitment to metal tolerance (O’Leary, 1994). Their original, and major, role was possibly to deal with excess concentrations of essential metal elements. The phytochelatins function by complexing the metal ions via thiolate coordination (Grill et al., 1985). They are derived from glutathione by a constitutive phytochelatin synthase (PCS) and have a general structure (γ -glutamate-cysteine)n glycine, where n ranges from 2 to 11 (Grill et al., 1989; Zenk, 1996). The gene encoding PCS has been cloned (Clemens et al., 1999; Ha et al., 1999; Vatamaniuk et al., 1999) and is activated by metal ions such as Cd2+ , Cu2+ , Ag+ , Hg2+ and Pb2+ . A wheat cDNA (TaPCS1) was identified (Clemens et al., 1999), which conferred a dramatic increase in cadmium tolerance when it was expressed in the yeast Saccharomyces cerevisiae. TaPCS1 encoded a 55-kDa protein (phytochelatin synthase – PCS) of no previously identified function, and homologues to it were identified in A. thaliana. Cells expressing PCS were cadmium tolerant and accumulated more cadmium than did cells that did not express PCS, and the tolerance that they conferred was lost if glutathione biosynthesis was inhibited (Clemens et al., 1999). Brassica juncea overexpressing a bacterial homologue of PCS showed increased tolerance to cadmium, accumulating more cadmium and growing better in the presence of cadmium, as compared to wild-type plants (Zhu et al., 1999). Arsenic is an efficient inducer of phytochelatins and an approximate ratio of three sulphydryl groups to one atom of arsenic is compatible with properties of As–glutathione complexes (Schm¨oger et al., 2000) The authors demonstrated by electrospray ionisation mass spectroscopy and other methods that arsenic-induced phytochelatins resulted in the complexation and detoxification of arsenic. Although phytochelatins have a role in cellular metal homeostasis, it has been questioned whether there is evidence that they are involved in differential wholeplant metal tolerance (Schat and Kalff, 1992). These authors cite inconsistent reports of whether there are or are not higher levels of phytochelatins in tolerant than sensitive plants/cell lines. They (Schat and Kalff, 1992) also criticise the interpretation of the effects of buthionine sulphoximine (an inhibitor of γ -glutamylcysteine synthesis, which reduces tolerance). The argument is that since phytochelatins are important in metal homeostasis in all plants, then all plants, regardless of whether they are or are not tolerant to excess metals, will be harmed by the inhibitor. In populations of S. vulgaris, concentrations of phytochelatins in the root apex were related to the toxicity of copper (e.g. that concentration of copper causing 50% inhibition of root elongation rate) and not to the absolute concentration of copper; therefore, the difference in tolerance between populations was not dependent on differential phytochelatin production (Schat and Kalff, 1992). There is much current work on phytochelatins. At present it would appear that their original function was in the homeostasis of metals that have a role as essential nutrients but are disruptive if present as their free ions at higher concentrations. The use of phytochelatins may have been extended to cope with low-level chronic exposure to toxic metals, but is not the major method of tolerance in those plants that deal with very elevated concentrations of metals in their tissues. The hyperaccumulators compartmentalise metals in the vacuole – probably as the free ions, simply because

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of the enormous quantities involved. There remains a major role for chelation in transporting metals around the plant. The concentration of free metals in the vacuole probably underlies the herbivore-defence role proposed for the evolution of the hyperaccumulation of toxic metals.

12.5.4

Function of hyperaccumulation

Of proposed explanations for the evolution of metal hyperaccumulators, the ‘elemental defence hypothesis’ has supporting evidence: plant tissues with high-metal concentrations have been shown to be toxic to generalist insect herbivores and pathogens or to generally deter herbivory (Boyd and Martens, 1998; Boyd et al., 2002). There is some evidence that metal accumulation below ‘hyperaccumulator’ levels may also serve as defence, establishing the possibility of an evolutionary pathway. Snails (Helix aspersa) feeding on hyperaccumulator (Senecio coronatus) leaves contained tenfold the quantity of nickel, and suffered greater mortality, than those feeding on non-hyperaccumulator leaves. The time to 50% mortality in the snail population was reduced from about 25 days at low concentrations of nickel to about 10 days for nickel concentrations (as a supplement to a cornmeal diet) of 1600 and 3200 μg g−1 dry mass; leaves of S. coronatus may contain up to 12 100 μg g−1 (Boyd et al., 2002). Browsing can lead to metal accumulation in large herbivores: tungsten and copper were found to be accumulated in bones of cattle that had fed on metalliferous plants in Australia (Pyatt and Pyatt, 2004) and such accumulation may have deleterious effects upon the animals. Recent observations have indicated that metal concentrations well below ‘hyperaccumulator’ levels are still effective at deterring herbivory. All of the metals, cadmium, cobalt, chromium, copper, manganese, nickel, lead and zinc, were toxic to moth larvae of the crucifer specialist Plutella xylostella, not only at ‘hyperaccumulator’ concentrations, but at lower ‘accumulator’ concentrations: cadmium, manganese, nickel, lead and zinc remained toxic at still lower concentrations (Coleman et al., 2005).

12.6

Concluding remarks

Plants possess mechanisms that aid the acquisition of nutrients that are present but are of limited availability in the soil. Plants possess mechanisms to detoxify metals that are present at phytotoxic concentrations and can achieve this both in the soil and in the plant. Plants use organic ligand-forming compounds ranging from simple organic acids to phytochelatins and phytosiderophores to achieve this. The systems incorporate specific transporters, for moving chelating agents out into the soil and for moving metal–chelate complexes into the plant. The systems also include other specific additions such as FeIII -chelate reductase. Evolutionary considerations argue that the tolerance to high concentrations of metals, including non-essential metals, is by recruitment and extension of original roles in homeostasis. A coordinated morphological, physiological and biochemical response (exemplified by proteoid or cluster roots) has evolved to maximise phosphate acquisition in soils with low

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availability of phosphate. Symbiotic associations with mycorrhizal fungi are an alternative ‘strategy’ for increasing access to phosphate.

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Persans, M.W., Nieman, K. and Salt, D.E. (2001) Functional activity and role of cation-efflux family members in Ni hyperaccumulation in Thlaspi goesingense. Proceedings of the National Academy of Sciences 98, 9995–10000. Pineros, M.A., Magalhaes, J.V., Carvalho Alves, V.M. and Kochian, L.V. (2002) The physiology and biophysics of an aluminium tolerance mechanism based on root citrate exudation in maize. Plant Physiology 129, 1194–1206. Proudhon, D., Wei, J., Briat, J.F. and Theil, E.C. (1996) Ferritin gene organisation: difference between plants and animals suggest possible kingdom-specific selective constraints. Journal of Molecular Evolution 42, 325–336. Pyatt, F.B. and Pyatt, A.J. (2004) The bioaccumulation of tungsten and copper by organisms inhabiting metalliferous areas in North Queensland, Australia: an evaluation of potential health implications. Journal of Environmental Health Research 3, 13–18. Roberts, L.A., Pierson, A.J., Panaviene, Z. and Walker, E.L. (2004) Yellow stripe 1. Expanded roles for the maize iron-phytosiderophore transporter. Plant Physiology 135, 112–120. Robinson, N.J., Proctor, C.M., Connolly, E.L. and Guerimot, M.L. (1999) A ferric-chelate reductase for iron uptake from soils. Nature Biotechnology 397, 694–697. Robinson, N.J., Tommey, A.M., Kuske, C. and Jackson, P.J. (1993) Plant metallothioneins. Biochemical Journal 295, 1–10. Roelofs, R.F.R., Rengel, Z., Cawthray, G.R., Dixon, K.W. and Lambers, H. (2001) Exudation of carboxylates in Australian Proteaceae: chemical composition. Plant, Cell and Environment 24, 891–904. Rorison, I.H. (1960) Some experimental aspects of the calcicole-calcifuge problem. II: The effects of mineral nutrition on seedling growth in nutrient solution. Journal of Ecology 48, 679–688. Ryan, P.R., Delhaize, E. and Jones, D.L. (2001) Function and mechanism of organic acid excretion from plant roots. Annual Review of Plant Physiology and Plant Molecular Biology 52, 527–560. Ryan, P.R., Skerrett, M., Findlay, G.P., Delhaize, E. and Tyerman, S.D. (1997) Aluminium activates an anion channel in the apical cells of wheat roots. Proceedings of the National Academy of Sciences 94, 6547–6552. Salt, D.E., Smith, R.D. and Raskin, I. (1998) Phytoremediation. Annual Review of Plant Physiology and Plant Molecular Biology 49, 643–668. Sarret, G., Saumitou-Laprade, P., Bert, V., et al. (2002) Forms of zinc accumulated in the hyperaccumulator Arabidopsis halleri. Plant Physiology 130, 1815–1826. Schaaf, G., Ludewig, U., Eronoglu, B., Mori, S., Kitahara, T. and von Wir´en, N. (2004) ZmYS1 functions as a proton-coupled symporter for phytosiderophore- and nicotianamine-chelated metals. The Journal of Biological Chemistry 279, 9091–9096. Schat, H. and Kalff, M.M.A. (1992) Are phytochelatins involved in differential metal tolerance or do they merely reflect metal-imposed strain? Plant Physiology 99, 1475–1480. Schikora, A. and Schmidt, W. (2001) Iron stress-induced changes in root epidermal cell fate are regulated independently from physiological responses to low iron availability. Plant Physiology 125, 1679–1687. Schm¨oger, M.E.V., Oven, M. and Grill, E. (2000) Detoxification of arsenic by phytochelatins in plants. Plant Physiology 122, 793–801. Sch¨utzend¨ubel, A. and Polle, A. (2002) Plant responses to abiotic stresses: heavy metal-induced oxidative stress and protection by mycorrhization. Journal of Experimental Botany 53, 1351– 1365. Shane, M.W. and Lambers, H. (2005) Cluster roots: a curiosity in context. Plant and Soil 274, 101–125. Shen, Z.G., Zhao, F.H. and McGrath, S.P. (1997) Uptake and transport of zinc in the hyperaccumulator Thlaspi caeruleans and the non-hyperaccumulator Thlaspi ochroleucum. Plant, Cell and Environment 20, 898–906. Sivaguru, M., Baluska, F., Volkmann, D., Felle, H.H. and Horst, J. (1999) Impact of aluminium on the cytoskeleton of the maize root apex. Short-term effects on the distal part of the transition zone. Plant Physiology 119, 1073–1082. Smith, F.W. (2002) The phosphate uptake mechanism. Plant and Soil 245, 105–114.

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13 Water-limited conditions Anthony Yeo

13.1

Introduction

This chapter is concerned with growth and survival in water-limited conditions, except specifically for the tolerance of desiccation (Chapter 15) and of salinity (Chapter 14), which are dealt with separately. Desiccation tolerance is a special case because it means the absence of, and not merely the reduced availability of, water. Salinity tolerance is a special case because it combines the reduced availability of water with exposure to very elevated concentrations of inorganic ions. Drought tolerance is an area in plant physiology in which there is the greatest confusion and discrepancy between the ‘ecological’ and the ‘agricultural’. What is ‘good for the plant’ in terms of ecological success (which is measured as surviving to complete the life cycle) is commonly not ‘good for the crop’ (which is measured as yield or yield stability). There is logic in looking at nature’s survivors for clues to how plants can deal with a stressful environment. But the caveat is, that which confers survival may not, and frequently does not, imply a yield advantage. In fact, the mechanisms conferring ecological success are often diametrically opposed to those linked to yield. This book is about plants, not specifically about crops, and so many mechanisms that are only of ecological advantage will be considered, while keeping in focus the caveat that these may be of zero, or negative, agricultural value. Water availability is a major factor in plant zonation. About 47% of the land surface is categorised as ‘dryland’ with a precipitation to potential evapotranspiration ratio (P/PET) of less than 0.65. This has ‘drier’ subdivisions and 17.7% of the land surface is categorised as semi-arid (P/PET 0.20–0.45), 12.1% as arid (P/PET 0.05– 0.20) and 7.5% as hyper-arid (P/PET < 0.05) (Hou´erou, 1996). Consequently the categories of ‘dryland’ cover a continuum of environments, from irrigated and waterlimited farmland to deserts. There is a difficult and inconsistent terminology referring to ‘avoidance’, ‘resistance’ and ‘tolerance’. Water deficit in some form is an inimical feature of plant life on land and something to which nearly all species are subjected, whether at some times of the day, some part of the growing season or at some seasons and not others. Plant water deficit arises whenever rates of water loss exceed rates of water gain. Water gain is most commonly limited by the availability of water in the soil. All species possessing stomata have control over water loss, so none can be deemed as totally lacking in any adaptation to deal with water deficit. How much water deficit a plant is exposed to, and to what extent it copes, exists as a continuum. Sinclair and Ludlow (1986) categorised water deficits in terms of stage I, stage II and stage III

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drought – subdivisions with a clear mechanistic and physiological basis. The stages of increasing soil water deficit correspond to: I. there is enough water for the stomata to be fully open, and water loss is dependent upon atmospheric conditions (temperature, humidity and windspeed); II. stomatal regulation can maintain plant water status (water availability cannot support maximum stomatal conductance but control of conductance can match loss of water to gain of water, although conductance will reduce throughout this stage) and; III. plants cannot maintain water status even with the stomata shut. The range of responses found in plants varies according to the severity and probability of drought; this extends from stomatal regulation to the more extreme anatomical adaptations occurring in species that have evolved in environments that are regularly arid. In this respect it has to be considered that some of these adaptations are pre-emptive and their function is to conserve soil water and thus prevent, or delay so long as possible, entry into stage III drought. Only the desiccation-tolerant ‘resurrection plants’ (Chapter 15) are adapted to crossing the boundary into stage III and back again in the vegetative state. Drought represents the interaction between atmospheric and soil physical characteristics, nutrient availability and biological factors (pests, pathogens and competing plants), making it difficult to define ‘typical drought’ (Price et al., 2002). Organisms that feed on roots can have substantial detrimental effect upon drought response (Audebert et al., 2000), and arbuscular mycorrhizae can increase plant tolerance to drought (Porcel and Ruiz-Lozano, 2004). This chapter addresses growth in water-limited conditions in the broadest sense, and this is functionally interrelated with heat tolerance and includes aspects of freezing tolerance. One major role of solutes is in osmotic adjustment to help compensate for low external water potentials. Some solutes may also have a role in protecting cellular structures and macromolecules from damage at low water potentials inside the cell. Another major role of solutes is in inter- and intracellular transport of carbon in C 4 and CAM (crassulacean acid metabolism) photosynthesis. Solute transport is central to changing stomatal aperture. Water-limited conditions arise principally through lack of water availability to the roots, usually because the soil is dry, but it may also be frozen or waterlogged. Plants may suffer water deficit through excessive demand: in hot, dry and windy conditions, water loss can exceed water supply via the xylem, even when the soil is wet. Plants moved from protected culture to open sites may wilt even when watered, because they do not have enough roots to cope with the increased atmospheric demand upon their leaves. Plants can also suffer water deficit through other causes such as dehydration by extracellular ice in freezing damage.

13.2

Plant responses to reduced water availability

Responses of plants to reduced availability of water fall broadly into a number of non-exclusive classes:

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1. rapid changes in stomatal conductance and leaf orientation to conserve water and minimise thermal load; 2. changes in stomatal conductance following perception and signalling of water deficit by the roots (including longer term effects upon stomata); 3. regulation of leaf area expansion and total leaf area to match growth to water availability (this may be by functional balance or be pre-emptive, using longdistance signalling of root water status). 4. enhanced nutrient uptake and/or biochemical adaptations that provide osmotic adjustment; 5. allocation of resources to roots to increase the exploration and exploitation of soil water (this may be by functional balance, by competition between sinks within the plant or by genetically mediated allocation); 6. anatomical adaptations to limit water loss and/or to reduce thermal load, and/or provide ‘water storage’; 7. biochemical adaptations that provide increased water-use efficiency, including C 4 and facultative/obligate CAM photosynthesis. The time frame of processes ranges from rapid to constitutive as follows: Rapid initiation, rapidly reversible: r regulation of stomatal conductance by plant water status; r regulation of axial conductance of water; r leaf movements to reduce thermal load/transpiration. Short-term initiation, medium- to long-term consequences, not always/completely reversible: r regulation of stomatal conductance by remote sensing/signalling; r effects of stomatal density upon leaf conductance; r regulation of leaf area via cell division and expansion (turgor/water availability); r regulation of leaf area via cell division and expansion (remote sensing/ signalling); r regulation of leaf area by abscission/reduced initiation; r functional reallocation of resources to root growth; r uptake and/or mobilisation and/or synthesis of osmotic solutes (osmotic adjustment); r reduction in root axial hydraulic conductance. Medium-term, long-term consequences, occasionally reversible: r regulation of indeterminate growth; r induction of CAM; r increased succulence; r changes in leaf type or characteristics. Constitutive adaptations, long-term to permanent consequences, frequently linked to slow growth rate, generally irreversible:

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r r r r r r r

317

deep roots (genetically controlled resource allocation); CAM or C 4 photosynthetic pathways; small dissected leaves; waxy, pubescent, silvery leaves; reduced/minimal leaf area; succulence/water storage; desiccation tolerance.

Within any category, it is often difficult or impossible to distinguish whether a response indicates particular sensitivity to drought or a highly responsive tolerance mechanism (Price et al., 2002), though the consequences of this interpretation have considerable implication in plant breeding programmes. Common themes around stress tolerance appear in the above lists. All levels of organisation, from the ion channels that mediate stomatal opening, to whole-plant anatomy, can be affected by, and can be adaptive to, water deficit. Almost everything probably interacts with almost everything else, and picking out ‘one key issue’ is fraught with danger. It may be relatively straightforward to specify what will be needed to allow a plant to survive in Death Valley. It is difficult to generalise as to what will maximise growth in water-limited conditions. It is even more difficult to generalise what will optimise the yield of a crop, in which practically any adaptation, except possibly C 4 photosynthesis, will carry a cost in terms of yield, in some, if not all, of the scenarios that year-to-year variance will throw at it. The practical ‘Catch-22’ is that there is a well-established dependency of economic yield (as a component of biomass production) upon the volume of water transpired during the growing season (Passioura, 1983), and so it is often argued that to maintain yield potential it is necessary to maintain transpiration under stress (Blum, 1989). At the least, maximising yield is linked to utilising all the water that is available. The agricultural agenda is therefore one of brinkmanship, taking water use to the limits. The ecological agenda may be more ‘cautious’, keeping some distance from the stage III edge, valuing the successful completion of the life cycle in bad years over productivity in good years. The needs of the subsistence farmer may be based on another risk analysis and fall somewhere between these two. A net increase in the quantity of osmotically active solutes (osmotic adjustment) allows water uptake from soils of lower water potential, and this may be assisted if changes in the cell walls allow expansion to occur with less hydrostatic (turgor) pressure. Osmotic adjustment is commonly reported and is achieved with inorganic ions, organic anions and neutral organic solutes. It follows that resources so committed cannot be used directly for yield in grain and seed crops, and in all cases the commitment of resources to adjust the whole cell with organic solutes is very great. The commitment is less in the case of compatible solutes that are localised in the cytoplasm. Here the nature and effective compartmentalisation of the solute are important and the absolute quantity of solute will be small if the necessary condition of effective compartmentalisation is satisfied. The two major adaptations of photosynthesis result in considerable gain in water-use efficiency and reduction in water loss. Both are mediated by shuttling fixed carbon, stored as an organic acid,

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either between different parts of the mesophyll (C 4 ) or in and out of storage (CAM). Storage of organic acid is the characteristic feature of the solute relations of CAM plants. Water-use efficiency and osmotic adjustment are responses commonly pursued for agricultural aims. Both can be thought of as related to the concept of ‘keeping going’, of maintaining transpiration and photosynthesis under stress and thus offering the potential to help sustain productivity. Biochemical mechanisms are particularly attractive because of their relative ease of study and manipulation. However, water-use efficiency may not help if it is not linked with water exploitation and the agricultural advantage of osmotic adjustment has been challenged. Fewer studies have concerned rooting properties, which are difficult to study and to manipulate, but are nonetheless at the forefront of increasing water gain.

13.3 13.3.1

Mechanisms to reduce water loss: regulation of stomata and regulation of leaf area Stomatal regulation

It has been recognised since early split-root system experiments (experiments in which the roots of a plant are divided between two containers having different availability of water) that leaf growth can be regulated by long-distance chemical signals from the roots. Drying only a part of the root system induced stomatal closure without any change in leaf water status (Blackman and Davies, 1985; Davies and Zhang, 1991). Cutting off roots that were in drying soil (an intervention that could not increase water supply but could remove chemical signals about water deficit) overrode the inhibition of leaf expansion in apple (Gowing et al., 1990). Abscisic acid (ABA) has frequently been identified as such a signal. Ethylene can also mediate effects upon leaf elongation (Sobeih et al., 2004). ABA-deficient mutants have been produced from arabidopsis, tobacco, tomato and maize. These mutants grow fairly normally in the absence of water stress and temperature stress but wilt and die under drought stress and grow poorly under salinity stress. ABA acts as a signal in the immediate response through guard cell regulation, and, in the longer term, by induction of proteins that are concerned with the tolerance of dehydration (Zhu, 2002). ABA accumulates upon osmotic stress as a result of both activation of its synthesis and inhibition of its degradation. The biosynthetic sequence for ABA goes from zeaxanthin to violaxanthin to neoxanthin to xanthoxanthin to ABA-aldehyde to ABA. Osmotic stress appears to regulate this process at several steps, though the rate-limiting enzyme is thought to be 9-cis-epoxycarotenoid dioxygenase (NECD) (Koornneef et al., 1998), which mediates the conversion of neoxanthin to xanthoxanthin. The degradation of ABA has not yet been so well characterised (Zhu, 2002) as the steps involved in its synthesis. Signalling between osmotic stress perception and ABA biosynthesis is presumed to involve protein phosphorylation cascades and Ca2+ signalling (Zhu, 2002). For ABA effects on guard cells, evidence has been presented (Levchenko et al.,

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2005) for direct activation of a cytosolic receptor: ABA rapidly activated anion channels but did not induce a rise in the background Ca2+ needed for anion channel activation, and the effect on the anion channels was not mimicked by a number of putative ABA-signalling intermediates. The mode of action of ABA on guard cells is thought to be mediated by changes in cytosolic free Ca2+ , which may arise from net influx across the plasma membrane or by release from intracellular storage (Hamilton et al., 2000). The pathways of internal release depend upon either phospholipase C or cyclic ADP-ribose (MacRobbie, 2000). Measurements with guard cell plasma membrane revealed a Ca2+ -selective channel whose open probability was dramatically increased by ABA and a close physical coupling between ABA perception and channel control at the plasma membrane (Hamilton et al., 2000). Stomatal closure depends upon the efflux of K+ from the vacuole, which is the step affected by Ca2+ signalling. Studies using inhibitors of these two pathways indicated that at high concentrations of ABA, vacuolar K+ efflux was dependent upon Ca2+ influx at the plasma membrane, while at lower ABA concentrations K+ efflux was dependent upon both available pathways of Ca2+ release from internal stores (MacRobbie, 2000), and this may represent different tonoplast K+ channels being influenced by different Ca2+ (and ABA and so stress) levels. All three routes to increasing cytosolic Ca2+ are activated by ABA. There is further discussion of ion transport in stomatal guard cells in Section 6.2.1. The ABA concentration in leaves is a result of synthesis, breakdown, redistribution and transport from the roots. Bulk leaf ABA concentration increases with water stress, and there is a good inverse correlation between stomatal aperture and concentration of ABA (Tardieu et al., 1996). The short-term effects of ABA (hours to days) on stomatal conductance are reversible and do not permanently affect stomatal function (Franks and Farquhar, 2001). For example, accumulation of ABA was strongly associated with rapid stomatal closure and reduction of leaf area growth in cassava at an early stage of water stress and this was also rapidly reversible (Alves and Setter, 2000). These rapid effects are mediated by direct manipulation of guard cell turgor. Longer term effects, which are not reversible, can also take place if water stress is prolonged. Apart from these rapid adjustments to fluctuations in water status taking place during the day or within a few days, there are further changes that take place when water stress means that the concentration of ABA remains elevated for longer periods of time. A distinction is made between short-term physiological and long-term anatomical causes of plant reaction to water stress (Spence et al., 1986). Treatment of Tradescantia virginiana plants with ABA led to their having smaller stomata at a higher unit area density and showing a greater degree of closure at the same guard cell turgor pressure than did controls without ABA treatment (Franks and Farquhar, 2001). Plants grown with ABA treatment had lower maximal stomatal conductance and operated at a lower stomatal conductance for a given guard cell turgor pressure. They also had lower C i (concentration of carbon dioxide in the air spaces inside the leaf) at a given atmospheric concentration of carbon dioxide (C a ), though photosynthetic capacity (defined as assimilation rate at a given C i ) was not reduced by treatment with ABA. It was proposed (Franks and Farquhar, 2001)

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that these changes in stomatal properties would produce an increase in water-use efficiency (because they were operating at a lower C i /C a ) under prolonged exposure to water deficit. Leaves from ABA-treated plants were ‘noticeably smaller’ (Franks and Farquhar, 2001), which could mean that (a) water status of the plant would also be improved by reduced total transpirational demand and (b) despite increased water-use efficiency there was still reduced growth.

13.3.2

Leaf area regulation

Reduction in leaf area is a line of defence in coping with limitation of water supply, in terms of both passive consequence and long-term adaptation. Lack of availability of water will tend to reduce turgor pressure and this will almost instantaneously reduce leaf expansion, since: Expansion = ([m] extensibility) multiplied by ([P] turgor pressure −[Y ] yield threshold) A modest osmotic shock to the roots results in a rapid and transient decrease in growth (measured as leaf elongation in monocotyledonous plants) followed by (partial) recovery. The rapidity of induction and recovery and independence of whether the shock is applied by inorganic ions or impermeant osmotica imply that these responses are entirely due to changes in cell water relations (Yeo et al., 1991; Munns, 2002). The ‘immediate’ responses, mediated by water relations alone, are distinguished from longer term responses in which root–shoot signalling may be implicated. Signalling can override recovery if changes are made to m and Y. Unlike changes in P, changes in m and Y can be adaptive, rather than a passive consequence, and can be longer term, rather than rapidly reversible. As well as by reduction in expansion rate and final area of individual leaves (due to cell size or cell number) reduction in total leaf area can be added to by reduction in numbers of leaves (by reduced rate of initiation and/or by increased rate of abscission). There is an aggregate loss of leaf area × time as a consequence of size, duration and delay. Leaf area expansion is controlled by both cell expansion and cell division, and water deficit affects these independently (Granier and Tardieu, 1999). In cassava, reduction in cell expansion due to mild water deficit was largely reversible on rewatering, and the loss of leaf area was due mostly to reduced cell division in leaves that were meristematic at the time of water deficit, and to developmental delay (Alves and Setter, 2004). Short-term modulation of leaf expansion can be achieved if the ABA signal is responsive to changes in the environment, and such modulation will suffice for plants in situations where water deficit is transient and in experimental systems following this pattern. Regions with ‘significant but erratic summer rainfall’ (Taiz and Zeiger, 1991) favour plants with indeterminate growth patterns allowing a flexible approach to varying conditions. Where there is a ‘predictable’ pattern of drought, there is an ecological value in pre-emptive water conservation. The progressive development of water deficit

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commonly characterises arid zones subject to only seasonal rainfall or seasonal flooding. When stress develops slowly, developmental processes can respond, and this usually includes a reduction in leaf area, increasing the probability that the plant will complete its life cycle before it depletes the available water. This may be facultative in character, leaf development and expansion being regulated by environmental signals in dry seasons, or constitutive in character, for species whose adaptations include having reduced, or vestigial, leaves. Extremely drought-tolerant species may abscise all their leaves and regrow them when water becomes available again.

13.3.3

Consequences: interaction with leaf temperature

(Partial) closure of the stomata will automatically reduce photosynthesis in the short term, will reduce transpirational cooling and is likely to make the leaf vulnerable to oxidation damage. Leaves in the sun gain heat from solar radiation. The heat gain of the leaf is offset by radiation, by sensible heat loss and by transpirational cooling. In broad leaves with adequate water supply, transpirational cooling predominates. Transpirational cooling depends upon the hydrogen bonding of water, which accounts for its large latent heat of vaporisation (see Section 3.2). Hydrogen bonding also gives water a comparatively large thermal capacity (a relatively large amount of energy is needed to raise its temperature). Despite this buffering, plants in high-irradiance environments with high air temperatures may still operate at acceptable leaf temperatures, but with little margin for error. Such plants will require constitutive adaptations to decrease the thermal load on the leaf and to aid cooling by radiation, conduction and convection. In hot, high-irradiance climates short exposures to damaging temperatures can be serious or lethal. Small and dissected leaves maximise boundary conductance because there is a greater ratio of edges to surface area and a greater amount of air turbulence than over the surface of larger, flatter leaves. The possession of small and dissected leaves is frequently combined with increasing the reflectivity of the leaves and reducing total leaf area (both to conserve water and to limit thermal load). Thick epicuticular wax is a shoot-related trait for which significant genetic variation has been reported (in upland rice; O’Toole and Cruz, 1983). In the short term, leaf rolling and leaf movements (responsive alignment and passive wilting) both reversibly (unless wilting leads to mechanical damage) reduce either effective leaf area or the angle of exposure to solar radiation and thus the heat load. Apart from these shortand long-term adjustments of leaf shape and form, stomata clearly play a critical role in the fine balance between carbon gain and water loss. This must take account of the critical role of transpiration in preventing lethal overheating as well as the need for carbon fixation. Another potential consequence of stomatal closure is oxidative damage. Reactive oxygen species are considered to be the cause of photoinhibition (Asada, 1999; Takahashi et al., 2002). The prospects of manipulating photosynthesis to improve crop performance in drought conditions are discussed by Horton (Horton, 2000). Stomatal regulation provides the short-term tuning that a temperate species needs in

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order to deal with the short-term fluctuations in the environment. Growth regulation offers a longer term adjustment to water supply. In unusually demanding seasons there will be more drastic consequences: annuals will flower prematurely and die, and trees will drop their leaves. In climates where such seasons are frequent the flora will shift to those with constitutive adaptations.

13.4 13.4.1

Mechanisms to maintain water potential gradients: osmotic adjustment Water potential of drying soil

As the soil dries, water retreats into interstices between soil particles. Mutual attraction between water molecules underlies the phenomenon of surface tension (or surface free energy; see Section 3.2). It is against these ‘matric’ forces, essentially the negative hydrostatic pressure of small menisci, that roots must compete for water as the soil dries. The water potential of the plant must be lower than that of the soil if the plant is to take up water. The hydraulic conductance of the soil also decreases as its water content decreases, as air replaces water in the spaces between the soil particles limiting the cross-sectional area available for mass flow. When the water potential of the soil is so low that the plant cannot maintain its water status on a 24-h aggregate, this is described as the permanent wilting point (wilting during daylight can no longer be recovered at night). Three stages can be distinguished in the response of plant transpiration to soil water deficit (Sinclair and Ludlow, 1986). Stage I is when water availability is sufficient for stomatal conductance to be maximal and thus transpiration to be dependent solely upon the atmospheric environment. When water availability is insufficient to support potential evapotranspiration, stomatal conductance is used to match transpiration to water uptake and maintain plant water balance: this is stage II. If water supply falls further, stomatal compensation has run out, and survival mechanisms take over (stage III). Processes correlated with crop yield are inhibited during stage II at the latest, and what happens in stage III drought has little if any effect upon yield, even in subsistence agriculture (Serraj and Sinclair, 2002). What happens in the ‘survival’ range is, however, crucial to those plants whose ecology includes these conditions. Plants adapted to these conditions may limit their potential transpiration from the outset and so never be capable of being highly productive (see Section 13.3.2). Plants can lower their water potential by increasing the concentration of osmotically active solutes or by decreasing turgor pressure; the latter will decrease expansion growth unless the yield threshold is also reduced. Large increases in elastic coefficients were seen in water-stressed white spruce allowing turgor to be maintained at water contents corresponding to turgor loss in well-watered controls (Marshall and Dumbroff, 1999). Corresponding changes in osmotic potential at full turgor and bulk tissue elastic modulus, in water-stressed cassava, meant that turgor loss occurred at the same relative water content in both stressed and unstressed leaves (Clifford et al., 1998).

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13.4.2

323

Osmotic adjustment

The term osmotic adjustment is used to imply an adaptive response: a net increase in the quantity of osmotically active solutes either by uptake or by synthesis. This must be over and above the consequential increase in concentration that will occur simply as a result of reduced hydration. There is the added factor that some increase in organic solutes may result from decreased consumption as a consequence of water deficit, rather than being an adaptive response to water deficit (Munns, 1988). Substantial genetic variation has been reported for osmotic adjustment in major crop species (Table 1 in Zhang et al., 1999) with a range from 0.1 to 1.7 Mpa. The upper limits are for genotypes within rice (1.7 Mpa), sorghum (1.7 Mpa), wheat (1.4 Mpa) and pigeon pea (1.3 Mpa). Osmotic adjustment can lower the plant water potential at which water movement into the plant can be maintained. Hence it is argued that since positive turgor is required for stomatal opening and expansion growth, osmotic adjustment is a mechanism that is of value to crop plants (Zhang et al., 1999). Even before the current wave of interest in manipulating solute accumulation in crop species, the value of osmotic adjustment was questioned (Munns, 1988). It has recently been questioned again by Serraj and Sinclair (2002) who asked whether there is any convincing evidence that osmotic adjustment increases crop yield and, furthermore, whether there is any theoretical expectation that it should. These authors (Serraj and Sinclair, 2002) conclude that a positive yield response has only rarely been demonstrated and then only in conditions of such stress that the yields obtained were extremely low and ‘irrelevant for most agricultural conditions’. Figures for experimental yields of wheat (Morgan, 1983) were equivalent to 0.44 and 0.29 tonnes ha−1 for high and low osmotically adjusting lines, respectively. There may be situations in subsistence farming where such yields and yield differences do matter, but this is a very different scenario from general applicability to dryland farming. Challenges to a theoretical expectation of a positive effect of osmotic adjustment on crop yield (Serraj and Sinclair, 2002) focus upon the conclusion that by the time osmotic adjustment comes into play, soil water has largely been depleted and thus: 1. it is already stage III drought and so is mostly irrelevant to agricultural contexts; 2. maintaining turgor and water use at this point, when water-conserving mechanisms ought to have priority, may be counterproductive; 3. there is not enough water left to extract from the soil to make much difference to yield. None of this detracts from the relevance of osmotic adjustment as an adaptation for plants that inhabit arid regions; it is whether increasing osmotic adjustment in crops has any value that is being questioned. Osmotic adjustment can be achieved with inorganic ions that are readily available in the soil, such as potassium, nitrate, chloride and sulphate. The importance of the uptake of inorganic ions in regulating turgor in roots exposed to osmotic stress is demonstrated in concomitant measurements of turgor pressure and changes in

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ion activity (Shabala and Lew, 2002). Immediate decrease in turgor was observed upon hyperosmotic stress, with recovery initiated within minutes and accompanied by increases in inorganic ion uptake leading to substantial recovery of turgor: voltage-gated plasma membrane potassium transporters were at least partially implicated (Shabala and Lew, 2002). Osmotic adjustment can be achieved with the synthesis of organic solutes. These may be neutral, such as sugars, or, for example, organic anions to balance potassium. Since osmotic adjustment will often require that the concentration of ions in the cell be in the inhibitory range for processes in the cytoplasm, compartmentalisation of ions may be necessary in order for osmotic adjustment to be effective. Ions can be used to adjust the bulk of the aqueous phase in the vacuole, and other solutes, which do not adversely affect protein integrity, could be localised in the cytoplasm. The latter are collectively known as ‘compatible solutes’, ‘osmolytes ‘or ‘osmoprotectants’, the last strictly implying that their presence has positive rather than merely neutral effect upon macromolecules.

13.4.3

Compatible solutes/osmolytes/osmoprotectants

When solutes dissolve in water all have effects known as colligative properties (properties related to the number of molecules or ions in solution; see Section 3.3.3), which affect, for example, the water potential, freezing point and boiling point of the solution. Solutes can also affect the solvent properties of water, and if they do so are known as cosolvents (see Section 3.2); because of this, certain solutes perform protective roles in dehydration resistance, and in dehydration tolerance, which are disproportional to their colligative properties. A number of ‘compatible solutes’, some with ‘osmoprotectant properties’, have been described in plants and other organisms. Glycine betaine and proline have received the most attention, but some other examples are glycerol, sucrose, ectoine, trehalose, glucosylglycerol and 2-sulphotrehalose (Oren, 1999). Work largely conducted with halophytes in relation to salinity (Chapter 14) points to an osmoregulatory role in the cytoplasm where, because of the small volume fraction, small absolute quantities of compatible solutes can have substantial osmotic effect. In halophytes there is evidence that glycine betaine is localised in the cytoplasm. ‘Compatible solutes’ have received much attention with regard to osmotic adjustment, perhaps because there is the potential to introduce their production by relatively simple genetic interventions. Considerable effort has been made to introduce or enhance the production of compatible solutes through transgenics (Cushman and Bohnert, 2000). However, the practical outcomes of extensive studies have been only marginal. Shabala and Lew (2002) report, ‘To our knowledge, there are no reports of any significant improvements in drought tolerance of any crop species in field trials. This is probably due to the complexity of whole-plant responses to water stress’. An important consideration in understanding the role of compatible solutes in vacuolate cells is where the solutes are distributed within the cell. Examining reports of the overexpression of osmolytes in transgenic plants, Zhang et al. (1999) calculate substantial concentrations (and osmotic potentials) in the cytoplasm in a

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number of these cases (apparently assuming complete cytoplasmic compartmentalisation and a cytoplasmic volume fraction of 0.05). However, Zhang et al. (1999) cite the data of Ishitani (1995) as an example that there was no effect on the overall osmotic potential despite the biosynthesis of proline being equivalent to 600 mM or –1.48 Mpa (presumably assuming cytoplasmic localisation). That there was no (measured) osmotic adjustment can be explained if the proline was, in fact, not compartmentalised, in which case the average change in concentration might have been only 30 mM and thus difficult to detect in terms of the value and variance of osmotic potential. The implication is that in the cases considered, the synthesis of the osmolyte was (a) not compartmentalised and so served little purpose and (b) not accompanied by the accumulation of inorganic ions (or anything else) in the vacuole, and so was not associated with osmotic adjustment. Where osmolyte synthesis is linked with osmotic adjustment (which is not the same as their role in dehydration tolerance), synthesis must be linked with osmotic adjustment of the cell as a whole. Osmolytes may well retain protective and other roles that can be achieved at lower concentrations, and with lesser quantities, if the condition of compartmentalisation is not met, but they will not osmotically adjust the cytoplasm and probably never contribute appreciably to the overall osmotic adjustment of vacuolate cells. Reports of increases in concentration of solutes, including proline, are, nonetheless, frequent in plants exposed to water deficit (Clifford et al., 1998). In many cases the role of proline is unproven. However, an unequivocal role of the accumulation of compatible solutes in appropriate situations is indicated by the accumulation of proline in the pollen of tomato plants during their dehydration phase. Proline content was 60-fold greater in flowers than in any other tissue and was confined predominantly to the pollen, where it represented over 70% of free amino acids: uptake was mediated by a transporter, LeProT1 (Schwacke et al., 1999). Plants synthesise glycine betaine from choline via betaine aldehyde (Hanson and Rhodes, 1983) utilising the enzymes choline mono-oxygenase and betaine aldehyde dehydrogenase. Salinity and water deficit are associated with an increase in betaine aldehyde dehydrogenase mRNA and protein in leaves and in roots and the concomitant accumulation of glycine betaine. Most attention has been on this enzyme. Water deficit was also associated with increased choline mono-oxygenase mRNA and protein and an increase in enzymatic activity, all of which were reversed upon rewatering (Russell et al., 1998). Both enzymes are thus inducibly expressed in roots and leaves within the Chenopodiaceae (Russell et al., 1998). Transgenic plants were made in arabidopsis, Brassica napus and tobacco, using a bacterial choline oxidase that converts choline to betaine aldehyde and, also though inefficiently, betaine aldehyde to betaine (Huang et al., 2000), and these produced only small quantities of betaine unless they were substantially supplemented with choline. This would suggest that choline supply also needed to be increased to yield physiologically significant amounts of betaine (Huang et al., 2000); however, consideration needs to be given to the kinetics of conversion of betaine aldehyde to betaine by the bacterial enzyme in contrast to the two-enzyme system present in native betaine-producing plants.

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13.4.4

PLANT SOLUTE TRANSPORT

Water movement from protoplast to apoplast in freezing injury

As the temperature falls below zero, cell contents first supercool (Taiz and Zeiger, 1991). Spontaneous ice nucleation in pure water does not occur until −38.1◦ C, but crystallisation is facilitated by nucleation sites, dust particles and impurities. But the cell contents are essentially ultrafiltrated via membrane uptake and, in frost-hardy plants, are capable of supercooling. Sometimes, supercooling may occur by only a few degrees below zero, but florets of Rhododendron japonicum remained unfrozen at −14 to −21◦ C, and the xylem of Acer japonicum remained unfrozen at −21◦ C (Ishikawa et al., 2000). In the shorter term, freezing of water on the leaf surface raises the leaf temperature due to the latent heat of fusion of ice. In the longer term, ice crystallisation extends into the apoplast and it is the low water potential of ice in the apoplast (extracellular ice) that dehydrates the protoplast, causing the same types of damage seen in heat or desiccation, and leading ultimately to ice nucleation within the protoplast and thus to intracellular freezing. Because of the relative slowness of such freezing, ice crystals are large on the scale of cellular organisation and cause irreparable mechanical damage. Delay of freezing damage involves many similarities with delay of desiccation damage. Protective solutes perform similar roles, and in addition there are a number of molecules that are thought to act as ‘antifreeze’. The dual aims are to prevent the formation of intracellular ice and to counter the desiccation effects of dehydration by extracellular ice. Tolerance of low temperatures (chilling or freezing) has, like desiccation tolerance (Chapter 15), a long timescale, in which changes in proteins and lipids are needed to confer stability at lower temperatures.

13.5

Mechanisms to acquire more water: root properties

It has been demonstrated that deep rooting does result in an increased ability to extract soil water from depth in the soil profile, improve plant water status and may increase yield under drought (Yoshida and Hasegawa, 1982; Mambani and Lal, 1983), and there appears to be convincing evidence that deeper, thick roots should contribute to drought resistance in at least some environments (Price et al., 2002). Many mechanisms are available that alter or maintain root growth under water deficit. In addition to osmotic adjustment to maintain turgor and wall loosening to allow expansion at lower turgor, there are fine changes within the root growth zone as well as effects of ABA (Sharp et al., 2004). In the apical 3 mm of the primary maize root, elongation is maintained under water deficit in roots that have sufficient concentration of ABA. In the next 3–7 mm, elongation is progressively inhibited by water deficit (Sharp et al., 2004).

13.5.1

Constitutive formation of deep roots

A constitutive genetic predisposition to allocating large resources to deep roots involves some advantage from that commitment. The resources are put into root growth on the ‘expectations’ that (1) there will be unexploited water at depth in the

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soil and (2) the competitive advantage of accessing this water will outweigh the cost in terms of above-ground productivity. Dry summers following (usually) regular winter rains and early-season-only irrigation provide situations where commitment to exploiting deep soil is likely to be advantageous in the agricultural context. Having a deep, thick root system allowing exploitation of water from deep down the soil profile is seen as a crucial character in drought resistance of upland rice, even though the cost of carbon allocation to roots rather than shoots is clearly recognised, and is a character for which there is substantial genetic variation within the genome (Price et al., 2002).

13.5.2

Facultative formation of deep roots

There is a functional balance underlying root/shoot ratio (Taiz and Zeiger, 1991). Put plainly, leaves can grow only as much as the water that roots provide will permit, and roots can grow and access only as much water as the photosynthate provided by the leaves will permit. Source/sink relationships can operate at this ‘supplyand-demand’ level. This can be overridden by genetic differences controlling the allocation of resources to roots in constitutively deep-rooted species and genotypes. It can also be overridden by (hormonal) regulation of leaf area to ‘conserve’ soil water supplies. The importance of root growth to species that inhabit predictably drought-prone environments is clear enough. Interpretation is more problematic for agricultural crops, where the compromise between maximising yield in good years and sustaining yield in bad years is a difficult call – whether or not to have a genetic commitment of more resources to roots. For upland rice it is noted that quantitative trait loci (QTLs) associated with root growth have so far made little contribution to drought avoidance (Price et al., 2002) despite the expectation that deeper, thicker roots should be of benefit in some environments. The problem for plant breeders is that in any cross they make there is a range of root and shoot traits, which may or may not be linked physiologically and/or genetically, either positively or negatively, all with relatively small and possibly incidental effects upon drought tolerance. The great hope is to pull out QTLs linked to agricultural merit from all of this. It is a hope, and not necessarily an expectation, that natural selection has provided some coordination whereby at least some of these traits are all pointing in the same direction. To what extent such ‘natural’ variation remains, following the intense human selection upon agricultural species since Neolithic times, is open to debate. There are many competing demands for resources, which are exacerbated by water deficit: left to a functional resolution the outcome may be variable and not optimised for yield; genetic pre-determination of the response can optimise the plant for water-limited conditions, probably at the expense of performance in more ideal conditions.

13.5.3

Root conductance

While the acquisition of water by roots depends on the ability of those roots to explore the soil profile for regions with obtainable water, the movement of water

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through roots is a function of their hydraulic conductance. This conductance depends upon the conductance of the root cortex and of its xylem. The hydraulic conductance of roots can be a limiting factor since, overall, the conductance of the root might be about half that of the stem and two-thirds that of the leaves (Nobel, 2005). There is evidence that root hydraulic conductance is under genetic control, as it has increased with ploidy level over the evolution and development of modern types of wheat (Zhao et al., 2005). An increase in ABA concentration and direct stimuli from drought and salinity have been observed to regulate expression of both tonoplast and plasma membrane aquaporins (Maurel et al., 2002). A-proteins from Mesembryanthemum crystallinum (proteins that mediated a flux of water when expressed in oocytes of Xenopus laevis) showed membrane redistribution in response to mannitol-induced water stress, and this redistribution was prevented by inhibitors of vesicle trafficking (Vera-Estrella et al., 2004). Arabidopsis plants expressing antisense for both PIP1 and PIP2 plasma membrane aquaporins exhibited reduced quantities of both of these proteins and substantially reduced osmotic hydraulic conductance in isolated protoplasts. The plants had a threefold reduction in unit mass root hydraulic conductance, which was compensated by a nearly equivalent decrease in shoot/root ratio (Martre et al., 2002). The hydraulic conductance of the whole plant was similar in normal and doubleantisense plants, which might suggest that the lesions introduced by antisensing the aquaporins were compensated for by a functional balance. An important feature of the transport of water from drying soil into roots is the development of an air gap between roots and soil, as both roots and soil shrink. This can increase the resistance to water flow and decrease water exchange (both uptake and loss) between the root and the soil (Nobel, 2005). Direct pressure-probe measurements have been made in the xylem in recent years to complement the more established measurements made with the Scholander bomb. There have been conflicting views of how these measurements support or question the cohesion-tension theory. Much of the debate revolves around whether the pressure bomb correctly estimates the pressure in the xylem (pressure bomb estimates can be much more negative than are direct measurements). Direct measurements of xylem pressure with a pressure probe in maize leaves were held to be consistent with the cohesion-tension theory (Wei et al., 1999). There have been many contentions that transpiration cohesion is not the only mechanism involved in upward movement of water in the xylem (and this is discussed in Section 9.2.6).

13.6

Mechanisms to increase water-use efficiency: C 4 and crassulacean acid metabolism (CAM)

Put very simply, productivity is water use times water-use efficiency, and agricultural yield is productivity times the harvest index. Yield is broadly proportional to water transpired, so increasing water-use efficiency per se is not an agronomic end in its own right, especially if it interferes with the maximum utilisation of available water (Jones, 1993). Nevertheless, water-use efficiency has received much

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attention because it is a feature that has the capability to carry little yield penalty. Uptake of carbon dioxide and loss of water occur along a common pathway via the stomata. The uptake of carbon dioxide is driven by the concentration difference between the air and the air spaces within the leaf. This concentration difference is maintained by the rate of net photosynthesis (assimilation minus mitochondrial respiration and photorespiratory losses). The absolute concentration of carbon dioxide in the atmosphere is small so this concentration difference is also small. In contrast, the concentration difference for water vapour, in the inside-to-outside direction, is large. Water vapour is at or near to saturation in the leaf air spaces and in almost all cases well below this in the surrounding air. The boundary layer offers some protection against steep gradients of water concentration but this is stripped away as air movement increases. Not only is the concentration difference large, but the absolute concentration of water vapour (near saturation in the leaf) is large, meaning that fluxes of water vapour out of the leaf are also large. If a plant is to grow it must assimilate carbon dioxide, which means that stomata must be open. If stomata are open then a large net flux of water vapour out of the leaf will occur, which will be greatest at high temperatures, at low atmospheric relative humidity and at high windspeed. In this narrow sense, plant water-use efficiency is concerned with gas exchange and with the relationship between carbon gained and water lost. On a crop basis, it includes manipulation of harvest index to partition more of the productivity into economic yield, and on a field basis it includes maximising the use of available water. This means utilising the maximal fraction of water available and minimising losses through evaporation from the soil, deep drainage and water unused at harvest (Condon et al., 2004). Although it appears intuitive that improvements in leaf wateruse efficiency would be beneficial in water-limited growth, there is no clear link between this and crop water-use efficiency or yield (Condon et al., 2004). Again it appears that focus on one particular trait, no matter how intuitively obvious its importance may seem, fails to be a reliable indicator of the path to improvement, certainly where economic yield is the objective. Some of the problems lie in the sensitivity of reproduction to water deficit and the prominence of successful reproduction in determining economic yield. Inhibition of photosynthesis, directly by leaf water deficit or indirectly via ABA, decreases supply of photosynthate to developing organs and triggers ovary abortion in maize (Boyer and Westgate, 2004), which can be prevented by feeding sucrose to stems. Pollen sterility is also a common feature of drought. Water deficit experienced late in the life cycle can therefore drastically reduce economic yield, even if the plant has thus far grown well. Compared with variation of water-use efficiency within C 3 species, much greater increases in water-use efficiency are achieved in C 4 photosynthesis and in CAM.

13.6.1

C 4 photosynthesis

Net carbon fixation is mediated by Rubisco, which has both carboxylase and oxygenase activity catalysed at the same active site, so there is both a carbon fixation

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cycle (the Calvin cycle) and photosynthetic carbon oxidation cycle. The oxygenase activity results in photorespiration, although the pathway responsible for the CO 2 release actually succeeds in recovering three quarters of the carbon lost through the oxygenase activity. The oxygenase activity is widely regarded as an evolutionary leftover, in the sense that Rubisco evolved in conditions where the CO 2 /O 2 ratio was so high that the oxygenase activity was irrelevant; there was ‘no need’ for better discrimination between the two substrates. Differences in specificity between Rubisco from different sources do exist (Yu et al., 2005), the specificity in cyanobacteria being lower than in marine non-green algae. The earliest marine (cyanobacterial) O 2 evolvers date back 3.45 Gy (Raven, 1997). By the time that atmospheric CO 2 was depleted towards current concentrations, Rubisco already dominated the scene. Only in certain conditions does there seem to have been enough selection pressure to modify this scenario, and this has been achieved by methods that alter the CO 2 /O 2 ratio presented to Rubisco, rather than by any fundamental change to Rubisco itself. Such conditions have been in situations that involve severe CO 2 depletion (very high canopy photosynthesis), high temperatures (increasing photorespiration) and where water-use efficiency has sufficient importance to have evolutionary consequences. C 4 photosynthesis and CAM (and some aquatic algae) operate a CO 2 concentrating system. At standard conditions, the partial pressures of CO 2 and O 2 in the atmosphere are in the ratio of about 1:600. The biochemical reactions of the photosynthetic pathways occur in aqueous solution and the concentrations of the gases in solution are determined by Henry’s law: the concentration in solution is proportional to the partial pressure above the solution and the Bunsen coefficient. Of great importance to the plant is that the Bunsen coefficient is much greater for CO 2 than it is for O 2 at the same temperature. Thus, while the ratio of partial pressures in the atmosphere is about 1:600, the ratio of concentrations in solution is only 1:22.6 at 25◦ C. The solubility of both gases decreases as the temperature increases, but the solubility of CO 2 falls off more steeply. As the leaf temperature increases the ratio of CO 2 to O 2 decreases and this acts in favour of the oxygenase activity of Rubisco, making photorespiration of increasing significance as the temperature rises. In C 4 photosynthesis the initial carbon assimilation step is via phosphoenolpyruvate carboxylase (PEPCase) rather than Rubisco. Although PEPCase activity does not in itself result in the net fixation of carbon, the malate or aspartate that is formed is transported to the site of Rubisco and broken down to release carbon dioxide. The spatial separation of carbon fixation and the Calvin cycle allows a ‘carbon dioxide pump’ to operate, raising the concentration of carbon dioxide presented to Rubisco, inhibiting its oxygenase activity and suppressing photorespiration. The affinity of PEPCase for CO 2 is greater than that of Rubisco, and this enables the internal concentration of carbon dioxide (C i ) to be depleted to a lower concentration in C 4 plants than in C 3 plants, which has the effect of increasing the concentration difference from C a to C i and thus facilitates the diffusion of CO 2 into the leaf. This increases the uptake of CO 2 for a given stomatal aperture and contributes to an increase in water-use efficiency. There is a clear anatomical differentiation that characterises C 4 plants with two distinct types of chloroplast-containing cells, the mesophyll and the bundle sheath.

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Variants of C4 Photosynthesis

C4 variant

C4 acid transported from mesophyll to bundle sheath

NADP-ME

Malate

NAD-ME

Aspartate

PEP-CK

Aspartate

Decarboxylation system Chloroplastic NADP-dependent malic enzyme Mitochondrial NAD-dependent malic enzyme Cytosolic phosphoenolpyruvate carboxykinase

C3 compound transported from bundle sheath back to mesophyll Pyruvate

Alanine

Alanine and pyruvate

CO 2 is shuttled from the atmosphere to the bundle sheath cells producing a higher concentration of CO 2 than could be achieved through equilibration of the leaf interior with the atmosphere. The key role of solutes is in shuttling the CO 2 fixed by PEPCase in the mesophyll to the bundle sheath (as a C 4 acid, malate or aspartate), and shuttling a C 3 acid (pyruvate or alanine – see below) back to the mesophyll from which the CO 2 acceptor (PEP) is regenerated. The latter varies with the three variants of C 4 photosynthesis (Table 13.1). NADP malic enzyme (NADP-ME) is the simplest case. In the mesophyll, PEP is carboxylated to oxaloacetate by PEPCase, is reduced to malate by NADP malate dehydrogenase, and transported to the bundle sheath. Here malate is decarboxylated by NADP-ME (choroplastic) to pyruvate, which is returned to the mesophyll. Here the pyruvate is phosphorylated by pyruvate orthophosphate dikinase to regenerate PEP to continue the cycle. The CO 2 produced by decarboxylation in the bundle sheath feeds into the C 3 photosynthetic carbon reduction (Calvin) cycle. NAD-ME and PEP carboxykinase (PEP-CK) types include additional cyclic steps in both mesophyll and bundle sheath cells involving aspartate and alanine aminotransferases. In both cases it is aspartate that is transported from mesophyll to bundle sheath where it is converted back to oxaloacetate prior to decarboxylation. In NAD-ME, alanine alone is returned to the mesophyll. In PEP-CK, the decarboxylation with PEP-CK regenerates PEP directly, which is returned, as well as alanine produced in the aspartate/alanine aminotransferase cycle.

13.6.2

CAM

Crassulacean acid metabolism provides a temporal separation of initial carbon fixation, which occurs at night via PEPCase in the cytosol (phase I), and the photosynthetic carbon reduction cycle, which occurs by day in the chloroplasts (phase III). This enables carbon to be fixed at night, when the stomata can be open with much less water loss than would occur during the day, and reassimilated to carbohydrates during the day when light is available, but the stomata are closed. There

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are stages after dawn and before dusk (phase II and phase IV) where stomata may be partially open in the light and some carbon fixation directly by the C 3 pathway can occur. CAM relies upon the intermediate storage of carbon as an organic acid, usually malate, in the vacuoles. Vacuolar compartmentalisation is needed to provide sufficient storage to fuel the Calvin cycle all day. The name ‘crassulacean acid metabolism’ reflects the earliest observation that the acidity of certain species of the Crassulaceae showed a diurnal cycle. The storage of malate presents a challenge in compartmentalisation within the vacuole: malate concentrations rise to 500 mM, which means a gradient of many pH units is maintained across the tonoplast. Malate transport, as an intermediate in the Krebs and glyoxylate cycles, has been investigated extensively. The mitochondrial dicarboxylate transporter exchanges malate for HPO 4 2− and is one of a range of transporters for di- and tricarboxylic acids situated in the inner membrane (see Section 7.3.2.2): the dicarboxylate transporter is driven by the phosphate gradient, which is in turn set up by the proton gradient (Siedow and Day, 2000). The vacuolar malate transporter (see also Section 7.6.2.1) has proven difficult to identify. Vacuolar malate transport in Kalanch¨oe daigremontiana was characterised by Ratajczak et al. (1994). The putative transporter had a K m for malate of about 3 mM and a V max that was sufficient to account for diurnal accumulation of malate. Transport activity was inhibited by citrate and was suggested to be a carboxylate uniporter with strong affinities for malate and citrate. A summary of the literature (L¨uttge et al., 2000) listed the identification of influx carriers and influx channels in a range of species (including K. daigremontiana) and an efflux channel in the CAM species Graptopetalum paraguayense; however, in no case had a definitive molecular identification been achieved (L¨uttge et al., 2000). A tonoplast dicarboxylate transporter has been described in arabidopsis (AttDT) using a candidate gene approach, considering mammalian kidney malate transporters (homologues of mammalian kidney aquaporins are found in plant membranes): this identified a single copy of a gene with 65.6% similarity and 38.4% identity to the human sodium/dicarboxylate co-transporter (Emmerlich et al., 2003). The arabidopsis gene, unlike the mammalian gene, was located on the tonoplast and was not energised by sodium, but by tonoplast proton pumps. An AttDT-knock-out showed much reduced malate transport and accumulated less malate in the leaves. However, the knock-out mutant and wild-type plants showed no phenotypic differences when grown under standard conditions, and it was recently concluded that there is at least one other carrier protein and a channel for dicarboxylate and citrate transport at the tonoplast (Hurth et al., 2005). In the earlier study, wild-type malate transport by AttDT was competitively inhibited by citrate but there was a residual citrate-insensitive activity that the authors (Emmerlich et al., 2003) speculated could correspond to the activity of a malate channel (Pantoja and Smith, 2002). Decarboxylation of the storage acid, and carbon dioxide release, behind closed stomata during the day increases the concentration of carbon dioxide in the intercellular spaces thus instigating stomatal closure (L¨uttge, 2002) and also increases CO 2 concentration around Rubisco thus reducing its oxygenase activity and, with that, photorespiration. The concentrating effect (C i /C a ) has been observed to range from 2-fold to 62.5-fold, with a median of 6.5-fold (data in L¨uttge, 2002). This

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CO 2 -concentrating effect leads to water-use efficiencies that are several times greater than either C 3 or C 4 plants under comparable conditions (Drennan and Nobel, 2000). Some CAM plants have ecological distributions that typically are very arid or have seasonal or intermittent water supply (Cushman, 2001). There are anatomical/ morphological correlates including succulence, thick cuticles and reduced stomatal frequency, which are associated with minimising water loss. As well as the more obvious water-limited conditions, a large proportion of CAM species are rainforest epiphytes: the question is posed (L¨uttge, 2002) as to whether this contradicts the view that CAM evolution is driven by water-use efficiency or whether epiphytes have equivalent problems of water supply. It is noted (Borland and Taybi, 2004) that CAM is useful to plants growing with either or both carbon and water limitation, and it is possible that forest epiphytes are also responding to daytime CO 2 depletion within the canopy. CAM is phylogenically very widespread, more so than is C 4 photosynthesis, occurring in some 7% of species of vascular plants (Winter and Smith, 1996) among 33 taxonomically diverse families. This makes it likely that CAM evolved on many independent occasions (Cushman, 2001), and this presumption is strongly supported by data using PEPCase sequences (Gehrig et al., 2001). CAM is not without adverse consequences: the CO 2 -concentrating effect in daylight is also O 2 concentrating with potential oxidative stress, and it is asked whether the shifts seen in CAM/C 3 intermediates reflect a better protection from irradiance stress in C 3 than in CAM photosynthesis (L¨uttge, 2002). CAM shows considerable plasticity, with the ‘classic’ symptoms of ‘day’ and ‘night’ separated by variable periods in which both PEPCase and Rubisco make simultaneous contributions to CO 2 assimilation (Cushman, 2001). The extreme demonstration of plasticity is seen in the C 3 –CAM intermediates in which the C 3 pathway is used when water is non-limiting but CAM is induced when water becomes limiting. M. crystallinum is a facultative CAM species showing a gradual and largely irreversible transition from C 3 to CAM following exposure to water deficit (Cushman et al., 1990) while other facultative CAM species exhibit more rapid changes that are also reversible (L¨uttge, 1996). The shift to CAM is affected by light, humidity and temperature as well as by water deficit and thus appears to be an integrated environmental response taking more than one cue. M. crystallinum has emerged as the ‘model’ because in this species CAM is induced by salinity also. CAM induction, however initiated, is controlled mostly through transcriptional activation (Cushman, 2001). CAM induction focuses on a single PEPCase transcript among the PEPCase gene family. Carbon flux through CAM is determined by diurnal oscillations in PEPCase activity, partly controlled by circadian rhythm changes in the phosphorylation state of the enzyme. The state of phosphorylation is due mostly to changes in the activity of PEP-CK, whose activity, in M. crystallinum, is induced at the same time as that of CAM-specific PEPCase (Li and Chollet, 1994). The abundance of the kinase is in turn regulated by changes in the level of its translatable mRNA in response to a circadian rhythm. This can be overridden, to provide further modulation, by variations in cytosolic malate: a high concentration of cytosolic malate reduces both

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kinase mRNA and the accumulation of the kinase protein itself (Borland et al., 1999; Borland and Taybi, 2004). Mechanisms for sensing and signalling of malate, carbohydrate and CO 2 status are discussed (Borland and Taybi, 2004). Sugar sensing and signalling has been reviewed recently (Rolland et al., 2002).

13.7

Gene regulation

Coverage of the extensive literature on gene regulation by water stress is beyond the scope of a book on solute transport, but there are some pointers to how plants generally respond to water deficit. The short- and long-term effects of water deficit on plants are numerous and ultimately affect numerous processes, and so it is not surprising that large numbers of genes may be up- or down-regulated. Nor will it be surprising that different genes are seen to be affected according to the severity, and realism, of the stress applied. Conveniently rapid treatments can bear little relevance to the way water deficit develops in the field and can lose physiological response in a forest of pathological effects. Three sets of data using different methods of applying stress (all in microarray experiments using Arabidopsis thaliana) were analysed to look for genes that were affected by all the treatments: the treatments compared were as follows (Bray, 2004): r Desiccation: 3-week-old agar-grown seedlings were placed on filter paper in dim light for varying periods of time (Seki et al., 2002). r Osmotic shock: 4-week-old solution-grown seedlings were shocked with 200 mM mannitol (Kreps et al., 2002). r Drought: Soil-grown plants were drought treated for 8 days, at which time relative water content was 65% (Kawaguchi et al., 2004). In all three treatments a total of just over 800 genes were induced. However, only a small proportion of these were commonly induced or repressed by all three treatments (Bray, 2004). There were only 27 commonly induced genes (induced by all three treatments, all but one of which were also induced by ABA) and these served diverse functions. The induced genes functioned in metabolism, transport, signalling and transcription, encoded hydrophilic proteins, and there were some with as yet unknown function. There were only three commonly repressed genes and they were all linked to reduced growth. More specifically, there was a global decrease in the expression of genes that would promote cell expansion (Bray, 2004). It is interesting to note that the most ‘realistic’ treatment (that in drying soil) induced a total of 57 genes (while the desiccation and osmotic shock treatments induced 271 and 372, respectively) and down-regulated a total of 654 (other treatments downregulated 143 and 501). Such data reinforce the picture that plant response to water stress is generally a controlled slowdown. The analysis (Bray, 2004) further makes it very clear that it is very easy to find genes that are affected by water deficit but not so easy to work out which ones matter.

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335

Concluding remarks

Plants have needed to deal with water deficit since they first colonised the land, and the angiosperms have been around for 400 million years. Productivity has a competitive advantage but this is nothing compared with the competitive disadvantage of dying before completing the life cycle. It is perhaps not surprising that so many of the mechanisms that plants have evolved to cope with water deficit are concerned primarily with harm minimisation and damage limitation. The more arid the environments to which plants are adapted the more their adaptations are linked to conserving water. The generalised response of plants to reduced water is to reduce growth. Stage III drought is not a good place to be in. This has profound implications for the choice of selection criteria in the agricultural context. Even ‘obvious’ choices (plant water-use efficiency, osmotic adjustment and compatible solutes) have not been shown to have a simple relationship with yield and their value as criteria has been questioned.

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Kreps, J.A., Wu, Y., Chang, H.-S., Zhu, T., Wang, X. and Harper, J.F. (2002) Transcriptome changes for Arabidopsis in response to salt, osmotic and cold stress. Plant Physiology 230, 2129– 2141. Levchenko, V., Konrad, K.R., Dietrich, P., Roelfsema, M.R.G. and Hedrich, R. (2005) Cytosolic abscisic acid activates guard cell anion channels without preceding Ca2+ signals. Proceedings of the National Academy of Sciences 102, 4203–4208. Li, B. and Chollet, R. (1994) Salt induction and the partial purification/characterisation of phosphoenolpyruvate carboxylase protein-serine kinase from an inducible crassulacean acid metabolism (CAM) plant, Mesembryanthemum crystallinum L. Archives of Biochemistry and Biophysics 314, 247–254. L¨uttge, U. (1996) Clusia: plasticity and diversity in a genus of C3/CAM intermediate tropical trees. In: Crassulacean Acid Metabolism: Biochemistry, Ecophysiology and Evolution, Vol. 114 (eds Winter, K. and Smith, J.A.C.), pp. 296–311. Springer-Verlag, Berlin. L¨uttge, U. (2002) CO 2 -concentrating: consequences in crassulacean acid metabolism. Journal of Experimental Botany 53, 2131–2142. L¨uttge, U., Pfeifer, T., Fischer-Schliebs, E. and Ratajczak, R. (2000) The role of vacuolar malatetransport capacity in crassulacean acid metabolism and nitrate nutrition. Higher malate-transport capacity in ice plant after crassulacean acid metabolism-induction and in tobacco under nitrate nutrition. Plant Physiology 124, 1335–1347. MacRobbie, E.A.C. (2000) ABA activates multiple Ca2+ fluxes in stomatal guard cells, triggering vacuolar K+ (Rb+ ) release. Proceedings of the National Academy of Sciences 97, 12361–12368. Mambani, B. and Lal, R. (1983) Response of upland rice varieties to drought stress. I: Relationship between root system development and leaf water potential. Plant and Soil 73, 73–94. Marshall, J.G. and Dumbroff, E.B. (1999) Turgor regulation via cell wall adjustment in white spruce. Plant Physiology 119, 313–319. Martre, P., Morillon, R., Barrieu, F., North, G.B., Nobel, P.S. and Chrispeels, M.J. (2002) Plasma membrane aquaporins play a significant role during recovery from water deficit. Plant Physiology 130, 2101–2110. Maurel, C., Javot, H., Lauvergeat, V., et al. (2002) Molecular physiology of aquaporins in plants. International Review of Cytology 215, 105–148. Morgan, J.M. (1983) Osmoregulation as a selection criterion for drought tolerance in wheat. Australian Journal of Agricultural Research 34, 607–614. Munns, R. (1988) Why measure osmotic adjustment? Australian Journal of Plant Physiology 15, 717–726. Munns, R. (2002) Comparative physiology of salt and water stress. Plant, Cell and Environment 25, 239–250. Nobel, P.S. (2005) Physicochemical and Environmental Plant Physiology , 3rd edn. Elsevier Academic Press, Amsterdam. Oren, A. (1999) Bioenergetic aspects of halophilism. Microbiology and Molecular Biology Reviews 63, 334–348. O’Toole, J.C. and Cruz, R.T. (1983) Genotypic variation in epicuticular wax of rice. Crop Science 23, 393–394. Pantoja, O. and Smith, J.A.C. (2002) Sensitivity of the plant vacuolar malate channel to pH, Ca2+ and amino-channel blockers. Journal of Membrane Biology 186, 31–42. Passioura, J.B. (1983) Roots and drought resistance. Agricultural Water Management 7, 265–280. Porcel, R. and Ruiz-Lozano, J.M. (2004) Arbuscular mycorrhizal influence on leaf water potential, solute accumulation, and oxidative stress in soybean plants subjected to drought stress. Journal of Experimental Botany 55, 1743–1750. Price, A.H., Cairns, J.E., Horton, P., Jones, H.G. and Griffiths, H. (2002) Linking drought-resistance mechanisms to drought avoidance in upland rice using a QTL approach: progress and new opportunities to integrate stomatal and mesophyll responses. Journal of Experimental Botany 53, 989–1004.

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Ratajczak, R., Kemna, I. and L¨uttge, U. (1994) Characteristics, partial purification and reconstitution of the vacuolar malate transporter of the CAM plant Kalancho¨e daigremontiana Hamet et Perrier de la Bˆathie. Planta 195, 226–236. Raven, J.A. (1997) The role of marine biota in the evolution of terrestrial biota: gases and genes. Biogeochemistry 39, 139–164. Rolland, F., Moore, B. and Sheen, J. (2002) Sugar sensing and signalling in plants. The Plant Cell (Suppl.), S182–S205. Russell, B.L., Rathinasabapathi, B. and Hanson, A.D. (1998) Osmotic stress induces expression of choline monooxygenase in sugar beet and amaranth. Plant Physiology 116, 859–865. Schwacke, R., Grallath, S., Breitkreuz, K.E., et al. (1999) LeProT1, a transporter for proline, glycine betaine, and gamma-amino butyric acid in tomato pollen. The Plant Cell 11, 377–391. Seki, M., Narusaka, M., Ishida, J., et al. (2002) Monitoring the expression profiles of 7000 Arabidopsis genes under drought, cold and high salinity stress using a full-length cDNA microarray. The Plant Journal 31, 179–192. Serraj, R. and Sinclair, T.R. (2002) Osmolyte accumulation: can it really help increase crop yield under drought conditions? Plant, Cell and Environment 25, 333–341. Shabala, S.N. and Lew, R.R. (2002) Turgor regulation in osmotically stressed Arabidopsis epidermal root cells. Direct support for the role of inorganic ion uptake as revealed by concurrent flux and cell turgor measurements. Plant Physiology 129, 290–299. Sharp, R.E., Poroyko, V., Hejlek, L.G., et al. (2004) Root growth maintenance during water deficits: physiology to functional genomics. Journal of Experimental Botany 55, 2343–2351. Siedow, J.N. and Day, D.A. (2000) Respiration and photorespiration. In: Biochemistry and Molecular Biology of Plants (eds Buchanan, B.B., Gruissem, W. and Jones, R.L.), pp. 676–728. American Society of Plant Physiologists, Rockville, MD. Sinclair, T.R. and Ludlow, M.M. (1986) Influence of soil water supply on the plant water balance of four tropical grain legumes. Australian Journal of Plant Physiology 13, 329–341. Sobeih, W.Y., Dodd, I.C., Bacon, M.A., Grierson, D. and Davies, W.J. (2004) Long-distance signals regulating stomatal conductance and leaf growth in tomato (Lycopersicon esculentum) plants subjected to partial root-zone drying. Journal of Experimental Botany 55, 2352–2363. Spence, R.D., Wu, H., Sharpe, P.J.H. and Clark, K.G. (1986) Water stress effect on guard cell anatomy and the mechanical advantage of the epidermal cells. Plant, Cell and Environment 9, 197–202. Taiz, L. and Zeiger, E. (1991) Plant Physiology. Benjamin/Cummings, Redwood City, CA. Takahashi, S., Tamashiro, A., Sakihama, Y., Yamamoto, Y., Kawamitsu, A. and Yamasaki, H. (2002) High-susceptibility of photosynthesis to photoinhibition in the tropical plant Ficus microcarpa L. f. cv. Golden Leaves. Available at: http://www.biomedcentral.com/1471-2229/2/2. Tardieu, F., Lafarge, T. and Simonneau, T.H. (1996) Stomatal conductance by fed or endogenous xylem ABA in sunflower: interpretation of correlations between leaf water potential and stomatal conductance in anisohydric species. Plant, Cell and Environment 19, 75–84. Vera-Estrella, R., Barkla, B.J., Bohnert, H.J. and Pantoja, O. (2004) Novel regulation of aquaporins during osmotic stress. Plant Physiology 135, 2318–2329. Wei, C., Tyree, M.T. and Sturdle, E. (1999) Direct measurement of xylem pressure in leaves of intact maize plants: a test of the cohesion-tension theory taking hydraulic architecture into consideration. Plant Physiology 121, 1191–1205. Winter, K. and Smith, J.A.C. (1996) An introduction to crassulacean acid metabolism. In: Crassulacean Acid Metabolism: Biochemistry, Ecophysiology and Evolution, Vol. 114 (eds Winter, K. and Smith, J.A.C.), pp. 1–13. Springer-Verlag, Berlin. Yeo, A.R., Lee, K-S., Izard, P., Boursier, P.J. and Flowers, T.J. (1991) Short- and long-term effects of salinity on leaf growth in rice (Orzya sativa L.). Journal of Experimental Botany 42, 881–889. Yoshida, S. and Hasegawa, S. (1982) In: Drought Resistance in Crops with the Emphasis on Rice (ed. IRRI.), pp. 83–96. International Rice Research Institute, Manila. Yu, G.-X., Park, B.-H., Chandramohan, P., Geist, A. and Samatova, N.F. (2005) An evolution-based analysis scheme to identify CO 2 /O 2 specificity-determining factors for ribulose 1,5 bisphosphate carboxylase/oxygenase. Protein Engineering Design and Selection 18, 589–596.

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14 Salinity Anthony Yeo

14.1

Introduction

Salinity is unusual amongst conditions that are described as ‘stresses’ because the plants that grow naturally in these conditions are not particularly stressed. Conditions of high soil salinity are a clearly defined natural ecosystem for which there is an adequate, well-adapted, natural flora (halophytes). Saline environments are often fertile environments, and consequently, salt marshes rank amongst the most productive ecosystems on the planet. This contrasts with a plant surviving in a desert or on mine waste – struggling against inadequacy or excess in a way that a halophyte growing in sea water does not. The requirements placed upon the solute transport of plants in saline conditions can be broadly subdivided according to 1. whether ‘salt exclusion’ alone could be sufficient (Section 14.2) or 2. whether there must be an adjustment of water potentials. The latter can be further subdivided by whether this adjustment can (Section 14.3) or cannot (Section 14.4) be achieved without using sodium chloride. In a saline environment, all plants effectively partition water from the salt solution around their roots. They ‘exclude’ most (generally well over 90%) of that salt by membranes that are much more permeable to water than they are to salt. Of the salt that is taken up, only percentage points separate necessity and toxicity. The solute relations of halophytes, particularly dicoyledonous halophytes, is about co-ordinating the accumulation of salt from the environment. Much of the solute relations of non-halophytes (glycophytes) in saline conditions is concerned with responses, sometimes too desperate and too costly to be regarded as ‘tolerance strategies’, to stem and counter the influx of salt. When discussing these cases below, we will see that what is ‘acceptable’ or ‘unacceptable’ to the plant will depend upon the context. In a natural environment, an unacceptable cost is one that carries enough competitive disadvantage to undermine the viability of that plant in that ecosystem. In an agricultural setting, an unacceptable cost will be interpreted in terms of an uneconomic reduction in yield. Again, what is considered uneconomic will vary according to whether the crop is grown for subsistence or for profit. This chapter focuses mostly on sodium rather than chloride, though the transport of chloride is also discussed where appropriate. The emphasis on sodium is because there is invariably a driving force for passive entry of sodium into the plant.

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There are many candidate transport processes to facilitate the ‘uncontrolled’ entry of sodium. This entry may be due to inherently low selectivity of processes not evolved to provide discrimination in a saline environment. Entry may also be due to the selectivity of processes for other ions being overwhelmed by the excess of sodium. There are situations where sodium uptake is beneficial to a glycophyte, such as where there is a deficiency of potassium, and so sodium transport performs a necessary function. But the regulation of that process may not be tight enough to stop sodium uptake if the external concentration of sodium rises. In most cases, sodium influx is countered by active efflux, rather than by regulation of sodium influx by the sodium content of the root. Chloride influx from non-saline concentrations will be an active process; only at high external, and low cytoplasmic, concentrations can there be passive influx of chloride through an anion channel. There is also evidence that chloride fluxes are regulated by the chloride content of the root (White and Broadley, 2001). Chlorine is an essential element for all higher plants, whereas for sodium the role is established in only some cases (see Section 12.1). Although the essential requirements for chlorine (and sodium where demonstrated) are in the ‘micro-nutrient’ range, both elements may serve major roles in osmotic adjustment and ionic balance. Particularly in saline conditions, sodium and chloride may be the predominant mineral elements in the plant, reaching concentrations of several mmol g−1 dry mass, or over 500 mM on a tissue water basis, in halophytes.

14.2

External concentration of salt up to about 50 mM NaCl

This external concentration of salt is not sufficient to cause unacceptable growth reduction for osmotic reasons. The osmotic implications of the external solute potential in a saline environment has been recognised since the concept of ‘physiological drought’. If the water potential difference from medium to plant is reduced, then the plant must adjust osmotically and/or grow more slowly, or wilt and die. Consequently, the first ‘response’ of plants to suddenly imposed salinity (which, like suddenly imposed drought, is not a particularly common event in the ‘real world’) is to a water deficit. This may stop growth for a short while, but at low salinity this stoppage is usually transient and a new steady-state growth rate is established (Yeo et al., 1991; Munns, 2002). Salinity damage is expressed after a longer period of time when the accumulation of salt leads to damaging consequences in the leaves – a two-phase growth response to salinity (Munns, 1993). A whole array of processes at the plant, tissue and cellular levels of organisation may come into play to mitigate the effects of salt uptake. Long-term damage is due to unnecessary salt accumulation (because at this low salinity, accumulation is greater than that required for osmotic adjustment). At the extreme, salt accumulation kills leaves: both death rate and initiation rate are affected and the duration of individual leaves is reduced. Leaf area of individual leaves is reduced by reduced expansion (whether due to osmotic constraints, toxicity or lack of resources). Photosynthesis is reduced by direct effects of salt and

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by osmotic effects upon gas exchange. Together, these effects reduce the aggregate product of photosynthesis × effective leaf area × leaf duration. Damage is usually considered to result from too much salt in the cytoplasm (Flowers et al., 1977; Greenway and Munns, 1980) or in the apoplast, causing local osmotic stress (Oertli, 1968). Both compartments are critical because their small solute-available space amplifies any small imbalance of fluxes into and out of the compartment. The cytoplasm is also critical because of specific ionic requirements of macromolecular processes. It has long been recognised that higher plants require relatively a low concentration of sodium and a relatively high concentration of potassium in the cytoplasm, stemming probably from conserved requirements of protein synthesis (Wyn Jones et al., 1979; Leigh and Wyn Jones, 1984). An initial constraint such as this would have obviated any advantage in developing sodium-tolerant cytosolic systems and set the pattern for the widespread existence of potassiumactivated enzymes, which in turn accounts for the limited capacity of sodium to substitute for potassium. Salt exclusion at the root, by greater selectivity of transport processes, and/or, at greater cost, by active salt extrusion, is necessary and may be sufficient to allow the plant to grow at low salinity if it alone reduces net salt entry to a level that can be contained by the root system. The capacity of the root system to contain sodium is limited (the quantity of sodium is limited by the product of volume and the concentration that can be tolerated) and generally small in comparison with net uptake in saline conditions. Salt exclusion is a relative term. All plants that grow in saline solution exclude sodium chloride to the extent that the concentration in the xylem is much less than the concentration in the solution. This exclusion is typically in the range of 94–99% (Munns et al., 2006). What this implies (see also below about the balance between influx and efflux) is that differences in salt tolerance are about the last percentage points of efficiency in ‘filtering’ salt out of the solution that replaces transpiration. Even the halophyte Suaeda maritima growing at its optimal salinity, and where growth is now limited by ion transport capacity (Yeo and Flowers, 1986a), excludes about 85% (based upon a xylem concentration of 56 mM measured at an external concentration of NaCl of 200 mM; Clipson and Flowers, 1987). Perhaps surprisingly, sodium loading to the xylem is active for salt-sensitive plants at low salinity. This is possibly a consequence of root homeostasis (Tester and Davenport, 2003) and so possibly has little need for specificity. Roots have some set-point of ion concentration above which movement into the xylem will occur, whether or not the ions are ‘needed’ in the shoot. Once the salt enters the xylem, it enters into a potential dead-end where it can accumulate over time in leaves until the concentration is high enough to cause damage. ‘Interception’ of salt by accumulation in older roots, leaf sheaths, petioles, older leaves and so on can have a protective role in conserving active leaf area, but the capacity is strictly limited by the volume of tissue that can be sacrificed and the quantity of salt that it can contain. Such re-absorption is likely to make much difference only if the initial xylem loading is only marginally in excess of the capacity (i.e. growth rate) of the leaf to dilute this, for example in Durum wheat (Davenport

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et al., 2005). The grasses have leaf sheaths in which to accumulate sodium, though accumulation in petioles in dicots has also been reported. Once salt reaches the leaves, effective compartmentalisation between protoplast and apoplast will delay or avoid damage caused by internal osmotic stress (originally described as the ‘Oertli hypothesis’ (Oertli, 1968; Flowers et al., 1991)). Leaving salt behind in the apoplast (or pumping salt out of the protoplast other than into a gland) is lethal for the simple reason that an ion in the apoplast contributes about 100 times the osmotic potential that it would in the protoplast. Effective compartmentalisation between cytoplasm and vacuole will delay or avoid the achievement of toxic concentrations in the cytoplasm. Recirculation of salt from younger to older leaf tissue may be effective if this serves to maintain photosynthetically active leaf area, which is a balance between protection and sacrifice. However, source–sink relationships will be very critical if the salt entering the phloem is not to go to the developing leaf tissue and make matters much worse. Recirculation in the phloem back to the root is a possibility as a tolerance mechanism, but the validity depends upon the ability of the root to efflux the extra salt load reaching the root symplasm. This is debatable, if it is presumed that the salt got only to the shoot because the root was unable to keep out enough sodium in the first place, and this is discussed in Section 14.11.

14.3

External concentration of salt up to about 100–150 mM NaCl

This external concentration of salt is sufficient to cause unacceptable growth reduction for osmotic reasons and uptake of ‘nutrient’ ions (not sodium) and/or organic synthesis can achieve osmotic balance at an acceptable cost. Compensation for the external salinity is required or there will be an osmotic component of salinity damage other than the transient effect described in Section 14.2. Unless this adjustment occurs, there will be a permanent substantial reduction in growth rate due to the osmotic effects of salinisation, which will exacerbate any long-term effects of salt entry. As the external salinity increases, the osmotic adjustment requiring intracellular compartmentalisation becomes essential. This is a double requirement: something to adjust the vacuole and something to adjust the cytoplasm. Long-term damage in the shoot can still be due to unnecessary salt accumulation (above that needed for osmotic adjustment) and also due to the inability to adequately compartmentalise the concentration of ions necessary for osmotic adjustment. Salt exclusion at the root is necessary (even halophytes exclude most of the salt) but is not sufficient since if osmotic adjustment is not going to be achieved with sodium chloride, then an increased accumulation of some osmotically active species is necessary to maintain the water balance. This may be achieved by the increased uptake of available inorganic ions (principally potassium) and/or by the synthesis of organic anions and/or neutral solutes. Both carry costs in terms of energy as well as resources (Yeo, 1983). As above, interception and ‘recirculation’ of salt can have a protective role, but this becomes less likely to be of consequence as the salt load in the shoot increases with external salinity. The greater the increase in osmotic

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pressure that is needed, the more demanding the requirements of compartmentalisation become. This relates to regulating both the ion concentration and the organic solute concentration in the cytoplasm – by keeping excess ions in the vacuole and keeping osmolytes in the cytoplasm. The production of such organic solutes is likely to be an acceptable cost only when such solutes are compartmentalised effectively within the cytoplasm. Excretion of salt via glands can limit the concentration of salt in the leaf (Section 14.8).

14.4

External concentration of salt above about 150–200 mM

This external concentration of salt is sufficient to cause unacceptable growth reduction for osmotic reasons and uptake of nutrient ions and/or organic synthesis cannot achieve osmotic balance at an acceptable cost. This moves into the range which is exemplified by the salt-accumulating halophytes. The survival depends on at least partial utilisation of sodium for osmotic adjustment. This is beyond substitution of sodium for potassium in conditions of potassium deficiency. It requires concentrations of sodium within the protoplast that would be toxic if not effectively compartmentalised within the vacuole. The large quantity of sodium being moved about the plant demands precise integration of sodium transport processes in all parts of the plant in order to maintain critical steady-state concentrations in the small cytoplasmic and apoplastic compartments. Such integration of sodium transport is probably the defining feature of the halophilic flora. The key features of halophytes at the cellular level are massive accumulation of sodium chloride in the central vacuole, often associated with the development of succulence and linked with the production and effective compartmentalisation of a compatible solute in the cytoplasm. The best studied example of a compatible solute in halophytes is glycine betaine. The intracellular localisation has been established by electron microscopy, following freeze substitution, of the complex of glycine betaine with iodoplatinic acid (see Sections 2.6.2 and 2.6.7). The results demonstrated that glycine betaine was restricted to the cytoplasm in the halophyte S. maritima and that the amount of glycine betaine present was proportional to the salinity in which the plants were grown (Hall et al., 1978). But even these cellular-level features would be nothing on their own. They must be combined with co-ordination at the plant level, which allows potentially lethal quantities of salt to be transported about at steady-state concentrations which, at optimal growth, approach the upper limits of safety (Yeo and Flowers, 1986a). It was an early hope that the study of halophytes would inform crop improvement in salt-sensitive species. However, an extensive literature for crop species relates greater tolerance to lesser sodium uptake (particularly in the key agricultural crops of the Poaceae) and this diametrically opposing finding is not encouraging. It does clearly show that tolerance is not the opposite of sensitivity. Salt sensitivity in glycophytes is associated with too much ‘accidental’ sodium entry. This would imply that tolerance would be achieved by minimising entry or maximising efflux. However, this is not the mechanism used by those plants that are ecologically

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successful in saline environments. True tolerance has nothing to do with minimising salt entry. There is another sense in which sensitivity may not be the opposite of tolerance, salt sensitivity may be due to unrelated processes failing rather than tolerance processes working poorly. The fundamental physiological problems of halophytes and glycophytes in saline environments were established and reviewed long ago (Flowers et al., 1977; Greenway and Munns, 1980; Flowers et al., 1986). Nearly all research in the past decades has focused on individual components, increasingly on individual gene products. This passed through a period of reductionism, focusing on single-cell systems (yeast and cell cultures). Since the disappointing outcome that it was very rare to be able to regenerate a tolerant plant from a tolerant cell (Smith and McComb, 1981; Dracup, 1991; Winicov, 1998), there has been increasing recognition that there is a quantum jump between the cellular and the whole-plant level of organisation and consequently that what works in a cell may not work in a plant. This has suggested a host of investigations where gene products have been inserted, under- and over- expressed, and blocked, antisensed and mutated, in planta. Research has dismantled plants and crops into cells and molecules and in many ways is now seeking to build them back up again. At the moment there is a host of information on the function of individual gene products in relative isolation (such as expressed in heterologous systems), but it is still difficult to fit all this into a picture of what needs to be put together to make a tolerant plant. The lack of understanding about how sodium acquisition is co-ordinated has impaired progress towards improving salt tolerance of crops (Pardo and Quintero, 2002). It is a major problem in writing about plant solute transport that the quality of plant physiological data associated with evaluating the molecular interventions in solute transport is so often poor. There have been many pleas for the physiological analysis of molecular interventions to become appropriate, relevant and realistic (e.g. Yeo, 1998; Munns, 2002; Tester and Davenport, 2003). A combination of harsh and irrelevant treatments which do the wrong thing, together with a halfhearted ‘snapshot’ approach to monitoring what happens, makes it always difficult, and frequently impossible, to deduce what the intervention did or did not do. The physiological and agricultural outcomes cannot be revealed by a casual photograph or by one or two ion analyses performed on dead and dying plants. But publication after publication persists in making the seemingly obligatory nod in the direction of ‘crop improvement’ to justify its existence and so aspires to plant and crop physiology. If any intervention is offered as having any relevance to agriculture, it must be possible to make some evaluation of its effect over the life cycle of the plant, using treatments that are representative of what may happen in the field.

14.5

‘Molecular’ tolerance

No higher plant is tolerant of salt in the molecular sense that the halobacteria are. The gene encoding malate dehydrogenase from the archaebacterium Haloarcula

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marismortui, when cloned in Escherichia coli, produced a protein indistinguishable from the native protein from H. marismortui, which was activated only when the concentration of NaCl was increased to 3 M (Cendrin et al., 1993). Metabolic enzymes of halobacteria may function only in extremely high salinity, but higher plants are not tolerant of salinity in their cytoplasm. This may indicate that despite the high (higher than today) salinity of early oceans, the progenitors of plant life on land developed in marginal environments of lower salinity (Knauth, 1998). But whatever the early history, plant life had moved too far down a path requiring defined, non-saline, conditions in the cytoplasm long before the angiosperms evolved (about 0.4 Gy ago) and so long before the development of any of the many families that currently have halophilic members. Currently, no plant, even the most salt tolerant of halophytes, does any more than slightly bend the rules: all die if they cannot control the concentration of salt in the cytoplasm.

14.6

Cellular tolerance

If we consider a single vacuolate cell in solution/suspension culture, then sodium can enter across the plasma membrane via any of a collection of transport processes (channels, secondary active pumps, both symport and antiport) of various selectivity, in response to a substantial electrochemical potential difference. The sodium can be actively pumped out again or it can be accumulated into the vacuole. The most critical factor for the cell is the concentration of sodium that can be tolerated in the cytoplasm (not more than about 100 mM; less in most glycophytes, a figure than might be modified by osmoprotectants). The volume fraction of cytoplasmic solute-available space is a small part of non-meristematic cells. Consequently, all the unidirectional fluxes at plasma membrane and tonoplast are required to be closely regulated if an acceptable steady-state concentration in the cytoplasm is not to be exceeded. The concentration that can be accommodated in the vacuole is dependent upon the concentration gradient that can be sustained across the tonoplast (and by implication the cost of sustaining it). If the total cation concentration to be held in the vacuole exceeds the total cation concentration that can be tolerated in the cytoplasm, then a cytoplasmic compatible solute, and appropriate compartmentalisation, are also necessary. The quantity of sodium that can be accommodated in the vacuole depends upon its concentration in the vacuole, the volume of the vacuole and the rate of expansion of the vacuole. Expansion at constant internal concentration would require that net uptake be linked to expansion rate, whichever controls which. It must also be considered that the main transport processes pumping sodium out of the cytoplasm across either membrane are probably proton antiporters and thus cytoplasmic pH as well as ionic homeostasis are inextricably linked: high-affinity sodium uptake in potassium-starved barley prevents a decline in tissue pH (Carden et al., 2003). Finally, all of this consumes energy, probably a great deal of energy, especially if sodium persists in leaking the wrong way (1:1 sodium/proton antiport is potentially a very expensive answer; Tester and Davenport, 2003).

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347

Moving on to a cell in a plant

The cytoplasm is no longer only a concentric ring around the vacuole, but is now part of a transport pathway from outside to xylem. Transport to the xylem exists in parallel with accumulation into root vacuoles, and relative priorities vary with species, with concentration and duration of exposure to salt. It is part of the function of roots to transport ions to the shoot. Transport to the xylem also becomes an option for homeostasis of root cells when they reach their capacity for sodium. Pumping sodium into the xylem will not be more energetically costly than pumping it out of the root, and pumping it into the dilute xylem with the transpiration stream is likely to be easier than pumping it into the concentrated outside against the transpiration stream. There may be conditions of high symplasmic sodium concentration and low xylem sodium concentration where movement of sodium into the xylem could be a passive consequence of electrochemical potential differences between symplasm and xylem; but such conditions are probably extremely rare. There is also a mass flow of solution through the root from outside to xylem, coupled solute and water flows and, in some cases, a dilution effect setting up and maintaining a concentration difference from symplasm to xylem. The surroundings of the cell are now much more complex than the single cell in solution. In the soil there are unstirred layers, depletion zones and advection zones. Sodium pumped out of the root does not go away, but rather it is constantly drawn back towards the root by the mass flow of soil solution. The salt concentration at the root surface probably increases above that of the bulk solution. The main additional consideration for leaf cells is that they are at the end of the line. The transpiration stream brings sodium up in the xylem and pure water evaporates to the air. The considerations about quantity and concentration and vacuolar volume are the same as for a root cell, and constancy of concentration can be achieved only if net import and relative growth rate are in balance. Once sodium reaches the leaves, there is nowhere else to go unless the plant has salt glands or salt hairs, in which case it may be secreted to the outside. Phloem recirculation is possible, but unlikely (Munns, 2005), presenting staggering problems of managing ion concentration in the apoplast of the leaf lamina: recirculation may be a real possibility only by xylem–phloem exchange within the vascular bundles. Overall, a cell in a root, and even more so a cell in a leaf in the air, has problems that are not faced by a single cell in solution or suspension culture. Furthermore, what may work for a cell in a solution (pumping sodium out) would be lethal for a cell in a leaf in the air (Yeo, 1998).

14.8

Salt glands

Some halophytes use salt-secreting glands as a successful method of removing excess ions from their leaves (Thomson et al., 1988), which reduces the demand for the tight coupling between net transport and growth discussed already for the succulent halophyte S. maritima (Yeo and Flowers, 1986a). Salt glands are found

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in both mono- and dicotyledons. In grasses, these glands are simple in structure, often called microhairs, and, with one exception, are bicellular (Flowers et al., 1990; Ramadan and Flowers, 2004). Most grasses (except species of the Pooideae) possess microhairs, though only in halophytes do these act as salt glands. In dicotyledons, salt glands range from the simple bladders present on the leaves of species of Atriplex to complex structures of between 8 and 40 cells in Tamarix and Aegilitis (Thomson et al., 1988). In the more complex glands, the secretory cells are isolated from the remainder of the apoplast by cuticularisation. Salt glands from different species have many common features despite the differences in complexity. The presence of numerous mitochondria and dense cytoplasm, together with reduced secretion in low temperature, anoxia, or the presence of metabolic inhibitors, all suggest that the secretion itself is an active process (Flowers and Yeo, 1992). Salt glands allow the plant to cope with net influx of salt being in excess of growth rate. However, the plant still has to co-ordinate all the ion fluxes along the path from uptake at the root plasma membrane to secretion by the gland cell(s).

14.9

Selectivity at the root

The external concentration in saline conditions is usually greater than the concentration in the symplasm and the electrical potential inside the cell is invariably and substantially negative. In a sodium-hyper-accumulating mutant of Capsicum annuum, membrane hyper-polarisation was sufficient to account for the increased inwards sodium current, and was likely to be mediated by control of a number of transporters rather than by mutation in a single transporter (Murthy and Tester, 2006). Initial rates of unidirectional influx at 50 mM NaCl (average 1 μmol g−1 fresh weight min−1 ) have been calculated to be equivalent to 180 nmol m−2 s−1 of root plasma membrane (Tester and Davenport, 2003). Net influx is typically a small fraction of this, implying that most of the sodium is pumped out again. As already mentioned, ‘most’ means a few percent short of 100%. Since this last few percent is so critical, it provokes the question of whether there is a law of diminishing returns in force here, and raising the last few percent, however ‘desirable’, is somehow unachievable or unaffordable. There are many candidate carriers and channels that may mediate sodium uptake under saline conditions. These are summarised briefly from extensive reviews (Amtmann and Sanders, 1999; Cushman, 2001; Tester and Davenport, 2003; RodriguezNavarro and Rubio, 2006). Historically, living systems developed a dependency on substantial potassium concentration (around 100 mM) and high potassium:sodium ratios in the cytoplasm at a time when potassium, though not the most abundant cation, was nevertheless in inexhaustible supply. There was consequently no reason to not use potassium for gross roles, such as osmotic adjustment of the vacuole, which do not specifically require potassium. Colonisation of land and freshwater environments required adaptation to situations in which the external concentration of potassium was very much lower and on occasions became depleted. This required the development of carriers with high enough affinity for potassium to accumulate it

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from the concentrations available on land. The fact that some gross roles of potassium are not potassium-specific allows for sodium, which may also be in low supply, to substitute for potassium, leading to an advantage of developing high-affinity sodium transporters as well (Rodriguez-Navarro and Rubio, 2006). Many transport processes been identified in recent years and some more than others have been implicated as pathways of sodium entry. HAK/KUP transporters are highly potassium selective and calcium insensitive, but not necessarily high-affinity. HKT transporters have been associated with both potassium and sodium uptake according to the species and heterologous expression system used. The wheat HKT1 was originally associated with potassium-proton symport (Schachtman and Schroeder, 1994) but could co-transport sodium and potassium when expressed in yeast or Xenopus laevis (Rubio et al., 1995). A family of HKT1 transporters were described in rice, with OsHKT1 proposed to mediate sodium uptake in roots in conditions of potassium starvation (Garciadeblas et al., 2003). The situation proposed in rice suggests that a process of advantage in a natural environment where sodium can ‘safely’ substitute for scarce potassium could become a liability if rice continues to transport sodium when planted in saline conditions to which it is not evolved. AtHKT1 (the arabidopsis homologue) can mediate sodium influx but the data are from heterologous systems (Uozumi et al., 2000) and its function in a plant in the field is uncertain. However, there are pronounced species differences, and HKT transporters appear to be expressed and mediate sodium uptake into root epidermal and cortical cells in rice but not in arabidopsis (Rodriguez-Navarro and Rubio, 2006). However, HKT transporters may be involved in movement of sodium within the plant, particularly for sodium-specific roles such as ‘defensive’ re-uptake of sodium from the xylem (Rodriguez-Navarro and Rubio, 2006). The model currently suggested for initial uptake into the root (RodriguezNavarro and Rubio, 2006) is that HAK1 is involved in high-affinity potassium and low-affinity sodium uptake and that HKT1 is a high-affinity sodium transporter, does not transport potassium and is inhibited by potassium. Rodriguez-Navarro and Rubio (2006) ascribe the conflict between results showing HKT transporters as high-affinity sodium transporters and the characteristics of the wheat HKT1 as due to the complexities of the heterologous expression systems. KIRCs (potassium inward rectifying channels) are a major pathway for lowaffinity potassium uptake, but appear to have little significance in sodium influx. KORCs (potassium outward rectifying channels), include SKOR, which is involved in releasing potassium to the xylem for transport to the shoot. Only in situations of high symplastic sodium and low xylem sodium is passive loading of sodium into the xylem via a channel likely. NSCCs (non-selective cation channels) have lower potassium:sodium selectivity than other channels and may function as a major pathway for sodium entry at high external salinity (Cushman, 2001). The most likely pathway for calcium-sensitive sodium influx are the NSCCs (Amtmann and Sanders, 1999; Davenport and Tester, 2000; Tester and Davenport, 2003). Amongst the NSCC group, the main candidates are CNGCs (cyclic nucleotide gated) and GLRs (glutamate activated) channels.

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However, at present, functions in planta are uncertain until the target membranes and cell types for these channels are established (Tester and Davenport, 2003). LCT1 (from wheat) elicited low-specificity cation uptake when expressed in yeast; however, this may have been through an effect on endogenous transport systems in the yeast and the activity of LCT1 is uncertain in plant systems. NSCCs and high-affinity potassium transporters have emerged as potential pathways of sodium entry together with a low-affinity cation transporter represented by the wheat protein LCT1 (Amtmann et al., 2001). LCT1 conferred potassium uptake capacity only at 1 mM potassium and above, indicating relatively low affinity. Heterologous expression in yeast increased salt sensitivity (Amtmann et al., 2001). In summary, the high-affinity potassium transporter (HKT1), low-affinity cation transporter (LCT1) and NSCCs are considered the most likely mediators of sodium entry (Rus et al., 2001). AtHKT1 is another potential sodium-influx system that is thought to be countered by the sos-pathway that leads to the SOS1-encoded sodium/proton antiporter (Rus et al., 2001). In some species, an apoplastic route to the xylem, with a proportion of the transpiration stream evading membrane control, is a significant source of entry for sodium ions. In rice, there is close agreement between the movement of sodium and a membrane-impermeant dye to the shoot, which can explain the large individual differences in sodium transport. Furthermore, the differences in both sodium and dye transport co-segregate during breeding (Yadav et al., 1996). The movement of dye through the apoplastic continuity is constrained more than is the movement of water and such dyes are not exact quantitative tracers for water movement (Yeo et al., 1987). However, corrections for relative mobility in the apoplast (Yeo et al., 1987) provide estimates of bypass flow of the order of 1% in rice, which is quite sufficient to be a key factor in salt toxicity of this relatively sensitive species because of its rapid transpiration per unit leaf water volume. A leakage of 1% has to be seen in the context that the total leakage of sodium in salt-sensitive glycophytes is only of the order of a few percent. Halophytes often have anatomical adaptations to minimise apoplastic leakage. Apoplastic leakage may be more pronounced in rice due to its ecology, but could be significant in other species, particularly where there is damage to the root by physical injury, pathogenicity, salt treatment or osmotic shock. It is also possible that the bypass flow serves no useful purpose and might be selected against without penalty. The simplest scenario, available only for salt-sensitive plants growing in fairly low-saline conditions, is just to keep sodium out – ‘salt exclusion’ in its most specific sense. If this could be achieved, then none of the problems of sodium getting into the cytoplasm or into the xylem stream would arise. The caveat is that there must be adequate potassium for osmotic needs, including any necessary osmotic adjustment to the external salinity. In these circumstances, particularly for agricultural crops liberally dosed with fertilisers, maximising potassium/sodium selectivity through modifying transporters and channels, even ‘knocking out’ some that seem to get in the way, can appear to be a potentially viable solution. However, the degree to which any component can be regarded as working ‘in isolation’ is as yet unknown, as are the purposes of some of the processes held responsible for accidental entry of sodium.

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The consequences upon regulation and homeostasis of ‘rocking the boat’ have yet to be evaluated over a useful timescale. A telling example, giving some hint of the network of interaction that will have to be contended with, is the finding that disabling a PIP aquaporin resulted in compensatory increase in root:shoot ratio to restore hydraulic conductance (Kaldenhoff et al., 1998; Martre et al., 2002). An interesting observation has been that over-expression of a PIP aquaporin in transgenic tobacco improved plant vigour under good growth conditions, but failed to have any benefit under salt stress and was in fact deleterious under drought stress (Aharon et al., 2003). Above all it is vital to recognise that, even if maximising exclusion were a possibility, it is not a generalisation. There are some agricultural situations in which salt exclusion alone would theoretically work, but as the external salinity rises this approach will stall and fail, through the cost and feasibility of excluding sodium against an ever-increasing potential influx and through the cost and feasibility of adjusting osmotically with something else. The utility also depends upon whether leakage of sodium can be reduced or whether most of the leaked sodium would have to be pumped out again – and at what cost. Consideration of unidirectional and net fluxes suggest that net sodium influx in glycophytes is only a fraction of the unidirectional influx and that this difference is accounted for by active sodium efflux (Tester and Davenport, 2003). Control of efflux appears to be more important than control of influx in regulating net sodium transport in glycophytes (Qiu et al., 2002). The difference between unidirectional influx and unidirectional efflux (i.e. the net influx) must be accommodated in the vacuoles of the root or loaded into the xylem and accommodated in the shoot. The cost implications of regulating sodium uptake in saline conditions principally by active sodium extrusion are enormous. If the stoichiometry of sodium-proton antiporter is 1:1, it will cost an ATP for each sodium ion extruded. It is straightforward arithmetic to calculate what will happen if unidirectional influx at the plasma membrane continues unchecked and unopposed: the only debate will be over exactly how many minutes the cell takes to die. It is perhaps significant that halophytes appear much less dependent upon sodium extrusion. A halophyte deals with salt by moving sodium directly to the shoot (see below). Glycophytes, obviously, do not. Another consideration which might limit the options of a halophyte for sodium extrusion is that the pH of seawater is 7.5 or above and many saline soils are sodic-saline with alkaline pH. This will affect the ability of the plant to maintain a concentration gradient of protons across the plasma membrane. Boundary effects will usually allow the proton pump to acidify the apoplast and the immediate surroundings, but this may be compromised where there is a substantial independent mass flow of soil solution, such as in tidal salt marshes and on irrigation with brackish/saline water. Such conditions could stall an electro-neutral exchanger (Garciadeblas et al., 2001). The short-term influx of sodium in arabidopsis, at 25 mM NaCl, of about 0.56 μmol g−1 fresh weight min−1 (Essah et al., 2003, Fig. 2) was very similar to that for S. maritima, where the value was 0.44 ± 0.06 μmol g−1 fresh weight min−1 (Wang and Flowers, unpublished results). An interesting outcome is the close agreement (considering the approximations) between the unidirectional influx of sodium in glycophytes at low (50 mM) NaCl and the flux of sodium (net

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with little if any efflux) in the halophyte S. maritima at high (340 mM) NaCl. The value of 180 nmol m−2 s−1 (Tester and Davenport, 2003) is only a little less than the 250 nmol m−2 s−1 , which would be needed to account for sodium uptake in the halophyte S. maritima at 340 mM based on similar assumptions of uptake occurring over the whole plasma membrane surface of the epidermis and cortex. The estimate for glycophytes is based on a general approximation (Pitman, 1963) and for Suaeda is based upon extensive stereological analysis (Yeo and Flowers, 1986a). As discussed for the S. maritima example, it is unlikely that salt would have equal diffusive access to the whole root membrane surface and so the unit area rate at uptake sites was expected to be many times greater. What is perhaps surprising is that the rate of sodium uptake by the halophyte growing at its optimal salinity is little more than a glycophyte at a fraction of that salinity. The difference appears to be largely about what halophytes and glycophytes do with this sodium, rather than massive differences in potential sodium uptake in the first place. In the glycophyte example, most of this sodium flux will be effluxed; in the halophyte, this is the sodium flux accounting for net transport to the shoot. There may be regulatory differences in halophytes, which warrant further investigation. The halophyte Spergularia marina showed linear uptake over a prolonged period without evidence of efflux, and its rate of sodium uptake was lower than that of wheat at the same concentration of NaCl (Cheeseman et al., 1985). A more salt-tolerant relative of arabidopsis, Thellungiella halophila, differed in that a low unidirectional influx of sodium contributed to a low net influx of sodium (Wang et al., 2006). Growth and net sodium transport were tightly linked in S. maritima. There is some suggestion that unidirectional sodium influx in halophytes could be regulated (which it is demonstrably not in glycophytes) though this is not yet proven. Investigation of control of sodium transport in Durum wheat indicated there was no difference between relatively tolerant and relatively sensitive genotypes in terms of unidirectional uptake of sodium by the root, but that differences resided in reduced xylem loading as well as in increased re-absorption from the xylem into the leaf sheaths in the more tolerant cultivar (Davenport et al., 2005). Although at steady state, the great majority of the unidirectional influx is effluxed, the rates of unidirectional influx in both varieties (about 65 nmol g−1 fresh weight min−1 ) were well below the average values calculated for glycophytes (Tester and Davenport, 2003). It is possible that regulating xylem loading of sodium will be a possibility only in situations where the unidirectional influx of sodium is relatively small and so can be opposed by root efflux if the salt is not transported to the shoot. In situations where unidirectional sodium uptake exceeds the capacity of the root to efflux or contain it, the xylem loading of sodium may be an inevitable (and necessary) consequence of root ionic homeostasis. Accumulation into root vacuoles both regulates cytoplasmic sodium and allows sodium to be used ‘harmlessly’ to ‘cheaply’ balance the vacuole osmotically. The capacity of the root to ‘store’ salt is limited and linked directly to relative growth rate. The capacity is limited because of the root:shoot ratio, because root cells do not commonly become succulent and because (for reasons as yet unknown) root cells are not found to accumulate such high concentrations of salt in their

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vacuoles as leaf cells. The other direction for salt in the root to go is into the xylem stream and be moved to the shoot. Concentrations of salt in roots are often relatively stable while the concentration in the shoot continues to rise with time.

14.9.1

Root selectivity for chloride

Chloride is negatively charged and so the driving force for passive movement at the plasma membrane is very different from that for sodium, and in most physiological conditions, the electrochemical potential gradient will drive chloride out of the cell (Felle, 1994). If we refer to Eq. 3.21, it is apparent that a concentration gradient of chloride from outside to cytoplasm of one or two orders of magnitude is needed to permit for passive influx at typical membrane potentials. Such gradients are possible on initial exposure to saline conditions when cytoplasmic chloride concentration is initially low, and perhaps for longer at very high external salinity. Such thermodynamic considerations imply that active chloride transport will certainly be necessary, at all but highly saline conditions. A proton-chloride symporter was identified in root hair cells of Sinapis alba (Felle, 1994) with a stoichiometry of two protons to one chloride. This is the transporter that drives active chloride uptake, is voltage independent and is driven by the pH gradient across the membrane. The only inwardly directed chloride channel described in the plasma membrane of root cells is the OR (or outward rectifying) channel (Skerrett and Tyerman, 1994) [(for a summary of chloride transporters – pumps and channels – see Fig. 5 in White and Broadley, (2001)]. The OR channel has low unitary conductance and is permeable to nitrate as well as to chloride and contributes to chloride influx at high external concentrations of chloride. There is evidence (White and Broadley, 2001) that there is a regulation of net chloride uptake: this may be by feedback via the chloride concentration in the root to decrease influx and/or increase efflux. There is also some evidence that chloride uptake is ‘demand-driven’, which suggests a degree of regulation that is not seen for sodium in glycophyte species.

14.10

Transport from root to shoot

Salt damages the shoots of glycophytes to varying degrees. Vigorous plants may do better because the development of shoot tissue may outstrip the net transport of sodium. But there is no clear evidence in glycophytes that net transport of sodium and relative growth rate are inter-related, other than in the sense that excess salt transport is lethal. Within the shoot, how much the damage occurs is due to an inability to adequately compartmentalise sodium in the vacuole, or due to an inability to regulate the delicate balancing act of getting the salt from the xylem across the apoplastic and cytoplasmic compartments and into the vacuole, is not known. It is important to know this, because one reason, or the other, or both, prevents glycophytes accommodating sodium in the shoot. So long as they cannot, and sodium extrusion is the main option for regulating net uptake, the options for agricultural improvement are quite limited.

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Since it is improbable that sodium leaks into the xylem in glycophytes it appears that it is put there ‘deliberately’. Whether this is because it is sodium or just because it is any cation which is ‘in excess’ in the root is not known, but this is of survival advantage. Because shoot:root ratios are generally greater than unity, the plant will die more quickly if sodium toxicity kills root cells than if excess sodium is moved to the shoot. SOS1 is a sodium-proton antiporter, is localised in the plasma membrane of epidermal and stelar cells of the root and is suggested to be involved, in different circumstances, either in xylem loading or recovery of sodium from the xylem (Shi et al., 2002). The sos mutants were identified by their inability to sustain root growth in NaCl. The sos1 mutant accumulates less sodium (Zhu et al., 1998). This would be compatible with xylem loading of sodium being an active process and that it serves a role in root homeostasis because the failure of xylem loading could lead to toxic accumulation of sodium in the symplasm of the root and might account for the lack of root elongation in the mutant. This would however be a picture ‘at a moment in time’; with continued exposure to salt, this root damage would be expected to lead to reduced shoot growth and so to catastrophic increase in sodium concentrations in the shoot. CHX21 is a putative sodium-proton antiporter from arabidopsis belonging to the CHX cation transporter family. Insertion of transposons into CHX21 resulted, in the presence of NaCl, in xylem sodium and leaf sap sodium concentrations being reduced, and CHX21 was concluded to be involved in loading sodium into the xylem (Hall et al., 2006). However, a cautionary note is sounded in that transposon insertion into CHX21 also produced gross changes in rosette size and flowering time. This is one example of how genes for potentially ‘unwanted’ sodium transport processes are linked into other things. The situation in the halophyte is also likely to be one of active transport of sodium into the xylem. Although it was speculated earlier that sodium might leak into the xylem in S. maritima, consideration of the concentration difference between root symplasm and xylem makes this unlikely. The concentration in the symplasm calculated to sustain radial movement is at least 80 mM (Yeo and Flowers, 1986a) and upper limits measured by X-ray microanalysis and compartmental efflux analysis are 100–150 mM (Harvey et al., 1981; Yeo, 1981). The concentration of sodium in the xylem, calculated from transpiration and net transport measurements, was 56 mM for plants growing at an external concentration of 200 mM NaCl (Clipson and Flowers, 1987). The sodium concentrations in the symplasm and xylem lumen are therefore unlikely to differ by more than threefold for plants growing in high salinity. Unless the electrical potential difference at the symplast/xylem boundary is unusually depolarised, loading of sodium into the xylem would have to be active. This flux would be about 1200 nmol m−2 s−1 averaged over the plasma membrane surface area of the stelar parenchyma, more if restricted to the xylem parenchyma, and this is based on observed net fluxes and measured cell areas (Yeo and Flowers, 1986a). Although this would be a considerable energy cost, the halophyte would not be using more energy to pump sodium into the xylem than the glycophyte would be using to pump it back out again, and to much better effect.

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There have many reports since the 1960s that sodium may be recovered from the xylem stream in old roots and in stems, petioles and leaf sheaths. For example, evidence was found of sodium and chlorine accumulation in xylem parenchyma cells of roots of Zea mays when exposed to 50 mM sodium chloride as well as ultrastructural changes within these cells (Yeo et al., 1977). However, unless the salt is going to be moved back to the root and pumped out again, these options have strictly limited capacity, but may serve, at low external concentrations and low rates of sodium uptake, to provide another layer of protection for a while to the expanding and photosynthetically active leaves. The efficiency of compartmentalisation, at all possible cellular and tissue levels of organisation, determines the average concentration in the shoot that can be sustained without causing a reduction in growth rate. So long as relative growth rate stays in step, or keeps ahead, of net sodium import, then the situation remains manageable. An appropriate analogy is Dickens’ Mr Micawber ‘Annual income twenty pounds, annual expenditure nineteen pounds nineteen and six, result happiness. Annual income twenty pounds, annual expenditure twenty pounds ought and six, result misery’ (Dickens, 1850). It is necessary to only translate ‘income’ to ‘growth rate’ and ‘expenditure’ to ‘net sodium import’. If the net import exceeds relative growth rate, the shoot ion concentration will rise and sooner or later growth will be reduced. Beyond this point the effects are nonlinear (when compound interest spirals the debt). Salt uptake invariably continues after growth rate first declines, if for no reason other than it takes a lot of energy to keep the salt out, and so shoot salt contents rise, growth decreases more, and so on. This situation is commonly ‘catastrophic’ – a positive feedback – where increasing salt concentration causes more growth reduction and so more increase in salt concentration. The concept of catastrophic failure is important because in measurements made beyond this point, as they too often are, it is impossible to separate pathology from physiology. It also follows on that however good ion compartmentalisation is in the leaf, it is only one more layer of defence. Unless net transport to the shoot can be linked, by whatever means, to relative growth rate, the concentration in the shoot will rise and the plant will inevitably fail. There is some suggestion that the growth of some halophytes may be limited by ion transport. This was the interpretation of the ‘growth stimulation’ curve for S. maritima (Yeo and Flowers, 1986a). Salt cannot be transported to the shoot any faster than achieved at the optimal salinity and so beyond this concentration growth was limited by inadequate osmotic adjustment. Treatments (exposure to physiological concentrations of aluminium) that increased root surface area could increase growth even at the growth optimum, consistent with the possibility that growth was limited by ion transport capacity (Yeo and Flowers, 1977). If true, this would remove the need to postulate the feedback control on sodium transport in the root by sodium concentration in the shoot – something that demonstrably is never seen in glycophytes and something that would therefore constitute a qualitative difference between the two groups of plants. Nonetheless, regulation of salt transport in relation to growth emerges as a defining characteristic of the halophyte, though the details of this are not yet understood.

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Transport of chloride to the xylem

Two types of anion channel are found in the plasma membrane of xylem parenchyma cells: X-SLAC and X-QUAC (K¨ohler and Raschke, 2000). These resemble the kinetics of the S-type (slowly activating) and R-type (rapidly activating) channels of guard cell plasma membranes, respectively. The X-QUAC channels appear to dominate chloride transport from xylem parenchyma to xylem in physiological conditions (White and Broadley, 2001). There is a substantial literature that in species that are susceptible to chloride toxicity in saline conditions (particularly soybean, grapevine and citrus), salt tolerance resides with the ability to restrict chloride transport to the shoot. This has been demonstrated in cases of crops that are usually grafted, using ‘tolerant’ and ‘sensitive’ rootstocks; the tolerant rootstocks being those that restrict transport to the shoot. It is necessary to contrast the case for chloride with the case with sodium. For sodium, the root generally does not have the capacity to restrict transport to the shoot, and xylem loading with sodium is perhaps a feature of root homeostasis. In contrast to the array of potential entry pathways for sodium at saline concentrations, and the almost universal existence of a driving force for its passive entry, passive uptake pathways for chloride are much more limited and the existence of a driving force for passive chloride entry is also much more limited. It therefore appears that the root system can accommodate sufficient chloride to benefit salt tolerance of chloride-sensitive species, while retention in the root is only one component in relation to the tolerance of sodium.

14.11

Transport from shoot to root

There have been reports of phloem recirculation of sodium in glycophytes, which have sometimes been interpreted as a ‘tolerance mechanism’ and sometimes as a ‘pathological consequence’. Chloride is relatively mobile in the phloem, and the ratio of fluxes in xylem and phloem have been reported as around 5:1 (White and Broadley, 2001). Measurements in phloem sap suggest that more salt in the phloem sap is associated with salt-sensitive species (Wolf et al., 1991), which is consistent with a general principle of excluding salt from the phloem so that it is not directed to growing tissues (Munns et al., 2006). Neither sodium nor chloride is phloemimmobile, though retranslocation of both 22 Na and 36 Cl were much less than of 86 Rb (as a tracer for potassium) in the halophyte S. maritima (Yeo, 1981). The question is whether sodium is ‘deliberately’ loaded into the leaf phloem as a mechanism to deplete shoot sodium or whether more sodium leaks into the phloem in salt-sensitive plants and in salt-damaged leaves than in more tolerant plants. There is a conceptual difficulty with envisaging a significant role for phloem recirculation of sodium in effectively reducing the salt burden in the shoot. Recirculation would have to be combined with subsequent active efflux of that sodium from the roots, and this would have to be occurring in a situation in which efflux from the root is already inadequate to limit sodium uptake. Otherwise excessive sodium uptake and the need for phloem export would not have arisen.

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Mutation of AtHKT1 was regarded as demonstrating involvement in phloem loading of sodium for the purpose of recirculation since the mutant (sas) with defective AtHKT1 over-accumulated sodium in the shoot. It is reported that sodium transport to the shoot was unaffected by the mutation but that export in the phloem was reduced. However, it is difficult to evaluate this from the data. Calculated from the shoot weight and ion concentrations (Fig. 8 in Berthomieu et al., 2004), the quantity in the shoot and thus the net transport (i.e. xylem input–phloem retranslocation) was quite similar between the wild type and the mutant – at 25 mM NaCl and at 50 mM NaCl there was about half as much sodium in the shoots of the mutant plants. Therefore, the quantity of sodium transported to and remaining in the shoots was not increased by the mutation. The mutant plants certainly grew less well, but there is insufficient information to separate any effect from the background situation that ‘plants that grow poorly accumulate more salt’ and ‘plants that accumulate more salt grow poorly’ and the two go around in circles until the plant dies. In Fig. 6, Berthomieu et al. (2004) compare the concentrations in the phloem by comparing the sas mutant grown at 5 mM NaCl with the wild type grown at 50 mM NaCl. The disparity of treatment undermines any interpretation. Although the leaf sodium concentrations are comparable, the treatments are not equivalent for examining a hypothesis of phloem retranslocation. The wild-type plants are growing in 10 times the external NaCl. This will limit the potential to unload the phloem at the root and more importantly the potential to pump the sodium out of the root to the medium – upon which the strategy depends. This, rather than differences in phloem loading, is at least as likely an account for a higher concentration of sodium in the phloem of wild-type plants in this particular experiment. The sources may be similar in this experiment, but the sinks are certainly not. It is relevant that retranslocation of labelled sodium was observed only when sodium was removed from the solution around the roots (Blom-Zandstra et al., 1998): i.e. in a situation which would establish a source–sink gradient. There are also discrepancies concerning the pattern of expression of AtHKT1, which was reported to be localised in phloem tissue (Berthomieu, 2004) but was associated with sodium entry at the plasma membrane of roots (Rus et al., 2001).

14.12

Leaf cells

Salt that arrives in the xylem in the shoot needs to be accumulated by leaf cells. A major difference between monocotyledonous and dicotyledonous halophytes is that the former rarely develop succulence. Succulence has a ‘storage’ role, but this is not the same as storage of water in desert succulents or of malate in CAM plants: salt stored in the vacuoles of halophytes is not for subsequent release. Succulence in halophytes would appear to serve a role as a ‘safety valve’ – a role similar to that of salt glands – accommodating any imbalance between net transport and requirement for osmotic adjustment. Sodium-proton antiport is again the prime contender for loading the vacuole, energised by both ATPase and PPiase in the tonoplast. The demands of a continued, long-term compartmentalisation are as important as initial

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uptake into the vacuole. Minimising leakage out of the vacuole appears to be a major factor in the tolerance of the halophytes: the importance is easy to comprehend when it is considered that once having got the salt onto the vacuole it has to be kept there at an acceptable cost, for a whole season in the case of an annual and for longer in an evergreen perennial. The major storage cells of succulent halophytes such as S. maritima have a tiny cytoplasmic volume fraction and sparse mitochondria, making a prima facie case that they are unlikely to be expending large resources in fighting an on-going battle against leakage. Re-translocation studies (Yeo, 1981), membrane analysis (Leach et al., 1990) and studies of ion channel gating (Maathuis et al., 1992) all suggest that the tonoplast of S. maritima is fairly leak-proof towards sodium. Once sodium is accumulated in the vacuoles of leaf cells there is little export or recirculation. The number of channels in the S. maritima tonoplast is similar to that in glycophytes and their individual cation selectivity no better. However, though the potential leakage current was 4000 pA, the leakage current at physiological conditions was only 5–8 pA, which could be opposed by no more than 30% of the capacity of ATP-driven proton pumping (Maathuis et al., 1992). Compartmentalisation does not, in this case at least, appear to be about different channels, but of gating these channels closed and minimising leakage across the bilayer. Work on animal membranes has shown that cholesterol is a key inhibitor of sodium leakage through the membrane bilayer (Haines, 2001). It is therefore significant that in the tonoplast of S. maritima, planar lipids including cholesterol were present. Addition of cholesterol stabilised artificial lipid bilayers (Davenport and Tester, 2000). Cholesterol was a significant lipid in oat (Sandstrom and Cleland, 1989). The vacuolar proteome of arabidopsis contains a cholesterol acyltransferase (Carter et al., 2004). Different phytosterols have primary function as inhibitors of proton leakage through the membrane bilayer (Haines, 2001). The vacuole is large and can contain a great quantity of sodium, if it can be put there and kept there at an affordable cost. Whether this is achieved by minimising leakage of sodium from vacuole to cytoplasm or by a continual pump-and-leak, as at the plasma membrane of glycophyte roots, will clearly impact the cost of compartmentalisation. Intracellular compartmentalisation of sodium in halophytes was recognised before any of the possible transport pathways were characterised, or even known. Improving compartmentation of sodium in the vacuole remains a clear and sensible goal. Sodium compartmentalisation in the vacuole is initiated by sodium-proton antiport powered by a proton gradient established by proton ATPases and pyrophosphatases in the tonoplast (Blumwald et al., 2000). The membrane potential at the tonoplast would permit a two- or threefold accumulation of chloride in the vacuole relative to the cytoplasm, but there is a proton-chloride antiporter as well as two types of bidirectional chloride channel in the tonoplast (White and Broadley, 2001). Over-expression of the tonoplast sodium-proton antiporter has been attempted, presumably on the expectation that native activity is the weak link in the chain. This could be either because native activity is inadequate to match net sodium influx at the leaf plasma membrane and/or because it is inadequate to match leakage of sodium out of the vacuole. If it cannot match leakage, the question is posed whether

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it is antiport activity or leakage that would be the better addressed. It must also be remembered that although the focus tends to be on the sodium-proton antiport activity, the antiporter appears to have a vital role in cation homeostasis and tissue development in the absence of sodium. A T-DNA insertional mutant of AtNHX1 (the arabidopsis sodium/proton antiporter) also had reduced potassium/proton exchange and altered leaf development (leaf area and frequency of large epidermal cells were reduced) (Apse et al., 2003). The conclusion was that ‘as many genes encoding these antiporters have been cloned from salt sensitive plants, it is likely that they function in some capacity other than salinity tolerance’ (Apse et al., 2003). A timely caution that an intervention may not only change the most obvious, or the most hoped-for. It seems possible that proteins such as AtNHX1 are constitutive cation/proton exchangers that are able to exchange sodium in saline conditions. How this relates to the strange results of over-expressing AtNHX1 remains to be explained. Despite papers and reviews (eg Horie and Schroeder, 2004) supporting the over-expression of NHX transporters, the whole question bears serious investigation. The most dramatic result concerns the over-expession of the arabidopsis AtNHX1, coding for a vacuolar sodium-proton antiporter, in tomato (Zhang and Blumwald, 2001). At first sight it suggests that this may work in practice to increase the salt tolerance of tomato, but the evidence is confusing. The treatment regime also very strongly implies that any effect of over-expressing this gene is concerned with survival of osmotic shock and is not related to the tolerance of salinity. Transgenic plants grew and set fruit about as well in 200 mM NaCl as they, or wild-type plants, did at 5 mM NaCl. The wild-type plants are described as having died at 200 mM NaCl (there is a small photograph by way of evidence but no measurements were made before the plants died). Since the salt treatment consisted of applying 200 mM NaCl in a single step at 14 days, plants most likely died from the results of osmotic shock and not as a consequence of salinity (Flowers, 2004). The AtNHX1 was expressed in tonoplast-enriched vesicle preparations, and transgenic plants showed greater sodium-proton antiport activity in these preparations than did the wild-type plants (Zhang and Blumwald, 2001). However, the ion content data presented do not support the argument that transgenic plants survived and grew because they used sodium for osmotic adjustment and compartmentalised it within the vacuole. The potassium concentration in mature leaves of transgenic plants growing in 200 mM NaCl was about 4 mg/100 mg dry weight (the units used by the authors) lower than transgenic or wild-type plants growing in 5 mM NaCl (Fig. 3 in Zhang and Blumwald, 2001; this is a decline of about 1 mmol potassium g−1 dry mass). The corresponding increase in sodium in plants growing at 200 mM NaCl was about 0.8 mg/100 mg (about 0.35 mmol sodium g−1 dry mass). This increase in sodium concentration was fairly similar to the increase in chloride (Fig. 3 in Zhang and Blumwald, 2001). Consequently, the combined concentration of sodium + potassium decreased substantially when the external concentration was increased from 5 to 200 mM NaCl. Sodium partially (about one third) compensated for the loss of potassium, but the cation (sodium plus potassium) concentration in plants growing in 200 mM NaCl was still considerably lower than it was in plants growing in 5 mM

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NaCl. This is not consistent with sodium being used for osmotic adjustment. There is no data on plant water content (only for fruit) but the fresh weight of plants was not reduced by salinity, suggesting that there was no substantial concentration effect to compensate for the reduction in concentrations on a dry weight basis. The results (Zhang and Blumwald, 2001) imply that over-expressing AtNXH1 had some benefit in their experimental conditions, but leaves totally unexplained the bewildering situation of how the plants grew at 200 mM NaCl with a lower ion content that they had at 5 mM NaCl and provides no evidence that over-expressing the antiporter did anything to help the plant’s ion relations. Methodology also poses difficulty in working out what is happening in Brassica napus over-expressing AtNHX1 (Zhang et al., 2001). There are no ion measurements on both wild-type and transgenic plants at the same salinity (only at 10 mM NaCl) and so there is no evidence that transgenic plants had increased vacuolar sodium accumulation. Unlike tomato, sodium plus potassium content in leaves of transgenic B. napus at 200 mM NaCl was greater than at 10 mM NaCl: the increase in sodium more than compensated for the loss of potassium resulting in an increase in sodium + potassium of about 1.4 mmol g−1 dry mass, which would compensate for the increase in external salinity if the average water content of the plant was a little under 7 g water per gram dry mass (calculated from Fig. 2 in Zhang et al., 2001). The wild-type plants, from their appearance in the unscaled photograph, barely started to grow. The failure of the wild-type plants to grow is puzzling as B. napus is classed as a salt-tolerant crop (Tanji and Kielen, 2002) and should have produced appreciable growth. This suggests that they did not suffer from salinity, but from osmotic shock when given 200 mM NaCl instantaneously at 14 days – an outrageous treatment which has no basis in agriculture. This interpretation agrees with the view that ‘osmotic stress rather than ion-specific stress seems to activate NHX activity at the vacuole’ (Pardo et al., 2006). That the over-expression of AtNHX1 enabled plants to overcome osmotic shock says something very interesting about the roles of these exchangers and the far-reaching consequences of their alteration, but may not say much about vacuolar compartmentalisation of sodium and salt tolerance. Even in situations where there is some expectation that a simple intervention could lead to improvement in salt tolerance, there is no clear evidence that it can work. The published results provide anecdotal evidence that over-expression of AtNHX1 can overcome osmotic shock. It is not clear that lack of sodium/proton antiport capacity is the key weakness in relatively tolerant species such as tomato and brassica, because there are no comparable measurements on wild-type plants and on plants over-expressing the gene for the antiporter. Implications from sodium entry across the plasma membrane of glycophyte roots and retention of sodium in halophyte leaf vacuoles suggest that the antiporters may function mostly by attempting to counter leaks. It is also not clear that a protein that has constitutive effects on ion homeostasis and leaf development in the absence of salinity can be grossly manipulated with impunity. In tomato at least the sums do not add up, and the ‘salt-treated’ plants were growing without enough ions to explain the fact that they were alive.

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Expression of the AtNHX1 in wheat was reported both to increase grain yield in saline soil and to reduce the concentration of sodium in the leaf (Xue et al., 2004). Whatever the mechanism by which the expression of AtNHX1 may have improved yield, it would not appear to have been via the ‘expected’ mechanism. Increased compartmental ability could allow more sodium to be used for osmotic adjustment, or the same amount to be compartmentalised more effectively and/or more cheaply. But is it not readily apparent why increased compartmental ability would reduce the concentration of sodium in the leaf. For understandable reasons, it is the arabidopsis homologues that have been used. It is pointed out in a review (Tester and Davenport, 2003) that arabidopsis is somewhat anomalous as a glycophyte model in salinity, since there is poor relationship between sodium accumulation and salt sensitivity and mutations that increase salt sensitivity in arabidopsis do not increase sodium accumulation.

14.13

Prospects

Conventional breeding, whatever the criteria used, is a slow process. It needs the prospect of substantial increase in yield over current genotypes to make it worth trying to breed a new one. Because the investment may be a decade a more, the gulf between showing that a plant may grow a little better in an experiment and demonstrating that this will convert to a measured increase in the (say) grain production of a crop in the field, with all the vicissitudes of soil and climate, is an immense one. Even when successful salt tolerant varieties are produced, their utility is often limited to certain soil types. It is increasingly recognised that saline soils present a mixture of physical and chemical problems combined with soil structure and climate as well as nutritional inadequacies: all of which run counter to an idea of a ‘universal’ variety. The consequence of this is that any elite salt-tolerant line must be capable of back-crossing into a wide range of locally adapted cultivars and then be evaluated in the real world. There are enormous demands upon the stability, combining ability and lack of side effects of any genetic recombination, whether produced by conventional sexual hybridisation or by any other means. This new line must pass not a few laboratory measurements, but the demanding business of comparative yield trials over a range of sites and seasons. Nearly all ‘promising’ lines will be rejected at or before this stage, and any detrimental side effect will guarantee failure. Any or all of these considerations contribute to the limitations of getting molecular biology into the field: other than for high-value crops (Delmer, 2005). The value of selecting for salt tolerance as a strategy has even been questioned. It is not easy, to separate the effect of growth rate (or plant vigour) from specific mechanisms relating to ion transport. An extreme stance was taken (Richards, 1992) that it was better to concentrate on selection for vigour and ignore selection for salt tolerance per se. A difficulty encountered in selection for salt tolerance, at least in rice, is that, unfortunately, greatest vigour is associated with an undesirable plant type of low yield potential; the problems of trying to disentangle the contribution of

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vigour and ion transport traits in rice has been discussed at length elsewhere (Garcia et al., 1995). Most conventional breeding work with salt-sensitive species to date has accepted the arduous strategy of optimising variation in many ‘physiological traits’ that have small incremental benefits for, but probably have only incidental or accidental relationship with, salt tolerance (Yeo et al., 1990; Garcia et al., 1995; Koyama et al., 2001 and references and discussion therein). The use of ‘physiological criteria’ increases information and exploits diversity within a crop species (Yeo and Flowers, 1986b; Wang et al., 2006; Cuartero et al., 2006; Munns et al., 2006). Such pyramiding approaches are slow and tedious even with marker-assisted selection; there is enormous difficulty in establishing phenotyping techniques that are manageable on the required scale (Munns et al., 2006) and also are non-destructive (since in a breeding programme it is mandatory that the plant survives the screening procedure!). Crossing and selection can only operate upon such variation for salt tolerance as is present in the genome of the species. If that variation was never present in the species, or has been lost during human selection, perhaps over millennia, for performance under ‘good conditions’, then some other approach is needed. Either other genetic information must be introduced (possibly, re-introduced) or a more tolerant crop must be chosen. Conventional breeding can facilitate the introduction of new genes via wide crossing (inter-specific, sometimes inter-generic, hybridisation). This has been practised in relation to salinity most extensively in the Triticeae. Thinopyrum bessarabicum, a salt tolerant wild relative of wheat, was hybridised with a monosomic line of Triticum aestivum cv Chinese Spring (for details see Forster, 1994). The amphiploid was substantially more salt tolerant than the parent wheat cultivar (Gorham et al., 1986). Such work led to the description of the Kna1 locus, a potassium/sodium discrimination factor located in the wheat D genome (Gorham et al., 1987). Wide crossing is itself a complicated task, especially if the parents do not hybridise freely, requiring techniques such as embryo rescue. However, the underlying practical problem is that wide crossing implies the mixing all the genes from a wild genome, of no agricultural merit, with the breeder’s prize cultivar. Back-crossing to recover the gene(s) of interest combined with agricultural worth, and ‘washing out’ the rest, is a daunting task. Developing non-destructive assays for salt tolerance (ones much more sophisticated that pouring on salt and seeing what dies) is only the first challenge. Any approach that might help speed up or simplify progress in breeding for salt tolerance is earnestly sought – provided it can satisfy the same demanding criteria. Simple transfer of ‘tolerance genes’ has not as yet shown much success – even in what might be anticipated to be favourable circumstances. There are comparatively salt tolerant relatives of the cultivated tomato, but it has been difficult to use the genes of wild species because of the large number of genes involved, their small individual effect in comparison with environmental effect and the cost of recovering the qualities of the ‘commercial’ parent. Despite promising early observations from genetic transformation, this has not led to the development of a salt-tolerant cultivar of tomato (Cuartero et al., 2006). The stage at which transformants are assayed is also a serious question. Primary transformants (TG1) are subject to epigenetic

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effects, TG2 are segregant, and comparison be should between homozygous and azygous lines at TG3 against appropriate controls (Cuartero et al., 2006). Attempts at genetic modification of salt tolerance in higher plants can most kindly be described as having produced confusing results, which are sometimes due to the treatments which have been used. Bray (2004) demonstrated the different patterns of gene activation and repression that could be achieved by imposing ‘water stress’ in different ways (see Section 13.7). Flowers (2004) cites only a handful of papers (4 out of 68 published between 1993 and 2003) where there was adequate and realistic evaluation of plant salt tolerance following interventions. Great hopes have been, and continue to be, expressed for quantitative trait loci (QTLs) which, rather than necessarily being single genes, are regions of chromosome that have substantial effect on a trait and can, in terms of plant breeding, be regarded as ‘black boxes’ (it is not strictly necessary to know what the genes are or what they do – most of the enormous achievements to date of plant breeders have been achieved on the basis of phenotype and performance alone). To be of more use than single genes, it is still necessary that a QTL accounts for enough of the phenotypic variation to allow reasonable breeding progress to be made. For salinity, there are as yet few hopeful findings. Studies with arabidopsis found six QTL associated with germination and five with fresh weight production on NaCl, each explaining between 5.1 and 13.7% of the variance (Quesada et al., 2002). The growth measurements were made on agar +/– 50 mM NaCl. It must be assumed that transpiration was unrealistically low, which makes interpretation limited. QTLs have been measured for the ability to germinate in NaCl, which is certainly of ecological, though less often of agricultural relevance. The genetic controls underlying responses to salinity at germination are different from those at early seedling growth (Lindsay et al., 2004) and correspondence between tolerance at germination and at vegetative stages was inconsistent and sometimes back-to-front (Mano and Takeda, 1997; Foolad, 1999; Quesada et al., 2002; Lindsay et al., 2004). In rice, individual QTL for sodium uptake, concentration and sodium:potassium ratio explained between 6.4 and 9.6% of variation. One of the three QTL for potassium uptake did explain 19.6% (Koyama et al., 2001). QTL have also been identified in a cross between Nona Bokra and an elite japonica. QTL for shoot sodium and potassium concentrations (Lin et al., 2004) were reported to explain 48.5 and 40.1%. What is a possible difficulty here is something that has long undermined the use of land races such as Nona Bokra as simple donors. Their tolerance (resulting salt concentration) is intimately associated with their (undesirable) non-dwarfed genotype and so the relationship is somewhat spurious. The data (Fig. 1 from Lin et al., 2004) show that there was very little difference between the two parents in the quantity of sodium in the shoots, but there were large differences in concentration, and the lower concentrations in Nona Bokra would be accounted for by its greater vigour. Breeders at IRRI and elsewhere have been trying to use donors of this type since the early days of the 1970s, but with considerable difficulty. If any single factor potentially explained half the variation in sodium transport (not shoot concentration), the problem should have been solved a generation ago. Studies in barley revealed that seven QTLs related to salt tolerance are co-located with one of the dwarfing

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genes (Ellis et al., 2002). It has been possible, using a cross involving Nona Bokra, to locate a gene (SKC1) corresponding to a QTL for potassium homeostasis, and that the gene product may function as a sodium-selective transporter expressed in the xylem parenchyma (Ren et al., 2005), which could be a component trait. In barley, a total of 16 primary QTL were associated with vegetative salt tolerance, all located with a good degree of confidence, but in total accounting for only 35% of phenotypic variation. Overall, results ‘indicate that it is not easy to identify reliable loci for salt tolerance or any of its traits’ (Lindsay et al., 2004). Although barley is one of the most salt-tolerant crops, its field tolerance may be largely due to its vigour and early maturation rather than a preponderance of genetic information for physiological components of salt tolerance – the well-known Kna1 locus being apparently absent in barley (Munns et al., 2006). In terms of prospects for plant breeding, few of these QTL yet offer much to go on. One of the best results is for sodium exclusion in wheat, accounting for 38% of the variation (Lindsay et al., 2004). This locus (Nax1) maps to chromosome 2AL. This is not homoeologous with Kna1, which would map to chromosome 4AL (Lindsay et al., 2004). The gene or genes associated with the Kna1 locus are yet to be identified (Munns et al., 2006). In Durum wheat, the salt exclusion resided with limited xylem loading and increased re-absorption from the xylem, not with differences in unidirectional sodium influx (Davenport et al., 2005). Genetic analysis also showed that sodium concentration was controlled by two genes with major effect (Munns et al., 2003) corresponding to the Nax1 and Nax2 loci (Davenport et al., 2005). A further difficulty with salinity is the degree of environmental variation over place and time and hence the degree of G∗ E (genotype∗ environment) interaction. A large (30%) proportion of QTLs showed G∗ E interaction, a figure that is expected to increase as the range of target environments increases, as in comparing control and stress experiments (Cuartero et al., 2006). Where studies have been made of QTL∗ E and QTL∗ QTL interactions (Juenger et al., 2005), the contributions (at 19% and 12%, respectively) are larger than most individual QTLs.

14.14

Concluding remarks

The gulf between halophytes and glycophytes is vast. It has an almost ‘qualitative’ character in that, while all the component processes may well all be the same, they are put together in opposing ways. Changes that reduce the sensitivity of glycophytes run counter to changes that lead to the tolerance of halophytes. Reducing the sensitivity of glycophytes seems to be about keeping, or getting, salt out. The tolerance of halophytes is about co-ordinating getting salt in. This presents an enormous conceptual problem for finding any simple way of significantly improving the salt tolerance of salt-sensitive crops. There are clear possibilities at the extremes. For very salt-sensitive plants grown in low salinity with adequate fertilisation, it appears potentially possible to reduce pathways of inward leakage of sodium without destroying the mineral nutrition of the plant. How far this can go in practice depends on how vital the targets for

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manipulation are in other ways, and how intimately they are linked into regulatory systems operating at higher levels of organisation. The apparently independent evolution of salt tolerance in many families that are not closely related (Flowers et al., 1977) affords a little more hope for working with species that are more tolerant to salt. For species allied to halophytes, it appears potentially possible to improve key features such as compartmentalisation in the leaves. Again, how well this will work in practice depends on the knock-on effects of boat-rocking for which there are already many examples of just how far-reaching and confounding this can be. From a plant physiologist’s viewpoint this is all fine – halophytes and glycophytes behave in different ways and we begin to have some understanding of how they differ (and eventually, maybe, of how it is all regulated). From an agriculturalists point of view this is worrying. Taking any crop from the ‘middle ground’ which is neither very sensitive nor very tolerant there is a dilemma. All the pointers are towards reducing salt uptake, even treating compartmentalisation as just one more way of coping with too much salt. The end point could be to have a glycophyte expending energy on sodium extrusion. On top of this, it expends energy on xylem loading. It may then have to step in with more energy for recovery of sodium from the xylem, moving sodium about in the phloem, and even more energy to pump sodium out again. It must also expend energy to accumulate more potassium for osmotic adjustment (or synthesise something else) to make up for the sodium it excludes. Every step along this path takes the plant further from being a halophyte. But halophytes are not just quaint oddities; they are extremely successful in their niche. What halophytes do instead looks remarkably efficient. It seems at the moment that halophytes do not present a ‘quick fix’ – salt tolerance gene(s) to be cherry-picked out and put into glycophytes. Halophytes must, however, hold the clues to how sodium transport is co-ordinated, because they demonstrably do this so well. All the questions that might be raised concerning the co-ordination of sodium transport are questions that halophytes have already solved; only we do not yet know many of their answers.

References Aharon, R., Shahak, Y., Wininger, S., Benzov, R., Kapulnik, Y. and Galili, G. (2003) Overexpression of a plasma membrane aquaporin in transgenic tobacco improves plant vigor under favourable growth conditions but not under drought or salt stress. The Plant Cell 15, 439–447. Amtmann, A., Fischer, M., Marsh, E.L., Stefanovic, A., Sanders, D. and Schachtman, D.P. (2001) The wheat cDNA LCT1 generates hypersensitivity to sodium in a salt-sensitive yeast strain. Plant Physiology 126, 1061–1071. Amtmann, A. and Sanders, D. (1999) Mechanisms of Na+ uptake by plant cells. Advances in Botanical Research 29, 75–112. Apse, M.P., Sottosanto, J.B. and Blumwald, E. (2003) Vacuolar cation/H+ exchange, ion homeostasis, and leaf development are altered in a T-DNA insertional mutant of AtNHX1, the Arabidopsis vacuolar Na+ /H+ antiporter. The Plant Journal 36, 229–239. Berthomieu, P., Conejero, G., Nublat, A., et al. (2004) Functional analysis of AtHKT1 in Arabidopsis shows that Na+ recirculation by the phloem is crucial for salt tolerance. EMBO Journal 22, 2004–2014.

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15 Desiccation tolerance Anthony Yeo

15.1

Introduction

Chapters 13 and 14 have dealt with situations that can be described as desiccation avoidance. This comprises various strategies (ecological, physiological, structural and biochemical) aimed at maintaining water uptake and/or minimising water loss as plant water status decreases. Desiccation avoidance was crucial in coping with water deficit however this was imposed – whether by lack of water supply, by salinity, heat, freezing, or by any process or combination of processes that lead directly or indirectly to tissue dehydration. The majority of that which is usually described as ‘drought tolerance’ is concerned with desiccation avoidance, that is, maintaining the presence of at least some bulk water (Hoekstra et al., 2001). In agricultural terms the caveat is not only to survive but to maintain productivity in water-limited environments, which means sustaining water use as far as is possible. The role of solutes in drought tolerance was principally in bulk osmotic adjustment and compartmental osmotic adjustment (involving compatible solutes and osmoprotectants). Solutes also had a central role in the photosynthetic adaptations of C 4 photosynthesis (malate shuttle) and in crassulacean acid metabolism (malate storage). Desiccation tolerance is concerned with situations in which water content falls to the point at which there is no longer any bulk cytoplasmic water: this occurs at a water content of around 23% or around 0.3 g water g−1 dry mass (Hoekstra et al., 2001). At this point the rules change qualitatively. The hydrophobic effect, which is a major factor in the assembly of biomembranes and in the maintenance of protein conformation, ceases to exist. It is no longer a question of prioritising water allocation in more and more vital places for more and more vital processes, but of surviving without water. This means maintaining structural integrity without the services that water provides, and at this critical point, the services that water provides in maintaining the spacing between and within macromolecules. In addition to surviving in the desiccated state, desiccation tolerance includes the ability to pass down through the stages of decreasing water availability, which is described as drought tolerance, and implies the ability to successfully rehydrate after a period of desiccation (Vertucci and Farrant, 1995). Both passages include stages at which metabolism runs, but runs ineffectually, typically with poor coupling between processes, giving rise to unusual intracellular conditions and the generation of reactive oxygen species likely to cause oxidative damage. A whole raft of interconnected processes are put in place during seed maturation or during the development (in angiosperms) of vegetative desiccation, in

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order to allow tolerance of the desiccated state as well as the passage into and out of it. Solutes and solute transport play key roles in the cellular and subcellular adaptation to desiccation, and particularly notable are the roles of sugars and soluble late-embryogenesis-abundant (LEA) proteins. Desiccation tolerance is of enormous importance in seed and pollen production. There is, however, very limited adoption of desiccation tolerance in the vegetative tissues of vascular plants. This could be due to there being very few niches in which desiccation tolerance confers a competitive advantage and/or because tolerance fails at higher (tissue and organism) levels of organisation before the cellular-based processes ever come into play.

15.2

Occurrence of desiccation tolerance

Desiccation tolerance occurs in two situations. In nearly all species, desiccation tolerance is achieved at some point during reproduction, since spores, pollen and embryos usually undergo dehydration. Less commonly, desiccation tolerance (anhydrobiosis) of vegetative tissues occurs in a range of organisms. Desiccation tolerance is common in cryptograms, including the bryophytes and lichens, and is thought here to be of early evolutionary origin. In vascular plants desiccation tolerance is found as a rare ecological adaptation. At present about 350 species of desiccation-tolerant vascular plants are recognised (less than 0.2% of the total number of species) with a phylogenic distribution that is wide, but thin and uneven (Proctor and Tuba, 2002). Proctor and Tuba (2002) contrast the strategies adopted by bryophytes and vascular plants. The bryophytes tend to poikilohydry in that their hydration responds simply to the relative humidity of the environment, passing rapidly through wetting/drying cycles, being either fully turgid or desiccated, and suspending metabolism when they are too dry. Vascular plants on the other hand tend to homiohydry – uptake and transport of water from the soil, combined with techniques to adjust water loss, leading to relative stability of leaf water content. There is some flexibility of response within the homiohydric group – some vascular species achieving desiccation tolerance (Proctor and Tuba, 2002). All desiccation-tolerant (anhydrobiotic) vascular plants are angiosperms; there are no gymnosperms (Bohnert, 2000). The expression of desiccation tolerance in vegetative tissue of vascular plants, although rare, has (re-) evolved independently in the genus Selaginella, in ferns and at least eight times in the angiosperms (Oliver et al., 2000). If this is taken to suggest that the capacity for the development of desiccation tolerance is widely available then the rarity with which it is found may indicate that the development of tolerance in vegetative tissue is subject to enormous constraints; this is considered later in the chapter.

15.3

Desiccation tolerance in seeds

Whether in seed maturation or in whole vascular plants, desiccation tolerance is a progressively coordinated programme of many responses, which is switched on by dehydration or by abscisic acid (ABA). In non-vascular plants desiccation tolerance

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is relatively common, is constitutive and can be expressed very quickly. The desiccation tolerance that is characteristic of seeds and vascular plants is progressive in nature (and cannot be expressed very quickly). Similar to processes such as ‘hardening off’ in response to falling temperatures, the development of desiccation tolerance is a relatively slow process. It involves a major, long-term commitment, which includes a complete programmed metabolic shutdown, cannot be instantly reversed and must be initiated before the plant water content falls to desiccation levels. In seed formation, desiccation tolerance is developed before maturation drying (Hoekstra et al., 2001). This is programmed action, and not response mode. The desiccation process is most extensively studied in seed maturation, and this will be considered first since the principles are, for the most part, common to the development of desiccation tolerance in vegetative tissues of vascular plants. ‘Orthodox seeds’ (Roberts, 1973) are seeds that can be dried; they are desiccation tolerant. They acquire tolerance to desiccation during the process of development and retain this tolerance for an ecologically significant period of time (one to many seasons), which may be greatly extended (perhaps to centuries) by storage in the defined conditions of a seed bank. All orthodox seeds can withstand dehydration to about 5% water g−1 dry mass. Any seed that does not behave in this way is not ‘orthodox’. ‘Non-orthodox’ seed has been described as ‘recalcitrant’ (Roberts, 1973) and ‘intermediate’ (Ellis et al., 1990) according to their behaviour in storage. Recalcitrant seeds do not develop tolerance to desiccation, and consequently cannot usually survive long before germination. A simple separation of seeds into orthodox and recalcitrant has been viewed as an oversimplification, and a continuum is seen, spanning a range from those that are fully desiccation tolerant to those that undergo little if any maturation drying. and remain desiccation sensitive both during development and after they are shed (Berjak and Pammenter, 2002). That non-orthodox seeds are not equally sensitive to desiccation implies that one or more of the processes comprising desiccation tolerance are, and to a variable extent, undeveloped or unexpressed. Identification of which seed is or is not recalcitrant is vital for seed conservation: direct determination is a daunting prospect but some progress is being made with probabilistic models with woody species (Daws et al., 2006). Drying is a progressive process. It follows that the early stages of dehydration must be successfully negotiated for the latter stages of the process to be relevant. The sources of damage and the methods used to tolerate them differ completely at different stages of dehydration. Damage falls broadly into four categories, which are sequential in imposition (Pammenter and Berjak, 1999; Berjak and Pammenter, 2002): 1. Mechanical damage – due to reduction in cell volume; 2. Metabolic damage – these are degradative processes occurring in aqueous solution at intermediate water contents and are consequent upon deranged metabolism; 3. Desiccation damage (in the strict sense, acquired during desiccation and not during drying out); 4. Rehydration damage.

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Recalcitrant seed that is artificially dried (e.g. by embryo excision and rapid drying techniques) may suffer rehydration damage – but it is difficult to ascribe relevance to this, as the seed would naturally have died long before reaching stage 4 (rehydration damage). Specifically, the process of developing desiccation tolerance includes (Pammenter and Berjak, 1999; Berjak and Pammenter, 2002): 1. intracellular physical characteristics; 2. intracellular de-differentiation, resulting in the minimisation of surface area of membranes and probably also of the cytoskeleton; 3. ‘switching-off’ the metabolism; 4. presence, and efficient operation, of antioxidant systems; 5. accumulation of putatively protective molecules (including LEAs, sucrose, etc.); 6. deployment of certain amphiphilic molecules (molecules that contain both hydrophobic and hydrophilic regions); 7. an effective peripheral oleosin layer around lipid bodies; 8. presence and operation of repair mechanisms during rehydration.

15.3.1

Intracellular physical characteristics

A large vacuolar volume fraction can be associated with mechanical damage (Farrant et al., 1992). There is a negative correlation between vacuolar volume and desiccation tolerance. With tolerance, there is a reduction in the volume fraction of vacuoles in the embryonic axis and an accumulation of insoluble material in the vacuoles of cotyledonary cells. In the examples used (Farrant et al., 1992, 1997) it is perhaps significant that the ‘sensitive’ species (the halophyte Avicennia marina) is one that has a compelling need for large vacuoles to contain dissolved salts, compromising the freedom to reduce the volume fraction of vacuoles. In tolerant species the cells shrink, but they do not collapse; tolerance must include the ability of the cell walls to shrink and fold (concertina) without strain, and for the cell wall-to-plasma membrane contact to be preserved (Proctor and Tuba, 2002). Many complex anatomical adaptations are associated with this, but overall, anatomy alone is not predictive of desiccation tolerance (Berjak and Pammenter, 2002; Proctor and Tuba, 2002). The failure of some recalcitrant seeds to reassemble the cytoskeleton on rehydration (Mycock et al., 2000) does imply that this is another necessary condition for tolerance and must be satisfied in orthodox seeds. The cytoskeleton is needed for organisation and structural support and to act as a scaffold for complex enzymatic processes. Although there is little information on how DNA, chromatin and nuclear architecture are preserved during desiccation (Proctor and Tuba, 2002) it is implicit that their stability, or stability plus repair mechanisms, are a necessary prerequisite for tolerance.

15.3.2

Intracellular de-differentiation

The minimisation and simplification of internal structures during seed maturation is described as ‘de-differentiation’, a process that is reversed upon rehydration in

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orthodox seeds (Bewley, 1979). Both the volume fraction of mitochondria and the complexity of their cristae are negatively correlated with desiccation tolerance: in orthodox seeds mitochondria de-differentiate. Importantly, they do so prior to the initiation of maturation drying (Farrant et al., 1997), implying that de-differentiation is a prerequisite of, rather than a consequence of, maturation drying.

15.3.3

‘Switching-off’ metabolism

Decreasing tissue water content is associated with disruption of metabolism – ion and metabolite concentrations increase, electrical potentials are affected and compartmentation is compromised, and so on. In the range from 0.45 to 0.25 g of water g−1 dry mass, the generation and action of free radicals is a consequence of the metabolic derangement (Vertucci and Farrant, 1995). Seeds that are to become desiccation tolerant must pass down through this stage intact. This is achieved by running down metabolism in general and by protection against metabolic damage. A decline in respiratory substrates precedes (causes?) the reduction in metabolism – again emphasising the sequential nature of the process. Respiratory substrates, metabolism and mitochondrial number/structure all decline prior to desiccation. At the same time there is an accumulation of non-reducing sugars. A coordinated control of energy metabolism at the onset of dehydration seems to be an essential postulate (Hoekstra et al., 2001). There is also a link with progress through the cell cycle, orthodox seeds holding at G1, with variation observed in the state of the cell cycle in recalcitrant seeds, depending upon whether or not there was any dormancy (Berjak and Pammenter, 2002).

15.3.4

Antioxidant systems

Orthodox seeds will pass through the ‘danger’ water content (0.45 to 0.25 g water g−1 dry mass range) in which the first phase of free-radical production is most likely to occur (Vertucci and Farrant, 1995), requiring effective antioxidant systems both during dehydration and upon rehydration. Water deficit, however imposed, can of itself impair the function of enzymatically based antioxidant systems, amplifying the damage caused by free-radical generation. The enzymatic systems (for example ascorbate peroxidase, glutathione reductase and superoxide dismutase) function only while the cytoplasm is still in a hydrated state. In desiccated conditions, only molecular antioxidants will work: examples include amphiphilic molecules such as tocopherol, quinones, flavenoids and phenolics. Lipid peroxidation is a common consequence of oxidative damage and has been observed in temperate recalcitrant species (Berjak and Pammenter, 2002). Metabolic damage therefore includes the production of free radicals and the ineffectiveness of antioxidant systems to deal with them. To limit metabolic damage, metabolism is ‘switched off’ – the pools of respiratory substrates are reduced, preceding, and presumed to be causal in, a reduction in respiration rate (Rogerson and Matthews, 1997). This furthermore allows the sugars to be converted to low-molecular-weight carbohydrates which have protective roles during dehydration and desiccation.

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Protective molecules

Protective solute molecules fall into three groups: 1. Compatible solutes (functional while free water is present). These are preferentially excluded from the protein surface and structural integrity is still maintained by keeping the surface preferentially hydrated; 2. LEAs (late-embryogenesis-abundant proteins); 3. Low-molecular-weight carbohydrates including non-reducing sugars and cyclitols (with the major role in maintaining stability in desiccated conditions). Sugars are the only solutes that can confer structural preservation below the desiccation limit (0.3 g water g−1 dry mass; Hoekstra et al., 2001). This may be by ‘water replacement’ and/or by their contribution to glass phases (see below). LEAs (dehydrin-type proteins) are a set of hydrophilic, heat-resistant proteins whose accumulation coincides with the acquisition of desiccation tolerance in orthodox seeds, pollen and anhydrobiotic plants. However, LEAs are also induced by other stresses, and homologues are found in organisms other than plants; although the proposed functions are largely intuitive, there is evidence of some effect on stress tolerance (Wise and Tunnacliffe, 2004). LEAs have a very high solubility in water, are very stable and often remain soluble after boiling (Bartels and Salamini, 2001). Group 2 and Group 3 LEA proteins include α-helical structures and, on a larger scale, filaments (Wise, 2003). A bioinformatic analysis supported localisation of some LEAs in the nucleus and associated with the cytoskeleton, and possibly a role in DNA binding (Wise, 2003). At least the major groups of LEA proteins are ‘natively unfolded’ or ‘intrinsically disordered’, but this does not imply lack of function. There are three ways in which such unfolded or disordered proteins can have a physiological function (Wise and Tunnacliffe, 2004): 1. They have defined ‘partner molecules’ upon binding to which folding occurs (true for most unfolded proteins but not yet identified for LEAs); 2. Folding can be induced by abiotic stress (there is some evidence for this – an unusual situation, since protein dehydration results usually in loss of structure, or in aggregation). Coiled coil and superhelical filament formation bears similarities to other proteins that form part of the cytoskeleton; 3. They may exert their function in the unstructured state (which could be consistent with a role in water replacement or as a ‘molecular shield’ between partially denatured proteins). LEAs are not restricted to the desiccation phase, and are found in recalcitrant seeds and in various tissues subjected to any stresses that result in water and/or temperature stress. Small HSPs (heat-shock proteins) have also been shown to coincide with the acquisition of desiccation tolerance and are currently accorded an overall protective effect (Hoekstra et al., 2001). The ability to express LEAs is thus not indicative of the ability to develop desiccation tolerance; this may be a necessary condition, but this is a further indicator that no single character is sufficient.

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The accumulation of non-reducing sugars has been heavily implicated in the development of desiccation tolerance. Early work (Steponkus, 1971) on frost hardening in ivy suggested that protein changes occurred first to allow sucrose to replace the water of hydration. There is no doubt that the development and maintenance of the desiccated state in orthodox seeds is linked to the accumulation of large quantities of sucrose and other oligosaccharides. The role of sugars in desiccation-tolerant seeds is ascribed to the ‘water-replacement hypothesis’ (Clegg, 1986; Crowe et al., 1986; Allison et al., 1999) and to vitrification (Koster and Leopold, 1988; Crowe et al., 1996). According to the water-replacement hypothesis, as the water shell around proteins is removed on desiccation, sugars are able to replace water. Sugars take over ‘hydration’ roles in proteins, satisfying the hydrogen-bonding needs of polar groups on the surface of the molecule. This protects individual proteins from aggregating together and also contributes to balancing the conformational entropy of the protein, a role that becomes increasingly important with the loss, on desiccation, of the hydrophobic effect. Such protein–sugar associations are very stable, and proteins stabilised in this way have been shown to survive heating to 150◦ C, and to retain their native secondary structures long after the seeds themselves have died. Because of the large number of polar residues that they possess, some LEAs are also thought to act as ‘replacement water’ (Cuming, 1999) by providing a layer of hydroxylated residues to interact with the surface hydrophilic groups of other proteins. In membranes, the final stages of dehydration result in the removal of water molecules associated with the polar head groups. This reduces the distance between these groups and thus also increases the packing of the hydrophobic chains in the membrane core, dramatically increasing the (gel to liquid crystal) phase transition temperature and effectively ‘gelling’ the membrane at normal temperatures. This increase in transition temperature is prevented by interaction between sugars and polar head groups thus maintaining the spacing and packing (Crowe et al., 1987). This also prevents lateral phase separation on drying which would cause damaging membrane leakage upon rehydration. The transition temperature in dry anhydrobiotic seeds and pollen is little different from that in the hydrated state, and this similarity is attributed to the abundant sugars (Hoekstra and Golovina, 1999). Human fibroblasts that had been engineered to accumulate trehalose were found to have thereby acquired a degree of desiccation tolerance (Guo, 2000). However, the lifespan of the desiccated cells was limited – one more indication that a single factor (sugar accumulation), though perhaps necessary, was not sufficient. At the desiccation point (0.3 g water g−1 dry mass) the molecular motility of the cytoplasm decreases 100 000-fold and at about 0.1 g water g−1 dry mass the cytoplasm vitrifies (enters a glassy state). Sucrose and certain oligosaccharides or galactosyl cyclitols form supersaturated solutions (Berjak and Pammenter, 2002). The development of glassy states is considered to ‘freeze’ the cytoplasm into a state of immobility, which protects macromolecules and minimises or prevents phase transitions in membranes (Leopold et al., 1994; Berjak and Pammenter, 2002). Vitrification is an amorphous metastable state but one that maintains the disorder and physical properties of a liquid. Carbohydrates are good glass formers and achieve

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this through hydrogen bonding at appropriate temperatures and water contents. Intracellular glasses have been detected in dried seeds and in pollen. Recalcitrant seeds can still vitrify if they are air-dried to low-enough water contents – though this can be of no survival value, since the seeds will have long since died at much higher water contents than the glass-phase transition (Buitink et al., 1996; Hoekstra et al., 2001). Yet again, there is evidence that no single attribute confers desiccation tolerance. The function of glasses is considered to be the preservation of stability, their enormously high viscosity preventing crystallisation of embedded chemical compounds, fusion of macromolecules and conformational change. The tissue needs to possess a large measure of desiccation tolerance to survive to the water content at which vitrification takes place. Sucrose-only glasses are not the most stable and the stability of cytoplasmic glasses is considered to be due to a mixture of sugars with higher molecular weight polysaccharides and other polymers, LEAs being possible candidates for a role in glass formation (Hoekstra et al., 2001). Storage longevity is inversely proportional to molecular mobility in the cytoplasm. The relationship allows the prediction of longevity even in archival storage where experimental determination is difficult. Both LEAs and sugars have also been implicated in protective functions during the dehydration phase as well as in survival in the desiccated state. A function of LEA/sugar complexes during dehydration may account for the observations of the production of sucrose and oligosaccharides in some recalcitrant seeds.

15.3.6

Amphiphilic molecules

As the water content of the cytoplasm decreases during dehydration the concentration of solutes increases. For amphiphilic molecules that have both hydrophobic and hydrophilic regions this leads to a change in their partitioning between aqueous and lipid phases within the cell, with an increased partitioning into lipid membranes. This can have both negative (Hoekstra et al., 2001) and positive (Berjak and Pammenter, 2002) effects. Partitioning-induced perturbations may impair the electron transport chain and increase the production of reactive oxygen species. Positive effects may include the facilitated insertion of amphiphilic antioxidants and phospholipase inhibitors into the membranes, which may result in increased fluidity and thus counter the phase transition to the gel state (Golovina et al., 1998). Berjak and Pammenter (2002) also review the evidence that reversible partitioning of amphiphilic molecules into the membrane serves a protective function through lowering the water content at which the gel-phase transition takes place. The reversal, the repartitioning of amphiphilic molecules back into the cytoplasm as it becomes aqueous again upon rehydration, is suggested to account for the transient leakage through membranes, which is usually seen when pollen and desiccation-tolerant seeds are rehydrated (Golovina et al., 1998). This is not the same as the prolonged leakage found in imbibition injury, which is observed even in desiccation-tolerant material if it is wetted too quickly. In the cases of extremely dry tissue that is rewetted rapidly, particularly at low temperatures, membranes disrupt prior to any uptake of liquid water because there is melting of gel-phase lipids. This is avoided by slow pre-hydration, such as

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where the seed testa intervenes to limit the movement of water, or by vapour-phase equilibration.

15.3.7

Oleosins

Oleosins are a unique class of proteins that have a domain that can interact with the hydrophobic surface of a lipid droplet, as well as a hydrophilic domain that can interact with the aqueous phase of the cytoplasm. This protein boundary allows the presence of discrete lipid droplets in the aqueous cytoplasm and is suggested to prevent their coalescence during dehydration (Leprince et al., 1998). Such coalescence is a frequent characteristic of seed deterioration, including deterioration of orthodox seeds in storage.

15.3.8

Damage repair

Some or all of the mechanisms thus far described may serve in damage limitation but may not achieve damage prevention: damage repair is also a necessity. There can be no ‘active’ damage limitation/repair in the desiccated state, only ‘passive’ methods of damage limitation such as glass phases and non-enzymatic antioxidants. The damage sustained during the final stages of dehydration and during desiccation cannot usually be repaired in the desiccated state, because such repairs necessitate the functioning of enzyme systems that can function only when hydrated. This includes replacement of damaged RNA and repair to the DNA and protein-synthesising systems. The generation of free radicals appears to continue in air-dried orthodox seeds. On rehydration there is a further period of danger as metabolism comes back in but is not yet fully integrated, leading to another phase of reactive oxygen production until the fluxes through electron transport chains and linked metabolic systems come into balance. Damage repair will be needed rapidly upon rehydration since this phase may itself be rapid; thus, it follows that the repair mechanisms must usually be put in place before desiccation. Berjak and Pammenter (2002) summarise that repair systems may be present in recalcitrant seeds but that the mechanisms themselves are also desiccation sensitive and so are unable to be at all effective if the seed is dried too far. The array of processes involved in dehydration, desiccation tolerance and rehydration are summarised in Figure 15.1.

15.4

Vegetative tissues

Desiccation-tolerant vascular plants, exemplified by the resurrection plants, such as Craterostigma species, are able to survive desiccation for periods of up to 1 or 2 years (Scott, 2000). Tolerance is phylogenically widespread and there is some overlap (co-tolerance) with other stresses (e.g. salinity).

Tissue dehydration

Mechanical

Reduce vacuolation by separation into smaller vacuoles and reduce vacuolar volume fraction by accumulation of insolubles.

Xylem

Cavitates as tissue dehydrates.

Systems are in place. Protect during dehydration. Non-enzymatic systems replace enzymatic systems in desiccated state.

Antioxidant

Constitutive mechanical properties allow the tissue to shrink. Limitations of scale.

Protective Sucrose, etc., produced from storage CHO and photosynthesis. LEAs produced. Contribute to ‘water replacement’ and glass-phase formation.

Respiratory substrates converted to sucrose, etc. Programmed shutdown. Chlorophyll may be degraded. Anthocyanins may be produced.

‘Prediction’

Metabolic

Coordinated response

Gene regulation

Signalling

Detection

Desiccation tolerance needed

Molecules with both hydrophobic and hydrophilic regions partition into membranes as their concentration in the aqueous phase increases.

Amphiphile

Repair may be active. Any systems needed during rehydration must be synthesised now.

Repair

The preparatory stages, unless constitutive, must be completed between detection and the tissue water content falling to the point at which they are needed, and where necessary, in the right sequence.

Damage limitation: reduce water loss at expense of growth (drought resistance, xeromorphic adaptation)

Avoidance/adaptation: maintain transpiration (rooting properties, C4 WUE)

Mobilise insolubles. Restore vacuole volume fraction and coalesce vacuoles. Expand tissue.

Mechanical

Protective Glass-phase transition. Sucrose redeployed. LEAs decline.

Metabolic Chlorophyll restored. Sucrose converted to storage CHO and respiratory substrates restored. Restore metabolism.

Amphiphile Molecules with both hydrophobic and hydrophilic regions repartition back into aqueous phase.

Return through stages of low and increasing content to the hydrated state

Enzymatic systems regain importance. Protection very important as metabolism comes back in.

Antioxidant

Repair of damage sustained during desiccation and rehydration.

Repair

Figure 15.1 The processes involved in desiccation and rehydration of the vegetative tissue of a vascular plant. A schematic representation of the sequence of events involved in the acquisition of desiccation tolerance, survival of the desiccated state and return to hydration. The scheme is presented for vegetative tissue, but most of the processes are applicable to dehydration and rehydration of orthodox seed. (WUE, water-use efficiency; CHO, carbohydrates; ROS, reactive oxygen species)

Vessels refilled.

Xylem

Coordinated reversal of dehydration process

Rehydration

Survive desiccation

Tissue dehydrates towards the desiccated state in which proteins and membranes are protected by sugars, LEAs and glass formation, and metabolism is more or less at a standstill. Non-enzymatic antioxidants protect against ROS. Any repair mechanisms needed for the rehydration phase are in place.

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PLANT SOLUTE TRANSPORT

Gene expression

A general (though far from perfect) separation can be made between homiohydry (typical of vascular plants) and poikilohydry (typical of crytograms – including algae, lichens and bryophytes), and this is reflected in the pattern of gene expression of the two groups. In vascular plants that survive desiccation if it is imposed slowly (e.g. Craterostigma species), nearly all observed changes in gene expression occur during dehydration, with many dehydration-specific but few rehydration-specific gene products (Bernacchia et al., 1996; Bartels and Salamini, 2001). Craterostigma plantagineum survives desiccation provided that the rate of drying is slow enough to permit a programme of gene expression to come into play and adapt the plant to desiccation. The ability to survive desiccation has to be developed: it is not ever-present, and is not acquired if plants dry too rapidly (Bartels and Salamini, 2001). This contrasts with bryophytes, for example Tortula ruralis, which survives rapid desiccation often in less than 1 h, and in which tolerance is based upon rehydration-induced cellular repair (Oliver and Bewley, 1997). The bryophytes have an evolutionarily earlier mechanism of tolerance, being prepared for desiccation at all times, expressing, largely constitutively, proteins whose focus is damage repair upon rehydration (Bohnert, 2000). The ability to re-establish ‘safe’ conditions rapidly is exemplified in comparison of lichens of differing tolerance to desiccation. Absolute concentrations of antioxidants did not necessarily correlate with adaptation to desiccation, but it was concluded that the ability to re-establish the species-specific normal concentration of antioxidants, rapidly during rehydration, was a characteristic of well-adapted species (Kranner et al., 2003). Any changes in gene expression are within the early hours of rehydration (Wood and Oliver, 1999). The differences between homiohydric and poikilohydric plants and between the mechanisms adopted by vascular plants and by bryophytes are general, but not exclusive. Craterostigma wilmsii that was dried slowly required no new transcription or translation during rehydration to achieve recovery. However, plants that had been dried rapidly required induction of repair mechanisms during rehydration in order to survive; they died if mRNA, or protein, synthesis was inhibited (Cooper and Farrant, 2002). This indicates that the repair mechanisms in the resurrection plant were not constitutively present and were normally put in place prior to desiccation, though they could be initiated upon rehydration.

15.4.2

Physical characteristics

The major physical attribute of desiccation tolerance in the more complex plants is shrinkage, to perhaps 15% of the original leaf area in the case of C. plantagineum (Scott, 2000). Considerable tissue variation in the degree of shrinkage is possible. The shrinkage of roots may be constrained by their intimate association with a relatively rigid soil matrix. The shrinkage of woody species is limited by the characteristics of thickened tissue, which is relatively rigid. Folding of cell walls is the most obvious structural response in the resurrection plant C. wilmsii (Cooper and Farrant, 2002) together with substantial subdivision of vacuoles.

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The other mechanical consideration is one of scale. There is a scale constraint to the development of desiccation tolerance because rehydration requires that the xylem be recharged, generally by capillary action and root pressure (Sherwin et al., 1998). In the shrub Myrothamnus flabellifolia a complex system of lipid wall linings and lipid inclusions are reported to both facilitate water movement (by surface tension gradients and interfacial flow) and control the spatial pattern of re-filling (Wagner et al., 2000; Schneider et al., 2003). In addition to the complex programme for achieving tolerance of its tissues, a further array of highly specific adaptations is needed to permit a woody species to join the select group of desiccation-tolerant vascular plants.

15.4.3

Metabolism and antioxidants

The characteristics of vegetative tolerance in vascular plants echo those of seed tolerance, and both differ markedly from the vegetative tolerance of bryophytes in being both progressive and pre-emptive. There is a coordinated shutdown of metabolism. The indispensable role of the antioxidant system is also emphasised in vegetative tolerance of the desiccation/rehydration process. Angiosperm resurrection plants are either homio- or poikilo-chlorophyllous according to whether they retain or lose their chlorophyll prior to desiccation (Scott, 2000). Loss of chlorophyll limits the production of reactive oxygen species (photo-oxidation occurs when photosynthetic electron transport runs in water-stressed conditions without proper coupling with the metabolic turnover to utilise it). Anthocyanins are synthesised in some cases and accessory pigments can help protect against damage from ultraviolet radiation. M. flabellifolia retains chlorophyll in the desiccated state, and both desiccation and rewatering triggered increases in antioxidant capacity, whose protection began to fail only after 8 months of desiccation (Kranner et al., 2002). In lichens, the symbiotic association has been found to underlie the tolerance of reactive oxygen species. In Cladonia vulcani, the algal and fungal symbionts in isolation suffered oxidative damage during desiccation. But in symbiosis, tolerance was orders of magnitude more effective; each component appearing able to up-regulate the protective systems of the other (Kranner et al., 2005).

15.4.4

Low-molecular-weight carbohydrates

As with seeds, a key characteristic of desiccation tolerance in vegetative tissue is the accumulation of carbohydrates, of which sucrose is the dominant moiety. Across a range of resurrection plant species sucrose concentrations increased (on a mole per unit dry mass basis) in dehydrated compared to hydrated leaves. The range of increase was extremely wide (from 1.2- to 28-fold with a median around 3-fold), as was the range in absolute concentrations in dehydrated tissue (from 72 to 2000 with a median around 200 μmol g−1 dry mass; data in Table 1 of Scott, 2000). The median value for sucrose represents about 7% of the dry mass. Accumulation of trehalose has been shown to be important in yeasts, and the molecule has an appropriate structure to interpolate between polar lipid head groups. In the few cases where trehalose

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was measured in resurrection plants there were large species differences in absolute contribution but no clear correlation with dehydration (Table 1 in Scott, 2000). The mechanisms already discussed for the protective value of sucrose in seeds, in water replacement and in vitrification also apply to vegetative tissue (Scott, 2000). The accumulation of sucrose may be a common feature of many resurrection plants; however, the metabolic routes by which this is achieved may differ (Bartels and Salamini, 2001). Possibilities are: 1. by conversion from 2-octulose; 2. by liberation from starch reserves; 3. from photosynthesis. The situation in vegetative tissue differs from that in seeds in a number of respects. Seed maturation is generally (1) a predictable and recurrent part of the life cycle, (2) a relatively slow process operating in (3) relatively small volumes of tissue. None of these is necessarily true of vegetative tissue facing drought. Although there must be some element of predictability (a coordinated response could not otherwise have been competitive) this is much less certain. The dehydration is imposed by the vicissitudes of the external environment, and not by a programmed seed development, and this may call for a more rapid reaction. Finally, the tissue volume involved is very much greater. For all these reasons the redirection of respiratory substrates into sucrose synthesis is quantitatively inadequate to meet demands. Protection via synthesis may be too slow (Scott, 2000) especially because water deficit per se inhibits photosynthesis and because planned metabolic shutdown is itself part of the desiccation tolerance process. C. plantagineum has reserves of 2-octulose, although this is not universal amongst resurrection plant species (Scott, 2000). In C. plantagineum, the concentration of 2-octulose shows diurnal variation, appearing to provide the same storage role as starch in most C 3 plants; although C. plantagineum does make starch this is minor in relation to 2-octulose (Norwood et al., 2000). 2-Octulose is accumulated during the day and utilised at night, and this was suggested to be under the control of a circadian rhythm since the accumulation/mobilisation appeared to anticipate, rather than follow, the changes between dark and light (Norwood et al., 2000). In photosynthetically active leaves this C-8 sugar may constitute 90% of all soluble sugars and up to 40% of the lyophilised dry mass (Bartels and Salamini, 2001). As the plant dehydrates, the concentration of octulose declines and of sucrose increases, and this is reversed upon rehydration, but within this shift the total concentration of sugars in hydrated and dehydrated tissue differs little (Bartels and Salamini, 2001). The use of 2-octulose for temporary storage is therefore constitutive and the system needs only to be utilised, rather than initiated, in desiccation tolerance; however, the control during the prolonged dehydration/rehydration cycle may be quite different from the diurnal cycle. The octulose–sucrose interconversion is restricted to the leaves. Although octulose is found in phloem sap (Norwood et al., 2000) it is the tetrasaccharide stachyose that is the dominant sugar in the roots, and the sugar content of roots is little affected by water availability (Bartels and Salamini, 2001).

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15.4.5

385

Hydrins or LEA proteins

Hydrins in vegetative tissue are the counterparts of LEA proteins in seeds (Bohnert, 2000) and have the same proposed functions – particularly interacting with sucrose in stabilising internal structure. The same difficulties in resolving function apply as with seeds – that the LEAs are produced in situations (in seeds and tissues of desiccationsensitive species) in which desiccation tolerance is irrelevant. However, proteins needed in the desiccation phase can be synthesised only prior to the desiccation phase; this part of the programme may work and tolerance may fail elsewhere. Also, LEAs have stress-related functions other than the desiccation tolerance phase of response to water deficit (Bohnert, 2000). It is also pointed out (Bartels and Salamini, 2001) that the contribution to tolerance by individual LEA proteins may be small and that it is the coordinated synthesis of a complement that may play a central role; hence, detection of individual LEA proteins signifies little. The production of LEAs may be one necessary component for desiccation tolerance, but it is not sufficient: the coordinated response may fail elsewhere; hence, while LEA production may be needed, production alone is not predictive of desiccation tolerance. The picture for vegetative tissue mirrors that of seeds in being a programmed response in which many components are necessary for tolerating dehydration, desiccation and rehydration; none alone is sufficient, and the process can potentially fail if any component is absent or inefficient. The acquisition of tolerance is also a slow process in vascular plants. When the resurrection plant C. wilmsii was dried rapidly, ultrastructural damage was seen, indicating that at least some of the protective mechanisms were not able to be put in place in these circumstances (Cooper and Farrant, 2002). Rapid desiccation may be a common feature of the life of bryophytes, but for vascular plants in natural conditions the development of severe water deficit is generally a gradual process.

15.4.6

Signals

Desiccation tolerance needs the combination of a range of constitutive (e.g. ability to shrink without damage) and facultative (e.g. metabolic shutdown and conversion of respiratory substrates to sucrose) properties. What is know about how this action is initiated and coordinated? Several hundred genes may be differentially expressed in response to dehydration (Bartels and Salamini, 2001), and this differential expression may take the form of an increase, a decline or a transient peak, during dehydration. These genes represent those encoding proteins with protective properties, membrane transport proteins, enzymes of carbohydrate metabolism, regulatory molecules and some whose role cannot be inferred from homology with known sequences – though sequence homology does not always imply functional identity (Bartels and Salamini, 2001). A protein with sequence homology with LEA D11 has a promoter that is influenced by desiccation treatment or by ABA (Michel et al., 1994). Scott (2000) discusses evidence that ABA is involved in the signalling path of desiccation tolerance in common with its general role in plants exposed to water stress, and its established role in seed maturation. It is proposed that roots detect water deficit first

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and that they synthesise and release ABA in common with the general theme. In desiccation tolerance, ABA activates particular genes – those leading to the accumulation of sucrose from one or more sources and also inducing the synthesis of various LEAs. Apart from these dedicated examples, the coordinated response also requires the shutdown of metabolism and, in some species at least, the degradation of chlorophyll. Other pigments may also be synthesised. For resurrection plants, rehydration can take place in 24 h; it is therefore likely that the tolerance of rehydration must be present before desiccation, forming another part of the coordinated sequence. The number of genes that are responsive and the range of events that may all be needed in the correct sequence to prepare for, undergo and survive desiccation and subsequent rehydration implicate a complex regulatory pathway. A range of molecules with possible transcriptional activities or signalling roles have been suggested, based on induction of their transcripts by early dehydration and/or by ABA treatment, and on sequence homology with other molecules of known activity. They include genes for Myb, a heat-shock transcription factor, the HDZIP (homeodomain Leu zipper) family, PLD (phospholipase D), a kinase and CDT-1 (possibly acting as a regulatory RNA) (Bartels and Salamini, 2001). These are single points in a complex network and in most cases the target genes have still to be identified. As an example, in C. plantagineum, PLD is induced rapidly and specifically by dehydration, and not by ABA (Frank et al., 2000), and catalyses the hydrolysis of phospholipids to phosphatidic acid. The phosphatidic acid then acts as a regulator of protein kinases or GTP-binding proteins. Two PLD transcripts exist, one constitutive and one dehydration responsive (Frank et al., 2000). Regulation is clearly complex, acting at both expression and cellular distribution, responsive to the water status of the leaf. The closest structural homologue to this PLD in Arabidopsis thaliana is linked to senescence and regulates stomatal closure, demonstrating that both function and pattern of expression have diverged between A. thaliana and Craterostigma, with the latter recruiting more genes to the task of tolerance (Bartels and Salamini, 2001). Although elucidation of the signalling and regulatory processes is at an early stage, the likelihood that the relevant genes may be present in all land plants seems to suggest that enhancing desiccation tolerance rests with regulation rather than biochemical hardware (Bohnert, 2000).

15.4.7

Constraints to the development of desiccation tolerance

It is concluded (Bartels and Salamini, 2001) that the genetic information for desiccation tolerance ‘is present in the genome of most if not all higher plants’ and that differences in the control of gene expression are probably responsible for the confinement of desiccation tolerance to seed development in species that are desiccation sensitive in the vegetative state. The reasons for this bear consideration, particularly with regard to any discussion of introducing desiccation tolerance to agricultural advantage. Not many angiosperm plant species have ‘failed’ to develop desiccation tolerance in relation to reproduction (and some of those may not have ‘failed’ but never

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‘needed to’). However, only 0.2% of angiosperm species have developed desiccation tolerance of their vegetative tissues. The restriction of desiccation tolerance to a few hundred species of defined ecology, when the information to achieve it is thought to be widely present, suggests that the acquisition of tolerance in vegetative tissues either is not easy and/or carries in most cases an unacceptable cost. Some constraints are: 1. ecology (the programme of pre-emptive, coordinated shutdown carries a cost, which must still yield a competitive advantage); 2. size (in vascular plants the xylem must be able to be re-filled); 3. mechanical properties (the cells and tissues have to be able to shrink; most leaves just wilt and die); 4. roots (they have received relatively little attention). The first question is an ecological one. In which circumstances will a resurrection plant have a competitive advantage over (say) a desert ephemeral? Are there only a few niches where the controlled expression of a whole gamut of processes works better than producing seed quickly and calling it a day? Resurrection plants are able to reproduce within 2 weeks, but some ephemerals would not be far behind. The commitment of resources, and particularly the commitment to a pre-emptive shutdown, may partly explain the rarity of desiccation tolerance in whole vascular plants. The requirement for desiccation tolerance in the pollen and seeds of most species is highly predictable and the competitive advantage is clear enough. For a whole plant the competitive advantage is a closer call. If early dehydration is predictive of a drought that will be long and severe then it may be a competitive advantage to shut down and survive while other species die, provided that reproduction can still keep pace with ephemeral species. But shutting down at early signs of drought will not be such an advantage if the signs are poorly predictive and the rest of the competition keeps going. Perhaps there are only a few niches in which desiccation tolerance is a competitive advantage. The second question lies in scale and complexity, at organism level. In addition to the cellular and subcellular responses two other factors are critical: recharging of the xylem and the mechanical ability to shrink. The work that has been described on xylem recharging points to another tier of adaptation being needed to cope with this in all but low-growing plants. Another tier of adaptations permitting cell walls to concertina and maintain wall–plasma membrane contact during shrinkage is also implicated to avoid irreparable mechanical damage. Structural pitfalls occur even in seeds. The large degree of vacuolation of embryonic tissue of A. marina is of structural, ecological and physiological importance to the species. Such species may fail to develop orthodox seeds regardless of the biochemical machinery with which they were equipped. Another area that has received little attention is what do the roots do? The constraints on root shrinkage have been recognised, but how do roots survive? Is there yet another package of adaptations or a restriction to a soil type that can permit shrinkage?

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The expression of the package of processes seen in seed maturation and in vegetative tissue might be only a matter of control – the higher level adaptations are not. If it proves that it is the availability of these adaptations that are limiting in most species then the low incidence of vegetative desiccation tolerance would be accounted for. While desiccation tolerance in lower plants, and in seeds and pollen of most higher plants, is unequivocally essential, desiccation tolerance in vegetative tissues of angiosperms may remain an ecological curiosity. Agricultural relevance, except in some perennial crops, will be slight, since the vegetative tolerance of desiccation is applicable only to stage III drought (see Chapter 13).

15.5

Concluding remarks

Extensive work on seed maturation has demonstrated that the acquisition of desiccation tolerance results from a coordinated package of responses with mechanical, physiological and biochemical components. The process of acquisition is relatively slow and proceeds with a defined sequence. Desiccation tolerance includes the ability to survive both dehydration and rehydration. Desiccation tolerance of vegetative tissue is common in bryophytes, which can be dehydrated rapidly, and whose tolerance consists largely of repair processes operating during rehydration. This is in complete contrast with both seed and vegetative tolerance of desiccation in angiosperms. Tolerance in angiosperms has much in common with tolerance in seeds. It is a coordinated response of many components and is both sequential and preemptive, and includes advance commitment to full metabolic shutdown. Although the genetic information for desiccation tolerance as found in seed maturation is considered to be present in nearly all angiosperm species, the occurrence of vegetative tolerance in very rare. Additional factors at tissue and organism levels of organisation that might limit or preclude the development of desiccation tolerance, despite the potential for the necessary biochemical machinery, are considered.

Acknowledgements Dr Ilse Kranner, RBG Kew, is thanked for helpful comments and suggestions on this chapter.

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Index 14-3-3 proteins, 111–13 and protein–protein interaction, 111–13 2-octulose, 384 2-oxoglutarate:malate antiport, 144 AAC (mitochondrial inner membrane ATP/ADP carrier), 157 AAP (amino acid permease), 257 AAS (atomic absorption spectrometry), 16 ABA, 100, 204 biosynthesis, 318 calcium signalling, 319 in desiccation tolerance, 373, 386 in stomatal regulation, 319 ABA-deficient mutant, 318 ABC (ATP binding cassette) transporters, 92, 147–8 mitochondrial inner membrane, 160–1 peroxisomal, 165–6 tonoplast, 172 abiotic stresses (and transcriptional regulation), 107 accumulation (in transport assays), 53–5 active transport, 77 activity coefficient, 18 of solute, 18 adhesion, 30, 219, 222 ADP/ATP transport in plastids, 146 aequorin, 57 affinity, 275 of transport processes, 275, 280, 282 aggregation (of macromolecules), 30 agonist, 84 ALMT1 (aluminium-activated malate channel), 302 aluminium (Al) in acid soil, 301–3 chelation by citrate, 302 ionic species and pH, 301 lipid peroxidation, 302

mitochondrial activity, 302 reactive oxygen species, 302 root elongation, 302 tolerance and toxicity, 301, 302 amino acids (in phloem), 242–3 amino acid transport, 159–60, 257 in mitochondria, 159–60 in phloem, 257 ammonium transporter expression of, 103 regulation of, 104 amphiphatic (amphiphilic), 49 AMT (ammonium transporter family), 82, 116 amyloplasts, 136 analysis inorganic elements, 15–18 material for, 15 anhydrobiotic plants, 372 antagonist, 84 antioxidants in desiccation tolerance, 375, 383 in phloem, 245–7 antiport (definition), 79 aphids (phloem sampling), 249 apoplast quantification of ions in, 56 salt accumulation in, 342 apoplastic pathway, 39–40, 350 aquaglyceroporins, 86 aquaporin, 85–7 dimensions of, 86 phloem, 256 post-translational regulation, 87 structure, 86 tonoplast, 171–2 transcriptional regulation, 87 transport rates through, 86 aqueous polymer two-phase, 61 arabidopsis (as model plant), 134–6 arsenic (As), 305

392 AtHKT1, 231 in phloem loading of sodium, 357 atmospheric pressure, 31 atomic absorption spectrometry, 16 ATPase, 77, 168–70 autoinhibition of calcium pumps, 111 of proton ATPases, 109–10 autoinhibitory domains, 110–11, 112 betaine aldehyde dehydrogenase, 325 bioavailability and chelation, 277 of nutrients, 276 of phosphate, 277 bioinformatics, 134 bordered pits, 219 boundary conductance (leaves), 321 boundary layer, 42 of roots in solution, 53 in soil, 278, 279 BT1 (ADP-glucose transporter), 146–7 bundle sheath, 331 Bunsen co-efficient, 330 bypass flow, 198 in halophytes, 350 and initiation of lateral roots, 198 and salinity, 350 C4 photosynthesis, 318, 329–30 types, 331 CAC (carnitine carrier), 160 calcicole/calcifuge, 291, 301 calcium active transport into vacuole, 175–6 apoplastic pathway of transport, 201–2 channels, 85 in mitochndrial inner membrane, 162 and sieve element structure, 247 signalling in guard cell opening, 100–1 uptake along root, 201–2 transport (inner membrane), 150 calcium-binding proteins, 108 calcium-dependent kinases, 108 callose (and plasmodesmatal aperture), 239 calmodulin (CaM), 113–14 binding, 113–14 and autoinhibition, 113

INDEX

family (targets for), 113 and phosphorylation, 113–14 and vacuole, 114 Calvin–Benson cycle, 149, 330 CAM (crassulacean acid metabolism), 318, 331 adverse consequences, 333 ecological distribution, 333 induction, 333 intermediates, 333 cAMP (cyclic AMP) (in signal transduction), 114–15 capacitance of membrane, 62–3 recording, 118 capacity of transport processes, 280 carbohydrate transport (at inner envelope), 145 carboxylates, 300 carrier definition, 78 transport rates of, 78–9 Casparian strip, 40, 197, 200–2 CASTOR, 150 CATE (compartmental analysis by tracer efflux), 55 cavitation, 217 and conduit radius, 217–18 CAX (proton:calcium antiporter), 175–6 CAX1 (regulation by SOS), 107 cell volume reversible and irreversible change, 100 regulation, 99–102 cell walls, 26, 39 enzymes of, 244 cellular shrinkage (in desiccation tolerance), 374, 382 cGMP (cyclic GMP) (in signal transduction), 114–15 chaotrope, 30–1 chaperones (RNA and sieve element), 239 chelates (release by rots), 193 chelation (and metal tolerance), 305–6 chemical gradient, 79 potential (of water), 31 potential gradient, 41 chlorine (as essential element), 341

INDEX

chloride regulation of uptake, 353 transport to xylem, 356 channels (CLCs), 85 selectivity in root, 353 transport and salinity, 353 transport and grafting experiments, 356 transport in guard cells, 100 chloroplast, see also plastid chloroplast, 136–152 structure, 136 choline oxidase, 325 CHX23 (plastid envelope sodium/potassium:proton exchanger), 150 CLC (chloride channel), 141 cluster roots, 194, 299 CNGCs (cyclic nucleotide gated channels), 115 CO2 -concentrating system, 330 CO2 -pump CO2 -shuttle, 331 cohesion, 30, 219 cohesion-tension theory, 219–224 controversies, 222–224 colligative properties, 33–4 companion cell, 251 gene expression in, 243 compartmental analysis, 22–3, 55 compartmentalisation, 3, 133, 344 engineering of, 358 and metal tolerance, 305–6 of salt, 343 of sodium, 106 energy cost of, 358 compatible solute, 324, 344 in desiccation tolerance, 376 genetic manipulation, 324 concentration (effect of water structuring on), 18 concentration-dependence (of ion uptake), 208 copper (Cu) (transport at inner envelope), 152 cortex (and nutrient uptake), 196 cortical cells (role in nutrient uptake), 199 cosolvant, 30–1, 324 coupling (water and solute fluxes), 44–5 Crafts and Broyer hypothesis, 224

393

crassulacean acid metabolism, see CAM cristae stacks (in mitochondria), 153 critical concentration in soil solution, 280 critical-point drying, 21 cryopreservation, 20, 21 cryptograms, 372 CTS/COMATOSE (peroxisomal), 165–6 cyclic nucleotides and channel gating, 115 in signal transduction, 114–15 cytoplasm molecular motility and desiccation, 378 volume fraction and salinity, 346 cytoplasmic calcium (as signal of salt stress), 106 cytoskeleton, 374 Darcy’s law, 39 databases, 63 Arabidopsis knock-out lines, 63–4 in plant research, 135 deficiency (mineral), 291 deletion mutants, 108 depletion zone, 196,199, 278, 282 depolarisation-activated channels, 84 desiccation avoidance, 371 point, 378 tolerance, 371–88 antioxidants, 375 constraints to development of, 386–7 co-ordinated response, 380–1 ecology of, 387 evolution of, 372 intracellular de-differentiation, 374–5 metabolic shut-down, 375 phylogenic distribution, 372 physical characteristics, 374, 382 processes involved, 374 scale effects, 383 seeds, 372–9 vegetative tissue, 379–88 DGDG, 49 D-genome (of wheat and salt tolerance), 362 dicarboxylate transporter, 144, 332 diffusion, 41–2, 80 across membranes, 41–2 coefficient, 41, 80

394 diffusion (Continued ) potential, 42–3 rate in soil, 277, 278 through membranes, 81–2 direct sampling (single cells), 22 divalent cation transporter (IRT1/2), 295, 296 DNA/RNA interaction (in phloem), 245 Donnan systems, 43–4 in root apoplast, 196–7 drought, 106, 315 3-stage model, 314, 312, 322 tolerance (desiccation avoidance), 371 tolerance (ecological and agricultural contexts), 314 dryland (definition), 314 DTC (mitochondrial dicarboxylate/tricarboxyate carrier), 158–9 dual isotherms, 209 efflux analysis, 23–4 radioisotope measurements, 206 elaioplasts, 136 electrical gradient, 79 potential gradient, 41 electrochemical potential, 53 gradient, 41 electrophysiology, 57–60, 100 technical requirements, 58 voltage-based measurements, 58–60 elongation zone, 201 embolism (of xylem), 217–18 endocytosis, 117 endodermis, 39, 197 epicuticular wax, 321 epidermal transfer cells (and iron uptake), 200 epidermis and nutrient uptake, 196 role in nutrient uptake, 199 essential elements, 290 metals, 303 exocytosis, 117

INDEX

exodermis, 39, 202 exploitation by roots, 279 exploration by roots, 279 F1 –F0 ATP synthase, 89 facilitated diffusion, 82–3 Fenton reaction, 293, 303, 304 ferric, see also iron chelate reductase, 295 physosiderophore transporter, 296, 298, 299 ferritin, 298 fertiliser losses from soil, 279 FES (flame emission spectrometry), 16 Fick’s first law, 41–2, 80 field capacity, 38 flame emission spectrometry, 16 flow (in phloem), 250–1 fluid mosaic model, 50 fluorescent probes, 22–3 AM esters, 56 membrane-impermeant, 56 ratiometric, 56 single-wavelength, 56 in transport assays, 55–7 protein (FP) nanosensors, 57 flux analysis, 23–4, 55 density of nutrient uptake, 278 FOL1 (folate carrier), 147 free energy, 31–2 freeze-fracture, 21 freeze-substitution, 21 freezing, 326 injury, 326 FRET (fluorescence resonance energy transfer), 57 FRO1/FRO2, 295 fungi, 194 fusicoccin (FC), 111–12 gene expression in companion cell, 243 in desiccation tolerance, 382, 385–8 in phloem, 243 regulation of nutrient transport, 102 in water deficit, 334 general diffusion pore, 154

INDEX

genetic manipulation (of compatible solute synthesis), 324 GEOCHEM, 52 Gibbs free energy, 31–2 glassy state, 377–8 glucose transporter (at inner envelope), 146 glutamate/malate transporter, 144–5 glycerophospholipids, 49 glycine betaine, 324, 344 compartmentalisation, 344 synthesis, 325 glycolipids, 49 glycophyte, 107, 340 Goldman Hodgkin Katz equation, 44, 81 GORK, 101 GPT (glucose-6-phosphate/phosphate translocator), 143 Gram-negative bacteria, 142 graphite furnace, 16 gravitational potential, 35 GS-GOGAT, 167 guard cell, 319 guttation, 222 Hagen–Poiseuille equation, 40, 216, 251 HAK/KUP transporters (and salinity), 349 halobacteria, 346 halophyte, 107, 340 halophytes (quick fix or model?), 365 hardening-off, 373 heavy metal ATPases (HMAs), 91 Henry’s Law, 330 heterologous expression in transport assays, 65–6 in Xenopus laevis, 66 in yeast, 65 heteromerisation (and regulation of transport), 116–17 high affinity phosphate transport (induction of), 102–3 phosphate transporters, 285 sulphate transport (induction of), 102–3 HKT transporters (and salinity), 349 HKT1, 82–3 homiohydry, 372, 382 HXK (hexokinase) signalling, 105 hydraulic conductance, 53 and soil water content, 322

395

hydrins, 385 hydrogen bond, 29–30, 221 strength of, 29 hydrophobic effect, 371 hydrophobicity (of xylem), 223 hydroponics advantages, 286 controlled environments, 287 definition, 286 disadvantages, 286 economic costs, 286 media, 286 hydrostatic pressure, 38 in xylem, 222 hyperaccumulation, 305–8 functions of, 308 and herbivory, 308 hyperaccumulator, 198, 292 hyperpolarisation-activated channels, 84 ice nucleation, 326 in silico techniques, 63–4 inhibitors (in transport assays), 53 inner envelope carriers in, 146–7 transport across, 142–152 inorganic elements (analysis), 15–18 ions extraction of, 15 in phloem, 241–2 interception (of nutrients by roots), 279 intermembrane space (in mitochondria), 153 inward-rectifying channels, 84 ion channel(s) anion, 356 in xylem loading, 227 definition, 78 gating of, 83 in xylem parenchyma, 225 mitochondrial inner membrane, 161–2 regulation by 14-3-3 proteins, 112 and stomatal opening, 100 at tonoplast, 173–5 transport rates through, 78–9 transport through, 83–5 chromatography, 17 selective electrode, 17, 21–2, 58–9

396 ion (Continued ) compartmental resolution with, 59 multi-barrelled, 59 transport in hyperaccumulators, 305 techniques of measurement, 207–9 transporters post-translational regulation, 99 regulation, 99, 107–16 by photosynthesis, 104–5 transcriptional regulation, 99 trafficking of, 117–20 uptake longitudinal distribution in root, 201 iron, see also ferric in alkaline soils, 293 availability, 193, 294 chelating agents (in plants), 298 concentrations in plant tissue, 293 as essential element, 293 mechanisms of uptake, 294–9 and phytotoxicity, 293 roles in the plant, 293 strategy-I plants, 294, 295, 296 strategy-II plants, 294, 296, 297, 298 transport at inner envelope, 152 irreversible thermodynamics, 44–5 IRT1/IRT2, 295, 296 K/Na selectivity, 350 KAT1, 119 kinetics Michaelis–Menten, 54, 208, 275, 281 of potassium uptake, 208 solute uptake by roots, 204–9 KIRCs (potassium inward rectifying) channels, 230–1, 349 Km , 275 KORCs (potassium outward rectifying) channels, 349 kosmotrope, 31 Krebs cycle, 158 KUP (and xylem unloading), 231 KUP/HAK gene family, 94 laser micro-dissection, 244 latent heat of vaporisation, 29 LCT1, 350

INDEX

LEA proteins, 372, 385 in desiccation tolerance, 376 intrinsically disordered, 376 mode of action in desiccation tolerance, 376 leaf area expansion, 320 area regulation, 320 temperature, 321 ligand-gated channels, 83–4 lignin (in xylem), 216 lipid classes of, 49 hydrophobic and hydrophilic regions, 49 rafts, 50 lipid–protein interactions, 51 liposomes (and patch clamping), 63 liquid-disordered (of membrane structure), 50 localisation by electron microscopy, 20–1 by freezing, 20–1 organic compounds, 25 by precipitation, 20 of solutes, 19–25 low MW carbohydrates (in desiccation tolerance), 376, 283 macromolecular trafficking (in phloem), 248 macronutrients, 290 magnesium active transport into vacuole, 176–7 transport at inner envelope, 152 major facilitator superfamily (MFS), 144–5 malate transport in CAM, 171, 332 at tonoplast, 170–1 hypotheses (of aluminium tolerance), 302 transporter, 332 manganese (and aluminium), 301 markers (for membrane fractions), 61 mass flow dependence on hydraulic conductance, 278 of nutrients to roots, 196

INDEX

matrix (in mitochondria), 153 maturation drying, 373 MCF (mitochondrial carrier family), 156 mechanosensitive channels, 83 membrane(s) as barrier, 75 capacitance, 62–3 composition, 47–50 first proposal of, 75 fluidity, 49, 51 fractionation, 61, 64 fraction identification, 61 functions of, 3, 47 isolation techniques, 60–1 major transport proteins of, 48 microdomains, 50 passive permeability of, 49 phase transition (in desiccation), 378 potential, 84 measurement, 58–60 roles of, 48 structure, 50–1 transport (terminology), 76–7 turnover, 117 vesicles (transport assays in), 61–3 meniscus, 30, 38 mercury (and aquaporins), 53 metabolic shut-down (in desiccation tolerance), 375 metal detoxification (methods), 304 tolerance, 303 and vacuolar accumulation, 306 toxicity, 303 acute and chronic, 306–7 metallophyte, 292 metallothioneins, 306 MEX1 (maltose transporter), 145–6 MFS family, 144–5 MGDG, 49 Michaelis–Menten kinetics, 54, 208, 275, 281 microdomains (in membranes), 50 micronutrients, 290 mineral analysis (optical methods), 16

397

deficiency, 293–300 definition, 292 efficiency (definition), 292 toxicity, 301–8 definition, 292 use efficiency (definition), 292 mineralisation, 277 MINTEQ, 52 mitochondria, 153–62 function of, 153–4 transport at inner membrane, 155–62 transport at outer membrane, 154–6 mobilisation (of nutrients) 284 mRNA (in phloem), 243 MRP (multi-drug-resistance-related protein) subfamily (of ABC transporters), 172–3 mucopolysaccharides (role in xylem), 223 mugeneic acid, 296, 297 synthesis of, 297 multi-barrelled electrode, 21–2, 59 multi-drug-resistance (and ABC transporters), 92 M¨unch hypothesis, 250–1 MYB-CC family (transcription factors), 108 mycorrhizae, 194, 282, 300 negative charge (of cell interior), 79 Nernst equation, 43, 81 potential, 43 Nernst-Planck electrodiffusion relationship, 80 net uptake (in transport assays), 53–4 NHX (sodium:proton antiporter), 95 osmotic or ionic regulation, 359–60 overexpresion in Arabidopsis, 359–60 difficulties and inadequacies of methodology, 359–60 in tomato, 359–60 in wheat, 361 transporter, 359 overexpression of, 359 NHX1, 176 calmodulin binding to, 114 regulation by SOS, 107 Ni, 305 nicotianamine synthase, 298

398

INDEX

nitrate critical concentration, 281 movement in soil, 194 in phloem, 243 transport, 94 inner envelope, 149 transporter (stoichiometry of), 94 nitrification, 277 nitrogen (in phloem), 242–3 nitrogen-fixing bacteria, 195 non-aqueous extraction, 22 non-essential metals, 303 non-orthodox seeds, 373 non-reducing sugars (in desiccation tolerance), 377 non-selective ion channels, 85 NORC channel (and xylem loading of sodium), 226 NRAMPs (as metal transporters), 177 NRT1/2 gene families, 94 NRT2 (high affinity nitrate transport) (induction of), 103 NSCCs (non-selective cation channels), 200, 349 NTT (ATP:ADP transporter in plastids), 146 nuclear magnetic resonance (NMR), 54 nutrient concentration in xylem, 281 flux density, 281 mass flow in soil, 278 mobility, 193 patches, 283 replacement, 290 solubility, 193 supply to root surface, 276 translocation in xylem independence of water movement, 224 regulation of, 224 transport regulation by plant nutrient status, 103–4 regulation of gene expression, 102 in xylem, 224–31 uptake flux density of, 278 and inter-specific competition, 284 link to carbon status, 104–5 link to other nutrient status, 105 by roots, 276

OEP (Outer Envelope Proteins), 138–42 family, 138–42 OEP16 (channel for amino acids and amines), 138, 140 OEP21 (channel for phosphorylated carbohydrates), 140–1 regulation of, 140 structure, 140 OEP24 (high conductance channel), 140–1 structure, 141 OEP37, 141 Oertli hypothesis, 342 Ohm’s law, 83 oleosins, 378 organic compounds (analysis), 18–19 orthodox seeds, 373 osmolyte, 324 osmoprotectant, 324 osmosis, 33–4 discovery of, 75 osmotic adjustment, 317, 322–3 and compartmentalisation, 324 and crop yield, 323 and ion uptake, 324 genetic variation in, 323 and salinity, 343–4 pressure (in sieve element), 249 water lifting, 223 outward-rectifying channels, 84 overexpression of phosphate transporters, 285 of target genes, 64 parasitic plants, 262 passage cells, 200 and calcium uptake, 200–1 passive transport, 77, 81–7 patch clamp electrophysiology, 62–3 cell attached, 62 inside-out, 62 outside-out, 62 whole cell, 62 patchy distribution (of nutrients), 194, 293 P-ATPase interaction with 14-3-3 proteins, 88 PMF produced by, 88 structure, 88

INDEX

PEP-CK (PEP carboxykinase), 333 PEPCase (PEP carboxylase), 330–1 perforation plate (in xylem), 214 permanent wilting point, 322 permeability, 53 coefficient, 42 peroxisomes, 162–7 function of, 163 peroxisomal membrane porin, 163–5 pH (of stroma), 149 phase transition (in membrane), 50 phloem anatomy, 236–9 carbohydrate composition of, 240–1 composition of translocate, 240–8 contents (variation in), 241 defence against herbivores, 261 exploitation of, 261–2 interaction with pathogens, 246 loading, 251–7 ecophysiological context, 253–4 symplastic or apoplastic, 251–2 transporters, 254–6 macromolecular trafficking, 248 resource partitioning, 260 sap (collection of), 240 transporters and loading, 254–6 unloading, 257–60 apoplastic pathway in, 259 developing fruits, 259 pathway of in roots, 257–8 seed coats, 259–60 symplastic pathway in roots, 258 viruses in, 261 phosphate. See also phosphorus bioavailability, 277 critical concentration, 281 concentration in cytoplasm, 284 in soil, 284 in vacuole, 285 translocator family, 142–4 transport (inner envelope), 149 transporters gene regulation, 285 and plant phosphate content, 285 uptake, 299, 300 and proton extrusion, 300

399

phospholipase D, 49, 386 phosphorus. See also phosphate acquisition, 284 availability, 193 concentration in soil, 299 rate of diffusion in soil, 299 and release of organic acids, 284 and release of phosphatases, 284 and root proliferation, 284 phosphorylation (and 14-3-3 binding), 112 photorespiration, 166–7, 169 photorespiratory pathway, 166–7, 169 photosynthate (efficiency of allocation), 235 Pht1 family, 285 Pht2 family, 285 PHT2;1 (inner envelope phosphate transporter), 149 physiological criteria (in breeding), 362 phytochelatin synthase, 307 phytochelatins, 306, 307 and homeostasis, 307 phytohormones (in xylem sap), 229 phytoplasmas (in phloem), 261 phytoremediation, 303 phytosiderophores, 296 phytotoxicity, 291 PiC (mitochondrial inner membrane phosphate carrier), 156–7 PIPs (Plasma membrane intrinsic proteins), 86–7, 256 pit(s) (of xylem), 218 field, 219 membrane, 215, 219–8 plant growth (for transport studies), 52 plasmodesmata, 197, 200 in phloem, 238–9 selectivity of, 238 frequency, 253 regulation of aperture, 239 structure, 238 plastids, 136–52 differentiation, 136 function of, 137 PMF (proton motive force), 88 PMP38 (peroxisomal ATP/ADP carrier), 165 Poacea (iron uptake in) 296, 297 poikilohydry, 372, 382 Poiseuille equation, 40, 216, 251

400

INDEX

POLLUX, 150 porin, 163–5 post-translational regulation, 109–11 potassium channel(s), 84–5 in mitochondrial inner membrane, 161–2 regulation by pH, 101 voltage dependence, 101–2 distribution at equilibrium, 93 in phloem, 241 sensing of concentration, 102 transport in guard cells, 100 and phloem loading, 256 transporter(s) and phloem loading, 256 regulation of, 103–4 uptake, 93–4 concentration dependence, 208 effects of ABA and FC, 113 xylem loading of, 225–6 potassium:sodium selectivity, 350 potassium:sucrose reciprocity (in phloem), 242 potassium:proton symport (stoichiometry), 94 potassium:proton antiport (and xylem unloading), 230–1 PPase, 168–70 PPF1 (role in calcium transport at inner envelope), 150–1 PPT (phosphoenolpyruvate:phosphate translocator), 143–4 pressure-flow hypothesis (in phloem), 249 pressure-probe (use in xylem), 223 primary active transport, 87–92 primary metal pumps, 90–2 primary proton pumps, 87–90 primary transport, 77 proline, 324–5 protein(s) folding of, 30 kinase cascade, 101 in phloem, 246 synthesis (and salinity), 342 tertiary structure of, 30 trafficking (in plasmodesmata), 238–9 protein-lipid interactions, 51

proteoid roots, 282, 299 proteomes (of organellar membranes), 134 proton ATPase and phloem loading, 255 traffic in, 117 turnover rate, 88 and xylem unloading, 231 ‘battery’, 77 extrusion and phosphate uptake, 300 and iron uptake, 295 protoplasts isolation of, 60 transport assays in, 61–3 P-type (plasma membrane) ATPase, 88–9 P-type calcium ATPase, 90–1 pulvini (volume change in), 100 pump definition, 78 transport rates through, 78 pyrophosphatase, 90 QTL analysis, 63 co-localisation of salt tolerance and vigour, 363 and gene/environmental interaction, 364 and salt tolerance, 363 and variance in salt tolerance, 363–4 radial transport across root, 197–201 apoplastic pathway, 197–8 symplastic pathway, 197–8 radioactive tracers (in transport assays), 54–5 radioisotopes in study of solute uptake, 204–7 technical problems, 207 re-absorption (of sodium from xylem), 342 recalcitrant seeds, 373–4 re-circulation (of salt), 343, 356 reflection coefficient, 34, 42 regulation molecular basis of, 107–17 post-translational, 109–11 transcriptional, 108–9

INDEX

of transporters, 107–16 under stress ‘acclimation or panic’, 107 rehydration, 371 damage repair, 378 reporter genes (in transport assays), 64 resistance gas phase, 37–8 water flow, 37 in xylem, 215–16 resurrection plants, 382, 384–5 re-translocation (of salt), 356 rhizobia, 195 rhizoferrin, 298 rhizoshpere, 193, 277 RNA movement through plasmodesmata, 239 transport (in phloem), 248 root, 326–7 apex (involvement in nutrient uptake), 201 behaviour in ‘real’ soils, 196 branching of, 202 contribution to uptake of different root types, 202 deep constitutive, 327 facultative, 327 exudates and chelation, 279 and phosphate uptake, 300 and soil pH, 279 hairs, 196, 198 connection to cortical cells, 200 and immobile nutrients, 198 and iron strategyI/II plants, 199 longitudinal differences in uptake, 201–2 and phosphorus deficiency, 198 proliferation, 198 radius and surface area, 282 morphological responses to nutrient, 194–5 pressure, 222 proliferation, 299 and nutrient uptake, 282–3 precision effects, 283 scale effects, 283

401

release of enzymes by, 193 secretion by, 193 selectivity (and salinity), 348 and water acquisition, 327 rubisco, 166–7, 330, 332 salinity, 196, 340–64 high concentrations, 344–5 low concentrations, 341–3 moderate concentrations, 343–4 and protein synthesis, 342 short- and long-term effects, 341 stress, 106 and water deficit, 341 salt accumulation in apoplast, 342 and leaf area reduction, 341 long term damage, 341 damage (and plant vigour), 353 exclusion, 340, 342–3, 350–1 consequences of manipulating, 351 genetic manipulation of, 351 glands, 348–9 tolerance, 226 at cellular level, 346 conventional breeding, 362 genetic manipulation, 362 difficulties, 362–3 in tissue culture, 345 limited success of, 362–3 wild relatives, 362 inclusion vs exclusion, 344–5 and level of organisation, 345 at molecular level, 345–6 prospects for breeding, 361 timescale for breeding, 361 and vigour, 361 whole-plant level, 347 transport energy cost of, 346 in leaf cells, 347 as limiting growth of halophytes, 355 from root to shoot, 353 processes, 346 and transpiration, 347 sap-feeding insects, 261–2 saturation kinetics, 280

402 secondary active transport, 92–6 transport, 77 wall (in xylem), 215 sectoriality, 202 seed(s) desiccation tolerance in, 372–9 maturation, 373 storage longevity, 378 selectivity filter (in ion channel), 83 SFC (succinate-fumarate carrier in mitochondria), 160 Shaker-type channels, 84, 116, 225 sieve element (sieve tube) anucleate, 236 longevity, 237 metabolism, 247–8 organelles in, 236 osmotic pressure in, 249 positive hydrostatic pressure in, 248 protein synthesis in, 236 size, 236 turgor pressure in, 248–50 water relations, 248–60 sieve plate, 237 blockage, 237, 247 and conductance, 237 sieve tube. See sieve element signalling (in guard cell opening), 100 signals (in desiccation tolerance), 385 Singer and Nicholson model, 50 single-cell sampling, 22, 54 size exclusion, 78 SKOR, 101–2 potassium channel, 225 and xylem loading, 225 slam freezing, 21 SNARE proteins, 118–19 and ABA signalling, 118–19 sodium active transport into vacuole, 176 ‘battery’, 77 concentration in cytoplasm and xylem, 354 efflux, 95 extrusion, 352 energetic cost of, 351 in halophytes, 351 flux to load xylem, 354

INDEX

influx (rates of in halophytes), 352 leakage out of vacuole, 358 loading of vacuole, 358 mode of entry at root, 349 pathway of uptake, 106 in phloem, 241 phloem re-circulation, 357 re-absorption from xylem, 355 transport co-ordination of, 345 regulation of, 352 unidirectional and net transport, 355 xylem loading of, 226, 227, 351–2, 354 sodium:proton antiport, 95, 354, 359 accumulation power of, 95 electroneutrality, 95 at tonoplast, 357 and xylem loading of sodium, 227 sodium-coupled secondary active transport, 95 soil heterogeneity, 193, 279, 284 soil pH (and nutrient availability), 194 soil-plant-atmosphere system, 31, 36–8 solute(s) accumulation historical reasons, 1 physical reasons, 1 structural reasons, 2 acquisition and uptake by root, 280–4 functions of, 25–6 localisation, 19–25 radial transport, 197–8 range found in plants, 2, 19 range transported, 76 supply to root surface, 276–9 transport active, 87–95 assay techniques, 52–60 isolated organelles, 133 of nutrients, 224–30 passive, 81–6 scales of, 2–3 solution design (for transport studies), 52 sos mutants, 354 SOS signalling pathway, 106 SOS1, 227, 354 SOS1/2, 106 source-sink, 260 SP2 domain, 118, 119

INDEX

sphingolipids, 50 split-root experiments, 204, 206, 318 stereological analysis, 19–20 sterols, 49 stomatal closure and leaf temperature, 321 and oxidative damage, 321–2 guard cells (volume change), 100 regulation, 318 stopped-flow spectrophotometry, 61–2 stromal pH (regulation by potassium/proton exchange), 150 structural genes (in phloem), 244 succulence (and salinity), 357 sucrose accumulation in desiccation tolerance, 384 in phloem, 240 synthase (in phloem), 245 transport at tonoplast, 171 transporter, 255, 259 sulphate transport (inner envelope), 149 uptake (link to carbon metabolism), 105 SULTR1/2 (sulphate transporter), 103–4 supercooling, 326 surface free energy, 30 surface tension, 30, 322 in apoplast, 222 driving force for water movement (in xylem), 219–20 and xylem, 219 SUT1/SUC2 (proton-sucrose co-transporter), 238, 254–5 symplast, 197 symplastic pathway, 39–40 TCA cycle (mitochondrial carriers), 158–9 tensile strength (of water), 30 tension (in xylem), 220–2 thermal capacity (of water), 29 thermodynamics irreversible, 44–5 second law, 77 three-compartment model, 23–4, 205–6 TIPs (tonoplast intrinsic proteins), 86–7, 171–2, 256 tonoplast, 167–8

403

leakage in halophytes, 358 permeability and sterol composition, 358 salt concentration across, 346 solute transport across, 170 ion channels, 173–5 ion transport, 173–5 and passive leakage of salt, 358 proton-ATPase, 168–70 proton-pyrophosphatase, 168–70 torus (bordered pits), 219 toxicity (mineral), 291 TPC1 (slow vacuolar channel), 15 TPK1/KCO1 (tonoplast potassium channel), 173–4 TPT (triose phosphate/phosphate translocator), 143 tracheary elements, 214, 217 tracheids, 214–5 transcellular pathway (of radial movement), 200 transcriptional regulation, 108–9 by abiotic stress, 107 transfer cells, 200, 228 transition metal transport at inner envelope, 152 transport at tonoplast, 177–8 transpiration, 221–2 and humidity, 222 and leaf temperature, 321 and xylem transport, 203 transpirational cooling, 29, 321 transport active, 77, 87–95 assays, combining techniques, 66–7 future developments, 66–7 techniques, 51, 52–60 mechanisms (classification of), 78–9 passive, 77, 81–6 processes (regulation by stress), 106–7 proteins clustering of, 51 functions, 99 multimeric, 51 in phloem, 245–6 water, 214–22 transporter (definition), 76 regulation of expression, 102 trans-root potential, 203

404

INDEX

tree (tallest), 219 turgor pressure (in sieve element), 249–50 regulation, 322 ubiquitin (in phloem), 244 UCP (mitochondrial uncoupling proteins), 157–8 unidirectional fluxes, 205–6 uniport (definition), 79 unstirred layer, 42 roots in solution, 53 in soil, 278, 279 vacuoles, 25–6, 167–78 functions of, 168 V-ATPase, 168–70 complexity of, 89 PMF produced by, 89 regulation of, 89 VDAC (voltage dependent anion channel porin), 54–6, 138, 142, 167 vesicle traffic, 117 and cell volume, 118 vessel elements, 214 viruses (in phloem), 261 viscocity (of phloem), 251 vitrification, 377–8 voltage clamp, 82, 100 in intact tissue, 60 voltage-gated channels, 83 volume flux (of water), 36–7 V-PPase, 168–70 V-type (vacuolar) ATPase, 89–90 water ascent in xylem, 219–24 availability and plant zonation, 314 in cell walls, 39 channels, 85–7 deficit, 314–35 and elongation growth in monocotyledons, 320 gene regulation in, 334 plant responses (timeframe), 316–17 plant responses, 316 terminology, 314 movement electrical analogy, 36–7

phloem, 40 radial across root, 39–40 through soil, 38–9 xylem, 40, 216–17 additional mechanisms, 222–4 driving forces, 221–2 polar nature of, 30 potential, 32–3, 39 of drying soil, 322 gradient (soil-plant-atmosphere), 221–2 gradient, 32–3 in soil-plant-atmosphere system, 36 properties of, 29–31 quasi-crystalline nature, 30 structure of, 29 structuring of, 30 relations (of cells), 34–5 replacement hypothesis, 377 water-use efficiency, 318 and C4 /CAM, 328–9 xenobiotics (removal by ABC transporters), 92 X-IRAC (anion channel), 227 XPT (xylulose-5-phosphate/phosphate translocator), 144 X-QUAC channels, 227, 356 X-ray crystallography, 51 X-ray fluorescence, 17 X-ray microanalysis, 20, 54, 207 X-SLAC channels, 227, 356 xylem concentrations in, 281, 282 effects of stress, 230 in conifers and angiosperms, 216 developmental state along root, 202 embolism, 217–18 evolutionary development, 217 functional development, 201 leak prevention, 214 loading, 203, 224–7 active models, 225 factors affecting, 229 gating factors, 227 leakage hypothesis, 224 signals, 204 with sodium, 342 mechanical strength, 214, 217

405

INDEX

parenchyma, 203, 224–5 peak velocity, 217 re-filling (in desiccation tolerance), 383 re-uptake from, 204 safety mechanisms, 217–18 sap analytical methods, 228 composition, 228–30 osmotic pressure of, 228 pH, 229 structure, 214–16 unloading

hormonal control of, 230 in leaves, 230–1 mechanisms involved, 230 vessel (evolution of), 217 yield and water transpired, 317 threshold, 322 zinc, 305 transport at tonoplast, 177–8 ZNT1 (metal transporter), 305

S Passive 10 100 transport mM mM S Active transport

(A)

ATP Primary H + - ADP H+ S1 Sec. + H+ active S2 Sec. S3 passive +

(B)

+ -

Primary pump

+Antiport

ADP

+ - Carriers

Channels

(C)

Symport

- +

Uniport + -

(D)

Colour plate 5.1 The major classification systems that pertain to solute transport mechanisms. (A) When the cellular electrochemical potential is higher than the external electrochemical potential, uptake of solute S is an active transport step whereas its efflux can proceed through passive transport. (B) Primary transport in plants is exemplified by a membrane ATPase that uses metabolic energy by hydrolysing ATP to extrude H+ in the extracellular medium. Secondary active transport can use the resulting proton motive force to drive extrusion S1 or uptake S2 of solutes by coupling solute transport to the inwardly directed proton flux. Passive secondary transport S3 does not require energy input. (C) Different classes of transporter are often categorised on the basis of kinetic mechanisms: (i) ‘Pumps’ such as the primary H+ -ATPase are fuelled by metabolic, light or redox energy. (ii) Carriers bind and de-bind their substrate on either side of the membrane and can be active or passive. (iii) Ion channels only weakly bind their substrate and form a diffusive pore for the rapid trans membrane movement of ions. (D) Mechanisms that transport one type of substrate only are ‘uniporters’ and include both ion channels and carrier type transporters. Carriers that couple flow of more than one substrate are defined as symporters when all substrates move in the same direction, or as antiporters when counterflow of substrates occurs.

CO2 CO2

+ + -

CO2

CO2

CO2 CO2

+ -

- + NH4+

(A)

- +

+ -

NH4+ LeAMT1.1 (B)

Colour plate 5.2 Diffusion and facilitated diffusion of solutes. (A) Uncharged gaseous solutes such as CO 2 and O 2 enter and leave cells by moving through the membrane lipid bilayer without the help of specific proteins. Such compounds diffuse in the direction of lower chemical potential. (B) Facilitated diffusion is mediated by specific membrane proteins that allow passive but regulated transport of solutes across the membrane. An example is the uptake of NH 4 + through the tomato ammonium transporter LeAMT1.1.

−→ Colour plate 5.3 Ion channel properties. (A) Ion channels are integral membrane proteins that contain a diffusive pore through which ions can cross the membrane. The pore contains a narrow region, the selectivity filter (SF), that determines which ions can pass and which ions are excluded. Opening and closing of the pore, gating, is under control of a gate (GT) that in voltage-dependent ion channels senses the membrane voltage via a voltage sensor (VS). The latter is typically a transmembrane domain with positively charged residues. (B) Single channel current recording through a voltage-dependent ion channel, showing channel activity only when the membrane is depolarised (positive potentials, denoted on the right hand side). The arrows on the left hand side denote the channel closed level and channel opening results in a discreet increase in current level. (C) Voltage dependence of the channel recorded in (B) plotted as the open probability versus the membrane potential. Note that at negative membrane potential, the open probability is virtually zero whereas it rapidly increases at positive membrane potentials. (D) Channel gating can also depend on factors other than the membrane potential, for example on the presence of a channel ligand. Binding of the ligand to specific sites on the channel protein causes a conformational change leading to opening of the pore. Other compounds that have similar channel opening properties are called agonists, whereas antagonists have effects opposite to that of the ligand.

outside

(A)

SF

cytoplasm

VS

GT

80

(B)

60

> > > > > > > > > >

40 20 −20 −40 −60 −80

100 Po (%) 60 20 −200

(C)

−100 V membrane (mV)

closed

(D)

binding site

100

200

open

agonist ligand antagonist

(A) 140

Outward K+

0 -140

K+ Inward

(B) outside + D S1 S2 S3 S4 S5 GY S6 + G NH2

cytoplasm

HO2C

Colour plate 5.4 Voltage dependent K+ channels. (A) Whole cell recording of inward and outward K+ currents, which respectively occur when the membrane potential is clamped at negative membrane potentials (−140–0 mV in 20 mV steps) and positive potentials (0–140 mV). Note the time dependence of both inward and outward currents. Vertical scale bar represents 0.1 nA and horizontal bar 0.1 sec. (B) The generalised secondary subunit structure of voltage-dependent K+ channels is similar to that of Drosophila Shaker type channels comprising six TMD with a voltage sensing region in the S4 domain that controls channel gating, and a GYGD motif in the pore region that confers K+ selectivity to the channel. Functional channels contain four 6 TMD subunits that form either hetero- or homotetramers.

(A)

outside S1 S2

P

S3

S4

S5

N A P S6

N A cytoplasm

NH2

(B)

HO 2C

S5

S6

S1

HO2C

N S2 A

P P

A N

S4 S3

NH2

Colour plate 5.5 The generalised subunit structure of aquaporins. (A) Similar to Shaker-type voltagedependent ion channels, water channel subunits contain six TMD. Two pore regions appear in each subunit, where lipophilic regions that contain the aquaporin signature motif ‘NPA’ partially enter the lipid bilayer. (B) The two ‘half pore’ regions are believed to be positioned opposite each other in a tertiary ‘hourglass’ structure that forms a ring-like ‘NPANPA’ selectivity filter at the narrowest point that is highly selective for water molecules. Four subunits depicted in (B) are needed to form functional channel tetramers.

outside S5 S6 S7 S8 S9 10

P T D A NH2

cytoplasm -3

S1 S2 S3 S4

HO2 C

3 14 -

Colour plate 5.6 General topology of P-type ATPases. P-type pumps function in the membrane as a single polypeptide of around 100 kDa and contain 10 TMD. The TMD4-TMD5 cytoplasmic loop contains the ATP binding site and a conserved aspartyl residue that becomes phosphorylated and dephosphorylated during the catalytic cycle. The C-terminal residues function as an autoinhibitory domain and can also interact with 14-3-3 proteins that activate the ATPase.

ATP

ADP A B A B A A V 1 B H E C D G

cytoplasm membrane

a

lumen

H+

H+

F d cc c c V0 c

Colour plate 5.7 The vacuolar V 1 V 0 H+ -ATPase. Subunits of the rotor (V 0 ) domain are shown in grey. The V 1 stalk domains are shown in light green and the V 1 catalytic subunits (A and B) are is shown in dark green. Hydrolysis of ATP to ADP occurs at the catalytic domain and causes a rotational conformation change. This rotational movement is transduced via the stalk subunits to the rotor, which moves H+ from the cytoplasm to the vacuolar lumen.

lumen S1S2S3S4 S5S6S7S8 S9 101112 cytoplasm NH2

HO2C

Colour plate 5.8 Secondary structure of NHX type cation: H+ exchangers. NHX antiporters typically contain 10–12 TMD with a mass of 120–130 kDa. A conserved region in the third TMD binds the diuretic inhibitor amiloride and TMD sixth to seventh are thought to be critical for transport activity. Alternative models have also been suggested (see text).