Plant solute transport
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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 ...

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.

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Plant solute transport Plant solute transport Document Transcript

  • 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. I. Yeo, A. R. II. Flowers, T. J. (Timothy J.) QK871.P53 2007 571.2–dc22 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 General introduction 1 ANTHONY YEO 1.1 Introduction 1 1.2 Synopsis 3 1.3 Concluding remarks 14 Reference 14 2 Solutes: what are they, where are they and what do they do? 15 TIM FLOWERS 2.1 Solutes: inorganic and organic 15 2.2 Analysis of inorganic elements 15 2.2.1 Obtaining material for analysis 15 2.2.2 Optical methods 16 2.2.3 Mass spectrometry 16 2.2.4 X-ray fluorescence 17 2.2.5 Ion-specific electrodes 17 2.2.6 Ion chromatography 17 2.3 Solute concentrations 17 2.4 Organic compounds 18 2.5 Range of solutes found in plants 19 2.6 Localisation 19 2.6.1 Stereological analysis 19 2.6.2 Inorganic elements and electron microscopy 20 2.6.3 Ion-specific microelectrodes 21 2.6.4 Direct sampling 22 2.6.5 Use of fluorescent dyes 22 2.6.6 Flux analysis 23 2.6.7 Organic compounds 25 2.7 What do they do? 25 2.7.1 Vacuoles 25 2.7.2 Organelles and the cytoplasm 26 2.7.3 Cell walls 26 2.7.4 Conclusions 26 References 27
  • iv CONTENTS 3 The driving forces for water and solute movement 29 TIM FLOWERS and ANTHONY YEO 3.1 Introduction 29 3.2 Water 29 3.3 Free energy and the properties of solutions 31 3.3.1 Free energy and chemical potential 31 3.3.2 Water potential and water potential gradients 32 3.3.3 Osmosis and colligative properties 33 3.4 Cell water relations 34 3.5 Water movement 35 3.5.1 Water movement through the soil 38 3.5.2 Water in cell walls 39 3.5.3 Water movement across a root (or leaf) 39 3.5.4 Water movement through the xylem and phloem 40 3.6 Solute movement 40 3.6.1 Chemical, electrical and electrochemical potentials and gradients 41 3.6.2 Diffusion – Fick’s first law 41 3.6.3 Diffusion potential 42 3.6.4 Nernst potential 43 3.6.5 Donnan systems 43 3.6.6 Goldmann equation 44 3.7 Coupling of water and solute fluxes 44 References 45 4 Membrane structure and the study of solute transport across plant membranes 47 MATTHEW GILLIHAM 4.1 Introduction 47 4.2 Plant membranes 47 4.2.1 Plant membrane composition 47 4.2.2 Plant membrane structure 50 4.3 Studying solute transport across plant membranes 51 4.4 Transport techniques using intact or semi-intact plant tissue 52 4.4.1 Plant growth 52 4.4.1.1 Solution design 52 4.4.1.2 Using inhibitors 53 4.4.2 Accumulation and net uptake 53 4.4.3 Radioactive tracers 54 4.4.4 Fluorescent solute probes 55 4.4.5 Electrophysiology 57 4.4.5.1 Voltage-based measurements (membrane potential and ion concentration) 58 4.4.5.2 Voltage clamping 60 4.5 Using isolated membranes for transport studies 60
  • CONTENTS v 4.5.1 Isolating membranes 60 4.5.2 Assaying transport activities of protoplasts and membrane vesicles 61 4.6 Using molecular techniques to inform transport studies 63 4.6.1 Revealing the molecular identity of transporters and testing gene function 63 4.6.2 Location of transport proteins 64 4.6.3 Heterologous expression 65 4.7 Combining techniques (an example of increasing resolution and physiological context) 66 4.8 Future development 66 4.9 Conclusions 67 Acknowledgements 67 References 67 5 Transport across plant membranes 75 FRANS J. MAATHUIS 5.1 Introduction 75 5.1.1 Plant solutes 76 5.1.2 Definitions and terminology 76 5.1.3 Some formalisms 79 5.2 Passive transport 81 5.2.1 Diffusion through membranes 81 5.2.2 Facilitated diffusion through carriers 82 5.2.3 Transport through ion channels 83 5.2.3.1 Potassium channels 84 5.2.3.2 Calcium channels 85 5.2.3.3 Non-selective ion channels 85 5.2.3.4 Chloride channels 85 5.2.4 Transport through water channels 85 5.3 Primary active transport 87 5.3.1 Primary proton pumps 87 5.3.1.1 P-type ATPases 88 5.3.1.2 V-type ATPases 89 5.3.1.3 The pyrophosphatase 90 5.3.2 Primary pumps involved in metal transport 90 5.3.2.1 P-type Ca2+ pumps 90 5.3.2.2 Heavy metal ATPases 91 5.3.3 ABC transporters 92 5.4 Secondary active transport 92 5.4.1 Potassium uptake 93 5.4.2 Nitrate transport 94 5.4.3 Sodium efflux 95 5.4.4 Non H+ -coupled secondary transport 95
  • vi CONTENTS 5.5 Concluding remarks 96 References 96 6 Regulation of ion transporters 99 ANNA AMTMANN and MICHAEL R. BLATT 6.1 Introduction 99 6.2 Physiological situations requiring the regulation of ion transport 99 6.2.1 Change of cell volume 99 6.2.2 Nutrient acquisition 102 6.2.3 Stress responses 106 6.3 Molecular mechanism of regulation 107 6.3.1 Transcriptional regulation 108 6.3.2 Post-translational regulation 109 6.3.2.1 Autoinhibition 109 6.3.2.2 14-3-3 proteins 111 6.3.2.3 Calmodulin 113 6.3.2.4 Cyclic nucleotides 114 6.3.2.5 Heteromerisation 116 6.4 Traffic of ion transporters 117 6.5 Conclusions and outlook 120 References 120 7 Intracellular transport: solute transport in chloroplasts, mitochondria, peroxisomes and vacuoles, and between organelles 133 KATRIN PHILIPPAR and J ¨URGEN SOLL 7.1 Introduction 133 7.1.1 Research to identify solute transport proteins in plant organelles 133 7.1.1.1 Benefits of a model plant: Arabidopsis thaliana 134 7.2 Chloroplasts 136 7.2.1 The function of plastids 137 7.2.2 Transport across the outer envelope: general diffusion or regulated channels? 137 7.2.2.1 A porin in the outer envelope of plastids? 138 7.2.2.2 OEPs, a family of channels with substrate specificity 138 7.2.2.3 Outer membrane channels and porins: evolutionary aspects in chloroplasts and mitochondria 142 7.2.3 Transport across the inner envelope: phosphate translocators, major facilitators and carriers 142 7.2.3.1 The phosphate translocator family 142
  • CONTENTS vii 7.2.3.2 Major-facilitator-mediated transport 144 7.2.3.3 Carriers in the inner envelope of plastids 146 7.2.4 Transport across the inner envelope: ABC transporters and ion transport 147 7.2.4.1 ABC transporters 147 7.2.4.2 Ion transport 149 7.2.4.3 Transport of metal ions 150 7.3 Mitochondria 153 7.3.1 The function of plant mitochondria 153 7.3.2 Transport across the outer membrane: the porin VDAC 154 7.3.3 Transport across the inner membrane: carriers 156 7.3.3.1 Transporters involved in ATP production 156 7.3.3.2 Carriers for transport of TCA cycle intermediates 158 7.3.3.3 Amino acid transport across mitochondrial membranes 159 7.3.3.4 Carriers involved in β-oxidation of fatty acids 160 7.3.4 Transport across the inner membrane: ABC transporters and ion channels 160 7.3.4.1 ABC transporters 160 7.3.4.2 Ion channels 161 7.4 Peroxisomes 162 7.4.1 Function of peroxisomes in plant metabolism 163 7.4.2 Solute transport across the peroxisomal membrane 163 7.4.2.1 A porin in the peroxisomal membrane 163 7.4.2.2 Specific transport proteins in the peroxisomal membrane 165 7.5 Photorespiration: transport between plastids, mitochondria and peroxisomes 166 7.6 Vacuoles 167 7.6.1 Generating a pH gradient across the tonoplast: H+ -ATPase and H+ -pyrophosphatase 168 7.6.2 Transport of malate and sucrose across the tonoplast 170 7.6.2.1 Malate 170 7.6.2.2 Sucrose 171 7.6.3 Aquaporins and ABC transporter in the tonoplast 171 7.6.3.1 Aquaporins in the vacuole are tonoplast-intrinsic proteins 171 7.6.3.2 ABC transporters in the tonoplast 172 7.6.4 Ion transport 173 7.6.4.1 Ion channels 173 7.6.4.2 Calcium, sodium and magnesium uptake involves active transport 175 7.6.4.3 Transport of transition metals 177 References 178
  • viii CONTENTS 8 Ion uptake by plant roots 193 ROMOLA J. DAVENPORT 8.1 Introduction 193 8.2 Soil composition 193 8.3 Root exploration of the soil 194 8.4 Physical factors affecting root uptake: depletion zones and Donnan potentials 196 8.5 Radial transport of solutes across the outer part of the root 197 8.5.1 The role of apoplastic barriers 197 8.5.2 Root hairs and cortical cells 198 8.6 Solute uptake from different root zones 201 8.7 Transport of solutes to the xylem 203 8.8 The kinetics of solute uptake into roots 204 8.8.1 Radioisotopic studies 204 8.8.2 Other methods 207 8.8.3 Kinetics of uptake in response to solute availability 207 8.9 Conclusion 209 References 209 9 Transport from root to shoot 214 SERGEY SHABALA 9.1 Introduction 214 9.2 Transport of water 214 9.2.1 Xylem structure 214 9.2.2 Physics of water flow and evolutionary aspects of conduit development 216 9.2.3 Water flow between xylem elements: safety mechanisms 217 9.2.4 Hydraulics of the sap lift: general overview 219 9.2.5 Driving force for water movement in the xylem 221 9.2.6 Controversies and additional mechanisms 222 9.3 Transport of nutrients 224 9.3.1 General features of xylem ion loading 224 9.3.2 Ionic mechanisms of xylem loading 225 9.3.2.1 Potassium 225 9.3.2.2 Sodium 226 9.3.2.3 Anion channels 227 9.3.2.4 Gating factors 227 9.3.3 Xylem-sap composition 228 9.3.4 Factors affecting ion concentration in the xylem 229 9.3.5 Xylem unloading in leaves 230 References 231 10 Solute transport in the phloem 235 JEREMY PRITCHARD 10.1 Introduction 235
  • CONTENTS ix 10.2 Phloem anatomy 236 10.2.1 Sieve tubes 236 10.2.1.1 Sieve tubes are anucleate 236 10.2.1.2 Sieve plate blockage 237 10.2.2 Plasmodesmata 238 10.2.2.1 Plasmodesmatal structure 238 10.2.2.2 Plasmodesmatal selectivity 238 10.3 Phloem composition 240 10.3.1 Carbohydrate 240 10.3.1.1 Sucrose 240 10.3.1.2 Other carbohydrates 240 10.3.2 Inorganic ions 241 10.3.2.1 Variation in sieve element composition 241 10.3.2.2 K+ /sucrose reciprocity 242 10.3.3 Nitrogen 242 10.3.4 mRNA 243 10.3.4.1 Protein metabolism message 244 10.3.4.2 Structural genes and cell-wall enzymes 244 10.3.4.3 Interaction with DNA/RNA 245 10.3.4.4 Carbohydrate metabolism 245 10.3.4.5 Redox–oxidative stress 245 10.3.4.6 Amino acid metabolism 245 10.3.4.7 Transport 245 10.3.4.8 Interaction with the environment 246 10.3.5 Proteins 246 10.3.5.1 Oxidative stress 246 10.3.5.2 Defence 247 10.3.5.3 Calcium and sieve element structure 247 10.3.5.4 Metabolism 247 10.3.6 Macromolecular trafficking 248 10.4 Sieve element water relations 248 10.4.1 Sieve element water relations 249 10.4.1.1 Sieve element osmotic pressure 249 10.4.1.2 Sieve element turgor pressure 249 10.4.2 Flow in the phloem 250 10.4.3 Phloem loading 251 10.4.3.1 Symplastic or apoplastic loading? 251 10.4.3.2 Transporters facilitating apoplastic loading 254 10.4.3.3 H+ /ATPase 255 10.4.4 Phloem unloading 257 10.4.4.1 Evidence for unloading pathway: root tips 257 10.4.4.2 Evidence for unloading pathway: developing fruits 259 10.4.4.3 Evidence for unloading pathway: seed coats 259
  • x CONTENTS 10.4.5 Resource partitioning through the phloem 260 10.5 Exploitation by other organisms 261 10.5.1 Micro-organisms and viruses 261 10.5.2 Sap-feeding insects 261 10.5.3 Plants 262 10.5.4 Other organisms 262 10.6 Conclusions 262 References 263 11 Factors limiting the rate of supply of solutes to the root surface 275 ANTHONY YEO 11.1 Introduction 275 11.2 Supply of nutrients to the root surface 276 11.2.1 Absence of the nutrient element in the growth medium in any form 276 11.2.2 Bioavailability of the element 276 11.2.3 Movement of nutrients towards roots 278 11.2.4 Homogeneity or heterogeneity (spatial and temporal) in availability 279 11.2.5 Losses 279 11.3 Acquisition and uptake of nutrients by the root 280 11.3.1 Affinity and capacity of transport processes in the roots 280 11.3.2 Exploration and exploitation of soil volume by roots 282 11.4 Acquisition of phosphorus 284 11.5 Protected cropping systems: hydroponics as an example of ‘ideally’ controlled conditions 286 11.6 Concluding remarks 287 References 287 12 Mineral deficiency and toxicity 290 ANTHONY YEO 12.1 Introduction 290 12.1.1 Terminology 291 12.2 Deficiency and efficiency: iron in alkaline soils 293 12.2.1 ‘Strategy I’: reduction-dependent iron uptake 295 12.2.2 ‘Strategy II’: phytosiderophores 296 12.3 Phosphate uptake in soils that are low in phosphate 299 12.3.1 Cluster roots and root exudates 299 12.3.2 Mycorrhizal symbiosis 300 12.4 Toxicity and tolerance–aluminium in acid soils 301 12.5 Toxicity and tolerance–essential and non-essential metals 303 12.5.1 Hyperaccumulation 304 12.5.2 Ion transport in hyperaccumulators 305 12.5.3 Phytochelatins 306 12.5.4 Function of hyperaccumulation 308
  • CONTENTS xi 12.6 Concluding remarks 308 References 309 13 Water-limited conditions 314 ANTHONY YEO 13.1 Introduction 314 13.2 Plant responses to reduced water availability 315 13.3 Mechanisms to reduce water loss: regulation of stomata and regulation of leaf area 318 13.3.1 Stomatal regulation 318 13.3.2 Leaf area regulation 320 13.3.3 Consequences: interaction with leaf temperature 321 13.4 Mechanisms to maintain water potential gradients: osmotic adjustment 322 13.4.1 Water potential of drying soil 322 13.4.2 Osmotic adjustment 323 13.4.3 Compatible solutes/osmolytes/osmoprotectants 324 13.4.4 Water movement from protoplast to apoplast in freezing injury 326 13.5 Mechanisms to acquire more water: root properties 326 13.5.1 Constitutive formation of deep roots 326 13.5.2 Facultative formation of deep roots 327 13.5.3 Root conductance 327 13.6 Mechanisms to increase water-use efficiency: C4 and crassulacean acid metabolism (CAM) 328 13.6.1 C4 photosynthesis 329 13.6.2 CAM 331 13.7 Gene regulation 334 13.8 Concluding remarks 335 References 335 14 Salinity 340 ANTHONY YEO 14.1 Introduction 340 14.2 External concentration of salt up to about 50 mM NaCl 341 14.3 External concentration of salt up to about 100–150 mM NaCl 343 14.4 External concentration of salt above about 150–200 mM 344 14.5 ‘Molecular’ tolerance 345 14.6 Cellular tolerance 346 14.7 Moving on to a cell in a plant 347 14.8 Salt glands 347 14.9 Selectivity at the root 348 14.9.1 Root selectivity for chloride 353 14.10 Transport from root to shoot 353 14.10.1 Transport of chloride to the xylem 356
  • xii CONTENTS 14.11 Transport from shoot to root 356 14.12 Leaf cells 357 14.13 Prospects 361 14.14 Concluding remarks 364 References 365 15 Desiccation tolerance 371 ANTHONY YEO 15.1 Introduction 371 15.2 Occurrence of desiccation tolerance 372 15.3 Desiccation tolerance in seeds 372 15.3.1 Intracellular physical characteristics 374 15.3.2 Intracellular de-differentiation 374 15.3.3 ‘Switching-off’ metabolism 375 15.3.4 Antioxidant systems 375 15.3.5 Protective molecules 376 15.3.6 Amphiphilic molecules 378 15.3.7 Oleosins 379 15.3.8 Damage repair 379 15.4 Vegetative tissues 379 15.4.1 Gene expression 382 15.4.2 Physical characteristics 382 15.4.3 Metabolism and antioxidants 383 15.4.4 Low-molecular-weight carbohydrates 383 15.4.5 Hydrins or LEA proteins 385 15.4.6 Signals 385 15.4.7 Constraints to the development of desiccation tolerance 386 15.5 Concluding remarks 388 Acknowledgements 388 References 388 Index 391 The colour plate section appears after page 78
  • 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 indi- vidual 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 in- organic 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 cel- lular 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 compartmentalisa- tion 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 con- version 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, syn- thesis 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 Mole- cular 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-Maximilians- Universit¨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, Pri- vate Bag 54, Hobart, Tasmania, 7001, Australia Professor J¨urgen Soll Department Biologie I, Botanik, Ludwig-Maximilians- Universit¨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,BN19QG,UKandSchoolofPlantBiology,35StirlingHighway,Crawley, Western Australia 6009, Australia. ARY current address: aryeo@aol.com
  • 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 substan- tially 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 concen- tration 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 concen- tration 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 no- win 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 pres- sure. 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 ma- jority 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 diox- ide, 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 wa- ter 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 accu- mulation 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 pro- cesses to go on within the same cell. The concentrations of soluble metabolic inter- mediates 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-to- alkaline 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 py- rophosphatases, 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 spec- troscopy), mass spectroscopy, X-ray fluorescence, with ion-specific electrodes, and by ion chromatography. Analysis of organic solutes is usually achieved by chro- matographic separation, often in conjunction with mass spectroscopy and nuclear magnetic resonance. Intracellular localisation can be achieved either via transmis- sion or scanning electron microscopy preceded by precipitation, freezing or freeze- substitution. 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 accu- mulate 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 deriva- tion 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 de- velops 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 so- lutes 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 cur- rently 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 be- ginning 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 em- phasis on design and composition of experimental solutions, particularly their os- molarity, 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 in- cluding 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 iso- lation 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 sec- ond), 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 chloro- plasts. 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 car- riers 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 potas- sium 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 degra- dation, 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 reg- ulation, 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 componentsofthetranscriptionalregulationofnutrienttransportersaresummarised. 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 or- ganelles 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 elec- tron 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 sul- phate 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 phos- phorylation – 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 mem- brane. 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 β- oxidation and are part of the photorespiratory cycle; they also generate reactive oxygen species and contain appropriate protective mechanisms. Glyoxisomes con- vert lipid reserves to sucrose. The peroxisome family has a ‘specific porin’ as well as transporter proteins including the peroxisomal ATP/ADP carrier. The pho- torespiratory pathway is split between the chloroplast, mitochondrion and per- oxisome. Vacuoles are multifunctional and are involved in the storage of different metabo- lites, quantitatively extreme examples being malate (in CAM) and sucrose (in stor- age 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 mediat- ing 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 nu- trients 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 transpira- tional 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 cy- toplasmic 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 inter- connected 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. Sam- pling 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 chan- nels. 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 againstoxidativedamage.Theothermaincomponentofthephloemisthecompanion 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 con- tents 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. En- try 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 vol- umes 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 Km values for high-affinity transporters are often in the μM range. Maintain- ing 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 detox- ify 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 home- ostasis for essential metals, can cope with low-level chronic exposure, the hyperac- cumulators function by compartmentalisation of metals in the vacuole. There is evi- dence 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 stomatalconductancetoconstitutiveanatomicalmodificationsseenindesertspecies.
  • 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 C4 pho- tosynthesis through the storage and transport of fixed carbon as malate. Both provide substantial increase in water use efficiency, and C4 photosynthesis is associated with high productivity. Water deficit has been shown to affect the expression of numerous genes. Sig- nificantly, the way in which deficit is applied accounts for most of the differences in expression. This underlines the essentiality of applying treatments that are phys- iologically relevant. Commonly-affected genes were generally involved in down- regulation of growth, again emphasising the difference between agricultural objec- tives 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 retransloca- tion) 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. Compartmentalisa- tion 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. Desic- cation 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 chem- istry 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 (shrink- age), 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 phys- ical 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. Pro- tective 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 des- iccation 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 an- tioxidants, and on solutes, including LEA proteins, that can act in water replacement and vitrification. Desiccation tolerance in both seeds and vascular plants is slow, co- ordinated 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 ex- plain 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.
  • 14 PLANT SOLUTE TRANSPORT 1.3 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 va- porising 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 mate- rial 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.
  • 16 PLANT SOLUTE TRANSPORT 2.2.2 Optical methods When vaporised (atomised) in a flame, atoms emit light, as electrons that have ab- sorbed 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 el- ements 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 pho- tomultiplier) 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 wave- length 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 detec- tor 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 NH4 + , Ba2+ , Ca2+ , Cd2+ , Cu2+ , Pb2+ , Hg2+ , K+ , Na+ , Ag+ , Br− , CO3 2− , Cl− , CN− , F− , I− , NO3 − , NO2 − , ClO4 − , 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 activi- ties (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) con- taining 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: aj = λj cj (2.1) where γj 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 inor- ganic 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 com- pounds. 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 energyorwavelength)canbeusedtoidentifyandquantifytheelementalcomposition 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 trans- mission 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 micro- scopes 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 po- sition 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 re- ports 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 retain- ing water-soluble solutes, the binding of solutes from other compartments to the chloroplast envelope is an important source of potential contamination. Interest- ingly, chloride is retained in the chloroplasts even after aqueous preparation (see Flowers, 1988), suggesting that the envelope keeps its ability to retain solutes dur- ing 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
  • SOLUTES: WHAT ARE THEY, WHERE ARE THEY AND WHAT DO THEY DO 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 ben- zofuran 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 membranesthatrepresentaresistancetothemovementofsolutes–theplasmalemma 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 PLANT SOLUTE TRANSPORT A B C 5.2 5.7 6.2 0 50 100 4 4.5 5 5.5 6 5 5.5 6 0 0.5 1 1.5 2 2.5 0 5 10 15 20 25 Time (minutes) Logarithmofremaining activity Logarithmofremaining activity Logarithmofremaining activity 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 CaCl2, 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 de- termine 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 arabidop- sis 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 de- tected 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 at- tachment of electron-dense gold particles to the antibody (see Hajibagheri, 1999). In most published cases, the technique has been used to localise proteins: recent exam- ples 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 salt- tolerant 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 io- dine 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 com- partments 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 sta- bility 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 mag- nesium 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 os- moprotectants 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. So- lutes 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). Under- standing 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.
  • 28 PLANT SOLUTE TRANSPORT 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 compart- mentation 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 Pub- lishing, Collingwood, Victoria, Australia. Riens, B., Lohaus, G., Heineke, D. and Heldt, H.W. (1991) Amino-acid and sucrose content determined inthecytosolic,chloroplastic,andvacuolarcompartmentsandinthephloemsapofspinachleaves. 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 embark- ing 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 non- polar, 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 incom- pressible, 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. Thermo- dynamic 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 Sec- tion 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 mem- brane 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 con- ditions, liquid water enters the system following rainfall – it may be distant rain- fall 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). Tem- perature 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 tempera- tures 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 (μw) is given by the equation: μw = ∂G ∂nw T,P,E,h,n j (3.1) where nw 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 μw 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 (aw), pressure (P) and height (h) in a gravitational field. This is expressed mathematically as: μw = μ∗ w + RT ln aw + ¯Vw P + mwgh (3.2) where R is the gas constant, T is the absolute temperature, aw is the activity of the water, ¯Vw is the partial molal volume of water (see below), P is the pressure, mw 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 aw = 1 (RT ln aw = 0), P = 1 and the height in the gravitational field is zero (mwgh = 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 ( ¯Vw) 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. ¯Vw = (∂V/∂nw)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 ¯Vw (3.3) The term ¯Vw (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 − + ρwgh (3.4) where is the osmotic pressure −(RT/ ¯Vw) ln aw and ρw is the density of water (mw/ ¯Vw).
  • 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 deep- rooted 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 – thosefavouredbyphysiologistssincethediscoveryofosmoticpressure.Moredetails 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/ ¯Vw) 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 cs 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 temper- ature 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 be- tween solvent (water) and solute. However, where solute passes through the mem- brane to some degree, the effective osmotic pressure is reduced. It is easy to imag- ine 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 semiper- meable membranes (plasma membrane and tonoplast) plus small organelles embed- ded 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 o w = 0 = i w = 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 = mwgh ¯Vw (3.8) where mw is mass per mole (18.016 g mol−1 ), g is the acceleration due to grav- ity (9.8 m s−2 ), h is the vertical height in m and ¯Vw 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 contin- uum 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 ¯Vw, 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 −0.1 Root −0.2 Shoot −0.5 Atmoshere 75% rh −38.9a Atmosphere 50% rh −93.6a a The relationship between the water potential of water vapour in the atmosphere and the relative humidity is given by: w = RT ¯Vw ln rh 100 (3.10) where rh is the relative humidity (%), R is the gas constant (8.314 J mol−1 deg−1 ), T is the temperature (K) and ¯Vw 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 hu- midity 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: JVw = ψw Rw (3.11a) JVw = w Lw (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 ), Lw is a water conductivity coefficient and Rw is a resistance to water flow. To calculate the total flux, JVw must be multiplied by the area A.
  • THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT 37 Atmospheric Atmospheric water potential resistance Leaf water potential Xylem Root water potential Root resistance resistance Soil water potential Ψ Ψ Ψ Ψ a Ra l Rx r s Rr w w w w 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 (mea- suredinMPasm−3 ;thereciprocalofconductance)inthevarioussegmentsoftheflux 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 Jv, the quantity of water entering equals that leaving, or Aroot Jin v = Aleaf Jout v . 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: Jroot V Aroot = xylem w − root w Rroot = J xyl V Axyl = root w − leaf w Rxylem = leaf w − atmosphere w Ratmosphere (3.12) Inserting representative figures from Table 3.1 and rearranging to solve for Ratmosphere: Ratmosphere = leaf w − atmosphere w shoot w − leaf w (Rroot + Rxylem) = −0.5 − (−38.9) −0.2 − (−0.5) (Rroot + Rxylem) = 113(Rroot + Rxylem) This simple calculation illustrates a very important aspect of flow through the con- tinuum that the resistance in the gas phase is significantly greater than the sum of the other plant resistances. Ratmosphere 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: P = −σ 1 r1 + 1 r2 (3.13) where σ is the surface tension of water and r1 and r2 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 (Jv) depends on the gradient of soil water potential with distance, viz.: Jv = −Ls d s dx (3.14)
  • THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT 39 where the soil water potential s is given by s = s p + s G – 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 s G and in dry soils by s p. 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 cellwallsoftheepidermalcells:theserepresentasimilarmediumtothesoilforwater 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 wellasforitslongitudinalflow.Themajorityofradialmovementoccursfromthesoil 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).Thesebarriersnotonlypreventunwantedingressofsolutes,butperhaps,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 8η δP δx (3.15) where Jv 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 posi- tive, 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.
  • THE DRIVING FORCES FOR WATER AND SOLUTE MOVEMENT 41 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: μj = μ∗ j + RT ln aj + 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 con- centration 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.: Jj = −Dj δcj δx (3.17) where δcj /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 −δcj /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. (co j − ci j )/ 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
  • 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 co j − ci j , where Kj 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: Jj = Pj co j − ci j (3.18) 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 coef- ficient 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 dimen- sions 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, Kj and x may be uncertain, Pj is a readily mea- surable 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 co j and ci j 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 PNa/PK is about 0.2 and PCl/PK, 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, μo j = μi j , whence, expanding using Eq. 3.16: μ∗ j + RT ln ao j + z j FEo = μ∗ j + RT ln ai j + z j FEi (3.20) The potential at equilibrium ENj (named after Nernst, who first derived this rela- tionship) is given by ENj = Ei − Eo . Rearranging Eq. 3.20, z j F(Ei − Eo ) = RT ln ao j − ln ai j = RT ln ao j ai j whence ENj = Ei − Eo = RT z j F ln ao j ai j = RT z j F 2.303 log ao j ai j At 25◦ C for a monovalent cation, this reduces to: ENj = 59.2 log co ci (3.21) Equation 3.21 demonstrates the poise between electrical potential and chemical con- centration 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+ (mM) 0.5 45 – – EN(K) (mV) 0 −57 EN(K) (mV) 0 −126 EN(K) (mV) 0 −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 (EM) is given by: EM = RT ln PKco K + PNaco Na + PClci Cl PKci K + PNaci Na + PClco Cl (3.22) 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: Jj = L j1 X1 + L j2 X2 + · · · + L jn Xn (3.23) where L is a coefficient. For fluxes of water (Jw) and solute (Js), the equations reduce to: Jw = Lww μw + Lws μs (3.24) and Js = Lsw μw + Lss μs (3.25) 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 coef- ficients is, in practice, 3 as it can be shown that Lws = Lsw. 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 (JV), the differences between the mean velocities of solute and water (JD) and the differences in hydrostatic pressure ( P) and turgor pressure ( ) across a membrane: JV = LP P + LPD (3.26) JD = LDP P + LD (3.27) where LP, LPD, LDP and LD are the Onsager coefficients and LP is the hydraulic conductivity of the membrane. These coefficients have been used to define an im- portant parameter, the reflection coefficient σ (=− LDP LP =− LPD LP ). This can be seen as the proportion of solute molecules ‘reflected’ by the membrane: for a perfect semiper- meable 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 Physi- ology. Cambridge University Press, Cambridge, UK. Kedem, O., Katchalsky, A. (1958) Thermodynamic analysis of the permeability of biological mem- branes 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 Publish- ers, 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 func- tion, 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 combi- nation 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 Plant membranes 4.2.1 Plant membrane composition All eukaryotic organisms have lipid- and protein-rich bilayers that delineate indi- vidual cells and compartmentalise intracellular regions into distinct organelles or other membrane-bound subcompartments. These membranes both form and pro- vide 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 intra- cellular 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 PLANT SOLUTE TRANSPORT Distinctive transport proteins Connectivity/ Membrane Main roles Trafficking Nuclear envelope RNA/protein transport Endoplasmic reticulum Synthesising, sorting and processing proteins Golgi Protein, vesicle trafficking Protein, vesicle trafficking Protein, vesicle trafficking Protein, vesicle trafficking Protein, vesicle trafficking trans-Golgi network Partially coated reticulum Multivesicular body Secretory vesicle Plasma membrane Signalling, cell wall synthesis, homeostasis P-type H+ -ATPase, PIP1 Endocytic vesicle Protein, vesicle trafficking Transport vesicle Protein, vesicle trafficking Tonoplast Storage vacuole Lytic Undefined Storage of metabolites and toxins, homeostasis, pigmentation Lysis TIPs, V-PPase TIP2 (α-TIP) TIP1 (λ-TIP) TIP3 (δ-TIP) not TIP1/2 Mitochondria ATP synthesis Inner ATP/ADP translocator Outer Porins Chloroplast envelope Photosynthesis Inner VDAC Outer Porin Thylakoid P-type Cu2+ -ATPase Symbiotic membrane, e.g. symbiosome N2 fixation NOD26-like Perioxisomes Lipid mobilisation, glycolate pathway, perioxidation Figure 4.1 Major plant membranes, their roles, major transport proteins and connectivity. —, con- nection 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 plasmodes- mata, 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 traf- ficking 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 (Staehe- lin 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], phosphatidylser- ine [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 com- ponents. 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 mem- branes, 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 compo- nents. PS and CL are the distinctive lipids of the mitochondria but in phosphate- deprived 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 ef- ficiency (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 24- methylcholesterol) 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 inti- mately 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-boundandfacilitatesthepotentialformationoflargesoluteconcentration gradients across the bilayer. Only highly lipid-soluble (e.g. ethanol, glycerol), small non-polar (e.g. O2, CO2) and some small polar (e.g. H2O, 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 andtheirpresencecontrolledbycytoticevents,bothperipheral(linkedbysaltbridges to other proteins or lipids) and lipid-linked proteins (e.g. fatty-acid-, prenyl-group- and 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 dif- fer 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 physio- logical conditions. The fluidity of lipid bilayers is naturally temperature-dependent (they will undergo a liquid-crystal to gel-like phase transition as temperature in- creases). 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 interac- tions 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 endomem- branes,resembleamosaicofmicrodomainswithaparticularmolecularcomposition. 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 ‘liquid- disordered’ 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
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 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 func- tional 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 regu- late 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, proto- cols and databases on the internet, as well as the recently sequenced and anno- tated genomes of multiple organisms, has provided the tools to mine data effec- tively 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 generaltechniquescanbeapplied,andsotheremainingchapterhasbeendividedinto 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
  • 52 PLANT SOLUTE TRANSPORT 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 consid- ered 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 orien- tated 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 (ax; see Section 2.3), especially at high concentrations (a square bracket is com- monly 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+ pas- sage 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 Vi- sual 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 differ- ence 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).
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 53 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 (Ps) and hydraulic conductivity (Lp) (Tyree et al., 2005). Furthermore, the electrostatic interactions of ions with the outer membrane surface charge (ψo 0 ) will affect the concentration of charged species at the membrane surface compared to that of the bulk solution (Kin- raide, 2004). ψo 0 , 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, highionicstrength,highlycharge-denseionsoralowpHwillinduceslightlypositive values (Kinraide, 2001). Most studies ignore the contribution of ψo 0 to Em 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 ψo 0 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 ap- proach 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 im- ply 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,
  • 54 PLANT SOLUTE TRANSPORT 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 differ- entiate 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 composi- tions) include nuclear magnetic resonance (NMR; see Section 2.4) and X-ray mi- croanalysis (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 perme- ability (Pf), Ps and Lp 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 con- taining a proportion of a particular solute as a radioactive isotope, e.g. 43 K, 86 Rb, 22 Na, 45 Ca, 36 Cl, 36 ClO3 (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 ki- netics 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 pa- rameters (Km and Vmax) (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 Vmax is high and Km is low, as the presence of a significant unstirred layer may
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 55 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 so- lute 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 mod- els 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 ques- tioned 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 mea- suring 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 autoflu- orescence (e.g. chlorophyll, NADH). The fluorescent properties of probes measure the activity of target solutes through two basic mechanisms following substrate
  • 56 PLANT SOLUTE TRANSPORT 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-) quantifica- tion after in vivo calibration. Ratiometric probes negate any heterogeneity in mea- surement 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 neces- sary 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 in- vestigated 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). Load- ing of probes into cells across the PM and cellular compartments provides a more significant challenge. Membrane-permeant probes can be loaded by vacuum filtra- tion 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 exper- iments, 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 mi- croinjection (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.,
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 57 1995), but is extremely technically demanding and has low throughput. Recent de- velopment 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 re- leases 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 com- partment (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 target- ing of apoaequorin to compartments. [Ca2+ ] can be imaged fairly non-invasively in whole plants, detached tissues or protoplasts, with relatively poor temporal res- olution (>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
  • 58 PLANT SOLUTE TRANSPORT series with resistors. The compound electrochemical potential gradient (as a conse- quence of ion concentration equalities) across a membrane (membrane potential – Em) 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 tech- nological instructions should then be adequate to carry out many procedures. Several standard texts are referred to for a complete grounding in electrophysiological the- ory 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 environ- ment 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. Un- fortunately, 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 elec- troneutral 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) Em (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. Em 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 Em (Ogden, 1994).
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 59 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 O2, 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 NO3 − and organic anions, and K+ from NH4 + . 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 Em in addition to [x]in. Therefore, both an ISE and Em electrode have to be inserted into the same membrane compartment and Em 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’ Em 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 use- ful to ascertain which membrane compartment the electrode has penetrated. As the vacuole usually inhabits the majority of the cell interior, impalements may pene- trate the tonoplast; therefore any Em reading will be a compound measurement of both that of the tonoplast and PM. A combined pH and Em 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 mea- sured in an extracellular K+ of 1 mM; cf. Section 2.6.3). Em measurement has also been informative in understanding the electrogenic basis of stimulus-invoked mem- brane 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 distanceawayfromthesourceoverasetperiodoftime,andtheparametersintegrated using Fick’s law of diffusion (Sections 3.6.2 and 5.1.3) or an equivalent electrochem- ical 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). Cau- tion 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 ex- ternal ISE-based flux measurement can be used to study the non-electrogenic move- ment of solutes across membranes if the relevant sensor is available.
  • 60 PLANT SOLUTE TRANSPORT 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 (rela- tively) 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 Em it is possible to observe the passage of charged solutes across membranes as current (Ii) over time (down to the level of a single channel). Vm can be fixed at a nominated level and compared to Em across the poles of a feedback amplifier, the difference being due to Ii (the flow of ions across the membrane). The feedback circuit in- jects a current (If) equal and opposite to Ii and all parameters can be recorded by computer. Ii can be similarly clamped to observe the change in voltage. Vm (or Ii) clamping can be performed on larger cells using two intracellular microelectrodes (TEVC) or by using one electrode (dSEVC). Vm clamping therefore allows the char- acteristics of transporters (conductance, gating and selectivity) to be identified over the physiological Em 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 un- derutilized in plant sciences, Ii clamping could be useful in studying ligand-gated ion channel activity in intact or isolated tissues. 4.5 Using isolated membranes for transport studies 4.5.1 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 obviousinfluenceinsignallingorthecreationofunstirredlayers)canberemovedand 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 protectagainstphenolicoxidases,polyvinylpyrrolidone,ascorbateandmetabisulfite 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
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 61 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 or- ganelles, or membranes can be fractionated directly and there are established pro- tocols 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, immuno- logical 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 scat- tering (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 trans- port 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 electropora- tion and detergent permeabilisation, although these often make membranes ‘leaky’. Stopped-flow spectrophotometers allow quick solution exchanges (<1 ms) in small
  • 62 PLANT SOLUTE TRANSPORT 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, Pf or Ps 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 inves- tigating 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 Vm, 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, Erev). 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 elec- trogenic 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 trans- porters). Patches can be in an either ‘inside-out’ (internal membrane side facing the bathing solution) or ‘outside-out’ (outside facing the bathing solution) configu- ration. A ‘cell-attached’ configuration, although having the advantage that cellular contents remain undiluted, is infrequently used as Vm 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 Erev (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 con- centrations whilst simultaneously controlling Vm. This can reveal whether certain membrane currents on particular membranes are responsible for specific ion in- creases (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
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 63 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 pro- tein 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 elec- trogenic 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 observ- ing 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 chargeorstructuraleffectsmayaffectchannelfunction;ithasalsobeenobservedthat some lipids can act as ionophores and conduct ion currents (M. Tester, unpublished data). 4.6 Using molecular techniques to inform transport studies 4.6.1 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, in- valuable 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-of- function’ (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
  • 64 PLANT SOLUTE TRANSPORT ‘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 criti- cal 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’ mu- tants 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 pheno- type 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 Sec- tion 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 pro- tein (in addition to the transport activity it catalyses) is its spatial and temporal expression. Fractionation has revealed valuable information about transport activi- ties 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 ex- traction 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
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 65 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 re- action 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. aqua- porins; 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 tech- niques 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 com- mon, but has been used to identify and characterise several plant proteins (e.g. Uozumi, 2001). The most utilised system has been yeast (Saccharomyces ceriv- isiae, 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).
  • 66 PLANT SOLUTE TRANSPORT Immature eggs (or oocytes) isolated from ovaries of the South African clawed toad (Xenopus laevis) provide another popular system for characterising trans- porters. X. laevis oocytes can synthesise proteins from injected cDNA (into the nucleus) or cRNA (cytoplasm) and target plant proteins to the PM (subcellular tar- geting 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 as- says 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 overex- pression 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 be- coming 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¨ornroth- Horsefield et al., 2006). However, it is becoming apparent that the signalling net- works that regulate many transport processes, when integrated into the physiological framework of a functional cell (or plant), are too complex to comprehend without an
  • SOLUTE TRANSPORT ACROSS PLANT MEMBRANES 67 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 physiol- ogy. 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, ex- pertise and equipment (or its accessibility). All of these may be solved through collaboration (although boundaries and expectations should be clearly defined be- fore entering into any such relationship). It would have been impossible to describe all techniques with sufficient detail for readers to fully understand any one tech- nique 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’ De- partment of Plant Sciences, University of Cambridge for being a source of knowledge and inspiration over the past 8 years. References Alexandre, J. and Lassalles, J.P. (1992) Intracellular Ca2+ release by InsP3 in plants and effect of buffers on Ca2+ diffusion. Philosophical Transactions of the Royal Society of London Series B-Biological Sciences 338, 53–61. Allen, G.J., Kwak, J.M., Chu, S.P., et al. (1999) Cameleon calcium indicator reports cytoplasmic calcium dynamics in Arabidopsis guard cells. Plant Journal 19, 735–747. Alleva, K., Niemietz, C.M., Maurel, C., Parisi, M., Tyerman, S.D. and Amodeo, G. (2006) Plasma membrane of Beta vulgaris storage root shows high water channel activity regulated by cyto- plasmic pH and a dual range of calcium concentrations. Journal of Experimental Botany 57, 609–621.
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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 physiol- ogy, 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 sug- gestion 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 in- terspersed 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, be- tween ‘life’ and ‘death’. The selectivity and transport functions of membranes en- sure 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, mito- chondrial (see Section 7.3), chloroplast (see Section 7.2) and peroxisomal (see Sec- tion 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
  • 76 PLANT SOLUTE TRANSPORT the other hand, xylem parenchyma cells that transfer minerals to the xylem for subsequenttranslocationtotheshoot,aremorelikelytomediateaneteffluxfunction. 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 similarirrespectiveofthecelltype,speciesoreventhekingdomsoflife.Forexample, the conversion of trans-membrane H+ gradients into chemical energy in the form of ATP is mediated by F1F0 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+ , NH4 + , Ca2+ and Mg2+ ) and anions (e.g. NO3 − , Cl− , PO4 3− and SO4 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. Of- ten 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 F0 part of the F1F0 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 conven- tion adopted here will be that any membrane protein involved in cross-membrane movement of substrates is a transporter.
  • TRANSPORT ACROSS PLANT MEMBRANES 77 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’.
  • 78 PLANT SOLUTE TRANSPORT 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 de- scribing 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 dif- ferent 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 frac- tion 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 consid- erably 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 geom- etry 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
  • TRANSPORT ACROSS PLANT MEMBRANES 79 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 sub- strates 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 chan- nels 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 ex- changes 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 funda- mental 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 extracel- lular 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 (Em) 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 cy- toplasmic 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.
  • 80 PLANT SOLUTE TRANSPORT 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) whichintuitivelystatesthattheflux Js ofasolutesdependsonitsdiffusioncoefficient 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 = −μsCs dE dx , (5.2) where μs is a constant analogous to Ds and dE/dx is the electrical gradient or elec- trical 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 dx − μsCs dE 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, zs, included to yield
  • TRANSPORT ACROSS PLANT MEMBRANES 81 a relationship that expresses ionic current as a function of concentration and voltage: Is = −zs FDs dCs dx + FzsCs RT dE dx , (5.4) 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): E = RT zs F ln S+ out [S+]in , (5.5) 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 EK for potassium or ENa for sodium. We can rearrange Eq. 5.5 and substitute room temperature and z = 1 to get: E = 58 log I+ out [I+]in , (5.6) 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 Em 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 fre- quently applied to assess selectivity of membranes or of specific transport systems (Chapter 3). The so called Goldman Hodgkin Katz voltage equation gives the mem- brane potential (Erev) 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 PK, PNa and PCl according to: Erev = RT zF ln PK[K]o + PNa[Na]o + PCl[Cl]i PK[K]i + PNa[Na]i + PCl[Cl]o . (5.7) 5.2 Passive transport 5.2.1 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
  • 82 PLANT SOLUTE TRANSPORT 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 in- clude important compounds like O2, CO2, H2O2 and NH3. For example O2–CO2 gas exchange in lungs and in photosynthesising plant tissues operates by this pro- cess (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 pro- teins (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. Organ- isms 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 rela- tively 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 (NO3 − ) or ammonium (NH4 + ) ions, or a mixture of the two. The translocation of NH4 + 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 NH4 + 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 NH4 + 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
  • TRANSPORT ACROSS PLANT MEMBRANES 83 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 func- tion 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 attrac- tive 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 conduc- tance, channel selectivity and channel gating (Hille 2001). Conductance G is the reciprocal of resistance R defined in Ohm’s law as: V = I R 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
  • 84 PLANT SOLUTE TRANSPORT 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 Em. A decrease in membrane polarisation, depolarisa- tion, 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 Em to chan- nel 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 Em, 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 Em, 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 (lit- erally meaning a ‘competitor’) is also frequently used since often different ligands can have similar actions and compete for the same binding site. In contrast, antago- nists 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 require- ment 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 Em to transit from closed to open state. Functional channels are made up of four subunits either as homomers
  • TRANSPORT ACROSS PLANT MEMBRANES 85 or as heteromers. Apart from Shaker-like channels, the major families of plant K+ channelsalsocontaintwoorfourTMD-structuredchannelsandvoltage-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 IP3, 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).Althoughsomenon-selectivecationchannelshavebeenshowntoberegulated 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 Em with a Po 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 NO3 − (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
  • 86 PLANT SOLUTE TRANSPORT 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 Shaker- type 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 selectiveforwatermolecules,whichmovethroughtheproteininasinglefile,andthat 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 smallnon-chargedsolutessuchasglycerolandurea.Sometimesadistinctionismade between water-selective aquaporins and those that can conduct water and small so- lutes, 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 mem- branes 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 (Jo- hanson 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
  • TRANSPORT ACROSS PLANT MEMBRANES 87 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 diur- nal 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 phosphoryla- tion 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 Inmanyconditions,cellsrequiresolutesinquantitiesthatcannotbeachievedthrough passive transport. Similarly, many waste products need to be removed or compart- mentalised often against their electrochemical potential. This transport can, by def- inition, not occur via passive transport and requires active systems that directly use energy.Afurtherfundamentalrequirementofalllivingcellsisthecapacitytoconvert chemical energy into electrochemical energy and vice versa. Primary active trans- port 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 respi- ration 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
  • 88 PLANT SOLUTE TRANSPORT 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 sub- sequent 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 en- zyme 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 ac- tivity 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 contain- ing 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 expres- sion 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
  • TRANSPORT ACROSS PLANT MEMBRANES 89 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, V1, where ATP is bound and split. The basal part, V0, of the complex protrudes through the membrane and forms the H+ channel. In F1–F0 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 (V1) and electrochemical (V0) energy is precisely structured in V-ATPases is unknown but it is believed that V-type ATPases are very similar to F1–F0 ATP synthases, the enzymes found in mitochondria and chloroplasts that convert redox energy into chemicalenergyintheformofATP(Klugeetal.,2003).V-typeATPasesarebelieved to function as rotary motors, similar to their F1–F0 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 non- ionic 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, Em 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 tissue- and 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
  • 90 PLANT SOLUTE TRANSPORT 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 over- expression 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 cofac- tors 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+
  • TRANSPORT ACROSS PLANT MEMBRANES 91 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 Km 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 mem- bers 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 CPx- ATPases 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+ AT- Pases (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.
  • 92 PLANT SOLUTE TRANSPORT 5.3.3 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. Struc- turally, 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 con- tains 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 re- moval 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 conun- drums 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. Energi- sation 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
  • TRANSPORT ACROSS PLANT MEMBRANES 93 Na+ gradient and hence secondary transport is generally Na+ coupled. Plants, how- ever, 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 reac- tions 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 nu- trients, 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: Em = 60 log K+ out 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 Em, the respective ion gradients, and the coupling ratio n: G = (n + 1)FEm + RT ln H+ n out K+ out H+ n in K+ in . (5.10) At equilibrium G = 0 and Em = RT (n + 1)F ln H+ n out K+ out H+ n in K+ in . (5.11)
  • 94 PLANT SOLUTE TRANSPORT 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 Em 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 ex- pression 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 NH4 + and NO3 − of which NO3 − 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 NO3 − 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 NO3 − . If unity is substituted for n then the overall charge becomes zero and NO3 − uptake has to rely entirely on the pH as the driving force. With a pH of 2 and typical cytoplasmic NO3 − 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 NO3 − concentrations. In addition, measurements of root cell membrane potentials and voltage clamp experiments with nitrate-transporter- expressing oocytes have shown that the membrane depolarises in response to an increase in external NO3 − . These observations strongly indicate that entry of nitrate into the symplast carries positive charge and it is therefore assumed that 2 H+ accompany each NO3 − taken up through a NO3 − :2H+ symporter (Meharg and Blatt, 1995). Two main classes of genes have been identified that are involved in various forms of NO3 − uptake. These are the NRT1 and NRT2 gene families, both encoding proteinswithasixplussixTMDstructure.Asacoarsegeneralisation,theNRT1gene products are involved in low-affinity NO3 − uptake, whereas NRT2 genes encode proteins that mediate both constitutive and inducible high-affinity uptake capacity (Orsel et al., 2002).
  • TRANSPORT ACROSS PLANT MEMBRANES 95 5.4.3 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 con- centrations 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: signifi- cantly 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 there- fore coupled to the H+ gradient. Both at the tonoplast and the plasmamembrane H+ :Na+ antiport mechanisms have been characterised that participate in export- ing 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 accumula- tion 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. How- ever, there is convincing evidence that it occurs for some substrates such as urea, NO3 − and K+ . Na+ :urea symport has only been observed in charophytes and never in higher plants (Walker and Sanders, 1991), whereas Na+ -coupled NO3 − 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
  • 96 PLANT SOLUTE TRANSPORT system that is driven through Na+ cotransport. The latter generally has a lower Km 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 physiolog- ical 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 con- trol 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 trans- port. 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 sub- strate 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 modula- tion 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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  • 98 PLANT SOLUTE TRANSPORT 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) NO3- 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 nutri- ents, 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 condi- tion. According to developmental stage, metabolic state, composition of the soil and environmental condition, the plant has to integrate uptake, allocation and redistribu- tion of solutes to maximise their usage for growth and reproduction. Such adaptation requires flexible and robust mechanisms to regulate solute transport. Developmen- tal, 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, mem- brane 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 chap- ter 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 molec- ular 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 Physiological situations requiring the regulation of ion transport 6.2.1 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
  • 100 PLANT SOLUTE TRANSPORT extentsCl− andorganicanions–astheprimaryosmoticallyactivesolutestomaintain 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 (Holdaway- Clarke 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 ef- fects 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 mem- brane. 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 (ICl) and the two dominant K+ channels – identified with the inward-rectifying (IK,in) and outward-rectifying (IK,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 ICl and IK,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
  • REGULATION OF ION TRANSPORTERS 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 ADP- ribose-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 IK,in and ICl 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 IK,in and ICl 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 IK,in and ICl, 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 (pHi) and on extracellular [K+ ] ([K+ ]o). Increasing pHi augments IK,out in a scalar fashion, consistent with a synergistic binding of two H+ (Grabov and Blatt, 1997) and the rise in pHi 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 pHi nor of its site of action, although its kinetics (and that of changes in IK,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 IK,out is unique to plant K+ chan- nels, 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 stom- atal 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).
  • 102 PLANT SOLUTE TRANSPORT 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 phos- phorus 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 ac- quisition 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 trans- port 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
  • REGULATION OF ION TRANSPORTERS 103 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 in- duced 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 theirfunctionashigh-affinitysystemsinnutrientuptake.Fine-tuningofthisresponse 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 re- spect to sulphur sensitivity of their transcriptional response. SULTR1;1 shows sharp induction between 100 and 1500 μM external S, whereas SULTR1;2 transcripts in- crease 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 induc- tion 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 en- vironment 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 glu- tamine (Zhuo et al., 1999; Vidmar et al., 2000). Glutamine seems to be involved in
  • 104 PLANT SOLUTE TRANSPORT SULTRPT NRTAMT HAK S P /+N -K Light Suc, Fru, Glu GSH, Cys OAS Gln, Asn, Arg H2PO4 SO4 2 NH4 + NO3 K+ ROS P1BS SURE PHR Sac3 Ckn /CRE1 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). Trans- porters 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, glu- tamine; 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 ammo- nium transporter AMT1;1 in response to resupply of ammonium nitrate occurs only if assimilation of ammonium into glutamine is inhibited by methionine sulphox- imine (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,
  • REGULATION OF ION TRANSPORTERS 105 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,thisisnotthecaseforlightregulationofnutrienttransporters (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 (Maruyama- Nakashita 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 re- sponds 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
  • 106 PLANT SOLUTE TRANSPORT 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 com- partmentalisation 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+ chan- nels 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 Chap- ter 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 non- selective 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 plasma- membrane-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 (Halfteret al., 2000). Mutations of the SOS2 protein in the N-terminal kinase activation loop and the C- terminal 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 phosphoryla- tion 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 be- tween 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).
  • REGULATION OF ION TRANSPORTERS 107 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). Microarrayexperimentshaverevealedthattranscriptionalregulationofiontrans- porters 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 regula- tion 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 intracel- lular Ca2+ and pH, protein kinases, transcription factors, metabolites and hormones have been confirmed to be essential components of the signalling pathways linking
  • 108 PLANT SOLUTE TRANSPORT 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 remain- der 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 re- gion 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 trans- porters 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 (Ru- bioetal.,2001;Toddetal.,2004).AtPHR1recognisesaGnATATnCmotif,theP1BS 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 (Amt- mann et al., 2006). Very little is known to date about upstream events linking changes in soil nu- trient 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 tran- script levels of several Ca2+ -binding proteins and Ca2+ -dependent kinases (CDPKs) (Armengaudetal.,2004).Transcriptionalregulationofsulphatetransportersappears to involve phosphorylation/dephosphorylation events; in arabidopsis, application of phosphatase blockers (okadaic acid, calyculin A) abolishes green fluorescent pro- tein (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 ex- pression 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 positionofethyleneandjasmonatesignalswithintheK+ starvationresponseremains
  • REGULATION OF ION TRANSPORTERS 109 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 (Maruyama- Nakashita 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 en- vironmental 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 bio- chemical 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 en- dogenous proton pump activity with plant H+ -ATPases was unsuccessful unless the C-terminus was removed (Palmgren and Christensen, 1993; Baunsgaard et al., 1996;
  • 110 PLANT SOLUTE TRANSPORT cyt out AHA/PMA2 Ca2+ cyt out/ER ACA2/8 cyt out KCO1 Ca2+ cyt out CNGC1/2 Ca2+ cyt vac NHX1 AID 14-3-3 BD 14-3-3 FC CaMBD CaM CNBD SelRD P P 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, 14- 3-3 protein-binding domain; FC, fusicoccin; CaM, calmodulin; CaMBD, calmodulin-binding domain; CNBD, cyclic nucleotide binding domain; SelRD, regulatory domain controlling ion selectivity). Indi- vidual 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 max- imal pump activity (Vmax), decreased the apparent Km 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 Km 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
  • REGULATION OF ION TRANSPORTERS 111 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 autoin- hibitory domains are important for autoinhibition of H+ and Ca2+ pumps (Mor- somme 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 autoin- hibition, 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 cytoplas- mic loops (Morsomme et al., 1996, 1998). Partial tryptic digestion showed that these mutations cause a conformational change that makes the C-terminus more accessi- ble 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 tar- get protein containing phosphoserine or phosphothreonine are bound in a conserved amphipathic region present in each monomer of a dimeric 14-3-3 protein. Hence, in- teractions 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
  • 112 PLANT SOLUTE TRANSPORT 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 Vmax, lower Km, shift of pH optimum; see above) suggesting that FC modifies the C-terminus (Johansson et al., 1993; Lan- fermeijer 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 C- terminus 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 (Ca- moni 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 obser- vation 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 sev- eral 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 bar- ley 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 KCO1- mediated 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
  • REGULATION OF ION TRANSPORTERS 113 (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 pro- toplasts derived from the radicle (ABA reduces Kin, FC reduces IK,out and stimulates IK,in). Inclusion of recombinant barley 14-3-3B protein in the patch pipette solution (cytoplasmic side) resulted in a fast decline of Kout currents by 60%, whereas Kin currents were increased. Furthermore, addition of a 14-3-3 binding protein to the pipette solution completely abolished IK,in, showing that residual Kin in the pre- vious 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 pro- tein 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+ an- tiporter NHX1 (Figure 6.2). The interaction between Ca2+ , CaM and Ca2+ -ATPases creates a negative feed- back 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 an- chors 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 bind- ing. 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 modu- lators 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
  • 114 PLANT SOLUTE TRANSPORT 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 pro- tein 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 con- served 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. Pro- gressive 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 iden- tical 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 pro- teinase 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 trans- formed 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 Vmax of Na+ /H+ antiport, whereas the Vmax 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 ev- idence 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
  • REGULATION OF ION TRANSPORTERS 115 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 in- cluding 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 α- and β-subunits (Maathuis, 2004). Nevertheless, heterologous expression has been achieved for AtCNGC1, CNGC2, AtCNGC4 and NtCBP4. Interestingly, these ex- periments 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 bind- ing. Indeed, whole-cell patch-clamp experiments showed that inclusion of AtCaM4 in the (cytoplasmic) pipette solution resulted in a decrease of cAMP-induced cur- rents 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 indi- vidual 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.
  • 116 PLANT SOLUTE TRANSPORT 6.3.2.5 Heteromerisation Membrane transporters are often assembled from several subunits. Hetero- oligomerisation involving several transporter isoforms is likely to be a common phenomenon but has only recently attracted interest from the point of view of trans- porter 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 activ- ity (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 re- vealed 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 het- erotetramers. 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 homote- trameric 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 sys- tems, 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 het- erotetramerisation 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 opportu- nity 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 ob- servations 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 chan- nel’. Nevertheless, disruption of the AtKC gene results in qualitative changes of the inward current affecting its Ca2+ and pH sensitivity (Reintanz et al., 2002) suggest- ing 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
  • REGULATION OF ION TRANSPORTERS 117 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 intercala- tion 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 mem- brane 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 vesi- cle dimensions derived from ultrastructural studies (Picton and Steer, 1983; Phillips et al., 1988) and from imaging studies using fluorescent styryl dyes to label in- ternalised 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 exocy- totic 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 deliveryofspecificiontransportproteinstovarioustargetmembraneswithinthecell. Vesicletraffichasbeenimplicatedinthespatialdistributionoftheauxineffluxcarrier 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 auxin- stimulated 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 demon- strated 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
  • 118 PLANT SOLUTE TRANSPORT V. faba guard cell protoplasts using capacitance recording and styryl dyes. Capaci- tance 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 con- firmed 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 sug- gest 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 move- ments (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 traffick- ing 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 sur- prising 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, es- pecially in guard cells. What is noteworthy, however, is that both SNAREs were initially identified in a screen for ABA receptors using an expression-cloning strat- egy. Furthermore, NtSyp121 was cleaved by the Clostridium botulinum neurotoxin BotN/C and, intriguingly, both neurotoxin treatments and the (so-called Sp2) do- main 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
  • REGULATION OF ION TRANSPORTERS 119 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 dominant- negative Rab1b mutant that blocks export to the Golgi apparatus (Batoko et al., 2000). This shift to endosomal accumulation implies a backlog of synthesised pro- tein 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 mem- brane 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 par- titions 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 obser- vations 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
  • 120 PLANT SOLUTE TRANSPORT 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 im- mediate 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 com- bination of electrophysiological, biochemical, molecular and genetic approaches has created a wealth of information on responses of ion transporters to environ- mental 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, phosphopro- teomics 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. References Adamo, H.P. and Grimaldi, M.E. (1998) Functional consequences of relocating the C-terminal calmodulin-binding autoinhibitory domains of the plasma membrane Ca2+ pump near the N- terminus. Biochemical Journal 331, 763–766. Aitken, A. (1996) 14-3-3 and its possible role in co-ordinating multiple signaling pathways. Trends in Cell Biology 6, 341–347. Alagem, N., Yesylevskyy, S. and Reuveny, E. (2003) The pore helix is involved in stabilizing the open state of inwardly rectifying K+ channels. Biophysical Journal 85, 300–312. Al-ghazi, Y., Muller, B., Pinloche, S., et al. (2003) Temporal responses of Arabidopsis root architec- ture to phosphate starvation: evidence for the involvement of auxin signaling. Plant, Cell and Environment 26, 1053–1066. Allen, G.J., Chu, S.P., Harrington, C.L., et al. (2001) A defined range of guard cell calcium oscillation parameters encodes stomatal movements. Nature 411, 1053–1057. Allen, G.J., Chu, S.P., Schumacher, K., et al. (2000) Alteration of stimulus-specific guard cell calcium oscillations and stomatal closing in Arabidopsis det3 mutant. Science 289, 2338–2342. Amtmann, A., Armengaud, P. and Volkov, V. (2004) Potassium nutrition and salt stress. In: Membrane Transport in Plants (ed. Blatt, M.R.), pp. 316–348. Blackwell, Oxford.
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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 rep- resented 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, in- volving 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, per- oxisomes 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 character- istics 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 pro- teins 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 fol- lowing 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 trans- port 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 conduc- tance, gating, voltage dependence, and selectivity. The biggest task thereafter was to
  • 134 PLANT SOLUTE TRANSPORT 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 knowl- edge 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 flu- orescent 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 tech- niques, plant organellar transporters have often been isolated from quite diverse species – e.g. pea and spinach chloroplasts allow good separation of outer and in- ner 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 en- tire 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 ara- bidopsis 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 com- prehensive 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 character- isation 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 rep- resents 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
  • INTRACELLULAR SOLUTE TRANSPORT 135 Table 7.1 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: Carteretal.(2004);Sazukaetal.(2004);Shimaokaetal.(2004);Szponarskietal.(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. b The April 2006 issue of the Journal of Experimental Botany is dedicated to Plant Proteomics, providing current knowledge about efforts on the proteomic field.
  • 136 PLANT SOLUTE TRANSPORT 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 or- ganelles, 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 biochem- ically, 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, im- provement 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 photo- synthesisandthusbuildupthebasisforalllifedependentonatmosphericoxygenand 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 co- operate 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 repre- sent 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 dark- grown plantlets develop during biogenesis into the mature, autotrophic chloroplast of the green leaf. In contrast, storage plastids are heterotrophic organelles that con- vert 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 assimila- tion, 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 dynam- ically interconvertable; hence, the name plastid originates from the Greek word ‘plastikos’, meaning ‘plastic or shapeable’.
  • INTRACELLULAR SOLUTE TRANSPORT 137 7.2.1 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 O2, thereby providing the basis for most of the life on earth. Via the reductive pentose phosphate cycle (Calvin–Benson cycle) in the stroma, atmospheric CO2 is converted into carbohydrates that are di- rectly used for starch and sucrose biosynthesis inside the chloroplast and cytosol, respectively. Furthermore, glyceraldehyde-3-phosphate can enter the chloroplas- tidic 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 or- ganelles (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 reac- tions 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 biogene- sis 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 cy- tochrome b6-f complex, ferredoxin and photosystem I, ensuring electron flow in the thylakoids. These manifold biosynthetic functions of plastids require the existence of dif- ferent 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 (NO2 − , SO4 2− and PO4 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 diffu- sion process through a general porin. But, as in peroxisomes (Section 7.4.2.1), this
  • 138 PLANT SOLUTE TRANSPORT 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 voltage- dependent 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 enve- lope proteins (see Figure 7.1). OEP16, OEP21, OEP24 and OEP37 – classified ac- cording to their molecular weight – were all isolated as protein bands from fraction- atedouterenvelopemembranesfrompeachloroplasts.Thus,theyrepresentabundant proteins in this membrane system. In addition to the preprotein-conducting chan- nel Toc75 (Hinnah et al., 1997), these OEPs have been functionally characterised invitrobyelectrophysiologicalmeasurementsinartificiallipidbilayers.Theyallrep- resent 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 mem- branes, 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 3- phosphoglycerate (3-PGA) or to sugars (glucose, fructose, sucrose and sorbitol) al- though 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,
  • INTRACELLULAR SOLUTE TRANSPORT 139 Solutes TP, 3-PGA, Pi OEP 21 ATP TP OEP24 OEP37 OEP16 Amino acids, amines Fatty acids, Amino acids? Cold acclimation Seed germination Embryo development Pollen development Export of photoassimilates: TP OE IE Stroma THY TPT Pi TP IMS GPT1 Gluc-6-P Pi Male + female gametophytes: import of carbon skeletons Cl- PPT1 PEP Pi Shikimate pathway: aromatic compounds XPTExchange of intermediates Pentose phosphate pathway Pi Xyl-5-P Pi Xyl-5-P Export of photoassimilates: TP ATP synthesis CLC 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 iden- tified; 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 phos- phate, 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.
  • 140 PLANT SOLUTE TRANSPORT 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 in- tramembrane space. OEP21 has been shown to form a rectifying, anion-selective channel, which is regulated by ATP and triose phosphates (TPs) from the intramem- brane 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 phosphory- lated 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). In- terestingly, 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 chan- nel properties of the slightly cation-selective OEP24 closely resemble those described for general diffusion pores (Nikaido, 2003). The 2.5-nm pore of the high- conductance (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
  • INTRACELLULAR SOLUTE TRANSPORT 141 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 ara- bidopsis, 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, un- published). 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 hourglass- shaped 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 mem- brane 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 accu- mulate 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 func- tion 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 characterisa- tion. 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 com- pounds 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.
  • 142 PLANT SOLUTE TRANSPORT 7.2.2.3 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 mem- brane 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¨olterandSoll,2001).AlthoughGram-negativebacteriaingeneralareconsid- erd 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 enor- mous 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 in- ner 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 as- signed 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 pro- teins 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 Transport across the inner envelope: phosphate translocators, major facilitators and carriers 7.2.3.1 The phosphate translocator family Members of this family are phosphate antiporters that exchange various phosphory- lated 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
  • INTRACELLULAR SOLUTE TRANSPORT 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 ex- pense 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 transis- tory 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, includ- ing starch breakdown and formation and export of free sugars (mostly maltose by MEX1, see below). Simultaneous knockout of both starch synthesis and TPT, how- ever, prevented this compensation and lead to impaired growth and photosynthesis. GPT, the glucose-6-phosphate/phosphate translocator. The glucose-6- phosphate/phosphate antiporter was isolated from heterotrophic pea root plastids (Kammerer et al., 1998). Since plastids of non-green tissue are not photosynthet- ically 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 ara- bidopsis, two GPT proteins are present, which represent functional proteins when reconstituted into liposomes (transport of Pi, glucose-6-phosphate, TPs and phos- phoenolpyruvate) 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 im- ports phosphoenolpyruvate into the plastid rather than export it (Fischer et al., 1997). In these cases, phosphoenolpyruvate is required as a precursor for the plastid- localisedshikimatepathway(Figure7.1).ThearabidopsisPPT1-mutantcue1islack- ing 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
  • 144 PLANT SOLUTE TRANSPORT 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 nitro- gen assimilation. Ammonia, generated by nitrite reduction in the cytosol and/or photorespiration (see Section 7.5), is directly assimilated into glutamate by the ac- tion 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 trans- port is mediated by a malate-coupled two-transporter system (Figure 7.2) in- volving 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 pro- tein (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 ex- pression 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 ox- aloacetate transporter. However, Renne et al. (2003) argue that under in vivo condi- tions 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 CO2 to survive, is defective in chloroplast dicarboxylate transport (Somerville and
  • INTRACELLULAR SOLUTE TRANSPORT 145 IE Carbon + nitrogen assimilation Amino acids, fatty acids, starch BT1 ADP-glucose AMP FOL1 DiT1 2-Oxoglutarate Glutamate Malate Malate DiT2 Photorespiration NTT ATP ADP G lcT Glucose MEX1Starch Maltose Monocots: starch THF Biosynthesis + Breakdown 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
  • 146 PLANT SOLUTE TRANSPORT 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 maltosetransporterthatisunrelatedtoothersugartransportersofthemajorfacilitator 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 andmutantsofthetwoNTTproteins(majorfacilitatorfamily)presentinarabidopsis, 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 undisturbeddevelopmentofyoungtissuesaswellasacontrolledcellularmetabolism 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 depen- dent 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 en- dosperm, however, have to import ADP-glucose, since they cannot convert glucose- 1-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; Shan- non 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 ex- changes ADP-glucose with AMP, is an MCF protein. Interestingly BT1 orthologues
  • INTRACELLULAR SOLUTE TRANSPORT 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 Transport across the inner envelope: ABC transporters and ion transport 7.2.4.1 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, vac- uolar 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 membrane- spanning domains of a classical eukaryotic ABC transporter (for a description of ABCtransporterstructureseeChapter5.3.3).TheyresembleprokaryoticABCtrans- port systems, in which a soluble ATP-binding domain interacts with the transmem- brane 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 protopor- phyrin 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.
  • 148 PLANT SOLUTE TRANSPORT IE Proto IX ? Proto IX ABC1 ABC1 IMS Far-red light Chlorophyll Lipid biosynthesis Lipids OE TGD1 TGD1 ? ? Nitrogen, sulphate reduction ? NO2 - NO2 - SO4 2- SO4 2- PHT 2;1 PO4 3- H+H+ CHX23 Na+/K+ H+ pH control Cl-, Pi, Glutamate ANTR2 POL CAS K+ ? Ca2+ signalling: formation of root nodules or mycorrhizal symbiosis 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 half- molecules 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 bacte- rial ABC transporter complexes (Higgins, 2001). TGD1 was identified in a high- throughputscreenformutantswithalteredlipidmetabolism.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 em- bryo abortion and accumulate triacylglycerols, oligogalactolipids and phosphati- date, 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 149 7.2.4.2 Ion transport Sulphate, nitrite, phosphate. Sulphate and nitrite reduction occur in plastids; there- fore, the SO4 2− and NO2 − 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, PO4 3− is transported in an alternative way. Douce and Joyard (1990) propose a carrier system, based on exchange of SO4 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, homo- logues 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 (ni- trate assimilation related) family in Chlamydomonas reinhardtii, containing trans- membrane 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 trans- port 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 chloro- plast 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, light- driven 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, concen- trations 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 pho- tosynthetic 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 com- plex (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 wild- type 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 simi- larity 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 CO2 fixation, while in the thylakoid lumen Ca2+ is required for the function of PSII.
  • INTRACELLULAR SOLUTE TRANSPORT 151 IEIMSOE MRS2-11 Ca2+ ? Mg2+ Metal Fe2+, Zn2+, Cu2+, Mn2+ H+ PPF1 PAA1 ATP ADP HMA1 ATP ADP Cytoplasmic Ca2+ Ca2+ signalling Light-dependent Mg2+ uptake PAA2 Photosynthetic electron transport Cu2+ SOD: detoxification Cu2+ Chlorophyll 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+ andcorrespondingsignallingevents.LocalisationtotheinnerenvelopeandCa2+ transportcapacity of PPF1, however, are not clarified yet. MRS2–11 (two to three α-helical domains) can mediate light- dependent 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 trans- genic 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, repre- sents 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 ara- bidopsis is localised to thylakoid membranes and required for the membrane inser- tion 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.
  • 152 PLANT SOLUTE TRANSPORT Magnesium. In chloroplasts, stromal magnesium, which increases upon illumi- nation, is involved in the regulation of key enzymes of carbon fixation and photo- synthesis and represents the central cation of the chlorophyll molecule (Berkowitz and Wu, 1993; Shaul, 2002; Ishijima et al., 2003). AtMRS2–11, a protein that be- longs 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 iron- enriched systems (PSII, PSI, cytochrome b6-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, zinc- finger 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 clus- ters 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 Lo- breaux, 2005; see also Section 12.2). Thus, plastids represent one of the most iron- enriched systems in the plant cell. However, the iron transporter in the plastid en- velope 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 en- velope, 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 con- tent and Cu/Zn superoxide dismutase activity in chloroplasts; thus, both ATPases represent the Cu uptake system across the inner envelope.
  • INTRACELLULAR SOLUTE TRANSPORT 153 7.3 Mitochondria Mitochondria are highly dynamic and complex semi-autonomous organelles, com- posed of a smooth outer membrane surrounding an extensively folded inner mem- brane 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 con- tact 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 dehy- drogenase complex, the tricarboxylic acid cycle, and for the oxidative decarboxyla- tion 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 mem- brane. In contrast to animals, the bulk of fatty acid oxidation in plants is confined to peroxisomes (cf. Douce and Neuburger, 1989). However, mitochondria are in- volved 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 reg- ulated 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 cofac- tors 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, thisβ-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 β- sheets, 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 pro- teins than humans (3) or yeast (2), giving the opportunity to diversify physiological functions. However, all expressed VDAC isoforms (one VDAC is a possible pseudo- gene) 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
  • INTRACELLULAR SOLUTE TRANSPORT 155 VDAC ≤5 kDa IM OM IMS matrix Solute fluxes across the OM PiC H+ Pi AAC ATP ADP ATP synthesis + OH-/ Pi UCP H+ Heat Cold acclimation ATP export DTC Di-, tricarboxylates TCA cycle Ammonia assimilation Arg BAC1 Orn Mobilisation seed storage proteins Arg breakdown Lipid mobilisation SFC Suc Fum 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.
  • 156 PLANT SOLUTE TRANSPORT (Wandrey et al., 2004). In summary, by improvement and usage of miscellaneous assays for subcellular localisation of membrane proteins, evidence is accumulat- ing 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 electro- chemical 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 carri- ers have been reported in arabidopsis (Millar and Heazlewood, 2003; Picault et al., 2004). However, in databases and publications, several MCF members have anno- tated 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 iden- tifying 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 regula- tion 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 (H2PO4 − )/proton symport or phosphate/hydroxyl ion antiport as well as the ex- change 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 mitochon- drial 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 be- low), 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). Pharma- cological studies have demonstrated that ATP/ADP transport in plant mitochondria involves an AAC similar to that of mammals (see Laloi, 1999, for details). Subse- quently,AACwaspurifiedfrommaizemitochondriaandshowntocatalyseATP/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 phospho- rylation 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 trans- port, 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 con- firmed by the identification of plastidic AACs from arabidopsis that belong to the major facilitator family and so are structurally different from the mitochondrial car- riers (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 activ- ity of mitochondrial carriers either for the import of respiratory substrates such as pyruvate, malate and oxaloacetate, or for the constant export of intermediates. Previ- ous 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 re- centlyidentifieddicarboxylate–tricarboxylatecarrier(DTC;seebelow).Ontheother hand, the MCF sequences in arabidopsis include three putative dicarboxylate carrier proteins (Picault et al., 2004), awaiting functional characterisation. Further, physio- logical 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 dicar- boxylates 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 elon- gation), 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 di- carboxylates (such as malate, oxaloacetate, oxoglutarate and maleate) and tricar- boxylates (such as citrate, isocitrate, cis-aconitate and trans-aconitate) by a counter- exchangemechanism(Figure7.5;Picaultetal.,2002).Furthermore,nitratesupplyto
  • INTRACELLULAR SOLUTE TRANSPORT 159 nitrogen-starved tobacco plants leads to an increase in DTC mRNA in roots and leaves.DTC,whichisubiquitouslyexpressedandwidelyfoundintheplantkingdom, 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 simi- larity 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 tricar- boxylate 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 in- volved 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 mi- tochondria, 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 photores- piratory cycle (see Section 7.5; Figure 7.8) requires an amino acid transporter. Al- though 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 phos- pholipid 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
  • 160 PLANT SOLUTE TRANSPORT 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 dicar- boxylate 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). Acetyl- CoA 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 suc- cinate 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 expres- sion in developing and germinating pollen suggests a role in ethanolic fermentation. 7.3.4 Transport across the inner membrane: ABC transporters and ion channels 7.3.4.1 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 arabidop- sis (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.
  • INTRACELLULAR SOLUTE TRANSPORT 161 Fe–S cluster K+ IM Matrix ATM3 ATM3 Iron homeostasis Fe–S cluster biogenesis Kin Sur ?K+ H+ Energy dissipation Cold acclimation, volume regulation, ... Ca2+? Ca2+ signalling Cl- ClC Nt1 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(Kin),coupledtoasulphonylureareceptor(Sur).TogetherwithapotentialK+ /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 potas- sium (energy dissipation), calcium (cell signalling) and anions/chloride. However, the molecular identities of the corresponding transport proteins still remain un- known. 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 impli- cations to the physiology of this organelle, because the existence of a K+ import
  • 162 PLANT SOLUTE TRANSPORT 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 soy- bean suspension cultures indicate the involvement of a K+ -ATP channel during the manifestation of PCD induced by H2O2 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 mi- tochondrial targeted aequorin that internal Ca2+ is modulated by physiological and environmental stimuli. It is speculated that as in animals, the main targets of mi- tochondrial 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 mem- brane. Initially, the term microbody was introduced by mammalian microscopists to describe a membrane-surrounded particle of unknown function. Since then, sub- classifications, such as peroxisomes and glyoxysomes, have been used for micro- bodies with different metabolic pathways and tissue distribution (Tolbert, 1971). The synonymously used term peroxisome refers to the primary function in compart- mentalisation 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 163 7.4.1 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 peroxisomesandunspecialisedperoxisomes(Beevers,1979).Theygeneratereactive oxygen species and contain defence mechanisms in the form of catalase (detoxifi- cation of H2O2), superoxide dismutase and the membrane-bound ascorbate perox- idase. 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 re- side in cells of storage tissue, such as endosperm, and in cotyledons during germina- tion of oilseed plants. They contain enzymes for fatty acid oxidation and for the gly- oxylate 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 inter- convert during cellular processes (Hayashi et al., 2000). In germinating seedlings, for example, glyoxysomes in cotyledons are functionally transformed into leaf per- oxisomes upon illumination. The reverse process has been observed when leaves undergo senescence. Apart from the primary functions in oxidative metabolism, β- oxidation of fatty acids, photorespiration (leaf peroxisomes) and lipid mobilisation (glyoxysomes), plant peroxisomes also play a significant role in nitrogen assimi- lation (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. How- ever, 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
  • 164 PLANT SOLUTE TRANSPORT 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 bac- teria. It was proposed that metabolism of peroxisomes is in general not compart- mentalised by the boundary membrane, but by the strictly organised arrangement of matrix enzymes in multienzyme complexes (Heupel and Heldt, 1994). In conse- quence, 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? CTS FA PMP38 ADP FA FA-CoA ß-oxidation CoASH AMP + PPi ATP CoASH ? (FA-CoA?), PMP22 ? ? Lipid mobilisation in glyoxysomes Hormone precursors Phytohormone biosynthesis 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.
  • INTRACELLULAR SOLUTE TRANSPORT 165 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 (per- oxisomes) 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 me- diated 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 mem- brane 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 fam- ily MCF. The peroxisomal localisation of AtPMP38 was verified by immunoblot analysis on fractionated organellar membranes and immunogold labelling. Since ex- pression 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
  • 166 PLANT SOLUTE TRANSPORT (‘COMATOSE’, for review see Theodoulou et al., 2006). Peroxisomal ABC trans- porters 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 or- ganellar 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 medi- ates 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 trans- port 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 mem- branes (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 CO2 and consume O2, 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). Pho- torespiration starts in the chloroplast stroma with the oxygenation of ribulose-1,5- bisphosphate 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 dephospho- rylated. 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
  • INTRACELLULAR SOLUTE TRANSPORT 167 Glt Glt2-Pglt 3-PGA Ru 1,5-BP Calvin cycle ? ? ? ? (i) Gox Gly (ii) Gly ? ?Ser (iii) Ser ? ?Glc (iv) Glc DiT2 ? ? GluGlu DiT1 Mal Mal 2-OG2-OG Gly NH4 + CO2 (v) ? (vi) Orn or Glu ? Cit or Gln ? O2 GOGAT GS Gln Glu Plastid Peroxisome Mitochondrion Rubisco NH3 DTC ? Figure 7.8 Solute transport during photorespiration. Photorespiration involves solute transport be- tween 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 mitochon- dria, 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 num- bers (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 2- oxoglutarate [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 CO2 in mitochondria is assumed to diffuse back to chloroplasts (dotted arrow), while for NH4 + 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 assim- ilated into glutamate by the action of glutamine synthetase/glutamate synthase (GS–GOGAT) cycle. Please note that the cytotoxic NH4 + /NH3 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 ox- idised 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 medi- ated by a porin (Reumann, 2000). Thus, transporters for glycine in the peroxisome and inner mitochondrial membrane have not yet been identified. In the mitochon- dria, 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.
  • 168 PLANT SOLUTE TRANSPORT 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. How- ever, 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 CO2 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 pho- torespiratory 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 com- partment 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 compartmen- talisation 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 cy- toplasm 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 biosyn- thetic 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, mat- uration 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 trans- port mechanisms requiring high-energy metabolites for their operation: the vacuo- lar 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 eukary- otic 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 cellu- lar pH homeostasis. The V-ATPase is a multisubunit enzyme complex, structurally related to F0F1-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 (Schu- macher 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).
  • 170 PLANT SOLUTE TRANSPORT V-PPase PPi Pi H+V-PPase Salt tolerance Cell growth pH gradient Storage pH homeostasis Cell turgor (growth) tDT Mal SUT4 Suc Photosynthesis TIP TIP H2O Osmoregulation Cell volume Urea Glycerol MRP Detoxification GS (Glutathione) -X X-Glucose X-Glucuronic acid Storage of xenobiotics, secondary metabolites Phytosiderophores? IDI7 IDI7 Iron acquisition? H+ ATP ADP V-ATPase 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 (F1F0-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 Transport of malate and sucrose across the tonoplast 7.6.2.1 Malate In plant cells, excess malate is stored within the large, central vacuole. Malate serves as a storage form of fixed CO2, as charge balance and as an osmolyte for maintenance of cell turgor. The transport of malate into the vacuole is crucial for
  • INTRACELLULAR SOLUTE TRANSPORT 171 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 pH- dependent 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 trans- ported 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 dif- fusion 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. Further- more, this proteomic analysis provides additional protein sequences, e.g. a potential vacuolar hexose transporter (Endler et al., 2006). 7.6.3 Aquaporins and ABC transporter in the tonoplast 7.6.3.1 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 oocytesofX.laevis,manyoftheTIPsfunctionaswaterchannels(aquaporins;Bethke
  • 172 PLANT SOLUTE TRANSPORT and Jones, 2000; Maeshima, 2001, and references therein). Aquaporins facilitate wa- ter transport across biomembranes in an osmotic pressure-dependent manner. TIPs as well as PIPs (plasma membrane intrinsic proteins) belong to the ubiquitous fam- ily 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 aquapor- ins 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 pro- tecting 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 tono- plast to other endomembrane fractions after hyperosmotic treatment with mannitol (Vera-Estrella et al., 2004). This redistribution provides insight into a novel mech- anism 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 Sec- tion 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 extru- sion protein (MATE) (Yazaki, 2005, and references therein). The MRP subfamily in arabidopsis contains 15 members and represents the best characterised plant ABC
  • INTRACELLULAR SOLUTE TRANSPORT 173 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 func- tions in detoxification of xenobiotics, which are conjugated to glutathione. How- ever, 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 flavoneglucuronidesintotonoplastvesicles.Thistransportisdependentonthemem- brane potential, pointing to a possible regulation by vacuolar proton pumps (Klein et al., 2000). Transport mechanisms for the major barley flavonoid saponarin, how- ever, 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 Ion transport 7.6.4.1 Ion channels The first ion channels described in the tonoplast of plant cells were the slow- activating (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 electro- physiologically 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).
  • 174 PLANT SOLUTE TRANSPORT 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 for- mation of the SV conductance (Sch¨onknecht et al., 2002). However, recent studies of TPK1 conductance in yeast vacuoles (Bihler et al., 2005) and the characterisa- tion 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). K+ TPK1 TPK1 Ca2+ BCA1 ATPADP Ca2+ CAX Ca2+ H+ NHX H+Na+ MHX H+ Mg2+, Zn2+ NRAMP3+4 Fe2+, (Zn2+, Mn2+) Fe mobilisation under iron starvation Zn2+ MTP 1+3K+ Ca2+ Volume regulation of vacuole + cell TPC1 TPC1 Ca2+ signalling Storage Ca2+ signalling Salt tolerance Storage Zn, metal tolerance homeostasis pH 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.
  • INTRACELLULAR SOLUTE TRANSPORT 175 Nevertheless, the K+ channel TPK1 might be involved in regulation of the SV- channel conductance in vacuoles. TPC1representstheslowvacuolarchannel.Recently,theCa2+ -dependentCa2+ - 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 demonstratethefunctionofvacuolarsolutetransportduringcellularCa2+ 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 estab- lished 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 vac- uoles 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 Ca2+ accumulation in the vacuole. Again, Ca2+ /H+ exchange across the vacuolar membranehasbeenmeasuredindiverseplantspeciesandthecorrespondingproteins 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
  • 176 PLANT SOLUTE TRANSPORT 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 at- tention 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 Sec- tion 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 overexpress- ing 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
  • INTRACELLULAR SOLUTE TRANSPORT 177 expression (zinc-finger proteins). AtMHX is an arabidopsis tonoplast transporter that can exchange protons with Mg2+ and Zn2+ ions (Shaul et al., 1999). Simi- lar 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 dismu- tases and catalysing electron transfer in mitochondria and chloroplasts. Zinc (Zn) is an essential plant micronutrient and functions as cofactor for many enzymes. How- ever, 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 iso- forms 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 α- helical 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 accumu- lation in roots (Thomine et al., 2003). It was proposed that NRAMP3 influences metal accumulation by mobilising vacuolar metal pools to the cytosol, in partic- ular 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 trans- genic 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
  • 178 PLANT SOLUTE TRANSPORT AtMTP1 have been implicated in the metal tolerance of hyperaccumulator plants (Desbrosses-Fonrouge et al., 2005, and references therein). All plant MTP1-like pro- teins complement zinc hypersensitivity of a yeast mutant, and AtMTP1 transports Zn2+ when heterologously expressed in yeast and X. laevis oocytes (Desbrosses- Fonrouge et al., 2005). Thus, MTP1 sequesters excess Zn into vacuoles. Onlyveryrecently,theMTP3proteininarabidopsiswaslocalisedtothetonoplast and shown to also exhibit Zn transport capacity (Arrivault et al., 2006). Overexpres- sion 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. 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. The Plant Cell 17, 1233–1251. Abrecht, H., Goormaghtigh, E., Ruysschaert, J.M. and Homble, F. (2000a) Structure and orientation of two voltage-dependent anion-selective channel isoforms. An attenuated total reflection fourier- transform infrared spectroscopy study. The Journal of Biological Chemistry 275, 40992–40999. Abrecht, H., Wattiez, R., Ruysschaert, J.M. and Homble, F. (2000b) Purification and characterization of two voltage-dependent anion channel isoforms from plant seeds. Plant Physiology 124, 1181– 1190. Aljamal, J.A., Genchi, G., De Pinto, V., et al. (1993) Purification and characterization of porin from corn (Zea mays L.) mitochondria. Plant Physiology 102, 615–621. Apse, M.P., Aharon, G.S., Snedden, W.A. and Blumwald, E. (1999) Salt tolerance conferred by overexpression of a vacuolar Na+ /H+ antiport in Arabidopsis. Science 285, 1256–1258. 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. Arrivault, S., Senger, T. and Kr¨amer, U. (2006) The Arabidopsis metal tolerance protein AtMTP3 maintains metal homeostasis by mediating Zn exclusion from the shoot under Fe deficiency and Zn oversupply. The Plant Journal 46, 861–879. Baldi, P., Grossi, M., Pecchioni, N., Vale, G. and Cattivelli, L. (1999) High expression level of a gene coding for a chloroplastic amino acid selective channel protein is correlated to cold acclimation in cereals. Plant Molecular Biology 41, 233–243. Balk, J. and Lobreaux, S. (2005) Biogenesis of iron–sulfur proteins in plants. Trends in Plant Science 10, 324–331. Barkla, B.J. and Pantoja, O. (1996) Physiology of ion transport across the tonoplast of higher plants. Annual Review of Plant Physiology and Plant Molecular Biology 47, 159–184. Barrieu, F., Marty-Mazars, D., Thomas, D., Chaumont, F., Charbonnier, M. and Marty, F. (1999) Desiccation and osmotic stress increase the abundance of mRNA of the tonoplast aquaporin BobTIP26-1 in cauliflower cells. Planta 209, 77–86. Beavis, A.D. and Vercesi, A.E. (1992) Anion uniport in plant mitochondria is mediated by a Mg(2+ )- insensitive inner membrane anion channel. The Journal of Biological Chemistry 267, 3079–3087. Bedhomme, M., Hoffmann, M., McCarthy, E.A., et al. (2005) Folate metabolism in plants: an Ara- bidopsis homolog of the mammalian mitochondrial folate transporter mediates folate import into chloroplasts. The Journal of Biological Chemistry 280, 34823–34831.
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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 de- scribes 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 consid- eration 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 solubil- ity 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
  • 194 PLANT SOLUTE TRANSPORT mobility of even very soluble cations such as K+ . By contrast, NO3 − 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 trans- port 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 strate- gies 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). Ram- ification 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 NO3 − , 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 major- ity 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 en- zymes and chelating and solubilising agents; Landeweert et al., 2001), and many decompose organic matter directly. They may also compete effectively with other
  • ION UPTAKE BY PLANT ROOTS 195 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 photo- synthates. 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 micro- habitats 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
  • 196 PLANT SOLUTE TRANSPORT 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 develop- ment 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 dif- fuse 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 Na2SO4 uptake rates are low compared to the concentration of salt in the soil solution (Tinker and Nye, 2000). Preferentialuptakeofparticularnutrientsandthedevelopmentofdepletionzones 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. Cell- wall components contain a high density of fixed, mainly anionic charges. Cell-wall residues preferentially bind particular ions (especially H+ and Ca2+ ). In addition,
  • ION UPTAKE BY PLANT ROOTS 197 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 Radial transport of solutes across the outer part of the root 8.5.1 The role of apoplastic barriers The Casparian strip of the endodermis forms a suberised barrier to apoplastic move- ment 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 mem- branes 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, usu- ally 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 con- nected by plasmodesmata, the symplast forms a continuum linking the cortical and cortex endodermisepidermis stele xylem Casparian strip apoplastic pathway transcellular pathway symplastic pathway 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).
  • 198 PLANT SOLUTE TRANSPORT 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 particularlyimportantinaccessingimmobilenutrientsandshowespeciallyextensive 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
  • ION UPTAKE BY PLANT ROOTS 199 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 etal.,2003;Shinetal.,2005).Othertypesofevidencefortheimportanceofroothairs 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 ex- tent 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 con- tribute 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 NH4 + and NO3 − (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 con- ditions 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
  • 200 PLANT SOLUTE TRANSPORT 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 (Clark- son, 1993). However, it is nevertheless possible that many solutes reach the stele via some combination of the symplastic pathway through plasmodesmata and symplast- apoplast-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. Non- selective 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 symplasti- cally 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 there- fore report a narrowing rather than complete closure of plasmodesmata (Duckett et al., 1994). However, electrophysiological studies of the electrical continuity be- tween 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 par- ticularly 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 exo- and endodermal barriers (especially for uptake of Ca, or solute absorption across
  • ION UPTAKE BY PLANT ROOTS 201 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 NO3 − 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 tox- icity 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 spe- cialised for Ca2+ influx, and the inner face for Ca2+ efflux. However in many species,
  • 202 PLANT SOLUTE TRANSPORT 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 com- pletely, 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 un- der field conditions, the late metaxylem vessels, which constitute the major high- conductance 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 trans- fer 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 hy- droponically, 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 de- scriptions 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.
  • ION UPTAKE BY PLANT ROOTS 203 8.7 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 im- mediately 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, suggest- ing that there may be substantial unloading from the symplast before the xylem parenchyma (Ma and Peterson, 2001). In this case, unloading may occur from peri- cycle 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. More- over, 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 pro- vide 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 sen- sitive 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
  • 204 PLANT SOLUTE TRANSPORT the shoot (Smith, 1991). Split-root experiments, where the shoot solute content is manipulated independently of root content, indicate that shoot nutritional require- ments 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+ concen- tration 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 de- cline 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 thebidirectionalexchangeofsolutesbetweenthexylemfluidandxylemparenchyma cells, caution must be exercised in determining whether transporter expression or transport activity, measured in xylem parenchyma cells, represent xylem loading mechanisms. 8.8 The kinetics of solute uptake into roots 8.8.1 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 NH4 + (Britto et al., 2001) and 20–50% of NO3 − 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
  • ION UPTAKE BY PLANT ROOTS 205 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 0 20 40 60 80 100 0 2 4 6 8 10 12 root total root vacuole shoot root cytosol time (min) tissue 22 Nacontent + 0 200 400 600 800 1000 0 20 40 60 80 shoot root total root vacuole root cytosol time (min) tissue 22 Na + content B A 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.
  • 206 PLANT SOLUTE TRANSPORT 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 tis- sues, 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 re- uptake 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
  • ION UPTAKE BY PLANT ROOTS 207 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 par- ticularly 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 neces- sarily 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 multi- valent 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 mea- sure 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 ion- selective 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 xylem- fluid composition measured using X-ray microanalysis, ion-sensitive microelec- trodes, 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 elec- trophysiological studies of isolated protoplasts, although these must then be contex- tualised with measures of membrane potentials in situ and estimates of the solute concentrations to which the cells are normally exposed. The roles of particular trans- port 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 ac- tivity 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.
  • 208 PLANT SOLUTE TRANSPORT 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 andchangesindiffusionratesandmembranepotentialatdifferentK+ 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), NH4 + in rice (Wang et al., 1993a,b; 1994) and NO3 − 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 PO4 − 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 solutetransportersandmutantslackingexpressionofthesetransportershasindicated that uptake of K+ , NH4 + and NO3 − 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 high- affinity 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, NH4 + 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 NH4 + 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 NO3 − appears to be active and H+ -coupled even at 20 mM external NO3 − (Glass et al., 1992). Nevertheless both high- and low-affinity NO3 − transporters have been identified,
  • ION UPTAKE BY PLANT ROOTS 209 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 ‘in- stantaneous’ 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 NO3 − transport is induced by detection of extra- cellular NO3 − . High-affinity uptake is reduced in K+ and NO3 − -replete roots, in the case of NO3 − by accumulation of products of NO3 − assimilation rather than by NO3 − 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 manipu- lated independently of the roots (see Section 8.7). 8.9 Conclusion Faced with patchy and fluctuating solute availability, plants utilise a variety of mech- anisms 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 ef- fects 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). References Ashley, M.K., Grant, M. and Grabov, A. (2006) Plant responses to potassium deficiencies: a role for potassium transport proteins. Journal of Experimental Botany 57, 426–436. Aslam, M., Travis, R.L. and Rains, D.W. (1996) Evidence for substrate induction of a nitrate efflux system in barley roots. Plant Phyisology 112, 1167–1175. Baldani, J.I., Caruso, L., Baldani, V.L.D., Goi, S.R. and D¨obereiner, J. (1997) Recent advances in BNF with non-legume plants. Soil Biology and Biochemistry 29, 911–922.
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