Biochemistry and Molecular Biology of Plants [Second edition] 9780470714218, 9780470714225, 0470714212, 9781118502211, 1118502213

Table of contents :
Title Page......Page 5
Copyright Page......Page 6
Brief Contents......Page 7
Contents......Page 8
The Editors......Page 13
List of Contributors......Page 14
Preface......Page 17
About the Companion Website......Page 18
PART I COMPARTMENTS......Page 19
1.1 Common properties and inheritance of cell membranes......Page 20
1.2 The fluid-mosaic membrane model......Page 22
1.3 Plasma membrane......Page 28
1.4 Endoplasmic reticulum......Page 31
1.5 Golgi apparatus......Page 36
1.6 Exocytosis and endocytosis......Page 41
1.7 Vacuoles......Page 45
1.8 The nucleus......Page 46
1.9 Peroxisomes......Page 49
1.10 Plastids......Page 50
1.11 Mitochondria......Page 57
Summary......Page 62
2.1 Sugars are building blocks of the cell wall......Page 63
2.2 Macromolecules of the cell wall......Page 69
2.3 Cell wall architecture......Page 91
2.4 Cell wall biosynthesis and assembly......Page 98
2.5 Growth and cell walls......Page 108
2.6 Cell differentiation......Page 117
2.7 Cell walls as sources of food, feed, fiber, and fuel, and their genetic improvement......Page 126
Summary......Page 128
3.1 Overview of plant membrane transport systems......Page 129
3.2 Pumps......Page 138
3.3 Ion channels......Page 146
3.4 Cotransporters......Page 160
3.5 Water transport through aquaporins......Page 164
Summary......Page 166
4.1 The cellular machinery of protein sorting......Page 169
4.2 Targeting proteins to the plastids......Page 171
4.3 Targeting proteins to mitochondria......Page 175
4.4 Targeting proteins to peroxisomes......Page 177
4.5 Transport in and out of the nucleus......Page 178
4.6 ER is the secretory pathway port of entry and a protein nursery......Page 179
4.7 Protein traffic and sorting in the secretory pathway: the ER......Page 193
4.8 Protein traffic and sorting in the secretory pathway: the Golgi apparatus and beyond......Page 200
4.9 Endocytosis and endosomal compartments......Page 206
Summary......Page 207
5.1 Introduction to the cytoskeleton......Page 209
5.2 Actin and tubulin gene families......Page 212
5.3 Characteristics of actin filaments and microtubules......Page 214
5.4 Cytoskeletal accessory proteins......Page 220
5.5 Observing the cytoskeleton: Statics and dynamics......Page 225
5.6 Role of actin filaments in directed intracellular movement......Page 228
5.7 Cortical microtubules and expansion......Page 234
5.8 The cytoskeleton and signal transduction......Page 237
5.9 Mitosis and cytokinesis......Page 240
Summary......Page 256
PART II CELL REPRODUCTION......Page 257
6.1 Composition of nucleic acids and synthesis of nucleotides......Page 258
6.2 Replication of nuclear DNA......Page 263
6.3 DNA repair......Page 268
6.4 DNA recombination......Page 273
6.5 Organellar DNA......Page 278
6.6 DNA transcription......Page 286
6.7 Characteristics and functions of RNA......Page 288
6.8 RNA processing......Page 296
Summary......Page 306
7.1 Amino acid biosynthesis in plants: research and prospects......Page 307
7.2 Assimilation of inorganic nitrogen into N-transport amino acids......Page 310
7.3 Aromatic amino acids......Page 320
7.4 Aspartate-derived amino acids......Page 336
7.5 Branched-chain amino acids......Page 344
7.6 Glutamate-derived amino acids......Page 348
7.7 Histidine......Page 351
Summary......Page 354
8.1 Structure and function of lipids......Page 355
8.2 Fatty acid biosynthesis......Page 362
8.3 Acetyl-CoA carboxylase......Page 366
8.4 Fatty acid synthase......Page 368
8.5 Desaturation and elongation of C16 and C18 fatty acids......Page 370
8.6 Synthesis of unusual fatty acids......Page 378
8.7 Synthesis of membrane lipids......Page 383
8.8 Function of membrane lipids......Page 391
8.9 Synthesis and function of extracellular lipids......Page 400
8.10 Synthesis and catabolism of storage lipids......Page 407
8.11 Genetic engineering of lipids......Page 413
Summary......Page 418
9.1 Genome structure: a 21st-century perspective......Page 419
9.2 Genome organization......Page 422
9.3 Transposable elements......Page 434
9.4 Gene expression......Page 440
9.5 Chromatin and the epigenetic regulation of gene expression......Page 448
Summary......Page 454
10.1 Organellar compartmentalization of protein synthesis......Page 456
10.2 From RNA to protein......Page 457
10.3 Mechanisms of plant viral translation......Page 465
10.4 Protein synthesis in plastids......Page 468
10.5 Post-translational modification of proteins......Page 475
10.6 Protein degradation......Page 481
Summary......Page 493
11.1 Animal and plant cell cycles......Page 494
11.2 Historical perspective on cell cycle research......Page 495
11.3 Mechanisms of cell cycle control......Page 500
11.4 The cell cycle in action......Page 506
11.5 Cell cycle control during development......Page 515
Summary......Page 524
PART III ENERGY FLOW......Page 525
12.1 Overview of photosynthesis......Page 526
12.2 Light absorption and energy conversion......Page 529
12.3 Photosystem structure and function......Page 537
12.4 Electron transport pathways in chloroplast membranes......Page 547
12.5 ATP synthesis in chloroplasts......Page 555
12.6 Organization and regulation of photosynthetic complexes......Page 558
12.7 Carbon reactions: the Calvin–Benson cycle......Page 560
12.8 Rubisco......Page 566
12.9 Regulation of the Calvin–Benson cycle by light......Page 569
12.10 Variations in mechanisms of CO2 fixation......Page 575
Summary......Page 583
Introduction......Page 585
13.1 The concept of metabolite pools......Page 588
13.2 The hexose phosphate pool: a major crossroads in plant metabolism......Page 589
13.3 Sucrose biosynthesis......Page 591
13.4 Sucrose metabolism......Page 595
13.5 Starch biosynthesis......Page 598
13.6 Partitioning of photoassimilates between sucrose and starch......Page 605
13.7 Starch degradation......Page 611
13.8 The pentose phosphate/triose phosphate pool......Page 615
13.9 Energy and reducing power for biosynthesis......Page 619
13.10 Sugar-regulated gene expression......Page 624
Summary......Page 626
14.1 Overview of respiration......Page 628
14.2 Citric acid cycle......Page 631
14.3 Plant mitochondrial electron transport......Page 638
14.4 Plant mitochondrial ATP synthesis......Page 650
14.5 Regulation of the citric acid cycle and the cytochrome pathway......Page 652
14.6 Integration of the cytochrome pathway and nonphosphorylating pathways......Page 653
14.7 Interactions between mitochondria and other cellular compartments......Page 657
14.8 Biochemical basis of photorespiration......Page 664
14.9 The photorespiratory pathway......Page 666
14.10 Role of photorespiration in plants......Page 670
Summary......Page 673
PART IV METABOLIC AND DEVELOPMENTAL INTEGRATION......Page 675
15.1 Selection pressures and long‐distance transport systems......Page 676
15.2 Cell biology of transport modules......Page 682
15.3 Short-distance transport events between xylem and nonvascular cells......Page 686
15.4 Short-distance transport events between phloem and nonvascular cells......Page 691
15.5 Whole-plant organization of xylem transport......Page 709
15.6 Whole-plant organization of phloem transport......Page 714
15.7 Communication and regulation controlling phloem transport events......Page 723
Summary......Page 728
16.1 Overview of nitrogen in the biosphere and in plants......Page 729
16.3 Enzymology of nitrogen fixation......Page 733
16.4 Symbiotic nitrogen fixation......Page 736
16.6 Nitrate uptake and transport......Page 753
16.7 Nitrate reduction......Page 757
16.8 Nitrite reduction......Page 762
16.10 Interaction between nitrate assimilation and carbon metabolism......Page 763
16.11 Overview of sulfur in the biosphere and plants......Page 764
16.12 Sulfur chemistry and function......Page 765
16.13 Sulfate uptake and transport......Page 768
16.14 The reductive sulfate assimilation pathway......Page 770
16.15 Cysteine synthesis......Page 773
16.16 Synthesis and function of glutathione and its derivatives......Page 776
16.17 Sulfated compounds......Page 781
16.18 Regulation of sulfate assimilation and interaction with nitrogen and carbon metabolism......Page 782
Summary......Page 785
17.1 Gibberellins......Page 787
17.2 Abscisic acid......Page 795
17.3 Cytokinins......Page 803
17.4 Auxins......Page 813
17.5 Ethylene......Page 824
17.6 Brassinosteroids......Page 828
17.7 Polyamines......Page 836
17.8 Jasmonic acid......Page 839
17.9 Salicylic acid......Page 844
17.10 Strigolactones......Page 848
Summary......Page 851
18.1 Characteristics of signal perception, transduction, and integration in plants......Page 852
18.2 Overview of signal perception at the plasma membrane......Page 856
18.3 Intracellular signal transduction, amplification, and integration via second messengers and MAPK cascades......Page 861
18.4 Ethylene signal transduction......Page 865
18.5 Cytokinin signal transduction......Page 868
18.6 Integration of auxin signaling and transport......Page 870
18.7 Signal transduction from phytochromes......Page 875
18.8 Gibberellin signal transduction and its integration with phytochrome signaling during seedling development......Page 879
18.9 Integration of light, ABA, and CO2 signals in the regulation of stomatal aperture......Page 884
Summary......Page 888
19.1 The transition from vegetative to reproductive development......Page 890
19.2 The molecular basis of flower development......Page 899
19.3 The formation of male gametes......Page 907
19.4 The formation of female gametes......Page 915
19.5 Pollination and fertilization......Page 920
19.6 The molecular basis of self-incompatibility......Page 926
19.7 Seed development......Page 931
Summary......Page 941
20.1 Types of cell death......Page 943
20.2 PCD during seed development and germination......Page 948
20.3 Cell death during the development of secretory bodies, defensive structures and organ shapes......Page 950
20.4 PCD during reproductive development......Page 955
20.5 Senescence and PCD in the terminal development of leaves and other lateral organs......Page 958
20.6 Pigment metabolism in senescence......Page 966
20.7 Macromolecule breakdown and salvage of nutrients in senescence......Page 969
20.8 Energy and oxidative metabolism during senescence......Page 975
20.9 Environmental influences on senescence and cell death I: Abiotic interactions......Page 979
20.10 Environmental influences on senescence and cell death II: PCD responses to pathogen attack......Page 982
20.11 Plant hormones in senescence and defense-related PCD......Page 992
Summary......Page 1000
PART V PLANT ENVIRONMENT AND AGRICULTURE......Page 1001
21.1 Pathogens, pests, and disease......Page 1002
21.2 An overview of immunity and defense......Page 1003
21.3 How pathogens and pests cause disease......Page 1007
21.4 Preformed defenses......Page 1027
21.5 Induced defense......Page 1030
21.6 Effector-triggered immunity, a second level of induced defense......Page 1040
21.7 Other sources of genetic variation for resistance......Page 1050
21.8 Local and systemic defense signaling......Page 1051
21.9 Plant gene silencing confers virus resistance, tolerance, and attenuation......Page 1060
21.10 Control of plant pathogens by genetic engineering......Page 1062
Summary......Page 1068
22.1 Plant responses to abiotic stress......Page 1069
22.2 Physiological and cellular responses to water deficit......Page 1072
22.3 Gene expression and signal transduction in response to dehydration......Page 1079
22.4 Freezing and chilling stress......Page 1086
22.5 Flooding and oxygen deficit......Page 1094
22.6 Oxidative stress......Page 1103
22.7 Heat stress......Page 1112
22.8 Crosstalk in stress responses......Page 1115
Summary......Page 1117
Introduction......Page 1119
23.1 Overview of essential mineral elements......Page 1120
23.2 Mechanisms and regulation of plant K+ transport......Page 1121
23.3 Phosphorus nutrition and transport......Page 1131
23.4 The molecular physiology of micronutrient acquisition......Page 1136
23.5 Plant responses to mineral toxicity......Page 1145
Summary......Page 1149
Introduction......Page 1150
24.1 Terpenoids......Page 1151
24.2 Biosynthesis of the basic five-carbon unit......Page 1153
24.3 Repetitive additions of C5 units......Page 1156
24.4 Formation of parent carbon skeletons......Page 1159
24.5 Modification of terpenoid skeletons......Page 1161
24.6 Metabolic engineering of terpenoid production......Page 1163
24.7 Cyanogenic glycosides......Page 1164
24.8 Cyanogenic glycoside biosynthesis......Page 1170
24.9 Functions of cyanogenic glycosides......Page 1175
24.10 Glucosinolates......Page 1176
24.11 Alkaloids......Page 1177
24.12 Alkaloid biosynthesis......Page 1182
24.13 Biotechnological application of alkaloid biosynthesis research......Page 1189
24.14 Phenolic compounds......Page 1196
24.15 Phenolic biosynthesis......Page 1203
24.16 The phenylpropanoid-acetate pathway......Page 1206
24.17 The phenylpropanoid pathway......Page 1213
24.18 Universal features of phenolic biosynthesis......Page 1220
24.19 Evolution of secondary pathways......Page 1223
Summary......Page 1224
Chapter 2......Page 1225
Chapter 4......Page 1226
Chapter 6......Page 1227
Chapter 9......Page 1228
Chapter 10......Page 1229
Chapter 13......Page 1230
Chapter 14......Page 1231
Chapter 16......Page 1232
Chapter 19......Page 1234
Chapter 20......Page 1235
Chapter 22......Page 1236
Chapter 23......Page 1237
Chapter 24......Page 1238
Index......Page 1240
EULA......Page 1283

Citation preview

BIOCHEMISTRY & MOLECULAR BIOLOGY OF PLANTS

BIOCHEMISTRY & MOLECULAR BIOLOGY OF PLANTS

Second Edition

EDITED BY

Bob B. Buchanan,Wilhelm Gruissem, and Russell L. Jones

This edition first published 2015 © 2015 by John Wiley & Sons, Ltd Registered Office John Wiley & Sons, Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Editorial Offices 9600 Garsington Road, Oxford, OX4 2DQ, UK The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK 111 River Street, Hoboken, NJ 07030‐5774, USA For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/wiley‐blackwell. The right of the author to be identified as the author of this work has been asserted in accordance with the UK 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. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. Limit of Liability/Disclaimer of Warranty: While the publisher and author(s) have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. It is sold on the understanding that the publisher is not engaged in rendering professional services and neither the publisher nor the author shall be liable for damages arising herefrom. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging‐in‐Publication Data are available. Paperback ISBN: 9780470714218 Hardback ISBN: 9780470714225 A catalogue record for this book is available from the British Library. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Cover image: The illustration on the cover shows a fluorescence image of an Arabidopsis epidermal cell depicting the localization of cellulose synthase (CESA, green) and microtubules (red). The overlying graphic shows how the synthesis of a cellulose microfibril (yellow) is related to the CESA complex, portrayed as a rosette of six light green particles embedded in the plasma membrane that are attached to a microtubule by a purple linker protein (CSI1). Fluorescent image courtesy of Chris Somerville and Trevor Yeats, Energy Biosciences Institute, University of California, Berkeley. Cover design by Dan Jubb. Complex illustrations by Debbie Maizels, Zoobotanica Scientific Illustration. Set in 10/12pt Minion by SPi Global, Pondicherry, India

1 2015

BRIEF CONTENTS I 1

COMPARTMENTS

IV

Membrane Structure and Membranous Organelles  2

METABOLIC AND DEVELOPMENTAL INTEGRATION

2 The Cell Wall  45

15 Long‐Distance Transport  658

3 Membrane Transport  111

16 Nitrogen and Sulfur  711

4 Protein Sorting and Vesicle Traffic  151

17 Biosynthesis of Hormones  769

5 The Cytoskeleton  191

18 Signal Transduction  834 19 Molecular Regulation of Reproductive

II

CELL REPRODUCTION

Development  872 20 Senescence and Cell Death  925

6 Nucleic Acids  240 7 Amino Acids  289 8 Lipids  337 9 Genome Structure and Organization  401

V

PLANT ENVIRONMENT AND AGRICULTURE

10 Protein Synthesis, Folding, and Degradation  438

21 Responses to Plant Pathogens  984

11 Cell Division  476

22 Responses to Abiotic Stress  1051 23 Mineral Nutrient Acquisition, Transport,

III

ENERGY FLOW

and Utilization  1101 24 Natural Products  1132

12 Photosynthesis  508 13 Carbohydrate Metabolism  567 14 Respiration and Photorespiration  610

v

CONTENTS The Editors  xi List of Contributors  xii Preface  xv About the Companion Website  xvi

I

COMPARTMENTS

1 Membrane Structure and Membranous Organelles  2 Introduction  2 1.1 Common properties and inheritance of cell membranes  2 1.2 The fluid‐mosaic membrane model  4 1.3 Plasma membrane  10 1.4 Endoplasmic reticulum  13 1.5 Golgi apparatus  18 1.6 Exocytosis and endocytosis  23 1.7 Vacuoles  27 1.8 The nucleus  28 1.9 Peroxisomes  31 1.10 Plastids  32 1.11 Mitochondria  39 Summary  44

2 The Cell Wall  45 Introduction  45 2.1 Sugars are building blocks of the cell wall  45 2.2 Macromolecules of the cell wall  51 2.3 Cell wall architecture  73 2.4 Cell wall biosynthesis and assembly  80 2.5 Growth and cell walls  90 2.6 Cell differentiation  99 2.7 Cell walls as sources of food, feed, fiber, and fuel, and their genetic improvement  108 Summary  110

vi

3 Membrane Transport  111 Introduction  111 3.1 Overview of plant membrane transport systems  111 3.2 Pumps  120 3.3 Ion channels  128 3.4 Cotransporters  142 3.5 Water transport through aquaporins  146 Summary  148

4 Protein Sorting and Vesicle Traffic  151 Introduction  151 4.1 The cellular machinery of protein sorting  151 4.2 Targeting proteins to the plastids  153 4.3 Targeting proteins to mitochondria  157 4.4 Targeting proteins to peroxisomes  159 4.5 Transport in and out of the nucleus  160 4.6 ER is the secretory pathway port of entry and a protein nursery  161 4.7 Protein traffic and sorting in the secretory pathway: the ER  175 4.8 Protein traffic and sorting in the secretory pathway: the Golgi apparatus and beyond  182 4.9 Endocytosis and endosomal compartments  188 Summary  189

5 The Cytoskeleton  191 Introduction  191 5.1 Introduction to the cytoskeleton  191 5.2 Actin and tubulin gene families  194 5.3 Characteristics of actin filaments and microtubules  196 5.4 Cytoskeletal accessory proteins  202 5.5 Observing the cytoskeleton: Statics and dynamics  207 5.6 Role of actin filaments in directed intracellular movement  210 5.7 Cortical microtubules and expansion  216 5.8 The cytoskeleton and signal transduction  219 5.9 Mitosis and cytokinesis  222 Summary  238

CONTENTS

II

CELL REPRODUCTION

6 Nucleic Acids  240 Introduction  240 6.1 Composition of nucleic acids and synthesis of nucleotides  240 6.2 Replication of nuclear DNA  245 6.3 DNA repair  250 6.4 DNA recombination  255 6.5 Organellar DNA  260 6.6 DNA transcription  268 6.7 Characteristics and functions of RNA  270 6.8 RNA processing  278 Summary  288

7 Amino Acids  289 Introduction  289 7.1 Amino acid biosynthesis in plants: research and prospects  289 7.2 Assimilation of inorganic nitrogen into N‐transport amino acids  292 7.3 Aromatic amino acids  302 7.4 Aspartate‐derived amino acids  318 7.5 Branched‐chain amino acids  326 7.6 Glutamate‐derived amino acids  330 7.7 Histidine  333 Summary  336

8 Lipids  337 Introduction  337 8.1 Structure and function of lipids  337 8.2 Fatty acid biosynthesis  344 8.3 Acetyl‐CoA carboxylase  348 8.4 Fatty acid synthase  350 8.5 Desaturation and elongation of C16 and C18 fatty acids  352 8.6 Synthesis of unusual fatty acids  360 8.7 Synthesis of membrane lipids  365 8.8 Function of membrane lipids  373 8.9 Synthesis and function of extracellular lipids  382 8.10 Synthesis and catabolism of storage lipids  389 8.11 Genetic engineering of lipids  395 Summary  400

9 Genome Structure and Organization  401 Introduction  401 9.1 Genome structure: a 21st‐century perspective  401 9.2 Genome organization  404 9.3 Transposable elements  416 9.4 Gene expression  422 9.5 Chromatin and the epigenetic regulation of gene expression  430 Summary  436

10 Protein Synthesis, Folding, and Degradation  438 Introduction  438 10.1 Organellar compartmentalization of protein synthesis  438 10.2 From RNA to protein  439 10.3 Mechanisms of plant viral translation  447 10.4 Protein synthesis in plastids  450 10.5 Post‐translational modification of proteins  457 10.6 Protein degradation  463 Summary  475

11 Cell Division  476 Introduction  476 11.1 Animal and plant cell cycles  476 11.2 Historical perspective on cell cycle research  477 11.3 Mechanisms of cell cycle control  482 11.4 The cell cycle in action  488 11.5 Cell cycle control during development  497 Summary  506

III

ENERGY FLOW

12 Photosynthesis  508 Introduction  508 12.1 Overview of photosynthesis  508 12.2 Light absorption and energy conversion  511 12.3 Photosystem structure and function  519 12.4 Electron transport pathways in chloroplast membranes  529 12.5 ATP synthesis in chloroplasts  537 12.6 Organization and regulation of photosynthetic complexes  540 12.7 Carbon reactions: the Calvin–Benson cycle  542

vii

viii

CONTENTS

12.8 Rubisco  548 12.9 Regulation of the Calvin–Benson cycle by light  551 12.10 Variations in mechanisms of CO2 fixation  557 Summary  565

13 Carbohydrate Metabolism  567 Introduction  567 13.1 The concept of metabolite pools  570 13.2 The hexose phosphate pool: a major crossroads in plant metabolism  571 13.3 Sucrose biosynthesis  573 13.4 Sucrose metabolism  577 13.5 Starch biosynthesis  580 13.6 Partitioning of photoassimilates between sucrose and starch  587 13.7 Starch degradation  593 13.8 The pentose phosphate/triose phosphate pool  597 13.9 Energy and reducing power for biosynthesis  601 13.10 Sugar‐regulated gene expression  606 Summary  608

14 Respiration and Photorespiration  610 Introduction  610 14.1 Overview of respiration  610 14.2 Citric acid cycle  613 14.3 Plant mitochondrial electron transport  620 14.4 Plant mitochondrial ATP synthesis  632 14.5 Regulation of the citric acid cycle and the cytochrome pathway  634 14.6 Integration of the cytochrome pathway and nonphosphorylating pathways  635 14.7 Interactions between mitochondria and other cellular compartments  639 14.8 Biochemical basis of photorespiration  646 14.9 The photorespiratory pathway  648 14.10 Role of photorespiration in plants  652 Summary  655

IV

METABOLIC AND DEVELOPMENTAL INTEGRATION

15 Long‐Distance Transport  658 Introduction  658 15.1 Selection pressures and long‐distance transport systems  658

15.2 Cell biology of transport modules  664 15.3 Short-distance transport events between xylem and nonvascular cells  668 15.4 Short‐distance transport events between phloem and nonvascular cells  673 15.5 Whole‐plant organization of xylem transport  691 15.6 Whole‐plant organization of phloem transport  696 15.7 Communication and regulation controlling phloem transport events  705 Summary  710

16 Nitrogen and Sulfur  711 Introduction  711 16.1 Overview of nitrogen in the biosphere and in plants  711 16.2 Overview of biological nitrogen fixation  715 16.3 Enzymology of nitrogen fixation  715 16.4 Symbiotic nitrogen fixation  718 16.5 Ammonia uptake and transport  735 16.6 Nitrate uptake and transport  735 16.7 Nitrate reduction  739 16.8 Nitrite reduction  744 16.9 Nitrate signaling  745 16.10 Interaction between nitrate assimilation and carbon metabolism  745 16.11 Overview of sulfur in the biosphere and plants  746 16.12 Sulfur chemistry and function  747 16.13 Sulfate uptake and transport  750 16.14 The reductive sulfate assimilation pathway  752 16.15 Cysteine synthesis  755 16.16 Synthesis and function of glutathione and its derivatives  758 16.17 Sulfated compounds  763 16.18 Regulation of sulfate assimilation and interaction with nitrogen and carbon metabolism  764 Summary  767

17 Biosynthesis of Hormones  769 Introduction  769 17.1 Gibberellins  769 17.2 Abscisic acid  777 17.3 Cytokinins  785 17.4 Auxins  795 17.5 Ethylene  806 17.6 Brassinosteroids  810 17.7 Polyamines  818 17.8 Jasmonic acid  821 17.9 Salicylic acid  826

CONTENTS

17.10 Strigolactones  830 Summary  833

18 Signal Transduction  834 Introduction  834 18.1 Characteristics of signal perception, transduction, and integration in plants  834 18.2 Overview of signal perception at the plasma membrane  838 18.3 Intracellular signal transduction, amplification, and integration via second messengers and MAPK cascades  843 18.4 Ethylene signal transduction  847 18.5 Cytokinin signal transduction  850 18.6 Integration of auxin signaling and transport  852 18.7 Signal transduction from phytochromes  857 18.8 Gibberellin signal transduction and its integration with phytochrome signaling during seedling development  861 18.9 Integration of light, ABA, and CO2 signals in the regulation of stomatal aperture  866 18.10 Prospects  870 Summary  870

19 Molecular Regulation of Reproductive Development  872 Introduction  872 19.1 The transition from vegetative to reproductive development  872 19.2 The molecular basis of flower development  881 19.3 The formation of male gametes  889 19.4 The formation of female gametes  897 19.5 Pollination and fertilization  902 19.6 The molecular basis of self‐incompatibility  908 19.7 Seed development  913 Summary  923

20 Senescence and Cell Death  925 Introduction  925 20.1 Types of cell death  925 20.2 PCD during seed development and germination  930 20.3 Cell death during the development of secretory bodies, defensive structures and organ shapes  932 20.4 PCD during reproductive development  937 20.5 Senescence and PCD in the terminal development of leaves and other lateral organs  940 20.6 Pigment metabolism in senescence  948

20.7 Macromolecule breakdown and salvage of nutrients in senescence  951 20.8 Energy and oxidative metabolism during senescence  957 20.9 Environmental influences on senescence and cell death I: Abiotic interactions  961 20.10 Environmental influences on senescence and cell death II: PCD responses to pathogen attack  964 20.11 Plant hormones in senescence and defense‐related PCD  974 Summary  982

V

PLANT ENVIRONMENT AND AGRICULTURE

21 Responses to Plant Pathogens  984 Introduction  984 21.1 Pathogens, pests, and disease  984 21.2 An overview of immunity and defense  985 21.3 How pathogens and pests cause disease  989 21.4 Preformed defenses  1009 21.5 Induced defense  1012 21.6 Effector‐triggered immunity, a second level of induced defense  1022 21.7 Other sources of genetic variation for resistance  1032 21.8 Local and systemic defense signaling  1033 21.9 Plant gene silencing confers virus resistance, ­tolerance, and attenuation  1042 21.10 Control of plant pathogens by genetic engineering  1044 Summary  1050

22 Responses to Abiotic Stress  1051 Introduction  1051 22.1 Plant responses to abiotic stress  1051 22.2 Physiological and cellular responses to water deficit  1054 22.3 Gene expression and signal transduction in response to dehydration  1061 22.4 Freezing and chilling stress  1068 22.5 Flooding and oxygen deficit  1076 22.6 Oxidative stress  1085 22.7 Heat stress  1094 22.8 Crosstalk in stress responses  1097 Summary  1099

ix

x

CONTENTS

23 Mineral Nutrient Acquisition, Transport, and Utilization  1101 Introduction  1101 23.1 Overview of essential mineral elements  1102 23.2 Mechanisms and regulation of plant K+ transport  1103 23.3 Phosphorus nutrition and transport  1113 23.4 The molecular physiology of micronutrient acquisition  1118 23.5 Plant responses to mineral toxicity  1127 Summary  1131

24 Natural Products  1132 Introduction  1132 24.1 Terpenoids  1133 24.2 Biosynthesis of the basic five‐carbon unit  1135 24.3 Repetitive additions of C5 units  1138 24.4 Formation of parent carbon skeletons  1141 24.5 Modification of terpenoid skeletons  1143

24.6 Metabolic engineering of terpenoid production  1145 24.7 Cyanogenic glycosides  1146 24.8 Cyanogenic glycoside biosynthesis  1152 24.9 Functions of cyanogenic glycosides  1157 24.10 Glucosinolates  1158 24.11 Alkaloids  1159 24.12 Alkaloid biosynthesis  1164 24.13 Biotechnological application of alkaloid biosynthesis research  1171 24.14 Phenolic compounds  1178 24.15 Phenolic biosynthesis  1185 24.16 The phenylpropanoid‐acetate pathway  1188 24.17 The phenylpropanoid pathway  1195 24.18 Universal features of phenolic biosynthesis  1202 24.19 Evolution of secondary pathways  1205 Summary  1206 Further reading 1207 Index 1222

The Editors Bob B. Buchanan A native Virginian, Bob B. Buchanan obtained his PhD in microbiology at Duke University and did postdoctoral research at the University of California at Berkeley. In 1963, he joined the Berkeley faculty and is currently a professor emeritus in the Department of Plant and Microbial Biology. He has taught general biology and biochemistry to undergraduate students and graduate-level courses in plant biochemistry and photosynthesis. Initially focused on pathways and regulatory mechanisms in photosynthesis, his research has more recently dealt with the regulatory role of thioredoxin in seeds, plant mitochondria and methane-producing archaea. The work on seeds is finding application in several areas. Bob has served as department chair at UC Berkeley and was president of the American Society of Plant Physiologists from 1995 to 1996. A former Guggenheim Fellow, he is a member of the National Academy of Sciences and the Japanese Society of Plant Physiologists (honorary). He is a ­fellow of the American Academy of Arts and Sciences, the American Society of Microbiology, the American Society of Plant Biologists, and the American Association for the Advancement of Science. His other honors include the Bessenyei Medal from the Hungarian Ministry of Education, the Kettering Award for Excellence in Photosynthesis, and the Stephen Hales Prize from the American Society of Plant Physiologists, a Research Award from the Alexander von Humboldt Foundation, the Distinguished Achievement Award from his undergraduate alma mater, Emory and Henry College, and the Berkeley Citation. Wilhelm Gruissem Wilhelm Gruissem was born in Germany where he studied biology and chemistry. After obtaining his PhD in 1979 at the University of Bonn in Germany and postdoctoral research at the University of Marburg in Germany and the University of Colorado in Boulder, he was appointed as Professor of Plant Biology at the University of California at Berkeley in 1983. He was Chair of the Department of Plant and Microbial Biology at UC Berkeley from 1993 to 1998, and from 1998 to 2000 he was Director of a collaborative research program between the Department and the Novartis Agricultural Discovery Institute in San Diego. In 2000 he joined the ETH Zurich (Swiss Federal Institute of Technology) as Professor of Plant Biotechnology in the Department of Biology and the Institute

of Agricultural Sciences. Since 2001 he has been Co-Director of the Functional Genomics Center Zurich. From 2006 to 2010 he served as President of the European Plant Science Organization (EPSO) and since 2011 as Chair of the Global Plant Council. From 2009 to 2011 he also served as Chair of the Department of Biology at ETH Zurich. In addition to his research on systems approaches to understand pathways and molecules involved in plant growth control, he directs a ­biotechnology program on trait improvement in cassava, rice, and wheat. In 2008 he founded Nebion, a bioinformatics company building the internationally successful Genevestigator database. He is an elected fellow of the American Association for the Advancement of Sciences (AAAS) and the American Society of Plant Biologists, he is Editor of Plant Molecular Biology, and he serves on the editorial boards of several journals and on advisory boards for various research institutions. He has received several prestigious awards, including a prize from the Fiat Panis Foundation in Germany and the Shang-Fa Yang award of Academia Sinica in Taiwan for his trait improvement work in cassava and rice. In 2007 he was elected lifetime foreign member of the American Society of Plant Biologists. Russell L. Jones Russell L. Jones was born in Wales and completed his BSc and PhD degrees at the University of Wales, Aberystwyth. He spent 1 year as a postdoctoral fellow at the Michigan State University Department of Energy Plant Research Laboratory with Anton Lang before being appointed to the faculty of the Department of Botany at the University of California at Berkeley in 1966. As Professor of Plant Biology at UC Berkeley he taught undergraduate classes in general biology and graduate courses in plant physiology and cell biology for over 45 years. He is now Professor Emeritus, Department of Plant and Microbial Biology at UC Berkeley. His research focuses on hormonal regulation in plants using the cereal aleurone as a model system, with approaches that exploit the techniques of biochemistry, biophysics, and cell and molecular biology. Russell was president of the American Society of Plant Physiologists from 1993 to 1994. He was a Guggenheim Fellow at the University of Nottingham in 1972, a Miller Professor at UC Berkeley in 1976, a Humboldt Prize Winner at the University of Göttingen in 1986, and a RIKEN Eminent Scientist, RIKEN, Japan, in 1996.

xi

LIst of CONTRIBUTORS Nikolaus Amrhein 

Institute of Plant Science,

Shaun Curtin  Department of Plant Pathology, University of Minnesota, St Paul, MN, USA

Julia Bailey‐Serres  Department of Botany and Plant Sciences, University of California, Riverside, CA, USA

David Day  Division of Biochemistry and Molecular Biology, Australian National University, Canberra, Australia

Tobias I. Baskin 

Stephen Day 

ETH Zurich, Switzerland

Department of Biological Science, University of Missouri, Columbia, MO, USA

Paul C. Bethke  Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA Gerard Bishop  Department

of Life Sciences, Imperial College London, London, United Kingdom

Elizabeth A. Bray  Erman University of Chicago, Chicago, IL, USA

Biology Center,

Karen S. Browning  Department of Chemistry and Biochemistry, University of Texas, Austin, TX, USA

Deceased

Emmanuel Delhaize  Lieven De Veylder 

CSIRO, Clayton, Australia

Universiteit Gent, Gent, Belgium

Natalia Dudareva  Horticulture and Landscape Architecture, Purdue University, West Lafayette, IN, USA David R. Gang  Institute of Biological Chemistry, Washington State University, Pullman, WA, USA Walter Gassmann  Division of Plant Sciences, University of Missouri, Columbia, MO, USA

John Browse  Institute of Biological Chemistry, Washington State University, Pullman, WA, USA

Jonathan Gershenzon Department of Biochemistry, MPI for Chemical Ecology, Jena, Germany

Judy Callis 

Ueli Grossniklaus  Institute of Plant Biology, University of Zurich, Zurich, Switzerland

University of California, Davis, CA, USA

Nicholas C. Carpita 

Department of Botany and Plant Pathology, Purdue University, Lafayette, IN, USA

Kim E. Hammond‐Kosack  Rothamsted Research, Harpenden, United Kingdom

Maarten J. Chrispeels 

Department of Biology, University of California, San Diego, CA, USA

Dirk Inzé 

Gloria Coruzzi  Department

Stefan Jansson 

York University, New York City, NY, USA

xii

of Biology, New

Universiteit Gent, Gent, Belgium

University, Umeå, Sweden

Umeå Plant Science Centre, Umeå

list of Contributors

Jan Jaworski  Department of Chemistry, Miami University, Miami, FL, USA Jonathan D. G. Jones  The Sainsbury Laboratory, John Innes Centre, Norwich, United Kingdom Michael Kahn 

Institute of Biological Chemistry, Washington State University, Pullman, WA, USA

Leon Kochian  U.S.

Plant, Soil and Nutrition Laboratory, Cornell University, Ithaca, NY, USA

Stanislav Kopriva  Department of Metabolic Biology, John Innes Centre, Norwich, United Kingdom Toni M. Kutchan 

Center, St. Louis, MO, USA

Robert Last  MA, USA

Donald Danforth Plant Science

Cereon Genomics LLP, Cambridge,

Ottoline Leyser  The Sainsbury Laboratory, University of Cambridge, Cambridge, United Kingdom Birger Lindberg Møller 

Center for Synthetic Biology, Plant Biochemistry Laboratory, Department of Plant and Environmental Sciences, University of Copenhagen, Copenhagen, Denmark and Carlsberg Laboratory, Copenhagen, Denmark

Sharon R. Long  Department

of Biological Sciences, Stanford University, Stanford, CA, USA

Richard

Malkin Department

of Plant and Microbial Biology, University of California, Berkeley, CA, USA

Maureen C. McCann 

Department of Biological Sciences, Purdue University, West Lafayette, USA

A. Harvey Millar 

Luis Mur  Institute of Biological, Environmental and Rural Sciences, Aberystwyth University, Aberystwyth, Wales, UK Krishna K. Niyogi  Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA John Ohlrogge 

Department of Botany, Michigan State University, East Lansing, USA

Helen Ougham  Institute of Biological, Environmental and Rural Sciences, University of Aberystwyth, Aberystwyth, Wales, UK John W. Patrick  School of Environmental and Life Sciences, University of Newcastle, Newcastle, Australia Natasha V. Raikhel 

MSU−DOE Plant Research Laboratory, Michigan State University, East Lansing , MI, USA

John Ralph 

Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI, USA

Peter R. Ryan 

Canberra, Australia

Division of Plant Industry, CSIRO,

Hitoshi Sakakibara RIKEN

Center, Yokohama, Japan

Plant Science

Daniel Schachtman 

Department of Agronomy and Horticulture, University of Nebraska, Lincoln, NE, USA

Danny Schnell 

Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA

Australian Academy of Science,

Julian L. Schroeder  Biological Sciences, University of California, San Diego, CA, USA

Research School of Biological Sciences, Australian National University, Canberra, Australia

Lance Seefeldt  Department of Chemistry and Biochemistry, Utah State University, Logan, UT, USA

Acton, Australia

Tony Millar 

xiii

xiv

list of Contributors

Mitsunori Seo  RIKEN

Plant Science Center,

Yi‐Fang Tsay  Institute

Kazuo Shinozaki  RIKEN Center for Sustainable Resource Science, Yokohama, Japan

Stephen D. Tyerman 

James N. Siedow 

Matsuo Uemura  Iwate

Yokohama, Japan

University, Durham, NC, USA

Department of Botany, Duke

School of Agriculture, Food and Wine, Adelaide University, Adelaide, Australia

Iwate, Japan

Ian Small  Plant Energy Biology, ARC Center of Excellence, The University of Western Australia, Crawley, Australia

Aart J. E. van Bel 

Chris Somerville 

Biotechnology, Milan, Italy

Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA

Linda Spremulli  Department

of Chemistry, University of North Carolina, Chapel Hill, NC, USA

L. Andrew Staehelin 

Department of Molecular and Cell Development Biology, University of Colorado, Boulder, CO, USA

Masahiro Sugiura 

Nagoya University, Japan

Yutaka Takeda 

Japan

Centre for Gene Research,

Okayama University, Okayama,

Howard

Thomas Institute of Biological, Environmental and Rural Sciences, University of Aberystwyth, Wales, UK

Christopher D. Town  San Diego, CA, USA

J. Craig Venter Institute,

of Molecular Biology,

Academia Sinica, Taiwan

University, Morioka,

Institute for General Botany, Justus‐Liebig‐University, Giessen, Germany

Alessandro Vitale  Institute

of Agricultural

John M. Ward 

College of Biological Sciences, University of Minnesota, MN, USA

Peter

Waterhouse School of Molecular Bioscience, The University of Sydney, Sydney, Australia

Frank Wellmer 

Smurfit Institute of Genetics, Trinity College, Dublin, Ireland

Elizabeth Weretilnyk  Department of Biology, McMaster University, Hamilton, Ontario, Canada Ricardo A.Wolosiuk  Instituto de Investigaciones Bioquímicas, Buenos Aires, Argentina Shinjiro Yamaguchi  RIKEN Center,, Yokohama, Japan

Samuel C. Zeeman 

ETH Zurich, Switzerland

Plant Science

Institute of Plant Science,

Preface

T

he second edition of the Biochemistry & Molecular Biology of Plants retains the overall format of the first edition in response to the enthusiastic feedback we received from users of the book. The first edition was organized into five sections dealing with organization and functioning of the cell (Compartments), the cell’s ability to replicate (Cell Reproduction), generation of energy (Energy Flow), regulation of development (Metabolism and Developmental Regulation), and the impact of fundamental discoveries in plant biology (Plant, Environment, and Agriculture). Although the section organization of the second edition remains unchanged, many of the chapters have been written by new teams of authors, reflecting the retirement of some of our colleagues, but also the dynamic development of plant biology during the last 20 years that was driven by a cohort of younger investigators, many of whom have contributed to this second edition. Changes in chapter authorship also reflect the impact that molecular genetics had on our field, and three chapters stand out in this regard: Chapter  9 on Genome Structure and Organization, Chapter  18 on Signal Transduction, and Chapter  19 on Molecular Regulation of Reproductive Development. Advances resulting from molecular genetics have been particularly dramatic in the field of plant hormones and other signaling molecules where the receptors for all of the major hormones and their complex signaling pathways have now been described in detail. Soon after publication of the first edition, Biochemistry & Molecular Biology of Plants was translated into Chinese, Italian, and Japanese, and a special low‐priced English‐­language version of the book was published in India. In this version the entire book was published in black and white, illustrating the costs involved in producing four‐color ­versions of textbooks. Another change that accompanied the writing and ­production of this second edition was the involvement of the publisher John Wiley and our interaction with the Editorial

Office in the United Kingdom. Wiley had entered into an agreement with the American Society of Plant Biologists to lead the publication of books written by ASPB members. The second edition of Biochemistry & Molecular Biology of Plants is one of the first of hopefully many books that will be published jointly by ASPB and Wiley. Production of this book required input from many ­talented people. First and foremost the authors, who patiently, in some cases very patiently, worked with the editors and developmental editors to produce chapters of remarkably high ­quality. The two excellent developmental editors, Justine Walsh and Yolanda Kowalewski, worked to produce a collection of chapters that read seamlessly; the artist Debbie Maizels ­ ­produced figures of exceptional technical and artistic quality; the staff at John Wiley, who worked tirelessly on this p ­ roject; and Dr Nik Prowse, freelance project manager, who efficiently handled the chapter editing and management during the production phase of the book.. Special thanks go to Celia Carden whose support, enthusiasm, and management across two continents have gone a long way to making this book successful. The support of ASPB’s leadership and staff, notably Executive Director Crispin Taylor and Publications Manager Nancy Winchester, are gratefully acknowledged. We also appreciate the continuing/ongoing support that we received from ASPB as this book was being developed. The contributing authors thank reviewers for commenting on their chapters. Most important, we want to express appreciation to our wives, Melinda, Barbara, and Frances, who during the past few years again tolerated and accepted the textbook as a demanding family member. Bob B. Buchanan Wilhelm Gruissem Russell L. Jones November, 2014 Berkeley, CA, and Zurich, Switzerland

Note: Following the common publishing convention, species names that appear in the italicized figure legends have been set in standard roman typeface so that they are easily identifiable. xv

ABOUT THE COMPANION WEBSITE

This book is accompanied by a companion website: www.wiley.com/go/buchanan/biochem This website includes: ●● ●●

PowerPoint slides of all the figures from the book, to download; PDF files of all the tables from the book, to download.

I

COMPARTMENTS

1 Membrane Structure and Membranous Organelles L. Andrew Staehelin Introduction Cells, the basic units of life, require membranes for their existence. Foremost among these is the plasma membrane, which defines each cell’s boundary and helps create and maintain electrochemically distinct environments within and outside the cell. Other membranes enclose eukaryotic orga­ nelles such as the nucleus, chloroplasts, and mitochondria. Membranes also form internal compartments, such as the endoplasmic reticulum (ER) in the cytoplasm and thylakoids in the chloroplast (Fig. 1.1). The principal function of membranes is to serve as a barrier to diffusion of most water‐soluble molecules. Cellular compart­ ments delimited by membranes can differ in chemical compo­ sition from their surroundings and be optimized for a particular activity. Membranes also serve as scaffolding for certain pro­ teins. As membrane components, proteins perform a wide array of functions: transporting molecules and transmitting signals across the membrane, processing lipids enzymatically, assembling glycoproteins and polysaccharides, and providing mechanical links between cytosolic and cell wall molecules. This chapter is divided into two parts. The first is devoted to the general features and molecular organization of mem­ branes. The second provides an introduction to the architecture and functions of the different membranous organelles of plant cells. Many later chapters of this book focus on metabolic events that involve these organelles.

1.1  Common properties and inheritance of cell membranes 1.1.1  Cell membranes possess common structural and functional properties All cell membranes consist of a bilayer of polar lipid mol­ ecules and associated proteins. In an aqueous environment, membrane lipids self‐assemble with their hydrocarbon tails clustered together, protected from contact with water (Fig. 1.2). Besides mediating the formation of bilayers, this property causes membranes to form closed compartments. As a result, every membrane is an asymmetrical structure, with one side exposed to the contents inside the compart­ ment and the other side in contact with the external solution. The lipid bilayer serves as a general permeability barrier because most water‐soluble (polar) molecules cannot readily traverse its nonpolar interior. Proteins perform most of the other membrane functions and thereby define the specificity of each membrane system. Virtually all membrane molecules are able to diffuse freely within the plane of the membrane, permitting membranes to change shape and membrane mol­ ecules to rearrange rapidly.

Biochemistry & Molecular Biology of Plants, Second Edition. Edited by Bob B. Buchanan, Wilhelm Gruissem, and Russell L. Jones. © 2015 John Wiley & Sons, Ltd. Published 2015 by John Wiley & Sons, Ltd. Companion website: www.wiley.com/go/buchanan/biochem

2

Chapter 1  Membrane Structure and Membranous Organelles

PM Nuclear membrane

Nucleus

Nucleolus

V N

M

G CW

A

ER

B

Vacuole

Peroxisome Golgi body

Smooth endoplasmic reticulum Chloroplast Air space Mitochondrion Rough endoplasmic reticulum

Middle lamella Plasma membrane

Cell wall

A

Cellulose/ hemicellulose wall Pectin-rich middle lamella

FIGURE 1.1  (A) Diagrammatic representation of a mesophyll leaf cell, depicting principal membrane systems and cell wall domains of a differentiated plant cell. Note the large volume occupied by the vacuole. (B) Thin‐section transmission electron micrograph (TEM) through a Nicotiana meristematic root tip cell preserved by rapid freezing. The principal membrane systems shown include amyloplast (A), endoplasmic reticulum (ER), Golgi stack (G), mitochondrion (M), nucleus (N), vacuole (V), and plasma membrane (PM). Cell wall (CW). Source: (B) Micrograph by Thomas Giddings Jr., from Staehelin et al. (1990). Protoplasma 157: 75–91.

1.1.2  All basic types of cell membranes are inherited Plant cells contain approximately 20 different membrane systems. The exact number depends on how sets of related membranes are counted (Table 1.1). From the moment they are formed, cells must maintain the integrity of all their membrane‐bounded compartments to survive, so all mem­ brane systems must be passed from one generation of cells to

the next in a functionally active form. Membrane inheritance follows certain rules: ●●

●●

●●

Daughter cells inherit a complete set of membrane types from their mother. Each potential mother cell maintains a complete set of membranes. New membranes arise by growth and fission of existing membranes.

3

4

Part I  COMPARTMENTS TABLE 1.1  Membrane types found in plant cells. Hydrophilic head group

Plasma membrane Nuclear envelope membranes (inner/outer) Endoplasmic reticulum Golgi cisternae (cis, medial, trans types)

Lipid micelle

Trans‐Golgi network/early endosome membranes Clathrin‐coated,COPIa/Ib*, COPII*, secretory and retromer vesicle membranes Autophagic vacuole membrane

Hydrophobic tail

Multivesicular body/late endosome membranes Tonoplast membranes (lytic/storage vacuoles) Peroxisomal membrane

Lipid bilayer

Glyoxysomal membrane Chloroplast envelope membranes (inner/ outer) Thylakoid membrane Mitochondrial membranes (inner/outer)

FIGURE 1.2  Cross‐sectional views of a lipid micelle and a lipid bilayer in aqueous solution.

1.2  The fluid‐mosaic membrane model The fluid‐mosaic membrane model describes the molecular organization of lipids and proteins in cellular membranes and illustrates how a membrane’s mechanical and physio­ logical traits are defined by the physicochemical character­ istics of its various molecular components. This model integrates much of what we know about the molecular properties of membrane lipids, their assembly into bilayers, the regulation of membrane fluidity, and the different mechanisms by which membrane proteins associate with lipid bilayers.

1.2.1  The amphipathic nature of membrane lipids allows for the spontaneous assembly of bilayers In most cell membranes, lipids and glycoproteins make roughly equal contributions to the membrane’s mass. Lipids  belong to several classes, including phospholipids,

*COP, coat protein.

glucocerebrosides, galactosylglycerides, and sterols (Figs. 1.3 and 1.4). These molecules share an important physico­ chemical property: they are amphipathic, containing both hydrophilic (“water‐loving”) and hydrophobic (“water‐ fearing”) domains. When brought into contact with water, these molecules spontaneously self‐assemble into higher‐ order structures. The hydrophilic head groups maximize their interactions with water molecules, whereas hydropho­ bic tails interact with each other, minimizing their exposure to the aqueous phase (see Fig.  1.2). The geometry of the resulting lipid assemblies is governed by the shape of the amphipathic molecules and the balance between hydro­ philic and hydrophobic domains. For most membrane lipids, the bilayer configuration is the minimum‐energy self‐assembly structure, that is, the structure that takes the least amount of energy to form in the presence of water (Fig. 1.5). In this configuration, the polar groups form the interface to the bulk water, and the hydrophobic groups become sequestered in the interior. Phospholipids, the most common type of membrane lipid, have a charged, phosphate‐containing polar head group and two hydrophobic hydrocarbon tails. Fatty acid tails con­ tain between 14 and 24 carbon atoms, and at least one tail has one or more cis double bonds (Fig. 1.6). The kinks introduced by these double bonds influence the packing of the molecules in the lipid bilayer, and the packing, in turn, affects the overall fluidity of the membrane.

Chapter 1  Membrane Structure and Membranous Organelles FIGURE 1.3  Plant membrane lipids.

O O Glycerol 1

CH2

CH

O

O

C

O C

3

Glucose Sphingosine

Glycerol

O 2

Galactose

O–

P

1

CH2

O

CH2

O 2

3

CH

O

O

C

O C

OH 3

CH2

O

CH

O 2

CH

CH

NH

CH

C

1

Polar (hydrophilic)

Choline, ethanolamine, or serine

CH2

O Nonpolar (hydrophobic)

Fatty acid Fatty acids

Fatty acids

Phospholipid

Galactosylglyceride

Cholesterol

Campesterol

Sitosterol

OH

OH

OH

Glucocerebroside

FIGURE 1.4  Sterols found in plant plasma membranes.

Stigmasterol OH

Hydrophilic Hydrophobic

1.2.2  Phospholipids move rapidly in the plane of the membrane but very slowly from one side of the bilayer to the other Because individual lipid molecules in a bilayer are not bonded to each other covalently, they are free to move. Within the plane of the bilayer, molecules can slide past each other freely. A membrane can assume any shape without disrupting the hydrophobic interactions that stabilize its structure. Aiding this general flexibility is the ability of lipid bilayers to close on themselves to form discrete compartments, a property that also enables them to seal damaged membranes.

Studies of the movement of phospholipids in bilayers have revealed that these molecules can diffuse laterally, rotate, flex their tails, bob up and down, and flip‐flop (Fig. 1.7). The exact mechanism of lateral diffusion is unknown. One theory sug­ gests that individual molecules hop into vacancies (“holes”) that form transiently as the lipid molecules within each mono­ layer exhibit thermal motions. Such vacancies arise in a fluid bilayer at high frequencies, and the average molecule hops ~107 times per second, which translates to a diffusional distance of ~1 μm traversed in a second. Both rotation of individual molecules around their long axes and up‐and‐down bobbing are also very rapid events. Superimposed on these motions is a constant flexing of the hydrocarbon tails. Because this flexing

5

Part I  COMPARTMENTS FIGURE 1.5  Organization of amphipathic lipid molecules in a bilayer.

Phosphatidylcholine

Phosphatidylethanolamine

Cholesterol

FIGURE 1.6  (A) Space‐filling model of a phosphatidylcholine molecule. (B) Diagram defining the functional groups of a phosphatidylcholine molecule.

Choline Polar head group

H

H

H

C

C H

O P

O H H

H C

H C H H C H H C H H C H H C H H C H H C H

C

A

increases towards the ends of the tails, the center of the bilayer has the greatest degree of fluidity. In contrast, spontaneous transfer of phospholipids across the bilayer, called flipping, rarely occurs. A flip would require the polar head to migrate through the nonpolar interior of the bilayer, an energetically unfavorable event. Some membranes contain “flippase” enzymes, which mediate movement of

O

Glycerol

C H O

O O

H C H C

H H

C

H H

C

H H

C

H H

C

H H

C

H H

H C H H

Phosphate

O-

H

C

O

Nonpolar tails

6

C

H C H H C H H C H H C H H C H H C H H C H cis double H C bond H C C H H H H C C H H H H C C H H H C H H

B

newly synthesized lipids across the bilayer (Fig. 1.8). Different flippases specifically catalyze translocation of particular lipid types and thus can flip their lipid substrates in only one direc­ tion. The energy barrier to spontaneous flipping and flippase specificity, together with the specific orientation of the lipid‐ synthesizing enzymes in the membranes, result in an asym­ metrical distribution of lipid types across membrane bilayers.

Chapter 1  Membrane Structure and Membranous Organelles Lateral diffusion Bobbing

FIGURE 1.7  Mobility of phospholipid molecules in a lipid bilayer.

Flexion Rotation Flip-flop

Membrane sterols in lipid bilayers behave somewhat ­ ifferently from phospholipids, primarily because the hydro­ d phobic domain of a sterol molecule is much larger than the uncharged polar head group (see Fig. 1.4). Thus, membrane sterols are not only able to diffuse rapidly in the plane of the bilayer, they can also flip‐flop without enzymatic assistance at a higher rate than phospholipids.

Phospholipid translocator (flippase)

1.2.3  Cells optimize the fluidity of their membranes by controlling lipid composition Like all fatty substances, membrane lipids exist in two differ­ ent physical states, as a semicrystalline gel and as a fluid. Any given lipid, or mixture of lipids, can be melted—converted from gel to fluid—by a temperature increase. This change in state is known as phase transition, and for every lipid this transition occurs at a precise temperature, called the tempera­ ture of melting (Tm, see Table 1.2). Gelling brings most mem­ brane activities to a standstill and increases permeability. At high temperatures, on the other hand, lipids can become too fluid to maintain the permeability barrier. Nonetheless, some organisms live happily in frigid conditions, whereas others thrive in boiling hot springs and thermal vents. Many plants survive daily temperature fluctuations of 30°C. How do organisms adapt the fluidity of their membranes to suit their mutable growth environments? To cope successfully with the problem of temperature‐ dependent changes in membrane fluidity, virtually all poikilo­ thermic organisms—those whose temperatures fluctuate with the environment—can alter the composition of their mem­ branes to optimize fluidity for a given temperature. Mechanisms exploited to compensate for low temperatures include shorten­ ing of fatty acid tails, increasing the number of double bonds, and increasing the size or charge of head groups. Changes in sterol composition can also alter membrane responses to ­temperature. Membrane sterols serve as membrane fluidity

FIGURE 1.8  Mechanism of action of a “flippase,” a phospholipid translocator.

“buffers,” increasing the fluidity at lower temperatures by dis­ rupting the gelling of phospholipids, and decreasing fluidity at high temperatures by interfering with the flexing motions of the fatty acid tails. Because each lipid has a different Tm, lower­ ing the temperature can induce one type of lipid to undergo a fluid‐to‐gel transition and form semicrystalline patches, whereas other lipids remain in the fluid state. Like all cellular molecules, membrane lipids have a finite life span and are turned over on a regular basis. This turnover enables plant cells to adjust the lipid composition of their membranes in response to seasonal changes in ambient temperature.

7

8

Part I  COMPARTMENTS

1.2.4  Membrane proteins associate with lipid bilayers in many different ways

research has led to the discovery of three additional classes of  membrane proteins—fatty acid-linked, prenyl grouplinked, and phosphatidylinositol‐anchored—all of which are attached to the bilayer by lipid tails (Fig. 1.10). By definition, peripheral proteins are water‐soluble and can be removed by washing membranes in water or in salt or acid solutions that do not disrupt the lipid bilayer. Peripheral proteins bind either to integral proteins or to lipids through

The different ways in which membrane‐bound proteins asso­ ciate with lipid bilayers reflect the diversity of enzymatic and structural functions they perform. The original fluid‐mosaic membrane model included two basic types of membrane proteins: peripheral and integral (Fig.  1.9). More recent

TABLE 1.2  The effects of fatty acid chain length and double bonds on the temperature of melting (Tm) of some defined phospholipids. Tm (°C) Types of chains*

Phosphatidylcholine

Phosphatidyl‐ethanolamine

Two C14:0

24

51

Two C16:0

42

63

Two C18:0

55

82

–22

15

Two C18:1 (cis)

Phosphatidic acid

67

 8

*The shorthand nomenclature for the fatty acyl chains denotes how many carbon atoms (first number) and double bonds (second number) they contain.

Outside cell

Oligosaccharide side chains Central plane of lipid bilayer

GPI lipid-anchored protein

Lipid bilayer

Inside cell (cytosol)

Hydrophobic integral membrane protein domains

Integral membrane proteins

Peripheral membrane proteins

Lipid-anchored protein

FIGURE 1.9  A modern version of the fluid‐mosaic membrane model, depicting integral, peripheral, and lipid‐anchored membrane proteins. Not drawn to scale.

Chapter 1  Membrane Structure and Membranous Organelles

C

N

Phosphatidylinositolanchored protein Ethanolamine

P

Galactose Glucosamine

Mannose

Inositol P

Ceramide

Extracellular

Lipid bilayer

NH O

Fatty acid-anchored proteins

Myristic acid C14

C

O

HN

Farnesyl C15

N

S

S

CH2

CH2

CH2

H C

O

O

O

Geranylgeranyl C20

S

Cys

O

Prenyl lipid-anchored protein

Palmitic acid C16

Gly

Amide bond

DiacylO glycerol

OH O

C

C

N

O

O

CH3

H

Thioether linkage

C

C

N

O

O

Cytosol

CH3

N

C N

FIGURE 1.10  Fatty acid-anchored, prenyl lipid-anchored, and glycosylphosphatidylinositol (GPI)‐anchored proteins.

salt bridges, electrostatic interactions, hydrogen bonds, or some combination of these, but they do not penetrate the lipid bilayer. Some peripheral proteins also provide links between membranes and cytoskeletal systems. In contrast, the amphipathic, transmembrane or partly embedded integral proteins are insoluble in water. Because the hydrophobic domains are sequestered in the hydrophobic interior of the bilayer, an integral protein can be removed and solubilized only with the help of detergents or organic solvents, which degrade the bilayer. Both the fatty acid-linked and the prenyl group‐linked proteins bind reversibly to the cytoplasmic surfaces of mem­ branes to help regulate membrane activities. Cycling between

the membrane‐bound and free states is mediated in most cases by phosphorylation/dephosphorylation or by GTP/ GDP binding cycles. The fatty acid‐linked proteins are attached either to a myristic acid (C14), by way of an amide linkage to an amino terminal glycine, or to one or more ­palmitic acid (C16) residues, by way of thioester linkages to cysteines near the carboxyl terminus. Prenyl lipid‐anchored proteins are attached to one or more molecules of farnesyl (C15; 3 isoprene units) or geranylgeranyl (C20; 4 isoprene units), which are also coupled to cysteine residues in carboxyl‐ terminal CXXX, CXC, and XCC motifs (Fig. 1.10). In contrast to the fatty acid‐ and the prenyl group‐linked proteins, the phosphatidylinositol‐anchored proteins are

9

10

Part I  COMPARTMENTS

bound to the lumenal/extracellular surfaces of membranes (Fig. 1.10). Interestingly, these proteins are first produced as larger, integral proteins with one transmembrane domain. Enzymatic cleavage between the transmembrane domain and the globular surface domain produces a new C terminus on the globular domain, to which the lipid is coupled by ER‐ based enzymes (see Chapter 4, Section 4.6.4). The remaining transmembrane domain is then degraded by proteases. Many arabinogalactan proteins (AGPs) appear to be linked to the plasma membrane via a glycosylphosphatidylinositol (GPI) anchor. These molecules can be enzymatically released from the cell surface by phospholipase C.

1.2.5  The fluid‐mosaic membrane model predicts structural and dynamic properties of cell membranes Although the original fluid‐mosaic membrane model was developed at a time when membrane researchers knew only of peripheral and integral proteins, slight modifications to its basic premises have accommodated more recent discoveries, including lipid‐anchored proteins and membrane protein– cytoskeletal interactions. Membrane fluidity involves the movement not only of lipid molecules, but also of integral proteins that span the bilayer and of the different types of surface‐associated mem­ brane proteins. This ability of membrane proteins to diffuse laterally in the plane of the membrane is crucial to the func­ tioning of most membranes: Collisional interactions are essential for the transfer of substrate molecules between many membrane‐bound enzymes and of electrons between the electron transfer chain components of chloroplasts and mitochondria (see Chapters 12 and 14). Such movements are also  critical for the assembly of multiprotein membrane complexes. In addition, many signaling pathways depend on  transient interactions among defined sets of integral membrane proteins and peripheral or lipid‐anchored proteins. Tethering structures regulate and restrict the movement of  membrane proteins, often limiting their distribution to defined membrane domains. This tethering can involve ­connections to the cytoskeleton and the cell wall, bridges between  related integral proteins, or junction‐type interac­ tions between proteins in adjacent membranes. A particularly striking example of the latter type of interaction occurs in the grana stacks of chloroplast membranes (see Section 1.10.4). Grana stack formation has been shown to affect the lateral distribution of all major protein complexes in thylakoid membranes and to regulate the functional activity of the pho­ tosynthetic reaction centers and other components of the photosynthetic electron transport chain. Another mechanism for generating transient membrane microdomains of different composition involves membrane lipids organized in the form of lipid rafts. GPI‐anchored pro­ teins are typically associated with such membrane domains, which have been defined by cell biologists as membrane

domains that are resistant to certain types of detergents. Biochemical analyses of these detergent‐resistant membrane fractions have shown that they contain over 100 proteins and are enriched for phytosterols, and that the degree of fatty acid unsaturation affects their stability. However, due to their transient nature, there is no consensus on their in vivo size and composition. Indirect evidence suggests that lipid rafts participate in membrane sorting and signaling functions.

1.3  Plasma membrane The plasma membrane forms the outermost boundary of the living cell and functions as an active interface between the cell and its environment (Fig. 1.11). In this capacity it controls the transport of molecules into and out of the cell, transmits sig­ nals from the environment to the cell interior, participates in the synthesis and assembly of cell wall molecules, and pro­ vides physical links between elements of the cytoskeleton and the extracellular matrix. In conjunction with specialized domains of the ER, the plasma membrane produces plasmodesmata, membrane tubes that cross cell walls and pro­ vide direct channels of communication between adjacent cells (Fig.  1.12). As a result of these plasmodesmal connec­ tions, almost all the living cells of an individual plant share a physically continuous plasma membrane. This contrasts sharply with the situation in animals, where virtually every

MT

CW

PM

FIGURE 1.11  The plasma membrane (PM) of a turgid plant cell is pressed tightly against a cell wall (CW). These adjacent cryofixed plant cells have been processed by techniques that preserve the close physical relationship between plasma membrane and cell wall. Cells preserved with chemical fixatives for observation under an electron microscope often demonstrate artifacts of specimen preparation, such as a wavy conformation of the plasma membrane and a gap between the membrane and the cell wall. Microtubule (MT). Source: TEM by A. Lacey Samuels, University of British Columbia, Vancouver, Canada.

Chapter 1  Membrane Structure and Membranous Organelles

PM

CW

ER

FIGURE 1.12  Longitudinal section through a plasmodesma. Plasma membrane (PM), endoplasmic reticulum (ER), cell wall (CW). Source: TEM by Lewis Tilney, from Tilney et al. (1991). J Cell Biol 112: 739–747.

cell has a separate plasma membrane, and cell‐to‐cell com­ munication occurs instead through protein channels known as gap junctions. Yet another important difference between plants and ani­ mals is that plant cells are normally under turgor pressure, whereas animal cells are isoosmotic with their environments. Turgor pressure forces the plasma membrane tightly against the cell wall (see Fig. 1.11).

1.3.1  The lipid composition of plasma membranes is highly variable Plasma membranes of plant cells consist of lipids, proteins, and carbohydrates in a molecular ratio of ~40:40:20. The lipid mixture contains phospholipids, glycolipids, and sterols, the same classes found in animal plasma membranes. In plant plasma membranes, the ratio of lipid classes varies remarkably

among the different organs in a given plant and among iden­ tical organs in different plants—in contrast to the far more constant ratios in animal cells. Barley (Hordeum vulgare) root cell plasma membranes, for example, contain more than twice as many free sterol molecules as phospholipids (Table 1.3). In leaf tissues this ratio is generally reversed, but varies: In barley leaf plasma membranes, the phospholipid to free sterol ratio is 1.3:1, whereas in spinach (Spinacia oleracea) it is 9:1. This striking variability, which continues to puzzle researchers, indicates that ubiquitous plasma membrane enzymes can function in widely different lipid environments. These results have led to the suggestion that the lipid compo­ sition of plant plasma membranes may have little bearing on their functional properties and that the only important lipid parameter is membrane fluidity. If this were true, it would mean that virtually all lipid classes are interchangeable so long as a given combination of lipids yields a bilayer of desired fluidity at a particular temperature. This provocative idea may well be an overstatement, reflecting our ignorance about the functional roles of specific lipid types; moreover, it seems to be contradicted by the finding that the activity of proton‐ translocating ATPase (H+‐ATPase) molecules from corn (Zea mays) root reconstituted into artificial membranes can be modulated by changes in sterol composition. More research is needed to clarify how different lipid classes contribute to plasma membrane function. The most common free sterols of plant plasma mem­ branes are campesterol, sitosterol, and stigmasterol (see Fig. 1.4). Cholesterol, the principal free sterol of mammalian plasma membranes, is a minor component in the vast major­ ity of plant species analyzed to date, oat (Avena sativa) being a notable exception to this trend. Sterol esters, sterol glyco­ sides, and acylated sterol glycosides are more abundant in plants than in animals. Sterol glycosylation, a reaction cata­ lyzed by UDP‐glucose:sterol glycosyltransferase, has been exploited as a marker for isolated plant plasma membranes. Sphingomyelin, another major type of lipid formed in mam­ malian plasma membranes, has yet to be found in plants. Interesting differences in the fatty acid tails of plant and mammalian plasma membrane glycerolipids have also been reported. Whereas plants principally utilize palmitic (C16:0), linoleic (C18:2), and linolenic (C18:3) acids, mammals use ­palmitic (C16:0), stearic (C18:0), and arachidonic (C20:4) acids.

TABLE 1.3  Lipid composition of plasma membranes from various non‐cold‐acclimated species and tissues (mole %). Lipid type

Barley root

Barley leaf

Arabidopsis leaf

Spinach leaf

Phospholipids

26

44

47

64

Free sterols

57

35

38

7

Steryl glucosides

7

–

5

–

Acylated steryl glycosides

–

–

3

13

Glucocerebrosides

9

16

7

14

11

12

Part I  COMPARTMENTS

1.3.2  Cold acclimation leads to characteristic changes in plasma membrane lipid composition Low temperature is one of the most important factors limit­ ing the productivity and distribution of plants. All plants able to withstand freezing temperatures possess the ability to freeze‐proof their cells by a process known as cold acclimation (see Chapter 22). This metabolic process involves alter­ ing the composition and physical properties of membranes, cytoplasm, and cell walls so that they can withstand not only freezing temperatures but also freeze‐induced dehydration. One of the most cold‐hardy woody species is the mulberry tree (Morus bombycis Koidz). After cold acclimation in mid­ winter, these trees can withstand freezing below –40°C, but in midsummer, when they are not cold‐acclimated, they can be injured by a freeze below –3°C. Among the most pronounced and critical alterations that occur during cold acclimation are changes in lipid composition of plasma membranes. One might expect cold acclimation‐ induced lipid changes to vary among species, given the differ­ ences in plasma membrane lipid composition already noted (Table 1.3). However, in all cold‐hardy herbaceous and woody species studied to date, cold acclimation has been reported to cause an increase in the proportion of phospholipids and a decrease in the proportion of glucocerebrosides. In addition, the mole percent of phospholipids carrying two unsaturated tails increases. Species in which the cold‐acclimated plasma membranes contain the highest proportion of diunsaturated phospholipids and the lowest proportion of glucocerebrosides tend to be the most cold hardy.

In plants, transmembrane signaling receptors (see Chapter  18) are essential for cell communication and for mediating interactions with the environment. They also play important roles in development and in orchestrating diverse defense responses. Receptors capable of responding to many types of signaling molecules, including hormones, oligosaccharins, proteins, peptides, and toxins have been identified, but only a small number of these have been char­ acterized to date. Plasma membrane proteins participate in a variety of interactions with the cell wall, including formation of physi­ cal links to cell wall molecules, synthesis and assembly of cell wall polymers, and creation of a highly hydrated, tissue‐ specific interfacial domain. The presence of physical connec­ tions between the plasma membrane and the cell wall was first deduced from the presence of thread‐like strands con­ necting the protoplasts of plasmolyzed cells to the cell wall (Fig. 1.13). These strands are known as Hechtian strands in honor of Kurt Hecht, who is credited with their discovery in 1912. During cold acclimation, the number of Hechtian strands increases, suggesting that increasing the strength of the protoplast–cell wall interactions helps protect protoplasts from the stress of freeze‐induced dehydration. Electron microscopic analysis has shown that these strands are thin

P

1.3.3  Plasma membrane proteins serve a variety of functions Among the prominent classes of proteins present in the plasma membrane are transporters, signal receptors, and proteins that function in cell wall interactions and synthesis. Most plasma membrane proteins involved in these trans­ membrane activities are of the integral type. However, these proteins often form larger complexes with peripheral pro­ teins. The extracellular domains of many integral proteins are glycosylated, bearing N‐ and O‐linked oligosaccharides. The plasma membrane H+‐ATPase (P‐type H+‐ATPase) couples ATP hydrolysis to the transmembrane transport of protons from the cytosol to the extracellular space. This pro­ ton pumping has two effects. First, it acidifies cell walls and alkalinizes the cytosol, thereby affecting cell growth and expansion (see Chapter 2) as well as many other cellular activ­ ities. Second, it produces an electrochemical potential gradi­ ent across the plasma membrane that can drive the transport of ions and solutes against their respective concentration gradients (see Chapters 3 and 23). The plasma membrane also contains specialized water‐conducting channels known as aquaporins (see Chapter 3).

A Retracted protoplast Hechtian strands Cell wall

H2O

B

Plasma membrane

FIGURE 1.13  (A) Light micrograph of plasmolyzed onion epidermal cells. Hechtian strands (“arrowheads”) connect the protoplasts (P) to the cell walls (CW). (B) Diagram illustrating features of plasmolyzed plant cells. Source: TEM by Karl Oparka, from Oparka et al. (1994). Plant Cell Environ 17: 163–171.

Chapter 1  Membrane Structure and Membranous Organelles

tubes of cytoplasm delineated by a plasma membrane that retains tight contacts with the cell wall. These strands remain continuous with the plasma membrane. Although the mole­ cules that link the plasma membrane to the cell wall have not yet been identified, indirect studies suggest they may be integrin‐type receptors that recognize the amino acid sequence Arg‐Gly‐Asp (RGD) in cell wall constituents. A protein known as WAK1, a plasma membrane receptor with kinase activity, is another candidate protein. AGPs, another class of cell surface proteins, are highly gly­ cosylated proteoglycans that derive >90% of their mass from sugar. Classical‐type AGPs appear to be anchored to the exter­ nal surface of the plasma membrane by means of GPI lipid anchors (see Section  1.2.4), providing a carbohydrate‐rich interface between the cell wall and the plasma membrane. The fact that AGPs are expressed in a tissue‐ and developmental stage‐specific manner suggests they may play a role in differ­ entiation. Additional plasma membrane proteins, the cellu­ lose synthase and callose synthase complexes, extrude cellulose (ß‐1,4‐linked glucose) and callose (ß‐1,3‐linked glu­ cose), respectively, directly into the cell walls (see Chapter 2).

1.4  Endoplasmic reticulum The ER is the most extensive, versatile, and adaptable orga­ nelle in eukaryotic cells. It consists of a three‐dimensional (3D) network of continuous tubules and flattened sacs that underlie the plasma membrane, course through the cyto­ plasm, and connect to the nuclear envelope but remain dis­ tinct from the plasma membrane. In plants, the principal functions of ER include synthesizing, processing, and sorting proteins targeted to membranes, vacuoles, or the secretory pathway as well as adding N‐linked glycans to many of these proteins and synthesizing a diverse array of lipid molecules. The ER also provides anchoring sites for the actin filament bundles that drive cytoplasmic streaming, and plays a critical role in regulating the cytosolic concentrations of calcium (Ca2+), which influence many other cellular activities. The classical literature distinguishes three types of ER membranes: rough ER, smooth ER, and nuclear envelope. However, researchers now recognize many more morpho­ logically distinct subdomains that perform a variety of differ­ ent functions (Fig.  1.14). Despite this functional diversity, virtually all ER membranes are physically linked and enclose a single, continuous lumen that extends beyond the bounda­ ries of individual cells via the plasmodesmata.

1.4.1  The ER gives rise to the endomembrane system The endomembrane system includes membranous orga­ nelles that exchange membrane molecules, either by lateral diffusion through continuous membrane or by transport vesicles that bud from one type of membrane and fuse ­

with  another (Fig.  1.15). The principal membrane systems connected in this manner include the nuclear envelope, membranes of the secretory pathway (ER, Golgi, trans‐Golgi network, multivesicular body, plasma membrane, vacuole, and different types of transport/secretory vesicles), and membranes associated with the endocytic pathway (plasma membrane, clathrin‐coated endocytic vesicles, trans‐Golgi network/early endosome/recycling endosome, multivesicular body/late endosome, vacuole, and transport vesicles). Extensive traffic between these compartments not only transports secreted molecules to the cell surface and vacuolar proteins to the vacu­ oles, but also distributes membrane proteins and membrane lipids from their sites of synthesis, the ER and Golgi cisternae, to their sites of action, all of the endomembrane organelles. A plethora of sorting, targeting, and retrieval systems regulate traffic between the different compartments, ensuring delivery of molecules to the correct membranes and the maintenance of organelle identity (see Chapter 4). All membranes of the endomembrane system are con­ nected by both anterograde (forward) and retrograde (backward) traffic (Fig.  1.15). The anterograde pathway ­usually delivers newly synthesized molecules to their desti­ nation. In the retrograde pathway, membrane molecules dispersed by transport processes are recycled to their sites of origin, and “escaped” molecules are returned to their normal site of action. Because the volume of membrane traffic is large and the accuracy of sorting is 10,000. Each Golgi stack‐TGN unit consists of a set of five to eight flattened Golgi cisternae that exhibit a distinct morphologi­ cal polarity and possess fenestrated and bulbous margins, and a Golgi‐associated TGN cisterna on the trans‐side of the stack (see Figs. 1.25, 1.26, and 1.28). Three types of Golgi cisternae, cis, medial, and trans, can be distinguished based on their position in a stack, their staining properties, the types of vesi­ cles that bud from their rims, and their biosynthetic func­ tions. According to the cisternal progression model of Golgi trafficking, COPII vesicles from the ER are assembled into new cisternae on the cis‐side of the stack (Figs. 1.25 and 1.28), and the trans‐most cisternae are converted to Golgi‐­ associated TGN cisternae before they are shed to become free TGN cisternae. This process leads to a net displacement of cargo‐ laden cisternae across the stack as new cisternae are added

Chapter 1  Membrane Structure and Membranous Organelles

G

TGN GS

FIGURE 1.29  Tomographic slice image of an Arabidopsis meristem Golgi stack (G) and TGN cisternae and their encompassing, ribosome‐excluding scaffold/matrix (GS) systems. Source: Micrograph by Byung‐Ho Kang from Staehelin, L.A. and Kang, B.-H. (2008). Plant Physiol 147: 1454–1468.

ER

GS

FIGURE 1.30  3D tomographic model of a dividing Golgi stack docked to an ER export site in an Arabidopsis meristem cell. The docking is mediated by structural links (arrowheads) between the cis‐side of the Golgi scaffold/matrix (GS) and the scaffolds on the budding COPII vesicles. See diagrams Figures 1.25A and 1.31. Source: Model by Byung‐Ho Kang, from Staehelin, L.A. and Kang, B.-H. (2008). Plant Physiol 147: 1454–1468.

and old cisternae are shed. During their assembly the cis‐ Golgi cisternae remain biochemically inactive. Most of the biosynthetic activities are confined to the medial and trans‐ Golgi cisternae and to a lesser extent to the Golgi‐associated TGN compartments. To maintain the characteristic enzymatic properties of the different cisternal types during translocation of cisternae across the stack, the cisternae bud recycling vesicles that

transport membrane proteins in a retrograde direction. These vesicles are known as COPI vesicles (see Fig.  1.25A). In plants, two types of COPI vesicles have been characterized: COPIa vesicles that recycle escaped ER proteins from cis‐ Golgi cisternae back to the ER, and COPIb vesicles that bud from medial‐ and trans‐Golgi and Golgi‐associated TGN ­cisternae and transport cisternal membrane proteins in a ­retrograde direction. This latter transport is responsible for maintaining the steady‐state distribution of enzymes within Golgi stacks. Intercisternal elements are another type of trans‐Golgi cisterna–associated structure (see Fig. 1.25B). These parallel protein fibers lie sandwiched between cisternae and may serve as anchors for the glycosyltransferases involved in syn­ thesis of large polysaccharide slime molecules, such as those secreted by the outermost cells of root caps.

1.5.2  The trans‐most Golgi cisternae give rise to TGN cisternae that sort and package Golgi products The function of the TGN is to sort and to package the prod­ ucts of the Golgi stacks into secretory vesicles and clathrin‐ coated vesicles. Secretory vesicles transport membrane and cargo molecules to the plasma membrane and cell wall, whereas clathrin‐coated vesicles produced by the TGN deliver membrane and soluble molecules to multivesicular bodies and vacuoles. The TGN cisternae arise from the trans‐most Golgi cisternae by a cisternal peeling process that yields free, independent TGN compartments (see Fig. 1.25). TGN cister­ nae in the process of peeling are called Golgi‐associated or early TGN cisternae, and those that have become physically separated from the originating Golgi stack, free or late TGN cisternae. During cisternal peeling, the cisternae undergo multiple changes in architecture, including a 30–35% decrease in cisternal membrane surface area, most likely due to the removal of membrane by COPIb‐type recycling vesicles. Concomitantly, the flattened cisternae are converted into grape‐like membrane compartments as the number of bud­ ding secretory and clathrin‐coated vesicles increases. This maturation process continues until all products have been sorted and packaged. Once this has happened, the fully matured free TGN cisterna breaks up, yielding free secretory and clathrin‐coated vesicles together with small residual frag­ ments of the cisternal membranes (see Fig. 1.25A). The fate of the residual membrane fragments is unknown. The ratio of secretory to clathrin‐coated vesicles budding from free TGN cisternae is highly variable, but typically secretory vesicles are more numerous. Even in a single root apical meristem cell, the ratio can vary from 5:1 to 1:3 on free TGN cisternae derived from adjacent Golgi stacks. This ­variability most likely reflects the composition of mRNAs (coding for secretory versus vacuolar proteins) that were translated by ER‐bound polysomes close to a given ER export site. Upon transfer of each batch of locally produced

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Part I  COMPARTMENTS

proteins to a forming cis‐Golgi cisterna of an individual Golgi stack, and after passing through the stack, the biased set of vacuolar or secretory proteins reaches the TGN. There, the ratio of secretory to clathrin‐coated vesicles formed is adjusted to the ratio of products in the cisternal lumen. This variability reflects the inherent flexibility of the sorting and packaging systems of the TGN cisternae. Free TGN com­ partments with a high ratio of clathrin‐coated vesicles were given the name “partially coated reticulum” by early electron microscopists, but in light of the new findings, this name has outlived its usefulness and should be abandoned. Trans‐ Golgi and TGN cisternae are the most acidic compartments of the Golgi apparatus. They are acidified by vacuolar (V‐ type) H+‐ATPases (see Chapter 3). The low pH environment of the cisternal lumen appears to regulate enzyme activities and cause an osmotic collapse of the lumen, which is of criti­ cal importance for the sorting and packaging of the Golgi products.

1.5.3  The Golgi scaffold/matrix originates on COPII buds and mediates ER‐to‐Golgi COPII vesicle transport The Golgi matrix is a fine, filamentous, cage‐like structure that completely surrounds all Golgi and TGN cisternae and is comprised of long coiled‐coil scaffolding proteins, hence the  more recent name Golgi scaffold. This scaffold/matrix excludes ribosomes from the immediate vicinity of Golgi and TGN cisternae, thereby facilitating its visualization in elec­ tron micrographs of ribosome‐rich cell types (Fig.  1.29). It also appears to prevent COPI‐type intercisternal recycling vesicles from escaping from the immediate surroundings of the Golgi stack and helps maintain structural integrity of Golgi stacks and associated TGN cisternae as they move along actin filaments through the cytoplasm. Analysis of Golgi stack dynamics has shown that stacks move in a stop‐and‐go manner. This has led to the hypothesis that stacks may pause at ER export sites to pick up cargo for processing by Golgi enzymes. The mechanism responsible for inducing Golgi stacks to pause at ER export sites has puzzled cell biologists for many years. It now appears that the Golgi scaffold/matrix is intimately engaged in this process (see Figs. 1.25 and 1.30). In particular, recent studies have shown that COPII vesicles originate with a ~40‐nm‐wide external layer of long, scaffold‐type proteins, termed COPII scaffold. As illustrated in the “dock, pluck and go model” of ER‐to‐ Golgi trafficking (Fig.  1.31), the scaffold layer of budding COPII vesicles appears to capture passing Golgi stacks by binding to the cis‐side of the Golgi scaffold/matrix. Once bound to a COPII vesicle, the wiggling motion of the Golgi stack appears to provide the force needed to pluck the ­budding vesicle from the ER export site. The plucked COPII vesicle together with its scaffold layer is then transferred to the cis‐ side of the Golgi scaffold/matrix where it contributes to the assembly of a new cis‐Golgi cisterna.

1.5.4  The sugar‐containing molecules produced in Golgi cisternae serve diverse functions The Golgi apparatus is involved in assembling the N‐linked and O‐linked glycans of glycoproteins and proteoglycans and in synthesizing complex polysaccharides (see Chapters 2 and 4). One of the principal functions of glycosylation is to protect proteins against proteolysis, thereby increasing their life span. Sugar groups increase protein solubility and may specify plasma membrane–cell wall interactions, prevent premature activation of lectins (highly specific sugar‐binding proteins), and contribute to protein folding or assembly of multiprotein complexes. Most proteins subject to N‐linked glycosylation perform enzymatic functions, whereas O‐glycosylated proteins often serve structural roles. For example, the O‐glycosylation of extensin molecules (see Chapter 2) is responsible for their rod‐shaped architecture. Both N‐ and O‐linked glycan side chains are present in many highly glycosylated AGPs that, at >90% sugars, are classified as proteoglycans. The complex cell wall polysaccharides produced by the Golgi apparatus perform structural functions and can bind water and heavy metals. In addition, these polysaccharides contain cryptic regulatory oli­ gosaccharide domains that can be released by specific enzymes to yield regulatory molecules known as oligosaccharins.

1.5.5  The Golgi apparatus is a carbohydrate factory The synthesis of N‐linked glycans starts in the ER with assem­ bly of a 14‐sugar oligosaccharide on a molecule of dolichol, a large lipid composed of 14–24 isoprene units. Once complete, this oligosaccharide is transferred by oligosaccharyl trans­ ferase to selected asparagine residues in nascent polypeptide chains. Most of the subsequent processing of the oligosaccha­ ride occurs in the Golgi, including systematic removal of mannose residues and the addition of other types of sugars (see Chapter 4). The enzymes involved in these reactions are not randomly distributed among Golgi cisternae but become localized to medial, or trans cisternae, depending on which sequential step they catalyze as a given N‐linked glycan moves through the stack from cis to trans. For example, the native enzyme mannosidase I, which mediates the first step of N‐glycan processing in the Golgi is localized in medial Golgi cisternae. However, when this enzyme is expressed in trans­ genic plant cells as a mannosidase I‐GFP fusion protein, the fusion protein is also seen in cis‐Golgi cisternae and even in the ER. Apparently, when too many Golgi membrane enzymes of a given type are produced, they pile up in upstream com­ partments of the secretory pathway. For this reason, localiz­ ing Golgi membrane proteins by means of GFP fusion proteins yields unreliable results. O‐linked glycans are important components of hydroxyproline‐ rich glycoproteins and AGPs, many of which serve structural

Chapter 1  Membrane Structure and Membranous Organelles

Polyribosome

ER export site

ER

Golgi stack

Golgi scaffold/ matrix

COP II bud

COP II scaffold

Actin filament

Myosin

A Go-phase (saltatory/directional movements) ER export site ER COP II vesicle

cis

Golgi scaffold/matrix

trans

On/off

B DOCK-and-PLUCK-phase (wiggling and slow directional movements) FIGURE 1.31  Dock, pluck, and go model of ER‐to‐Golgi vesicle trafficking. (A) Go-phase: Golgi stacks travel along actin filaments propelled by myosin motors. (B) Dock‐and‐pluck phase: The scaffold of a budding COPII vesicle attaches to the cis‐side of the Golgi scaffold/matrix of a passing Golgi stack and pulls the Golgi off the actin track. Once the COPII scaffold binds to the Golgi matrix, the wiggling motion of the Golgi stack facilitates the release of the COPII vesicle from the ER by a “plucking” mechanism. Once it has harvested the COPII vesicle from the ER export site, the Golgi is free to resume its movement along the actin track. Source: Adapted from Staehelin, L.A. and Kang, B.-H. (2008). Plant Physiol. 147: 1454–1468.

roles. The sugars, mostly arabinose and galactose, are attached to amino acids that contain hydroxyl groups, such as hydroxy­ proline, serine, and threonine. Very little is known about the synthesis of O‐linked glycans. On newly synthesized proteins destined for O‐glycosylation, selected proline residues are con­ verted to hydroxyproline in the ER, but where the enzymes are located that add the first arabinose sugars to those hydroxypro­ lines remains to be determined. The matrix polysaccharides of plant cell walls are structur­ ally complex molecules that play a central role in determining cell size and shape (see Chapter 2). Defined fragments of such molecules also function as signaling molecules in pathways that control plant growth, organogenesis, and the elicitation of defense responses. Unlike the linear polymers cellulose and callose, all branched cell‐wall polysaccharides are synthesized by enzymes in Golgi and Golgi‐associated TGN cisternae. The molecular details of the assembly pathways have yet to be elu­ cidated. However, an outline of the spatial organization of the xyloglucan and pectic polysaccharide pathways in the  Golgi stacks has been developed by using immunolocalization to

track in which cisternae specific carbohydrate groups are added. Probably the most striking result of these studies is that assembly of pectic polysaccharides appears to involve enzymes localized in medial and trans‐Golgi and possibly Golgi‐associated TGN cisternae, whereas enzymes that pro­ duce xyloglucan appear to be confined to trans‐Golgi and Golgi‐associated TGN cisternae. To date, very few native enzymes and nucleotide sugar transporters involved in com­ plex polysaccharide biosynthesis have been immunolocalized to specific types of Golgi cisternae.

1.6  Exocytosis and endocytosis Exocytosis is the process by which secretory vesicles derived from the TGN fuse with the plasma membrane, releasing their contents into the extracellular space (Fig.  1.32A). In growing cells, this process delivers proteins and lipids needed

23

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Part I  COMPARTMENTS

PM

PM

A

B

FIGURE 1.32  Cross‐sectional images of secretory vesicles. (A) TEM of a secretory vesicle in the process of initiating membrane fusion with the plasma membrane (PM). (B) TEM of a secretory vesicle that has fused with the plasma membrane and discharged its contents. The disc‐shaped infolding of the membrane shown here is observed only in turgid cells. Source: (A) Micrograph by L. Andrew Staehelin, from Staehelin et al. (1990). Protoplasma 157: 75–91. (B) Micrograph by Yoshinobu Mineyuki, University of Hyogo, Himeji, Japan.

CCV

CP

A

B

FIGURE 1.33  Tomographic slice images of clathrin‐coated vesicles. (A) A clathrin‐coated vesicle (CCV) in the process of budding from a late‐ stage cell plate (CP) membrane. The arrow points to a dynamin motor protein ring that mechanically separates the vesicle from the membrane. (B) Higher magnification view of a cross‐sectioned clathrin‐coated vesicle. Source: (A) Micrograph by Jose‐Maria Segui‐Simarro, Universidad Politécnica de Valencia, Spain. (B) Micrograph by Mathias Gerl, University of Colorado, Boulder.

for expansion of the plasma membrane as well as complex polysaccharides, glycoproteins, and proteoglycans required for cell wall growth. Because of the large surface‐to‐volume ratio of secretory vesicles, exocytosis delivers more mem­ brane to the cell surface than is needed for expansion of the plasma membrane. Excess plasma membrane molecules are returned to the cytoplasm by two mechanisms: (1) endocytosis mediated by clathrin‐coated vesicles (Fig.  1.33) that contain membrane proteins and lipids, and (2) non‐vesicular, intermembrane lipid transport at plasma membrane–ER interaction sites associated with fused vesicle membrane domains (see domain 15 in Fig.  1.14). Endocytosis is also used to turn over plasma membrane molecules and remove activated receptors from the cell surface. A recycling pathway allows endocytosed, but still functional plasma membrane proteins to be recycled back to the plasma membrane. In animal cells, endocytosis plays a major role in the uptake of nutrient molecules, but little evidence suggests such a role in plants.

1.6.1  In plants, turgor pressure affects membrane events associated with exocytosis, endocytosis and membrane recycling When a secretory vesicle in an animal cell fuses with the plasma membrane, its contents are expelled to the extracel­ lular space and the vesicle membrane becomes part of the plasma membrane. As this occurs, the plasma membrane is expanded slightly, an expansion that can be readily accom­ modated by changes in surface architecture of the animal cell. Turgor pressure prevents this from happening in plant cells. In turgid plant cells, the plasma membrane is pressed tightly against the cell wall (see Fig. 1.11) and cannot expand unless the cell wall expands as well. When a spherical secretory vesi­ cle fuses with the plasma membrane of a turgid cell, turgor pressure not only forces the vesicle contents into the cell wall, but also flattens the vesicle into a disc‐shaped infolding of the

Chapter 1  Membrane Structure and Membranous Organelles

Cell wall

Plasma membrane PM

1

1

2

3

CCV ER

Lipid transfer protein Golgi/TGN

FIGURE 1.34  Three mechanisms for removing excess membrane from a plasma membrane infolding generated by the fusion of a secretory vesicle with the plasma membrane (see Fig. 1.32B) of a turgid plant cell: 1. cell expansion; 2. formation of clathrin‐coated endocytic vesicles (CCV); 3. direct transfer of lipid molecules from the plasma membrane to adjacent ER membrane via lipid transfer proteins. Source: Adapted from Staehelin, L.A. and Chapman, R.L. (1987). Planta 171: 43–57.

plasma membrane (Fig.  1.32B). Because the plasma mem­ brane cannot expand, the infolding remains in place until excess membrane is removed. How then is excess membrane removed from plant plasma membranes? Plants, like animal cells, can recycle excess membrane from the plasma membrane via endocytosis of clathrin‐coated vesicles (Fig. 1.34 and Section 1.6.2). However, endocytosis demands much more energy in plants than in animal cells because sizable hydrostatic pressure forces must be overcome to form plasma membrane invaginations. In contrast, retrieval of lipid molecules from the plasma mem­ brane via non‐vesicular, intermembrane lipid transport— lipid “hopping”—mediated by lipid transfer proteins (Fig. 1.34) circumvents this energy problem, and reduces the number of endocytotic vesicles needed for the removal of excess plasma membrane material. Evidence for lipid hopping has come from both lipid uptake research and electron microscopic studies of ultra­ rapidly frozen cells. To investigate lipid uptake, a fluorescent analog of the membrane lipid phosphatidylcholine was added to the outer surface of the plasma membrane. This molecular marker was quickly translocated to peripheral ER  cisternae with no evidence of vesicular intermediates. However, this uptake required that an extracellular enzyme convert phosphatidylcholine to diacylglycerol, a membrane lipid that lacks a large, polar headgroup and can readily flip‐ flop across the plasma membrane before being transferred to the ER. Electron micrographs of cryofixed cells suggest that this rapid lipid transfer is mediated by unique ER membrane extensions that form tight caps over the plasma membrane appendages left behind by the fused secretory vesicles (Figs. 1.34 and 1.35; see also domain 15 in Fig. 1.14). During this process the disc‐shaped appendages tip over to form characteristic horseshoe‐shaped plasma membrane invagi­ nations. Once capped by an ER extension, the invagination

ER

FIGURE 1.35  TEM showing lipid‐recycling ER domains. The ER cisterna has characteristic extensions that form specialized contact sites (arrows) with freshly fused and collapsed secretory vesicles in the plasma membrane (PM) of a cryofixed pea root tip cell. These structures correspond to domain 15 in Figure 1.14. Source: Micrograph by Stuart Craig, from Craig, S. and Staehelin, L.A. (1988). Eur J Cell Biol 46: 81–93.

shrinks until excess membrane is gone. The ER then retracts from the plasma membrane.

1.6.2  The membrane compartments associated with endocytosis can be identified by following the uptake of tracer molecules The process of endocytosis can be visualized by exposing cells to tracer molecules that are internalized by clathrin‐coated endocytic vesicles. Two classes of molecular markers have been used in such investigations. Compounds in one group serve as membrane markers and bind to the plasma mem­ brane (e.g., cationized ferritin); members of the other class, known as fluid‐phase markers, are internalized with the aqueous phase (e.g., lanthanum nitrate). The widely used fluorescent endocytosis marker FM4‐64 is generally portrayed as a membrane marker, but based on published micrographs it also appears to bind to cell‐wall components. Compared to the vast literature on endocytosis in animal systems, the num­ ber of comparable plant studies remains remarkably small. Two factors have contributed to the slow development of the field: (1) the low level of endocytic activity of plant cells, and (2) the presence of cell walls that greatly impede access of tracer molecules to the plasma membranes, making experi­ mentation more difficult. Protoplasts can be used for tracer uptake studies, but the concentrated sucrose medium that protects protoplasts from bursting also affects cell function, yielding results that may not be representative of what occurs in healthy turgid cells. A schematic model of our current understanding of the endocytic pathway in plants is depicted in Figure 1.36. Much

25

26

Part I  COMPARTMENTS FIGURE 1.36  Schematic diagram of the endocytic pathway of plant cells. Plasma membrane material internalized via clathrin‐coated vesicles is delivered to compartments called early endosomes, most of which appear to correspond to free TGN cisternae (see Fig. 1.25A) where the different types of proteins are sorted and packaged into either secretory vesicles (SV) that recycle materials back to the plasma membrane or clathrin‐coated vesicles (CCV) that deliver molecules to multivesicular bodies (MVBs), which serve as late endosomes. Some MVBs deliver molecules to lytic vacuoles, whereas others send materials to protein storage vacuoles.

Endocytosis

Plasma membrane

Exocytosis ?

CCV

Non-clathrin vesicle ?

Lytic

SV Protein storage CCV Golgi

Golgiassociated early TGN

of the recent progress in plant endocytosis research has resulted from following the uptake of fluorescent tracer ­molecules into cells. The first compartments to receive endo­ cytosed molecules are known as early endosomes and/or recycling endosomes, and those that receive the molecules later as late endosomes. In plants, the early endosomes ­correspond to TGN cisternae, and the late endosomes to multivesicular bodies (MVBs). As discussed above, TGN cisternae serve as sorting and packaging compartments in the secretory pathway. The endocytosis pathway makes use of these capabilities to sort recyclable molecules from the plasma membrane into secretory vesicles so that they can be returned to the plasma membrane, and molecules destined for destruc­ tion into clathrin‐coated vesicles for transport to MVBs and  ultimately to lytic vacuoles. MVBs are spherical mem­ brane compartments that contain distinct internal vesicles (Fig.  1.37A). Two types of vesicles bud from the boundary membrane of MVBs. Those that bud towards the cytoplasm and recycle membrane proteins back to the TGN are known as retromer vesicles. In contrast, budding of the boundary membrane into the MVB lumen gives rise to the internal vesicles. This latter process is employed to sequester mem­ brane proteins—most notably activated, internalized plasma membrane receptors—into the interior of MVBs so that they can be functionally silenced and subsequently degraded by lytic vacuolar enzymes. Delivery of the vesicular contents of MVBs to storage or lytic vacuoles occurs by fusion of the MVBs with the vacuolar membranes (Fig. 1.37B). Two types of structurally similar MVBs can be distin­ guished based on their cellular location and functional attrib­ utes. One type is seen in close proximity to the TGN, with both types of membrane compartments encompassed by a continuous TGN/MVB scaffold system. In developing Arabidopis embryo cells the proteolytic processing of seed storage proteins starts in these TGN‐associated MVBs, which

Free TGN/ early endosome/ recycling endosome

MVB/late endosome

Vacuoles

A

MVB

V

B FIGURE 1.37  (A) TEM of one large and one small multivesicular body in the cytoplasm of a tobacco cell cultured in suspension. (B) Multivesicular body (MVB) that has just fused with a vacuole (V) and is in the process of delivering its internal vesicles to the vacuolar lumen. Source: (A) Micrograph by L. Andrew Staehelin. (B) Micrograph by Byung‐Ho Kang, University of Florida, Gainesville, FL.

Chapter 1  Membrane Structure and Membranous Organelles

deliver the processed storage proteins to the protein storage vacuoles. MVBs that contain cargo molecules destined for vacuoles have also been called prevacuolar compartments, but because the definition of MVBs is more precise, the pre­ vacuolar compartment nomenclature is now being phased out. MVBs not contained within the TGN/MVB scaffold sys­ tem are capable of moving around cells in an independent manner and do not appear to process storage proteins. These independent MVBs, which receive endocytosed molecules, are also surrounded by a scaffold system, and deliver their contents to lytic vacuoles.

1.7  Vacuoles Vacuoles, fluid‐filled compartments encompassed by a tonoplast membrane, are conspicuous organelles of most plant cells: They typically occupy ~30% of the cell volume (Fig. 1.38), but in some cells, the space occupied by the vacuolar compartment(s) can approach 90%, with most of the cyto­ plasm confined to a thin peripheral layer connected to the nuclear region by transvacuolar strands of cytoplasm. During the cell cycle (see Chapter  11), vacuoles of apical meristem cells undergo major changes in shape and size. During inter­ phase (G1 phase of the cell cycle), numerous small vacuoles are dispersed throughout the cytoplasm (see Fig. 1.1B). These vacuoles fuse into larger units during S and G2 phases and prometaphase. Coinciding with formation of the phragmo­ plast during cell plate formation, vacuolar volume shrinks by up to 80% and vacuoles assume a tubular configuration. This

C

shrunken state persists until cytokinesis is complete and the two daughter cells enter the G1 phase. It has been postulated that the transient reduction in vacuole volume during early telophase provides a means for increasing the volume of the cytosol to accommodate the forming phragmoplast micro­ tubule arrays and associated cell plate‐forming structures.

1.7.1  Plants use vacuoles to produce large cells cheaply One of the major challenges faced by plants during evolution was to produce large solar collectors at a metabolic cost that could be recovered by the energy trapped and utilized by chloroplasts in a growing season. This problem was solved by increasing the volume of the vacuolar compartment to drive cell enlargement while keeping the amount of nitrogen‐rich cytoplasm constant. This latter point is particularly important for plants, whose growth is often limited by N availability. By filling a large volume of the cell with “inexpensive” vacuolar contents, mostly water and minerals, plants are able to drasti­ cally reduce the cost of making expanded structures such as leaves, which are essentially throwaway solar collectors. Plant cell expansion is driven by a combination of osmotic uptake of water into the vacuoles and altered cell wall extensi­ bility. The water taken into vacuoles generates turgor pressure which expands the primary cell wall and creates stiff, load‐­ bearing structures in conjunction with the walls. This exploita­ tion of internal hydrostatic pressure to stiffen thin primary cell walls resembles the use of air pressure in an inner tube to ­convert a pliable, flat bicycle tire into a stiff circle capable of supporting heavy loads. Wilting and the associated softening of plant organs are caused by the loss of water from the vacuoles. To maintain turgor pressure of continuously expanding cells, solutes must be actively transported into the growing vacuole to maintain its osmolarity. An electrochemical poten­ tial gradient across the tonoplast membrane provides the driving force for uptake of solutes. The gradient, in turn, is produced and maintained by two electrogenic proton pumps: V‐type H+‐ATPase and vacuolar H+‐pyrophosphatase (H+‐ PPase). The principal solutes in vacuoles include the ions K+, Na+, Ca2+, Mg2+, Cl‐, SO42‐, PO43‐, and NO3‐, and primary metabolites such as amino acids, organic acids, and sugars. Movement of water across the tonoplast membrane is medi­ ated by aquaporin channels (see Chapter 3).

V

1.7.2  Plant vacuoles are multifunctional compartments FIGURE 1.38  TEM showing a cross‐sectional view of spongy mesophyll cells in a bean leaf illustrating the large amount of cell volume occupied by the central vacuoles (V). Chloroplasts (C). Source: Micrograph by L. Andrew Staehelin.

Vacuoles play several metabolic roles in addition to promot­ ing cell expansion. Storage: In addition to the solutes and primary metabolites mentioned above, plants also store large amounts of pro­ teins in their vacuoles, especially in seeds. Stored reserves

27

28

Part I  COMPARTMENTS

can be retrieved from vacuoles and used in metabolic path­ ways to sustain growth. Interestingly, most of the flavors of fruits and vegetables can be traced to compounds that are stored in vacuoles. Digestion: Vacuoles have been shown to contain the same types of acid hydrolases found in animal cell lysosomes. These enzymes, which include proteases, nucleases, gly­ cosidases, and lipases, together allow for the breakdown and recycling of nearly all cellular components. Such recy­ cling is needed not only for the normal turnover of cellular structures, but also for the retrieval of valuable nutrients during programmed cell death associated with develop­ ment and senescence (see Chapter 20). pH and ionic homeostasis: Large vacuoles serve as reservoirs of protons and metabolically important ions, such as Ca2+. Typically, plant vacuoles have a pH between 5.0 and 5.5, but the range extends from ~2.5 in vacuoles of lemon (Citrus limon) fruit to >7.0 in unactivated protein storage vacuoles. By controlling the release of protons and other ions into the cytosol, cells can regulate not only cytosolic pH but also the activity of enzymes, the assembly of cytoskeletal structures, and membrane fusion events. Defense against microbial pathogens and herbivores: Plant cells accumulate an amazing variety of toxic com­ pounds in their vacuoles, both to reduce feeding by herbi­ vores and to destroy microbial pathogens. These compounds include: ●● phenolic compounds, alkaloids, cyanogenic glycosides, and protease inhibitors to discourage insect and animal herbivores ●● cell wall‐degrading enzymes such as chitinase and ­glucanase, and defense molecules such as saponins to destroy pathogenic fungi and bacteria ●● latexes, wound‐clogging emulsions of hydrophobic ­polymers that possess insecticidal and fungicidal prop­ erties and also serve as antiherbivory agents Sequestration of toxic compounds: Plants cannot escape from toxic sites. Nor can they efficiently excrete toxic elements (e.g., heavy metals) and metabolites (e.g., oxa­ late). Instead, plants sequester these compounds into vacuoles. For example, to remove oxalate, specific cells develop vacuoles containing an organic matrix within which oxalate is allowed to react with Ca2+ to form cal­ cium oxalate crystals. In other plant cell types, members of the ABC family of transporters (see Chapter  3) are used to transport xenobiotics (foreign compounds) from the cytoplasm into vacuoles. Accumulation of toxic compounds in leaf vacuoles is one reason leaves are shed on a regular basis. Pigmentation: Vacuoles that contain anthocyanin pig­ ments are found in many types of plant cells. Pigmented flower petals and fruits are used to attract pollinators and seed dispersers, respectively. In leaves, some vacuolar pigments screen out UV and visible light, helping pre­ vent photo‐oxidative damage to the photosynthetic apparatus.

1.7.3  Plants generate different types of vacuoles by means of vacuole transformation pathways A fundamental question in plant vacuole research is whether plant cells produce multiple populations of functionally differ­ ent types of vacuoles (multiple vacuole hypothesis), or if they produce one basic type of vacuole system that becomes spe­ cialized in response to developmental and/or physiological signals. Until recently, most researchers subscribed to the mul­ tiple vacuole hypothesis, which postulates that the large central vacuoles in root tip cells arise from the fusion of protein stor­ age vacuoles and pre‐existing lytic vacuoles. However, several recent investigations have yielded data that contradict this hypothesis. In particular, these new studies have demonstrated that the root tip cells of freshly germinated seedlings contain only protein storage vacuoles, which provide nutrients to the growing cells. As these nutrients are released from the vacu­ oles, the protein storage‐type vacuoles are gradually converted into lytic vacuoles by means of cell type‐specific transforma­ tion pathways. The transformation process includes changes in vacuole architecture as well as changes in composition of the vacuole membrane. Most notably, the protein storage vacuole marker protein α‐TIP, an aquaporin‐type protein, is replaced by the lytic vacuole aquaporin homolog, γ‐TIP. Formation and maintenance of the different vacuole types also involves differ­ ent types of vacuole targeting signals (see Chapter 4). Senescing leaves of Arabidopsis and soybean (Glycine max) have been shown to produce two types of acidic vacuole sys­ tems in mesophyll and guard cells, a large central vacuole sys­ tem with lytic properties, and smaller senescence‐associated vacuoles. These latter vacuoles are more acidic than the ­central vacuole, accumulate a senescence‐specific cysteine protease, and lack the γ‐TIP of the central vacuole. Yet to be determined is whether these vacuoles arise de novo, or if they originate by budding from the central vacuole. Autophagic vacuoles, also known as autophagosomes, are transient, large, double membrane vesicles responsible for delivering cytoplasmic materials to lytic vacuoles/lysosomes for degradation. The process begins with the entrapment of a targeted region of cytoplasm, including whole organelles, by cisterna‐like membranes derived from collapsed vacuolar membranes and possibly ER membranes, followed by fusion with a lytic vacuole or lysosome. Autophagic vacuole forma­ tion is used to recycle old or damaged organelles, to provide nutrients during starvation, to generate large central vacuoles in root tip cells during germination, and to mobilize the cyto­ plasm during programmed cell death (Fig. 1.39).

1.8  The nucleus The nucleus contains most of the cell’s genetic information and serves as the center of regulatory activity (Fig. 1.40). Although the DNA–protein complexes that make up chromosomes are

Chapter 1  Membrane Structure and Membranous Organelles

V1 V2 NE NU

FIGURE 1.39  Light micrograph of an aleurone protoplast stained with a fluorescent dye. Two types of vacuoles are depicted: large protein storage vacuoles (V1) and smaller lytic/autophagic‐type vacuoles (V2) that may be involved in autophagy‐associated programmed cell death. Source: Micrograph by Paul C. Bethke, University of Wisconsin, Madison, WI.

evident only as an irregular network of chromatin during interphase, individual chromosomes nevertheless occupy dis­ crete domains within the nucleus throughout this part of the cell cycle. Interphase is the most important stage of the cell cycle for gene expression because during this period chromo­ somes are actively transcribed. A typical interphase plant cell nucleus also contains one to several nucleoli (singular: nucleolus) lying free in the nuclear matrix, or nucleoplasm. These prominent, densely staining, often spherical bodies house the cytoplasmic ribosome man­ ufacturing machinery. A brief introduction is given below to the ultrastructure and activities of the nucleolus as well as to  the structure of nuclear pores, through which nucleolar products must move to reach the cytoplasm.

N

FIGURE 1.40  TEM showing the nucleus (N) of a bean root tip cell. Note the two membranes of the nuclear envelope (NE) and the large central nucleolus (NU). Source: Micrograph by Eldon Newcomb.

NE

NP

1.8.1  The nuclear envelope is a dynamic structure with many functions The nuclear envelope forms the outer boundary of the nucleus. It consists of two concentric membranes, the inner nuclear membrane and the outer nuclear membrane, separated by the perinuclear space (Fig. 1.41). The two principal functions of the nuclear envelope are to separate the genetic material in the nuclear compartment from the enzyme systems in the cytoplasm, and to regulate the exchange of molecules that traf­ fic between the two compartments via the nuclear pores (Figs. 1.41 and 1.42). The inner and outer envelope membranes

FIGURE 1.41  TEM of a freeze‐fractured nuclear envelope (NE) with nuclear pores (NP). The continuity of the inner and outer membranes becomes apparent where the membranes are seen in cross section. Source: Micrograph by L. Andrew Staehelin.

are connected to each other via the bridging bilayer regions of the pores. The outer membrane of the nuclear envelope is con­ tinuous with membranes of the ER through narrow (~20 nm in diameter) connections (see domain 3 in Fig.  1.14) and

29

30

Part I  COMPARTMENTS

Cytosol Outer nuclear membrane

Cytoplasmic filament

Cytoplasmic ring

Luminal ring

Inner nuclear membrane Central channel

Scaffold Nuclear ring

B Nuclear basket

Nucleus

A FIGURE 1.42  (A) Diagram of a nuclear pore complex in a nuclear membrane. (B) TEM showing a tangential thin section through nuclear pore complexes of a tobacco root tip cell. Arrows indicate pores in which the central transporter plug depicted in (A) is clearly seen. Source: (B) Micrograph by Takashi Murata, National Institute of Basic Biology, Okazaki, Japan.

resembles the ER in having functional ribosomes on its cyto­ plasmic face. The perinuclear space is therefore continuous with the ER lumen. A meshwork of 10‐nm‐diameter filaments, called the nuclear lamina, underlies the inner envelope mem­ brane (see domain 1 in Fig. 1.14). The lamina links the nuclear pore complexes and anchors and organizes the interphase chromatin at the nuclear periphery.

1.8.2  Nuclear pore complexes function as both molecular sieves and as active transporters The density of nuclear pores embedded in the envelope varies considerably, depending on the type of cell. In plant cells, pores occupy from 8% to 20% of the envelope surface, at a pore density of 6–25 µm–2. The pattern of pore distribution over the envelope varies in different organisms and cell types. Each pore consists of an elaborate macromolecular assem­ blage known as the nuclear pore complex (Fig. 1.42). Nuclear pore complexes have an octagonal symmetry and appear similar in size and architecture throughout the plant and ani­ mal kingdoms. A nuclear pore complex is ~50 nm in diame­ ter, has an estimated molecular mass of 125 MDa, and is composed of multiple copies of at least 30 different proteins, nucleoporins. Nuclear pore complexes regulate trafficking between cyto­ plasm and nucleus. They permit rapid diffusion of small (90% sequence identity, but each recognizes a different race of powdery mildew. The recognition specificity of distinct MLA alleles is conferred by their LRR domains. Only a few of the first‐discovered R proteins directly bind the cognate effector. Many R proteins recognize the effects of pathogen effectors on host proteins (Fig. 21.45) rather than the effectors directly. As described in Section  21.2.4, plants respond to pathogens by activating PTI, and in turn, some pathogens can overcome these defenses by producing effector molecules to modify or suppress host responses. Each R protein might act as a guard of a host component (the “guardee”). When the pathogen effector molecule modifies this component, the modification is recognized by the R protein and ETI is activated. This guard hypothesis predicts that multiple pathogen effector proteins may interact with a common host target that can be guarded by more than one R protein. The CC‐NB‐ LRR R proteins RPS2 and RPM1 recognize AvrRpt2, or AvrRpm1 and AvrB, respectively. The Arabidopsis RIN4 protein is targeted by these three effectors, which are delivered into plant cells via the bacterial type III secretion system (Fig.  21.46). RIN4 is a 211 amino acid plasma membrane‐ associated p ­ rotein, which interacts with the nonrace‐specific disease resistance 1 protein NDR1 that triggers SA production. The AvrB and AvrRpm1 proteins cause phosphorylation of RIN4, whereas the AvrRpt2 protein is a protease that degrades it. Arabidopsis plants that carry the RPM1 gene are

Virulence target

Susceptible response to favour pathogen growth and development

AVR

A Compatible interaction R2

R1 AVR1 Resistance by direct interaction

Virulence target AVR2 Resistance by guarding

B Incompatible interaction FIGURE 21.45  Guarding of pathogen virulence targets by plant R proteins. (A) A virulence target exists in a susceptible host plant. Upon pathogen infection, the Avr effector binds to its cognate virulence target, resulting in modifications to the target. These modifications lead to pathogen virulence and host susceptibility, thereby generating a compatible interaction. (B) Incompatible interactions conferred by a resistant host plant can arise in two ways. (Left) The R1 protein directly recognizes the Avr1 effector itself. An example of this type of interaction occurs between rice blast Avr‐Pita and rice Pi‐ta. (Right) The R2 protein is a guard protein, recognizing the modified plant virulence target caused by the earlier binding of the Avr2 effector. An example of this type of interaction occurs between AvrB/AvrRpm1 (Avr), RIN4 (the virulence target) and RPM1 (R2). Direct recognition by R1 may be circumvented by alterations of the Avr1 effector without modifying its virulence function. By contrast, recognition mediated by the guarding R2 protein cannot be circumvented by alterations of the Avr2 effector without affecting its virulence function.

Chapter 21  Responses to Plant Pathogens

Extracellular Resistant host

AvrRpm1 or AvrB

Susceptible host

Resistant host

A

RIN4 P P P

Pto

Prf

No PRF or PTO (prf or pto plants) Increased virulence owing to AvrPto and AvrPtoB activity on unknown targets

N DR

N DR

Cell wall Pto

1

AvrPto or AvrPtoB

1

Susceptible host

Pseudomonas syringae

RIN4 P P P

RPM1

No RPM1 (rpm1 plants)

HR Less pathogen growth Cytosol

B

Increased virulence owing to AvrRpm1 activity on RIN4 and other targets

HR Less pathogen growth

Cf-2 Susceptible host

Rcr3

1

RIN

4

C

Rcr3

DR RIN4

RIN

Increased virulence owing to AvrRpt2 activity on RIN4 and other targets

Avr2 Rcr3

N

1 DR N RIN4

No RPS2 (rps2 plants)

Avr2

Resistant host

AvrRpt2

Cf-2

4

RPS2 RPS2 HR Less pathogen growth

Inhibition of Rcr3 protease activity by the Avr2 protease inhibitor contributes to disease

Activation of Cf-2-dependent defense response

Susceptible host

Resistant host

D

FIGURE 21.46  Direct and indirect recognition of pathogen effectors triggers plant defenses. (A) Pto is a tomato serine/threonine protein kinase that requires the NB‐LRR protein Prf. The two proteins form a molecular complex in which Prf guards Pto. Pto is the target of two unrelated P. syringae effectors, AvrPto and AvrPtoB, delivered into the cell by the type III secretion system (T3SS; brown and green syringe). Each effector contributes to pathogen virulence. (B) Arabidopsis RPM1 is a peripheral plasma membrane NB‐LRR protein. It is activated by either the AvrRpm1 or the AvrB effector proteins. AvrRpm1 enhances the virulence of some P. syringae strains on Arabidopsis as does AvrB on soybeans. AvrRpm1 and AvrB are modified by eukaryote‐specific acylation once delivered into the cell by the T3SS and are thus targeted to the plasma membrane. The biochemical functions of AvrRpm1 and AvrB are unknown, although they target RIN4, which becomes phosphorylated (+P), and activate RPM1. In the absence of RPM1, AvrRpm1 and AvrB presumably act on RIN4 and other targets to contribute to virulence. Light blue unlabeled ovals in this and subsequent panels represent unknown proteins. (C) RPS2 is a plasma membrane NB‐LRR protein. It is activated by the AvrRpt2 cysteine protease type III effector from P. syringae. AvrRpt2 is a third effector that targets RIN4. Cleavage of RIN4 by AvrRpt2 leads to RPS2‐mediated ETI. In the absence of RPS2, AvrRpt2 presumably cleaves RIN4 and other targets as part of its virulence function. (D) The transmembrane R protein Cf‐2 guards the extracellular cysteine protease Rcr3. Cf‐2 recognizes the C. fulvum extracellular effector Avr2, which encodes a cysteine protease inhibitor. Avr2 binds and inhibits the tomato Rcr3 cysteine protease. Mutations in Rcr3 result in the loss of Cf‐2‐ dependent recognition of Avr2. Hence, Cf‐2 seems to monitor the state of Rcr3 and activates defense if Rcr3 is inhibited by Avr2. Remarkably, a nematode effector also targets Rcr3 and triggers Cf‐2‐dependent resistance. Note that in D the plant cell wall is not shown for clarity.

resistant to  infection by P. syringae strains carrying the avrRpm1 or avrB genes, because RPM1 recognizes phosphorylated RIN4. RPM1 then activates a signaling pathway that leads to defense activation. Similarly, Arabidopsis plants that carry RPS2 are resistant to infection by P. syringae carrying avrRpt2. RPS2 is normally found in a complex with RIN4, and degradation of RIN4 by the AvrRpt2 protease releases RPS2, rendering the R protein capable of activating a defense response. Thus, the guarding of RIN4 by at least two different

R proteins assures continuous monitoring of the RIN4 protein status. The tomato Mi‐1.2 protein also belongs to the CC‐NB‐ LRR class of R proteins, but is extended at the C‐terminus with a Solanaceae-specific domain. Remarkably, Mi‐1.2 confers resistance to the root‐knot nematode Meloidogyne incognita, the potato aphid Macrosiphium euphorbiae, the sweet potato white fly Bemisia tabaci, and the tomato psyllids insect Bactericerca cockerelli. These diverse resistances conferred by

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Part V  PLANT ENVIRONMENT AND AGRICULTURE

Apoplast Pseudomonas syringae Avr Rpm1

Avr Rps4 Cell wall

1

AvrB

N DR

1030

Plasma membrane

RIN4 EDS1

Nucleus RPS4 RPM1

RRS1 W

RPS2 Salicylic acid Resistance

Cytosol

FIGURE 21.47  The Nonrace‐Specific Disease Resistance 1 protein (NDR1) and the ENHANCED DISEASE SENSITIVITY 1 protein (EDS1) mediate, respectively, the defense signaling abilities of CC‐NB‐LRR and TIR‐NB‐LRR disease resistance proteins. (Left) NDR1 spans the plasma membrane and interacts with the RIN4 protein that resides in the cytoplasm in close proximity to the plasma membrane. RIN4 is targeted by three bacterial effectors delivered into plant cells via the type III secretion system. The guarding of RIN4 by at least two different R proteins (RPM1 and RPS2) assures continuous monitoring of the RIN4 protein status. RPM1 or RPS2 then activate a signaling pathway that leads to defense activation. (Right) EDS1 is required for resistance mediated by the TIR‐NB‐LRR class of R proteins (RPS4 and RRS1) as well as basal defense against many virulent pathogens. EDS1 is a lipase‐like protein that always functions in association with either of two other lipase-like proteins, PAD4 or SAG101. Both the cytoplasmic and nuclear pools of EDS1 are required to activate defense signaling. At least two bacterial effectors interact with the WRKY domain of the RRS1 protein. Both RRS1 and RPS4 cooperate to activate plant defenses. Although the NDR1 and EDS1 proteins function in two different signaling pathways, these must later converge, because all these resistances require the induction of salicylic acid synthesis.

one R gene might be due to each invertebrate targeting a shared guardee that is guarded by Mi‐1.2. The Prf protein (a NB‐LRR protein) from tomato (Solanum lycopersicum) may recognize the direct interaction between Pto and AvrPto. The Pto protein kinase confers resistance in  tomato to strains of Pseudomonas bacteria that express AvrPto. Bacteria deliver AvrPto protein into the plant cell cytoplasm via a T3SS (see Section  21.3.3). When AvrPto interacts with the Pto kinase, both proteins are directed to the plasma membrane via an N‐terminal myristoylation site. The direct interaction between the Pto and AvrPto proteins is recognized by Prf, which then activates various downstream defense responses (Fig. 21.46A). The discovery of the direct Pto–AvrPto interaction was important because cytoplasmically localized protein kinases were previously unknown to function as receptors in any organism and the same domain in the Pto kinase confers both recognition and signaling capacity. Another example of R protein guarding is provided by the tomato Cf‐2 protein. Originally identified as mediating resistance to specific races of the fungus C. fulvum expressing the effector Avr2 (Table 21.3), Cf‐2 also confers resistance to the cyst nematode Globodera rostochiensis. Cf‐2 enables the

plant to recognize binding of the Avr2 proteinase inhibitor (PI) of C. fulvum to the plant apoplastic protease Rcr3 (Fig.  21.46D), which then triggers defense responses. The presence of Cf‐2 also senses binding of the venom allergen‐ like effector protein Gr‐VAP1 of G. rostochiensis to the Rcr3 protease. In the absence of Cf‐2, variants of Rcr3 encoded by specific alleles increase susceptibility of tomato plants to G. rostochiensis, revealing a key role for Rcr3 as a virulence target for this cyst nematode species. Some alleles of Rcr3 trigger Cf‐2‐dependent defense, even in the absence of Avr2, suggesting that selection must constrain guardee alleles to avoid inadvertent R protein‐dependent defense activation. The TIR‐NB‐LRR class of R proteins can confer resistance to biotrophic fungi, bacteria, and viruses. How might they signal? The presence of the conserved TIR domain has raised the possibility that immune responses in plants, mammals, and invertebrates utilize an ancient, conserved mechanism, or arose via convergent evolution (see Fig. 21.40). Intriguingly, the protein kinases PELLE and IRAK that function downstream of the Drosophila Toll and human IL‐1R receptors, respectively, show strong homology to the PTO resistance protein (Fig. 21.46, see also Fig. 21.40).

Chapter 21  Responses to Plant Pathogens Lid open ATP

ATP

Lid closed ATP ATP

ATP

Catalytic Arg

CH 2 ATP

Mature NLR

Complex dissociation

Formation of intramolecular interaction

Immature NLR TPR SGS CS

TPR SGS CH2 CS ADP

Closed conformation

1

Open conformation

ADP

ATP

CH

CH2

CH1

TPR SGS

CH1

Lid swinging

CH2 CS

Catalytic Arg

CH1

ATP hydrolysis

Complex assembly

FIGURE 21.48  A proposed model for NB‐LRR (NLR) protein maturation by the HSP90–SGT1–RAR1 ternary complex. The three proteins are HEAT SHOCK PROTEIN 90 (HSP90, brown), SUPPRESSOR OF THE G2 ALLELE OF SKP1 (SGT1, purple), and REQUIRED FOR MLA12 RESISTANCE 1 (RAR1, blue). This model shows HSP90 as the central protein of the cycle that involves the following interacting domains: the N‐terminal domain of HSP90 containing the ATP‐binding site (yellow half‐circle), the TPR, SGS and CS domains of SGT1, and cysteine‐ and ­histidine‐rich domain 1 and 2 (CH11 and CH2) of RAR1. In this model, interactions occur sequentially in a clockwise direction. Upon ATP binding, the lid segments of the HSP90 dimer form a closed conformation to create an active ATPase enzymatic site with the key arginine in the catalytic (green triangle). When the RAR1 CH1 domain binds to one of the ATP‐loaded HSP90 lids this prevents the formation of the lid‐closed ­conformation. Binding of the RAR1‐CH1‐domain then facilitates the interaction of the RAR1 CH2‐domain with the other HSP90 lid of the dimer. The bound RAR1 CH‐domains promote the formation of the HSP90 complex with SGT1 and an immature NLR protein (orange). The stable HSP90‐RAR1‐SGT1‐NLR ternary complex then triggers the hydrolysis of ATP by the catalytic Arg to facilitate the intramolecular reorganization of the NLR protein. After release of the ADP, the RAR1‐SGT1‐NLR complex dissociates from HSP90 and releases the mature NLR protein (red).

21.6.5  Other plant proteins are required for R protein action Mutagenesis of resistant plant lines carrying an R gene followed by screening for disease‐sensitive mutants has revealed other plant genes required for disease resistance. In Arabidopsis, mutations in the EDS1 (ENHANCED DISEASE SENSITIVITY 1) gene eliminate race‐specific resistance mediated by the TIR‐NB‐LRR class of R proteins, and also reduce basal defenses against many virulent pathogens. EDS1 is required for many (but not all) RPP gene‐­ mediated resistances to various races of the downy mildew oomycete (Hyaloperonospora arabidopsidis), and for RPS4‐ and RRS1‐mediated resistance to P. syringae carrying AvrRps4. EDS1 is a member of a family of proteins that carry an atypical lipase motif, and has been located to both the nuclear and cytoplasmic compartments. EDS1 likely functions as a heterodimer with either of two related proteins PAD4 and SAG101. PAD4 mutants are also more disease‐­susceptible. Although mutations in EDS1 and NDR1 (see section  21.6.4)

c­ om­promise function of TIR‐NB‐LRR and CC‐NB‐LRR signaling, both proteins are required for SA production, indicating that the signal transduction pathways involving these proteins later converge (Fig. 21.47 and see Section 21.8.4). In barley and Arabidopsis, mutations in either RAR1 (required for Mla‐mediated resistance 1) or SGT1 (suppressor of the G2 allele of skp1) result in susceptibility to powdery mildew fungi or downy mildew, respectively, even when the plant genotypes carry functional copies of various race‐specific R genes and the fungal races express the correct cognate Avr effector. These two proteins, in conjunction with HSP90 (heat shock protein 90) (Fig.  21.48), are required for resistance mediated by multiple R proteins recognizing viral, bacterial, oomycete, or fungal pathogens. RAR1 contains two highly conserved zinc‐binding domains called CHORD‐I and CHORD‐II (cysteine‐ and histidine‐rich domain). The CHORD‐I domain binds to the molecular chaperone HSP90. Both RAR1 and HSP90 stabilize certain NB‐LRR proteins and thereby ensure the R proteins can assume a conformation that is competent to receive

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pathogen signals. Both RAR1 and HSP90 bind to SGT1 (Fig. 21.48), which is required for the function of R proteins of the TIR‐NB‐LRR class, the CC‐NB‐LRR class, the eLRR class, and RPW8. SGT1 contains three distinct domains: a tetratricopeptide repeat domain (TPR), the CS motif (present in CHP and SGT1 proteins), and the SGS motif (an SGT1‐ specific sequence). The CS motif of SGT1 is sufficient to bind to the ATPase domain of HSP90 as well as interact with the CHORD‐II domain of RAR1. The structure of the HSP90–SGT1–RAR1 complex has been elucidated, and a dynamic model for the interaction of the three components proposed (Fig. 21.48). Overall, this HSP90– SGT1–RAR1 chaperoning complex is proposed to maintain the steady state levels of different R proteins and ensure that R proteins are correctly folded and maintained in a recognition‐ competent state in the appropriate cellular compartment. It may be required for the conformational changes involved in resistance activation. Misfolded R proteins can inappropriately activate defense and therefore must be immediately inactivated and degraded. This chaperon complex is structurally and functionally conserved in other eukaryotes. Some NB-LRR proteins require other NB-LRR proteins to function. The N REQUIRED GENE 1 (NRG1) is required for function of the tobacco TIR‐NB‐LRR protein N, which confers resistance against TMV through recognition of the TMV p50 protein. NRG1 is a CC‐NB‐LRR type R protein. Similarly, the TIR‐NB‐LRR protein RPS4 also requires the TIR‐NB‐ LRR‐WRKY protein RRS1 for its function (and vice versa) (see Section  21.5.5). In some CC‐NB‐LRRs, the CC domain resembles that present in the atypical Arabidopsis R protein RPW8 (see Section  21.6.3). This is referred to as the CCR domain. CCR‐NB‐LRR‐encoding genes are present in most plants; examples include the Nicotiana benthamiana NRG1 protein and the Arabidopsis activated disease resistance gene 1 (ADR1) protein. CCR domains are sufficient for the induction of defense responses, and this activity is SGT1‐­independent. Both CCR domain genes are unique to genomes that carry TIR‐NB‐LRR genes, suggesting CCR domain proteins may be required for signaling by TIR‐NB‐LRR R proteins.

21.6.6  Many R genes can function in heterologous plant species Most R gene sequences confer resistance when introduced into a susceptible genotype of the original host or a closely related species. For example, the tobacco N gene is active in tomato, where it restricts TMV infections to localized necrotic lesions, as it does in tobacco. Thus, Avr‐dependent R protein signaling cascades are conserved between related plant species. Pto, Cf‐9, and Bs2 from Solanaceous plants do not confer recognition of their corresponding effectors in Arabidopsis. However, barley Mla can confer function in Arabidopsis, and Arabidopsis RPS4 and RRS1 can function in Solanaceae. Overexpression of some R genes in heterologous plants results

in necrotic symptoms in the absence of pathogen attack, or plants with altered development. These data highlight how finely tuned the relationship is between R proteins and signaling partners in their native ­species. The evolution of R gene sequences is most likely constrained not only by selection for pathogen recognition, but also for functional interactions with guardee proteins, and for  avoidance of recognition of related endogenous plant ­proteins that control plant development.

21.7  Other sources of genetic variation for resistance 21.7.1  A few plant R proteins are unique A few R genes provide broad‐spectrum resistance to multiple pathogenic species, and functional homologs of these R genes have not been reported in other plants. Potentially each R protein could provide a plant species‐specific function, but it is more likely that the corresponding R gene needs to be ­formally identified in other plant species. RPW8.2 from Arabidopsis confers broad‐spectrum resistance to several powdery mildew species, including Golovinomyces orontii. RPW8 proteins feature an N‐terminal transmembrane domain and one or more coiled‐coil (CCR) domains (homologous to the ADR1/NRG1 subclass of CC‐ NB‐LRR proteins), and they lack an LRR domain. The wheat Lr34 gene provides durable adult plant resistance to the basidiomycetes Puccinia triticina and P. striiformis, which respectively cause leaf rust and stripe rust, and to the ascomycete powdery mildew species Blumeria graminis f. sp tritici. Through plant breeding, the Lr34 gene is now present in most wheat cultivars globally. The Lr34 gene sequence encodes a putative adenosine triphosphate binding cassette (ABC) transporter that may translocate toxic compounds stored in plant cells (possibly in the vacuole) to the point of fungal ingress, arresting growth. Two major quantitative loci, rhg1 and Rhg4, control root infections in soybean (Glycine max) crops caused by the cyst nematode Heterodera glycines. Roots of plants carrying the Rhg genes are penetrated by infective juvenile nematodes, but the feeding nematodes die before reaching adult stages (see Section 21.1.5). The Rhg4 locus encodes a serine hydroxymethyltransferase, an enzyme that is ubiquitous in nature and structurally conserved across kingdoms. It is responsible for interconversion of serine and glycine and essential for one‐ carbon metabolism. Transcriptional and metabolic profiling studies indicate the syncytial feeding cells support a demand for folate one‐carbon metabolism. Nematodes, like other animals, acquire folate from their diet; therefore, impaired folate production at induced feeding site may lead to a nutritional deficiency that starves the nematode.

Chapter 21  Responses to Plant Pathogens

21.7.2  Insensitivity to host‐selective toxic proteinaceous effectors and metabolites is important in plant defense against necrotrophs Two closely related necrotrophic fungal pathogens, Stagnospora nodorum and Pyrenophora tritici‐repentis, produce host‐selective toxins that induce susceptibility only in wheat genotypes that harbor the corresponding toxin sensitivity gene Tsn1. The Tsn1 protein has several R protein‐like features, including serine‐threonine kinase, NB, and LRR domains, which are required for ToxA sensitivity and disease susceptibility. Wheat genotypes that are insensitive to ToxA lack the entire gene sequence, and Tsn1 does not seem to directly bind the ToxA effector. These findings suggest that necrotrophic pathogens may subvert the resistance mechanisms acquired by plants to combat other pathogen types by making molecules that provoke cell death, and have also evolved ways to thrive in environments that would be detrimental to biotrophic pathogens.

21.7.3  Recessively inherited resistance genes provide resistance against bacteria, fungi, and viruses in cereal and noncereal species Recessively inherited resistance is rarely reported to control either bacterial or fungal infections. However, this mode of inheritance is more frequently encountered in the successful control of plant viruses. Some of the different types of recessive resistance (r) genes and how they function are described below and in Table 21.7. Resistance to some bacteria involves sequence variation in proteins targeted by virulence effectors. In rice, nine of the 30 documented resistance genes against the bacterial blight pathogen Xanthomonas oryzae pv. orzyae are recessively inherited. The xa13 gene occurs as a series of natural alleles of Os‐8N3, whose expression is induced by strains of Xanthomonas ory­ zae pv oryzae carrying the gene pthXo1, which encodes the transcription activator‐like (TAL) effector PthXo1 described in Section 21.3.8. The xa13 alleles are unresponsive to PthXo1, and plants with xa13 are resistant to strains of the pathogen that rely solely on PthXo1 as the essential effector for virulence. Os‐8N3 encodes a member of the NODULIN3 gene family of SWEET proteins that transport sugar. Recessive (mlo) alleles of the Mlo locus in barley, Arabidopsis, tomato, and pea confer broad spectrum resistance to various powdery mildew species. The deduced MLO ≈60‐kDa gene product is predicted to be anchored in the plasma membrane by a conserved seven membrane‐spanning-domain protein. Two mlo mutant alleles are deployed commercially to reduce the risk of mildew infection in barley (Hordeum vulgare).

Recessively inherited resistance against viruses often confers non‐HR immunity, as opposed to an HR‐associated defense. Virus genomes are small and code for just a few functional proteins (see Section  21.3.4; see Chapter  10). To complete their life cycle, viruses rely on host plant proteins involved in DNA replication, transcription, and translation. Therefore, recessive antiviral resistance genes are frequently mutant alleles of genes that encode plant host proteins required at specific steps in the virus life cycle. For example, mutations in the eukaryotic translation initiation factor 4E (eIF4E) or its isologs eIF(iso)4E occur in many cereal and noncereal plant species, where they confer resistance primarily against RNA viruses belonging to the Potyviridiae family (Fig. 21.49). These viruses encode VPg, a small protein that is attached to the 5′ end of the viral RNA genome. VPg is a likely functional equivalent of the eukaryotic mRNA cap structure, which is important for translation initiation and recruitment of ribosomes via interaction with eIF4E or eIF(iso)4E (see Chapter 10). Both the potyviral VPg protein and the eIF4E/eIF(iso)4E proteins are multifunctional and are involved in processes other than initiation of protein synthesis. The exact purpose of the VPg–eIF4E interaction has still to be discovered; however, specific mutations in the host eIF4E gene lead to a failure of the potyvirus to complete their life cycle (i.e., translation, replication, or cell‐to‐cell trafficking), which results in resistance without HR formation.

21.8  Local and systemic defense signaling Within minutes of pathogen, insect, or nematode attack, plant defense responses are activated locally, and within hours, defense responses are elaborated in tissues distant from the invasion site and even in neighboring plants. Remarkably, the type of induced systemic response is determined by the identity of the initial attacking organism. As shown in Figure 21.50, induced systemic responses to fungi, bacteria, and viruses are distinct from those to insects. Nematodes appear to induce a mixture of the two. The adaptive significance to plants of orchestrating systemic responses is enormous. They ensure that distant plant tissues can deal more effectively with subsequent attackers.

21.8.1  Plants activate systemic responses to pathogens Fungi, bacteria, and viruses systemically activate a defense response known as systemic acquired resistance (SAR). For SAR to occur, the initial infection must result in formation of necrotic lesions, either as part of the HR or as a symptom of disease. SAR is induced strongly by ETI and weakly by PTI, and during SAR, a specific subset of pathogenesis‐related (PR) genes (described later) is activated in the plant tissues distant to the initial pathogen infection site. SAR activation

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Arabidopsis

Rice (Oryza sativa)

Rice

rrs1‐r

xa5

xa13 (Os‐8N3)

Bacterial blight pathogen Xanthomonas oryzae pv. oryzae

Bacterial blight pathogen Xanthomonas oryzae pv. oryzae

Extracellular/multiple organs/all tissue layers

Extracellular/multiple organs/all tissue layers

Extracellular/multiple organs/all tissue layers

Obligate biotroph/ multiple organs/ epidermal specific

Various powdery mildew fungal species

Barley (Hordeum vulgare), Arabidopsis and tomato

mlo

Bacterial blight pathogen Ralstonia solanacearum

Systemic intracellular infections throughout the plant

RNA viruses mostly belonging to the Potyviridiae family

Multiple cereal and noncereal species

eIF4E or its isologs eIF(iso)4E

Various extracellular and intracellular lifestyles on leaves

Infection type/organ/ cell layer attacked

Enhanced resistance to Pseudomonas syringae, Hyaloperonospora arabidopsidis, powdery mildew Golovinomyces cichoracearum, but enhanced susceptibility to Colletotrichum higginsianum and the necrotrophic Alternaria brassicicola

Pathogen(s)

Arabidopsis

Plant

edr1

Gene

TABLE 21.7 Recessively inherited resistance genes.

A member of the NODULIN3 gene family of SWEET proteins that transport sugar

Subunit of the general transcription factor TFIIA required by RNA polymerase II

Yes

Yes

Yes

The mutant xa5 protein, which has only two residue changes, retains its basic transcription factor function and may possess enhanced ability to interact with the bacterial effector Avrxa5 xa13 alleles are unresponsive to the TAL effector PthXo1

No

Negative regulator of plant defense and/or PCD

Plant‐specific seven transmembrane helices protein

Upon physical interaction with bacterial effector Pop2, RRS1 cooperates with another TIR‐NB‐LRR protein RPS4 to activate defense

Mostly yes Mutations result in abolition of interaction with the viral Vpg protein attached to 5’ end of the viral RNA genome. As a result, virus fails to hijack various host processes to complete its life cycle

Eukaryotic translation initiation factor. Multifunctional and involved in processes other than the initiation of protein synthesis

TIR‐NB‐ARC‐LRR‐ WRKY

No

Race specific?

Mutation creating a premature stop codon eliminates the kinase domain. Proposed to be a negative regulator of plant defense

Proposed/known protein function

A Raf‐like mitogen‐ activated protein kinase kinase kinase (MAPKKK)

Protein type

Chapter 21  Responses to Plant Pathogens

Susceptible host

AUG VPg 4E PA B

P

3

4A

4G

4B

Resistant host 40s ribosomal subunit 4E PA B

Viral RNA

P

3

4A

4G

4B

A AAAAAAAAA

AUG VPg

A AAAAAAAAA

Viral VPg incorporates into translation initiation complex

No interaction between VPg and translation initiation complex

Viral proliferation

Resistance

Cytosol

FIGURE 21.49  R proteins and defense activation. Recessive resistance to potyviruses is conferred by mutant alleles of a eukaryotic transla­ tion ­initiation factor 4E (eIF4E). Initiation of translation in plants (and other eukaryotes) uses a multi‐protein complex consisting of initiation factors 3, 4A, 4B, 4E, 4G, poly(A)‐binding proteins (PABPs), 40S ribosomal subunit and several other minor components. An interaction between the mRNA cap structure (m7GpppG) and eIF4E is required for efficient translation. Potyviruses produce a small protein called VPg, which is ­covalently attached to the 5′‐end of their RNA genomes and is likely to play a role similar to the mRNA cap structure during translation i­nitiation (left). Some naturally occurring structural variants of eIF4E confer resistance to potyviruses in many plant species, probably because of their inability to bind potyviral VPg and recruit potyviral RNA into the translation initiation complex (right).

also leads to a significant reduction in disease symptoms after subsequent infection by many different pathogen species. For example in tobacco, N‐gene mediated resistance against TMV protects the plant against later infection by most, but not all, tobacco pathogens, including an identical TMV strain. SAR can convert normally genetically compatible plant–pathogen interactions into incompatible ones. Therefore, by understanding how SAR works, it may be possible to engineer broad‐­spectrum disease control into crops (see Section 21.10). A requirement for SA in SAR activation was demonstrated by expression of the bacterial NahG SA hydroxylase in tobacco and Arabidopsis plants. These plants do not accumulate free SA and are incapable of activating SAR. So is SA the translocated signal? In vivo 14C‐SA labeling studies revealed that, following TMV infection on N tobacco, as much as 70% of the SA increase in uninfected tissue results from its translocation from the infected leaves; a series of grafting experiments between wild‐type and NahG plants, however, indicated that SA is only needed in the distal plant organs for SAR to be induced (Fig. 21.51). Therefore, SA is probably not the mobile signal in tobacco. Grafting experiments using tobacco showed that SA methyltransferase activity, which converts SA into methylsalicylic acid (MeSA), is required in the tissue that generates the mobile signal. Conversely, MeSA esterase activity, which converts MeSA back to SA is required for signal perception in the distal tissues. However, in Arabidopsis, SA is required for SAR, whereas MeSA is dispensable for SAR. Thus, MeSA may have a signaling role for SAR induction in tobacco, but not in all plant species.

SA induces or potentiates many plant defense responses and induces a set of pathogenesis‐related (PR) proteins. Seventeen families of PR proteins have been classified (Table 21.8), and the same types of PR protein families occur in most plant species. Transcripts of PR genes accumulate within minutes to hours of pathogen attack and PTI or ETI activation, and they are also induced in compatible inter­ actions, but much more weakly and slowly. Induction of PR proteins distal to the initial site of infection frequently occurs. Several PR proteins are also induced as plants start to flower or experience abiotic stress. Some PR proteins function as chitinases and glucanases, degrading fungal cell wall structural polysaccharides and impairing fungal growth. SA regulates the transcriptional activation of many PR genes. Also, ethylene and SA can act synergistically, enhancing PR gene expression even further. Other PR defense proteins, such as lipoxygenase, may contribute to defense by generating secondary signal molecules, such as jasmonic acid (JA) and lipid peroxides, and by producing an array of toxic volatile and nonvolatile secondary metabolites with antimicrobial activity. Various synthetic chemicals induce SAR. Two of the most  potent are 2,6-dichloroisonicotinic acid (INA) and benzo(1,2,3) thiodiazole‐7‐carbothionic acid S‐methyl ester (BTH). Both compounds act as SA mimics and activate SAR when applied to NahG plants. To understand the molecular and biochemical basis of SAR, mutagenesis screens using Arabidopsis revealed the SAR‐deficient mutant dir1‐1 (defective in induced resistance). The dir1‐1 mutant plants can activate defenses locally,

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Phloemmobile signal

Systemic wound response proteins and proteinase inhibitors accumulate in distant tissues

Pathogenesis-related protein induction in distant tissues

MeSA

SA Increase in salicylic acid

MeJA Virus, fungus, or bacteria

SA

JA

Transient C2H4 synthesis

Nematode

A Systemic acquired resistance

Chewing insects or mechanical wounds

Nematode

B Systemic proteinase inhibitor/wound response No pathogenesis-related protein induction in distant tissues

Transient synthesis of JA and C2H4

Nonpathogenic rhizobacteria

C Induced systemic resistance FIGURE 21.50  Systemic responses to plant pathogens. (A) Viruses, fungi, and bacteria activate systemically a subset of defense responses in a phenomenon known as systemic acquired resistance (SAR), in which local necrosis formation at the initial site of pathogen invasion triggers both a local increase in salicylic acid (SA) and formation of a phloem‐mobile signal. Subsequently, in distal plant tissue, SA concentrations increase and volatile methyl‐SA (MeSA) is released. Together, these signals induce synthesis of various pathogen‐related proteins in non‐invaded parts of the plant. (B) In contrast, attack by chewing insects or mechanical wounding activates a different protective response, the systemic PI/wound response, wherein initial tissue damage causes a transient increase in the synthesis of ethylene and jasmonic acid (JA). Volatile methyl jasmonate (MeJA) and another phloem‐mobile signal called systemin then activate the systemic responses, which include the accumulation of proteinase inhibitors (PIs) and other systemic wound response proteins (SWRPs). Root‐attacking nematodes appear to induce a mixture of both the SAR and systemic PI/wound responses. (C) Induced systemic resistance (ISR) caused by soil‐inhabiting nonpathogenic rhizobacteria colonizing plant roots. ISR requires both JA‐ and ethylene‐mediated signaling to induce protective defense responses in the distant leaf tissue. This form of defense does not involve accumulation of pathogenesis-related (PR) proteins or require SA.

i­ndicating DIR1 is only required for the systemic responses. DIR1 is predicted to encode an apoplastic lipid‐transfer protein, and this protein was detected in the vascular exudates of wild‐type plants after localized pathogen inoculation. Therefore, the DIR1 protein has a role in either the generation

of the systemic signal or transport of another lipid‐based signal to distal tissues. A mutant impaired in the biosynthesis of glycerol 3‐­phosphate (G3P) also fails to activate SAR. The SAR response can be rescued by the addition of either G3P or vascular exudates from

Chapter 21  Responses to Plant Pathogens Control grafts

Scion

Stock Xanthi/Xanthi

nahG/nahG

Reciprocal grafts

nahG/Xanthi

Xanthi/nahG

FIGURE 21.51  Grafting experiments demonstrate that salicylic acid (SA) is unlikely the phloem‐mobile signal that activates SAR. An absolute requirement for SA in systemic acquired resistance (SAR) was demonstrated using transgenic tobacco (Nicotiana tabacum) and Arabidopsis expressing the bacterial nahG gene, in which the NahG SA hydroxylase degrades SA to catechol, CO2, and water. These plants did not accumulate free SA and were incapable of activating SAR. Then a series of grafting experiments between wild‐type and transgenic nahG‐expressing plants revealed that, to induce SAR, SA only needed to be present in the distal plant organs. Reciprocal and control grafts were generated using two types of tobacco plants, which expressed either the N resistance gene alone (cultivar Xanthi) or the N gene in combination with the nahG transgene to remove inducible SA (nahG). The four types of grafted plants were inoculated with TMV on the lower rootstock leaves; 7 days later, the same TMV isolate was inoculated onto the upper scion leaves. The photographs show the infection types on the scion leaves five days after the second TMV ­inoculation. The nahG scions grafted onto the Xanthi rootstocks (nahG/Xanthi) were unable to mount an SAR response. In contrast, Xanthi scions grafted onto nahG rootstocks (Xanthi/nahG) demonstrated SAR responses similar to those of the control Xanthi/Xanthi grafts not expressing the nahG transgene. The nahG/nahG grafts lacking SA were unable to mount an SAR response. Source: (Photos) Vernooij et al. (1994). Plant Cell 6:959–965.

induced wild‐type plants to the distal tissues. Thus, G3P may also be a signaling molecule in SAR that requires DIR1. In addition, analysis of vascular exudates revealed azelaic acid as a mobile signal capable of inducing SAR. Application of azelaic acid leads to induction of the AZI1 (AZELAIC ACID‐INDUCED 1) gene, which is predicted to encode a secreted lipid‐transfer protein. Reciprocal application of vascular exudates from wild‐ type and azi1 mutant plants show that AZI1 is involved in the production or translocation of a mobile signal. Together, these studies reveal that multiple mobile signals are transported through the vasculature to systemic, uninfected parts of the plant, where they induce the accumulation of SA, a signal molecule for SAR. These mobile signals in Arabidopsis and their roles and potential interplay are depicted in Figure 21.52.

21.8.2  NPR1, NPR3, and NPR4 mediate SA induction of PR genes A key protein in SAR is NPR1 (NONEXPRESSOR OF PATHOGENESIS‐RELATED GENES 1) (Fig.  21.53). Arabidopsis npr1 mutant plants fail to induce PR genes upon

treatment with chemical inducers, such as SA and INA, and are compromised in SAR. The npr1 mutant does, however, show a stronger HR and greater SA accumulation in response to genetically incompatible pathogens, whereas NPR1 over‐ expressing Arabidopsis lines exhibit a weaker HR. Therefore NPR1 is a positive regulator of SAR but a negative regulator of effector triggered HR (ETI). NPR1 encodes an ankyrin repeat protein that positively regulates SA signaling and is a member of a small gene family that includes NPR3 and NPR4 (see later). Similar ankyrin repeats are found in numerous eukaryotic proteins involved in protein–protein interactions. In healthy plant cells, NPR1 is retained in an oligomeric form in the cytoplasm by intermolecular disulfide bridges, and S‐nitrosylation by nitric oxide (NO) facilitates NPR1 oligomerization. The NO moiety becomes covalently attached to the thiol side chain of cysteine, resulting in protein S‐nitrosylation (NPR1 oligomer). Upon pathogen infection, thioredoxins also located in the cytoplasm counteract the S‐nitrosylation of NPR, leading to monomer release and migration of this transcription cofactor to the nucleus (NPR1 monomer). SA affects cellular redox state and NPR1 nuclear translocation. Once inside the nucleus, NPR1 binds to TGA (TGACG‐motif binding) transcription factors, enhancing

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Part V  PLANT ENVIRONMENT AND AGRICULTURE TABLE 21.8 Recognized families of pathogenesis‐related proteins. Family

Type member

Properties

PR‐1

Tobacco PR‐1a

Unknown

PR‐2

Tobacco PR‐2

β‐1,3‐glucanase

PR‐3

Tobacco P, Q

Chitinase type I,II,IV,V,VI,VII

PR‐4

Tobacco ‘R’

Chitinase type I,II

PR‐5

Tobacco S

Thaumatin‐like

PR‐6

Tomato Inhibitor I

Proteinase‐inhibitor

PR‐7

Tomato P69

Endoproteinase

PR‐8

Cucumber chitinase

Chitinase type III

PR‐9

Tobacco “lignin‐forming peroxidase”

Peroxidase

PR‐10

Parsley “PRI”

Ribonuclease‐like

PR‐11

Tobacco “class V” chitinase

Chitinase type I

PR‐12

Radish Rs‐AFP3

Defensin

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their binding to SA‐responsive promoters. This then leads rapidly to the transcriptional activation of a wide repertoire of defense responses and pathogen resistance. The NPR1 protein carries a Broad‐complex, Tramtrack, and Bric‐a‐brac BTB domain, also found in various proteins  that make E3 ligase complexes with CULLIN3 (see Chapter 10). In addition, once within the nucleus NPR1 is phosphorylated on two serine residues, and this facilitates the interaction between NPR1 and CULLIN to enhance NPR1 degradation. NPR1 protein turnover (degradation) by the 26S proteasome is required for full induction of the NPR1‐induced genes, the mechanism of which is still under investigation. It is hypothesized that NPR1 turnover facilitates the “refreshment” of the transcription initiation complex. NPR1 is responsible for heightening defense responses in SAR‐induced plants during subsequent pathogen attacks. Two receptors for SA have been identified in Arabidopsis: the low‐affinity NPR3 protein and the high‐affinity NPR4 protein. At the site of infection, NPR1 must be degraded in

the nucleus via interaction with NPR3 to allow PCD and ETI to occur. In a healthy plant, if NPR1 enters the nucleus, it is degraded by NPR4, whose interaction with NPR1 does not require SA. At a distance from the site of infection, NPR4 functions in conjunction with NPR1 and promotes defense gene induction without host cell death. In npr3npr4 double mutant plants, local NPR1 levels remain high and no SAR occurs. Both NPR3 and NPR4 proteins have two functions, as SA receptors and as adaptor proteins that link the NPR1 protein to CULLIN3. Upon binding SA, NPR3 promotes the NPR1–NPR3 interaction, whereas NPR4 disrupts the NPR1–NPR4 interaction. At the high SA concentrations typically induced locally at the sites of initial pathogen infection, the low‐­affinity NPR3 receptor mediates degradation of the cell death suppressor NPR1, thereby permitting activation of ETI and cell death to occur. In contrast, at the lower SA concentrations typically found in distal tissues, SA is unable to bind to NPR3, and cell death is blocked. In these distal tissues, SA instead binds to the high‐affinity

Chapter 21  Responses to Plant Pathogens

O OH

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DIR1/AZI1 O O

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Infected area

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FIGURE 21.52  Systemic activation of plant defense responses. Several proteins and metabolites with various functions are required to establish and maintain the systemic acquired resistance (SAR) response. DIR1 and AZI1 are predicted to be apoplastic lipid‐transfer proteins that have a role in either generation of the systemic signal or transport of various systemic signals to distal tissues. The metabolites glycerol 3‐phosphate (G3P) and azelaic acid are two systemically mobile signals that accumulate in vascular exudates in activated plants. When the mobile signals reach the systemic tissues, accumulation of salicylic acid (SA) occurs and induces or potentiates many plant defense responses, including the induction of a set of pathogenesis‐related (PR) proteins. The induced volatile metabolite methylsalicylic acid (MeSA) may have a SAR signaling role in tobacco, but not in all plant species.

Noninfected area

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NPR1

SA Programmed cell death

Cell survival

Local defense

Systemic defense

FIGURE 21.53  SA‐mediated control of plant cell death and survival. Upon microbial infection, SA levels increase, its concentration decreasing gradually with increasing distance from the site of infection. At high SA concentrations—typically found in infected areas of the plant—the receptor NPR3, which binds SA with low affinity, mediates degradation of the cell‐death suppressor NPR1 (left), thereby favoring programmed cell death and local effector‐triggered immunity. However, at the lower SA concentrations typically found in cells distant from the infection site, SA cannot bind to the low‐affinity receptor NPR3, so cell death is blocked. In these cells, SA instead binds to the high‐affinity receptor NPR4 (right), blocking degradation of NPR1, thereby favoring cell survival and expression of genes associated with activation of systemic defenses.

receptor NPR4, blocking the degradation of NPR1 to favor cell survival and the expression of genes associated with SAR (Fig. 21.53).

21.8.3  Systemic responses to mechanical wound and insect attack occur in plants Many herbivorous insects mechanically wound plant tissue while feeding and induce the rapid accumulation of proteinase inhibitors (PIs) and other systemic wound response proteins (SWRPs) throughout the plant (see Fig. 21.50). The systemic

response to chewing insects is frequently referred to as the wound response, because similar molecular and biochemical events are triggered after plant tissue is mechanically wounded. In tomato, although high concentrations of oligogalacturonides released from damaged plant cell walls induce PI and SWRP genes locally, these oligogalacturonides do not move ­systemically. Instead an 18 amino acid polypeptide called systemin (see Chapter 17) is the mobile systemic signal. Systemin can initiate defense responses at a few femtomoles (10–15 moles) per plant. It is synthesized by cleavage from the C‐terminus of pro‐systemin, a 200 amino acid precursor protein, in the cytoplasm at the site of wounding. Systemin is released from damaged cells, moves throughout the damaged

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leaf within 30 minutes, and is transported via the phloem to upper unwounded leaves within 60–90 minutes. The importance of systemin to plant defense against insect attack has been demonstrated using transgenic tomato plants that produce either continuously high or lower than normal systemin levels in response to wounding. When systemin levels are high, the tomato plants constitutively express defense mechanisms even in the absence of wounding, and are more resistant to herbivory by larvae of the tobacco hornworm Manduca sexta. When systemin production is lower than normal, plants have a severely reduced systemic induction of defense mechanisms and are more susceptible to M. sexta larvae. Once systemin reaches the target tissue in tomato plants, its perception activates several signaling pathways that lead to expression of various defense mechanisms. One pathway is similar to that elicited by P/MAMPs, involving ion fluxes across the membrane, an oxidative burst, and the activation of MAP kinases (see Fig. 21.30). This activates a lipid‐based signaling cascade and JA production (see Chapter  17). JA induces transcription of PI and other SWRP genes; however, for JA induction of PI genes to occur, a third signal molecule is required, the gaseous hormone ethylene. Ethylene transiently accumulates within 30–120 minutes of systemin addition to the translocation stream. When ethylene synthesis is blocked pharmacologically or by reverse genetics using transgenic plants expressing an antisense ACC oxidase gene (see Chapter  17), neither wounding nor systemic or JA treatments can induce PI gene expression. A tomato mutant in JA biosynthesis called defenseless 1 (def1) has a compromised insect defense response (Fig. 21.54). SA inhibits both the octadecanoid pathway and ethylene biosynthesis. The SA inhibition of the wound response explains why PI and SWRP genes are not induced during SAR to pathogens, when high levels of SA accumulate.

Some, but not all, features of the defense response to wounding are conserved across plant species. For example, in both Arabidopsis and tomato, JA signaling plays a crucial role in initiation of the wound response; however, the systemin signaling pathway is found only in the Solanaceae. Arabidopsis has no genes that encode proteins resembling prosystemin; therefore, in Arabidopsis, JA is most likely the signal that directly induces the systemic wound response.

21.8.4  Insect herbivores induce production of proteinase inhibitors In addition to activating the systemic wound response, attacking insect herbivores activate synthesis of a different  group of defense proteins to microbial pathogens that ­interfere with the insect digestive system. They include serine, cysteine, and aspartyl PIs and polyphenol oxidases. These proteins interact with proteins and proteinases in the herbivore gut and inhibit proteolysis of the ingested food. The result is reduced availability of essential amino acids, which either retards herbivore growth and development or kills them. Nematodes induce PI accumulation, which results in similar adverse effects.

21.8.5  Jasmonic acid and ethylene mediate resistance to necrotrophs Plant defensins are defense‐related proteins with antimicrobial activity. This growing family of basic cysteine‐rich peptides with a molecular mass below 7 kDa is of particular

1 cm

A

B

FIGURE 21.54  The tomato mutant defenseless (def1, A, left) is deficient in the biosynthesis of the octadecanoid pathway‐derived signal, jasmonic acid. The defenseless plant has a compromised systemic wound response compared with that of the wild‐type plant (A, right). When larvae of the tobacco hornworm insect (Manduca sexta) feed on defenseless plants (B, left), their growth rate is faster than when feeding on wild‐type tomato plants, which can systemically synthesize PI and other systemic wound response proteins in response to feeding by larvae (B, right). Source: Howe et al. (1996). Plant Cell 8:2067–2077.

β–1

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interest for three reasons. First, unlike SA‐mediated defense responses, plant defensin accumulation in vegetative plant tissues is controlled by a combination of JA and ethylene. Second, JA levels rise in systemic tissues. Third, insects, birds, and mammals produce structurally and functionally similar defensin peptides following microbial attack. For example, the fruit fly Drosophila melanogaster produces a peptide called drosomycin that resembles plant defensins from ­ radish. Both peptides exhibit potent antifungal activity (Fig.  21.55). This conserved structure/function ­ relationship indicates an ancient conserved defense ­strategy against microbial attack involving extracellular peptide production. Necrotrophic fungal, oomycete, and bacterial pathogens, as well as insects, trigger defense responses that are mediated by JA in combination with ethylene (Fig.  21.56). In Arabidopsis, the thionin gene Thi2.1 and plant defensin gene PDF1.2 are locally and systemically activated after infection with a necrotizing pathogen or exogenous application of ethylene and methyl jasmonate, but not after the leaves have been treated with SA. Coronatine is the toxin effector produced by the bacterium P. syringae to mimic JA and thereby assist infection (see Section  21.3.3). JA can become conjugated to different amino acids. In particular, JA-isoleucine is the JA derivative that is recognized by COI1, and is crucial for JA signaling. Most JA responses are mediated by the F‐box protein COI1 (CORONATINE INSENSITIVE 1). In Arabidopsis, JA‐insensitive coi1 mutants are more susceptible to necrotrophic pathogens, such as Alternaria brassicicola and Botrytis cinerea. A key regulator of JA responses in plant–pathogen interactions is the basic helix–loop–helix transcription factor AtMYC2. In addition, JAZ (JASMONATE ZIM‐domain‐containing) proteins in Arabidopsis negatively regulate JA signaling by binding AtMYC2. In the presence of JA‐Ile, these JAZ ­proteins interact with the COI1 protein, promoting ubiquitination of the JAZ proteins and leading to their degradation by the 26S proteasome, releasing AtMYC2 to activate the JA‐ responsive genes (Fig.  21.56). The crystal structure of the Arabidopsis JA receptor has revealed the JA responses are mediated by a complex formed between COI1 and JAZ proteins that requires the presence of both the signal JA‐Ile and inositol 6‐phosphate (see Chapter 17). JA and ethylene signaling molecules often work in combination during the defense response. Ethylene is frequently synthesized during both compatible and incompatible interactions. In Arabidopsis, the ethylene signal is perceived by five ethylene receptors, each of which contains a protein kinase domain (see Chapter 18). These receptors are negative regulators of ethylene signaling and interact with another negative regulator, CTR1, which negatively regulates downstream components of the ethylene‐signaling pathway and is inactivated by ethylene. This inactivation by ethylene abolishes negative regulation of the downstream ethylene‐signaling components EIN2 and EIN3. EIN2 is a membrane protein

β–3

Chapter 21  Responses to Plant Pathogens

PDF1-2 PR-1

C Inducing signals FIGURE 21.55  Plant defensins are small, cysteine‐rich peptides that accumulate at the periphery of the plant plasma membrane and are frequently found in dry plant seeds. They are induced during the defense response in growing plants. Plant defensin peptides are structurally and functionally similar to those produced by insects, birds, and mammals after a microbial attack. (A) The Drosophila drosomycin defensin peptide (left) resembles MeDef1 a 45 amino acid protein from the seed of Medicago sativa (right). (B) MsDef1 inhibits in vitro growth of a filamentous fungus, Fusarium graminearum, at micromolar concentrations (right). It prevents normal filamentous hyphal growth; instead, only a few short and highly swollen hyphae form. (C) In Arabidopsis, induction of the defensin PDF1‐2 gene transcript is regulated by defense signaling cascades that require ethylene and methyl jasmonate (MeJA). Application of either of these two signaling molecules does not induce the pathogenesis‐related PR‐1 gene, which is activated by salicylic acid (SA) or dichloroisonicotinic acid (INA). Source: (B) Sagaram et al (2011). PLoS One 6(4):e18550.

with a C‐terminal domain that is released by proteolysis upon activation of ethylene signaling, and leads to elevated levels of EIN3, a short‐lived transcription factor protein that accumulates in the nucleus after ethylene levels rise. EIN3 regulates the expression of many ethylene target genes, for example ERF1 that encode ethylene‐response element binding (AP2/ EREBP) transcription factors involved in defense against

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necrotrophs. Overexpression of ERF1 enhances resistance against Botrytis cinerea and increases susceptibility to the hemibiotroph P. syringae pv. tomato.

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JA/ET necrotroph inducible genes

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FIGURE 21.56  Comparison of the three main defense signaling p­ athways locally activated following pathogen infection or pest attack. The signaling pathways are activated by jasmonic acid‐­ isoleucine (JAILE) (left), a combination of JAILE and ethylene (ET) (center), and salicylic acid (SA) and reactive oxygen species (ROS) (right). The JAILE and combined JAILE/ET response pathways are important for defense against necrotrophic fungal, oomycete, and bacterial ­pathogens as well as insects. In contrast, the SA defense pathway is crucial for defense against biotrophic and hemibiotrophic ­pathogens. (Left) Most JA responses are mediated by the F‐box‐­ containing ­CORONATINE INSENSITIVE 1 protein (COI1). COI1 functions as a jasmonate receptor and forms Skp1/Cullin1/F‐box protein COI1 (SCFCOI1) complexes. In Arabidopsis, key regulators of JA that ­mediate defense activation are the basic helix‐loop‐helix ­transcription factor AtMYC2 and JAZ proteins (JASMONATE ZIM‐domain‐­containing), which bind to AtMYC2 and ­negatively regulate JA ­signaling. When JAIle levels rise, the JAZ proteins ­interact with the COI1 protein, promoting ubiquitination of the JAZ p­ roteins and their degradation by the 26S proteasome. The released A ­ tMYC2 ­transcription factor goes on to activate the JA‐responsive genes. (Center) The crosstalk and integration of the JAILE and ET signals in plant defense requires two members of the ETHYLENE RESPONSE FACTOR (ERF) family of transcription factors, namely ORA59, which contains an APETALA2/ERF domain and ERF p­ rotein 1 (ERF1). Cooperation between ORA59 and ERF1 activates the ­expression of several defense‐related genes, including plant defensins (for example PDF1.2) and basic chitinases (for example ChiB). (Right) SA mediated defense signaling can be further enhanced when reactive oxygen species levels are elevated. SA signaling requires the protein NONEXPRESSOR OF PATHOGENESIS‐RELATED GENES 1 (NPR1). NPR1 is an ankyrin repeat protein that exists in healthy plants in the cytoplasm in an oligomeric form. Upon pathogen ­infection, due to a change in redox status, monomers of NPR1 are released and migrate to the nucleus (see Section 21.8.2). Once inside the nucleus, NPR1 binds to TGA (TGACG‐motif binding) transcription factors, e­ nhancing their binding to SA‐ responsive promoters. This in turn leads to the activation of multiple defense‐related genes, including the pathogenesis-related proteins (for example PR1). In certain s­ ituations, the SA defense signaling pathway can suppress the JAILE/ET signaling pathway. Plant defense signaling can also be influenced by the plant hormones gibberellic acid and abscisic acid.

21.8.6  Salicylic acid and jasmonic acid/ethylene are mutually antagonistic signals and also intersect with other signaling pathways SA signaling triggers resistance to biotrophic and hemibiotrophic pathogens, whereas a combination of JA and ethylene activates defense against necrotrophic pathogens. These two pathways are mutually antagonistic: elevated SA levels and resistance to biotrophs are often correlated with increased necrotroph susceptibility, and elevated levels of resistance to necrotrophs are often correlated with enhanced susceptibility to biotrophs. This classical view requires revision because other hormones contribute to pathogen virulence and to plant defense. A complex crosstalk influences the levels of these induced hormones, and the eventual biological outcomes are influenced by both pathogen lifestyle and plant genetics. Gibberellic acid (GA) signaling is regulated by the DELLA proteins (see Chapter 18). In Arabidopsis, loss of DELLA proteins results in increased resistance to P. syringae pv. tomato, and this is associated with increased SA biosynthesis and signaling. DELLAs can also interact with the JA signaling pathway by interacting with the JAZ proteins that prevent AtMYC2 activation; DELLAs thus promote JA signaling. This explains why Gibberella fujikuroi, the rice necrotrophic fungal pathogen and cause of “foolish seedling” disease, makes gibberellin: removal of DELLAs attenuates JA signaling. The binding of GA to the DELLA proteins leads to their degradation and enables JAZ proteins to reimpose suppression of AtMYC2. This also explains the increased resistance of the DELLA mutants to biotrophs and hemibiotrophs. In summary, plant signaling molecules such as SA, JA, ­ethylene, and GA can influence the balance and level of ­activation of different facets of the plant defense response via complex mechanisms that remain an active topic of investigation. It is also clear from pathogen genome sequencing projects and targeted biochemical studies that certain pathogenic species produce some of these plant hormones and can, therefore, influence this crosstalk.

21.9  Plant gene silencing confers virus resistance, tolerance, and attenuation Neither the HR nor the SAR is responsible for limiting the systemic spread of viruses through the plant vascular system. Thus, other defense mechanisms must be responsible for controlling virus infections.

Chapter 21  Responses to Plant Pathogens

21.9.1  The systemic long-distance spread of viruses is restricted Arabidopsis mutants have provided genetic evidence for the involvement of specific host genes in controlling vascular movement of a tobamovirus or a potyvirus. At least three loci, RTM1 (restricted TEV movement 1), RTM2, and RTM3, are required for restriction of long‐distance tobacco etch virus (TEV) movement in the Arabidopsis ecotype Col‐0. The RTM1 protein is similar to the lectin jacalin. The RTM2 protein contains several domains, including an N‐terminal region with similarity to plant small heat shock proteins and a C‐terminus that includes a predicted transmembrane ­spanning domain. These two proteins of the RTM system function within the phloem sieve elements to restrict long‐ distance movement of TEV.

21.9.2  RNA silencing is important for resistance to many virus species Plants infected with some viruses can recover and produce new shoots that do not show viral disease symptoms (Fig.  21.57). This is because plants can attenuate the ­accumulation of many virus species by a mechanism called RNA silencing (see Chapter  6). Although plant viruses are highly diverse in sequence, morphology, genome organization, protein production, and host range, all virus species  accumulate viral RNA as a part of their life cycle. Accumulation of viral RNA, particularly double‐stranded viral RNA (dsRNA), provides the signal that activates the silencing mechanism. The formation of dsRNA can be part of the normal virus replication process, the result of an inverted repeat sequence within the single‐strand RNA (ssRNA) that folds into a hairpin structure, or through the presence of a complementary RNA product from overlapping bidirectional transcription. RNA silencing is an effective defense mechanism that has three main characteristics. First, it is homology‐dependent and, therefore, specific to viral RNA. It is driven by small inhibitory RNAs (siRNAs) derived from a double‐stranded form of the viral RNA; host‐encoded RNAs are not affected by the viral RNA‐induced silencing mechanism. Second, there is enormous potential for amplification, because siRNAs can serve as primers for production of secondary long viral dsRNAs from ssRNA templates by plant‐encoded RNA‐ dependent RNA polymerases (see Chapter 6). Third, siRNAs also function as a mobile signal that moves ahead of or with the advancing virus. This signal ensures the virus is not able to escape the effects of silencing by movement between cells or in the phloem. To counter this efficient RNA silencing mechanism, many plant viruses encode proteins that act as suppressors of RNA silencing. Silencing suppressors from different virus species are mostly unrelated in sequence or structure. Under the

FIGURE 21.57  RNA silencing is a general antiviral plant defense mechanism. The bottom leaves of a tobacco plant were inoculated with tobacco ringspot virus (TRSV). By 23 days post inoculation, strong ringspot symptoms developed on the lower leaves; there is, however, a gradual decline in the development of ringspot symptoms on the upper leaves, and the top leaves appear normal and are virus free. The virus causing the initial symptoms activated RNA silencing, which inhibited spread of the infection into the upper leaves and also caused them to be immune to secondary infection by the same virus. Source: Wingard (1928). J. Agric. Res. 37:127–153.

strong selection pressure of RNA silencing, several different mechanisms appear to have evolved independently that fulfill the same function. This is an example of convergent evolution. Suppressor proteins often provide the virus with additional, unrelated roles in virus replication, and their ability to suppress RNA silencing has evolved as an additional feature. The P19 protein of tomato bushy stunt virus binds siRNAs, and this prevents formation of an active RISC (see Chapter 6). In contrast, the 2b protein of ­cucumber mosaic virus appears to inhibit the endonuclease activity of Argonaute (see

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EFR EF-Tu

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FIGURE 21.58  Transfer of a pattern‐recognition immune (PTI) receptor to a crop confers resistance to several bacterial pathogens. (A) ­Engineering broad‐spectrum resistance in tomato, a member of the Solanaceae, by transferring a pattern‐recognition receptor from Arabidopsis (family Brassicaceae). Elongation factor Tu receptor (EFR; purple) is absent from the solanaceous species N. benthamiana and tomato, and these plants are normally susceptible to infection by phytopathogenic bacterial from different genera. Transgenic expression of Arabidopsis EFR increases resistance to these bacterial pathogens, presumably by activating a signaling cascade that confers resistance to a range of bacteria expressing its cognate P/MAMP, elongation factor Tu (EF‐Tu; orange). This interfamily transfer of EFR‐mediated disease resistance suggests N. benthamiana and tomato contain the components necessary for EFR signaling other than the receptor. MAPK, mitogen‐ activated protein kinase. (B) Wild‐type (EFR–; left) and transgenic plants expressing EFR (EFR+; middle) tomato plants infected with a virulent isolate of Ralstonia solanacearum. The time course of disease symptom formation (right) reveals the dramatic reduction in disease severity in the transgenic ERF plants. Source: (B) Lacombe et al. (2010). Nat. Biotechnol. 28:365–369.

Chapter 6). The formation of some virus disease symptoms, especially plant stunting and abnormal development, may be the result of interference by viral suppressor proteins in siRNA and microRNA processing.

21.10  Control of plant pathogens by genetic engineering Exciting opportunities exist to use plant PRRs, plant R and other genes, and pathogen Avr and effector genes to improve crop disease resistance. In addition, processes required for

pathogenesis can be targeted for novel plant‐specified inhibitory mechanisms.

21.10.1  Pattern‐recognition receptors can elevate pathogen resistance in plants Pattern‐recognition receptors (PRRs) detect microbes by recognizing conserved pathogen/microbe‐associated molecular patterns (P/MAMPs, see Section  21.5). However, different plant families carry distinct repertoires of PRRs that perceive distinct P/MAMPs. The Arabidopsis PRR protein EFR perceives the acetylated N‐terminus of the bacterial elongation factor‐Tu (EF‐Tu) (Section 21.5.1), but bacterial EF‐Tu is not perceived in the

Chapter 21  Responses to Plant Pathogens

Traditional breeding method

Transgenic approach

P1 (Parent 1) Resistant

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C

A, B, C

Backcross 7 is resistant but otherwise has predominantly P2 genes

Multiple resistance in a single transgenic line

×

Susceptible cultivar for improvement

Fewer backcrossing generations required before a useful new line is identified

FIGURE 21.59  Traditional breeding compared with transformation technology approaches to introgress desirable genes into commercial crop plants. In a traditional breeding method (left), as much as 0.4% of the genome complement from each donor parent can remain in the seventh backcross generation along with the R gene of interest (originally from parent 1). In the transgene transformation approach, multiple R genes from several initial sources are first assembled into a single Ti plasmid. After T‐DNA integration into the plant genome, these R genes co‐segregate in all subsequent breeding steps, greatly simplifying the subsequent backcrossing program for introducing multiple new traits into a cultivar. When the transgenic transformation approach is used, the entire sequence of the introduced DNA is known, whereas in the traditional breeding program, neither the total extent of the introgressed DNA nor its sequence identity is known.

Solanaceae. Transfer of EFR from Arabidopsis into tobacco (Nicotiana benthamiana) and tomato (Solanum lycopersicum) confers responsiveness to EF‐Tu and elevates resistance to phytopathogenic bacteria from different genera, including A. tumefaciens, P. syringae, Ralstonia solanacearum, and Xanthomonas perforans (Fig.  21.58). These results suggest that extending the P/MAMP recognition repertoire of crop plants can elevate their pathogen resistance.

21.10.2  R genes can confer broad‐spectrum resistance Traditionally plant breeders have introgressed, via lengthy breeding programs, novel R genes from wild relatives of crop species into elite commercial cultivars to control diseases

(Fig.  21.59 and also see Fig.  21.39, Section  21.6.1). During breeding, the R gene must be recombined away from any linked unwanted alleles of other genes that reduce crop performance (linkage drag). The availability of cloned R genes opens the possibility of their direct transfer into elite lines by genetic transformation (see Box 21.2). Genetic transformation enables transfer of several different R genes that may be effective against a single pathogen species in a “cassette” of linked genes that cannot be separated by recombination (Fig. 21.59). This should slow pathogen evolution to overcome resistance, because the various R genes could only be overcome if, in a single pathogen isolate, several effector genes were simultaneously mutated. The introduction of R genes by transformation also means that interplant species fertility barriers can be crossed, thereby increasing the repertoire of plant species from which useful novel resistance sources can be obtained.

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B FIGURE 21.60  Comparison of severity of late blight disease caused by Phytophthora ­infestans in Rpi‐vnt1.1‐transgenic and ­non‐transgenic var. Desiree potato plants. (A) Percentage of late blight infected leaf tissue during the vegetative growing stage. (B) ­Photograph of the control Desiree (left) and transgenic (right) plants almost one month after first symptoms of infection on Desiree plants were observed. No symptoms of late blight were present on transgenic plants during the growing season or as the crop matured. Source: (B) Jones et al. (2014). Phil. Trans. R. Soc. Ser. B 369:doi:10.1098/rstb.2013.0087.

R genes that confer resistance to all or most races of a pathogen species are attractive candidates for interspecies transfer, because they may confer the same broad‐spectrum resistance as occurs in their native genomes. For example, most accessions of the wild diploid potato species Solanum bulbocastanum are resistant to all known races of Phytophthora infestans. Using map‐based cloning the resistance gene RB/ Rpi‐blb1 was isolated. Transgenic cultivated potato plants (Solanum tuberosum) expressing the RB gene show a high level of resistance to nearly all P. infestans isolates, including a European “super race” that can overcome all 11 known R genes in cultivated potato. Two other R genes conferring broad‐spectrum resistance to P. infestans have been isolated: Rpi‐blb2 from S. bulbocastanum and Rpi‐vnt1.1 from S. ven­ turii. Transgenic cultivated potato lines expressing both ­Rpi‐blb1 and Rpi‐blb2 provided durable control of late blight disease in several years of field trials in the United Kingdom and mainland Europe (Fig. 21.60). Isolating R genes from wild relatives and introducing them into elite lines, whether by breeding or transformation, is laborious; therefore, it is important to prioritize those R genes that are most likely to be durable, and analysis of pathogen effector complements can help. If an R gene recognizes an

effector that is conserved in all known races of a pathogen, it is more likely to be durable than an R gene that recognizes an effector that is absent from many races of the pathogen. Some effectors are essential for full virulence of the pathogen (see Box 21.3). Emerging knowledge of differences in the effector complement between pathogen races can thus be helpful for  choosing R genes for prioritization. Already useful in the  potato/P. infestans interaction, these approaches are deployed to identify durable resistance genes against the Ug99‐derived races of the wheat stem rust fungus Puccinia graminis f. sp. tritici, which threatens wheat harvests in Africa and Asia (see Box 21.3 and Box 21.1). Tomato and banana are susceptible to the closely related leaf mold (Cladosporium ful­ vum) and black Sigatoka (Mycosphaerella fijiensis) diseases, respectively. Homologs of C. fulvum Avr4 and Ecp2 effectors are present in M. fijiensis. Since tomato Cf‐4 confers recognition of Avr4 homologs from both pathogens, it may prove useful for M. fijiensis resistance in banana. To control the wheat glume blotch pathogen Stagonospora nodorum, the ToxA and other related toxins recovered from fungal culture filtrates have been used globally to screen both existing commercial elite wheat germplasm and potential donor breeding lines for the lack of a necrotic response. This

Chapter 21  Responses to Plant Pathogens

BOX 21.3

Effector‐guided R gene deployment Effector‐guided R gene deployment is a disease control strategy based on the identification of core pathogen effectors that lack sequence polymorphism. It is currently being used in potato (Solanum tuberosum) to mitigate the impact of the late blight pathogen Phytophthora infestans. This pathogen undergoes major population shifts in agricultural systems via the successive emergence and migration of asexual lineages (A). A new aggressive lineage in the European P. infestans population termed 13_A2 is rapidly displacing other older lineages. Comparative whole-genome and transcriptome sequencing (see Chapter 9) of P. infestans pathogen races has revealed the core RXLR effector repertoire found in isolates from the new aggressive lineage of P. infestans 13_A2 and two older European P. infestans isolates. All three P. infestans isolates carry intact and in planta induced Avrblb1, Avrblb2, and Avrvnt1 effector genes that trigger resistance in potato lines carrying the

corresponding disease resistance genes Rpi‐blb1, Rpi‐ blb2, and Rpi‐vnt1.1 (B). R gene stacking can be achieved by conventional plant breeding with marker‐assisted selection or by plant genetic engineering with T‐DNA ­cassettes containing the desired R gene combinations. This core effector‐guided approach helps to exclude any R genes from the elite potato germplasm improvement scheme that are already defeated by the pathogen because the cognate core effectors contain sequence polymorphisms that evade plant recognition. (A) Disease‐causing abilities of historic and current Phytophthora infestans (Pi) isolates. (B) The conserved core effectors Avrblb1, Avrblb2 and Avrvnt1 are present in all Pi isolates. These are recognized when the corresponding R genes Rpi‐blb1, Rpi‐blb2, and Rpi‐vnt1.1 are expressed in the host plant. Source: Cooke et al. (2010). PloS Pathog. 8(10):e1002940. Emerging aggressive isolates from the 13_A2 lineage

Older, low aggression isolates 2_A1

6_A1

8_A1

10_A2

13_A2

A Rpi-blb1

Rpi-blb2

Rpi-vnt1.1

R gene present, no disease No R gene, full disease B

ensures that the corresponding Tsn locus, which confers S. nodorum disease susceptibility, is eliminated from all commercial wheat breeding programs. Some R genes confer resistance to all races of a single pathogenic species, for example the recessive barley mlo resistance gene. Other R genes confer resistance to multiple unrelated pathogenic species, for example the tomato Mi gene, which confers both nematode and whitefly and psyllid resistance.

T30-4 06_3928A

Different Pi isolates

Certain R genes confer resistance to different but taxonomically closely related fungal species, for example the wheat Lr34 gene, which confers resistance to leaf rust and stripe rust fungal pathogens (see Section 21.7.1). By analyzing the increasing amount of genome and transcriptome sequence information available for crop species, highly homologous R gene sequences can be isolated and tested for their ability to confer broad‐ spectrum resistance spectrum in different crop plant species.

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Part V  PLANT ENVIRONMENT AND AGRICULTURE FIGURE 21.61  TALEN technology has been used to edit a specific pathogen susceptibility (S) gene in rice to thwart the virulence ­strategy of Xanthomonas oryzae pv. oryzae (Xoo) and engineer ­heritable genome modifications for resistance to bacterial blight. The rice bacterial blight susceptibility gene Os11N3 (also called OsSWEET14) was targeted for TALEN‐ based disruption. (A) The Os11N3 promoter contains an effector‐binding element (EBE) for AvrXa7, overlapping with another EBE for PthXo3 and with the TATA box. Two pairs of designer TALENs (pair 1 and pair 2) were independently deployed to induce mutations in these ­overlapping EBEs of the Os11N3 promoter and thereby interfere with the ­virulence function of AvrXa7 and PthXo3, but not the ­developmental function of Os11N3. Each designer TALEN contained 24 repeat units for recognition of a specific set of 24 contiguous nucleotides at the target sites. (B) Severity of disease damage to wild‐type and ­transgenic rice plants caused by AvrXa7‐, PthXo3‐, and PthXo1‐dependent Xoo strains. Lengths of leaf lesions in wild‐type plants (ck1), a nontransgenic plant (ck2) and transgenic plants ­homozygous for Os11N3 promoter mutations of 6 bp (–6b), 9 bp (–9a), 15 bp (–15) and 4 bp (–4d) deletions, respectively, measured 14 days post inoculation with different Xoo strains expressing three different TAL effectors. (C) Resistance phenotype displayed to Xoo (avrXa7) by two transgenic rice plants (carrying either the –4d or –9a deletion constructs) compared with the disease susceptibility phenotype of a wild‐type plant (wt). Leaves were inoculated at the left end and the extent of visible disease symptom formation is indicated by the red arrowheads. Source: Ting Li et al. (2012). Nat. Biotechnol. 30(5):390.

Os11N3

ACTATATAAACCCCCTCCAACCAGGTGCTAAGCTC

AvrXa7 EBE PthXo3 EBE TATA box

ATAAACCCCCTCCAACCAGGTGCTAA ATATAAACCCCCTCCAACCAGGTGCTAAG TATAAA Pair 1

Os11N3

Chr 11 Pair 2

A

Exon 1

18

AvrXa7

16

PthXo3

PthXo1

14 Lesion length (cm)

1048

12 10 8 6 4 2

B

0

ck1

ck2

–6b

–9a

–15

–4d –4d

–9a wt

C

Many race‐specific R proteins are temperature‐sensitive. For example, N gene‐mediated tobacco mosaic virus (TMV) resistance fails at temperatures above 30°C and permits the  virus to spread systemically throughout the plant. With climate changes now being experienced in many global regions, R genes that fail at higher temperatures may give inconsistent pathogen control. A lack of correct R protein folding or incorrect cellular localization may explain this phenomenon.

21.10.3  TAL effectors can be used for engineering resistance Transcription activator‐like (TAL) effectors of Xanthomonas oryzae pv. oryzae (Xoo) contribute to pathogen virulence by transcriptionally activating specific rice disease‐susceptibility (S) genes (see Section  21.1.8). TAL effector nucleases (TALENs) are fusion proteins that combine the DNA recognition repeats of native or customized Xoo TAL effectors and the DNA cleavage domains of FokI nuclease. These TALENs are useful for precise genome engineering.

0

1

2

3

4

5

6

8 7 cm

9

10

11 12

13 14 15

For example, to engineer rice plants (Oryza sativa) resistant to Xanthomonas bacterial infection, the promoter of the rice bacterial blight susceptibility gene Os11N3 (also called OsSWEET14) was targeted for TALEN‐based disruption. This gene encodes a member of the SWEET sucrose‐efflux transporter family. The Xoo TAL effectors AvrXa7 or PthXo3 activate this gene and thus divert sugars from the plant cell to the bacteria. The promoter of Os11N3 contains binding sites for both TAL effectors near the TATA box. TALENs were designed to mutate DNA sequences required for TAL effector binding, but not the development function of Os11N3. The modified Os11N3 gene was no longer inducible when the rice plants were infected by pathogenic Xoo strains that could deliver either the AvrXa7 or PthXo3 TAL effectors into the rice cells. The stable TALEN‐­modified plants showed strong (though recessive) resistance to infection by the AvrXa7‐ or PthXo3‐dependent Xoo strains, but  not the PthXo1‐dependent pathogenic Xoo strain (Fig. 21.61). So far, natural polymorphisms in the Os11N3 gene that would prevent induction by AvrXa7‐ and PthXo3‐dependent Xoo strains and also confer disease resistance have not been identified in rice germplasm. By using TALENs to edit a rice gene

Chapter 21  Responses to Plant Pathogens

A

B

FIGURE 21.62  Papaya ringspot virus (PRSV) is a limiting factor in the production of papaya worldwide. By constitutively overexpressing the coat protein of this virus in papaya, several highly resistant varieties are now in commercial use in Hawaii and elsewhere. (A) Transgenic papaya line (left) showing resistance to PRSV compared to infected non‐transgenic papaya (right). (B) Aerial view of transgenic papaya test field in Hawaii, showing a block of healthy transgenic papaya plants surrounded by severely infected non‐transgenic papaya. Source: Gonsalves, AgBioForum 7(1 & 2), http://www.agbioforum.org/v7n12/v7n12a07-gonsalves.htm.

required for Xoo susceptibility, desirable alleles conferring resistance can be obtained that can then be used in rice breeding programs.

21.10.4  Virus sequences can induce resistance to virus infection Virus sequence‐induced resistance was based on the concept that a transgenic plant engineered to express viral proteins could interfere with the normal pathogenicity program of the virus. Coat protein (CP)‐mediated resistance was proposed to operate through high CP levels, which inhibit disassembly of the virus particles in the initially infected cells, whereas the overexpression of a nonfunctional mutant movement protein (MP) was proposed to lead to competition for plasmodesmatal binding sites between the nonfunctional and the wild‐ type MP of the infecting virus. However, molecular analysis of transgenic CP or MP plants has revealed that resistance is RNA‐mediated and sequence‐specific, and likely functions by post‐transcriptional silencing of viral gene expression (see Section 21.9.2). CP sequence‐induced virus resistance was one of the early success stories of plant genetic engineering and led to the development of virus‐resistant papaya and squash cultivars for commercial production. For example, papaya crops in

Hawaii derived from transgenic lines carrying the CP gene of papaya ringspot virus (PRSV) provide effective control of this virus (Fig. 21.62).

21.10.5  Genetically engineered disease control is a knowledge‐based approach The transgenic disease control options described above have each taken immediate advantage of fundamental discoveries that had significantly advanced our overall understanding of host–pathogen interactions. In each example, a target of potential pathogen vulnerability was exposed through the acquisition of new knowledge, various intervention options were proposed and tested in the laboratory, growth room, or glasshouse, and only the best solutions went on to rigorous field testing and, finally, commercial release. In addition, many of the single gene‐engineered disease control solutions that failed to make the grade, either in early testing or later when tested under field conditions, have provided considerable useful and novel insight into the importance of other plant components and information on the ways in which pathogens can rapidly evolve to overcome plant defenses.

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Summary

P

lants are resistant to most plant pests and pathogens. Every plant cell can defend itself from attacking pathogenic microorganisms and invertebrates. Some defenses, such as antimicrobial secondary metabolites, are constitutive and located in specific cellular compartments ready to be released upon cell damage. Other defense responses, such as those induced by pathogen invasion, require detection of the pathogen by the plant. Defense activation is correlated with rapid activation of defense‐related genes and often culminates in the hypersensitive response that causes localized cell death to impair pathogen spread. Plant resistance to pathogens can be mediated by dominant resistance (R) genes in plants that are complementary to avirulence (Avr) genes in pathogens. Avr effector proteins exhibit extensive sequence diversity, and their function in the pathogen and in the host plant is still poorly understood. In contrast, plant R proteins are strikingly similar in structure, sharing such motifs as leucine rich repeats (LRRs), a nucleotide binding or a serine/ threonine protein kinase domain, among others. R proteins both detect pathogens and initiate signal transduction to activate defense mechanisms. In addition, R loci/ genes can evolve new R gene specificities to keep pace with the evolution of virulence in pathogen populations.

By investigating the genomes of numerous pathogens and pests, the full repertoires of effector genes thought to function either in the suppression or inactivation of plant defenses have been predicted and their specific roles experimentally tested. Plant defense reactions involve complex biochemical pathways and multiple signal molecules, including ROS, NO, SA, JA, and ethylene, to provoke the induction of proteins, secondary metabolites, and often cell wall fortification reactions—both at the infection site and systemically throughout the attacked plant. Specialist defenses against plant viruses include post‐ transcriptional gene silencing (PTGS); those against insects involve proteinase inhibitor proteins. Many aspects of induced plant defense appear to be conserved in other eukaryotes, perhaps indicating the existence of an ancient defense strategy that evolved early against pathogen attacks. The genetic engineering of plants has started to achieve broad‐spectrum and durable pest and pathogen control in crops. However, a better understanding of the factors and mechanisms involved in plant–pathogen interactions is still needed and better resistance strategies still need to be deployed to reduce crop losses.

Responses to Abiotic Stress

22

Kazuo Shinozaki, Matsuo Uemura, Julia Bailey‐Serres, Elizabeth A. Bray, and Elizabeth Weretilnyk

Introduction Plants are frequently exposed to environmental stress, ­external conditions that adversely affect growth, development, or ­ productivity. Stress can be biotic, imposed by other  organisms (Chapter  21), or abiotic, arising from an excess or deficit in the physical or chemical environment. Environmental conditions that cause damage to plants include flood, drought, high or low temperatures, excessive soil salinity, inadequate mineral nutrients, and excess or insufficient light. Phytotoxic compounds, such as ozone, can also damage plant tissues. Stress triggers a wide range of plant responses, from ­alterations in gene expression and cellular metabolism to changes in growth rate and crop yield. The duration, severity, and rate at which a stress is imposed all influence how a plant responds. Several adverse conditions in combination can elicit a response that is different from that caused by a single stress. A response can be triggered directly by a stress, such as drought, or result from stress‐induced injury, such as loss of membrane integrity. Moreover, resistance and sensitivity to stress vary according to species, genotype, developmental stage, and organ or tissue type (Fig. 22.1). The molecular mechanisms underlying plant responses to abiotic stress, including changes in gene expression and ­cellular signal transduction, have been analyzed ­extensively, and various transcription factors and signaling molecules are now known to play important roles in cellular homeo-

stasis under stress conditions. The functions of these ­factors are described in this chapter.

22.1  Plant responses to abiotic stress 22.1.1  Plant stress diminishes crop yield As human populations grow, agricultural systems must feed more people while competing with urban development for premium arable land. This increasing demand, coupled with shrinking resources, has fueled research into elucidating mechanisms by which plants respond to stress and manipulating these mechanisms to enhance plant productivity in suboptimal environments. The impact of the environment on plant productivity becomes apparent when record yields are compared to ­average yields. If record yields are assumed to represent plant growth under ideal conditions, then the losses associated with biotic and abiotic stress can reduce average productivity by 65–87%, depending on the crop. Successful application of biotechnological and classical breeding techniques may lead to stress‐tolerant crop plants that enhance world food s­ upplies and provide considerable economic benefit. These crops may promote survival during periods of intense or prolonged stress or maintain high plant productivity under conditions of moderate environmental stress.

Biochemistry & Molecular Biology of Plants, Second Edition. Edited by Bob B. Buchanan, Wilhelm Gruissem, and Russell L. Jones. © 2015 John Wiley & Sons, Ltd. Published 2015 by John Wiley & Sons, Ltd. Companion website: www.wiley.com/go/buchanan/biochem

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Part V  PLANT ENVIRONMENT AND AGRICULTURE FIGURE 22.1  Many factors determine how plants respond to environmental stress: the genotype and developmental conditions of the plant, the d ­ uration and severity of the stress, the number of times the plant is subjected to stress, and any additive or ­synergistic effects of multiple stressors. Plants respond to stress through a variety of mechanisms. Failure to compensate for severe stress can result in plant death.

Environmental stress

Stress characteristics

Severity

Duration

Number of exposures

Combination of stresses

Organ or tissue in question Plant characteristics Genotype

Stage of development

Response

Resistance

Susceptibility

Result

Survival and growth

Death

22.1.2  Resistance mechanisms allow organisms to avoid or tolerate stress Stress resistance mechanisms can be grouped into two ­general categories: avoidance mechanisms, which prevent exposure to stress, and tolerance mechanisms, which enable the plant  to  withstand stress (Fig.  22.2). Many desert plants are xerophytic—tolerant of water deficits—because their ­ ­morphological characteristics facilitate survival under arid conditions. A drought‐avoidance mechanism—deep roots— provides another class of plants (phreatophytes) with improved access to groundwater. In contrast, desert ephemerals evade drought by germinating and completing their life cycles while adequate water is available. Sunken stomata, light‐reflective spines, and deep roots are among the ­constitutive, genotypically determined traits for stress resistance that are expressed whether the plants are stressed or not. They constitute adaptations, evolutionary improvements that enhance the fitness of a population of organisms. Other resistance mechanisms are achieved through acclimation, the adjustment of individual organisms in response to changing environmental factors. During acclimation, an organism alters its homeostasis, its steady‐state physiology, to accommodate shifts in its external environment. A period of acclimation before stress is encountered may confer ­resistance to an otherwise vulnerable plant. For example, ­during the summer, trees in the northern latitudes cannot withstand freezing but many can acclimate and eventually

withstand freezing in winter. Whether based on acclimation or adaptation, successful mechanisms for stress resistance support ­survival under lethal conditions or maintain productivity under circumstances that impair crop yields.

22.1.3  Gene expression patterns often change in response to stress Stress‐induced changes in metabolism and development can often be attributed to altered patterns of gene expression. A stress response is initiated when a plant recognizes a stress at the cellular level. Stress recognition activates signal ­transduction pathways that transmit information within individual cells and throughout the plant (Fig. 22.3). Ultimately, changes in gene expression are integrated into a response by  the whole plant that modifies growth and development and that may even influence reproductive capabilities. The ­duration and severity of the stress dictate the scale and timing of the response. Little is known about how plants recognize stress. Our best insights come from yeast and bacterial proteins that initiate signal transduction in response to abiotic stress, such as low osmotic potential. Though plants likely contain similar ­proteins, comparable functions have not yet been demonstrated. The intricate signaling pathways that are assumed to participate in altering plant gene expression in response to stress have yet to be elucidated; however, there is considerable

Chapter 22  Responses to Abiotic Stress Saguaro

Honey mesquite

Abiotic stress

Spinach

Acclimation Mohave desert star

Black spruce

Resistance • Stress avoidance • Stress tolerance

A

B

FIGURE 22.2  Stress resistance can involve tolerating or avoiding the stressful condition. (A) Some resistance mechanisms are constitutive and active before exposure to stress. In other cases, plants exposed to stress alter their physiology in response, thereby acclimating themselves to an u ­ nfavorable environment. (B) Examples of constitutive mechanisms of drought resistance include the succulent, photosynthetic stem of the saguaro cactus Cereus giganteus, a drought‐tolerant species; the deep roots of the honey mesquite Prosopis glandulosa var. glandulosa, a drought‐avoiding species; and the wet‐season life cycle of the Mohave desert star, Monoptilon bellioides. Examples of acclimation mechanisms include osmotic adjustment (see Section 22.2.3) in plants such as spinach (Spinacia oleracea) and freezing tolerance (see Section 22.4) in cold‐hardy trees such as black spruce, Picea mariana Mill. Source: Epple & Epple (1995). A Field Guide to the Plants of Arizona. Falcon Press Publishing, Helena, MT.

Ozone

Stress recognition

Extreme temperatures

Signal transduction

Flooding Drought

FIGURE 22.3  Plants respond to stress both as collections of cells and as whole ­organisms. Stressors constitute ­environmental signals that are received and recognized by the plant. After stress ­recognition, the signal is communicated within cells and throughout the plant. Transduction of environmental signals typically results in altered gene expression at the cellular level, which in turn can influence metabolism and development of the whole plant.

Physiological and developmental event Salt Altered cellular metabolism

evidence that regulation of plant stress responses involves hormones—especially abscisic acid (ABA), jasmonic acid (JA), and ethylene—and secondary messengers such as Ca2+ (see Chapters 17 and 18). In response to stress, some genes show increases in ­expression, while others are repressed. Protein products of

stress‐induced genes often accumulate in response to ­unfavorable conditions. The functions of these proteins and the mechanisms that regulate their expression are currently a central topic of research in stress physiology. Although most studies have focused on transcriptional activation of gene expression, accumulation of gene products may also be

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i­nfluenced by post‐transcriptional regulatory mechanisms that increase levels of specific protein‐coding mRNAs or ­noncoding regulatory RNAs, enhance translation, stabilize proteins, and alter protein activity by various types of modifications. Using molecular genetic techniques, researchers have ­dissected plant responses associated with exposure to specific abiotic stress. The stresses addressed in this chapter include water deficit, low and high temperatures, hypoxia, and environmental oxidants. Other sources of abiotic stress ­ described elsewhere in this text include nutrient deficiencies and toxic concentrations of aluminum (see Chapter  23), ­cadmium (see Chapter 16) or other metals. Plant responses to these disparate stresses can incorporate similar or even ­overlapping strategies, such as the activation or repression of gene expression, production of proteins or compatible ­solutes, activation of transport activity, etc. In addition, plants are often exposed to simultaneous or sequential abiotic stresses, such as heat and drought, or flooding followed by drought. Interactions between different types of stress, whether ­synergistic or antagonistic, likely influence the success in coordinating the activity of many genes that together increase abiotic stress survival in crops.

22.2  Physiological and cellular responses to water deficit 22.2.1  Many environmental conditions can lead to water deficit Water‐related stress results if the environment contains excess water or if the quantity or quality of water available is insufficient to meet basic needs. Though periods of little or no rainfall can lead to drought, water deficit can also occur in environments in which water is not limited. In saline habitats, for example, high salt concentrations make it more difficult for plant roots to extract water from the environment. In addition, low temperatures can also result in water stress; exposure to freezing temperatures, for example, can lead to cellular dehydration as water leaves the cell and forms ice crystals in intercellular spaces (see Section 22.4). Occasionally, even plants that have been watered well show periodic signs of water deficit, such as a transient loss of turgor at midday. In this case, wilting indicates that transpirational water loss has exceeded the rate of water absorption. Many factors can affect the response of a plant to water deficit, including the duration of water deficiency, rate of onset, and possible acclimation to water stress by previous exposure. The response of a plant to water deficit can be complex and incorporate many of the strategies and mechanisms of tolerance and resistance used in response to other types of stress; therefore it is addressed first.

22.2.2  Two parameters that describe the water status of plants are water potential and relative water content Water can be described thermodynamically in terms of its free energy content, also known as its chemical potential. Plant physiologists use a related parameter, water potential (Ψw; see Chapter  15, Box  15.1, for a more quantitative ­discussion of this topic). Such measurements can be used to evaluate the extent to which a cell, organ, or whole plant is “hydrated.” Equation 22.1: Water potential w

s

p

g

m

The Ψw of a plant equals the sum of various component potentials. Solute potential, Ψs, is dictated by the number of particles dissolved in water. Water potential decreases as solute concentration increases. Pressure potential, Ψp, ­ reflects physical forces exerted on water by its environment. When water is subjected to negative pressure (tension), Ψp is less than 0 MPa (megapascals) and Ψw is diminished. (Note that water potential is typically defined in units of pressure rather than energy.) In contrast, water potential is increased by positive pressure (turgor, Ψp > 0 MPa). Gravitational potential, Ψg, can have a substantial effect when water is transported over vertical distances greater than 5–10 meters, but this term can be omitted when describing transport between cells or within small plants. A fourth factor, matric potential, Ψm, accounts for how solid surfaces (e.g., cell walls and colloids) interact with water and depress Ψw. Because Ψm values are small and difficult to measure, however, its impact on plant water potential is usually ignored. For conditions under which Ψg and Ψm are insignificant, the water potential equation is frequently simplified as follows: Equation 22.2: Water potential (simplified) w

s

p

Water potential can be used to predict the movement of liquid water into or out of a plant cell. The difference in water ­potential across a membrane determines the direction of flow: water moves spontaneously from regions of high water potential to adjoining regions of low water potential. Physiological or metabolic changes detected in water‐ stressed plants are not always correlated with changes in plant Ψw measurements. To address these issues, a second ­parameter frequently used to assess water status—relative water content (RWC)—is often reported in conjunction with plant Ψw measurements. Equation 22.3: Relative water content RWC

fresh wt. dry wt.

turgid wt. dry wt.

100

Chapter 22  Responses to Abiotic Stress

Ψp = +0.5 MPa

FIGURE 22.4  Osmotic adjustment occurs when the concentrations of solutes within a plant cell increase to maintain positive turgor pressure within the cell. As the cell actively accumulates solutes, Ψs drops, promoting the flow of water into the cell. In cells that fail to adjust osmotically, solutes are concentrated passively, but turgor is lost.

0 MPa –1.2 MPa = Ψs Ψw = –1.2 MPa Ψp =

Ψs = –2.0 MPa Ψw = –1.5 MPa

Water deficit

Soil Ψw = –1.2 MPa

Osmotic adjustment

No osmotic adjustment

When water uptake by roots closely matches water loss by leaves, the RWC of transpiring leaves typically ranges from 85% to 95%. If the RWC for an organ drops below a critical value, tissue death follows. The value for critical RWC ­varies among species and tissue types, but it is frequently less than 50%.

22.2.3  Osmotic adjustment is a biochemical mechanism that helps plants acclimate to dry or saline soil While some plants are highly sensitive to water stress and wilt (dehydrate), others can endure dry or saline conditions ­without evident loss of turgor. To extract water from the soil, a plant root must establish a water potential gradient so that water flows toward the root surface from the soil (i.e., the water potential must be lower in the root than in the surrounding soil). Many drought‐tolerant plants regulate ­ their solute potential (Ψs) to compensate for transient or extended periods of water stress. This process, called osmotic adjustment, results from a net increase in the number of ­solute particles in the plant cell. The osmolalities achieved through osmotic adjustment exceed those from passive ­concentration of solutes by dehydration. By decreasing plant Ψs, osmotic adjustment can drive root Ψw to values lower than soil Ψw, thereby allowing water to move from soil to plant down a potential gradient (Fig. 22.4). Osmotic adjustment is believed to play a critical role in helping plants acclimate to conditions of drought or high salinity.

22.2.4  Compatible solutes and aquaporins play roles in osmotic adjustment One mechanism for osmotic adjustment is the accumulation of compatible solutes, or compatible osmolytes, a ­chemically diverse and highly soluble group of organic compounds that do not interfere with cellular metabolism, even at high ­concentrations (Fig. 22.5). Synthesis and accumulation of these compounds are widespread in plants, but their distribution varies among species. For example, whereas one such compatible solute, the amino acid proline, is accumulated by a taxonomically diverse set of plants, the quaternary ammonium compound β‐alanine betaine appears to be confined to representatives of a few ­genera of Plumbaginaceae (Leadwort). Stress can trigger the irreversible synthesis of these compounds (for example, ­glycine betaine; see Section 22.3.4), alter the balance between their synthesis and catabolism (e.g., proline), or trigger their release from polymeric forms (e.g., monomeric sugars like glucose and fructose can be released from their polymeric forms, starch and fructans). Once the stress is removed, these monomers can be repolymerized to facilitate rapid and reversible osmotic adjustment, or they can be metabolized to produce primary metabolites or energy. Unlike many inorganic solutes, which may become toxic at high concentrations, the organic character of compatible ­solutes greatly diminishes their potential toxicity. Many ions found in cells adversely affect metabolic processes at high concentrations, possibly by binding to and altering the

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Amino acid:

Tertiary sulfonium compound: H3C

+ N H

COO

S+

–

CH2

CH2

COO–

H3C

H

Dimethylsulfoniopropionate

Proline

Quaternary ammonium compounds: H3C N+

H3C

(CH2)n

COO–

H3C

COO–

+ N H3C

n = 1, Glycine betaine n = 2, β-Alanine betaine

OH

H

H H OCH3 H

H H OH

OH

HO

C

H

HO

C

H

H

C

OH

H

C

OH

CH2

CH2

O

S

O –

O

Choline-O-sulfate

HO

CH2 O H

HO OH

CH2OH Pinitol

N+

H3C

Proline betaine

CH2OH OH

H3C

CH3

Polyhydric alcohols:

HO

O

H3C

Mannitol

O OH HO

CH2 O H

HO

CH2

OH

H

O HO

O OH

OH

H2C OH

Raffinose

FIGURE 22.5  Chemical structures of several important compatible solutes (osmolytes) that accumulate in plant cells in response to water deficit. The organic character and nonionic or zwitterionic nature of compatible solutes greatly diminishes their potential toxicity.

­ roperties of cofactors, substrates, membranes, and enzymes. p Furthermore, many ions can enter the hydration shells of a protein and promote its denaturation. In contrast, compatible solutes tend to be neutrally charged at physiological pH, either nonionic or zwitterionic (dipolar, with spatially ­separated positive and negative charges), and are excluded from hydration shells of macromolecules (Fig. 22.6). In addition to stereotypical charge characteristics, ­compounds active in osmotic adjustment show distribution patterns that support water potential equilibria among the various membrane‐bound compartments of the cell. Vacuoles tend to accumulate charged ions and solutes that would ­perturb metabolism if present in the cytoplasm. Compatible solutes in the cytoplasm, however, allow the cytosol to achieve osmotic balance with the vacuole. Another possible mechanism for osmotic adjustment is increasing movement of water into the cell. The hydrophobic nature of the lipid bilayer presents a considerable barrier to the free movement of water into the cell and between intracellular compartments. However, plasma membranes ­ and tonoplasts can be rendered more permeable to water by

proteinaceous transmembrane water channels called aquaporins (see Chapter 3). Water movement through aquaporins can be modulated rapidly, and evidence suggests these ­channels may facilitate water movement in drought‐stressed plant tissues and promote rapid recovery of turgor upon watering. The abundance of aquaporin mRNA is correlated with turgor changes in leaves subjected to osmotic stress. Higher transcript levels, enhanced translation, and activation of existing proteins may constitute multiple mechanisms for regulating aquaporin abundance and activity, which may be advantageous for plants coping with water deficit.

22.2.5  Some compatible solutes may have protective functions in addition to osmotic adjustment The role that compatible solutes play in osmotically stressed plants is often defined as “osmoprotection”—solute accumulation is believed to have a protective function against water

Chapter 22  Responses to Abiotic Stress

22.2.6  Genetic engineering provides an opportunity to test the adaptive significance of compatible solutes

Perturbing ions

Na+

Cl

–

Pro

Pro Cl–

Cl

–

Cl–

Cl–

Cl–

Disrupted protein (Fewer ordered H2O molecules bound to protein, entropy high)

Compatible solutes (e.g. proline)

Pro

Pro

Pro

Pro

Pro

Pro Pro

Pro

Pro

Intact protein (Highly ordered H2O molecules surround protein, entropy low)

FIGURE 22.6  Hydration shells of macromolecules are not disrupted by compatible solutes. Depicted is a protein with a hydration shell (i.e., surrounded by ordered H2O molecules). Ions such as Na+ and Cl– can penetrate these shells and interfere with the noncovalent ­interactions that maintain the structure of the protein. Unlike ions, compatible solutes, such as proline and glycine betaine, do not p­ enetrate the protein’s hydration shell, so protein and solute do not come into direct contact. The hydration shells of the ions and ­compatible solutes are not shown.

deficit. However, this term should be applied with discretion. Historically, direct physiological evidence for osmoprotection has been obtained for bacteria, but not for plants. Salt‐­ sensitive bacteria, such as Escherichia coli, can be induced to grow in high‐salt media supplemented with specific compounds, including the osmolytes shown in Figure  22.5. Therefore, the presence of concentrated compatible solutes in drought‐tolerant and halophilic (“salt‐loving”) plants has been taken as compelling, albeit indirect, evidence for the role of osmolyte in plant osmoprotection. Compatible solutes that accumulate in plants may also act as antioxidants to minimize the impact of abiotic stress in plants. In vitro, many of these compounds directly offset the deleterious, perturbing effects of ions. For example, glycine betaine prevents salt‐induced inactivation of ­ Rubisco and destabilization of Photosystem II (PSII). Sorbitol, mannitol, myo‐inositol, proline, and raffinose can also scavenge hydroxyl radicals in vitro, though glycine betaine cannot. This antioxidant activity suggests a protective role for these compounds in osmotic stress tolerance distinct from osmotic adjustment.

Drought‐ or salt‐sensitive plants can be transformed with genes encoding enzymes that are critical for the synthesis of a putative osmoprotectant. Transformed plants can then be assayed for enhanced accumulation of compatible solutes and ability to adjust osmotically to conditions of drought and high salinity. In fact, biotechnological approaches for e­ nhancing drought tolerance typically involve dissection of the biosynthesis pathways of specific osmolytes and subsequent manipulation of enzymes operating in those pathways. Among the compounds studied to date are proline, glycine betaine, oligosaccharides such as raffinose, and polyhydric alcohols, including mannitol and pinitol. The results from these studies indicate a role for these compounds not only in responses to water and salinity stress, but to other types of abiotic stress as well. Upon exposure to abiotic stress, many plants accumulate carbohydrates such as mannitol, trehalose, myo‐inositol, fructan, galactinol, and raffinose. These metabolites not only serve as energy storage but are thought to play roles in b ­ alancing osmotic strength, stabilizing macromolecules, and preserving the membrane. Genetic manipulations of key enzymes involved in the biosynthetic pathways of these c­arbohydrates can improve abiotic stress tolerance in transgenic plants. Metabolite profiling using different types of mass spectrometry has revealed that various types of m ­ etabolite, including proline and branched chain amino acids such as valine, leucine, and isoleucine, accumulate in response to abiotic stress and serve a protective function in cells (see Fig. 22.6). The mechanisms that facilitate proline accumulation in response to environmental conditions are discussed in Chapter 7. Glycine betaine (N,N,N‐trimethylglycine, GB) is synthesized under various types of environmental stress and functions as a major osmoprotectant. It stabilizes the ­ ­quaternary structures of the PSII protein‐pigment under high salinity and maintains the ordered state of ­membranes at extreme temperatures. GB is synthesized mainly from choline via betaine aldehyde (BA), with the first and second steps in the pathway catalyzed by choline monooxygenase (CMO) and betaine aldehyde dehydrogenase (BADH), respectively (Fig.  22.7). A wide variety of plant species ­accumulate GB, but Arabidopsis, rice (Oryza sativa), and tobacco (Nicotiana) are considered “nonaccumulators.” Genetic engineering to ­introduce GB‐biosynthetic pathways into nonaccumulator species is a promising approach to increase abiotic stress t­olerance. Transgenic rice plants ­overexpressing the barley BADH gene convert applied BA into GB more efficiently than wild‐type plants and exhibit significant tolerance to salt, cold, and heat stress. Raffinose family oligosaccharides (RFOs), such as raffinose and stachyose and their precursor galactinol, play important roles in drought stress tolerance in plants and seeds. RFOs are important for reducing reactive oxygen species (ROS) that accumulate and cause oxidative stress under conditions of

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Betaine synthesis pathway +

H3N

CH2

CH2OH

Ethanolamine O

ATP

Ethanolamine kinase ADP

H3N

CH2

CH2O P

+

H 3C

Phosphoethanolamine

O

C

R

HC

O

C

R’

O

H3C

+

H2C

N

CH2

CH2

O

H3C

O

P

CH2

O

–

O

Phosphatidylcholine S-Adenosylmethionine (3)

Phospho-base N-methyltransferases

S-Adenosylhomocysteine (3)

Phosphocholine phosphatase

H3C H3C

N+

CH2

P

Phosphocholine

H3C N+

H 3C

CH2O P

H3C

Phospholipase D

Phosphatidate

i

CH2

CH2OH

H3C Choline

O2 + 2 H+ + 2 Fdxred Choline monooxygenase 2 H2O + 2 Fdxox O

H3C H 3C

N

+

CH2

C H

H3C Betaine aldehyde NAD+

Betaine aldehyde dehydrogenase

NADH

H3C N+

H 3C

CH2

COO–

H3C Glycine betaine

FIGURE 22.7  Proposed biosynthetic pathways of glycine betaine from choline. Introduction of such pathways into nonaccumulator species is a promising approach for increasing abiotic stress tolerance.

abiotic stress. They are synthesized from UDP‐galactose (Fig.  22.8), which is converted into galactinol by galactinol synthase (GolS). Raffinose synthase then produces raffinose from the galactinol. Both raffinose and galactinol accumulate in Arabidopsis in response to drought, high salinity, and cold stress. These stressors induce expression of the genes for GolS and raffinose synthase, resulting in the production of RFOs. Overexpression of the stress‐inducible GolS gene improves drought tolerance by activating the accumulation of galactinol and raffinose in transgenic Arabidopsis plants.

22.2.7  Some proteins protect macromolecules and membranes against damage Large numbers of proteins, such as late embryogenesis abundant (LEA) proteins and heat shock proteins (HSPs), protect plant cells directly against water or osmotic stress damage. They exert their protective effects either as highly hydrophilic proteins that retain water or as molecular

Chapter 22  Responses to Abiotic Stress

HO OH

CH2OH

HO

OH

O O-UDP

OH O

GolS

OH

OH

O

Galactinol synthase

HO

+

CH2CH OH

OH

HO

OH

+

OH

UDP

OH

OH

UDP-galactose

1059

HO

myo-Inositol

OH

Galactinol CH2OH

HO HO

CH2OH OH

OH

O

O CH2OH OHO

OH

OH

+

OH

OH

CH2OH O

O

HO

O

HO

HO

HO HOCH2

Sucrose

OH

OH OCH2

Galactinol

HO HO

CH2OH OH

O

OH OCH

Stachyose synthase

2

OH OH

+

HO HOCH2

OH HO

O

OH

myo-Inositol CH2OH

OH

O

OH

O

HO

OH

CH2OH

Raffinose

OH

O

OH HO HOCH2

HO

+

2

OH

OH

HO OH OCH

OH O

OH OH

OH O

O

OHOCH 2 HO

O

Galactinol

O

HO

+

Raffinose

CH2OH OH

CH2OH OH

OHO

O

OH

HO

OH

HO

OH CH2OH

OH

OH

OH

Raffinose synthase

O

OH

OH

OH O

myo-Inositol

HO CH2OH

Stachyose

FIGURE 22.8  Biosynthetic pathway for raffinose family oligosaccharides (RFOs). RFOs are osmolytes that also reduce reactive oxygen species that cause oxidative stress.

c­haperones that prevent denaturation of macromolecules and protect membranes. Plants exposed to abiotic stress usually express or accumulate higher levels of these proteins, suggesting they are necessary in plant abiotic stress protection in general. LEA proteins were first reported as a group of proteins produced in abundance during late embryogenesis, with maximum expression during seed desiccation. Their genes have been identified and their mRNAs shown to accumulate during the seed maturation process. Some LEA family genes are also induced by cold, osmotic stress, or exogenous ABA. Although their precise functions remain unknown, LEA proteins accumulate in stressed plants and are thought to be protective molecules that confer stress tolerance. They are associated with desiccation tolerance in seeds, pollen, and anhydrobiotic plants and may be essential for the survival of plant cells under such conditions. LEA proteins may function

in an unstructured state, with folding and conformational changes induced under conditions of water deficit. LEA ­proteins become more folded and develop a significant α‐­ helical component when dried. Another hypothesis is that LEA proteins may decrease the collisions between partially denatured proteins, thereby reducing aggregation of exposed hydrophobic domains. Drought conditions may also induce accumulation of HSPs, which accumulate in plants in response to high ­temperature stress (see Section 22.7); some genes encoding HSPs are also induced by drought or osmotic stress. HSPs are molecular chaperones with important roles in the ­folding and assembly of proteins, stabilization of proteins and ­membranes, protein refolding, and removal of nonfunctional and degraded proteins under conditions of stress. Manipulation of HSP gene expression not only enhances thermotolerance of transgenic plants, but also increases water stress tolerance.

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Part V  PLANT ENVIRONMENT AND AGRICULTURE FIGURE 22.9  Ion channels and transporters help maintain ionic homeostasis under conditions of salt stress. Salt treatment increases the activity of Na+/H+ antiporters, which then move Na+ either out of the cell or into the vacuole. For example, salt stress triggers changes in Ca2+, which regulate SOS3, which then regulates SOS2 protein kinase to activate the plasma membrane SOS1 Na+/H+ antiporter. Changes in ABA levels also regulate the NHX1 Na+/H+ antiporter and other ion transporters. Other transporters that mediate Na+ transport into the cell include non‐selective cation channels (NSCC) and HKT1/HKT2, the low‐affinity cation transporter (LCT1), and K+ transporters of the KUP/HAK/KT and AKT families. Note that the cell wall is not shown for clarity.

Salt stress

HKT1/ HKT2

KUP/ HAK/KT

ABA

Na+

LCT1

Ca2+ Na+

K+

NHX1 SOS3

AKT

H+ Vacuole

K+ K+

SOS2

Na

+

H+

NSCC

Cytosol SOS1

Na+

22.2.8  Water deficit and salinity affect transport across membranes Drought and salt stress both involve acclimation to low water potential; however, plants growing under high salinity must also cope with potentially toxic amounts of specific ions. Although ion uptake may provide a means of osmotic ­adjustment, high cytosolic concentrations of some ions such as Na+ perturb metabolism. Thus, regulating the concentration, composition, and distribution of ions within the cell can be an essential feature of tolerance to osmotic stress. Plants or cultured cells exposed to high NaCl concentrations tend to accumulate Na+ as a result of Na+ influx through channels and transporters (Fig.  22.9). Electrophysiological studies have indicated that nonselective cation channels (NSCCs) mediate primary influx of Na+ under saline ­conditions, but no definitive candidate molecules have been identified. Other Na+ transporters identified in plants include the high-affinity K+ transporter HKT1, which controls the internal Na+ distribution between the root and the shoot under saline conditions, and HKT2, which plays a role in p ­ rimary Na+ uptake under K+ ­deprivation. Transporters such as the low‐affinity cation transporter 1 (LCT1) and K+ t­ ransporters of the K+ uptake permease/ high-affinity K+ uptake/K+ transport (KUP/HAK/KT) and Arabidopsis K+ transporter (AKT) families may also ­participate in Na+ uptake. Na+ inhibits a variety of biological processes, including K+ absorption. Under conditions of salt stress, ­therefore, plants exclude Na+ and accumulate K+ in shoots to maintain a high cytosolic K+/Na+ ratio, especially in leaves. Cytoplasmic Na+ accumulation is partially prevented by the active export of cytosolic Na+ across the plasma ­membrane to the extracellular space and across the tonoplast into the vacuole. In the vacuole, it can accumulate to concentrations that would otherwise have a marked effect on the osmotic

balance of plant cells. Salt treatment increases the activity of vacuolar Na+/H+ antiporters in several plant systems, such as sugar beet (Beta vulgaris) cultures and barley (Hordeum ­vulgare) roots. Operation of these carriers requires an electrochemical potential gradient across the tonoplast, usually generated through the action of H+ pumps such as the plasma membrane H+‐ATPase and vacuolar H+‐ATPase and H+‐ pyrophosphatase (H+‐PPase) (see Chapter 3). Arabidopsis SALT OVERLY SENSITIVE 1 (SOS1) is a plasma membrane Na+/H+ antiporter that mediates the p ­ assive transport of H+ into and active transport of Na+ out of the cell (Fig. 22.9). Arabidopsis sos1 mutants show an NaCl‐hypersensitive phenotype, whereas overexpression of SOS1 improves salt stress tolerance. Transgenic plants treated with increasing Na+ concentrations show significant salt tolerance compared to wild‐type controls, and this tolerance is due to reduced accumulation of Na+. Na+ compartmentalization into vacuoles is another mechanism of avoiding high Na+ toxicity, and the vacuolar Na+/H+ antiporter Na+/H+ exchanger 1 (NHX1) is important in this process (Fig.  22.9). Transgenic plants overexpressing NHX1 show increased Na+ transport into vacuoles and can grow in salty water. Overexpression of NHX1 in transgenic Arabidopsis improves salinity stress tolerance, allowing transgenic plants to grow and produce seeds even in the presence of 200 mM NaCl hydroponic solution (control plants in the same solution die). NHX overexpression confers salt tolerance in a wide range of plant species.

22.2.9  The hormone ABA plays an important role in plant response to water deficit In addition to its roles in seed maturation and germination, the plant hormone abscisic acid (ABA) also plays important roles in plant responses to water deficit (see Chapter 17). ABA

Chapter 22  Responses to Abiotic Stress

Cytosol Chloroplast 9-cis-Epoxycarotenoids NCED Xanthoxin

Biosynthesis

Xanthoxin ABA2 ABAId AAO, ABA3 ABA

Catabolism

CYP707A

8’-OH ABA

Phaseic acid

ABAGT-ase

AtBG1

ABA-GE

Recycling

ABA-GE

Vacuole apoplast

FIGURE 22.10  ABA biosynthesis and degradation in response to a­ biotic stress. In the final process of the ABA biosynthetic pathway, xanthoxin is cleaved from 9‐cis‐epoxycarotenoids by NCED and released from the chloroplast into the cytoplasm. Then, ABA is ­produced through abscisic aldehyde (ABAld) formation. In ABA catabolism, the major pathway seems to be an oxidative pathway that is triggered by ABA 8′‐hydroxylation catalyzed by the CYP707A family. There are other pathways for ABA inactivation, such as ­glucosylation. Red letters indicate drought stress‐­responsive ­regulation. CYP707As are shown in orange, which indicates ­dehydration and rehydration‐responsive regulation. For more details on ABA metabolism see Chapter 17. NCED, 9‐cis‐epoxycarotenoid ­dioxygenase; ABA2, short‐chain dehydrogenase/reductase; AAO, abscisic aldehyde oxygenase; ABA3, molybdenum cofactor sulfurase; CYP707A, ABA 8′‐hydroxylase; ABA‐GTase, ABA glucosyltransferase; AtBG1, β‐glucosidase; ABA‐GE, ABA glucosyl ester.

is produced in response to water deficit and is involved in ­stomatal closure, which is required to prevent water loss from leaves under conditions of dehydration. The responses of ­stomata to water deficit have been studied extensively (see Chapter 3). ABA production is also essential for the cellular accumulation of various metabolites and proteins that have protective roles in water deficit resistance. Moreover, accumulated ABA induces a number of stress genes, the products of which are important for plant responses and tolerance to

dehydration. Endogenous ABA levels—determined by the balance between ABA biosynthesis and catabolism—increase significantly in response to drought or high salinity. ABA is primarily synthesized de novo in response to drought and high salinity, and the genes involved in ABA biosynthesis and catabolism have been identified mainly ­ through genetic and genomic analyses (see Chapter  17). Xanthoxin, a C15 precursor of ABA, is produced by direct cleavage of C40 carotenoids by 9‐cis‐epoxycarotenoid dioxygenase (NCED) in plastids. This step is critical for ABA stress responses (Fig. 22.10, see Chapter 17). NCED is encoded by a multigene family, and the stress‐inducible NCED3 gene plays a key role in ABA biosynthesis under stress conditions. In Arabidopsis, overexpression of NCED3 increases endogenous ABA levels and improves drought stress tolerance, whereas disruption results in defective ABA accumulation under drought stress and impairs drought stress tolerance. At least two regulatory pathways exist for ABA ­catabolism: the oxidative pathway and the sugar conjugation pathway. The oxidative pathway is catalyzed by ABA C‐8′ hydroxylase to produce phaseic acid. This enzyme belongs to a class of  cytochrome P450 monooxygenases, the CYP707As (Fig. 22.10), of which there are four members in Arabidopsis. Among the four CYP707As, CYP707A3 is a major enzyme for ABA catabolism during the osmotic stress response. The CYP707A3 gene is induced by rehydration after exposure to conditions of dehydration, and CYP707A3 knockout mutants show increased endogenous ABA and dehydration tolerance. In the sugar conjugation pathway, ABA is inactivated in sugar‐conjugated forms, such as ABA glucosyl ester, and stored in vacuoles or apoplastic pools (Fig.  22.10). Under conditions of dehydration, ABA is released from the glucosyl ester form by β‐glucosidase. Regulation of the genes involved in ABA synthesis and catabolism by transgenic technology can improve drought tolerance. ABA transport is also thought to be important for plant responses to abiotic stress. NCED3 is mainly expressed in ­vascular tissues, and endogenous ABA is mainly synthesized in vascular tissue of leaves. ATP binding cassette (ABC) transporters function as ABA transporters in both the export and import of ABA. Regulation of ABA transport is important for both inter‐ and intracellular signaling in plants.

22.3  Gene expression and signal transduction in response to dehydration Water deficit or dehydration can induce or repress the ­expression of thousands of plant genes with diverse functions. Many products of dehydration‐inducible genes function in stress tolerance and responses at the cellular level, and their overexpression can improve stress tolerance, indicating important functions in plant responses and adaptation to dehydration or drought.

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22.3.1  Microarray and other techniques provide insights into the expression of dehydration‐inducible genes and their predicted functions Microarray technology employing cDNAs or oligonucleotides (Chapter  9) is a powerful tool for analyzing the gene expression profiles of plants exposed to abiotic stress. Using this technology, many stress‐inducible genes have been ­identified in various plants, and most of these are common among different plants, indicating their general roles in plant stress responses. Analysis of their functions is important for understanding the molecular mechanisms governing plant stress responses and tolerance. Moreover, basic knowledge regarding their functions is useful for enhancing stress ­tolerance in crops through transgenic technology. Microarrays and more recently RNA sequencing technologies have been used to identify genes in Arabidopsis that are differentially expressed in various organs under various growth conditions, including stress conditions and phytohormone treatment. In addition, coexpression analysis of different trans­ criptome datasets p ­ rovides a new method for the discovery of genes with similar expression profiles based on bioinformatics analysis; such genes often have similar functions, and this facilitates ­elucidation of stress‐induced genes of unknown function. Results from these investigations show that many dehydration‐inducible genes are also induced by high salinity and ABA treatment, suggesting significant crosstalk between the drought, high‐salinity, and ABA response pathways. In ­contrast, only a small number of drought‐inducible genes are also induced by cold stress. The products of the dehydration‐inducible genes identified through recent microarray analyses can be classified into two major groups: ●●

●●

proteins and enzymes that function in abiotic stress ­tolerance, such as chaperones, LEA proteins, osmotin, antifreeze proteins, RNA‐binding proteins, key enzymes for osmolyte biosynthesis, water channel proteins, sugar and proline transporters, detoxification enzymes, and ­various proteases regulatory proteins, such as protein factors involved in further regulation of gene expression and signal transduction. These include transcription factors, protein kinases, protein phosphatases, enzymes involved in phospholipid metabolism, other signaling molecules, and factors involved in posttranscriptional regulation, such as RNA‐ processing enzymes.

Many transcription factor genes are stress inducible, ­suggesting that various transcriptional regulatory ­mechanisms may function in regulating dehydration, cold, or high‐salinity stress signal transduction pathways. These transcription ­factors control expression of stress‐inducible genes, either cooperatively or independently, and may constitute regulatory networks.

Transcriptome analyses at the whole‐genome level have also been carried out using high‐throughput DNA sequencing and genome tiling arrays (see Chapter  9), and novel mechanisms of transcriptional regulation by noncoding RNAs or micro‐RNAs have been reported. Micro‐RNAs are involved in post‐transcriptional regulation in abiotic stress responses by regulating mRNA stability (see Chapter  6). Sense and antisense RNAs for stress‐responsive mRNAs have been reported to function in stress responses. Moreover, gene  expression driven by stress often depends on DNA methylation or posttranslational histone modification. ­ Modifications of DNA or histones may play important roles in gene expression and plant growth under stress conditions. Therefore, in addition to gene expression, epigenetic ­regulation (see Chapter 9) may play an important role in plant adaptation to abiotic stress.

22.3.2  Water deficit triggers changes in gene expression at the transcriptional level Various transcription factors are important regulatory factors in stress‐responsive gene expression. Several act as master switches for the expression of various sets of downstream genes through specific binding to the cis‐acting elements in their promoters. Corresponding transcription factors have been isolated that bind to cis‐elements and initiate transcription of target genes. This type of transcription unit is called a “regulon.” Analysis of the expression mechanisms of osmotic and cold stress‐responsive genes has revealed multiple ­regulons in transcription. Many plant transcription factors, such as AP2/ERF, bZIP, MYB, MYC, Cys2His2 zinc‐finger, and NAC, constitute multigene families involved in stress‐ responsive gene expression. Dehydration triggers de novo production of ABA, which in turn induces expression of stress‐related genes (see Section  22.2.7). There is evidence for both ABA‐dependent and ‐independent regulatory systems governing stress‐­ inducible gene expression. Many dehydration‐ and salinity‐ inducible genes can also be activated by exogenous ABA treatment, but a number of genes are also unaffected by ABA treatment. Promoter regions of dehydration‐inducible genes have been analyzed extensively to identify regulatory cis‐­acting elements. Both cis‐ and trans‐acting regulatory elements functioning in ABA‐independent and ‐dependent gene expression induced by water deficit have been studied in detail.

22.3.3  Endogenous ABA accumulation under dehydration stress also controls gene expression Analysis of ABA‐responsive promoters has revealed a diverse range of potential cis‐acting regulatory elements, but most ubiquitous cis‐elements share the (C/T)ACGTGGC

Chapter 22  Responses to Abiotic Stress Osmotic stress

Heat stress

1063

Cold stress

ABA-independent pathway

JA

ABA

Jasmonic acid

ABA-dependent pathway

MYC2

MYB2

DREB2 (AP2/ERF)

CBF/DREB1 (AP2/ERF)

P

ZAT12

AREB/ABF (bZIP)

ABRE

CE

DRE/CRT

Target stress-inducible genes

Stress tolerance

FIGURE 22.11  Overlapping transcriptional regulatory networks in plant responses to osmotic, heat, and cold stress. Transcription factors and cis‐acting elements are shown in ovals and boxes, respectively. ABA‐dependent pathway (shown in red): ABRE is a key cis‐acting element of genes regulated by ABA‐dependent stress responses. The coupling element (CE) is necessary for ABA‐dependent transcription. AREB/ABF are major transcription factors in the ABA response, and are phosphorylated for their activation (see Fig. 22.12). MYC2 is not only regulated by ABA, but also by jasmonic acid (JA) and functions in crosstalk between ABA and JA responses (see Section 22.8). MYB2 is induced by ABA and functions in cooperation with MYC2 in dehydration‐inducible gene expression. Other transcription factors have been identified in abiotic stress response but are not shown. ABA‐independent pathways (shown in blue): DRE/CRT is a key cis‐acting element of genes regulated by ABA‐independent stress responses. CBF/DREB1s are major transcription factors in the cold stress response, whereas DREB2 is mainly regulated by osmotic and heat stress. CBF/DREB1 is explained in Section 22.4.7. The roles of ZAT12 in the cold stress response and DREB2 in the heat response are described in ­Sections 22.4.7 and 22.7.5, respectively.

consensus sequence, the ABA‐responsive element (ABRE; PyACGTGGC). ABRE is a major cis‐acting element in ABA‐ responsive gene expression (Figs.  22.11 and 22.12) that requires another cis‐acting element, the coupling element (CE), for function. The basic leucine zipper (bZIP) transcription factors AREB and ABF can bind to the ABRE cis‐ acting element to activate ABA‐dependent gene expression. The AREB and ABF transcription factors require an ABA‐ mediated signal for activation, as indicated by their reduced activity in the ABA‐deficient aba2 and ABA‐­insensitive abi1 mutants and their enhanced activity in the ABA‐hypersensitive era1 mutant of Arabidopsis. This is due to the ABA‐ dependent phosphorylation of AREB and ABF proteins by the SNF1‐related protein kinases SnRK2s (Fig.  22.12 and see Section 22.3.5). Transgenic plants expressing an active form of AREB1 with multi‐site mutations show induction of many ABA‐responsive genes without application of exogenous ABA. These observations indicate an important role of ­ protein phosphorylation in the activation of AREB and ABF transcription factors in response to increased levels of endogenous ABA under osmotic stress conditions. In addition to the AREB and ABF bZIP ­ transcription factors that function as key regulators of ABA‐dependent gene expression, various ­

t­ ranscription factors with MYB2, MYC2 (bHLH), NAC, HD‐ ZIP, HD, HB, AP2, and B3 domains are involved in ABA responses under osmotic stress conditions (see Section 22.8). Among them, MYB2 and MYC2 are involved in stress‐­ regulated gene expression after the accumulation of ABA. MYC2 is also important in jasmonic acid (JA)‐regulated gene expression (see Fig. 22.11).

22.3.4  ABA‐independent pathways can also influence the abiotic stress response and DREB2 transcription factors The promoters of genes such as RD29A and COR15, which induced by dehydration, high salinity, and cold, contain two major cis‐acting elements, ABRE and DRE (dehydration‐ responsive element)/CRT (C‐RepeaT), both of which are involved in stress‐inducible gene expression (Figs.  22.11 and  22.12). Whereas ABRE functions in ABA‐dependent pathways, DRE/CRT functions in ABA‐independent gene ­ expression in response to abiotic stress (Fig. 22.12); it also contains the conserved cis‐acting promoter motif A/GCCGAC.

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Part V  PLANT ENVIRONMENT AND AGRICULTURE Cold

Heat

Dehydration, salinity

ABA PYR/RCAR ICE1

CAMTA

HSF

PP2C CBF/DREB1

DREB2 SnRK2

CBF/DREB1

DREB2

DRE/CRT

DRIP (E3 ligase) CE

P

AREB/ABF ABRE

TATA

FIGURE 22.12  Function of cis‐acting elements DRE/CRT and ABRE in ABA‐independent and ‐dependent induction of RD29A, respectively. AP2/ERF transcription factors CBF/DREB1 and DREB2 bind DRE/CRT in response to different types of stress: CBF/DREB1 functions in cold‐ inducible gene expression, and their genes are regulated by the transcription factors ICE1 and CAMTA (see Section 22.4.7). DREB2 is involved in dehydration‐ and salinity‐inducible gene expression and is activated by protein stabilization. AREB/ABFs are bZIP transcription factors that bind ABRE/CE and mediate ABA‐dependent gene expression (see Fig. 22.11). Major ABA signaling cascades (involving ABA receptors PYR/RCARs, PP2C phosphatases, and SnRK2 protein kinases) function upstream of AREB/ABF transcription factors (see Section 22.8). The function of CBF/DREB 1 in cold response is also described in Section 22.4.7, and the function of DREB2 in heat response is also described in Section 22.7.5.

Two transcription factors belonging to the AP2/ERF family, CBF/DREB1 and DREB2, bind to the DRE/CRT element. The genes encoding CBF/DREB1 are rapidly and transiently induced by cold stress, and their products activate expression of target stress‐inducible genes (see Section 22.4.7); ­expression of DREB2, on the other hand, is mainly induced by dehydration, high salinity, and heat stress. Overexpression of CBF/DREB1 in transgenic plants increases tolerance to drought, freezing, and salt stress, ­suggesting the CBF/DREB1 proteins function in the development of cold stress tolerance without posttranslational modification (Fig. 22.13A). Overexpression of DREB2 in transgenic Arabidopsis plants, though, does not improve stress tolerance, which suggests the involvement of post‐translational activation of DREB2 proteins. Deletion of a region of 30 amino acids adjacent to the AP2/ERF DNA binding domain, named the DREB2A negative regulatory domain (NRD), transforms DREB2A into the ­constitutively active form DREB2A‐CA, which interacts with an E3 ubiquitin ligase (DRIP: DREB2 interacting protein 1). Posttranslational regulation may be necessary for stabilization of the DREB2A protein. Arabidopsis DREB2A target genes are induced to a significantly greater extent by dehydration stress than by cold stress (see Fig. 22.12).

Preference of DNA‐binding activities has been compared between the DREB2A and DREB1A protein. Whereas DREB2A binds preferentially to the sequence ACCGAC, DREB1A shows affinity for the A/GCCGACNT sequence. These data indicate that the function of the DREB2‐type ­transcription factor in the drought stress response is different from that of the DREB1 protein involved in the cold stress response (Fig. 22.13B). DREB2A is also involved in responses to heat and drought stress, as discussed in Section 22.8. Transgenic plants that overexpress various transcription factors have more abiotic stress tolerance. For example, ­overexpression of the rice (Oryza sativa) NAC transcription factor SNAC1 improves drought tolerance of transgenic rice  and induces expression of a large number of stress‐­ regulated genes. Another good example involves the NF‐Y ­transcription factor family; transgenic maize (Zea mays) plants overexpressing ZmNF‐YB2 exhibit tolerance to drought based on various stress‐related parameters including chlorophyll ­content, stomatal conductance, leaf temperature, reduced wilting, and maintenance of photosynthesis. Moreover, maize plants overexpressing ZmNF‐YB2 have better grain yield under conditions of drought stress (Fig. 22.14). Constitutive or conditional expression of transcription factors has great potential for the development of drought‐tolerant crops.

Chapter 22  Responses to Abiotic Stress Constitutive active DREB2A overexpressors wt

DREB1A-b

DREB2A CA-a DREB2A CA-b DREB2A CA-c

Before treatment

Freezing

Drought

A Drought High salinity

Cold

Modification

DREB transcription factors

DRE/CRT Cis-elements

Target genes

DREB2A

DREB1A/CBF3

DREB2A

DREB1A/CBF3

ACCGACNA/G/C

ACCGACNT

A/GCCGACNT

MT2A (metalothionein-like) At1g52690 (LEA) AtGolS1 HSPs HSFs etc.

RD29A RD17/COR48 LEA14 At2g223120 AtGolS3 At5g62350 etc.

COR15A COR15B KIN1 KIN2 AtCOR413-TM1 At2g02100 etc.

B

FIGURE 22.13  Arabidopsis CBF/DREB1 and DREB2 mediate overlapping and unique pathways of resistance to freezing, drought, and salt stress. (A) Transgenic Arabidopsis plants that overexpress DREB1 or the constitutively active form of DREB2 (DREB2‐CA) show tolerance to both drought and freezing stress. (B) Model of the induction of genes regulated by DREB1 and DREB2A under drought, salinity, and cold stress. The genes downstream of DREB1 and DREB2A are categorized into three groups. The center group contains genes shared by DREB1A and DREB2A, and the other groups consist of DREB1A‐ and DREB2A‐specific genes. Source: Sakuma et al. (2006). Plant Cell 18:1292–1309.

22.3.5  Osmotic stress triggers various signaling pathways in plants Though changes in transcriptional regulation in response to osmotic stress have been analyzed extensively, their cellular signal transduction pathways are not yet fully understood. Our current understanding of cellular signal transduction in the osmotic stress response has been based largely on ­knowledge obtained from analysis of yeast and other microorganisms. Protein kinase homologs have been identified based on their functions in stress signaling pathways (Fig.  22.15). Genetic

screening of mutants showing abnormal responses to abiotic stress, either resistance or sensitivity, has been performed to isolate factors involved in stress responses and signaling pathways. Stress signaling involves modification of regulatory p ­ roteins through phosphorylation and dephosphorylation, ubiquitination, and calcium binding. These groups of r­ egulatory proteins are modified or activated to fulfill their functions quickly. Although the complexity of this signal transduction network is still not completely understood, ­several types of protein kinase and calcium‐sensing proteins have been characterized, and their functions have also been investigated in transgenic plants.

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A

B

FIGURE 22.14  Transgenic maize (Zea mays) plants that overexpress ZmNF‐YB2 (plants at right) grown in a greenhouse (A) and a test field (B) show significantly improved drought tolerance. The rice (Oryza sativa) actin gene promoter was used to overexpress ZmNF‐YB2 cDNA.

Osmotic stress sensors have been analyzed in bacteria and yeast. In yeast, the His‐Asp phosphorelay system functions in sensing osmotic stress. The membrane‐bound histidine kinase Sln1p functions as an osmosensor, and the Arabidopsis histidine kinase AHK1 can complement a yeast sln1 mutant, ­indicating AHK1 also functions as an osmosensor in yeast. In addition, AHK1 protein functions as a positive regulator in water deficit and osmotic stress signaling in plants (Fig.  22.15). Transcriptional profiling analysis of ahk1 knockdown mutants has revealed that some target genes of AREB1, ANAC, and DREB2A transcription factors are downregulated in the mutant. In contrast, transgenic Arabidopsis overexpressing AHK1 driven by its own ­promoter show markedly improved drought stress tolerance. The response regulator‐like factors (ARRs) are thought to function downstream of AHK1 in osmotic stress signaling pathways like the yeast His‐Asp osmosensing phosphorelay system Sln1p‐Ypd1p‐Ssk1p.

The mitogen‐activated protein kinase (MAPK) cascade functions in various cell signaling cascades (see Chapter 18), including osmotic stress responses in eukaryotes. It is characterized by a three‐step phosphorylation relay by MAPK, MAPK kinase (MAPKK), and MAPKK kinase (MAPKKK) on ­conserved threonine, tyrosine, and serine residues (see Chapter 18). In yeast, the HOG1 pathway (Ssk2/22p‐Pbs1p‐ Hog1p) is a MAPK cascade that functions downstream of Sln1p‐Ypd1p‐Ssk1p. In plants, MAPK cascades are rapidly activated by various environmental stresses and pathogen infection. The Arabidopsis MEKK1‐MKK2‐MPK4/MPK6 cascade is activated by salinity and also cold stress, and plays a role in cellular stress signaling pathways (Fig. 22.15). These observations suggest that MAPK cascades function in abiotic as well as biotic stress signaling. However, the upstream signaling components in plants have yet to be determined. ROS are produced in response to reduced metabolic activities of chloroplasts and mitochondria in biotic and abiotic stress responses (see Section 22.6). Factors involved in signal transduction pathways may function as ROS sensors to ­control downstream events. ROS regulate transcription ­factors, such as the heat shock factors (HSFs) involved in the heat stress response (see Section  22.7) and NPR1 involved in ­disease resistance. Protein kinases such as SOS2 and ANP1 and heterotrimeric G proteins are also regulated by ROS. Therefore, ROS can modulate abiotic stress signaling p ­ athways and function in crosstalk in stress signaling (see Section 22.8). Of the many second messengers involved in cell signaling pathways, Ca2+ is the most ubiquitous (see Chapter  18). In plants, Ca2+ levels increase transiently in response to various stimuli, including water deficit. Osmotic stress can trigger rapid and dynamic oscillations in cytosolic Ca2+ levels that are  mediated by various Ca2+ channels and transporters (Fig. 22.15). The Ca2+‐dependent stress signaling pathway is important in plants and is mediated by calcineurin B‐like (CBL) proteins and their downstream CBL interacting protein kinases (CIPK). CBL proteins bind Ca2+ via a helix– loop–helix structure motif (the EF hand), and function as Ca2+ signal sensors under conditions of stress in plants. The specific interactions of d ­ ifferent CBLs with CIPKs can transmit signals to ­downstream proteins such as transcription factors by protein phosphorylation to activate the plant stress response. The major ­component in this signaling pathway is SOS3/CBL4 and its interactive partner SOS2/CIPK24. Disruption of either or both of these genes results in the severe “salt overly sensitive” (SOS) phenotype, suggesting an important role of the SOS3‐SOS2 protein kinase cascade in salinity stress ­signaling ­pathways (see Fig. 22.9). Calcium‐dependent protein kinases (CDPKs) are well characterized and are also of particular interest in plant ­calcium‐mediated signal transduction (Fig.  22.15). CDPKs have both a kinase domain and a calmodulin‐like domain in a single protein. CDPKs phosphorylate their respective ­substrates to transduce perceived signals, and binding of ­calcium can stimulate their protein kinase activity. The identification of protein kinases that activate AREB and ABF transcription factors is important for understanding

Chapter 22  Responses to Abiotic Stress High salinity Ionic stress

Ion Oxidative stress sensors ROS sensors

1067

Dehydration Osmotic stress

Osmosensors (AHK1, etc.)

Ca2+ channel

Ca2+ Phospholipids

ABA ABA receptor Protein phosphatase (PP2C)

Protein kinases (MPK4/6, MPK3)

Protein kinases (CDPK, CIPKs)

Transcription factors

Channels Transporters

Protein kinases (SnRK2)

Transporters

Target stress genes

FIGURE 22.15  Cellular signal transduction cascades in response to high salinity and water deficit. Osmotic change, ionic stress, cold stress (not shown) and reactive oxygen species trigger stress responses, and many stress sensors have been identified. ABA is an important mediator of stress signals, and Ca2+ and phospholipids are second messengers in stress signaling pathways. Protein phosphorylation plays important roles in stress and ABA signal transduction pathway, and involves a number of different protein kinases, including SnRK2 in the ABA response, and MAP kinase, calcium‐regulated protein kinases, and two‐component histidine kinase in the osmotic stress response. ­Transcriptional regulation is most important for the regulation of stress genes, whose products function in both stress response and stress ­tolerance. In addition, the activities of various types of transporters are required for homeostasis during stress conditions.

upstream ABA signaling pathways. One ABA‐activated SnRK2 protein kinase (OST1/SRK2E/SnRK2.6) functions in the ABA signal transduction pathway controlling stomatal closure (Fig. 22.15). SnRK2 is a member of the SNF1‐ related protein kinase family, which contains ten members in Arabidopsis and rice. SnRK2s are activated by dehydration, salinity, and ABA. Several SnRK2s of various plants can phosphorylate AREB and ABF or related proteins in vivo and in vitro, indicating that SnRK2s are upstream ­factors of AREB and ABF. A triple loss‐of‐function mutant of Arabidopsis SnRK2.6, 2.2, and 2.3 exhibits a strong growth defect, which is related to ABA responses such as germination, stomatal closure, and root growth. These three SnRK2 proteins are repressed by ABI1‐related PP2C protein phosphatases, which are major negative components in the ABA signaling pathway. The ABI1‐related PP2Cs directly and negatively regulate the activities of these three SnRK2 via dephosphorylation. The pyrabactin resistance (PYR)/PYR‐related (PYL)/­ regulatory component of ABA receptor (RCAR) family of ABA receptors (PYR/PYL/RCAR protein family) have ABA‐binding activity and inhibit PP2C activity by ABA‐

mediated interaction (Fig.  22.16). Furthermore, ABA‐­ activated SnRK2 protein kinases are inhibited in loss‐of‐ function mutants of PYR/PYL/RCARs. A signaling complex of PYR/PYL/RCAR, PP2C, and SnRK2 is important for ABA perception and the initial signal transduction process. PYR/PYL/RCARs function as cytoplasmic ABA receptors and inhibit PP2C‐mediated inactivation of SnRK2 in an ABA‐dependent manner (Fig. 22.16). Under normal growth conditions ­without ABA, PP2C inhibits SnRK2 by direct binding and dephosphorylation of SnRK2. In response to abiotic stress, endogenous ABA is synthesized and binds to the PYR/PYL/RCAR receptor, and ABA‐bound PYR/PYL/ RCAR inhibits PP2C activity. This activates SnRK2, which then phosphorylates its target proteins, such as AREB/ABF bZIP transcription factors and ion transporters. The structure of the ABA‐PYR/PYL/RCAR‐PP2C complex was determined by X‐ray ­diffraction (Fig.  22.17). An ABA‐bound receptor is capable of competitively inhibiting the phosphatase activity of PP2C. Perception of ABA and its stream signal transduction are fully understood at the molecular level to illustrate the precise molecular switch of ABA ­signaling (Fig. 22.16).

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SnRK3 protein kinase was identified (see Section  22.3.5). In  addition, genetic screens have been carried out using ­various reporter gene constructs to identify mutants with altered expression of stress‐induced genes (see Box 22.1).

Environmental stimuli Developmental cues

PYR/PYL/ RCAR

ABA

22.4  Freezing and chilling stress

ABA

22.4.1  Freezing and chilling stress have both similarities and differences P

P

AB

A

PP

2C

P P

SnRK2

SnRK2

PP

2C

P

Other factors

AREB/ABF

ABRE Target genes

P

Other responses (e.g. ion transport)

ABA responses Stress responses Seed dormancy etc.

FIGURE 22.16  Proposed model of the early process of ABA p­ erception and signal transduction. PYR/PYL/RCAR functions as an ABA receptor, and PYR/PYL/RCAR, PP2C phosphatase and SnRK2 protein kinase form a signaling complex. In the absence of ABA, PP2C binds to and inactivates SnRK2. In the presence of ABA under abiotic stress conditions, PYR/PYL/RCAR binds to ABA and inhibits PP2C activity. Then, SnRK2 is released from the inhibition by PP2C and activated to phosphorylate downstream transcription factors, including AREB/ABFs and transporters. These activated proteins then function in cellular and molecular responses to abiotic stress in an ABA‐dependent manner.

22.3.6  Genetic screens have identified stress signaling proteins Genetic analyses have also revealed genes involved in the ­signal transduction pathways that control stress‐responsive gene expression. Various mutants with tolerance to abiotic stressors have been isolated and some of the mutant genes have been identified. For example, SOS mutants were isolated by screening salinity‐hypersensitive mutants with root response to salinity stress, and the gene encoding a CIPK/

Freezing and chilling stress impose both direct and indirect effects on plant health. Direct effects include solidification of membrane lipids and reductions in enzymatic reaction rates, and these occur over a relatively short time. Indirect (or ­secondary) injury symptoms, on the other hand, appear g­radually over time  and include solute leakage from cells, respiration and ­photosynthesis imbalance, ATP depletion, accumulation of toxic substances, and wilting by water loss (see Chapter 8). Both freezing and chilling stress are associated with some form of cellular injury caused by the direct effects of low ­temperature; however, freezing stress has several additional indirect effects due to ice crystal formation and growth in extracellular compartments that can damage plants at the whole‐plant, tissue, and cellular levels. In addition, freezing of water in the extracellular spaces results in osmotic dehydration and, therefore, increases in solute concentrations inside the cell. These damaging effects together markedly affect plant performance, and susceptible plants will die. Another difference between the two forms of temperature is the time effect: while chilling injury occurs some time after exposure of plants to the stress (the period differs between plants, organs, tissues, and developmental stage), freezing injury may be visible instantly after freezing occurs. Thus, these two low‐temperature stressors must be considered distinct and analyzed individually with suitable methods.

22.4.2  Chilling stress causes membrane destabilization and metabolic dysfunction Membrane lipids maintain a fluid (liquid crystalline) phase at normal, warm temperatures, which ensures maintenance of cellular function (see Chapter 8). When the temperature decreases, however, lipids with high melting temperatures begin to solidify (gel phase) and become phase‐separated within the membrane. Membranes become leaky or otherwise dysfunctional, intracellular water and solutes are lost, and membrane‐associated ­reactions such as carrier‐mediated transport, enzyme‐mediated processes, and receptor function are inactivated. Although low temperature affects plant cells in many ­different ways, one of the most important is its effect on photosynthesis. Low temperature markedly impairs the ­ electron transport chain while showing little effect on ­

Chapter 22  Responses to Abiotic Stress

PYR/PYL/RCAR

Trp

ABA

Latch loop Gate loop

Trp

FIGURE 22.17  Three‐dimensional s­ tructure of the complex of ABA‐bound PYR/PYL/RCAR (ABA receptor) and ABI1 (PP2C). PP2C is represented by both a ribbon (left) and surface (right) model. ABA‐bound PYL1 can competitively inhibit the ­phosphatase activity of PP2C using the gate loop like a plug.

Gate loop Catalytic site

PP2C

­ hotoenergy reception. As a result, chloroplasts are exposed p to excess excitation energy, and photoreduction of oxygen molecules occurs with concomitant production of ROS (see Section  22.7). The resultant oxidative stress with increased free radical levels damages membrane lipids and proteins as well as macromolecules in chloroplasts. The development of injury symptoms sometimes becomes more apparent after a return to normal temperatures. At low temperature, the development of symptoms requires a long time, and injury may not advance due to the slow rate of the dysfunctional process. Several studies have shown that ­chilling injury occurs due to the lack of or impaired cellular r­ ecovery functions. Although the D1 protein, a major component of the PSII reaction center, is affected at low temperature, the major cause of chilling injury to the PSII is the lack of recovery from damage caused by ROS after returning to warmer temperatures. ROS scavengers such as glycine betaine, a compatible solute (Figure 22.5), may increase the efficiency of PSII metabolic turnover during the recovery process from abiotic stressors. This hypothesis is supported by genetic engineering studies that indicate overexpression of ROS‐scavenging enzymes improves low temperature tolerance.

22.4.3  Freezing also causes membrane destabilization and damage due to osmotic and mechanical stress As explained in Section 22.4.1, freezing has a marked impact on cellular water and causes a type of water deficit stress that distinguishes it from the effects of chilling. The chemical

potential of ice is lower than that of unfrozen water. In addition, the vapor pressure of extracellular ice is lower than that of the water in the cytoplasm or v­ acuole. As ice forms in extracellular compartments, the c­ ellular water moves down the water potential gradient, across the plasma membrane and toward the extracellular ice (Fig. 22.18). As a result, cell volume decreases and intracellular solute concentration increases; extracellular ice formation also distorts the shape of cells. These types of stress persist as long as freezing continues, and they even increase when ­temperature decreases, causing further injury to plant cells. The plasma membrane is the primary site of freezing‐ induced injury, and much of the damage involves membrane destabilization caused by cellular dehydration. Membrane structures and interactions are altered when freezing‐induced dehydration brings the plasma membrane into close ­apposition with the membranes of organelles, such as the chloroplast, leading to membrane destabilization. Membrane destabilization results in distinct forms of injury in some plants, such as expansion‐induced lysis, loss of osmotic responsiveness associated with lamellar‐to‐hexagonal II phase transition in less tolerant (nonacclimated) cells, and loss of osmotic responsiveness associated with fracture‐jump lesion in more tolerant (cold‐acclimated) cells (Fig. 22.19). Membrane destabilization also results from the osmotic and mechanical stress imposed during freeze–thaw cycles. Osmotic dehydration increases solute concentrations in the cytoplasm and other intracellular compartments, and this can inactivate membrane‐associated enzyme and transporter activities. Direct interaction of solutes with the membrane results in dissociation of membrane proteins due to changes in electrostatic and hydrophobic interactions. In addition,

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Freezing temperature H2O

Cell wall

Plasma membrane

Ice formation in cell wall

FIGURE 22.18  Exposure of plants to freezing temperatures causes a cellular water deficit as water travels down its potential gradient, crossing the plasma membrane into the cell wall and intercellular spaces. When the rate of freezing is sufficiently slow to prevent formation of ice crystals in the cytoplasm, the cell dehydrates and freezing occurs in the apoplast.

Freeze dehydration

Thawing

Contraction

Expansion

A Expansion-induced lysis (EIL) with endocytotic vesiculation

Freeze dehydration

Thawing

Contraction

No expansion

B Loss of osmotic responsiveness with endocytotic vesiculation and hexagonal II phase (LOR-HII)

Freeze dehydration

Thawing

Contraction

No expansion

C Loss of osmotic responsiveness with exocytotic extrusions and the fracture-jump lesion (LOR-FJL)

FIGURE 22.19  Freezing‐induced lesions associated with the plasma membrane in isolated protoplasts. (A) When a suspension of protoplasts isolated from nonacclimated plants is frozen, freezing‐induced dehydration occurs in the protoplast, gradually reducing protoplast volume. To maintain membrane tension, the plasma membrane must be removed from the surface of the protoplast, and this is accomplished through formation of endocytotic vesicles that are not continuous with the plasma membrane. Under mild but injurious dehydration conditions, the protoplasts cannot expand during thawing due to the lack of the plasma membrane materials, resulting in expansion‐induced lysis. (B) When severe freezing‐induced dehydration occurs at lower temperature, the plasma membrane of the protoplast is brought into close apposition to endomembranes, ultimately resulting in irreversible ultrastructural changes of the lipid bilayers (such as lamellar‐to‐hexagonal II phase transition) due to membrane–membrane interactions. Consequently, the protoplast does not respond osmotically when thawed (loss of osmotic responsiveness). (C) In protoplasts isolated from cold‐acclimated plants, freezing‐induced dehydration results in the reduction of their volume, but the plasma membrane forms exocytotic extrusions that are continuous with the plasma membrane. No lysis occurs upon thawing of protoplasts that were frozen at any temperature. Membrane– membrane interactions occur when the protoplast is subjected to severe freezing‐induced dehydration, but a distinct structure associated with the plasma membrane, the fracture‐jump lesion (FJL), occurs. The FJL is a result of membrane–membrane fusion and is manifested as a jump of the ­fracture plane from one membrane to the other in freeze‐fracture electron micrographs. Consequently, the protoplast does not respond osmotically when thawing (loss of osmotic responsiveness associated with the fracture‐jump lesion, but not with the hexagonal II phase formation).

Box 22.1

A genetic screen was used to identify mutants with altered RD29A gene expression in response to abiotic stress

G

polyphosphate 1‐phosphatase, which is involved in inositol‐1,4,5‐triphosphate signaling. FRY1 functions as a negative regulator of ABA and stress signaling, suggesting the involvement of phospholipids in stress signaling pathways. Another HOS gene, HOS15 encodes a WD40 repeat protein similar to   TBL1, which is associated with histone deacetylation in mammals. HOS15 is likely involved in histone H4 deacet­ ylation to regulate stress tolerance through chromatin remodeling. Repressor of silencing (ros) has been identified in other mutants. ROS1 encodes a DNA glycosylase that demethylates DNA by base excision repair and can counteract RNA‐dependent DNA methylation. ROS3 contains an RNA recognition motif and may direct sequence‐specific demeth­ ylation by ROS1 and related DNA demethylases. Therefore, this mutant screening approach provides a powerful tool not only for analysis of upstream signal transduction pathways, but also for studies of post‐transcriptional regulation of gene expression and chromatin regulation.

enetic screens have been used to identify genes involved in the plant abiotic stress response. Arabidopsis plants were transformed with the firefly luciferase gene fused to the RD29A promoter, which is induced by drought, cold, and ABA treatment. Arabidopsis plants containing the RD29A promoter‐luciferase (LUC  ) transgene were then mutated by EMS or T‐DNA insertion for mutant screening. Mutants were isolated that exhibited constitutive expres­ sion (cos), high expression (hos), or low expression (los) of luciferase in response to cold and salt stress treatments (see figure). The proteins encoded by the mutated genes function not only as transcription factors upstream of RD29A gene activation, but also in posttranscriptional regulation of the transcription factors. Many factors involved in stress signaling were identified among the mutants isolated. For example, LOS5 and LOS6 were identified as ABA3 (encoding molybdenum cofactor sulfurase) and ABA1 (encoding zeaxanthin epoxidase), respectively. These are involved in ABA biosynthesis. One of the HOS genes, named FRY1 (FIERY 1), encodes inositol

Source: Ishitani et al. (1997). Plant Cell 9:1935–1949. No stress

hos to cold

los to cold hos and to NaCI NaCI and ABA los to cold

los to all

hos to all

cos WT

A

D

B

Cold

E

C

ABA

F

NaCI

Luminescence images of wild‐type (RD29A promoter:LUC transgenic Arabidopsis plant) and mutants (hos, los, and cos) under cold, osmotic, and ABA stress. (A) Arrangement of the wild‐type (WT) and mutant lines. Clockwise: wild‐type RD29A‐LUC transgenic parent; cos, mutants with constitutive RD29A expression; los to all, mutants with reduced responses to cold, ABA, and osmotic stress; hos to cold, mutants with enhanced response to cold; los to cold and NaCI, mutants with reduced responses to cold and osmotic stress; hos to NaCI and ABA, mutants with enhanced response to ABA and osmotic stress; los to cold, mutants with reduced response to cold stress; hos to all, mutants with enhanced response to cold, ABA, and osmotic stress. (B) Morphology of the seedlings of WT and mutants. (C) Luminescence from plants before stress and ABA treatments. Note the constitutive luminescence of the cos mutant. (D) Luminescence from plants exposed to cold conditions for 2 days. (E) Luminescence from plants after ABA treatment. The plants were placed at room temperature (20–22°C) for 2 days to allow the luminescence to drop to pre‐cold treatment levels before spraying with 100 μM of ABA and imaging 3 hours later. (F) Luminescence from plants under osmotic stress. Three days after ABA treatment, luminescence from the plants dropped to pretreatment levels. The plants were then treated with high salt for 5 hours by flooding the plate with 300 mM NaCI. Some of the cos mutant seedlings died after cold and ABA treatment and so did not emit luminescence upon NaCI treatment.

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solute concentration affects lipid phase behavior due to the electrostatic interactions with charged headgroups of bilayer lipid molecules. Mechanical stress is another factor that destabilizes the membrane; ice formation in the extracellular space results in deformation of the cell, and ice crystals can directly damage the plasma membrane. Freezing stress also causes other types of damage in cells. Cell wall–plasma membrane interactions are altered due to freezing; for example, in some plants, survival of intact cells is less than that of isolated protoplasts, which do not have a cell wall. This is because the presence of the cell wall can add extra stress from extracellular ice crystals on cells. Further, several specific proteins become detached from the plasma membrane when intact cells with a cell wall are frozen, suggesting the physicochemical interactions between the cell wall and the plasma membrane occurring during freezing cause extra damage to the plasma membrane. Freezing leads to acidification of the cytoplasm, probably as a result of disturbance of H+‐transport systems associated with the vacuolar membrane (tonoplast). This acidification markedly affects metabolic reactions in the cytoplasm. Taken together, these ­observations indicate that freezing exposes cells to very complex multifaceted stress, and the rate of freezing and the temperature and rate at which ice crystals form can strongly affect the extent of damage to the cell.

22.4.4  Plants have multiple low‐ temperature sensing systems To adapt to low temperature, plants must first perceive changes in temperature. The cyanobacterium Synechocystis PCC6803 has a putative low‐temperature sensor, the h ­ istidine kinase Hik33. Low temperature activates Hik33 by s­ignals mediated by changes in membrane fluidity, leading to autophosphorylation and subsequent phosphate transfer to the response regulator Rre26. Microarray data have shown that HiK33 regulates the expression of 21 of 36 cold‐­regulated genes. Thus, Hik33 is considered a low temperature regulator that activates expression of the majority of cold‐regulated genes, in addition to another temperature‐sensing pathway involved in cellular responses of the cyanobacterium to low temperature. Although no Hik33 orthologs have been found in higher plants, two‐component systems like the combination of His and Rre are involved in responses to some ­phytohormones (see Chapter 18). The participation of these two‐component systems in low temperature sensing in plants remains to be determined. There are a number of candidate low temperature sensors in plants, although these have not been confirmed. For ­example, changes in membrane fluidity are considered to play a role in sensing a temperature drop outside the cell. Alterations in membrane fluidity of alfalfa cells using treatments with dimethylsulfoxide as a membrane fluidizer and benzyl alcohol as a membrane stabilizer result in changes in both freezing tolerance and cold‐induced gene expression.

Calcium influx and cytoskeleton organization are also affected under these conditions, so changes in membrane ­fluidity may act as sensors and induce a series of signal ­transduction events at low temperatures.

22.4.5  Plants can acclimate to freezing by increasing membrane stability The ability to survive temperatures below freezing is genotype‐specific. Whereas many important crop plants, ­ including corn (Zea mays), tomato (Lycopersicon esculentum), and rice (Oryza sativa), are unable to withstand freezing temperatures, many temperate and sub‐arctic plants can ­ ­survive freezing temperatures, and some can even survive temperatures below –40°C. However, these plants are not ­tolerant to freezing throughout the growing season. Freezing tolerance develops in a process known as cold acclimation, a response to low but nonfreezing temperatures that occur before freezing. As an example, Arabidopsis can normally survive temperatures as low as –5°C, but after ­exposure to temperatures in the range of 1°C to 5°C for one to seven days, it can survive much lower temperatures of –8°C to –13°C. This makes Arabidopsis a good model for studying the mechanisms underlying freezing tolerance. A number of seemingly disparate responses occur during cold acclimation, including alterations in membrane composition and accumulation of compatible solutes, but these changes ultimately contribute to protection of the plasma membrane, the p ­ rimary site of freezing injury in plants. The most notable changes in the plasma membrane after cold acclimation are the increase in proportion of plasma membrane phospholipids (see Chapter 8), which is widely observed in ­various herbaceous and woody plants, and concomitant decrease in the proportion of glucocerebrosides. These cold‐induced changes in lipid composition improve water ­retention at the membrane surface, which can prevent thermotropic and lyotropic lipid phase alterations in the membrane bilayers during freezing. Comprehensive lipid analysis, however, does not support the suggestion that there is a single lipid component that predominantly affects maintenance of plasma ­membrane integrity during freeze–thaw cycles. Rather, altered lipid–lipid (or lipid–protein) interactions after cold acclimation should contribute to the increased plasma ­membrane stability against freezing stress. Plasma membrane proteins and microdomains also respond dynamically to cold acclimation. Cold‐responsive proteins in the plasma membrane include osmotic and other stress‐related proteins, proteolysis‐associated proteins, and membrane trafficking proteins. Plasma membrane microdomains are enriched in glucocerebrosides and sterol lipids, and the proportions of both lipids change after cold acclimation. Cold acclimation also results in changes in microdomain ­proteins, such as those associated with membrane transport, membrane trafficking, and cytoskeleton–plasma membrane interactions. Thus, microdomain analysis is a promising

Chapter 22  Responses to Abiotic Stress

Ca2+

Ice

Ca2+

Ice

Ca

2+

Ca2+

Ca2+

Ice

Ca2+ Ca

Ice Ca2+

2+

SYT1

C2A

Ca2+

Resealing

C2B

C2A

AntiSYT1

Ca2+

C2B

C2A Punctured plasma membrane

Ca2+

C2B

C2B

C2A

Cell wall

Cytosol

A

B

C

D

FIGURE 22.20  Model for Ca ‐ and SYT1‐induced membrane resealing occurring during freeze/thawing. (A) The plasma membrane is mechanically punctured by ice crystals, and (B) Ca2+ moves from the extracellular space into the cytoplasm through the damaged sites. (C) Endomembranes may then fuse at the site of the damaged plasma membrane via Ca2+ binding SYT1. (D) The damaged site is resealed. 2+

means of determining the molecular and functional responses of the plasma membrane to low temperature and freezing ­tolerance in plants. Although not analyzed in detail, some cold‐responsive proteins in the plasma membrane do affect freezing tolerance. Phospholipases influence freezing tolerance probably through alterations in the plasma membrane lipid composition and phospholipid‐mediated signaling. In Arabidopsis, antisense suppression of phospholipase Dα1 (PLDα1), the most abundant plant phospholipase, increases freezing tolerance. The difference in freezing tolerance is probably because suppression of PLDα1 inhibits an increase in the proportion of ­phosphatidic acid that promotes destabilizing phase ­transition of the membrane bilayers. In contrast, Arabidopsis ­phospholipase Dδ (PLDδ) knockout lines show reduced freezing tolerance, and overexpressors show increased f­reezing tolerance. PLDδ modifications do not change the proportion of major phospholipids in the plasma membrane during freezing but selectively increase molecular species of phosphatidic acid, suggesting that specific phosphatidic acid species act as signal transducers of low temperature. Another plasma membrane‐associated protein that affects freezing tolerance is synaptotagmin1 (SYT1). Synaptotagmins are a family of membrane‐trafficking proteins that function as calcium sensors in plasma membrane vesicle fusion p ­ rocesses mediated by the SNARE protein complex. The level of Arabidopsis SYT1 increases rapidly in the plasma ­membrane in parallel with the development of freezing tolerance during cold acclimation. As Arabidopsis freezing tolerance is enhanced by exogenous application of calcium, and both Arabidopsis SYT1‐RNAi lines and T‐DNA insertion mutants clearly have less ­freezing tolerance than wild‐type plants, SYT1‐dependent membrane resealing in the presence of c­ alcium is considered a vital component of plant freezing t­olerance (Fig. 22.20). Genetic screens for mutants showing enhanced osmotic sensitivity have also identified SYT1 as an important component in osmotic tolerance.

22.4.6  Cold‐induced changes in metabolite profiles have important roles in cold acclimation Compatible solutes, such as simple sugars (sucrose, glucose, fructose, raffinose, and stachyose), proline, and glycine betaine, accumulate in conjunction with the development of freezing tolerance in many plant species. The deduced m ­ olecular functions of compatible solutes in freezing stress tolerance are similar to those in osmotic and dehydration stress (see Sections 22.2.3 and 22.2.4). The Arabidopsis mutant eskimo1, which is constitutively freeze‐tolerant, overproduces proline at warm temperatures. The ESKIMO1 gene encodes a 57‐kDa protein of unknown function belonging to a large gene family of DUF231 proteins. The eskimo1 mutation alters the expression of genes that overlap with those regulated by other abiotic stressors, including salt, osmotic, and cold stress, but that are apparently independent of those mediated by CBF/DREB1 transcription factors (see Section  22.4.7). Sugars have been suggested to protect membranes and contribute to improved freezing tolerance after cold acclimation. In fact, exogenous sugar application has been used to recover the impaired increase in freezing tolerance of Arabidopsis sensitive‐to‐freezing 4 (sfr4) mutant. However, in some cases, the accumulation of sugars alone is not sufficient for the development of freezing tolerance. Several single‐gene mutants of Arabidopsis that are defective in freezing tolerance nonetheless accumulate sugars normally in response to low‐temperature stress.

22.4.7  Freezing tolerance involves changes in gene expression In most cases, the cold acclimation‐induced alterations described above are mediated by changes in gene expression at low temperature. Since the mid‐1980s, when changes in gene expression in response to cold acclimation were first

1073

HVA1 (Hordeum vulgare ABA‐induced) D‐7 (cotton)

Group 3 (D‐7 family*)

LE25 (tomato) D‐113 (cotton)

Group 5 (D‐113 family*)

*The protein families are named for seed proteins from cotton.

D‐95 (soybean)

Group 4 (D‐95 family*)

+ + + + + + + + + + + +

DHN1 (maize) D‐11 (cotton)

Group 2 (D‐11 family*)

+ + + + + + + + + + + +

Em (early methionine‐ labeled protein, wheat)

Representative proteins

Group 1 (D‐19 family*)

Hypothetical structures

TABLE 22.1 The five groups of LEA proteins.

Shares sequence homology at the conserved N terminus, is predicted to form α‐helix C‐terminal domain is predicted to be a random coil of variable length and sequence Rich in alanine, glycine, and threonine

Hydropathy plots are unremarkable and slightly hydrophobic N‐terminal region contains a possible amphipathic α‐helix

Contain repeated motifs of the consensus sequence TAQAAKEKAXE Predicted to contain amphipathic α‐helices Predicted to form dimers

Structure variable Includes one or more conserved lysine‐rich regions that may form α‐helices. The consensus sequence is EKKGIMDKIKELPG. The number of repeats per protein varies May or may not contain a poly(serine) region Contain regions of variable length rich in either glycine and threonine or glutamate and lysine

Most (70%) protein conformation is random coil with some predicted short α‐helices Rich in charged amino acids and glycine

Structural characteristics and shared motifs

D‐113 is abundant in cotton seeds (up to 0.3 mM)

A gene encoding a similar protein in tomato is expressed in response to nematode feeding

D‐7 is abundant in cotton embryos (0.25 mM) Putative dimer of D‐7 may bind as many as 10 inorganic phosphates and their counterions

Most members localize to cytoplasm and nucleus, but also associated with plasma membrane

More hydrated than most globular polypeptides

Properties

May bind membranes or proteins to maintain structural integrity May sequester ions to protect cytosolic metabolism LE25 confers salt and freezing tolerance to yeast

HVA1 promotes stress tolerance in transgenic plants

May stabilize macromolecules under conditions of reduced water content

Binds water to minimize loss of cellular water content Overexpression confers water‐deficit tolerance on yeast cells

Proposed function

Chapter 22  Responses to Abiotic Stress

observed in spinach (Spinacia oleracea), numerous genes responsive to low temperature treatments have been ­identified. Unfortunately, no clear functional roles have yet been determined for many of these genes. Several of the genes induced by low temperature are also induced by water deficit or ABA (see Section  22.3). Gene products that are hydrophilic and soluble when boiled, including Group 2 and Group 3 LEA proteins (Table  22.1) and the products of other unique genes, accumulate during cold acclimation. One such unique gene product is COR15a, a small, nuclear‐encoded hydrophilic protein that is targeted to the chloroplast. When imported into the chloroplast, the protein is processed to form the mature protein, COR15am, a 9.4‐kDa polypeptide. Protoplasts of nonacclimated ­transgenic Arabidopsis plants that constitutively express the COR15a gene show increased freezing tolerance. COR15am has been suggested to alter the intrinsic curvature of the chloroplast envelopes and protect chloroplast proteins through direct associations with each other. As described in Section 22.3, many of the cold‐regulated genes contain a DRE or CRT cis‐acting DNA element in their promoter regions that binds CBF/DREB1 transcriptional activators (see Fig. 22.12). Three CBF/DREB1 genes (CBF1/ DREB1b, CBF2/DREB1c, and CBF3/DREB1a), which each contain sequences encoding a 60 amino acid DNA binding AP2/ERF domain, are present in tandem array on Arabidopsis chromosome 4 and are induced rapidly (within 15 minutes) upon exposure to low temperature, with peak expression after 2 hours. Constitutive or stress‐induced CBF/DREB1 expression (with constitutive CaMV 35S promoter or dehydration‐ and cold‐inducible RD29A promoter, respectively) in Arabidopsis increases the abundance of COR and other DRE/ CRT‐containing target gene transcripts in nonacclimated plants, which in turn increases freezing tolerance. Overexpression of the transcriptional activator CBF/ DREB1s enhances freezing tolerance to a greater extent than overexpression of COR15a alone. Further, CBF/DREB1 genes are widely conserved in plants, and the number of these genes and the extent of their expression are correlated with the degree of freezing tolerance in some plant groups. Taken together, these results suggest a role of cold‐regulated genes in the development of tolerance to low temperature and have facilitated new research on protection of crop plants against freezing using overexpression of transcription factors. Microarray analysis of plants overexpressing CBF/DREB1 has identified its target genes. The number of direct CBF/ DREB1 target genes is around 40, and about 100 genes have been assigned to the CBF/DREB1 regulon. The CBF/DREB1 regulon genes encode proteins that fall into two groups: proteins that function in stress tolerance and those that are involved in further regulation of the expression of other genes. COR proteins, along with enzymes involved in proline metabolism (Δ1‐pyrroline‐5‐carboxylate synthase) and raffinose synthesis (galactinol synthase, GolS), belong to the CBF/ DREB1 regulon and participate in enhancement of freezing tolerance. The presence of proteins with regulatory functions in the CBF/DREB1 regulon, such as transcription factors,

suggests there may be some subregulons within the CBF/ DREB1 regulon. In fact, microarray experiments and ­subsequent bioinformatics analysis have revealed that approximately 20% of the CBF/DREB1 regulon genes do not have the core DRE/CRT sequence (CCGAC) in their promoter sequences. Thus, there must be complex, elaborate gene ­regulatory networks within the CBF/DREB1 regulon that control the development of freezing tolerance during cold acclimation. The circadian system regulates the expression of CBF/ DREB1 as well as some CBF/DREB1 regulon genes. The extent to which CBF/DREB1 transcripts accumulate upon exposure to low temperature varies with the time of day that the plants are transferred to low temperature. Cold‐induced CBF/DREB1 transcript accumulation reaches the maximum level at 4 hours and the minimum at 16 hours after subjective dawn. Accordingly, CBF/DREB1 target genes do in some cases show circadian rhythmic oscillation. This is of interest because freezing tolerance shows rhythmic changes within the day. However, the detailed mechanisms underlying the circadian rhythmicity of plant freezing tolerance remain to be determined. The Inducer of CBF expression (ICE1) gene has been ­identified as a regulator of CBF3/DREB1A gene expression. ICE1 encodes a MYB‐like basic helix–loop–helix (bHLH) transcription factor and is expressed constitutively, ­suggesting that cold‐induced posttranslational modification is necessary to activate ICE1 for CBF3/DREB1A expression. ICE1 overexpression enhances expression of the CBF/DREB1 ­ ­regulon genes and increases freezing tolerance in plants after cold acclimation (see Fig. 22.12). Thus, ICE1 functions as an upstream regulatory protein to control CBF3/DREB1A ­transcription. ICE1 also functions in the regulation of s­ tomata numbers in leaf epidermal cell development. However, it is not involved in the cold induction of other CBF/DREB1 genes. It has been proposed that the involvement of calcium in CBF2/DREB1C expression is mediated by a calmodulin‐ binding transcription factor (CAMTA) (see Fig. 22.12). The CBF/DREB1 cold‐responsive pathway likely plays an important role in cold acclimation in plants, but additional pathways must also be involved in activating cold‐induced reactions in response to low temperature. The eskimo1 mutant, which is constitutively freezing tolerant, does not show constitutive CBF/DREB1 gene expression. Similarly, the sfr4 mutant, which shows impaired freezing tolerance enhancement during cold acclimation, does not differ in CBF/DREB1 gene expression on exposure to low temperature (see Section 22.4.5). The ZAT12 cold‐responsive pathway also plays a role in cold ­acclimation. Overexpression of ZAT12, which encodes a zinc-­ finger transcriptional repressor protein, alters gene expression profiles and increases freezing tolerance of plants. The ZAT12 cold‐responsive pathway appears to interact with the CBF/ DREB1 cold‐responsive pathway: some target genes are shared by the two pathways and some genes in the CBF/DREB1 pathway are downregulated by ZAT12 overexpression, which suggests complicated regulatory systems for genes associated with freezing tolerance.

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22.5  Flooding and oxygen deficit Flooding (waterlogging of soil to complete submergence) can also trigger a number of different stress responses in plants. Whereas oxygen concentrations in well drained, porous soil are nearly equal to atmospheric concentrations (20.6% ­oxygen, 20.6 kPa), the diffusion coefficient of oxygen in water is four orders of magnitude lower than that in air. When flooding occurs, soil gases are replaced with water, thereby reducing entry of oxygen into the soil and making it difficult for roots and other organs to carry out respiration. Like most eukaryotic organisms, plants are obligate ­aerobes that derive most of their energy from mitochondrial respiration of hexose sugars to ATP. Under normal aerobic conditions, plants oxidize 1 mol of hexose sugar through ­glycolysis, the citric acid cycle, and oxidative phosphorylation to yield 30–36 mol of ATP (see Chapters 13 and 14). In the absence of oxygen, however, plants produce ATP mainly by glycolysis, which yields only 2–4 mol of ATP per mole of ­hexose sugar. The function of a noncyclic flux mode of citric acid cycle can boost the ATP output by 1 mol per pyruvate metabolized (see Section  22.5.2). Cytoplasmic ATP/ADP ratios decline as mitochondrial ATP production is inhibited. Ironically, oxygen deficit associated with flooding can also prevent plants from obtaining adequate water from the soil due to gating of root cell aquaporins; this reduces the permeability of root cells to water and limits transport of water to aerial tissue. The supply of oxygen to root cells is influenced by several factors, including soil porosity, water content, temperature, root density, and the presence of competing algae and aerobic TABLE 22.2 Impact of oxygen deprivation on respiratory metabolism. Oxygen status

Effect on metabolism

Normoxic (aerobic)

Aerobic respiration proceeds normally; almost all ATP production results from oxidative phosphorylation

Hypoxic

Partial pressure of O2 limits ATP production by oxidative phosphorylation; glycolysis and a non‐cyclic flux mode of the citric cycle account for a larger percentage of ATP yield than under normoxic conditions; metabolic and developmental changes are stimulated that result in adaptation, acclimation to a low‐oxygen environment

Anoxic (anaerobic)

ATP is produced only by glycolysis and a non‐cyclic flux mode of the citric cycle; cells exhibit low ATP content, diminished protein synthesis, and impaired division and elongation. If anoxic conditions persist, many plant cells die

microorganisms. Oxygen concentrations in root tissues also vary according to root depth, root thickness, the volume of intercellular gaseous spaces, and cellular metabolic activity. Oxygen gradients exist within other organs and tissues, including meristems, tubers, and developing seeds. Plant or cellular oxygen status can be defined as normoxic, hypoxic, or anoxic (Table 22.2). To survive short‐term flooding, plants must generate sufficient ATP, regenerate NADP+ and NAD+, and avoid accumulation of toxic metabolites. Periods of oxygen deficit can trigger developmental responses that promote acclimation to hypoxic or anoxic conditions.

22.5.1  Plants vary in their ability to tolerate flooding Plants can be generally classified as wetland, flood tolerant, or flood sensitive, according to their ability to withstand periods of soil flooding or submergence (Table 22.3). Wetland plants possess anatomical, morphological, and physiological features that permit survival in waterlogged soils and partial submergence. Growth in a wetland environment promotes formation of a thickened root hypodermis to reduce O2 loss to the anaerobic soil. To facilitate transport of O2 from aerial structures to submerged roots and thereby maintain aerobic metabolism and growth, some plants develop specific structures: aerenchyma (continuous, ­columnar intracellular spaces formed in root cortical tissues (Fig.  22.21; see also Section  22.5.4 and Chapter  20), ­adventitious roots from the hypocotyl or stem (Fig.  22.22), lenticels (openings in the TABLE 22.3 Plant species categorized by sensitivity to flooding. Wetland

Flood tolerant

Flood sensitive

Acorus calamus

Arabidopsis thaliana

Glycine max (soybean)

Echinochloa crus‐galli (rice grass)

Echinochloa crus‐pavonis (barnyard grass)

Lycopersicon esculentum (tomato)

Echinochloa phyllopogon (barnyard grass)

Rumex acetosa (sorrel)

Pisum sativum (pea)

Erythina caffra (coral tree)

Solanum tuberosum (potato)

Rumex palustrus (marsh dock)

Zea mays (corn)

Oryza sativa (rice)

Triticum aestivum (wheat)

Phragmites australis (common reed)

Hordeum vulgare (barley)

Chapter 22  Responses to Abiotic Stress

A

Endodermis

Hypodermis

B

Lanuca

Endodermis

Hypodermis

FIGURE 22.21  Aerenchyma development in the root cortex of maize (Zea mays L.) after oxygen deprivation. Photomicrographs of transverse s­ ections of maize roots maintained under aerobic conditions (A) or under 72 hours of hypoxia (B) demonstrate the formation of cortical ­aerenchyma in hypoxic roots. The hypodermis and endodermis remain intact, and the lacunae created by the death of the central cortical cells form columnar gas‐conducting chambers. Source: He et al. (1996). Plant Physiol. 112:463–472.

Lenticels

Adventitious root

FIGURE 22.22  Adventitious roots and prominent (hypertrophied) l­enticels on the stem of young ash (Fraxinus pennsylvanica Marshall) after flooding. The black arrow indicates the water depth during flooding. Source: Kozlowski (1984) Physiological Ecology. Academic Press, New York.

­ eriderm that allow gas exchange, Fig. 22.22), shallow roots, p and pneumatophores (shallow roots that grow with negative geotrophy out of the aquatic environment, Fig. 22.23). Other adaptive strategies include elongation of stems or leaf petioles towards the water surface (Fig. 22.24) and thinning of leaves to improve underwater photosynthesis. The morphological and anatomical characteristics that improve tissue aeration can be constitutive or induced by flooding. When inundated by water, wetland species alter cellular metabolism in root and aerial organs to increase survival. Partial submergence of deepwater rice (Oryza sativa L. var. Indica) ­promotes formation of adventitious roots and accelerates internodal stem elongation, which enables stems and leaves to be maintained above the aquatic environment

FIGURE 22.23  Pneumatophores of mangrove (Avicennia nitida) develop from roots submerged in estuarine mud. Source: Bowes (1996). A Colour Atlas of Plant Structures. Manson Publishers.

(Fig.  22.25). This escape strategy is associated with ATP ­production through active consumption of available carbohydrates coupled to glycolysis, fermentation, and a partially functioning citric acid cycle. Rice seeds are among the few that can germinate in an anaerobic environment, and germinated rice seedlings ­promote coleoptile elongation towards light and oxygen over root growth. In this situation, a calcineurin B‐like interacting protein kinase (CIPK) and sucrose nonfermenting 1 related

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protein kinase 1A (SnRK1A) sense changes in ATP or sugar homeostasis to promote synthesis of starch catabolizing amylases. In addition, mitochondrial morphology is altered (Fig. 22.26), but functional electron transport chain c­ omplexes of the inner membrane and enzymes of the citric acid cycle in the matrix are maintained. This rapid coleoptile elongation is an escape strategy that is effective if photosynthetic ­activity  can be established before seed starch reserves are fully consumed.

Grown in air

Submerged

FIGURE 22.24  The leaf petiole of Rumex palustris elongates in response to submergence. Source: Photo by Rens Voesenek.

In contrast, other wetland plants, such as the marshland monocot Acorus calamus, respond to flooding by downregulating their metabolism, which allows them to maintain a nearly quiescent state for several months while utilizing the starch reserves stored in their rhizomes. This quiescence strategy, a restriction of metabolism to survive a long and deep flood, is observed in a number of wetland species and submergence-tolerant accessions of rice. Flood‐tolerant plants can endure flooding and anoxia only temporarily. Like wetland species, these plants generate ATP through anaerobic metabolism during short‐term flooding. In most cases, root elongation is inhibited, overall rates of protein synthesis are diminished, and patterns of gene expression are markedly altered. Arabidopsis ecotypes vary in the length of time they can endure complete submergence. Whereas Arabidopsis plants survive two weeks or more, young seedlings of maize, for example, can survive only 3–5 days of submergence, depending on genotype and developmental age. Survival of the low oxygen stress ­associated with waterlogging is prolonged if maize seedlings become hypoxic before experiencing anoxia. Hypoxia ­promotes formation of adventitious and nodal roots with cortical aerenchyma as an acclimation to soils with low ­oxygen content (see Fig. 22.22) and enhances the ability to transport lactate out of roots. Flood‐sensitive plants exhibit an injury response to anoxia. As with flood‐tolerant species, flood‐sensitive plants carry out anaerobic respiration; these plants, however, rapidly ­succumb to flooding because of cytoplasmic acidification. When deprived of oxygen, flood‐sensitive species exhibit greatly diminished protein synthesis, degradation of mitochondria, inhibited cell division and elongation, disrupted ion transport, and cell death within root meristems. Typically, these plants do not develop root aerenchyma and cannot ­survive more than 24 hours of anoxia.

200 Water depth (cm)

1078

100

0 Preflood 2–3 months

A

Flood 4–5 months

Postflood usually 1–2 weeks

B

FIGURE 22.25  Growth responses of seedlings of deepwater rice (Oryza sativa L. var. Indica) to flooding. (A) Seedlings are established before the annual flood. Submergence promotes rapid internodal elongation and development of adventitious roots. Once the flood waters recede, the ­adventitious roots grow into the soil and aerial portions of the plant grow upward. (B) Photographs comparing internode elongation in aerobic (left) and submerged (right) plants. Arrows indicate positions of nodes. Source: (B) Kende et al. (1998). Plant Physiol. 118:1105–1110.

Chapter 22  Responses to Abiotic Stress

A

1079

B

FIGURE 22.26  Photomicrographs comparing mitochondria from rice (Oryza sativa L.) seedlings germinated aerobically and exposed to aerobic (A) or anaerobic (B) treatments for 48 hours. A mitochondrion from anoxic seedlings has well‐developed cristae but its matrix is less dense. Source: Couée et al. (1992). Plant Physiol. 98:411–421.

22.5.2  During short‐term acclimation to anoxic conditions, plants generate ATP through breakdown of carbohydrates, glycolysis, and fermentation Flooding stimulates an increase in glycolytic flux known as the Pasteur effect, where sucrose or glucose from the phloem is directed toward glycolysis in flooded organs. However, in some cases, flooding can restrict the translocation of p ­ hotosynthate from leaves; flooded organs of some ­species hydrolyze stored starch (see Chapter 13) to obtain additional sugars. The availability and mobilization of carbohydrate reserves is essential in oxygen‐deprived organs. Starch is slowly ­hydrolyzed by amylases in rhizomes of flood‐tolerant Acorus calamus and in germinating seeds and leaves of rice plants under oxygen constraints; however, many species must rely on soluble carbohydrate reserves for energy production under hypoxia and anoxia. Sucrose breakdown generally follows an increase in production of UDP‐sucrose synthase (SUS) (Fig.  22.27). Utilizing SUS rather than ATP‐dependent invertase reduces the investment in ATP to produce ­glucose‐6‐phosphate and fructose‐6‐phosphate. Use of pyrophosphate‐dependent phosphofructokinase (PFP) by rice and other species during oxygen deprivation further maximizes glycolytic ATP output. The energy‐yielding steps of glycolysis generate low amounts of ATP while reducing NAD+ to NADH (see Chapter 13). To support ongoing glycolysis in the absence of mitochondrial respiration, the glycolytic substrate NAD+ must be regenerated. The principal end products of glycolysis in oxygen‐deprived plant tissues are lactate and ethanol (Fig. 22.27); alanine, succinate, and γ‐aminobutyrate (GABA) are also be formed. Alanine and succinate are the principal products of a noncyclic flux mode of the citric acid cycle that generates 1 additional mol ATP per mol pyruvate metabolized.

This consumes 1 mol of glutamate and pyruvate to generate 1 mol alanine and 2‐oxoglutarate. The 2‐oxyoglutarate can be used for substrate level ATP production through conversion of succinyl CoA to succinate. Alternatively, 2‐oxoglutarate is shunted to GABA. Oxaloacetate reduced to malate and then fumarate, can produce NAD+ necessary to maintain succinyl CoA ligase. This alternative pathway of pyruvate consumption yields additional ATP. The relative abundance of specific end products varies according to the plant species, genotype, and organ as well as the duration and severity of oxygen deprivation. Both lactate‐ and ethanol‐producing fermentations yield NAD+, but lactate lowers cytosolic pH, whereas ethanol does not. According to the Davies–Roberts lactate dehydrogenase (LDH)/pyruvate decarboxylase (PDC) pH‐stat hypothesis, anaerobic metabolism is regulated by the activities of pH‐­ sensitive enzymes. According to this model, LDH, an enzyme with an alkaline pH optimum, converts pyruvate, which is produced initially by glycolysis, to lactate, and the lactate produced reoxidizes NADH but also lowers the cytoplasmic pH. As the cytosol acidifies, LDH is progressively inhibited and a second enzyme, PDC, is activated. The pH optimum for PDH is lower than the normal cytoplasmic pH; therefore, the accumulation of lactate ultimately stimulates the conversion of pyruvate to acetaldehyde. Alcohol dehydrogenase (ADH) subsequently reduces the acetaldehyde to ethanol while oxidizing NADH to NAD+. Unlike lactate, ethanol is an uncharged molecule at cellular pH and can diffuse across plasma membranes. As a result, the switch to ethanol production stabilizes cytoplasmic pH at a slightly acidic value. Cytosolic pH may also be maintained by operation of the noncyclic flux mode of the citric acid cycle, which is coupled with production of the zwitterionic species alanine from pyruvate. In some plants, the zwitterionic species GABA is formed from pyruvate. Both alanine and GABA

Starch

Sucrose H2O

UDP

SUS

INV

+

Fructose

UDP-glucose

Hexokinase

UTP

+

ATP

P P

ADP

UTP

Glucose-6P

UDP

i

Fructokinase

1 Glucose-1-P UTP

ATP

UDP

Fructose-6P ADP

P P P

ATP

Glucose-1-P

Fructose

or

ADP

i

Glucose

Glucose

Fructokinase

P

Amylases

or

ATP ADP

i

i

Fructose-1,6P2 ADP

Glycolysis

ATP

NADH

Phosphoenol pyruvate

NADH

ADP

AMP + P P

ATP

ATP + P

Lactate

i

NADH

i

Alanine

Pyruvate LDH PDC

Glutamate

NADH

Ethanol

Acetaldehyde

2-Oxoglutarate

Aspartate

Oxaloacetate

MDH

ADN + H

2-Oxoglutarate Acetyl CoA

NH4+ NADP(H)

NADH

Citrate

Malate

GDH Isocitrate

Fumarate

Glutamate H+

NADH

2-Oxoglutarate

Succinate ATP

ATP ATP ATP

NADP(H)

Oxidative phosphorylation + O2

ADP

Succinyl CoA

NAD(H)

GAD

GABA

SCS NADH

Succinic semialdehyde Amino acid

α-Ketoacid

FIGURE 22.27  Metabolic acclimation under O2 deprivation. Plants have multiple routes of sucrose catabolism, ATP production, and NAD(P)+ r­ egeneration. These include ethanol and lactate production as well as a modified noncyclic citric acid flux mode that is both an alanine and 2‐oxoglutarate shunt and a γ‐aminobutyric acid (GABA) shunt. Blue arrows indicate reactions that are promoted during anaerobic stress, and gray dashed lines indicate reactions that are inhibited during the stress. Metabolites indicated in brown boxes are major or minor end products of metabolism under hypoxia. Metabolites indicated in orange boxes decrease under hypoxia. ADH, alcohol dehydrogenase; GAD, glutamic acid decarboxylase; GDH, glutamate dehydrogenase; INV, invertase; LDH, lactate dehydrogenase; MDH, malate dehydrogenase; PDC, pyruvate decarboxylase; SCS, succinyl CoA synthetase; SUS, sucrose synthase.

Chapter 22  Responses to Abiotic Stress

can rapidly re‐enter the citric acid cycle upon reoxygenation, minimizing the net loss of carbon (Fig. 22.27). A different hypothesis predicts that concentrations of pyruvate, rather than cytoplasmic pH, regulate ethanolic ­fermentation. This PDC/pyruvate dehydrogenase (PDH)‐stat hypothesis is based on the observation that the Km of ­cytosolic PDC ranges from 0.25 to 1.0 mM pyruvate, while that of mitochondrial PDH ranges from 50 to 75 μM. Since cellular

Anoxia 7.4

Cytoplasmic pH

7.2 7.0 Wild type

6.8 6.6 6.4 6.2

ADH1– 0 40 min

5h

Lactate fermentation

10

15

25

20

30

Time Ethanol fermentation

Cytoplasmic acidosis

Response of wild type

FIGURE 22.28  Response of root tips of wild‐type maize (solid line) and mutants deficient in ADH1 (dashed line) to anoxia. Wild‐type root tips experience a rapid decrease in cytoplasmic pH, followed by a partial recovery; however, the wild‐type root cells ultimately succumb to cytoplasmic acidosis. Root tips of ADH1‐deficient seedlings show rapid cytoplasmic acidosis and cell death.

No pretreatment

Root death by cytoplasmic acidosis

Glucose

Anoxia Time (h)

24

concentrations of pyruvate are usually less than 0.4 mM, an increase in pyruvate under hypoxia or anoxia stimulates PDC activity. This model is supported by several observations. First, ethanol production can be stimulated under aerobic conditions that inhibit mitochondrial PDH activity. Second, lactate production does not accompany ethanolic fermentation in certain species. Finally, roots of transgenic tobacco plants that have twice the normal activity of PDC produce more ethanol under anoxia than do wild‐type roots. The LDH/PDC pH‐stat and PDC/PDH‐stat hypotheses are not mutually exclusive; the regulation of the switch to ethanolic fermentation may vary according to the species, organ and the conditions of low‐oxygen stress. Plants generally increase cellular levels of PDC and ADH in response to flooding. Mutants of maize and other plants that are deficient in ADH are more sensitive to flooding than are isogenic wild‐type genotypes. In maize, genotypes severely deficient in ADH1 exhibit acetaldehyde accumulation and a more rapid and sustained drop in cytoplasmic pH than does the wild‐type in response to anoxia (Fig.  22.28). However, plants engineered to overproduce PDC or ADH and flood‐ sensitive species such as pea (Pisum sativum) that can rapidly and dramatically upregulate ADH activity still succumb to flooding as a result of exhaustion of carbohydrate reserves and eventual cytoplasmic acidosis. In nature, hypoxia frequently precedes anoxia in flooded roots. If maize or rice seedlings are transferred to hypoxic conditions (3 kPa O2) before transfer to anoxia (0 kPa O2), their survival rates increase considerably and the ability to continue cell elongation is greatly improved. This hypoxic pretreatment promotes acclimation by increasing glycolytic flux and ATP production. An important feature of this a­cclimation is development of the capacity to export lactate from the cytoplasm to the surrounding medium (Fig.  22.29), a further indication that avoidance of cytoplasmic acidosis is a major factor in survival of low‐oxygen conditions.

Ethanol

Diffusion

Ethanol

Lactate accumulation

Hypoxic pretreatment

Glucose Hypoxia

Anoxia ≥4

Time (h)

Prolonged survival

Ethanol Lactate

Diffusion

Efflux

Ethanol

Lactate

FIGURE 22.29  Effects of hypoxic p­ retreatment and acclimation on survival of anoxia: avoidance of cytoplasmic acidosis by lactate efflux. Flooding typically results in hypoxia, followed by anoxia. Lactate levels and ethanolic fermentation increase in both anoxic and hypoxic cells. Exposure of maize seedlings to hypoxia for several hours before being transferred to anoxia increases the capacity of roots for lactate efflux and ­prolongs survival.

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22.5.3  Shifting from aerobic metabolism to glycolytic fermentation involves changes in gene expression

including phosphorylation of translation factors and ribosomal proteins, may allow for discrimination in translation of cellular mRNAs in oxygen‐deprived cells.

Generally, anoxia causes a rapid and dramatic shift in gene expression patterns, including a significant reduction in total protein synthesis. Protein synthesis is also reduced and altered under hypoxia, but to a lesser extent. Modifications in the pattern of protein synthesis observed in anoxic and hypoxic tissues result from transcriptional and post‐transcriptional regulation of gene expression. Although most gene transcription is generally repressed in response to oxygen deprivation, an important subset of genes is upregulated. These include transcription factors, metabolic enzymes, and uncharacterized proteins. A number of the proteins synthesized in high amounts in hypoxic and anoxic organs are enzymes involved in sucrose and starch degradation, glycolysis, ethanol fermentation, and alanine production. The ADH gene promoters of maize and Arabidopsis ­possess functional cis‐acting elements that are required for expression in hypoxic and anoxic cells. These elements include G‐box type motifs and the anaerobic response e­lement (ARE), which is found in the promoter region of many genes that are transcriptionally induced by low oxygen in monocots and dicots. Transcription factors that interact with Arabidopsis ADH1 have also been identified. Dimethyl sulfate footprinting and DNase I hypersensitivity analyses of ADH promoters have demonstrated both constitutive and dynamically modified DNA–protein interactions that correlate with transcriptional activity. The submergence of rice seedlings causes histone H3 lysine methylation (H3K4) in both 5′ and 3′ regions of ADH1 that is associated with decreased chromatin compaction and increased gene transcription. Together, these observations indicate the higher‐order structure of the chromatin of ADH genes is influenced by oxygen availability. Arabidopsis ADH1 is also transcriptionally induced by application of ABA, dehydration, and cold stress. Changes in gene expression cannot be attributed solely to transcriptional controls. In maize and Arabidopsis, many cytosolic transcripts accumulate at approximately the same amount under normal and oxygen deprivation conditions, but are poorly translated under oxygen deprivation. At least in maize, these mRNAs are constitutively transcribed. The majority of the stress‐induced mRNAs, including ADH1, are translated efficiently in anoxic/hypoxic cells. The translation of maize ADH1 mRNA under low‐oxygen conditions depends on the presence of particular sequences in the 5′ and 3′ untranslated regions. The reduced synthesis of many normal cellular proteins reflects the failure of their mRNAs to effectively maintain initiation of translation during stress. These transcripts are limited in translation and protected from degradation by storage in an mRNA ribonucleoprotein particle, also called stress granules (Fig.  22.30), as a means of conserving cellular energy, as evidenced by their rapid relocation to polyribosomes upon reoxygenation. Distinctions in mRNA binding proteins and changes in translational machinery,

22.5.4  The plant hormone ethylene promotes long‐term acclimative responses in wetland and flood‐tolerant species Flooding or submergence stimulates production and limits outward diffusion of the gaseous hormone ethylene, a key trigger for adaptive responses to submergence and low oxygen levels (