Molecular Responses

Kazuo Shinozaki andKazuko Yamaguchi-Shinozaki

Molecular Responses toCold, Drought, Heatand Salt Stress in Higher Plants

BIOTECHNOLOGY I N T E L L I G E N C E U N I T 1

R.G. LANDESC O M P A N Y

BIOTECHNOLOGYINTELLIGENCEUNIT 1

Kazuo Shinozaki, Ph.D.Laboratory of Plant Molecular Biology

Tsukuba Life Science CenterThe Institute of Physical and Chemical Research (RIKEN)

Tsukuba, Japan

Kazuko Yamaguchi-Shinozaki, Ph.D.Biological Resources Division

Japan International Research Centerfor Agricultural Sciences (JIRCAS)

Tsukuba, Japan

Molecular Responsesto Cold, Drought, Heatand Salt Stress in Higher Plants

AUSTIN, TEXAS

U.S.A.

R.G. LANDESCOMPANY

AUSTIN, TEXAS

U.S.A.

Cold, drought, heat and salt stress in higher plants / [edited by] Kazuo Shinozaki,Kazuko Yamaguchi-Shinozaki.

p. cm. -- (Biotechnology intelligence unit)ISBN 1-57059-563-1 (alk. paper)1. Plants, Effects of stress on—Molecular aspects. 2. Plant molecular genetics.I. Shinozaki, Kazuo. II. Yamaguchi-Shinozaki, Kazuko. III. Series.QK754.C65 1999571.9'52—dc21 99-33224

CIP

Cold, Drought, Heat and Salt Stress in Higher Plants

ISBN: 1-57059-563-1

Library of Congress Cataloging-in-Publication Data

BIOTECHNOLOGY INTELLIGENCE UNIT

R.G. LANDES COMPANYAustin, Texas, U.S.A.

Copyright ©1999 R.G. Landes Company

All rights reserved.No part of this book may be reproduced or transmitted in any form or by any means,electronic or mechanical, including photocopy, recording, or any information storage andretrieval system, without permission in writing from the publisher.Printed in the U.S.A.

Please address all inquiries to the Publishers:R.G. Landes Company, 810 South Church Street, Georgetown, Texas, U.S.A. 78626Phone: 512/ 863 7762; FAX: 512/ 863 0081

While the authors, editors and publisher believe that drug selection and dosage and thespecifications and usage of equipment and devices, as set forth in this book, are in accord withcurrent recommendations and practice at the time of publication, they make no warranty,expressed or implied, with respect to material described in this book. In view of the ongoingresearch, equipment development, changes in governmental regulations and the rapid accumulation ofinformation relating to the biomedical sciences, the reader is urged to carefully review and evaluatethe information provided herein.

CONTENTS

1. Genetic Approaches to Abiotic Stress Responses .................................... 1M. Koornneef and A.J.M. Peeters

The Genetic Approach in Stress Physiology—General Principals ........ 2Genetic Differences in the Response to Low Temperatures .................. 4ABA Related Mutants .............................................................................. 5Conclusions ............................................................................................. 7

2. Molecular Responses to Drought Stress ................................................ 11Kazuo Shinozaki and Kazuko Yamaguchi-Shinozaki

A Variety of Functions of Drought-Inducible Genes .......................... 12Regulation of Gene Expression by Drought ........................................ 14Signal Perception and Signal Transduction in Drought

Stress Response .................................................................................. 18Conclusion and Perspectives ................................................................ 25

3. Molecular Mechanisms of Salinity Tolerance ....................................... 29Hans J. Bohnert, Hua Su and Bo Shen

Osmolytes, Osmoprotectants, Compatible Solutes, OsmoticAdjustment ........................................................................................ 30

Cellular Mechanisms of Salt Tolerance—the Fungal Model .............. 31Molecular Mechanisms of Salt Tolerance in Plants ............................. 37Metabolic Engineering of Glycophytic Plants for Increased

Salt Tolerance .................................................................................... 47Perspectives ............................................................................................ 48

4. Plant Cold Tolerance ............................................................................... 63Michael F. Thomashow and John Browse

Chilling tolerance .................................................................................. 63Freezing Tolerance ................................................................................ 69Conclusions and Perspectives ............................................................... 77

5. Molecular Responses to Heat Stress ....................................................... 83Fritz Schöffl, Ralf Prändl and Andreas Reindl

Heat Shock Proteins and Thermotolerance ......................................... 84Links to Other Abiotic Stresses ............................................................. 87Transcriptional Regulation ................................................................... 89The Regulation of HSF .......................................................................... 90Conclusions and Perspectives ............................................................... 93

6. Cellular Responses to Water Stress ...................................................... 101Michael R. Blatt, Barbara Leyman and Alexander Grabov

The Stomatal Situation ........................................................................ 102Transport Mechanics ........................................................................... 103Transport Coordination in the Face of Stress .................................... 105Interaction of Signaling Elements ....................................................... 114Initial Events in ABA Stimulus Perception ........................................ 115Perspectives and Conclusion .............................................................. 117

Acknowledgments ............................................................................... 118

7. Role of Glycine Betaine and Dimethylsulfoniopropionatein Water-Stress Tolerance ..................................................................... 127Douglas A. Gage and Bala Rathinasabapathi

Stress Protection by Glycine Betaine and DMSP In Vivoand In Vitro ..................................................................................... 128

Biosynthesis of DMSP ......................................................................... 134Conclusion ........................................................................................... 147

8. Osmotic Stress Tolerance in Plants: Role of Proline and SulfurMetabolisms ........................................................................................... 155Desh Pal S. Verma

Osmoregulation in Microorganisms .................................................. 155Osmosensing and Signal Transduction Machinery ........................... 156Osmotic Stress Tolerance in Plants .................................................... 158Accumulation of Other Osmolytes ..................................................... 160Accumulation of Proline in Transgenic Plants Expressing Elevated

Levels of P5CS ................................................................................. 161Proline Accumulation Confers Osmoprotection............................... 163Role of Sulfur metabolism in Osmotic stress Tolerance ................... 164A Possible Role of DPNPase in Salt Tolerance .................................. 166Overexpression of Plant HAL2 Gene Confers Reduction in Free Radical

Production and in Heavy Metal Toxicity ....................................... 166

Kazuo Shinozaki, Ph.D.Laboratory of Plant Molecular Biology

Tsukuba Life Science CenterThe Institute of Physical and Chemical Research (RIKEN)

Tsukuba, JapanChapter 2

Kazuko Yamaguchi-Shinozaki, Ph.D.Biological Resources Division

Japan International Research Centerfor Agricultural Sciences (JIRCAS)

Tsukuba, JapanChapter 2

EDITORS

CONTRIBUTORSMichael R. Blatt, Ph.D.Laboratory of Plant Physiology and

BiophysicsUniversity of London, Wye CollegeWye, England, U.K.Chapter 6

Bo Shen, Ph.D.Departments of Plant SciencesThe University of ArizonaTucson, Arizona, U.S.A.Chapter 3

Hans J. Bohnert, Ph.D.Departments of Biochemistry,

Molecular and Cellular SciencesThe University of ArizonaTucson, Arizona, U.S.A.Chapter 3

John Browse, Ph.D.Institute of Biological ChemistryWashington State UniversityPullman, Washington, U.S.A.Chapter 4

Douglas A. Gage, Ph.D.Department of BiologyMichigan State UniversityEast Lansing, Michigan, U.S.A.Chapter 7

Alexander Grabov, Ph.D.Laboratory of Plant Physiology and


Hua Su, Ph.D.Departments of Plant SciencesThe University of ArizonaTucson, Arizona, U.S.A.Chapter 3

M. Koornneef, Ph.D.Laboratory of GeneticsWageningen Agricultural UniversityWageningen, The NetherlandsChapter 1

Barbara Leyman, Ph.D.Laboratory of Plant Physiology and


A.J.M. Peeters, Ph.D.Laboratory of GeneticsWageningen Agricultural UniversityWageningen, The NetherlandsChapter 1

Ralf Prändl, Ph.D.Lehrstuhl Allgemeine GenetikUniversität TubingenTubingen, GermanyChapter 5

Bala Rathinasabapathi, Ph.D.Hort Sciences DepartmentUniversity of FloridaGainesville, Florida, U.S.A.Chapter 7

Andreas Reindl, Ph.D.Lehrstuhl Allgemeine GenetikUniversität TubingenTubingen, GermanyChapter 5

Fritz Schöffl, Ph.D.Lehrstuhl Allgemeine GenetikUniversität TubingenTubingen, GermanyChapter 5

Michael F. Thomashow, Ph.D.Department of Crop and Soil Sciences,

Department of MicrobiologyMichigan State UniversityEast Lansing, Michigan, U.S.A.Chapter 4

Desh Pal S. Verma, Ph.D.Department of Molecular Genetics and

Plant Biotechnology CenterOhio State UniversityColumbus, Ohio, U.S.A.Chapter 8

PREFACE

T he genetic improvement of tolerance of crops to environmental stresses, suchas drought, high salinity, low temperature and heat, is an important

problem for the future of agriculture. Classical breeding methodologies toselect stress tolerant cultivars have already made some progress. Biotechnologyhas the potential to improve environmental stress tolerance of crops usingtransgenic plant technology. The limiting factor for developing this technologyis the isolation of genes that can improve drought tolerance and the preciseunderstandings of molecular process of stress tolerance and plants’ responsesto environmental stresses. Plants respond to environmental stresses, such asdrought, high salinity, low temperature and heat, through a number ofphysiological and developmental changes. Recently, higher plants respond tothese stresses at gene expression level. A variety of stress-inducible genes havebeen cloned and analyzed concerning to their expression and function in stresstolerance and stress responses. Recently, many mutants have been isolated thatare resistant or hypersensitive to environmental stresses, and cloning of theirgenes is now in progress. Molecular and genetic analyses of the regulation ofgene expression and signal transduction cascades proceed extensively, and willgive us more precise insight on the plants' responses to environmental stressesand their adaptation processes. These stress-related genes are thought tobecome useful resources to produce stress tolerant crops using genemanipulation. In this book, recent progresses on molecular mechanisms ofplant responses and tolerance to drought, salt, cold and heat stresses arereviewed by active researchers in this field. I hope that this book stimulatesyoung students and researchers to become interested in new plant science basedon molecular biology and new plant biotechnology.

Molecular Responses to Cold, Drought, Heat and Salt Stress in Higher Plants, edited by KazuoShinozaki and Kazuko Yamaguchi-Shinozaki. ©1999 R.G. Landes Company.

Genetic Approachesto Abiotic Stress ResponsesM. Koornneef and A.J.M. Peeters

Plants grow in almost any part of the world and under a wide variety of nutrient andclimatic conditions differing in temperature, light quantity and quality and availability

of water. Plants that grow in a specific environment are adapted to these different localconditions, and can also cope with changes in these conditions which might be adverse fortheir growth and development. Adaptation is required because plants cannot escapeunfavorable conditions due to their sessile growth habit. This implies that species differgenetically in their adaptation and resistance to abiotic stresses. Although more restricted,genetic variation for the adaptation to abiotic stresses can also be present within species andhas been used for plant breeding practice. Examples of how plants deal with extremetemporarily adverse conditions are the so-called resurrection plants, which can lose morethan 90% of their water content, but still are able to revive when supplied again with water.Examples of plants that can grow under extreme low temperatures are those that grow inarctic regions or at high altitudes.

Plants can be preadapted to stress conditions but often various protection mechanismsare induced by the stress treatments itself. This implies that plants are able to perceive stresssignals and that after perception signal transduction events take place. As a consequence,these lead to changes in gene expression, as indicated by the many situations whereupregulation of genes is observed after the application of various types of abiotic stress(reviewed by Zhu et al1). Ultimately various cellular mechanisms are set in place, whichallow the plant to cope with the stress imposed. These mechanisms are for instance osmo-adjustment and osmo-protection, changes in pathways affecting ion and water fluxes,production of protection proteins etc.2,3 In case of osmo-adjustment the osmotic potentialof the cell is lowered to favor water uptake and maintenance of turgor. Osmoprotectantsstabilize proteins and membranes when present in high concentrations and include a variety ofcompounds such as amino acids (proline), quaternary ammonium compounds(betaines), polyols (pinitol, mannitol), sugars such as fructans2 and specific proteins such asdehydrins.5 The introduction of genes leading to increased levels of such compounds intransgenic plants has resulted in increased stress tolerance.2,4 The genes used for this wereoften of microbiological origin. Certain gene products might also be involved in the repairof damage caused by the stress.

In addition to the cellular content, membranes also play an important role inadaptation. Especially, the degree of saturation of the membrane lipids is an importantfactor in this.6 When studying the response to stress, one should take into account thatorgans can differ in this respect. As an example seeds, and often pollen also, can surviveextreme desiccation, whereas the vegetative parts and flowers are susceptible to such

CHAPTER 1

Molecular Responses to Cold, Drought, Heat and Salt Stress in Higher Plants2

conditions. This response allows the seeds to survive in a dry state and is also present inthose species that grow under favorable conditions. The acquisition of this desiccationtolerance during seed maturation is very similar to vegetative responses to water deficits.

In addition to these cellular mechanisms, plants can control their water status bycontrolling water uptake and water loss. Water uptake can be regulated by the architectureand physiology of the root system, whereas water loss is regulated not only by controllingmorphological modifications to avoid excessive loss of water such as specific surfacestructures found in succulent plants but also by the strict control of stomatal aperture. Whensuch adaptations result in stress resistance the mechanism has been called “avoidance.”

Despite variation in the nature of adverse conditions, it should be emphasized thatabiotic stresses can have components in common. Insufficient water supply can result froman excessive loss of water, or an insufficient uptake of water. The latter can also result froma high concentration of osmotic material in water, which usually is salts. Chilling and freezingmay also lead to osmotic stress due to reduced water absorption and cellular dehydration.It is likely that for coping with other types of abiotic stresses such as UV light, heat, touch,wounding and hypoxia, plants have different mechanisms available. As anexample Reactive oxygen species (ROS) are involved in the damage due to ozone but alsohave been implicated in the damage that occurs from drought and chilling stress.7 Plantpossess a number of mechanisms and enzyme systems to scavenge ROS. However, theprotection of ROS targets is also a mechanism to deal with such damage and recently Shen etal7 showed that mannitol can play such a role by protecting the enzyme phosphoribulokinaseagainst oxidative inactivation.

Another type of abiotic stress, which has its specific mechanisms and genetics, deals withheavy metals. The latter topic is beyond the scope of this review.

The Genetic Approach in Stress Physiology—General PrincipalsThe genetic approach in stress biology is based on finding genetic differences in stress

responses and to relate this to the structure and function of the genes involved. The usegenetic variation with a “detectable” phenotype is an important tool to identify genesrelated to stress and stress tolerance, because it allows the identification of the respectivegenes by various techniques such as map based cloning or tagging. The feasibility of thesetechniques in model species such as Arabidopsis explains why much of the genetic analysisfocuses on this species.

Genetic variation can be generated by mutations, but also exists in nature or, in case ofcrop plants, among cultivated varieties. Genetic variation within species is often of a differenttype than that found in mutant screens. In contrast to mutants, which can be grown inprotective environments to survive even when they are weak growing plants, natural variantsneed to survive under normal growth conditions and therefore in any case should be wellgrowing plants.

In order to understand the underlying biochemical and molecular bases of geneticvariation present in nature, it is important that the genetic differences in stress tolerancecan be correlated with traits and genes that confer this difference in tolerance. For this it isnot sufficient to see a correlation in the parents, but this correlation between traits shouldbe analyzed in segregating populations. However, a situation of close linkage still does notprove that these traits are due to the same gene (pleiotropism). Pleiotropism cannot bedistinguished easily from close linkage by segregation analysis, unless very large populationsare used.

Since stress tolerance genetically behaves as a quantitative trait with large environmentaleffects on the parameters to be analyzed and often is under polygenic control, detailedgenetic analyses are difficult. However, the advent of molecular markers, and the use of

Genetic Approaches to Abiotic Stress Responses 3

specific mapping populations and developments in statistical methods, have improved theso called QTL (quantitative trait loci) analysis very much and make this also suitable for thegenetic analysis of stress tolerance.8 The more genes with known functions are placed onthe various genetic maps, the more candidate genes for stress tolerance can be identified bythe co-localisation of QTLs and candidate genes.9 The relation between the map location ofthe various dehydrin genes with the map position of a number of genes related to stresstolerance and other physiological traits in cereals is an example of this candidate geneapproach.10 However, confirmation of a causal relationship should preferentially come fromgene transfer experiments with alleles cloned from the various parental genotypes. In caseswhere individual QTLs have an effect that is large enough, it may be possible to identify therespective gene by map-based cloning approaches.

Additional genetic variation can be generated by the introduction of specific geneswhich originate from other plant species or even from completely different organisms. Thelatter has opened completely new ways to modify stress tolerance in crop plants.2,4

Furthermore, genetics and gene transfer technology not only allow the modification ofstress resistance but also are important tools as functional tests of components involved inthe response of plants to abiotic stresses.2

Since most mutations reflect damages within the gene or its controlling elements, oneexpects that mutants lacking a function are most frequent. However, even loss of functionmutants can result in an increase in the functioning of pathways when repressors of suchpathways are mutated. Not all genes can be identified by looking for mutant phenotypes. Areason for this is that for some traits, either the phenotype is relatively subtle, or thephenotype is too general, e.g., reduction of plant size or vigor. To solve the problem ofsubtle phenotypes more sophisticated screens, for instance by using reporter genes, arebeing developed and will be described hereafter. Another reason for not finding specificmutants is that for many genes genetic redundancy exists. This means that mutations insuch genes do not result in an obviously visible phenotype since the redundant counterpartproduces enough product to (partially) substitute the function of the mutated gene. Recently anumber of additional methods which include enhancer or gene trapping and reversegenetics11,12 have become available and allow the analysis of phenotypes associated withthe (loss of) function of specific genes. Reverse genetics uses gene sequences that are identifiedby “molecular” approaches and in large scale sequencing projects. When collections of plantswith insertions of T-DNA or transposable elements are available, one can use these toidentify insertions in the gene of interest. Plants with insertions in such genes areidentified by the ability to amplify DNA fragments in polymerase chain reaction (PCR).One PCR primer is based on the tagging-DNA, whereas the second is based on the sequencefor which an insertion mutant is sought. In case no mutant phenotype is observed when therespective gene is disrupted, due to the redundancy mentioned above, one can expectmutant phenotype, when (insertion) mutations in duplicated genes are combined bycROSsing. In addition to loss of function mutations, gain of function mutants can beisolated by introducing enhancer sequences next to relevant genes resulting in activation ofsuch genes (activation tagging).13 The study of transgenic plants in which cloned genesare over-expressed often has given clues about the function of the respective genes.

The development of appropriate mutant screens is a crucial aspect in any geneticapproach. One can focus on the stress trait itself, which might be considered as theend point of a signal transduction chain. However, one can also pay attention to specificcomponents known to be involved in stress responses. Examples of the latter are the searchand analysis of mutants affected in abscisic acid (ABA) biosynthesis or ABA response andthe analysis of mutants affecting specific stress related genes, either by finding mutations inthose genes or by finding mutations that modify the expression of such genes. The ease of


finding mutants also depends on the effectiveness of the screening system. For this, resistanceto, for instance, salt seems a more attractive screen than the isolation of salt susceptible geno-types. However, screens of the latter type have been successfully applied in case of salt14 andfrost.15

A number of these direct and indirect approaches to select mutants affected in stressresponses will be described, together with the results obtained in analyzing “natural”genetic differences. In what way these analyses have led and are expected to lead to abetter understanding of the responses to abiotic stress will also be discussed.

Genetic Differences in Salt ToleranceSalt overly sensitive (sos) Arabidopsis mutants were isolated by the inability of their roots

to grow on 50 mM NaCl.14 It was shown that these mutants were defective in the high-affinity K+ uptake system. These mutants are hypersensitive to salt stress because relativelyhigh Na+ concentrations inhibit the residual low affinity system of Na+ and K+, whichresults in potassium deficiency, which is the cause of the growth defect. The mutants alsoaffect Na+ uptake and consequently accumulate less.15 In contrast to sos1 mutant, the sos3mutant can be rescued by high external Ca2+.16 Attempts to isolate salt tolerant mutants inArabidopsis resulted in the rss mutants, that express this tolerance only at the seed-germinationlevel.17,18 NaCl tolerance at the germination level was also found to be a characteristic ofABA deficient mutants, which trait in these mutants was considered as reflecting thereduced seed dormancy characteristic of ABA deficient mutants.19 No seed dormancy orABA related phenotype was reported for the rss mutants, which locus maps at a differentposition than the three known ABA deficient (aba) mutants.

Genetic variation for salt tolerance has been described in many crop plants and theirwild relatives. In a number of cases, these have been associated with selective ion uptake.20

Hexaploid wheat (Triticum aestivum) is more salt tolerant than tetraploid durum wheat(T. turgidum). It was shown that in hexaploid wheat a single gene (Kna1) is responsible foraccumulating less Na+ and more K+ in expanding and young leaves. This gene is located onchromosome 4D, which is lacking in tetraploid wheats.21

The examples mentioned indicate that ion uptake and ion transport can affect stresstolerance. However, in addition, genetic differences in salt tolerance have also beenassociated with properties that relate to other osmotic mechanisms. For example, Saneokaet al20 found that near-isogenic maize lines differing in glycine betaines also differed insalt tolerance. Moons et al23 showed that ABA accumulation upon salt stress was muchmore in a salt tolerant rice cultivar as compared with a sensitive variety. The accumulationof a number of ABA induced LEA dehydrin type proteins was also higher in the tolerantvarieties. Whether the latter correlations are causal could not be established, since the varioustraits were not analyzed in segregating populations.

Salt tolerance seemed an attractive system for selection at the cell and tissue culturelevel. However, success of this approach appeared limited to a few successful examples wheresalt tolerant plants could also be obtained.24

Genetic Differences in the Response to Low TemperaturesThe ability of plants to tolerate low temperatures differs much between species.

Distinctions are often made between chilling (temperatures <10˚C) and frost at subzerotemperatures. Exposure to low non-freezing temperatures can strongly increase the freezingtolerance of a species, which process is called acclimation. This interaction between thesetwo temperature regimes complicates the analysis of cold tolerance.


Screens for cold sensitive mutants have been applied in Arabidopsis, where a number ofloci were identified. In a direct screen for lack of growth at 10˚C-13˚C chilling sensitive (chs)mutants were found. The chs1 mutant has been described in some detail and was shown tobe defective in the accumulation of newly synthesized chloroplast localized polypeptides atlow temperatures.25 However, the primary defect of this mutant is not known.

It has also been well established that in plants the level of unsaturated fatty acids in theglycerolipids of membranes changes with changes in growth temperature.6 The alterationof the level of enzymes controlling this trait, in transgenic plants, modifies cold tolerance.6

Furthermore, it was expected that genotypes with reduced levels of acyl-lipid desaturasesmight be more sensitive to low temperatures. This was shown for various Arabidopsismutants.26,27 An exception to this appeared to be the fab1 mutant of Arabidopsis (defective inpalmitoyl-ACP elongase).28 However, since chlorosis was observed at 2˚C in continous light,it appears that damage depends on the light conditions in which the plants were tested. Athigh light intensities the unsaturated species of phosphatidylglycerol accelerate therecovery process from the low temperature inactivated state of the photosystem II complex.6

This indicates the complexity of the various physiological interactions and therebythe difficulty in designing the proper test conditions to identify genetic differences.

Frost sensitive mutants were isolated in direct screens for such mutants and resulted inthe finding of seven different complementation groups, named sfr1-sfr7 (sensitivity tofreezing).15 Mutants at the loci sfr1, sfr2, sfr3 and sfr5 had no obvious pleiotropic effectssuch as slow growth at the seedling stage, which was observed in the sfr4 and sfr7 mutants.The sfr6 mutant had several additional pleiotropic defects.15 Thus far the biochemicaldefect or the nature of the genes is not known for any of these genes.

Relatively large genetic differences have been reported, especially in the grass family,where, as expected, winter cereals are clearly more frost tolerant than summer cereals. Thisled to the identification of some single gene differences for winter hardiness, which map atsimilar genetic positions in wheat, barley and rye.29 Although some recombinants have beenfound, thereby excluding pleiotropism, it is of interest that responsiveness to vernalization,which affects flowering time and thereby winter versus spring habit, and traits such as ABAaccumulation30 and some dehydrin genes map to a similar position.10 Therefore, causalrelationships of some properties can not be excluded but need confirmation.

ABA Related Mutants

The Isolation of ABA Related MutantsThe plant hormone abscisic acid (ABA) seems a crucial component of the response of

plants to stress. ABA levels increase rapidly, especially upon water stress, and result instomatal closure. This is caused by changes in gating properties of ion channels, which leadto changes in the turgor of the two surrounding guard cells. It is assumed that rapid changesin cell turgor (cell shrinkage) lead to this increase in ABA biosynthesis. The effect of coldand osmotic stress on ABA levels may be due to the effect on the water status of the cellscaused by these stresses. This signal induces gene expression of the cleavage enzyme gene ofABA biosynthesis,31 but how this is achieved is not known. However, the role of ABA in theacclimation process, which is associated with changes in gene expression may be slowerthan the effects on stomatal closure, as was shown by the ability of ABA to mimic theacclimation treatment.32,33 Recent progress on the signal transduction pathway of ABA hasbeen reviewed by Leung and Giraudat34 and Shinozaki and Yamaguchi-Shinozaki.35

In addition to effects on water relations and stress tolerance, ABA also plays a crucialrole during seed development and germination. This property allowed the efficientscreening for ABA related mutants at the seed germination level.


The lack of seed dormancy due to ABA deficiency is shown by a high percentage ofgermination of freshly harvested seeds and of seeds in darkness.36 Furthermore, the gibberellin(GA) requirement for Arabidopsis seeds to germinate is abolished or strongly reduced inABA-deficient and ABA-insensitive mutants. This allows seeds to germinate in the presenceof inhibitors of GA biosynthesis such as tetcyclacis and paclobutrazol. These propertieshave been the basis for the selection of ABA-deficient mutants at the ABA1, ABA2 and ABA3loci in Arabidopsis19,36 ABA-insensitive (abi1-abi5) mutants were selected as seeds thatgerminated at ABA concentrations that normally inhibit germination.37,38 These mutantsaffect many ABA responses (abi1 and abi2) or only seed germination (abi3, abi4, and abi5).31,38

The cool mutant of barley39 is an example of a mutant in which ABA insensitivity isspecific for stomatal closure upon drought stress. ABA-insensitive mutants for whichinsensitivity is restricted to the growth response were isolated as mutants for which rootgrowth is not inhibited by ABA. This class of mutants is called gca (growth control via ABA)and comprises at least 8 loci.40 However, the gca1 and gca2 mutants also show defectsin stomatal closure. Mutants which are hypersensitive to ABA are the era1-era3 (enhancedresponse to ABA) mutants, which do not germinate at ABA concentrations that normallypermit germination of the wild type.41 The era1 mutants are also affected in severaladult plant responses.

The Molecular Function of ABA, ABI and ERA GenesSeveral procedures allow the cloning of genes based on mutants. Map-based cloning

has been used to clone the ABI142,43 and ABI3 loci.44 T-DNA tagging was used to cloneERA1.41 Mutants tagged with transposable elements were used to clone the ABA2 gene inN. plumbaginifolia45 and the VP14 gene in maize,31 which both encode steps in ABAbiosynthesis. Through homology with the Nicotiana ABA2 gene, the Arabidopsis ABA1 gene,encoding zeaxanthin epoxydase was isolated.45 VP14 controls the cleavage of carotenoids,which is considered to be the key regulatory step in ABA biosynthesis.31,46 The ABI1 geneencodes a serine/threonine protein phosphatase 2C enzyme42,43,47 and by homology withABI1, the ABI2 gene was isolated.48 The similarity between ABI1 and ABI2 indicates thatthese genes have overlapping functions. The null mutants at either ABI1 and ABI2 may haveno, or a very mild, phenotype and only mutants with a dominant negative effect wouldblock the phosphatases encoded by both genes. The dominance of both mutations and thevery similar amino acid substitution present in both mutants are in accordance with thishypothesis.48

The amino acid sequence of ABI344 revealed that this gene is homologous to the maizeViviparous-1 (Vp1) gene,49 which is a transcription factor that is specific for seed development.The amino acid sequence of ERA1 showed that this gene encodes the β subunit of a farnesyltransferase. The era1 mutant lacks peptide farnesylation activity and it is suggested that theERA1 gene acts as negative regulator by modifying ABA signal transduction proteins formembrane localization.41

The Isolation of Mutants Affected in the Response to Stressand/or ABA Induced Genes and Processes

Novel ways to identify genes interacting with stress induced genes apply the expression ofreporter genes which are under the control of promoters of these genes.

Mutants with an altered expression of these reporters are expected to also be affectedalso in the expression of the endogenous promoter. When this has been confirmed thechanges in physiological properties of the mutant should indicate the relevance of theexpression of this and related genes that might be down or upregulated. Reporter genesthat are most efficient are those where the reporter gene assay is non-destructive, as, e.g.,


in the case of luciferase. That this approach can yield numerous mutants of which many haveno dramatic phenotypic effects was shown by Ishitani et al,50 who used the rd29A promoterdriving luciferase. The promoter of this gene contains both an ABA independent droughtresponsive (DRE) and an ABA dependent (ABRE) element. Screens based on this approachwere very productive and resulted both in mutants with constitutive expression, and inmutants with low and high expression of osmotically responsive genes named respectivelycos, los and hos mutants.50 Within the los and hos class, subclasses could be distinguishedaccording to defects in their responses to one or a combination of stress and ABA signals.

An approach that also makes use of reporter genes is the use of enhancer or gene trapprocedures, where insertions of reporter genes with a minimal or without a promoter mightresult in expression specifically after stress induction.12 This approach might also allow theidentification of genes that are redundant and therefore cannot be found in mutant screens.

Furini et al51 used activation tagging to identify a gene in ABA signaling. In this systemthe T-DNA contains enhancers of gene expression and was used to select inserts that wouldconfer dehydration tolerance to callus of Craterostigma plantagineum, which normallyrequires ABA to obtain this property. The cloning of the sequences involved revealedunusual features and resembled transposon-like sequences.51

Genetic Variation for Morphological Traits Related to Stress Tolerance Morphological adaptations can also be the basis of genetic variation in both water loss

and water uptake. With respect to the latter, root morphology and rooting patterns areimportant. A genetic analysis of these root characters in a segregating population derivedfrom a cross between a drought resistant upland rice cultivar and a drought sensitive lowlandvariety is described by Champoux et al.52 The same material allowed also the detection ofQTLs controlling drought tolerance in a shoot specific way through osmotic adjustment.53

Following a similar QTL approach, Price et al54 investigated leaf rolling, stomatalbehavior and heading date as examples of morphological and physiological traits related tostress tolerance in rice. The correlation found in rice between smaller leaves and ABAaccumulation observed in similar crosses was not confirmed to be pleiotropic in a refinedQTL analysis, but due instead to linkage.55

ConclusionsGenetic variants tell which intrinsic properties, such as chemical composition of cells,

are important for stress tolerance. Both mutants and transgenic plants have confirmed anumber hypotheses based on earlier observations and physiological experiments. Mutantsaffected in specific regulatory factors such as ABA showed that this compound mediates theexpression of many but not all stress induced genes. This indicated that different pathwaysare involved.35 The complexity of this stress-induced signal transduction pathway is alsosuggested by the large number of mutant classes identified by Ishitani et al.50 Such mutantswill be useful to identify the various steps in this pathway and will complement themolecular approaches.1,3,4,35 A further challenge of genetics is to identify, up to themolecular level, the genes controlling natural differences in stress tolerance. Although theeffects of individual genes in some cases may be limited, it can be expected that thesedifferences are useful in application of cloned genes because they operate with limitedpleiotropic effects. This approach to the identification of the genes for which variation ispresent in nature will complement the mutant approaches and molecular approaches. Itcan be expected that in a number of cases the same genes will be identified. However, inother situation they are likely to be different. Together, these genes will tell us how plantscope with and respond to abiotic stresses.


AcknowledgmentsOur research program is supported by the Human Frontier Science Program

(RG-303/95) and the BIOT4 program of the European Union (BIO4-CT96-0062).

References1. Zhu JK, Hasegawa PM, Bressan RA. Molecular aspects of osmotic stress in plants. Crit Rev

Plant Sci 1997; 16:253-277.2. Bohnert HJ, Nelson DE, Jensen RG. Adaptations to environmental stresses. Plant Cell 1995;

7:1099-1111.3. Bray EA. Plant responses to water deficit. Trends Plant Sci 1997; 2:48-54.4. Ingram J, Bartels D. The molecular basis of dehydration tolerance in plants. Annu Rev

Plant Physiol Plant Mol Biol 1996; 47:377-403.5. Close TJ. Dehydrins: A commonalty in the response of plants to dehydration and

low temperature. Physiol Plant 1997; 100:291-296.6. Nishida I, Murata N. Chilling sensitivity in plants and cyanobacteria: The crucial contri-

bution of membrane lipids. Annu Rev Plant Physiol Plant Mol Biol 1996; 47:541-568.7. Shen B, Jensen RG, Bohnert HJ. Mannitol protects against oxidation by hydroxyl radicals.

Plant Physiol 1997; 115:527-532.8. Koornneef M, Alonso-Blanco C, Peeters AJM. Genetic approaches in plant physiology. New

Phytol 1997; 137:1-8.9. Prioul JL, Quarrie S, Causse M et al. Dissecting complex physiological functions through

the use of molecular quantitative genetics. J Exp Bot 1997; 48:1151-1163.10. Campbell SA, Close TJ. Dehydrins: Genes, proteins, and associations with phenotypic traits.

New Phytol 1997; 137:61-74.11. Azpiroz-Leehan R, Feldmann KA. T-DNA insertion mutagenesis in Arabidopsis: Going back

and forth. Trends Genet 1997; 13:152-156.12. Sundaresan V. Horizontal spread of transposon mutagenesis: New uses for old elements.

Trends Plant Sci 1996; 1:184-190.13. Walden R, Fritze K, Hayashi H et al. Activation tagging: A means of isolating genes

implicated as playing a role in plant growth and development. Plant Mol Biol 1994;26:1521-1528.

14. Wu SJ, Ding L, Zhu JK. SOS1, a genetic locus essential for salt tolerance and potassiumacquisition. Plant Cell 1996; 8:617-627.

15. Warren G, McKown R, Marin A et al. Isolation of mutations affecting the development offreezing tolerance in Arabidopsis thaliana (L.) Heynh. Plant Physiol 1996; 111:1011-1019.

16. Liu J, Zhu JK. An Arabidopsis mutant that requires increased calcium for potassiumnutrition and salt tolerance. Proc Natl Acad Sci USA 1997; 94:14960-14964.

17. Saleki R, Young PG, Lefebvre DD. Mutants of Arabidopsis thaliana capable of germinationunder saline conditions. Plant Physiol 1993; 101:839-845.

18. Werner JE, Finkelstein RR. Arabidopsis mutants with reduced response to NaCl andosmotic stress. Physiol Plant 1995; 93:659-666.

19. Leon-Kloosterziel KM, Gil MA, Ruijs GJ et al. Isolation and characterization of abscisicacid-deficient Arabidopsis mutants at two new loci. Plant J 1996; 10:655-661.

20. Foolad MR. Genetic basis of physiological traits related to salt tolerance in tomato,Lycopersicon esculentum Mill. Plant Breed 1997; 116:53-58.

21. Dubcovsky J, Santa MG, Epstein E et al. Mapping of the K+/Na+ discrimination locus Kna1in wheat. Theor Appl Genet 1996; 92:448-454.

22. Saneoka H, Nagasaka C, Hahn DT et al. Salt tolerance of glycinebetaine-deficientand-containing maize lines. Plant Physiol 1995; 107:631-638.

23. Moons A, Bauw G, Prinsen E et al. Molecular and physiological response to abscisic acidand salts in roots of salt-sensitive and salt-tolerant indica rice varieties. Plant Physiol 1995;107:177-186.

24. Winicov I. Characterization of salt tolerant alfalfa (Medicago sativa L.) plants regeneratedfrom salt tolerant cell lines. Plant Cell Rep 1991; 10:561-564.


25. Schneider JC, Nielsen E, Somerville C. A chilling-sensitive mutant of Arabidopsis isdeficient in chloroplast protein accumulation at low temperature. Plant Cell Environment1995; 18:23-32.

26. Hugly S, Somerville C. A role for membrane lipid polyunsaturation in chloroplast biogenesisat low temperature. Plant Physiol 1992; 99:197-202.

27. Miquel M, James DJ, Dooner H et al. Arabidopsis requires polyunsaturated lipids for lowtemperature survival. Proc Natl Acad Sci USA 1993; 90:6208-6212.

28. Wu J, Browse J. Elevated levels of high-melting-point phosphatidylglycerols do not inducechilling sensitivity in an Arabidopsis mutant. Plant Cell 1995; 7:17-27.

29. Galiba G, Quarrie SA, Sutka J et al. RFLP mapping of the vernalization (Vrn1) and frostresistance (Fr1) genes on chromosome 5A of wheat. Theor Appl Genet 1995; 90:1174-1179.

30. Quarrie SA, Gulli M, Calestani C et al. Location of a gene regulating drought-inducedabscisic acid production on the long arm of chromosome 5A of wheat. Theor Appl Genet1994; 89:794-800.

31. Tan BC, Schwartz SH, Zeevaart JAD et al. Genetic control of abscisic acid biosynthesis inmaize. Proc Natl Acad Sci USA 1997; 94:12235-12240.

32. Heino P, Sandman G, Lang V et al. Abscisic acid deficiency prevents development of freezingtolerance in Arabidopsis thaliana (L.) Heynh. Theor Appl Genet 1990; 79:801-806.

33. Gilmour SJ, Thomashow MF. Cold acclimation and cold-regulated gene expression in ABAmutants of Arabidopsis thaliana. Plant Mol Biol 1991; 17:1233-1240.

34. Leung J, Giraudat J. Abscisic acid signal transduction. Annu Rev Plant Physiol Plant MolBiol 1998; 49:199-222.

35. Shinozaki K, Yamaguchi-Shinozaki K. Gene expression and signal transduction in water-stress response. Plant Physiol 1997; 115:327-334.

36. Koornneef M, Jorna ML, Brinkhorst-van der Swan DLC et al. The isolation of abscisicacid (ABA) deficient mutants by selection of induced revertants in non-germinatinggibberillin sensitive lines of Arabidopsis thaliana (L.) Heynh. Theor Appl Genet 1982; 61:385-393.

37. Koornneef M, Reuling G, Karssen CM. The isolation and characterization of abscisic acid-insensitive mutants of Arabidopsis thaliana. Physiol Plant 1984; 61:377-383.

38. Finkelstein RR. Mutations at two new Arabidopsis ABA response loci are similar to theabi3 mutations. Plant J 1994; 5:765-771.

39. Raskin I, Ladyman JAR. Isolation and characterization of a barley mutant with abscisic-acid-insensitive stomata. Planta 1988; 173:73-78.

40. Benning G, Ehrler T, Meyer K, Leube M, Rodriguez P, Grill E. Genetic analysis ofABA-mediated control of plant growth. In: Abscisic acid signal transduction in plants.Madrid: Juan March Foundation, 1996:34.

41. Cutler S, Ghassemian M, Bonetta D et al. A protein farnesyl transferase involved in ABAsignal transduction in Arabidopsis. Science 1996; 273:1239-1241.

42. Leung J, Bouvier-Durand M, Morris PC et al. Arabidopsis ABA response gene ABI1:Features of a calcium-modulated protein phosphatase. Science 1994; 264:1448-1452.

43. Meyer K, Leube MP, Grill E. A protein phosphatase 2C involved in ABA signal transductionin Arabidopsis thaliana. Science 1994; 264:1452-1455.

44. Giraudat J, Hauge BM, Valon C et al. Isolation of the Arabidopsis ABI3 gene by po-sitional cloning. Plant Cell 1992; 4:1251-1261.

45. Marin E, Nussaume L, Quesada A et al. Molecular identification of zeaxanthin epoxidaseof Nicotiana plumbaginifolia, a gene involved in abscisic acid biosynthesis and correspondingto the ABA locus of Arabidopsis thaliana. EMBO J 1996; 15:2331-2342.

46. Schwartz SH, Tan BC, Gage DA et al. Specific oxidative cleavage of carotenoids by vp14 ofmaize. Science 1997; 276:1872-1874.

47. Bertauche N, Leung J, Giraudat J. Protein phosphatase activity of abscisic acid insensitive 1(ABI1) protein from Arabidopsis thaliana. Eur J Biochem 1996; 241:193-200.

48. Leung J, Merlot S, Giraudat J. The Arabidopsis ABI2 gene is a homologue of ABI1 andimplicates redundant protein serine/threonine phosphatases 2C in abscisic acid signaltransduction. Plant Cell 1997.


49. McCarty DR, Hattori T, Carson CB et al. The Viviparous-1 developmental gene of maizeencodes a novel transcriptional activator. Cell 1991; 66:895-906.

50. Ishitani.M., Xiong L, Stevenson B et al. Genetic analysis of osmotic and cold stress signaltransduction in Arabidopsis: Interactions and convergence of abscisic acid-dependent andabscisic acid-independent pathways. Plant Cell 1997; 9:1-16.

51. Furini A, Koncz C, Salamini F et al. High level transcription of a member of a repeatedgene family confers dehydration tolerance to callus tissue of Craterostigma plantagineum.EMBO J 1997; 16:3599-3608.

52. Champoux MC, Wang G, Sarkarung S et al. Locating genes associated with root morphologyand drought avoidance in rice via linkage to molecular markers. Theor Appl Genet 1995;90:969-981.

53. Lilley JM, Ludlow MM, McCouch SR et al. Locating QTL for osmotic adjustment anddehydration tolerance in rice. J Exp Bot 1996; 47:1427-1436.

54. Price AH, Young EM, Tomos AD. Quantitative trait loci associated with stomatalconductance, leaf rolling and heading date mapped in upland rice (Oryza sativa). NewPhytol 1997; 137:83-91.

55. Quarrie SA, Laurie DA, Zhu JH et al. QTL analysis to study the association between leafsize and abscisic acid accumulation in droughted rice leaves and comparisons across cereals.Plant Mol Biol 1997; 35:155-165.

CHAPTER 2


Molecular Responses to DroughtStressKazuo Shinozaki and Kazuko Yamaguchi-Shinozaki

Plant growth is greatly affected by environmental abiotic stresses, such as drought, highsalinity and low temperature. Plants respond and adapt to these stresses in order to sur-

vive against abiotic stress. Among these abiotic stresses, drought or water deficit is the mostsevere limiting factor of plant growth and crop production. Drought stress induces variousbiochemical and physiological responses in plants. Recently, a number of genes have beendescribed that respond to drought at the transcriptional level.1-4 Their gene products arethought to function in stress tolerance and response (Fig. 2.1). Recently, stress-induciblegenes were used to improve stress tolerance of plants by gene transfer. It is important toanalyze functions of stress-inducible genes not only for further understanding of molecularmechanisms of stress tolerance and response of higher plants but also for improvement ofstress tolerance of crops by gene manipulation.

The plant hormone abscisic acid (ABA) is produced under water deficit conditionsand plays important roles in response and tolerance to dehydration. Most of the genes thathave been studied to date are also induced by ABA.5 It appears that dehydration triggersthe production of ABA, which, in turn, induces various genes. Several reports havedescribed genes that are induced by dehydration but are not responsive to exogenous ABAtreatments. These findings suggest the existence of ABA-independent as well as ABA-dependent signal-transduction cascades between the initial signal of drought stress andthe expression of specific genes.1-4 To understand the molecular mechanisms of geneexpression in response to drought stress, cis- and trans-acting elements that function inABA-independent and ABA-responsive gene expression by drought stress have beenprecisely analyzed. A variety of transcription factors are involved in stress responsive geneexpression, which suggests the involvement of complex regulatory systems in molecularresponses to drought stress.

Expression and functions of stress-inducible genes have been studied at molecularlevel as described in this chapter. Complex mechanisms seem to be involved in geneexpression and signal transduction in response to drought stress. However, genetic analyses ofdrought-resistant or drought-sensitive mutants have not been extensively performed. There-fore, details of molecular mechanisms of regulating plant genes to drought stress remainto be solved concerning signal transduction cascades. These include the sensing mechanismsof osmotic stress, modulation of the stress signals to cellular signals, transduction of thecellular signals to the nucleus, second messengers involved in stress signal transduction,roles of ABA in the signaling process, transcriptional control of stress-inducible genes, andthe function and cooperation of stress-inducible genes allowing drought stress tolerance(Fig. 2.1). In this article we describe recent progress mainly on gene expression and


signal transduction in drought stress response. Promoter analysis of several droughtinducible genes suggests that there are at least four signaling cascades in drought stressresponses. Several approaches to improve stress tolerance of plants by gene transfer ofstress-inducible genes are also described.

A Variety of Functions of Drought-Inducible Genes

Two Classes of Drought-Inducible GenesVarious genes respond to drought stress in various species, and functions of their

gene products have been predicted from sequence homology with known proteins. Manydrought-inducible genes are also induced by salt stress (see chapter 3) and low temperature(see chapter 4), which suggests the existence of similar mechanisms of stress responses.Genes induced during drought-stress conditions are thought to function not only inprotecting cells from water deficit by the production of important metabolic proteins butalso in the regulation of genes for signal transduction in the drought stress response.1,2,4

Thus, these gene products are classified into two groups (Fig. 2.2). The first group includesproteins that probably function in stress tolerance, such as chaperones, LEA (late embryo-genesis abundant) proteins, osmotin, antifreeze proteins, mRNA binding proteins, keyenzymes for osmolyte biosynthesis, water channel proteins, sugar and proline transporters,detoxification enzymes and various proteases. LEA proteins, chaperones and mRNA bindingproteins have been analyzed biochemically and shown to be involved in protecting macro-molecules like enzymes, lipids and mRNAs from dehydration. Proline, glycine betaine andsugars function as osmolytes and in protecting cells from dehydration (see chapter 8). Keyenzymes of several osmolytes have been cloned and analyzed biochemically. Water channel

Fig. 2.1. Schematic representation of molecular responses to drought stress in plant cells.Molecular and cellular responses to drought stress include perception of dehydration signal,signal transduction to cytoplasm and nucleus, gene expression, and responses and tolerance todrought stress.

13Molecular Responses to Drought Stress

proteins, sugar transporters and proline transporters are thought to function in transportof water, sugars and proline through plasma membranes and tonoplast to adjust osmoticpressure under stress conditions. Detoxification enzymes such as glutathione S-transferase,superoxide dismutase, and soluble epoxide hydrolase are involved in protection of cellsfrom active oxygens. Proteases including thiol proteases, Clp protease, and ubiquitin arethought to be required for protein turnover and recycle of amino acids.

The second group contains protein factors involved in further regulation of signaltransduction and gene expression that probably function in stress response: proteinkinases, transcription factors and enzymes in phospholipid metabolism.1,2,4 Genes for avariety of transcription factors that contain typical DNA binding motifs, such as bZIP,MYB, MYC, EREBP/AP2 and zinc fingers, have been demonstrated to be stress inducible.4

These transcription factors function in further regulation of various functional genes understress conditions. Various protein kinases, such as MAP kinases, calcium dependent proteinkinases (CDPK), SNF1 related protein kinase and ribosomal S6 kinase, were demonstratedto be induced or upregulated by dehydration.4,7 Stress-inducible genes for protein phos-phatases are reported.8 These protein kinases and phosphatases may be involved inmodification of functional proteins and regulatory proteins involved in stress signal trans-duction pathways. Phospholipid, such as inositol-1,4,5-triphosphate, diacylglycerol andphospahtidic acid are believed to be involved in stress signaling processes in plants.Enzymes involved in phospholipids metabolism whose genes are stress-inducible may playimportant roles in stress signaling as well.

Fig. 2.2. Drought stress-inducible genes and their possible functions in stress tolerance andresponse. Gene products are classifed into two groups. The first group includes proteins thatprobably function in stress tolerance (function proteins; open boxes), and the second groupcontains protein factors involved in further regulation of signal transduction and gene expressionthat probably function in stress response (regulatory proteins; shadowed boxes).


Existence of a variety of drought-inducible genes suggests complex responses of plantsto drought stress. Their gene products are involved in drought stress tolerance and stressresponses.

Improvement of Stress Tolerance Using Gene TransferRecently, several different approaches were attempted to improve stress tolerance of

plants by gene transfer of stress-inducible genes.9 Stress-inducible genes for functionalproteins such as key enzymes for osmolyte biosynthesis, LEA proteins and detoxificationenzymes were overexpressed in transgenic plants to produce stress tolerant phenotype ofthe plants, which indicates that their gene products really function in stress tolerance. Genesused for transformation were those encoding enzymes required for biosynthesis of variousosmoprotectants, such as Escherichia coli mannitol 1-phosphate dehydrogenase formannitol,10 mothbean ∆1-pyrroline-5-carboxylate synthetase for proline,11 Arthrobacterglobiformis choline dehydrogenase for glycine betaine,12 barley LEA protein13 and Nicotianaplumbaginifolia detoxification enzyme.14 In all these experiments, a single gene for a protectiveprotein or an enzyme was overexpressed under the control of the CaMV 35S constitutivepromoter in transgenic plants, although a number of genes have been shown to function inenvironmental stress tolerance and response. In the next step, I think it important tomanipulate several genes to achieve strong stress tolerance for the application of thistechnology to the development of stress tolerant transgenic crops.

Overproduction of genes for stress-induced transcription factors in transgenic plantsactivated the expression of many target genes involved in stress tolerance under unstressednormal conditions and significantly improved stress tolerance to drought and freezing.15,16

These results suggest that regulatory genes involved in stress response can be also used forthe improvement of stress tolerance by gene transfer.

Regulation of Gene Expression by Drought

Complex Regulatory Systems for Gene ExpressionThe expression patterns of genes induced by drought were analyzed by RNA gel-blot

analysis. Results indicated broad variations in the timing of induction of these genes underdrought conditions. All the drought-inducible genes are induced by high salinity stress.Most of the drought-inducible genes also respond to cold stress but some of them do not,and vice versa. Many genes respond to ABA whereas some others do not.1,2,4 ABA-deficientmutants were used to analyze drought-inducible genes that respond to ABA. Several geneswere induced by exogenous ABA treatment, but were also induced by cold or drought inABA-deficient (aba) or ABA-insensitive (abi) Arabidopsis mutants. These observationsindicate that these genes do not require an accumulation of endogenous ABA under coldor drought conditions, but do respond to ABA. There are ABA-independent as well asABA-dependent regulatory systems of gene expression under drought stress. Analysis ofthe expression of ABA-inducible genes showed that several genes require protein biosynthesisfor their induction by ABA, suggesting that at least two independent pathways existbetween the production of endogenous ABA and gene expression under stress conditions.

As shown in Figure 2.3, it is now hypothesized that at least four independent signaltransduction pathways function in the activation of stress-inducible genes under dehydrationconditions: Two are ABA-dependent (Pathways I and II) and two are ABA-independent(Pathways III and IV).4 One of the ABA-dependent pathways requires protein biosynthesis(Pathway I). Cis- and trans-acting factors involved in ABA-induced gene expression havebeen extensively analyzed in one of the ABA-dependent pathway that does not require denovo protein biosynthesis (Pathway II). One of the ABA-independent pathways overlaps


with that of the cold response (Pathway IV). There are several drought-inducible genesthat do not respond to either cold or ABA treatment, which suggests that there is a fourthpathway in the dehydration stress response (Pathway III). Recently, based on genetic analysis ofArabidopsis mutants with the rd29A promoter—luciferase transgene, the existence ofdrought-, salt- and cold- specific signaling pathways in stress-response was suggested, butcrosstalks between these signaling pathways were also observed (see chapter 1).

Major ABA-Independent Regulatory System of Gene Expression duringDrought and Cold Stress (Pathway IV): Important Roles of DRE/CRT Cis-ActingElement and its DNA Binding Proteins

A number of genes are induced by drought, salt, and cold in aba (ABA-deficient) orabi (ABA-insensitive) Arabidopsis mutants. This suggests that these genes do not requireABA for their expression under cold or drought condition.2,4,17 Among these genes, theexpression of a drought-inducible gene for rd29A/lti78/cor78 was extensively analyzed.18

At least two separate regulatory systems function in gene expression during drought andcold stress; one is ABA-independent (Fig. 2.3, Pathway IV) and the other is ABA-dependent(Fig. 2.3, Pathway II). A 9bp conserved sequence, TACCGACAT, named the dehydrationresponsive element (DRE), is essential for the regulation of the induction of rd29A under

Fig. 2.3. Signal transduction pathways between the perception of drought stress signal and geneexpression. At least four signal transduction pathways exist (I-IV): Two are ABA-dependent (Iand II) and two are ABA-dependent pathways (III and IV). Protein biosynthesis is required inone of the ABA-dependent pathways (I). In another ABA-dependent pathway, ABRE functionsas an ABA-responsive element and does not require protein biosynthesis (II). In one of the ABA-independent pathways, DRE is involved in the regulation of genes not only by drought and saltbut also by cold stress (IV). Another ABA-independent pathway is controlled by drought andsalt, but not by cold (III).


drought, low-temperature, and high-salt stress conditions, but does not function as anABA-responsive element (Fig. 2.4). The rd29A promoter contains ABRE, which functionsin ABA-responsive expression. DRE-related motifs have been reported in the promoterregions of many cold- and drought-inducible genes.3,4,17 These results suggest that DRE-related motifs including C-repeat (CRT) and low temperature responsive element (LTRE),which contain a CCGAC core motif, are involved in drought- and cold-responsive butABA-independent gene expression (see chapter 5).

Protein factor(s) that specifically interact with the 9bp DRE sequence were detectedin nuclear extract prepared from either dehydrated or untreated Arabidopsis plants.18

Recently, five independent cDNAs for DRE/CRT-binding proteins have been cloned usingthe yeast one hybrid screening method.15,19 All the DRE/CRT binding proteins (DREBsand CBFs) contain a conserved DNA binding motif that has also been reported in EREBPand AP2 proteins (EREBP/AP2 motif) that are involved in ethylene-responsive geneexpression and floral morphogenesis, respectively. These five cDNA clones that encodeDRE/CRT binding proteins are classified into two groups, CBF1/DREB1 and DREB2.Expression of the DREB1A gene and its two homologs (DREB1B = CBF1, DREB1C) wasinduced by low-temperature stress, whereas expression of the DREB2A gene and its singlehomolog (DREB2B) was induced by dehydration.15,64 Overexpression of the DREB1A cDNAin transgenic Arabidopsis plants not only induced strong expression of the target genesunder unstressed conditions but also caused dwarfed phenotypes in the transgenic plants.These DREB1A transgenic plants also revealed freezing and dehydration tolerance, whichwas also shown in the CBF1 transgenics.16 In contrast, overexpression of the DREB2AcDNA induced weak expression of the target genes under unstressed conditions and causedslight growth retardation of the transgenic plants. These results indicate that two independentfamilies of DREB proteins, DREB1 and DREB2, function as transacting factors in twoseparate signal transduction pathways under low-temperature and -dehydration conditions,respectively (Fig. 2.4).15

Fig. 2.4. A model of the induction of the rd29A gene and cis- and trans-acting elements involvedin stress-responsive gene expression.15 Two cis-acting elements, DRE/CRT and ABRE, areinvolved in the ABA-independent and ABA-responsive induction of rdnaA, respectively. Twodifferent DRE-binding proteins, DRBE1 and DREB2, separate two different signal transductionpathways in response to cold and drought stresses, respectively. ABRE-binding proteins encodebZIP-transcription factors.


Overproduction of the DREB1A and CBF1/DREB1B cDNAs driven by the 35S CaMVpromoter in transgenic plants significantly improved stress tolerance to drought andfreezing.15,16 However, the 35S-DREB1A transgenic plants revealed severe growth retardationunder normal growth conditions. The DREB1A cDNA driven by the stress-inducible rd29Apromoter was expressed at low level under unstressed control conditions and stronglyinduced by dehydration, salt and cold stresses. The rd29A promoter minimized negativeeffects on growth of plants, whereas the 35S-CaMV promoter caused severe growthretardation under normal growth conditions.15, 20 Moreover, this stress-inducible promoterenhanced tolerance to drought, salt and freezing at higher levels than that of the 35S-CaMVpromoter.

Drought-Specific ABA-Independent Regulatory System (Pathway III)There are several drought-inducible genes that do not respond to either cold or ABA

treatment, which suggests the existence of another ABA-independent pathway in thedehydration stress response (Fig. 2.3, Pathway III). These genes include rd19 and rd21 thatencode different thiol proteases, and erd1 that encodes a Clp protease regulatory subunit.21,22

The ERD1 protein is targeted to chloroplasts whereas the RD19 and RD21 proteins seem tofunction in cytoplasm. The catalytic subunit of the Clp protease (Clp P) is encoded on thechloroplast genome. The erd1 gene is not only induced by dehydration but also upregulatedduring natural senescence and dark-induced senescence.23 The erd1 and rd21 genes werealso identified as senescence-associated genes.24 Promoter analysis of the erd1 gene intransgenic plant indicates that erd1 promoter contains cis-acting element(s) involved in notonly ABA-independent stress responsive gene expression but also senescence-activated geneexpression.23 Further promoter analysis of these genes will give us more information onPathway III.

Major ABA-Dependent Regulatory System (Pathway II): Important Roles ofABRE Cis-Acting Element and its bZIP DNA Binding Proteins

Most drought-inducible genes are upregulated by exogenous ABA treatment. The levelsof endogenous ABA increase significantly in many plants under drought and high salinityconditions.1,2,4 In one of the ABA-dependent pathways (Fig. 2.3, Pathway II), drought-stressinducible genes do not require protein biosynthesis for their expression.4,5 Thesedehydration-inducible genes contain potential ABA-responsive elements (ABREs;PyACGTGGC) in their promoter regions. ABRE functions as a cis-acting DNA elementinvolved in ABA-regulated gene expression.5 ABRE was first identified in wheat Em and ricerab genes, and its DNA-binding protein EmBP1 was shown to encode a bZIP protein. TheG-box resembles the ABRE motif and functions in the regulation of plant genes in a varietyof environmental conditions, such as red light, UV light, anaerobiosis, and wounding. cDNAsfor ABRE and G-box binding proteins have been isolated and shown to have a basic regionadjacent to a leucine zipper motif (bZIP) and constitute a large gene family. Nucleotidesaround the ACGT core motif have been shown to be involved in determining the bindingspecificity of bZIP proteins. Furthermore, a coupling element (CE) is required to specifythe function of the ABRE, constituting an ABA-responsive complex in the regulation of theHVA22 gene.25 However, it has not been resolved how ABA activates bZIP proteins to bindsto ABRE and initiate transcription of ABA-inducible genes. Further studies are necessaryfor the precise understanding of the molecular mechanisms of ABA-responsive geneexpression that requires ABRE as a cis-acting element.

Several bZIP transcription factors from rice, maize and Arabidopsis plants respond tocold, dehydration, and to exogenous ABA treatment.4 These bZIP proteins bind to G-box-like sequences. These results suggest that ABA-inducible bZIP proteins are also involved in


one of the ABA-dependent pathways (Fig. 2.3., Pathway I) or in the enhancement of theABA-dependent gene expression (Fig. 2.3., Pathway II).

There are several cis-acting elements other than ABRE that function in ABA-responsivegene expression, not only under drought conditions but also in seed desiccation. The Sphbox and GTGTC motifs regulate ABA- and VP1-dependent expression of the maize C1gene, whose product is a MYB-related transcription factor and functions as a controllingelement in anthocyanin biosynthesis during seed.26 VP1 encodes a transcriptional activatorand is thought to cooperate with bZIP proteins. Arabidopsis ABI3 has sequence andfunctional similarity with maize VP1. Recently VP1 was demonstrated to have a DNA-binding activity to Sph box. EmBP1 and VP1 were shown to interact with 14-3-3 proteinsand form a transcription complex.27 This complex also interacted with ABRE on the Empromoter. A similar system is thought to function in ABA-responsive gene expression indrought stress response as well as in seed maturation.

Roles of MYC and MYB Homologs in ABA-Dependent Gene Expressionthat Requires Protein Biosynthesis (Pathway I)

Biosynthesis of novel protein factors is necessary for the expression of ABA-induciblegenes in one of the two ABA-dependent pathways (Fig. 2.3, Pathway I). The induction of anArabidopsis drought-inducible gene, rd22, is mediated by ABA, and requires proteinbiosynthesis for its ABA-dependent expression.28 A 67bp region of the rd22 promoter isessential for this ABA-responsive expression, and contains several conserved motifs of DNA-binding proteins, two MYC and one MYB recognition sequences, but this region has noABREs. First MYC and MYB recognition sequences are essential for the ABA- and drought-responsive expression of the rd22 gene.29 A cDNA for a transcription factor MYC homo-logue, named rd22BP1, was cloned by the DNA-ligand binding method, using the 67bpDNA as a probe. The rd22BP1 gene is induced by drought and salt stress. These resultssuggest that a drought- and salt-inducible MYC homologue function in the ABA-inducibleexpression of rd22 (Fig. 2.5). The ATMBY2 gene that encodes a MYB-related protein isinduced by dehydration and ABA treatment.30 Recombinant ATMYB2 protein binds to theMYB recognition sequence in the 67bp region of the rd22 promoter. Moreover, these MYCand MYB proteins transactivate the rd22 promoter GUS fusion gene in transient expressionsystem using leaf protoplasts.29 Therefore, the ATMYB2 protein might also cooperativelyfunction with the rd22BP1 protein as a transcription factor that controls the ABA-dependentexpression of the rd22 gene (Fig. 2.5).

Many stress- and ABA-inducible genes encoding various transcription factors havenow been reported. These contain conserved DNA binding motifs, such as MYB, MYC,bZIP and zinc finger. These transcription factors are thought to function in the regulationof ABA inducible genes, which respond to drought stress rather slowly after the production ofABA-inducible transcription factors (Fig. 2.3., Pathway I).

Signal Perception and Signal Transduction in Drought StressResponse

Complex Signal Transduction PathwaysSignal-transduction pathways, from the sensing of dehydration or osmotic change to

the expression of various genes, and the signaling, molecules that function in stress signalinghave not been extensively studied in plants. Signal transduction pathways in drought stressresponse have been studied in yeast and animal systems (Fig. 2.6). Two componentsystems function in sensing osmotic stress in bacteria and yeast. Plants as well ascyanobacteria contain many genes encoding sensor histidine kinases and response regulator


homologues, which suggests the involvement of similar osmosensing mechanisms in higherplants. Of course, other sensing mechanisms may function during drought stress responses,such as mechanical sensors in cytoskeltons and sensors for superoxides produced by stress.Stomatal closure is well characterized as a model system in the responses of plant cells todehydration stress and ABA treatment.4,6 During stomatal closure, the level of cytoplasmicCa2+ is increased, which suggests that Ca2+ functions as a second messenger in the osmoticstress response. In animal cells, inositol-3-phosphate (IP3) is involved in the release ofCa2+ into the cytoplasm from intracellular stores, and it may play a similar role in plantcells. Ca2+ and IP3 are the most probable candidates as second messengers in drought-stress responses in plant cells.31

ABA plays important roles in drought stress responses. ABA is involved in not onlystomatal closure but also induction of many genes.6 Several mutants in ABA signalinghave been identified and their genes encode protein phosphatases and farnesyl transferase.These suggest that protein dephosphorylation and protein farnesylation are involved inABA signaling. However, various signaling molecules seem to be involved in ABA signal-ing, such as phosphatidic acid and cyclic ADP ribose. Various protein kinases and enzymesinvolved in phospholipid metabolism have been reported in plants and are thought to func-tion in signal-transduction pathways including drought-stress and ABA responses (Fig. 2.6).4

MAP kinase cascades and calcium-dependent protein kinase were suggested to be involvedin drought stress response and ABA signaling. Complex signaling cascades are thought tofunction in molecular responses to drought stress. Molecular analysis of the signaling process isin progress based on genetics and gene cloning. In this chapter we will describe recent progressof signal transduction cascades from sensing of dehydration stress to gene expression.

Fig. 2.5. A model of the ABA-independent induction of the rd22 gene by drought stress. Droughtstress triggers the biosynthesis of ABA, which induces the expression of two genes for transcriptionfactors, MYC (rd22BP1) and MYB (ATw2) homologues. These transcription factors thenactivate the expression of the rd22 gene.


Sensing of Osmotic stress: The Two-Component SystemIn bacteria, the two-component system functions in sensing and response to osmotic

stress.32 The two-component system is composed of two types of proteins, a sensory histidinekinase and a response regulator. The osmosensing signaling pathway in Escherichia coli iscomposed of one of the two-component system, EnvZ and OmpR. The EnvZ proteinfunctions as an osmosensor, transmits a signal to the histidine kinase domain and activates thekinase. The activated histidine kinase autophosphorylates the histidine residue. The phosphateon the histidine is then transferred to an aspartate residue in the receiver domain of OmpR,and activates the OmpR transcription factor. The activated OmpR regulates the transcriptionof the OmpF and OmpC genes. OmpF and OmpC are porin proteins and form differentpores in the outer membrane. These two porin proteins control cellular osmotic pressurein E. coli. Cyanobacteria also contain numbers of two-component systems, one of which isthought to be involved in osmosensing.

In yeast, exposure to high osmolarity activates a MAPK cascade that includes Ssk2/Ssk22 (MAPKKK), Pbs2 (MAPKK) and Hog1 (MAPK), and then activates several genesinvolved in the biosynthesis of glycerol, which is an important osmoprotectant (Fig. 2.7).32

Three gene products (Sln1, Ypd1, and Ssk1) that act in an early phase of the hyperosmolaritystress response encode signaling molecules that constitute a prokaryote-type two-componentregulatory system.33 Sln1 is thought to act as a sensor protein, phosphorylating Ypd1 andSsk1 response regulator proteins under conditions of high osmolarity. The three proteinfactors perform a four step phosphorelay (His-Asp-His-Asp). At high osmolarity,unphosphorylated Ssk1 activates Ssk2 or Ssk22 (MAPKKKs), which results in the activation ofPbs2 (MAPKK) by Ser/Thr phosphorylation.34 Then, phosphorylated Pbs2 activates Hog1(MAPK) by Thr/Tyr phosphorylation (Fig. 2.7).

Recently, we isolated an Arabidopsis cDNA (ATHK1) encoding the two-componenthistidine kinase, a yeast osmosensor Sln1 homologue, by PCR. ATHK1 has a typicalhistidine kinase domain and a receiver domain like Sln1, and has a different structure in

Fig. 2.6. Second messengers and factorsinvolved in the signal perception and thesignal transduction in drought stressresponse. Two-component histidinekinase is thought to function as anosmosensor in plants. Ca2+ and IP3 aremost probable second messengers of thedehydration signal. Phosphorylationfunctions in water stress and ABA signaltransduction pathways. PI turnover isalso involved in drought stress response.ABA plays inportant roles in the regulationof gene expression and in physiologicalresponses during water stress. SeveralABA signal transduction pathways arereported.


the N-terminal domain from that of ETR1, an ethylene receptor (Fig. 2.7, Urao, Yamaguchi-Shinozaki, Hirayama and Shinozaki, submitted). Overexpression of cATHK1 suppressedthe lethality of a temperature-sensitive osmosensing-defective yeast sln1 mutant (sln1-ts).By contrast, cATHK1, with substitution of conserved His or Asp residues, failed to complementthe sln1-ts mutant, indicating that ATHK1 functions as a histidine kinase. Introduction ofcATHK1 into a yeast mutant lacking two osmosensors (sln1∆sho1∆) suppressed its lethality inhigh salinity media, and activated the HOG1 MAP kinase. The ATHK1 transcript was moreabundant in roots than other tissues under normal growth conditions and accumulated inconditions of high salinity and low temperature. Histochemical analysis of β-glucuronidase(GUS) activities driven by the ATHK1 promoter further indicates that the ATHK1 gene isresponsive to changes in external osmolarity at the transcriptional level. These resultssuggest that ATHK1 might function in signal perception during drought stress in Arabidopsis(Fig. 2.7). A similar osmosensing mechanism might operate in higher plants in responseto a water deficit. In higher plants, a two-component histidine kinase, ETR1, is a receptorin ethylene signal transduction,35 and another two-component histidine kinase, CKI1, isinvolved in cytokinin signaling.36 Two-component histidine kinases may function as sensorsor receptors in various signal transduction pathways in plants.

In Saccharomyces cerevisiae, Sln1 acts as an osmosensory protein, sequentially phos-phorylating a phosphorelay intermediator, Ypd1, and a response regulator, Ssk1, underconditions of normal osmolarity.32 These three protein factors perform a four-stepphosphorelay (His-Asp-His-Asp). Recently, we have cloned four cDNAs encoding aresponse regulator,37 and three cDNAs encoding a phosphorelay intermediator containingan HPt domain65 from Arabidopsis. The existence of response regulators was also reportedin Arabidopsis and maize.38, 39 These observations suggest that plants have an osmosensingand signaling system similar to that of yeast (Fig. 2.7).

Fig. 2.7. Possible roles of the two-componentsystem and MAP kinase cascades in plants incomparison with yeast osmosensing system. Inyeast, Sln1p is thought to act as a sensor proteinphosphorylating Ypd1 and Ssk1p, responseregulator proteins under conditions of highosmolarity. A MAP kinase cascade (Ssk2p/Ssk22p-Pbs2p-Hog1p) functions downstreamof the two-component system in osmotic stressresponse. In Arabidopsis ATHK1, a Sln1phomologue, functions the same as anosmosensor in yeast, which suggests that asimilar two-component system and MAPkinase cascade are involved in drought stresssignal transduction (see text).


Roles of MAP Kinase Cascades in Stress ResponseMitogen-activated protein kinase, or MAP kinase (MAPK), is involved in the signal

transduction pathways associated not only with growth factor-dependent cell proliferation butalso with environmental stress responses in yeast and animals. Many cDNAs for proteinsinvolved in MAPK cascade—MAPK, MAPKK, MAPKKK—have been cloned in plants (Fig.2.7). We have isolated more than nine genes for MAPK in Arabidopsis. There are at leastfour subfamilies of MAPK based on phylogenetic analysis.7 One of the Arabidopsis MAPKgenes, ATMPK3, is induced at the mRNA level by drought, low temperature, high salinityand touch.40 Moreover, two genes for protein kinases involved in the MAPK cascade,MAPKKK (ATMEKK1) and ribosomal S6 kinase (RSK; ATPK19), are induced by similarstresses. We demonstrated rapid and transient activation of MAP kinase activities ofATMPK4 in Arabidopsis plants by low temperature, humidity change, wounding and touch(Ichimura, Mizoguchi, Shinozaki, unpublished data). Recently, alfalfa MAPK, MMK4, wasdemonstrated to be activated at posttranslational levels by a variety of stresses includingdrought, low temperature and mechanical stimuli.41 The MMK4 gene is also induced bythese stresses at the transcriptional level. These observations indicate that certain MAPkinase cascades might function in the signal transduction pathways in drought stressresponse (Fig. 2.5). Recently, a protein phosphatase, 2C, has been suggested to function asa negative regulator of MAP kinase pathways in plants.42 Their roles in drought stressresponse have not yet been elucidated.

Roles of Phospholipids Metabolism and Calcium in Drought Stress SignalingIn animal systems, a variety of phosphoinositides that are metabolrites of membrane

lipids function as second messengers in various signaling processes. Phospholipase C (PLC)digests phosphatidyl-inositol-4, 5-bisphosphate (PIP2) to generate two second messengers,inositol-1,4,5-triphosphate (IP3) and diacylglycerol (DG). IP3 induces the release of Ca2+

into the cytoplasm, which in turn causes various responses in the cytoplasm. DG and PIP2

also function as second messengers and control various cellular responses. In plants, similarsystems are thought to function in drought stress response.43 A gene for phospholipase C,AtPLC1s, is rapidly induced by drought and salt stresses in Arabidopsis.44 Recently, a genefor phosphatidylinositol-4-phosphate-5-kinase (PIP5K) was demonstrated to be inducedby drought stress like the AtPLC1s gene.45 In animals, PIP5K catalyzes the production ofphosphatidylinositol-4,5-bisphosphate (PIP2) from phosphatidylinositol-4-phosphate(PIP). The stress-inducible PLC and PIP5K might function in the signal-transductioncascade under drought stress (Fig. 2.8).

Two genes for calcium-dependent protein kinases (CDPK), ATCDPK1 and ATCDPK2,were induced by drought and high salt.46 These CDPKs may be activated by calcium inresponse to osmotic stress (Fig. 2.8). Recently, coexpression of the constitutively activecatalytic domain of a stress-inducible CDPK, ATCDPK1, has been demonstrated toinduce the expression of an ABA-inducible HVA1 promoter-reporter fusion gene in maizeprotoplasts.47 The HVA1 promoter is also activated not only by cold, high salt and ABAtreatment but also by calcium in protoplasts. These observations also support the idea thatCa2+ might function as a second messenger and that ATCDPK1 functions as a positiveregulator in the ABA signal-transduction pathways under drought-stress conditions inplants.

Recently we isolated two genes for diacylglycerol kinase (DGK) and phosphatidic phosphatase(PAP) that are involved in PI turnover (Fig. 2.8).48 These two genes were demonstrated to beupregulated by dehydration (Katagiri, Shinozaki, unpublished observation). PAP synthesizesdiacyglycerol (DG) by dephosphorylating phosphatidic acid (PA). DGK converts DG into PA.PA is produced from phosphatidylcholine (PC) by phospholipase D (PLD), whereas DG is


produced from PIP2 by PLC. However, the roles of DG and PA as second messengers have notbeen extensively studied in plants. Recently, PLD was shown to be activated by ABA treatment. PAalso increased transiently and is involved in triggering the subsequent ABA response ofaleurone cells.49 These results also support that phospholipids metabolism including IP3 andPA as second messengers is involved in drought stress response.

ABA Signal TransductionThe role of ABA in drought-stress signal transduction has been analyzed genetically

with ABA-insensitive mutants in various species. Maize vp1 and Arabidopsis abi1, abi2, abi3,abi4 and era have been extensively characterized and their genes have been cloned (see chapter1).50 Among them, ABI1 and ABI2 gene products function mainly in vegetative tissues, andalso participate to some extent in seed development. Because of the wilty phenotype of abi1and abi2 mutants, ABI1 and ABI2 are thought to have important roles in ABA-dependentsignal-transduction pathways during drought stress. The ABI1 and ABI2 genes have been

Fig. 2.8. Possible roles of phospholipids metabolism in drought stress and ABA signal transduc-tion pathways. Many genes involved in phospholipids metabolism (PI turnover and PC turn-over), such as PI-PLC, PIP5K DGK and PAP, have been identified and shown to be upregulatedby drought and salt stress. IP3, PIP2, DG and PA are functions as second messengers. IP3 is involvedin release of calcium into cytoplasm to activate calcium dependent signaling moleculesincluding CDPK.


cloned and shown to encode proteins that are related to type 2C protein serine/threoninephosphatases (PP2Cs).51, 52, 53 The ABI1 gene product functions in stomatal closure, and theabi1 plant reveals the wilty phenotype.54 ABI1 was demonstrated to function as a negativeregulator in ABA dependent gene expression in a transient expression experiment usingmaize protoplasts.47 By contrast, the dehydration-inducible ATCDPK1, encoding calcium-dependent protein kinase functions as a positive regulator. These results indicate that aprotein phosphorylation and dephosphorylation process might be involved in ABA-responsive signaling during water deficit. ABI3 and ABI4 were shown to encode transcriptionfactors. Another Arabidopsis mutant, era, that confers an enhanced response to exogenousABA, has mutations in the ERA1 gene encoding the β subunit of farnesyl transferase.55 Thissuggests that a negative regulator of ABA sensitivity may requires farnesylation to function.

Several different signal transduction pathways are suggested to be involved in ABAresponse. ABA was demonstrated to induce a rapid and transient activation of MAPK inbarley aleurone protoplasts.56 Correlation between ABA-induced MAPK activation andABA-induced gene expression implicates that MAPK might be involved in ABA signaltransduction (Fig. 2.3). Recently, cyclic ADP ribose (cADPR) is shown to be involved inABA signal transduction as a second messenger and activate ABA responsive gene expression.57

cADPR is thought to function in the release of Ca2+ in plants. Several reports suggest thatspecific protein kinases, including MAP kinase, are activated in response to ABA treatment inaleulone cells, guard cells and cultured cells.7 Recently, ABA was shown to activate theenzyme PLD to produce PA, which is involved in triggering the subsequent ABA responsesof barely aleulone cells.47 These suggest the existence of several different signal transductioncascades in ABA response. Identification of ABA receptor(s) is important to furtherunderstanding of the signaling process and to understand which pathways are essential inABA signaling.

Regulation of ABA BiosynthesisABA is produced under drought stress conditions de novo, which requires protein

biosynthesis. As mentioned above, this process is important for drought-inducible geneexpression. ABA is thought to be synthesized from xanthophylls via violaxanthin, xanthoxinand ABA-aldehyde (C40 pathway) and the conversion of violaxanthin to xanthoxin is ratelimiting in the ABA biosynthesis under drought stress.58 The C15 pathway found in fungimay also function in plants. Genetic evidence supports only the C40 pathway, and bio-chemical studies suggest that the cleavage of 9-cis-xanthophylls is the key regulatory step.

Many ABA-deficient mutants that do not produce ABA have been isolated in variousplants. An ABA-deficient tobacco mutant, aba2, was isolated by transposon-tagging usingthe maize Ac transposon.59 ABA2 cDNA encodes a chloroplast-imported protein thatexhibits zeaxanthin epoxidase activity, which functions in the first step of the ABA biosyn-thesis pathway. The tobacco ABA2 gene corresponds to the Arabidopsis ABA1 gene. Twosteps in the conversion of xanthoxin to ABA-aldehyde and oxidation of ABA-aldehyde toABA are defined by Arabidopsis aba2 and aba3 mutants, respectively.

An ABA-deficient viviparous mutant of maize, vp14, has been isolated and itscorresponding gene has been cloned by transposon mutagenesis.60 VP14 encodesneoxanthin cleavage enzymes which are involved in the production of xanthoxin from9-cis-xanthophyll in ABA biosynthesis. The VP14 gene is expressed in embryo and rootsand is induced in leaves by drought stress.61 There are several VP14 related genes in maize.We also isolated a VP14 homologue as a drought-inducible gene in cowpea (Iuchi,Yamaguchi-Shinozaki, Shinozaki, unpublished data). These genes are likely to play keyroles in the ABA synthesis in seed development and stress response. Promoter analysis of


the VP14 gene will give us the precise molecular mechanism of the regulation of ABAbiosynthesis under stress conditions.

Conclusion and PerspectivesMolecular mechanisms of drought stress response and tolerance have been actively

studied over the past ten years. Many genes that are regulated by drought stress have beenreported in a variety of plants. Analyses of stress-inducible gene expression have revealedthe presence of multiple signal transduction pathways between the perception of droughtstress signal and gene expression. At least four different transcription factors have beensuggested to function in the regulation of dehydration-inducible genes; two are ABA-re-sponsive and two are ABA-independent. This variety explains the complex stress responseobserved after exposure of plants to drought stress. Genetic analysis of Arabidopsis mutantswith the rd29A promoter—luciferase transgene also suggests complex signaling pathwaysin drought, salt and cold stress responses (see chapter 1).62 Some genes are rapidly inducedby drought stress in 10 minutes, whereas others are slowly induced in a few hours after theaccumulation of endogenous ABA. Several genes for various transcription factors areinduced by drought stress and ABA at transcriptional levels, which might be involved in theregulation of slowly expressed genes whose products function in stress tolerance andadaptation. In addition, many genes for factors involved in the signal transduction cascades,such as protein kinases and enzymes involved in PI turnover, are upregulated by a droughtstress signal.4,7 These signaling factors might be involved in the amplification of the stresssignals and the adaptation of plant cells to drought stress conditions. Molecules that functionas osmosensors and ABA-receptors have not been identified. Based on the knowledge ofosmosensors in yeast and bacteria, cloning of homologues of the two-component histidinekinase as an osmosensor is in progress in higher plants. Molecular analyses of these factorsshould provide a better understanding of the signal transduction cascades during droughtstress. Transgenic plants that modify the expression of these genes will give more informationon the function of their gene products.

Sequencing of the Arabidopsis genome is now in progress and will be completed bythe year 2004, which means that structures of all 20,000 Arabidopsis genes will be determined ina few years.63 All the stress-inducible genes will be identified by systematic analysis of geneexpression. In the next decade, it will be important to develop novel methods to analyzecomplex networks of stress responses of higher plants. We are now constructing insertionmutant lines using T-DNA and transposons to analyze functions of disrupted genes inplants. Reverse genetics approaches as well as classical genetics will become more important tounderstanding not only functions of stress-inducible genes but also the complex signalingprocess in environmental stress responses. An efficient gene disruption method as well astransgenic approaches using antisense or sense constructs will also contribute to moreprecise understanding of the molecular mechanisms of stress response.

AcknowledgmentsThis work was supported by the Program for Promotion of Basic Research Activities

for Innovative Bioscience, the Special Coordination Fund of the Science and TechnologyAgency of the Japanese Government, a Grant-in-Aid from the Ministry of Education,Science and Culture of Japan, and The Human Frontier Science Program.

References1. Ingram J, Bartels D. The molecular basis of dehydration tolerance in plants. Annu. Rev.

Plant Physiol. Plant Mol Biol 1996; 47:377-403.2. Bray EA. Plant responses to water deficit. Trends Plant Sci 1997; 2:48-54.


3. Shinozaki K, Yamaguchi-Shinozaki K. Molecular responses to drought and cold stress.Current Opinion in Biotech 1996; 7:61-167.

4. Shinozaki K, Yamaguchi-Shinozaki K. Gene expression and signal transduction in waterstress response. Plant Physiol 1997; 115:327-334.

5. Giraudat J, Parcy F, Bertauche N, Gosti F, Leung J, Morris P-C, Bouvier-Durand M,Vartanian N. Current advances in abscisic acid action and signaling. Plant Mol Biol 1994;26:1557-1577.

6. Bohnert HJ, Nelson DE, Jensen RG. Adaptations to environmental stresses. Plant Cell 1995;7:1099-1111.

7. Mizoguchi T, Ichimura K, Shinozaki K. Environmental stress response in plants: The roleof mitogen-activated protein kinases (MAPKs). Trends Biotech 1997; 15:15-19.

8. Xu Q, Fu H-H, Gupta R, Luan S. Molecular characterization of a tyrosine-specific proteinphosphatase encoded by a stress-responsive gene in Arabidopsis. Plant Cell 1998; 10:849-857.

9. Holmberg N, Bülow L. Improving stress tolerance in plants by gene transfer. Trends PlantSci 1998; 3:61-66.

10. Tarczynski M, Bohnert H. Stress protection of transgenic tobacco by production of theosmolyte mannitol. Science. 1993; 259:508-510.

11. Kavi Kishor PB, Hong Z, Miao G-U, Hu C-AH, Verma DPS. Overexpression of ∆1-pyrroline-5-carboxylate synthetase increases proline production and confers osmotolerancein transgenic plants. Plant Physiol. 1995; 108:1387-1394.

12. Hayashi H, Mustardy L, Deshnium P, Ida M, Murata N. Transformation of Arabidopsisthaliana with the codA gene for choline oxidase; accumulation of glycine betaine andenhanced tolerance to salt and cold stress. Plant J. 1997; 12:1334-142.

13. Xu D, Duan X, Wang B, Hong B, Ho T-HD, Wu R. Expression of a late embryogenesisabundant protein gene, HVA1, from barley confers tolerance to water deficit and salt stressin transgenic rice. Plant Physiol. 1996; 110:249-257.

14. McKersie BD, Bowley SR, Harjanto E, Leprince O. Water-deficit tolerance and fieldperformance of transgenic alfalfa overexpressing superoxide dismutase. Plant Physiol. 1996;111:1177-1181.

15. Liu Q, Sakuma Y, Abe H, Kasuga M, Miura S, Yamaguchi-Shinozaki K, Shinozaki K. Twotranscription factors, DREB1 and DREB2, with an EREBP/AP2 DNA binding domain,separate two cellular signal transduction pathways in drought and low temperature-responsive gene expression, respectively, in Arabidopsis. Plant Cell 1998; 10:1391-1406.

16. Jaglo-Ottosen KR, Gilmour SJ, Zarka DG, Schabenberger O, Thomashow MF. ArabidopsisCBF1 overexpression induces coe genes and enhances freezing tolerance. Science 1998;280:104-106.

17. Thomashow MF. Arabidopsis thaliana as a model for studying mechanisms of plant coldtolerance. In E. Meyrowitz and C. Sommerville, eds., Arabidopsis. 1994; Cold SpringHarbor Laboratory Press, Cold Spring Harbor, pp. 807-834.

18. Yamaguchi-Shinozaki K, Shinozaki K. A novel cis-acting element in an Arabidopsis gene isinvolved in responsiveness to drought, low-temperature, or high-salt stress. Plant Cell 1994;6:251-264.

19. Stockinger EJ, Glimour SJ, Thomashow MF. Arabidopsis thaliana CBF1 encodes an AP2domain-containing transcription activator that binds to the C-repeat/DRE, a cis-acting DNAregulatory element that stimulates transcription in response to low temperature and waterdeficit. Proc. Natl. Acad. Sci. USA 1997; 94:1035-1040.

20. Kasuga M, Liu Q, Miura S, Yamaguchi-Shinozaki K, Shinozaki K. Improving drought, salt,freezing tolerance by gene transfer of a single stress-inducible transcription factor.Submitted.

21. Koizumi M, Yamaguchi-Shinozaki K, Tsuji H, Shinozaki K. Structure and expression oftwo genes that encode distinct drought-inducible cysteine proteases in Arabidopsis thaliana.Gene 1993; 129:175-182.

22. Kiyosue T, Yamaguchi-Shinozaki K, Shinozaki K. Characterization of cDNA for adehydration-inducible gene that encodes a Clp A, B like protein in Arabidopsis thaliana.Biochem Biophys Res Comm. 1993; 196:1214-1220.


23. Nakashima K, Kiyosue T, Yamaguchi-Shinozaki K, Shinozaki K. A nuclear gene, erd1,encoding a chloroplast-targeted Clp protease regulatory subunit homolog is not onlyinduced by water stress but also developmentally upregulated during senescence inArabidopsis thaliana. Plant J. 1997; 12: 851-861.

24. Weaver LM, Gan S, Quirino B, Amasino M. A comparison of the expression of the patternof several senescence-associated genes in response to stress and hormone treatment. PlantMol Biol 1998: 37:455-469.

25. Shen Q, Ho THD. Functional dissection of an abscisic acid (ABA)-inducible gene revealstwo independent ABA-responsive complexes each containing a G-box and a novel cis-acting element. Plant Cell 1995; 7:295-307.

26. McCarty DR. Genetic control and integration of maturation and germination pathways inseed development. Annu Rev Plant Physiol Plant Mol Biol. 1995; 46:71-93.

27. Schultz TF, Medina J, Hill A, Quatrano RS. 14-3-3 proteins are part of an abscisic acid-VIVIPAROUS1 (VP1) response complex in the Em promoter and interact with VP1 andEmBP1. Plant Cell 1998; 10:837-847.

28. Yamaguchi-Shinozaki K, Shinozaki K. The plant hormone abscisic acid mediates thedrought-induced expression but not the seed-specific expression of rd22, a gene responsiveto dehydration stress in Arabidopsis thaliana. Mol Gen Genet. 1993; 238:17-25.

29. Abe H, Yamaguchi-Shinozaki K, Urao T, Iwasaki T, Hosokawa D, Shinozaki K. Role ofArabidopsis MYC and MYB homologs in drought- and abscisic acid-regulated geneexpression. Plant Cell 1997; 9:1859-1868.

30. Urao T, Yamaguchi-Shinozaki K, Urao S, Shinozaki K. An Arabidopsis myb homologue isinduced by dehydration stress and its gene product binds to the conserved MYB recognitionsequence. Plant Cell 1993; 5:1529-1539.

31. McAinsh MR, Hetherington AM. Encoding specificity in Ca2+ signaling systems. TrendsPlant Sci. 1998; 3:32-36.

32. Wurgler-Murphy SM, Saito S. Two-component signal transducers and MAPK cascades.Trends Biochem Sci. 1997; 22:172-176.

33. Posas F, Wurgler-Murphy SM, Maeda T, Witten TC, Thai TC, Saito H. Yeast HOG1 MAPkinase cascade is regulated by a multistep phosphorelay mechanism in SLN1-YPD1-SSK1'two-component' osmosensor. Cell 1996; 86:865-875.

34. Maeda T, Takehara M, Saito H. Activation of yeast PBS2 MAPKK by MAPKKKs or bybinding of an SH3-containing osmosensor. Science 1995; 269:554-558.

35. Chang C. The ethylene signal transduction pathway in Arabidopsis: An emerging paradigm?Trends Biol Sci. 1996; 21:129-133.

36. Kakimoto T. CKI1, a histidine kinase homologue implicated in cytokinin signal trans-duction. Science 1996; 274:982-985.

37. Urao T, Yakubov B, Yamaguchi-Shinozaki K, Shinozaki K. Stress-responsive expression ofgenes for two-component response regulator-like proteins in Arabidopsis thaliana. FEBSLett. 1998; 427:175-178.

38. Imamura A, Hanaki N, Umeda H, Nakamura A, Suzuki T, Ueguchi C, Mizuno T.Response regulators implicated in His-to-Asp phosphor transfer signaling in Arabidopsis.Proc Natl Acad Sci USA. 1998; 95:2691-2696.

39. Sakakibara H, Suzuki M, Takei K, Deji A, Taniguchi M, Sugiyama T. A response-regulatorhomologue possibly involved in nitrogen signal transduction mediated by cytokinin inmaize. Plant J 1998; 14:337-344.

40. Mizoguchi T, Irie K, Hirayama T, Hayashida N, Yamaguchi-Shinozaki K, Matsumoto K,Shinozaki K. A gene encoding a MAP kinase is induced simultaneously with genes for aMAP kinase and an S6 kinase by touch, cold and water stress in Arabidopsis thaliana. Proc.Nalt. Acad. Sci. USA 1996; 93:765-769.

41. Jonak C, Kiegerl M, Ligterink W, Barker PJ, Huskisson NS, Hirt H. Stress signaling inplants: A MAP kinase pathway is activated by cold and drought. Proc. Natl. Acad. Sci.USA 1996; 93:11274-11279.


42. Meskiene I, Boerge L, Glaser W, Balog J, Brandstoetter M, Zwerger K, Ammerer G, HirtH. MP2C, a plant protein phosphatase 2C, functions as a negative regulator of mitogen-activated protein kinase pathways in yeasts and plants. Proc Natl Acad Sci USA. 1998;95:1938-1943.

43. Munnik T, Irvine RF, Musgrave A. Phospholipid signaling in plants. Biochem Biophys Acta1998; 1389:222-272.

44. Hirayama T, Ohto C, Mizoguchi T, Shinozaki K. A gene encoding a phosphatidylinositol-specific phospholipase C is induced by dehydration and salt stress in Arabidopsis thaliana.Proc. Nalt. Acad. Sci. USA 1995; 92:3903-3907.

45. Mikami K, Katagiri T, Iuchi S, Yamaguchi-Shinozaki K, Shinozaki K. A gene encodingphoaphatidylinositol-4-phosphate 5-kinase is induced by water stress and abscisic acid inArabidopsis thaliana. Plant J 1998; in press.

46. Urao T, Katagiri T, Mizoguchi T, Yamaguchi-Shinozaki K, Hayashida N, Shinozaki K. Twogenes that encode Ca2+-dependent protein kinases are induced by drought and high-saltstresses in Arabidopsis thaliana. Mol Gen Genet 1994; 224:331-340.

47. Sheen J. Ca2+-dependent protein kinase and stress signal transduction in plants. Science1996; 274:1900-1902.

48. Katagiri T, Mizoguchi T, Shinozaki K. Molecular cloning of a cDNA encodingdiacylglycerol kinase (DGK) in Arabidopsis thaliana. Plant Mol Biol 1996; 30:647-653.

49. Ritchie S, Gilroy S. Abscisic acid signal transduction in the barley aleurone is mediatedby phospholipase D activity. Proc Natl Acad Sci USA 1998; 95:2697-2702.

50. Bonetta D, McCourt P. Genetic analysis of ABA signal transduction pathways. TrendsPlant Sci 1998; 6:231-235.

51. Leung J, Bouvier-Durand M, Moris PC, Guerrier D, Chefdor F, Giraudat J. ArabidopsisABA response gene ABI1: Features of a calcium-modulated protein phosphatase. Science1994; 264:1448-1452.

52. Meyer K, Leube MP, Grill E. A protein phosphatase 2C involved in ABA signal transductionin Arabidopsis thaliana. Science 1994; 264:1452-1455.

53. Leung J, Merlot S, Giraudat J. The Arabidopsis ABSCISIC ACID-INSENSITIV2 (ABI2)and ABI1 genes encode homologous protein phosphatase 2C involved in abscisic acidsignal transduction. Plant Cell 1997; 9:759-771.

54. Armstrong F, Leung J, Grabov A, Brearley J, Giraudat J, Blatt MR. Sensitivity to abscisicacid of guard-cell K+ channels is suppressed by abi1-1, a mutant Arabidopsis gene encoding aputative protein phosphatase. Proc Natl Acad Sci USA 1995; 92:9520-9524.

55. Culter S, Ghassemian, Bonetta D, Cooney S, McCourt P. A protein farnesyl transferaseinvolved in abscisic acid signal transduction in Arabidopsis. Science 1996; 273:1239-1241.

56. Knetsch MLW, Wang M, Snaar-Jagalska BE, Heimovaara-Dijkstra S. Abscisic acidinduces mitogen-activated protein kinase activation in barley aleurone protoplasts. Plant Cell1996; 8:1061-1067.

57. Wu Y, Kuzma J, Maréchal E, Graeff R, Lee CH, Foster R, Chua N-H. Abscisic acidsignaling through cyclic ADP-ribose in plants. Science 1997; 278:2126-2130.

58. Kende H, Zeevaart JAD. The five "classical" plant hormones. Plant Cell. 1997; 9:1197-1210.59. Marin E, Nussaume L, Quesada A, Gonneau M, Sotta B, Hugueney P, Frey A, Marion-Poll

A. Molecular identification of zeaxanthin epoxidase of Nicotiana plumbaginifolia, a geneinvolved in abscisic acid biosynthesis and corresponding to the ABA locus of Arabidopsisthaliana. EMBO J 1996; 15:2331-2342.

60. Schwartz SH, Tan BC, Gage DA, Zeebaart JAD, McCarty DR. Science 1997; 276:1872-1874.61. Tan BC, Schwartz SH, Zeevaart JAD, McCarty DR. Genetic control of abscisic acid biosyn-

thesis in maize. Proc. Natl. Acad. Sci. USA 1997; 94:12235-12240.62. Ishitani M, Xiong L, Stevenson B, Zhu JK. Genetic analysis of osmotic and cold stress signal

transduction in Arabidopsis: Interactions and convergence of abscisic acid-dependent andabscisic acid-independent pathways. Plant Cell 1997; 9:1-16.

63. Bevan M, Ecker J, Theologis S, Federspiel N, Davis R, McCombie D, Martienssen R, Chen E,Waterston B, Wilson R, Rounsley S, Ventor C, Tabata S, Salanoubat M, Quetier F, CherryJM, Meinke D. The complete sequence of a plant genome. Plant Cell 1997; 9:476-478.

CHAPTER 3


Molecular Mechanisms of SalinityToleranceHans J. Bohnert, Hua Su and Bo Shen

Plants have evolved complex mechanisms allowing for adaptation to osmotic stress causedby drought and to osmotic and ionic stress caused by high salinity. These mechanisms

can be classified into two categories: One includes developmental, morphological, andphysiological mechanisms; the other includes biochemical mechanisms. Developmental,morphological, and physiological mechanisms are usually complex and require the functionsof many gene products. Examples of complex changes initiated by stress are the switchfrom the C3 photosynthetic pathway to Crassulacean acid metabolism (CAM) inMesembryanthemum crystallinum following salt stress,1 the development of salt glands inLimonium sp.,2 salt-storing epidermal bladder cells in Mesembryanthemum crystallinum3,4

and changes leading to increased water use efficiency in the development of the C4photosynthetic pathway.5

Biochemical mechanisms, in contrast, are relatively simple, typically involving theaction of only a few gene products. For example, the accumulation of compatible solutes,such as glycine betaine, proline, ectoine or polyols, only requires one to three enzymes forextending a main metabolic pathway into the branch pathway of metabolite accumulation.6-9

Similarly, adjustments in ion uptake seem to be controlled by an equally small number ofgene products.10,11 With the current knowledge of plant genetics and biochemistry, thegenetic engineering of biochemical mechanisms is possible, but the engineering of morecomplex traits is still beyond our capabilities. Once all relevant genes are known andfunctionally characterized, it should be possible to manipulate complex developmentalmechanisms, such as flower development, vegetative growth or seed formation. This goalis within reach for the genetic make-up of Arabidopsis thaliana at least,12 but the understandingof how the 21,000 Arabidopsis genes function and their biochemical and physiologicalinteractions lies far in the future. In this review, we will focus mainly on biochemicalmechanisms that lead to cellular and whole-plant adaptations caused by the combinedosmotic and ionic disturbance of metabolism resulting from salt stress.

Over evolutionary time, plants colonized most places on earth, being excluded onlyfrom high latitudes, the highest mountains and true deserts, which are cold and/or lackwater completely. Xerophytic and halophytic adaptations evolved in response to long-termclimate changes which allowed plants to tolerate all but the most extreme habitats. Plantswhich have adapted to stressful environments provide paradigms for biochemical andphysiological tolerance mechanisms, and they provide genes for pathways that couldbecome incorporated into crop plants, which are typically stress-sensitive, having originatedfrom species in subtropical or tropical areas.13-15 Species adapted to extreme habitats arenot equally distributed among all orders or families of the angiosperm lineage. They


appear more frequently in orders which include few crop species but many speciesrestricted to stressful environments. These orders can be considered quarries for obtainingnovel genes for alternative biochemical pathways, and paradigms for understanding howthese pathways interact physiologically.

What constitutes tolerance or resistance to salinity stress has many facets, but issurprisingly simple in principle. For both growth and development of reproductiveorgans, plants must have water for photosynthesis to continue under stress; each one ofthe many diverse mechanisms which evolved in an order-, family- or species-specific fashionmust be subordinate to this essential goal. We will review the molecular mechanisms forwhich the evidence is clear:

1. Scavenging of radical oxygen species,2. Controlled ion uptake,3. The “burning” of accumulated reducing power, and4. Adjustments in carbon/nitrogen allocation.From the confusing multitude of physiological data, a few principles emerge (for

recent reviews, see refs. 11, 14-20, and other articles in this volume). Biophysical andbiochemical principles that govern stress and plant stress responses are outlined by Levitt.21

Osmolytes, Osmoprotectants, Compatible Solutes, OsmoticAdjustment

High salinity disturbs uptake and conductance of water. Salt stress and other environ-mental factors that affect water supply lead to changes in stomatal opening which can, ifstress persists, set in motion a chain of events originating from a decline in the leaf-internalCO2 concentration, consecutively inhibiting the carbon reduction cycle, light reactions,energy charge, and proton pumping.22 Other pathways are affected by increased shuttling ofcarbon through the photorespiratory cycle.14,15 Eventually, carbon and nitrogen allocation andstorage require readjustment, reactions that lead to the consumption of reducing powerbecome favored, and development and growth may become altered. During the past years,the complex interrelationship of biochemical pathways that change during salt stress hasbecome appreciated, although we are far from understanding this complexity.

The accumulation of metabolites, acting as osmolytes, in response to external changesin osmolarity is probably universal.23-25 The generally accepted view is that osmolytes mustbe compatible,26,27 not inhibiting normal metabolic reactions, and that their accumulationleads to “osmotic adjustment” as the major element in accomplishing tolerance.6,7,24,28

Typically, compatible solutes are hydrophilic, giving rise to the view that they couldreplace water at the surface of proteins, protein complexes, or membranes—we might callthem osmoprotectants in this case. The terms carry physiological meaning, but do notexplain the biochemical function(s) such solutes carry out. There may be more than onefunction for a particular solute29,30 and, based on results from in vitro experiments,31-34

different compatible solutes seem to have different functions.The main function of compatible solutes may be stabilization of proteins, protein

complexes or membranes under environmental stress. In in vitro experiments, compatible sol-utes at high concentrations have been found to reduce the inhibitory effects of ions onenzyme activity,24,35-37 to increase thermal stability of enzymes,38-40 and to preventdissociation of the oxygen-evolving complex of photosystem II.41 One argument oftenraised against these studies is that the effective concentration necessary for protection invitro is very high, approximately 500 mM, concentrations which are usually not found invivo. However, when we consider the high concentration of proteins in cells, the concentrationnecessary for protection can, we think, be much lower than that required for protection inin vitro assays. In addition, it may not be the solute concentration in solution that is

31Molecular Mechanisms of Salinity Tolerance

important. Glycine betaine (which may be present in high or low amounts), for example,protects thylakoid membranes and plasma membranes against freezing damage or heatdestabilization,42-44 indicating that the local concentration on membranes or proteinsurfaces may be more important than the absolute concentration.

Two theoretical models have been proposed explaining protective or stabilizingeffects of compatible solutes on protein structure and function. The first is termed the“preferential exclusion model”45 which assumes the solutes are largely excluded from thehydration shell of proteins. Exclusion leaves a water shell around proteins which stabilizesprotein structure, or promotes or maintains protein/protein interactions. In this model,the solutes would not disturb the native hydration shell of proteins, but would interactwith the bulk water phase in the cytosol. The “preferential interaction model”, in contrast,emphasizes interactions between solute and proteins.46 During water deficit, compatiblesolutes may interact directly with hydrophobic domains of proteins and prevent theirdestabilization, or they may substitute for water molecules in the vicinity of such regions.While the two models seem to be mutually exclusive at first, the actual function may in factbe explained by both models. The structures of different solutes could accommodatehydrophobic, van-der-Waals interactions, as well as electrostatic interactions, butadditional biophysical studies will be necessary to gain a better insight into the stabilizingeffects documented by in vitro experiments.

Cellular Mechanisms of Salt Tolerance—the Fungal Model

Osmotic AdjustmentThe unicellular eukaryotic Saccharomyces cerevisiae, baker’s yeast, is an ideal model

for studying cellular and molecular mechanisms of salt tolerance in higher plants. Its smallgenome, which has been sequenced,47-49 adds to several other advantages. First, yeast is salttolerant and the cells have stress responses similar to halophytic plants. Yeast cellsaccumulate compatible solutes, mainly glycerol and some trehalose, to counteract highexternal osmolarity during salt stress;36,50-53 this is similar to the reactions of higher plants.For example, halophytic plants, such as Plantago maritima and Mesembryanthemumcrystallinum, accumulate high concentrations of sorbitol and methylated inositols,respectively, under salt stress.54,55 Second, both yeast and plants use proton gradients asthe driving force of secondary transport systems which control ion fluxes under stress.56, 57

Many ion flux mechanisms are highly conserved in yeasts and plants. In fact, a number ofplant membrane proteins, such as the potassium, amino acid, and sugar transporters, havebeen isolated by functional complementation of yeast mutants.58-61 Third, the accumulationof glycerol is essential for salt tolerance52,53 and glycerol-deficient mutants are available forevaluating the functions of other sugar polyols in stress tolerance. Finally, yeast providesan unsurpassed system for genetic analysis, transformation and functional characterization ofcell-specific functions, especially in light of the recent completion of the yeast genomesequence.48,62

Yeast cells employ two main mechanisms for adaptation to salt stress: accumulationof a polyol, glycerol, and maintenance of ion homeostasis. When exposed to NaCl the cellsexperience both osmotic stress and ion toxicity. To respond to a low external osmoticpotential, the accumulating glycerol seemingly compensates for the difference between theextra- and intra-cellular water potential.36 For reducing sodium toxicity, yeast cells have tomaintain low cytosolic Na+ concentrations and this is achieved by several mechanisms: byrestricting Na+ influx, rapidly extruding Na+ and/or efficiently compartmentalizingsodium into vacuoles. The genetic evidence indicates both mechanisms are essential foryeast salt tolerance.63-65


Glycerol AccumulationYeast cells accumulate glycerol as the major compatible solute when exposed to high

ion concentrations (Fig. 3.1).36 High osmolarity is perceived as a signal by two membraneosmosensors: the protein products of Sln1 and Sho1. The signal is then transferred via aMAP-kinase cascade66-68 and finally enhances the expression of the glycerol biosyntheticpathway. Glycerol is synthesized from dihydroxyacetone phosphate. The first reaction iscatalyzed by glycerol-3-phosphate dehydrogenase which is encoded by two genes, GPD1and GPD2. The second reaction converts glycerol-3-phosphate to glycerol by glycerol-3-phosphatase, encoded by GPP1 and GPP2.52,53,69 The osmotic induction of both GPD genesis mediated by the HOG-MAP kinase signaling pathway. In addition to induced glycerolproduction, yeast cells may decrease membrane permeability to glycerol, which leads to anincreased retention of glycerol in the cells under osmotic stress. In fact, the salt-tolerantyeast Zygosaccharomyces rouxii achieves glycerol accumulation by increased retention ofglycerol within the cell, and probably by active uptake of glycerol rather than by increasedproduction of glycerol during osmotic stress.36 In contrast, Saccharomyces cerevisiaeappears to increase glycerol production, while it fails to significantly alter membranepermeability for glycerol retention during osmotic stress. To maintain high glycerolconcentrations in the cell requires a high energy cost, which seems to limit furtherincreases in salt tolerance in Saccharomyces cerevisiae. A glycerol transport protein (FPS1)which shows homology with MIP-like water channel proteins has been recently isolated.The expression of FPS is not regulated by the HOG-MAP kinase signaling pathway.70

Replacing Glycerol in YeastAlthough the correlation between accumulation of glycerol and yeast osmotolerance

has been established, and although the essential role of glycerol in the adaptation toosmotic stress has been demonstrated by analysis of mutants deficient in glycerolproduction,52,53,71 the mechanism(s) by which glycerol can confer such tolerance is notclear. One obvious possibility is that glycerol is involved in osmotic adjustment tomaintain water flux into the cell. To test whether osmotolerance could be generated by thepresence of polyols other than glycerol, which would support the osmotic adjustmentconcept, we introduced the coding regions of genes encoding enzymes for mannitoland sorbitol production into a glycerol-deficient mutant (Fig. 3.1). However, accumulation ofeither sorbitol or mannitol was not able to replace glycerol function (Shen B, Hohmann S,Bohnert HJ, unpublished). Both foreign polyols accumulated to approximately the sameconcentration as glycerol in wild type, and both were retained by the cell better thanglycerol, but both polyols provided only marginal protection. Growth inhibition of 50%(I50) was 0.6 M NaCl for sorbitol/ mannitol producers in comparison to 0.4 M for themutant and 1.2 M for wild type. By reintroducing one of the deleted yeast GPD genes, asignificant increase in tolerance resulted, and the I50 increased from 0.4 M to 0.9 MNaCl.72 If osmotic adjustment through glycerol is sufficient for salt adaptation, an equalconcentration of sorbitol or mannitol would be expected to confer very similar protection.The results suggest that the concept underlying the term “osmotic adjustment” may not bevalid, or valid only if the synthesis of a metabolite for osmotic adjustment fullfills species-specific requirements. The consequence of our results, then, is that glycerol might have specificprotective functions which mannitol and sorbitol cannot replace. Evolutionary adaptationsmight have altered yeast proteins such that glycerol, but not other osmolytes, could exert aprotective role. Alternatively, the pathway through which an osmolyte is produced could bemore important than the end-product. Finally, the minimal protection by mannitol or sorbitolcould be caused by a difference in their intracellular distribution compared to glycerol.


Whether and how these polyols are compartmentalized in yeast cells is not known, butglycerol seems to be evenly distributed.73

A major difference exists between glycerol and mannitol/ sorbitol synthesis andaccumulation with respect to energy expenditure. Glycerol biosynthesis, which is also arequirement for the removal of excess NADH during anaerobiosis,71 is more costly thanmannitol/ sorbitol generation. While NADH oxidation is, in principle, also accomplishedby the mannitol and sorbitol metabolic pathways, which leads to NAD+ increase, the costis different. During salt stress, more than 95% of the glycerol produced leaked from thecells and accumulated in the medium. In contrast, sorbitol and mannitol did not significantlyexit from the cells. We calculated that glycerol biosynthesis under stress conditionsconsumed at least 10 times more carbon and NADH than sorbitol/ mannitol biosynthesis.72

Thus, it may be that “burning” of excess reducing power via glycerol biosynthesis is asimportant as the increasing osmotic potential provided by the steady-state glycerolconcentration in the cytosol.

Fig. 3.1. Genes involved in yeast osmotic stress signal transduction, and replacement of glycerolsynthesis by foreign osmolytes. The schematic drawing of a yeast cell includes the membraneosmosensors (Sln1 and Sho1) which transmit signals to a MAP kinase cascade. Specifictranscription is initiated, which leads to the synthesis of several proteins, among them glycerol-3-phosphate dehydrogenase (GPD) and glycerol phosphatase (GPP). This results in glycerolsynthesis and accumulation. The glycerol facilitator protein, FPS1, a MIP-type channel, is lesspermeable under stress than under normal growth conditions. Replacement of both genesencoding GPD by mannitol-6-P dehydrogenase or sorbitol-6-P dehydrogenase leads to theaccumulation of mannitol or sorbitol to a concentration approximately equal to glycerolaccumulation, but the two foreign polyols only marginally improve salt tolerance of the cells. 72


Osmosensing and SignalingThe signaling pathways by which yeast cells respond to external osmolarity changes

has been identified (Fig. 3.1). High osmolarity is perceived as a signal by two membraneosmosensors which are the protein products of Sln1 and Sho1. The Sln1 protein contains anextracellular sensor domain, a cytoplasmic histidine kinase domain, and a receiver domain.YPD and Ssk1 receive sensor signals in the cytosol. Sln1 and YPD/Ssk1 function like bacterialtwo-component systems.67,74,75 Sho1 is a transmembrane protein containing a cytoplasmicSH3 domain which can directly activate a MAP kinase kinase, Pbs2, by interaction betweenits SH3 domain and a proline-rich motif of Pbs2.68 The signal from Sln1 is transmitted via aMAP kinase cascade encoded by MAP kinase kinase kinase (Ssk2/22), MAP kinase kinase(Pbs2), and MAP kinase (Hog1).66-68 The cascade initiates the expression of the glycerolbiosynthetic pathway including the GPD1 and GPP2 (phosphatase) genes.52,69 In additionto glycerol biosynthesis, genes for other stress responses, such as CCT1 encoding catalase T76

and HSP12 encoding a small heat-shock protein,77,78 are induced by this signaling pathway.In contrast to hyperosmotic stress, hypoosmotic stress initiates a second MAP kinase

cascade called the protein kinase C1 (PKC1) pathway. The MAP kinase in this PKC1pathway is phosphorylated when cells are transferred from high osmolarity to low osmolarity.Protein kinases downstream of PKC1 include BCK1/SLK1, MKK1/MKK2 and MPK1/SLT2.79

PKC1 mutants exhibited a lytic phenotype due to defects in cell wall biosynthesis. Thelytic phenotype can be suppressed by the addition of osmolytes like sorbitol into themedium.80 How the two osmosensing pathways are coordinated remains to be determined.

Ion RelationsThe yeast genome contains approximately 5,800 genes which potentially encode proteins.48

About 250 genes show significant similarity to membrane transport proteins characterized inyeast and other organisms. Among those, a (partial) functional characterization existedfor only about 60 genes prior to the completion of the sequence, which amply documentsboth the value of sequencing projects and our relative ignorance of membrane transportprocesses in general.81 Many of these membrane transport proteins are involved in iontransport and carry out essential functions in salt tolerance.

Potassium TransportPotassium plays an important role in yeast salinity tolerance. The osmotic potential

generated by high internal potassium concentrations (e.g., in halobacteria) can alleviatesodium toxicity.36 Three membrane proteins are involved in potassium transport acrossthe plasma membrane, TRK1, TRK2, and TOK1.82-84 The TRK proteins are involved in K+

influx and TOK1 controls K+ efflux. TRK1 and TRK2 are required for high affinity andlow affinity potassium uptake, respectively. Importantly, TRK proteins can also transportNa+ but both have a higher affinity for K+. Under high external Na+ concentrations, Na+

can inhibit K+ uptake and enter the cell through the potassium channels. The capacity fortransporting potassium into cells and restricting sodium influx by increasedK+discrimination over Na+ is an essential element for salt tolerance acquisition.85-87

Because the high affinity K+ transport system shows a higher K+/Na+ discrimination thanthe low affinity system, under salt stress yeast cells may shift from low to high affinity K+

uptake, allowing the cells to accumulate more K+ than Na+ and to maintain a low Na+/K+

ratio.85,88 Increased K+/Na+ discrimination of a high affinity potassium transporter (HKT1)from wheat has been shown to increase salt tolerance of yeast strains deficient in potassiumuptake.86

Two halotolerance genes, HAL1 and HAL3, have been isolated by screening for genesthat enhance salt tolerance when overexpressed.89,90 Both have been implicated in the


regulation of K+ concentrations. Overexpression of HAL1 and HAL3 resulted in a strongeraccumulation of K+ under salt stress and increased salt tolerance. The beneficial effects arespecific for NaCl stress, and cannot moderate osmotic stress by sorbitol or excess KCl,suggesting that high K+ in a physiological range may specifically alleviate sodium toxicity.Yeast double mutants with deletions of the high affinity K+ transporter (TRK1) and theNa+-ATPase (ENA1) genes are sensitive to Na+ because of poor Na+/K+ discriminationand decreased Na+ efflux.85

Similarly, long-term salt-adapted tobacco cells showed increased capacity for K+

uptake compared to wild type cells,91 suggesting better Na+/K+ discrimination by the K+-uptake system as a significant element for salt tolerance. Likely in the same category,Arabidopsis sos1 mutants were hypersensitive to salt stress due to a defect in the high affinity K+

uptake system, highlighting the important role of K+ for salt tolerance in plants.92

H+-ATPasesPlasma membrane and vacuolar proton ATPases are essential for generating and

maintaining membrane proton gradients and for pH regulation in yeast and plants.93-96

They must be able to sense and respond to external acidification. The yeast plasma membraneH+-ATPases (P-ATPase), encoded by the gene PMA1, is predominantly responsible forproton gradient maintenance, while the product of the PMA2 gene is induced at low pHwhen the PMA1 protein cannot function properly.97 Regulation of activity by calcium-dependent protein kinases, in response to glucose levels, weak organic acids, heat shockand salt stress, has been shown.98,99 Mutants hypersensitive to the immunosuppressantscyclosporin A and FK506 were shown to be defective in assembly of the vacuolar H+-ATPase(V-ATPase). Their characterization indicated involvement of the calcineurin signaltransduction pathway in synthesis, endomembrane transport, assembly and activityregulation.98,100,101

Sodium Transport Across MembranesMaintaining low intracellular sodium amounts during salt stress is essential for yeast.

Low sodium concentrations in the cytosol could be achieved by decreased Na+ uptake,increased Na+ efflux, transport of Na+ into vacuoles or a combination of such activities. InS. cerevisiae, a Na+-ATPase encoded by a family of 4 or 5 ENA genes has been shown to beinvolved in sodium efflux. The expression of the ENA1 gene is induced by salt stress whilethe other genes are expressed constitutively and weakly. Mutants defective in the ENAfunction are sensitive to sodium and lithium.65,102,103 Also, an Na+/H+ antiport protein,encoded by Nha1, was found during sequencing of the genome81 and later functionallycharacterized.104 NHA1 seems to play a minor role in sodium efflux, but it may be important inan environment of acidic external pH which would affect the transmembrane protongradient.

Based on the results with yeast it was surprising, however, that in Schizosaccharomycespombe and Zygosaccharomyces rouxii the major sodium efflux is via such a Na+/H+-antiporter encoded by the SOD2 gene. SOD2 was initially identified by selection forincreased LiCl tolerance in fission yeast105 and the homologous SOD2 was isolated fromZ. rouxii.106 Functional expression of ENA1 in a sod2 mutant of S. pombe restored Na+

efflux and salt tolerance. Recently, the activity of the SOD2 Na+/H+ antiporter was con-firmed using microphysiometry, indicating reversible sodium transport, dependent on theNa+ and H+ gradient across the membrane.107 Based on this property, it should be possibleto utilize SOD2 to transport Na+ into vacuoles by targeting the protein to the tonoplast inhigher plants.


Although mechanisms of Na+ uptake in yeast are still not understood, mutantanalysis has clearly demonstrated an essential role for membrane-located processes.Disruption of the LIS1/ERG6 gene, encoding a SAM-dependent methyltransferase of theergosterol pathway, resulted in increased sodium uptake and decreased salt tolerance. Themutation seems to affect cation transport indirectly by changing membrane composition.108

Several other uncharacterized mutants showing high internal sodium were salt sensitivedespite normal glycerol accumulation.63,64 Halophytic plants usually sequester Na+ intovacuoles to lower the concentration of Na+ in the cytoplasm.3 Whether such a mechanismexists in yeast is unknown, but evidence exists for the vacuole’s important role in salttolerance. Mutants defective in vacuole morphology and vacuolar protein targeting aresalt-sensitive.109 A mutant in subunit C of the vacuolar ATPase shows increased sensitivityto Na+ and Li+.85 The essential function may be associated with both compartmentationof ions and osmoregulation.

Calcineurin SignalingThe signal transduction pathway regulating ion homeostasis remains unknown in

detail, but it is known to be different from the HOG pathway. Recent studies revealed thatcalcineurin and protein phosphatase PPZ seem to be involved in the regulation of ionfluxes.88,110-112 Calcineurin, a protein phosphatase 2B consisting of a catalytic subunit (CNA)and a regulatory subunit (CNB), requires Ca2+ and calmodulin for activity.113 Nullmutants of calcineurin fail to recover from G1-arrest in the presence of α-pheromone, butshow normal growth rates under normal growth conditions. Under salt stress, however,the mutants exhibited a salt-sensitive phenotype,88,110 caused by reduced expression of theENA1 gene which is regulated by calcineurin. Also, calcineurin mutants cannot shift fromlow- to high-affinity potassium transport under salt stress.88 In contrast, deletions of genesfor the protein phosphatases PPZ1 and PPZ2 increased salt-tolerance due to enhancedexpression of ENA1, suggesting an essential role of these phosphatases in yeast ionhomeostasis.112 At low salt concentrations, the HOG-MAP kinase pathway appears to beinvolved in regulation of ion fluxes, while at high salt concentrations ion balance is mainlycontrolled by calcineurin.114 Interestingly, calcineurin signaling seems to interact with theMAP kinase pathway. Disruption of the calcineurin gene (Ppb1) in fission yeast resulted insensitivity to chloride. High copy number of the Pmp1 gene, encoding a phosphatase,suppresses this sensitivity to chloride. The PMP1 phosphatase dephosphorylates PMK1,the third MAP kinase in fission yeast. As expected, deletion of Pmk1 also suppresses thechloride sensitivity of calcineurin mutants.115 Other components in the calcineurinsignaling pathway remain to be identified.

In addition to calcineurin and PPZ, HAL1 and HAL3 are involved in the regulation ofintracellular Na+ and K+ concentrations.90,116 The effect of HAL1 and HAL3 on intracellularNa+ is mediated by expression of the ENA1 gene. Calcineurin plays a role in the inductionof ENA1 expression by sodium, while HAL1, HAL3 and PPZ determine the basal level ofENA1. HAL1 and HAL3 are then required for the maximal expression of ENA1 under saltstress. Overexpression of HAL1 or HAL3 partially suppressed the salt sensitivity ofmutants with a non-functional calcineurin. Increased K+ by overexpression of HAL1 isindependent of the action of TRK1 and TOK1, probably due to decreased export of K+

during salt stress.116 Clearly, multiple regulatory pathways and control circuits govern ionresponses in a complex interaction depending on external signals.

In higher plants, a calcineurin-like protein phosphatase activity has been found inthe regulation of the K+-channel in guard cells of fava bean.117 FK506 and cyclosporinA, immunosuppressants which bind to cellular receptors, are strong and specificinhibitors of calcineurin.118 When FK506-receptor complexes were added to guard cells,


the Ca2+-induced inactivation of K+ channels was inhibited. A Ca2+-dependentphosphatase activity which is sensitive to complexes of FK506 and its binding proteinand to cyclophilin-cyclosporin A was also identified in guard cells.117 The gene whichencodes cyclophilin has been cloned,119 suggesting an important role of calcineurin inhigher plants. In addition, by complementation of yeast calcineurin mutants, two cDNAs(STO and STZ) which suppress the calcineurin deficient phenotype in yeast have beenisolated from plants, but the predicted protein sequence did not show significanthomology to phosphatases.120 Until now, the plant calcineurin gene has not beenidentified. The important function of calcineurin in salt tolerance has recently beendemonstrated by overexpression of a truncated yeast calcineurin in transgenic tobacco.Constitutive expression of this yeast calcineurin increased salt tolerance of transgenictobacco plants (Bressan RA, Pardo J, personal communication).121

Molecular Mechanisms of Salt Tolerance in Plants

Metabolite AccumulationAccumulation of compatible solutes during osmotic stress is a ubiquitous biochemical

mechanism, present in all organisms from bacteria, fungi and algae to vascular plants andanimals.24,36 The accumulating metabolites include amino acids, their derivatives (proline,glycine betaine, β-alanine betaine, proline betaine), tertiary amines, sulfonium compounds(choline o-sulfate, dimethylsulfoniopropionate), the raffinose series of sugars, and polyols(glycerol, mannitol, sorbitol, trehalose, fructans, and methylated inositols).6,14,15, 122,123

Enzymes from halophytes do not show remarkably higher salt resistance than thosefrom glycophytes, nor do they require sodium for optimal activities. In fact, the activity ofenzymes from both is generally strongly inhibited by high concentrations of either NaClor KCl.3 Although halophytes and glycophytes use similar compatible solute strategies todeal with osmotic stress,124 they use different strategies to cope with ion toxicity. Halophytestake up sodium and sequester ions into the vacuole. High osmotic potential in vacuoles isbalanced by accumulating compatible solutes in the cytoplasm. Because the cytoplasmicvolume is relatively small compared to the large volume of the vacuole, low concentrationsof compatible solutes suffice to reach the same osmotic potential in the cytoplasm. Incontrast, glycophytes usually attempt to limit sodium uptake or transport sodium to oldleaves as an alternative way to extrude sodium out of plants.3,125,126 The halobacteria deviatefrom the general compatible solute strategy, accumulating K+ as the osmolyte rather thanorganic solutes to counteract high external osmotic potential. Halobacterial enzymesrequire high ion concentration for their optimal activity.36 This adaptation required changesin protein structure. During evolution, this type of stress adaptation was abandoned,possibly because it proved inflexible to changing environments, and mechanisms becamefavored which utilized organic solutes, likely because they could be synthesized throughpathways attached to basic metabolism.

Several common features characterize the different compatible solutes. First, they caneasily be synthesized from compounds diverted from basic metabolism by novelenzymatic or regulatory reactions. For example, glycine betaine in higher plants issynthesized from choline via two reactions catalyzed by choline monooxygenase andbetaine aldehyde dehydrogenase,6 and pinitol is synthesized from myo-inositol in tworeactions catalyzed by inositol o-methyltransferase and ononitol epimerase.4,8,15 Bothcholine and myo-inositol are high-flux metabolites and are tightly regulated during growth.Second, accumulation of compatible solutes under osmotic stress is an active process, ratherthan an incidental consequence of other stress-induced metabolic changes. The biosyntheticpathway for a particular osmolyte is coordinately up-regulated during osmotic stress. For


example, two key genes, Inps1 and Imt1, are transcriptionally enhanced by salt stress, andhigher enzyme amounts lead to increased carbon flux through myo-inositol into pinitolbiosynthesis in stressed Mesembryanthemum.8,127-129 Genes involved in the degradation ofcompatible solutes are down-regulated under osmotic stress. This is, for example, the casefor proline oxidase in Arabidopsis thaliana. Stress-dependent lower expression of thisenzyme, at least in part, may explain the increases in proline during salinity and droughtstress.130 Third, many accumulating compounds are end-products of a branch pathwayrather than active intermediates in so far as one enzyme in the pathway catalyzes only theforward reaction. Examples for this point are DMSP synthesis in marine algae,9,131 pinitolsynthesis in Mesembryanthemum4,55 and glycinebetaine synthesis.6,132-134 Equally, prolinebiosynthesis has received much attention, because proline accumulation is a nearly universalreaction of plants to osmotic stress.135-137 Its true role in stress protection is, however, notclear—we consider the accumulation of proline a consequence of the necessity forreadjusting carbon nitrogen balance under stress.138 The biosynthesis of ectoine (tetra-hydropyrimidine and derivatives), an accumulating osmolyte in bacteria, has received

Table 3.1 Transgenes with Effects on Salt-, Drought- and Low TemperatureTolerance

ROS Scavenging Enzymes 1991 SOD, catalase, GST/GSX overexpressionleading to enhanced stress tolerance.20,181,232,233,236,238

Mannitol Synthesis 1992 Protection against salt stress.251,252,253

Fructan Accumulation 1995 Enhanced drought tolerance.254

Proline Accumulation 1995 Enhanced salt stress tolerance.135

Glycine betaine Synthesis 1997 Enhanced temperature stress, salt stress tolerance.255

LEA Protein Synthesis 1996 Salinity and drought stress protection.256

Potassium Transporter 1994 Enhanced Na+/K+-discrimination in yeast.86,207

Trehalose Synthesis 1996 Enhanced drought tolerance.141

Glutathione Cycle 1997 Altered redox control, salt and low temperatureEnhancement protection.192

Mannitol as a Hydroxyl 1997 Enhanced salt tolerance, mannitol synthesis inRadical Scavenger chloroplast.29,30

Inducible Ononitol 1997 Enhanced drought and salt tolerance; inducibilityAccumulation based on changes in substrate amounts.138

Extreme Sorbitol 1998 High accumulation, >600 mM sorbitol, leading toAccumulation necrotic lesions in sink leaves.227


attention recently.130,140 Expression of the three enzymes leading to ectoine in bacteriaconfers significant salinity tolerance. Figure 3.2 shows schematically selected pathways thatlead to the synthesis of polyols (mannitol, sorbitol, ononitol and pinitol) and to trehalosesynthesis.141 Apart from the pinitol biosynthetic pathway,8,11 the pathways shown areengineered pathways (Table 3.1) and may be different from pathways existing in someplant species naturally. The scheme indicates clearly how the addition of a single gene canbe exploited for metabolic engineering.

Water ChannelsWater channels, aquaporins (AQP), are found in all organisms as members of a

super-family of membrane proteins, 26-30 kDa in size, termed MIP (major intrinsicprotein).142,143 The proteins are characterized by six membrane-spanning domains and apore-domain with a characteristic sequence signature, NH3-NPAXT-COOH. Aquaporinsenhance membrane permeability to water in both directions depending on osmotic pressuredifferences across a membrane, but other members of the gene family in yeast and vertebratesencode glycerol-facilitators.143 Other MIPs, animal and plant—among them a nodulation-specific protein, may mediate ion transport and transport of other neutral metabolites, such asurea.144,145

Fig. 3.2. Pathways for the synthesis of selected compatible solutes. Biochemical pathways originatingfrom glucose-6-P or sorbitol-6-P whose presence in some stress-tolerant species or after genetransfer into transgenic tobacco is correlated with increased osmotic stress tolerance. Genes/enzymes used in transgenic experiments are PGM (phosphoglucomutase), INPS (myo-inositol1-P synthase), IMT (myo-inositol O-methyltransferase), GPDH (sorbitol-6-P dehydrogenase),MtlDH (mannitol-1-P dehydrogenase), TPS (trehalosephosphate synthase). Pase indicatesunspecific phosphatases. OEP (ononitol epimerase) is found in Mesembryanthemum, but thegene has not yet been cloned. IMP (myo-inositol monophosphatase) is not regulated inMesembryanthemum during stress and has not been included in transgenic plants.


Complexity of Plant IsoformsIn human DNA, five MIP genes have been characterized among a total of seven MIP-like

genes. They are expressed in different tissues, most highly in erythrocytes, kidney cells andthe brain. In contrast, Arabidopsis contains at least 23 MIP-like coding regions.146 Sequencesignatures of the Arabidopsis MIP indicate two large sub-families of 10 to 12 proteins eachwhose members are either plasma membrane-located (PIP) or tonoplast-located (TIP),and one MIP which diverges from the others has not been characterized.146 While some ofthe genes might encode facilitators for diverse small metabolites or ions, eight MIPproteins have already been identified as aquaporins. Why are there so many plantaquaporins? We discuss four possibilities which might explain the high number.

1. MIP-intrinsic functional variations might allow AQP to be active at differentmembrane osmotic potentials. Yet, all we know is that the intrinsic water permeabilitydistinguishes four human AQP and one glycerol facilitator by a factor of ~100,147 andthat plant AQP can be either sluggish or effective water transporters whenexpressed in Xenopus oocytes.143 There is no report about a functional plant modelthat would allow mechanistic studies on AQP. By antisensing with a plasma membraneAQP coding region148 which supposedly targeted all expressed PIP, the decrease inAQP amounts led to a decline in water uptake in plants. Such antisense AQPtransgenics increased the root to shoot ratio, suggesting a feedback mechanismbetween water uptake and root mass (Kaldenhoff R, personal communication).Protoplasts from the antisense-expressing plants did not burst as fast as wild typecells when transferred to hypoosmotic solutions.148

2. Functional differences could have evolved for fine tuning water flux through theplant—with high conductance AQP located in the root cortex and vascular tissueswhich accommodate bulk fluxes and low conductance channels betweenmesophyll cells, for example, or even within the cell cytosol and organelles and thevacuole.

3. Without assuming functional diversification, the number of AQP arising throughgene duplications could have changed gene and protein expression, half-life, andturnover such that AQP amount shows a gradient that follows the water transportgradient. In this scenario, the gene number—requiring different promoters, RNA-stability and translation characteristics and protein half-life regulation—would bedetermined by the necessity of cell-specific differences in accommodating waterflux and not by the water transport function per se. This explanation is similar tothe following one, and both find precedence in the presence of, for example, alarge number of genes encoding plasma membrane H+-ATPases, AHA, which aredifferentially expressed throughout the plant.95,149,150 Deletion of several AHA genesdid not produce a phenotype under normal growth conditions, but affected growthsignificantly under diverse growth and stress conditions, low temperature, salt stressand external high acidity, for example (Sussman MR, personal communication).

4. Last, AQP/MIP multiplications and diversifications could have been dictated bythe need for a flexible response to environmental changes in water supply orevaporation, demanding the presence of several sets of AQP. This assumes evolution ofone set of AQP genes for stress responses and that this set is different from others.It is conceivable that a set of Mip genes exists to take care of the business of cellexpansion following meristematic activity—and this function (missing from animals)might require regulatory circuits different from those necessary in genes that performhousekeeping (set 2) and stress-response functions (set 3). Although the data arenot complete with respect to AQP protein expression and cell-specificity, alignmentsof sequences indicate that sub-families of two to four closely related sequences


exist146,151 which might represent the three sets of genes. MIP associated with cellexpansion,152 developmental specificity 153,154 and stress functions151,155-158,257 havebeen described.

Mechanisms of RegulationMost important to the topic here is how MIP gene expression, protein amount and

aquaporin activity are controlled during development and under environmental stress.Regulation is by gene expression and protein amount, and possibly also by post-translationalmodification—but we have very little information on mechanistic details in plants.

Weig et al146 used quantitative PCR amplification for the 23 Arabidopsis MIP andfound differences in mRNA amounts spanning several orders of magnitude. Differencesin RNA amounts for each MIP in roots, leaves, bolts and the flowers and siliques wereequally pronounced. No signals were detected for at least three MIP, suggesting that thesemight be expressed under conditions not found during normal growth or that they areexpressed in a few cells only or at very low levels. The analysis of such a large gene family,once all genes are known, can best be done by in situ hybridization, immunocytology withspecific antibodies and DNA microarray analysis through which the amount, location andregulation of the genes during development and under different environmental conditionscan be monitored. For several MIP in a number of organisms, salt stress altered mRNAamounts have been reported. AQP expression also responds to drought and low temperature,hormone treatment (ABA, cytokinine, GA), light, and pathogen infection.143,157,158

Promoter studies have been performed with several MIP, but cell-specificity ismost likely the essential distinguishing factor between AQP and must receive moreattention in order to understand water transport in plants. The promoter for Rb7a159 fromtobacco conveys root-specificity, leads to differential expression in the root in a cell-specific manner and is induced by nematode feeding.156,159 The Mesembryanthemum MipBpromoter showed highest expression of the gene in roots;151 after transfer into tobaccoand observation of GUS expression, broader specificity was observed, with highest expressionin all meristematic cells and in vascular tissues.160

Even less complete is the information about protein amount, localization and changesduring development and under stress conditions. One essential consideration is that thelarge number of genes and high sequence identity among PIP and TIP, respectively,require excellent controls for avoiding cross-hybridizations between transcripts andimmunological cross-reactivity between antisera. For example, generation of anti-peptideantibodies against six Mesembryanthemum MIP resulted in distinguishable signals todifferent cells.161 However, in the absence of probes for all MIP for this species, it cannotbe excluded that some of the antibodies react to more than one MIP whose sequence is notyet known, but shares homology with the selected peptide domain.

Regulation has been documented at the level of post-translational modification, mostlyin animal systems. Salt stress conditions in kidney cells lead to changes in protein expres-sion, which may be controlled by oligomerization, glycosylation, or phosphoryla-tion.162,163 In addition, the presence in the cell membrane and the half-life of AQP isdetermined by the hormone vasopressin in animal cells. Increased vasopressin leads to thedeposition of AQP from internal stores, endosome vesicles, to the outer membrane, andlower hormone levels lead to cycling of membrane patches through endosomes.164 Clearly,such traffic and its control would constitute the fastest, most economic way of regulatingwater flux. Similar observations remain to be made with plant MIP, but patches of invaginatedplasma membrane regions, termed “plasmalemmasomes” that contain abundant AQPprotein have been found in plants,165 possibly the functional equivalent of animalendosomes. Our preliminary experiments indicate that PIP from Mesembryanthemum


sediments in different gradient fractions depending on whether salt-stressed or unstressedcells were used,161 which might indicate that similar membrane shuttle mechanisms existin plant cells.

Evidence for plant AQP regulation comes from studies which measured AQP phos-phorylation.153,166 Regulatory sites for phosphorylation have been mapped in severalMIP/AQP.143 Also, effects of pharmaceutical agents on water flow in Chara cells, forexample, point towards an association of water flux and the integrity of the cytoskeleton (seeref. 143). Spinach leaf PIP are reversibly phosphorylated in response to the apoplasticwater potential and calcium.166

The discovery and preliminary characterization of AQP in plants has provided morequestions than answers. Their existence cannot be questioned and they act as waterchannels. It is then intuitively obvious that control over their action should be importantunder stress conditions. Although there are few data available, it is equally clear that regulationduring stress is complex, involving transcriptional and post-translational controls whichseem to involve synthesis, membrane traffic and reversible insertion into membranes,complex assembly and MIP protein half-life.

Salt Stress and Radical Scavenging

Reactive Oxygen Species and Radical Scavenging SystemsProduction of Reactive oxygen species (ROS) is an unavoidable process in photosynthetic

tissues, but ROS are also produced in mitochondria and cytosol. ROS including singletoxygen, superoxide, hydrogen peroxide, and hydroxyl radicals react with and can damageproteins, membrane lipids, and other cellular components.33,167,168 Some ROS also serveas signaling molecules,20 for example, in the initial recognition of attack byfungal pathogens and the transmission of signals after a primary infection.169,170 Focusing onchloroplasts, superoxide is abundantly produced from photoreduction of oxygen. Oxygenconcentration as high as 300 mM can be photoreduced to superoxide by photosystem I viaa Mehler reaction.171,172 The production of superoxide has been estimated to be approximately30 mmol (mg chl)-1 h-1 in intact chloroplasts,173 and the rate of production in isolatedthylakoids was increased 1.5-fold by the addition of ferredoxin and decreased 50% byaddition of NADP+.174 Most of this thylakoid lumen-produced superoxide diffuses to thestroma.173 H2O2 in chloroplasts is predominantly generated by disproportionation ofsuperoxide by SODs. In peroxisomes, H2O2 originates directly from glycollate oxidaseactivity. Hydroxyl radicals derive from an interaction between hydrogen peroxide andsuperoxide or directly from hydrogen peroxide in the presence of transition metals such asFe+2 and Cu+ by a Fenton- or Haber-Weiss-reaction. The oxidized metal ions can bere-reduced by superoxide, glutathione, or ascorbate. Trace amounts, lower than the amountpresent in chloroplasts, of metal ions are needed to catalyze the Fenton reaction.168,173 Ithas in fact been shown that elevated amounts of iron lead to increased oxidative stress.175

These Reactive oxygen species are scavenged by resident enzyme systems andnonenzymatic antioxidants.176 Non-enzymatic detoxification mechanisms includemorphological features such as waxy surfaces and leaf or chloroplast movement, non-photochemical quenching processes by various compounds, for example, the violaxanthin-zeaxanthin cycle, and photorespiration. Non-enzymatic antioxidants include flavonones,anthocyanins, α-tocopherol, ascorbate (at a concentration of ~10 mM in chloroplasts),glutathione, carotenoids, phenolics and polyols.20,32,168,177 Botanical sources of suchantioxidants not only play important roles in plant stress adaptation, but also retard agingand diseases related to oxidative damage in animals.178


The enzyme systems involved include SODs which catalyze the reaction from superoxideto hydrogen peroxide, and ascorbate peroxidases (APX) responsible for the conversion ofhydrogen peroxide to water. Both SOD and APX are represented by isoforms localized tothe stroma and the thylakoid membrane. Ascorbate can be regenerated by the ascorbate-glutathione cycle. The level of reduced glutathione is maintained by glutathione reductaseusing NADPH.168,179,180 In addition, catalase has recently been demonstrated as a sink forH2O2 in C3 plants.181 In contrast to the detoxification systems for H2O2 and O2

-, anenzyme system that could deal with the short-lived, extremely toxic hydroxyl radical hasnot been identified and, in fact, might not have evolved.167,168,179,180 The best way ofdetoxifying hydroxyl radicals is to prevent their formation by reducing the concentrationof H2O2 and free metal ions. Once produced, however, protection depends on the presenceof antioxidants in the vicinity of the formation site. Together these systems provide sufficientprotection under normal growth conditions; in fact, the scavenging systems are able tohandle moderate increases of ROS, unless long-term stress exceeds the detoxificationcapacity.20,179,182 In chloroplasts, oxidative damage includes first a decline in CO2 fixation,and then inhibition of photochemical apparatus, loss of pigments, oxidation of proteins,and lipid peroxidation.183,184

ROS and Environmental StressSeveral lines of evidence support the toxicity of ROS during drought,20 chilling stress184

and salt stress.29,30 First, superoxide production is enhanced, as detected by EPR signals indrought stressed wheat and sunflower.185,186 Equally, H2O2 content increased about three-foldduring drought and low temperature.187-189 Enhanced production of ROS resulted in anincrease in lipid peroxidation, as documented by a more than 5-fold increase ofmalonaldehyde production in wheat.190 Second, the concentration of free transition ironincreased under drought stress,190,191 which stimulated production of hydroxyl radicals inthe presence of high concentrations of H2O2 via a Fenton reaction. Compared tosuperoxide and H2O2, hydroxyl radicals oxidize a variety of molecules at near diffusion-controlled rates. Finally, levels of non-enzymatic radical scavengers, such as ascorbate,carotenoids, flavonoids, sugar polyols, and proline,183 increase and may complementenzyme protection systems.

Excellent evidence for a protective effect of ROS scavenging systems has recently beenprovided by the overexpression of an enzyme with the combined activities of glutathioneS-transferase, GST, and glutathione peroxidase, GPX.192 By doubling the GST/GSX activity intransgenic tobacco, the seedlings and plants showed significantly faster growth than wildtype during chilling and salt stress episodes. The increased enzyme activities resulted inhigher amounts of oxidized glutathione (GSSG) in the stressed plants, indicating that theoxidized form could provide an increased sink for reducing power.

Another set of experiments shed light on the relationships between ROS and theaccumulation of polyols. When a bacterial gene (mtlD) encoding mannitol-1-phosphatedehydrogenase was modified so that the enzyme was expressed in chloroplasts, transgenictobacco contained approximately 100 mM mannitol in the plastids. Using transgenic plants,freshly prepared cells and a thylakoid in vitro system, the protective effect exerted bymannitol on photosynthesis characteristics could be shown.29, 30 The presence of mannitolresulted in increased resistance to oxidative stress generated by methylviologen, and cellsexhibited significantly higher CO2 fixation rates than controls during stress. Afterimpregnation of tissue and cells with dimethyl sulfoxide, a hydroxyl radical generator,mannitol-containing cells showed a lower rate of methane sulfinic acid production thanwild type, indicating that mannitol acted specifically as a hydroxyl radical scavenger. Itcould be shown that the primary damage was to enzymes of the Calvin cycle and not to


components of the light harvesting and electron transfer systems,30 a confirmation ofearlier reports.22 At present, the interpretation which we favor is that mannitol interfereswith either hydroxyl radical production or damage, but it is unknown whether theprotective mechanism is by exclusion of hydroxyl radicals from protein surfaces, a chemicalinteraction between mannitol and hydroxyl radicals, or by inhibiting or reducing theamount of hydroxyl radicals produced in the Fenton reaction.

Plant Ion Uptake and Compartmentation

H+-ATPases and Vacuolar PyrophosphatasePlasma membrane and vacuolar proton transporters play essential roles in plant

salinity stress tolerance by maintaining the transmembrane proton gradient that assurescontrol over ion fluxes and pH regulation (Fig. 3.3).101,193 Three proteins/protein complexesexist for this purpose: the plasma membrane (H+)-ATPase (P-ATPase) and two vacuolartransport systems, a (H+)-ATPase (V-ATPase) and a pyrophosphatase (PPiase).

The plant P-ATPase is represented by a gene family of more than 10, encodingproteins of ~100 kDa, with homology to the yeast PMAs.95,150 As the main proton pumpin the outer cell membrane it is essential for many physiological functions.194 Increasedactivity of the proton pump has been shown to accompany salt stress. Halophytic plantshave been shown to increase pump activity under salt stress conditions more drasticallythan glycophytes,56,195 but little is know about the regulatory circuits that lead to eitherincreased protein amount or activity during salt stress.

The V-ATPase, a multi-subunit complex homologous to organellar, yeast (VMA) andbacterial F0F1-ATPases, has already been shown to be important in plant salinity tolerance.Electrophysiological studies revealed increased activity of this ATPase when cells or tissuesfrom stressed plants were analyzed.196,197 Transcripts for several subunits of the V-ATPaseare upregulated following salt shock.198,199 In Mesembryanthemum, V-ATPase activityincreases several-fold following stress.200,201 In a Mesembryanthemum cell culture model ithas now been shown, based on immunological data, that the V-ATPase (and possibly theP-ATPase) activity does not increase due to more protein being present, but an unknownmechanism stimulates activity 2 to 3-fold.201 The response is specific for NaCl and couldnot be elicited by mannitol-induced osmotic stress.

PPiase genes and tonoplast-located PPiase proteins have been characterized in de-tail.94 Contrary to previous assumptions, the enzyme has now been authenticated asalso residing in the plasma membrane.202 Its function, if any, under salt stress conditionsis little known. A few reports have indicated that PPase activity declines under salt stress insome species,203,204 but increases in others.205

Potassium Transporters and ChannelsOne possible passage for sodium across the plasma membrane is through transport

systems for other monovalent cations. Among those the most significant is the uptakesystem for potassium, the most abundant cation in the cytosol, with important roles inplant nutrition, development and physiological regulation. Many studies have focused onidentifying components involved in K+-transport. Physiological observations indicating abiphasic uptake of K+ into roots206 gave rise to the assumption that two uptake entitiesshould be involved, a high-affinity system functioning at µM concentrations of externalK+ and a low-affinity system active in the mM range of potassium. Several plant K+

transporter and K+ channel genes have been isolated by functional complementation ofyeast mutants deficient in K+ uptake58,59,207 or by sequence homology with known K+

transporter or channel genes.208-210 Electrophysiological studies in heterologous


expression systems, such as Xenopus oocytes or yeast cells, indicated that some of themmay function at both affinity ranges.211

Inward-rectifying potassium channels function in the mM range, following theelectrochemical gradient at the plasma membrane and are categorized as low-affinitysystems.212,231 The AKT1- and KAT1-types of plant channels, similar to the Shakerchannels in animals, contain a pore-forming region conferring ion selectivity. In contrastto earlier assumptions, these channels are highly selective against Na+,213 and evidence islacking for specific regulation under salt stress. We think that the potassium channels playa minor role in salinity tolerance.

In contrast, K+ transporters which operate at low external potassium may mediateentry of sodium in saline soil. A high-affinity K+-transporter is known from yeast.214 Someof the cloned transporters take up potassium with dual-affinity.209,211 A high-affinity K+

transporter from wheat, HKT1, was indicated as a K+/Na+ symporter86 with high-affinitybinding sites for both K+ and Na+. Point mutations, which increased K+ selectivity overNa+, in one of the 12 transmembrane domains of HKT1 conferred increased salt toleranceof yeast. Another line of evidence for the involvement of high-affinity K+ uptake system insalt tolerance came from the study of salt-sensitive mutants. The sos1 mutant of Arabidopsisthaliana was characterized as hypersensitive to Na+ and Li+ and was unable to grow on low

Fig. 3.3. Transport proteins implicated in plant salinity stress tolerance. The schematic depictionof a plant cell includes the vacuole, chloroplast (cp), mitochondrion (mt) and cell wall (shaded).Transmembrane proton gradients established by proton-ATPases and pyrophosphatase areindicated (+/-). Under NaCl stress, Na+ and Cl- are sequestered to the vacuole, and K+ andosmolytes are present in high concentrations in the cytosol. Symbols for several membrane-locatedtransporters and channels are identified by the ion or proton transported and by the direction ofmovement. For organelles (mt and cp) no transporters have been characterized throughmolecular techniques. A Na+-ATPase, included in the plasma membrane is hypothetical, and aNa+/H+-antiporter in the plasma membrane has not been detected.


potassium.92 86Rb uptake experiments showed that sos1 was defective in high-affinitypotassium uptake, and it became deficient in potassium when treated with NaCl. Interestinglybut not surprisingly, expression of the wheat Hkt1 in sos1 mutant plants alleviated the salt-sensitive phenotype (Schroeder JI, Zhu J-K, personal communication). Further support isprovided by the expression characteristics of a rice homolog of wheat HKT1 in twovarieties that are distinguished by their salinity tolerance. The tolerant variety decreasedexpression of the root-specific HKT1 and efficiently excludes sodium, while a salt-sensitivevariety maintained high expression of the HKT1 in the presence of high NaCl.210,215

Irrespective of the indices pointing to the involvement of HKT1-type transporters, orhigh-affinity potassium uptake systems in general, in salt tolerance, there are other equallylikely scenarios. First, the presence of sodium is known to interfere with potassium uptake,as shown for several of the cloned transport proteins, and protective effects exerted byincreased potassium might be based on the nutritional value, and not on a sodiumexclusion mechanism. High sodium sensitivity, as for example shown by the sos1 mutant,might be due to growth interference when K+ uptake is reduced by the presence ofsodium. In this respect, the improved selectivity of K+ transport systems may increase salttolerance, while it is not involved in Na+ detoxification or osmotic adjustment. Othertransport systems, finally, might act in sodium uptake. How, for example, the calcium-regulated outward-rectifying K+-channel KCO1,216 or the regulation of other channelsand transporters, react under sodium stress conditions is unknown. It has been suggestedthat sodium might enter through outward-rectifying cation channels.217 Among the manypossibilities, evidence for significant sodium currents through a calcium transporter, LCT1,exists,218 and hexose and amino acid transporters may also let sodium pass.

Sodium Transport SystemsHow sodium enters plant cells, how it enters the plant circulatory system to be

selectively transported over long distances, and how it is partitioned to the vacuole is notknown in detail. Most information is available for the last step in this series: sodiumtransport from cytosol to vacuole is accomplished by a sodium/proton antiporter. Aprotein of approximately 170 kD219 is a candidate for this tonoplast-located antiporterbased on immunological studies and inhibition of the ameloride-regulated antiportactivity in the presence of the antibody. It will be important to characterize the protein indetail and to obtain the gene(s), because, when judged by protein size, the putative antiporterseems to be different from the proteins in bacteria, yeast and vertebrate organisms.Increased sodium/proton antiport activity during salt stress has been measured in severalmodel systems, tissues, cells and isolated vacuoles.96,220,221 The increase parallels anincrease in the V-ATPase activity.96,200

Our own data indicate that yet another pathway for sodium uptake may exist. Whenanalyzing the induction of myo-inositol synthesis in Mesembryanthemum, a surprisingdecline of the rate-limiting INPS (myo-inositol-1-phosphate synthase) enzyme in rootswas observed, but the concentration of myo-inositol remained constant in the roots.128,129

This is due to drastically enhanced transport of myo-inositol from the leaves through thephloem. In addition, myo-inositol is recycled to the leaves through the xylem and themyo-inositol amount in xylem vessels is correlated with sodium amounts.129 We have cloneda transcript with homology to vertebrate sodium/myo-inositol and yeast proton/myo-inositolsymporters222 and characterized its activity by complementation of a yeast mutantdefective in myo-inositol uptake.223 It seems possible that such a symporter is responsiblefor the excretion of sodium into the xylem, but it is equally possible that sodium/myo-inositol symport internalizes sodium from the apoplast of the root. The detection of such asymport mechanism is particularly attractive, considering that the passage of myo-inositol


through the plant circulatory system connects photosynthesis competence with sodiumuptake and transport to mesophyll cells of the leaf.

The Essentiality of CalciumIncreasing calcium improves salinity tolerance of crop plants. Physiological

experiments indicated that the effect is mediated through an increase of intracellular calcium,changes in vacuolar pH and activation of the vacuolar Na+/H+-antiporter.224, 225 The strictcontrol over calcium concentrations in the cytosol and calcium storage in a number oflocations (vacuole, mitochondria, endoplasmic reticulum) assign a crucial role to calciumin plant salinity stress responses.

Recently, an Arabidopsis mutant, sos3, with hypersensitivity to NaCl has beencharacterized. The mutant is different from other salt-sensitive mutants92 in that thephenotype can be masked by the external addition of calcium.226 This phenotype representsthe first mutant with an altered response to calcium in higher plants. The phenotypereveals the link between calcium and salinity stress tolerance, although the mechanismthrough which hypersensitivity and remediation by calcium are connected is not known.One attractive hypothesis is that a signaling system that responds to calcium spikes at lowcalcium concentrations—for example a homolog of the yeast calcineurin-type system—isdefective, and that at higher calcium concentrations a second sensing system can supportthe signal and elicit stress defense responses (Zhu JK, personal communication).

Metabolic Engineering of Glycophytic Plants for IncreasedSalt Tolerance

In increasing numbers, experiments are reported using transgenic plants for testingconcepts originating from the correlative evidence of physiological analyses. Table 3.1summarizes some of these reports. The concepts tested target four aspects of toleranceacquisition:

1. ROS scavenging,2. Compatible solutes and osmotic adjustment—carbohydrate biosynthesis and

synthesis of charged molecules,3. Ion balance—potassium uptake vs. sodium uptake, and4. The synthesis of specific, putatively protective proteins.A note of caution must be added with respect to the over-expression and accumulation

strategies that have been followed up to now. Too high an accumulation of metabolites, ortoo efficient scavenging of H2O2, for example, may not be desirable. When analyzingtransgenic tobacco plants that accumulated sorbitol to extremely high concentrations inthe cytosol, we observed stunted growth and the formation of necrotic lesions that reducedbiomass production, although the plants showed increased salinity and salt stress tolerance.227

The importance of radical oxygen scavenging for preventing oxidative stress in plantshas been demonstrated by genetic engineering of several enzymes into transgenicplants.179,180,228 Overexpression of superoxide dismutase (Cu/Zn-SOD and Mn-SOD),ascorbate peroxidase, catalase and glutathione reductase in transgenic plants has alreadybeen shown to lead to increased resistance to oxidative stress.181,229,230,232-238 The mostdramatic protective effect, up until now, was observed after enhancement of the glutathionecycle.192 In contrast, overexpression of an Fe-SOD in transgenic tobacco neither enhancedtolerance to chilling-induced photoinhibition in leaf discs nor increased tolerance to saltstress in whole plants,240 suggesting that isoforms of SOD may have different roles.

Noctor and Foyer20 provided a lucid assessment of the relatively marginal protectionthat has been observed in many transgenic plant studies, whether with respect to ROSscavenging or otherwise. It would certainly be premature to consider the protection


provided by the overexpression of SOD, ASX, or enzymes of the ascorbate/glutathionecycle as the final word. Protection has typically been observed in strictly controlledenvironments, and protective effects have often been marginal. We would like to provideone consideration as to why this is to be expected. In the case of ASX, at least six differentisoforms exist which are located in mitochondria, in chloroplasts (several, in differentsub-compartments/membranes), soluble in the cytosol, and in the cytoplasmicendomembrane system.241 A similarly complex distribution has been seen for SOD isoformswhich are found in the cytosol (Cu/Zn-SOD), mitochondria (Mn-SOD) and plastids(Fe-SOD and Cu/Zn-SOD). Transgenic modifications of single enzymes are likely to havea minimal effect because of the multitude of compartments that require protection.Irrespectively, these experiments have clearly shown that—in practically every study—the engineered expressed transgene elicited some protection. It is now necessary to adoptmulti-gene transfer strategies that alter several components of the stress tolerance system:

1. Targeting, for example, ROS scavenging enzymes to several compartments;2. Assembling gene constructs that target sodium exclusion and enhanced potassium

uptake;3. Generating transgenomes in which different pathways are satisfied, for example,

ion homeostasis, carbon allocation, and protein protection simultaneously;4. Generating transgenomes with strategies that take into account cell-, tissue-,

organ- and developmental specificity.The last point is particularly important, because little attention has typically been

paid to the “when,” “where,” and “how much” of transgene expression in the presentlyconcluded transgenic experiments. Significantly more attention needs to be directed tothe promoter elements that drive transgenes. Most attempts have targeted the metabolicengineering of carbon and nitrogen allocation: ectopic enzyme expression leading to thesynthesis of uncharged carbohydrates—mannitol, sorbitol, trehalose, fructan, andononitol—and to glycinebetaine and proline accumulation (Table 3.1). The underlyingmechanism is becoming apparent for some of these strategies, e.g., in the hydroxyl radicalscavenging function of mannitol.29,30 The mechanisms of protection underlying thesynthesis or presence of chaperones or specific LEA proteins remain to be determined.

Within a very short time, all genes that are essential for the salt tolerance phenotypeshown by some species and all genes that support damage avoidance in sensitive specieswill be available. The task remaining, however, is understanding in which metabolic andsignaling pathways the gene products function and in which developmental context stressprotection is necessary. This task will require new approaches. We consider two approaches:

1. Multi-gene transfer into model species—yeast, Arabidopsis, tobacco and rice areour suggestions; and

2. A focus on metabolic control analysis.The first strategy utilizes the transfer of all genes, controlled by appropriate promoter

elements, for one or several biochemical pathways to generate protection which can be ana-lyzed. Through the second approach, a biochemical description of flux in a multitude ofpathways, we will be able to gauge the cost of enzymes/pathways that enhance tolerance incomparison to the cost and benefits of resident pathways.

PerspectivesHigh salinity is a major factor responsible for the loss of crop biomass.242 Salinity

caused by irrigation affects many productive agricultural areas. The degeneration of stillproductive soils will become a more severe problem in the future. Development ofdrought- and salt-tolerant crops has been a major objective of plant breeding programsfor decades in order to maintain crop productivity in semiarid and saline lands. Although


several salt-tolerant varieties have been released, the overall progress of traditionalbreeding has been slow and has not been successful.13 The lack of success is mainly due tothe quantitative trait character of salinity tolerance which has to be reconciled withanother multigenic trait, high productivity, which is the ultimate goal of any breedingprogram. Marginal progress has equally been grounded in our poor understanding of themechanisms of salt tolerance, while the collected body of physiological data has focusedour attention more on details in a large variety of species and less on the principles.

This has changed over the last few years. Biochemical pathways that lead to theproduction of compatible solutes such as proline, glycine betaine, DMSP, or pinitol havebeen studied and most of the pathway genes have been characterized.6,7,9,14,15 We have thefirst glimpses of how the resulting metabolites from such pathways function in protection.Similarly, the principles of how radical oxygen species act and the principles, genes andproteins which deter radical damage have emerged. Membrane channels, transporters andpores are now available through which cells exert control over ion, carbohydrate, aminoacid or water fluxes.58,59,61,146,207,243 We owe most of this recent progress to the power ofthe yeast and Arabidopsis thaliana molecular genetic systems. Finding the genes whosedisruptions generate the various mutant phenotypes becomes rapidly easier as additionalmapping data and genomic DNA sequences from Arabidopsis are made available.12

Finally, plant stress perception, and inter- and intracellular signaling of salt stress hasbeen advanced greatly. Mutants in signal transduction pathways and components ofseveral signal transduction pathways have been found and are being characterized atpresent.244-249 Future studies can follow the blueprint of signaling components isolatedfrom yeast10,66,68,88,250 for finding and characterizing homologs of the essentialsignaling intermediates in plants. If we accept that a major objective of plant stressresearch is application, transgenic crops can be engineered not only for expression of novelbiochemical characters, but also for stress signal transduction that enhances the stressresponse inherent to all plants.

AcknowledgmentsBecause of space constraints a number of references could not be included, and we

apologize. We thank Ms. Pat Adams for help with the manuscript. Different projects have,off and on, been supported by the US National Science Foundation (Integrative PlantBiology and International Programs), Department of Energy (Biological Energy), andDepartment of Agriculture (NRI). Additional support has been provided by the ArizonaAgricultural Experiment Station, Japan Tobacco Inc., Rockefeller Foundation (New York)and New Energy Development Organization (Tokyo).

References1. Cushman JC, Bohnert HJ. Molecular biology of CAM. Plant Physiol 1997; 113:667-676.2. Shachar-Hill Y. Osmotic coupling and chloride transport in the limonium salt gland.

Proc Roy Soc London-Ser. B: Biol Sci 1990; 241:159-163.3. Flowers TJ, Troke PF, Yeo AR. The mechanism of salt tolerance in halophytes. Annu Rev

Plant Physiol 1977; 28:89-121.4. Adams P, Nelson DE, Yamada S et al. Growth and development of Mesembryanthemum

crystallinum (Aizoaceae). New Phytol 1998; 138:171-190.5. Langdale JA, Nelson T. Spatial regulation of photosynthetic development in C4 plants.

Trends Genet 1991; 7:191-196.6. McCue KF, Hanson AD. Drought and salt tolerance: Towards understanding and

application. Biotechnol 1990; 8:358-362.7. Delauney AJ, Verma DPS. Proline biosynthesis and osmoregulation in plants. Plant J

1993; 4:215-223.


8. Ishitani M, Majumder AL, Bornhouser A et al. Coordinate transcriptional induction ofmyo-inositol metabolism during environmental stress. Plant J 1996; 9:537-548.

9. Gage DA, Rhodes D, Nolte KD et al. A new route for synthesis of dimethyl-sulfoniopropionatein marine algae. Nature 1997; 387:891-894.

10. Niu X, Bressan RA, Hasegawa PM et al. Ion homeostasis in NaCl stress environments.Plant Physiol 1995; 109:735-742.

11. Nelson DE, Shen B, Bohnert HJ. Salinity tolerance-mechanisms, models, and the metabolicengineering of complex traits, In: Setlow JK, ed. Genetic Engineering, Principles andMethods. New York: Plenum Press, in press. 1998; 20:153-176.

12. Bevan M, Bancroft I, Bent E et al: Analysis of 1.9 Mb of contiguous sequence fromchromosome 4 of Arabidopsis thaliana. Nature 1998; 391:485-488.

13. Flowers TJ, Yeo AR. Breeding for salinity resistance in crop plants: Where next? Aust Jof Plant Physiol 1995; 22:875-884.

14. Bohnert HJ, Jensen RG. Metabolic engineering for increased salt tolerance-the next step.Aust J Plant Physiol 1996; 23:661-667.

15. Bohnert HJ, Jensen RG. Strategies for engineering water-stress tolerance in plants. TrendsBiotech 1996; 14:89-97.

16. Bartels D, Nelson DE. Approaches to improve stress tolerance using molecular genetics.Plant Cell Envir 1994; 17:659-667.

17. Serrano R. Salt tolerance in plants and microorganisms: Toxicity targets and defenseresponses. Intl Rev Cytol 1996; 165:1-52.

18. Jain RK, Selvaraj G. Molecular genetic improvement of salt tolerance in plants. BiotechAnnu Rev 1997; 3:245-267.

19. Zhu J-K, Hasegawa PM, Bressan RA. Molecular aspects of osmotic stress in plants. CritRev Plant Sci 1997; 16:253-277.

20. Noctor G, Foyer CH. Ascorbate and glutathione: Keeping active oxygen under control.Annu Rev Plant Physiol Plant Mol Biol 1998; 49:in press.

21. Levitt J. Responses of Plant to Environmental Stress Chilling, Freezing, and HighTemperature Stresses 2nd ed New York:Academic Press, Inc., 1980.

22. Kaiser WM. Reversible inhibition of the Calvin cycle and activation of oxidative pentosephosphate cycle in isolated chloroplast by hydrogen peroxide. Planta 1979; 145:377-382.

23. Bieleski RL. Sugar alcohols. In: Loewus FA, Tanner W, eds. Encyclopedia of Plant PhysiolVol 13A, Plant Carbohydrates I. Berlin:Springer Verlag. 1982:158-192.

24. Yancey PH, Clark ME, Hand SC et al. Living with water stress: Evolution of osmolytesystem. Science 1982; 217:1214-1222.

25. Ford CW. Accumulation of low molecular weight solutes in water stress tropical legumes.Phytochem 1984; 23:1007-1015.

26. Brown AD, Simpson JR. Water relations of sugar-tolerant yeasts: The role of intracellularpolyols. J Gen Microbiol 1972; 72:589-591.

27. Borowitzka LJ, Brown AD. 1974. The salt relations of marine and halophilic species ofthe unicellular green alga, Dunaliella. The role of glycerol as a compatible solute. ArchMicrobiol 1974; 96:37-52.

28. Le Rudulier D, Strom AR, Dandekar et al. Molecular biology of osmoregulation. Science1984; 224:1064-1068.

29. Shen B, Jensen RG, Bohnert HJ. Increased resistance to oxidative stress in transgenicplants by targeting mannitol biosynthesis to chloroplasts. Plant Physiol 1997;113:1177-1183.

30. Shen B, Jensen RG, Bohnert HJ. Mannitol protects against oxidation by hydroxyl radicals.Plant Physiol 1997; 115:527-532.

31. Schobert B, Tschesche H. Unusual solution properties of proline and its interaction withproteins. Biochem Biophys Acta 1978; 541:270-277.

32. Smirnoff N, Cumbes QJ. Hydroxyl radical scavenging activity of compatible solutes.Phytochem 1989; 28:1057-1060.

33. Halliwell B, Gutteridge MC. Role of free radicals and catalytic metal ions in humandisease: An overview. Meth Enzymol 1990; 186:1-85.


34. Orthen B, Popp M, Smirnoff N 1994. Hydroxyl radical scavenging properties of cyclitols.Proc Royal Soc Edinburgh 1994; 102B:269-272.

35. Pollard A, Wyn Jones RG. Enzyme activities in concentrated solutes of glycine betaineand other solutes. Planta 1979; 144:291-298.

36. Brown AD. Microbial Water stress Physiology, Principles and Perspectives. New York:John Wiley & Sons, 1990.

37. Solomon A, Beer S, Waisel Y et al. Effects of NaCl on the carboxylating activity of Rubiscofrom Tamarix jordanis in the presence and absence of proline-related compatible solutes.Plant Physiol 1994;. 90:198-204.

38. Back JF, Oakenfull D, Smith MB. Increased thermal stability of proteins in the presenceof sugars and polyols. Biochem 1979; 18:5191-5196.

39. Paleg LG, Douglas TJ, van Daal A et al. Proline, betaine and other organic solutes protectenzymes against heat inactivation. Aust J Plant Physiol 1981; 8:107-114.

40. Galinski EA. Compatible solutes of halophilic eubacteria: Molecular principles,water-solute interaction, stress protection. Experientia 1993; 49:487-496.

41. Papageorgiou G, Murata N. The unusually strong stabilizing effects of glycine betaine onthe structure and function of the oxygen-evolveing photosystem II complex. PhotosynRes 1995; 44:243-252.

42. Coughlan SJ, Heber U. The role of glycine betaine in the protection of spinach thylakoidsagainst freezing stress. Planta 1982; 156:62-69.

43. Jolivet Y, Larher F, Hamelin J. Osmoregulation in halophytic higher plants: The protectiveeffect of glycine betaine against the heat destabilization of membranes. Plant Sci Lett1982; 25:193-201.

44. Zhao Y, Aspinall D, Paleg LG. Protection of membrane integrity in Medicago sativa L. byglycine betaine against effects of freezing. J Plant Physiol 1992; 140:541-543.

45. Arakawa T, Timasheff SN. The stabilization of proteins by osmolytes. Biophys J 1985;47:411-414.

46. Schobert B. Is there an osmotic regulatory mechanism in algae and higher plants? J TheorBiol 1977; 68:17-26.

47. Dujon B. The yeast genome project: What did we learn? Trends Genet 1996; 12:263-270.48. Goffeau A, Barrell BG, Bussey H et al. Life with 6000 genes. Science 1996; 274:546-567.49. Wodicka L, Dong H, Mittman M et al. Genome-wide expression monitoring in Saccha-

romyces cerevisiae. Nature Biotech 1997; 15:1359-1367.50. Gadd GM, Chalmers K, Reed RH. The role of trehalose in dehydration resistance of

Saccharomyces cerevisiae. FEMS Microbiol Lett 1987; 48:249-254.51. Mackenzie KF, Singh KK, Brown AD. Water stress hypersensitivity of yeast: Protective

role of trehalose in Saccharomyces cerevisiae. J Gen Microbiol 1988; 134:1661-1666.52. Albertyn J, Hohmann S, Prior BR. Characterization of the osmotic-stress response in

Saccharomyces cerevisiae: Osmotic stress and glucose repression regulate glycerol-3-phosphate dehydrogenase independently. Curr. Genet 1994; 25:12-18.

53. Albertyn N, Hohmann S, Thevelein JM et al. GPD1, which encodes glycerol-3-phosphatedehydrogenase, is essential for growth under osmotic stress in Saccharomyces cerevisiae,and its expression is regulated by the high-osmolarity osmolarity glycerol responsepathway. Mol Cell Biol 1994; 14:4135-4144.

54. Ahmad I, Larher F, Stewart GR. Sorbitol, a compatible osmotic solute in Plantagomaritima. New Phytol 1979; 82:671-678.

55. Adams P, Thomas JC, Vernon DM et al. Distinct cellular and organismic responses tosalt stress. Plant Cell Physiol 1992; 33:1215-1223.

56. Niu X, Narasimhan ML, Salzman RA et al. NaCl regulation of plasma membrane H+-ATPasegene expression in a glycophyte and a halophyte. Plant Physiol 1995; 103:713-718.

57. van der Rest ME, Kamminga AH, Nakano A et al. The plasma membrane of Saccharomycescerevisiae:Structure, function, and biogenesis. Microbiol Rev 1995; 59:304-322.

58. Anderson JA, Huprikar SS, Kochian LV et al. Functional expression of a probableArabidopsis thaliana potassium channel in Saccharomyces cerevisiae. Proc Natl Acad SciUSA 1992; 89:3736-3740.


59. Sentenac H, Bonneaud N, Minet M et al. Cloning and expression in yeast of a plantpotassium ion transport system. Science 1992; 256:663-665.

60. Frommer WB, Ninnemann O. Heterologous expression of genes in bacterial, fungal,animal, and plant cells. Annu Rev Plant Physiol Plant Mol Biol 1995; 46:419-444.

61. Rentsch D, Hirner B, Schmelzer E et al. Salt stress-induced proline transporters and saltstress-repressed broad specificity amino acid permeases identified by suppression of ayeast amino acid permease-targeting mutant. Plant Cell 1996; 8:1437-1441.

62. Oliver SG. From gene to screen with yeast. Curr Opin Genet Dev 1997; 7(3):405-409.63. Yagi T, Tada K. Isolation and characterization of salt-sensitive mutants of a salt-tolerant

yeast Zygosaccharomyces rouxii. FEMS Microboil. Lett. 1988; 49:317-321.64. Ushio K, Ohtsuka H, Nakata Y. Intracellular Na+ and K+ contents of Zygosaccharomyces

rouxii mutants defective in salt tolerance. J Fermen Bioeng 1992; 73:11-15.65. Garciadeblas B, Rubio F, Quintero FJ et al. Differential expression of two genes encoding

isoforms of the ATPase involved in sodium efflux in Saccharomyces cerevisiae. Mol GenGenet 1993; 236:363

66. Brewster JL, De Valoir T, Dwyer ND et al. An osmosensing signal transduction pathwayin yeast. Science 1993; 259:1760-1763.

67. Maeda T, Wurgler-Murphy SM, Saito H. A two-component system that regulates anosmosensing MAP kinase cascade in yeast. Nature 1994; 369:242-245.

68. Maeda T, Takekawa M, Saito H. Activation of yeast PBS2 MAPKK by MAPKKKs or bybinding of an SH3-containing osmosensor. Science 1995; 269:554-558.

69. Norbeck J, Pahlman A, Akhtar N et al. Purification and characterization of two isoenzymesof DL-glycerol-3-phosphatase from Saccharomyces cerevisiae. J Biol Chem 1996;271:13875-13881.

70. Luyten K, Albertyn J, Skibbe WF et al. Fps1, a yeast member of the MIP family of channelproteins, is a facilitator for glycerol uptake and efflux and is inactive under osmotic stress.EMBO J 1995; 14:1360-1371.

71. Ansell R, Granath K, Hohmann S, Thevelein JM et al. The two isoenzymes for yeastNAD-dependent glycerol 3-phosphate dehydrogenase encoded by GPD1 and GPD2 havedistinct roles in osmoadaptation and redox regulation. EMBO J 1997; 16:2179-2187.

72. Shen B, Hohmann S, Jensen RG et al. Roles of sugar alcohols in osmotic stress adaptation-Replacement of glycerol in yeast by mannitol and sorbitol. To be submitted, 1998.

73. Blomberg A, Adler L. Physiology of osmotolerance in fungi. Adv Microbiol Physiol 1992;33:145-212.

74. Ota IM, Varshavsky A. A yeast protein similar to bacterial two-component regulators.Nature 1993; 262:566-569.

75. Posas F, Wurgler-Murphy SM, Maeda T et al. Yeast HOG1 MAP kinase cascade is regulatedby a multistep phosphorelay mechanism in the SLN1-YPD1-SSK1 two-componentosmosensor. Cell 1996; 86:865-875.

76. Schuller C, Brewster JL, Alexander MR et al. The HOG pathway controls osmoticregulation of transcription via the stress response element (STRE) of the Saccharomycescerevisiae CTT1 gene. EMBO J 1994; 13:4382-4389.

77. Hirayama T, Maeda T, Saito H et al. Cloning and characterization of seven cDNAsfor hyperosmolarity-responsive (HOR) genes of Saccharomyces cerevisiae. Mol GenGenet 1995; 249:127-138.

78. Varela JCS, Praekelt UM, Meacock PA et al. The Saccharomyces cerevisiae: Hsp12 gene isactivated by the high-osmolarity glycerol pathway and negatively regulated by proteinkinase A. Mol Cell Biol 1995; 15:6232

79. Davenport KR, Sohaskey M, Kamada Y et al. A second osmosensing signal transductionpathway in yeast: Hypotonic shock activates the PKC1 protein kinase-regulated cellintegrity pathway. J Biol Chem 1995; 270:30157-351

80. Shimizu J, Yoda K, Yamasaki M. The hypo-osmolarity-sensitive phenotype of theSaccharomyces cerevisiae hop2 mutant is due to a mutation in PCK1, which regulatesexpression of β-lucanase. Mol Gen Genet 1994; 242:641-648.


81. Andre B. An overview of membrane transport proteins in Saccharomyces cerevisiae. Yeast1995; 11:1575-1611.

82. Ko CH, Buckley AM, Gaber RF. TRK2 is required for low affinity K+ transport inSaccharomyces cerevisiae. Genetics 1990; 125:305-312.

83. Ko CH, Gaber RF. TRK1 and TRK2 encode structurally related K+ transporters inSaccharomyces cerevisiae. Mol Cell Biol 1991; 11:4266-4271.

84. Ketchum KA, Joiner WJ, Sellers AJ et al. A new family of outwardly rectifying potassiumchannel proteins with two pore domains in tandem. Nature 1995; 376:690-695.

85. Haro R, Banuelos M, Quintero FJ et al. Genetic basis of sodium exclusion and sodiumtolerance in yeast. Plant Physiol 1993; 89:868-874.

86. Rubio F, Gassman W, Schroeder JI. Sodium-driven potassium uptake by the plantpotassium transporter HKT1 and mutations conferring salt tolerance. Science 1995;270:1660-1663.

87. Gomez MJ, Luyten K, Ramos J. The capacity to transport potassium influences sodiumtolerance in Saccharomyces cerevisiae. FEMS Microbiol Lett 1996; 135:157-160.

88. Mendoza I, Rubio F, Rodriguez-Navarro A et al. The protein phosphatase calcineurin isessential for NaCl tolerance of Saccharomyces cerevisiae. J Biol Chem 1994; 269:8792-8796.

89. Gaxiola R, de Larrinoa IF, Villalba JM et al. A novel and conserved salt-induced proteinis an important determinant of salt tolerance in yeast. EMBO J 1992; 11:3157-3164.

90. Ferrando A, Kron SJ, Rios G et al. Regulation of cation transport in Saccharomycescerevisiae by the salt tolerance gene HAL3. Mol Cell Biol 1995; 15:5470-5481.

91. Watad AE, Reuveni M, Bressan RA et al. Enhanced net K+ uptake capacity of NaCl-adapted cells. Plant Physiol 1991; 95:1265-1269.

92. Wu S, Ding L, Zhu J. SOS1, a genetic locus essential for salt tolerance and potassiumacquisition. Plant Cell 1996; 8:617-627.

93. Nelson N. The vacuolar H+-ATPase: One of the most fundamental pumps in nature. JExp Biol 1992; 172:19-27.

94. Rea PA, Poole RJ. Vacuolar H+-translocating pyrophosphatase. Annu Rev Plant PhysiolPlant Mol Biol 1993; 44:157-180.

95. Sussman MR. Molecular analysis of proteins in the plant plasma membrane. Annu RevPlant Physiol Plant Mol Biol 1994; 45:211-234.

96. Barkla BJ, Pantoja O. Physiology of ion transport across the tonoplast of higher plants.Annu Rev Plant Physiol Plant Mol Biol 1996; 47:127-157.

97. Carmelo V, Bogaerts P, Sa-Correia I. Activity of plasma membrane H+-ATPase andexpression of PMA1 and PMA2 genes in Saccharomyces cerevisiae cells grown at optimaland low pH. Arch Microbiol 1996; 166:315-320.

98. Estrada E, Agostinis P, Vendenheede JR et al. Phosphorylation of yeast plasma membraneH+-ATPase by casein kinase I. J Biol Chem 1996; 271:32064-32072.

99. Braley R, Piper PW. The C-terminus of yeast plasma membrane H+-ATPase is essentialfor the regulation of this enzyme by heat shock protein HSP30, but not for stressactivastion. FEBS Lett 1997; 418:123-126.

100. Hemenway CS, Dolinski K, Cardenas ME et al. vph6 mutants of Saccharomyces cerevisiaerequire calcineurin for growth and are defective in vacuolar H+-ATPase assembly.Genetics 1995; 141:833-844.

101. Stevens TH, Forgac M. Structure, function and regulation of the vacuolar (H+)-ATPase.Annu Rev Cell Dev Biol 1997; 13:779-808.

102. Haro R, Garciadeblas B, Rodriguez-Navarro A. A novel P-type ATPase from yeastinvolved in sodium transport. FEBS Lett 1991; 291:189-191.

103. Wieland J, Nitsche AM, Strayle J et al. The PMR2 gene cluster encodes functionally distinctisoforms of a putative Na+ pump in the yeast plama membrane. EMBO J 1995;14:3870-3882.

104. Prior C, Potier S, Souciet J et al. Characterization of the NHA1 gene encoding aNa+/H+-antiporter of the yeast Saccharomyces cerevisiae. FEBS Lett 1996; 387:89-93.


105. Jia ZP, McMullough N, Martel R et al. Gene amplification at a locus encoding a putativeNa+/H+ antiport confer sodium and lithium tolerance in fission yeast. EMBO J 1992;11:1631-1640.

106. Watanabe Y, Miwa S, Tamai Y. Characterization of Na+/H(+)-antiporter gene closelyrelated to the salt-tolerance of yeast Zygosaccharomyces rouxii. Yeast 1995; 11:839-838.

107. Hahnenberger KM, Jia Z, Young PG. Functional expression of the Schizosaccharomycespombe Na+/H+ antiporter gene, sod2, in Saccharomyces cerevisiae. Proc Natl Acad Sci USA1996; 93:5031-5036.

108. Welihinda AA, Beavis AD, Trumbly RJ. Mutations in LIS1 (ERG6) gene confer increasedsodium and lithium uptake in Saccharomyces cerevisiae. Biochem Biophys Acta 1994;1193:107-117.

109. Latterich M, Watson MD. Isolation and characterization of osmosensitive vacuolarmutants of Saccharomyces cerevisiae. Mol Microbiol 1991; 5:2417-2426.

110. Nakamura T, Liu Y, Hirata D et al. Protein phosphatase type 2B (calcineurin)-mediated,FK506-sensitive regulation of intracellular ions in yeast is an important determinant foradaptation to high salt stress conditions. EMBO J 1993; 12:4064-4071.

111. Hirata D, Harada S, Namba H et al. Adaptation to high-salt stress in Saccharomycescerevisiae is regulated by Ca2+/calmodulin-dependent phosphoprotein phosphatase(calcineurin) and cAMP-dependent protein kinase. Mol Gen Genet 1995; 249:257-264.

112. Posas F, Camps M, Arino J. The PPZ protein phosphatases are important determinantsof salt tolerance in yeast cells. J Biol Chem 1995; 270:13036-13041.

113. Cyert MS, Kunisawa R, Kaim D et al. 1991. Yeast has homologs (CNA1 and CNA2 geneproducts) of mammalian calcineurin, a calmodulin-regulated phosphoprotein phosphatase.Proc Natl Acad Sci USA 1991; 88:7376-7380.

114. Marquez JA, Serrano R. Multiple transduction pathways regulate the sodium-extrusiongene PMR2/ENA1 during salt stress in yeast. FEBS Lett. 1996; 382:89-92.

115. Sugiura R, Toda T, Shuntoh H et al. Pmp1(+), a suppressor of calcineurin deficiency,encodes a novel MAP kinase phosphatase in fission yeast. EMBO J 1998; 17:140-148.

116. Rios G, Ferrando A, Serrano R. Mechanisms of salt tolerance conferred by overexpressionof the HAL1 gene in Saccharomyces cerevisiae. Yeast 1997; 13:515-528.

117. Luan S, Li W, Rusnak F, Assmann SM et al. Immunosuppressants implicate proteinphosphatase regulation of K+ channels in guard cells. Proc Natl Acad Sci USA 1993;90:2202-2206.

118. Liu J. FK506 and cyclosporin, molecular probes for studing intracellular signaltransduction. Immunol Today 1993; 14:290-295.

119. Luan S, Lane WS, Schreiber SL. pCyP B: A chloroplast-localized, heat shock-responsivecyclophilin from fava bean. Plant Cell. 1994; 6:885-892.

120. Lippuner V, Cyert MS, Gasser CS. Two classes of plant cDNA clones differentiallycomplement yeast calcineurin mutants and increase salt tolerance of wild type yeast. JBiol Chem 1996; 271:12859-12866.

121. Mendoza I, Quintero FJ, Bressan RA et al. Activated calcineurin confers high toleranceto ion stress and alters the budding pattern and cell morphology of yeast cells. J BiolChem 1996; 271:23061-23067.

122. Hanson AD, Rathinasabapathi B, Rivoal J et al. Osmoprotective compounds in theplumbaginaceae: A natural experiment in metabolic engineering of stress tolerance. ProcNatl Acad Sci USA 1994; 91:306-310.

123. Trossat C, Nolte KD, Hanson AD. Evidence that the pathway of dimethylsulfoniopropionatebiosynthesis begins in the cytosol and ends in the chloroplast. Plant Physiol 1996;111:965-973.

124. Bray EA. 1997. Plant responses to water deficit. Trends Plant Sci 1997; 2:48-54.125. Greenway H, Munns R. Mechanisms of salt tolerance in nonhalophytes. Annu Rev Plant

Physiol 1980; 31:149-190.126. Bohnert HJ, Nelson DE, Jensen RG. Adaptations to environmental stresses. Plant Cell

1995; 7:1099-1111.


127. Vernon DM, Bohnert HJ. A novel methyl transferase induced by osmotic stress in thefacultative halophyte Mesembryanthemum crystallinum. EMBO J 1992; 11:2077-85.

128. Nelson DE, Rammesmayer G, Bohnert HJ. Cell-specific myo-inositol metabolism andtransport in plant salinity tolerance. Plant Cell 1998; 10:753-764.

129. Nelson DE, Koukoumanos M, Bohnert HJ. Myo-inositol is necessary for efficient NaCluptake and transport in Mesembryanthemum crystallinum. Plant Physiol 1998; in press.

130. Kiyosue T, Yoshiba Y, Yamaguchi-Shinozaki K et al. A nuclear gene encoding mito-chondrial proline dehydrogenase, an enzyme involved in proline metabolism, isupregulated by proline but downregulated by dehydration in Arabidopsis. Plant Cell 1996;8:1323-1335.

131. Rhodes D, Hanson AD. Quaternary ammonium and tertiary sulfonium compounds inhigher plants. Annu Rev Plant Physiol Plant Mol Biol 1993; 44:357-384.

132. Ishitani M, Nakamura T, Han SY et al. Expression of the betaine aldehyde dehydrogenasegene in barley in response to osmotic stress and abscisic acid. Plant Mol Biol 1995;27:307-315.

133. Saneoka H, Nagasaka C, Hahn DT et al. Salt tolerance of glycine betaine-deficientand -containing maize lines. Plant Physiol 1995; 107:631-638.

134. Russell BL, Rathinasabapathi B, Hanson AD. Osmotic stress induces expression ofcholine monooxygenase in sugar beet and amaranth. Plant Physiol 1998; 116:859-865.

135. Kishor PBK, Hong Z, Miao G et al. Overexpresion of pyrroline-5-carboxylate synthetaseincreases proline production and confers osmotolerance in transgenic plant. Plant Physiol1995; 108:1387-1394.

136. Yoshiba Y, Kiyosue T, Katagiri T et al. Correlation between the induction of a gene forpyrroline-5-carboxylate synthetase and the accumulation of proline in Arabidopsis thalianaunder osmotic stress. J Plant 1995; 7:751-760.

137. Verbruggen N, Hua XJ, May M et al. Environmental and developmental signals modulateproline homeostasis: Evidence for a negative transcriptional regulators. Proc Natl AcadSci USA 1996; 93:8787-8791.

138. Sheveleva E, Chmara W, Bohnert HJ et al. Increased salt and drought tolerance byD-ononitol production in transgenic Nicotiana tabacum L. Plant Physiol 1997; 115:1211-1219.

139. Malin G, Lapidot A. Induction of synthesis of tetrahydropyrimidine derivatives inStreptomyces strains and their effect on E. coli in response to osmotic and heat stress. JBact 1996; 178:385-395.

140. Louis P, Galinski EA. Characterization of genes for the biosynthesis of the compatiblesolute ectoine from Marinococcus halophilus and osmoregulated expression in Escherichiacoli. Microbiol 1997; 143:1141-1149.

141. Holmström KO, Mäntyla E, Welin B et al. Drought tolerance in tobacco. Nature 1996;379:683-684

142. Chrispeels MJ, Agre P. 1994. Aquaporins: Water channel proteins of plant and animalcells. Trends Biochem Sci 1994; 19:421-425.

143. Maurel C. Aquaporins and water permeability of plant membranes. Annu Rev PlantPhysiol Plant Mol Biol 1997; 48:399-429.

144. Ishibashi K, Sasaki S, Fushimi K et al. Molecular cloning and expression of a member ofthe aquaporin family with permeability to glycerol and urea in addition to waterexpressed at the basolateral membrane of kidney collecting duct cells. Proc Natl Acad SciUSA 1994; 91:6269-6273.

145. Weaver CD, Shomer NH, Louis CF et al. Nodulin 26, a nodule-specific symbiosomeembrane protein from soybean, is an ion channel. J Biol Chem 1994; 269:17858-17862.

146. Weig A, Deswarte C, Chrispeels JJ. The major intrinsic protein family of Arabidopsishas 23 members form three distinct groups with functional aquaporins in each group.Plant Physiol 1997; 114:1347-1357.

147. Yang B, Verkman AS. Water and glycerol permeabilities of aquaporins 1-5 and MIPdetermined quantitatively by expression of epitope-tagged constructs in Xenopus oocytes.J Biol Chem 1997; 272:16140-16146.


148. Kaldenhoff R, Källing A, Meyers J et al. The blue light-responsive AthH2 gene ofArabidopsis thaliana is primarily expressed in expanding as well as in differentiatingcells and encodes a putative putative channel protein of the plasmalemma. Plant J1995; 7:87-95.

149. DeWitt ND, Sussman MR. Immunocytological localization of an epitope-tagged plasmamembrane proton pump (H+-ATPase) in phloem companion cells. Plant Cell 1995;7:2053-2067.

150. DeWitt ND, Hong B, Sussman MR et al. Targeting of two Arabidopsis H+-ATPase isoformsto the plasma membrane. Plant Physiol 1996; 112:833-844.

151. Yamada S, Katsuhara M, Kelly WB et al. A family of transcripts encoding water channelproteins: Tissue-specific expression in the common ice plant. Plant Cell 1995; 7:1129-1142.

152. Ludevid D, Hoefte H, Himmelblau E et al. The expression pattern of the tonoplastintrinsic protein γ-TIP in Arabidopsis thaliana is correlated with cell enlargement. PlantPhysiol 1992; 100:1633-1639.

153. Maurel C, Kado RT, Guern J et al. Phosphorylation regulates the water channel activityof the seed-specific aquaporin γ-TIP. EMBO J 1995; 14:3028-3035.

154. Ikeda S, Nasrallah JB, Dixit R et al. An aquaporin-like gene required for the Brassicaself-incompatibility response. Science 1997; 276:1564-1566.

155. Yamaguchi-Shinozaki K, Koizumi M, Urao S et al. Molecular cloning and characterizationof 9 cDNAs for genes that are responsive to desiccation in Arabidopsis thaliana: Sequenceanalysis of one cDNA clone that encodes a putative transmembrane channel protein.Plant Cell Physiol 1992; 33:217-224.

156. Opperman CH, Taylor CG, Conkling MA. Root-knot nematode-directed expression of aplant root-specific gene. Science 1994; 263:221-223.

157. Phillips AL, Huttly AK. Cloning of two gibberellin-regulated cDNAs from Arabidopsisthaliana by subtactive hybridization: Expression of the tonoplast water channel,gamma-TIP, is increased by GA3. Plant Mol Biol 1994; 24:603-615.

158. Kaldenhoff R, Kaling A, Richter G. Regulation of the Arabidopsis thaliana aquaporin geneAthH2 (PIP1b). J Photochem Photobiol B 1996; 36:351-354.

159. Yamamoto YT, Cheng CL, Conkling MA. Root-specific genes from tobacco andArabidopsis homologous to an evolutionarily conserved gene family of membranechannel proteins. Nucl Acids Res 1990; 18:7449-7457.

160. Yamada S, Nelson DE, Ley E et al. The expression of an aquaporin promoter fromMesembryanthemum crystallinum in tobacco. Plant Cell Physiol 1997; 38:1326-1332.

161. Kirch H-H, Michalowski CB, Bohnert HJ. Anti-MIP antibodies recognize distinct cell typesin roots and leaves from Mesembryanthemum crystallinum. Manuscript in preparation, 1998.

162. Verkman AS. Water channels. Austin:RG Landes Company, 1993.163. King LS, Agre P. Pathophysiology of the aquaporin water channels. Annu Rev Physiol

1996; 58:619-648.164. Nielsen S, Chou C-L, Marples D et al. Vasoprssin increases water permeability of kidney

collecting duct by inducing translocation of aquaporin-CD water channels to plasmamembrane. Proc Natl Acad Sci USA 1995; 92:1013-1017.

165. Robinson DG, Sieber H, Kammerloher W et al. PIPq aquaproins are concentrated inplasmalemmasomes of Arabidopsis thaliana mesophyll. Plant Physiol 1996; 111:645-649.

166. Johansson I, Larsson C, Ek B et al. The major integral proteins of spinach leaf plasmamembranes are putative aquaporins and are phosphorylated in response to Ca2+ andapoplastic water potential. Plant Cell 1996; 8:1181-1191.

167. Buxton GV, Greenstock CL, Helman WP et al. Critical review of rate constants for reactionsof hydrated electrons, hydrogen atoms and hydroxyl radicals in aqueous solution. J PhysChem Ref Data 1988; 17:512-579.

168. Asada K. Production and action of active oxygen species in photosynthetic tissues. In:Foyer CH and Mullineaux PM, eds. Causes of photooxidative stress and amelioration ofdefense systems in plants. London: CRC Press, 1994; 77-104.

169. Tenhaken R, Levine A, Brisson LF et al. Function of the oxidative burst in hypersensitivedisease resistance. Proc Natl Acad Sci USA 1995; 92:4158-4163.


170. Lamb C, Dixon RA:The oxidative burst in plant disease resistance. Annu Rev Plant PhysiolPlant Mol Biol 1997; 48:251-275.

171. Mehler AH, Brown AH. Studies on reactions of illuminated chloroplasts. III. Simultaneousphotoproduction and consumption of oxygen studied with oxygen isotopes. Arch BiochemBiophys 1952; 38:365-370.

172. Robinson JM. Does O2 photoreduction occur within chloroplasts in vivo? Plant Physiol1988; 72:666-680.

173. Asada K, Takahashi M. Production and scavenging of active oxygen species inphotosynthetic tissues. In: Kyle DJ, Osmond CB, Arntzen CJ, eds. Photoinhibition.Amsterdam: Elsevier Publ, 1994:227-287.

174. Hodgson RA, Raison JK. Superoxide production by thylakoids during chilling and itsimplication in the susceptibility of plants to chilling-induced photoinhibition. Planta 1991;183:222-228.

175. Kim CS, Jung J. The susceptibility of mung bean chloroplasts to photoinhibition isincreased by an excess supply of iron to plants: A photobiological aspect of iron toxicityin plant leaves. Photochem Photobiol 1993; 58:120-126.

176. Elstner EF. Metabolism of activated oxygen species. In: Davies DD, ed. Biochemistry ofMetabolism, Vol 11. San Diego: Academic Press Inc, 1987:253-315.

177. Fryer MJ. The antioxidant effects of thylakoid vitamin E (α-tocopheral). Plant Cell Eviron1992; 15:381-392.

178. Dalton DA. Antioxidant defense of plants and fungi. In: Ahmad S, ed. Oxidant-inducedStress and Antioxidant Defenses in Biology. New York: Chapman & Hall, 1995:298-355.

179. Foyer CH, Descourvieres P, Kunert KJ. Protection against oxygen radicals: An importantdefense mechanism studied in transgenic plants. Plant, Cell Environ 1994; 17:507-523.

180. Foyer CH, Lelandais M, Kunert KJ. Photooxidative stress in plants. Physiol Plant 1994;92:696-717.

181. Willekens H, Chamnongpol S, Davey M et al. Catalase is a sink for H2O2 and isindispensable for stress defence in C3 plants. EMBO J 1997; 16:4806-4816.

182. Osmond CB, Grace SC. Perspectives on photoinhibition and photorespiration in the field:Quintessential inefficiencies of the light and dark reactions of photosynthesis? J Exp Bot1995; 46:1351-1362.

183. Smirnoff N. The role of active oxygen in the response of plants to water deficit anddesiccation. New Phytol 1993; 125:27-58.

184. Wise RR. Chilling-enhanced photooxidation: The production, action and study ofreactive oxygen species produced during chilling in the light. Photosyn Res 1995;45:79-97.

185. Price AH, Atherton N, Handry GAF. Plants under drought-stress generate activated oxygen.Free Radical Research Comm 1989; 8:61-66.

186. Quartacci MF, Navari-Izzo F. Water stress and free radical mediated changes insunflower seedlings. J Plant Physiol 1992; 139:621-625.

187. Chowdhury SR, Choudhuri MA. Hydrogen peroxide metabolism as an index of waterstress tolerance in jute. Plant Physiol 1985; 65:503-507.

188. Prasad TK, Anderson MD, Martin BA et al. Evidence for chilling-induced oxidative stressin maize seedling and a regulatory role for hydrogen peroxide. Plant Cell 1994; 6:65-74.

189. Prasad TK. Mechanisms of chilling-induced oxidative stress injury and tolerance indeveloping maize seedlings: Changes in antioxidant system, oxidation of proteins andlipids, and protease activities. J Plant 1996; 10:1017-1026.

190. Price AH, Handry GAF. Iron-catalysed oxygen radical formation and its possiblecontribution to drought damage in nine native grasses and three cereals. Plant CellEnviron 1991; 14:477-484.

191. Moran JF, Becana M, Iturbe-Ormaetxe I et al. Drought induces oxidative stress in peaplants. Planta 1994; 194:346-352.

192. Roxas VP, Smith RK Jr, Allen ER et al. Overexpression of glutathione S-transferase/glutathioneperoxidase enhances the growth of transgenic tobacco seedlings during stress. NatureBiotech. 1997; 15:988-991.


193. Guern J, Mathieu Y, Kurkdjian A et al. Regulation of vacuolar pH in plant cells. PlantPhysiol 1989; 89:27-36.

194. Michelet B, Boutry M. The plasma membrane H+-ATPase. Plant Physiol 1995; 108:1-6.195. Weiss M, Pick U. Primary structure and effect of pH on the expression of the plasma

membrane H+-ATPase from Dunaliella acidophila and Dunaliella salina. Plant Physiol1996, 112:1693-1702.

196. Reuveni M, Bennett AB, Bressan RA et al. Enhanced H+ transport capacity and ATPhydrolysis activity of the tonoplast H+-ATPase after NaCl adaptation. Plant Physiol 1990;94:524-530.

197. Ayala F, O’Leary JW, Schumaker KS. Increased vacuolar and plasma membrane H+-ATPaseactivities in Salicornia bigelovii Torr. in response to NaCl. J Exp Bot 1996; 47:25-32.

198. Loew R, Rockel B, Kirsch M et al. Early salt stress effects on the differential expressionof vacuolar H+-ATPase genes in roots and leaves of Mesembryanthemum crystallinum.Plant Physiol 1996; 110:259-265.

199. Tsiantis M, Bartholomew DM, Smith JAC. Salt regulation of transcript levels for thec-subunit of a leaf vacuolar H+-ATPase in the halophyte Mesembryanthemum crystallinum.Plant J 1996; 9:729-736.

200. Barkla BJ, Zingarelli L, Blumwald E, Smith JAC. Tonoplast Na+/H+ antiport activity andits energization by the vacuolar H+-ATPase in the halophytic plant Mesembryanthemumcrystallinum. Plant Physiol 1995; 109:549-556.

201. Vera-Estrella R, Barkla BJ, Bohnert HJ et al. Salt stress in Mesembryanthemum crystallinumsuspension cells activates adaptive mechanisms identical to those observed in the wholeplant. Planta, in press.

202. Robinson DG. Pyrophosphatase is not (only) a vacuolar marker. Trend Plant Sci 1996;1:330.

203. Bremberger C, Luettge U. Dynamics of tonoplast proton pumps and other tonoplastproteins of Mesembryanthemum crystallinum L. during the induction of crassulacean acidmetabolism. Plant 1992; 188:575-580.

204. Matsumoto H, Chung GC. Increase in proton-transport activity of tonoplast vesicles asan adaptive response of barley roots to NaCl stress. Plant Cell Physiol 1988; 29:1133-1140.

205. Zingarelli L, Anzani P, Lado P. Enhanced K+-stimulated pyrophosphatase activity inNaCl-adapted cells of Acer pseudoplatanus. Physiol Plant 1994; 91:510-516. 141.

206. Epstein E (1966) Dual pattern of ion absorption by plant cells and by plants. Nature212:1324-1327.

207. Schachtman DP, Schroeder JI. Structure and transport mechanism of a high-affinitypotassium uptake transporter from higher plants. Nature 1994; 370:655-658.

208. Santa-Maria GE, Rubio F, Dubcovsky J et al. The HAK1 gene of barley is a member of alarge gene family and encodes a high-affinity potassium transporter. Plant Cell 1997;9:2281-2289.

209. Kim EJ, Kwak JM, Uozumi N et al. AKUP1: An Arabidopsis gene encoding high-affinitypotassium transport activity. Plant Cell 1998; 10:51-62.

210. Golldack D, Su H, Bennett J et al. Differential expression of HKT1-type potassiumtransporters in salt-sensitive and salt-tolerant rice lines. Manuscript in preparation, 1998.

211. Fu H-H, Luan S. AtKUP1: A dual affinity K+ transporter from Arabidopsis. Plant Cell1998; 10:63-73.

212. Maathuis FJM, Verlin D, Smith FA et al. The physiological relevance of Na+-coupledK+-transport. Plant Physiol 1996; 112:1609-1616.

213. Bertl A, Anderson JA, Slayman CL et al. Use of Saccharomyces cerevisiae for patch-clampanalysis of heterologous membrane proteins: Characterization of Kat1, an inward-rectifying K+ channel from Arabidopsis thaliana, and comparison with endogeneous yeastchannels and carriers. Proc Natl Acad Sci USA 1995; 92:2701-2705

214. Gaber RF, Styles CA, Fink GR. TRK1 encodes a plasma membrane protein required forhigh-affinity potassium transport in Saccharomyces cerevisiae. Mol Cell Biol 1988;8:2848-2859.


215. Golldack D, Kamasani U, Quigley F et al. Salt stress-dependent expression of a HKT1-typehigh affinity potassium transporter in rice. Plant Physiol 1997; S114:118.

216. Czempinski K, Zimmermann S, Ehrhardt T et al. New structure and function in plantK+-channels: KCO1, an outward rectifier with a steep Ca2+ dependency. EMBO J 1997;16:2565-2575.

217. Schachtman DP, Tyerman SD, Terry BR. The K+/Na+ selectivity of a cation channel inthe plasma membrane of root cells does not differ in salt-tolerant and salt-sensitive wheatspecies. Plant Physiol 1991; 97:598-605.

218. Schachtman DP, Kumar R, Schroeder JI et al. Molecular and functional characterizationof a novel low-affinity cation transporter (LCT1) in higher plants. Proc Natl Acad SciUSA 1997; 94:11079-11084.

219. Barkla BJ, Apse MP, Manolson MF et al. The plant vacuolar Na+/H+ antiport. Symp SocExp Biol 1994; 48:141-153.

220. Blumwald E, Poole RJ. Salt tolerance in suspension cultures of sugar beet: Induction ofNa+/H+ antiport activity at the tonoplast by growth in salt. Plant Physiol 1987; 83:884-887.

221. Garbarino J, DuPont FM. NaCl induces Na+/H+ antiport in tonoplast vesicles frombarley roots. Plant Physiol 1988; 86:231-236.

222. Cammarata PR, Xu GT, Huang L et al. Inducible expression of Na+/myo-inositolcotransporter mRNA in anterior epithelium of bovine lens: Affiliation with hypertonicityand cell proliferation. Exp Eye Res 1997; 64:745-757.

223. Nelson DE, Bohnert HJ. Characterization of the sodium/myo-inositol symporter fromMesembryanthemum crystallinum. Manuscript in preparation, 1998.

224. Colmer TD, Fan TWM, Higashi RM et al. Interactions of Ca2+ and NaCl stress on theion relations and intracellular pH of Sorghum bicolor roots tips: An in vivo 31P-NMRstudy. J Exp Bot 1994; 45:1037-1044.

225. Martinez V, Lauchli A. Effects of calcium on the salt-stress response of barley roots asobserved by in vivo phosphorus-31 nuclear magnetic resonance and in vitro analysis.Plata 1993; 190:519-524.

226. Liu J, Zhu JK. An Arabidopsis mutant that requires increased calcium for potassiumnutrition and salt tolerance. Proc Natl Acad Sci USA 1997; 94:14960-14964.

227. Sheveleva E, Marquez S, Zegeer A et al. Sorbitol dehydrogenase expression in transgenictobacco: High sorbitol accumulation leads to necrotic lesions in immature leaves. PlantPhysiol 1998; 117:831-839.

228. Allen RD. Dissection of oxidative stress tolerance using transgenic plants. Plant Physiol1995; 107:1049-1054.

229. Aono M, Kubo A, Saji H et al. Resistance to active oxygen toxicity of transgenicNicotiana tabacum that expresses the gene for glutathione reductase from E. coli. PlantCell Physiol 1991; 32:691-697.

230. Aono M, Kubo A, Saji H et al. Enhanced tolerance to photooxidative stress of transgenicNicotiana tabacum with high chloroplastic glutathione reductase activity. Plant Cell Physiol1993; 34:129-135.

231. Maathuis FJM, Ichida AM, Sanders D et al. Roles of higher plant K+ channels. PlantPhysiol 1997; 114:1141-1149.

232. Bowler C, Slooten L, Vandenbranden S et al. Manganese superoxide dismutase canreduce cellular damage mediated by oxygen radicals in transgenic plants. EMBO J 1991;10:1723-1732.

233. Bowler C, Van Montagu M, Inze D. Superoxide dismutase and stress tolerance. AnnuRev Plant Phys Plant Mol Biol 1992; 43:83-116.

234. Gupta AS, Heinen JL, Holaday AS et al. Increased resistance to oxidative stress intransgenic plants that overexpress chloroplastic Cu/Zn superoxide dismutase. Proc NatlAcad Sci USA 1993; 90:1629-1633.

235. Van Camp W, Wilekens H, Bowler WH et al. Elevated levels of superoxide dismutaseprotect transgenic plants against ozone damage. Biotechnol 1994;.12:165-168.

236. McKersie BD, Chen Y, de Beus M et al. Superoxide dismutase enhances tolerance offreezing stress in transgenic alfalfa (Medicago sativa L.). Plant Physiol 1993; 103:1155-1163.


237. Foyer CH, Souriau N, Perret S et al. Overexpression of glutathione reductase but notglytathione synthetase leads to increase in antioxidant capacity and resistance tophotoinhibition in poplar trees. Plant Physiol 1995; 109:1047-1057.

238. McKersie BD, Bowley SR, Harjanto E et al. Water-deficit tolerance and field performanceof transgenic alfalfa overexpressing superoxide dismutase. Plant Physiol 1996;111:1177-1181.

239. Pitcher LH, Zilinskas BA. Overexpression of copper/zinc superoxide dismutase in thecytosol of transgenic tobacco confers partial resistance to ozone-induced foliar necrosis.Plant Physiol 1996; 110:583-588.

240. Van Camp W, Capiau K, Montagu M et al. Enhancement of oxidative stress tolerance intransgenic tobacco plants overproducing Fe-superoxide dismutase in chloroplasts. PlantPhysiol 1996; 112:1703-1714.

241. Jespersen HM, Kjaersgard IV, Ostergard L et al. From sequence analysis of three novelascorbate peroxidases from Arabidopsis thaliana to structure, function and evolution ofseven types of ascorbate peroxidase. Biochem J 1997, 326:305-310.

242. Boyer JS. Plant productivity and environment. Science 1982; 218:443-448.243. Sauer N, Stolz J. SUC1 and SUC2: Two sucrose transporters from Arabidopsis thaliana;

expression and characrerization in baker’s yeast and identification of the histidine-taggedprotein. Plant J 1994; 6:67-77.

244. Nishihama R, Banno H, Shibata W et al. Plant homologues of components of MAPK(mitogen-activated protein kinase) signal pathways in yeast and animal cells. Plant CellPhysiol 1995; 36:749-757.

245. Kakimoto T. CKI1, a histidine kinase homolog implicated in cytokinin signal transduction.Science 1996; 274:982-985.

246. Ishitani M, Xiong L, Stevenson B et al. Genetic analysis of osmotic and cold stress signaltransduction in Arabidopsis: Interactions and convergence of abscisic acid-dependent andabscisic acid-independent pathways. Plant Cell 1997; 9:1935-1949.

247. Hirt H: Multiple roles of MAP kinases in plant signal transduction. Trends Plant Sci1997; 2:11-15.

248. Mizoguchi T, Irie K, Harashida N et al. A gene encoding a mitogen-activated proteinkinase kinase kinase is induced simultaneously with genes for a mitogen-activatedprotein kinase and an S6 ribosomal protein kinase by touch, cold, and water stress inArabidopsis thaliana. Proc Natl Acad Sci USA 1996; 93:765-769.

249. Mizoguchi T, Ichimura K, Shinozaki K. Environmental stress response in plants: Therole of mitogen-activated protein kinases. Trends in Biotech 1997; 15:15-19.

250. Shinozaki K, Yamaguchi-Shinozaki K. Gene expression and signal transduction in water-stress response. Plant Physiol 1997; 115:327-334.

251. Tarczynski MC, Jensen RG, Bohnert HJ. Expression of a bacterial mtlD gene in transgenictobacco leads to production and accumulation of mannitol. Proc Natl Acad Sci USA1992; 89:2600-2604.

252. Tarczynski MC, Jensen RG, Bohnert HJ. Stress protection of transgenic tobacco byproduction of the osmolyte, mannitol. Science 1993; 259:508-510.

253. Thomas JC, Sepahi M, Arendall B et al. Enhancement of seed germination in high salinity byengineering mannitol expression in Arabidopsis thaliana. Plant Cell Environ 1995;18:801-806.

254. Pilon-Smits EAH, Ebskamp MJM, Paul MJ et al. Improved performance of transgenicfructan-accumulating tobacco under drought stress. Plant Physiol 1995; 107:125-130.

255. Hayashi H, Alia, Mustardy L et al. Transformation of Arabidopsis thaliana with the codAgene for choline oxidase; accumulation of glycinebetaine and enhanced tolerance to saltand cold stress. Plant J 1997; 12:133-142.

256. Xu D, Duan X, Wang B et al. Expression of a late embryogenesis abundant protein gene,HVA1, from barley confers tolerance to water deficit and salt stress in transgenic rice.Plant Physiol 1996; 110:249-257.

257. Jones JT, Mullet JE. Developmental expression of a turgor-responsive gene that encodesan intrinsic membrane protein. Plant Mol Biol 1995; 28:983-996.

CHAPTER 4


Plant Cold ToleranceMichael F. Thomashow and John Browse

Plants vary greatly in their responses to cold temperatures. At one extreme are many plantsfrom tropical and subtropical regions which suffer injury when exposed to low nonfreezing

temperatures. These include economically important plants such as cotton, soybean, maize,rice, and many tropical and subtropical fruits. Such chilling-sensitive plants undergo sharpreductions in growth rate and development at temperatures between 0˚ and 12˚C.1,2 Thephysical and physiological changes in chilling-sensitive plants that are induced by exposure tolow temperatures, together with the subsequent expression of stress symptoms, are termedchilling injury. The symptoms that are associated with chilling injury include reduced orretarded germination and seedling emergence, wilting and chlorosis of leaf tissue, electrolyteleakage and tissue necrosis.

In sharp contrast to plants of tropical origin, those from temperate regions are notonly chilling-tolerant, but many are able to survive freezing. Herbaceous plants fromtemperate regions can survive freezing temperatures ranging from -5˚ to -30˚C, depending onthe species, while trees from boreal forests routinely survive winter temperatures below -30˚C.Significantly, the maximum freezing tolerance of these plants is not constitutive, but isinduced in response to low nonfreezing temperatures (below ~10˚C), a phenomenon knownas “cold acclimation.” For instance, rye plants grown at normal warm temperatures are killedby freezing below about -5˚C, but after cold acclimation can survive freezing temperaturesdown to about -30˚C.

What accounts for the differences in cold tolerance among plant species? Why arecucumber and rice plants injured at chilling temperatures while cold-acclimated cabbageand wheat survive freezing below -15˚C? The answers to these questions are of basic scientificinterest and have potential practical applications. Cold temperatures limit the geographicallocations where crop and horticultural plant species can be grown and periodically causesignificant losses in plant productivity. Greater knowledge of the molecular basis of chillingand freezing tolerance could potentially lead to the development of new strategies toimprove plant cold tolerance, resulting in increased plant productivity and expandedareas of agricultural production. Here we summarize the current understanding of themolecular basis for chilling and freezing injury and discuss recent advances in theidentification of genes involved in cold tolerance.

Chilling Tolerance

Role of Membranes in Chilling InjuryThe majority of chilling-sensitive plants share a similar threshold for the onset of

low-temperature damage and exhibit a common assembly of symptoms. These observations

62 Molecular Responses to Cold, Drought, Heat and Salt Stress in Higher Plants

have been interpreted by many investigators as indicating that there is a single primary lesion,or trigger, that initiates cell damage at some critical temperature and leads to a cascade ofsecondary events that are the more readily appreciated consequences of chilling damage.2

Several primary lesions have been proposed, but the most widely studied hypothesesinvolve temperature-dependent changes in membrane lipid structure.3,4 Early suggestionsenvisioned a mechanism in which the lipids in membranes underwent an overall phasetransition from the liquid crystalline (Lα) state to the gel (Lβ) state.1,5 According to thisproposal, the transition from liquid crystalline phase to gel phase would result in alterations inthe metabolism of chilled cells and lead to injury and death of the chilling-sensitive plants. Itwas quickly recognized that such a mechanism was an oversimplification6 but it was morethan ten years before a more sophisticated version of the membrane hypothesis wasarticulated. Raison and Wright7 observed that small additions of disaturated phospholipids topreparations of wheat polar lipids could produce entropy changes during differential scanningcalorimetry that were quantitatively similar to those observed for polar lipid extracts fromchilling-sensitive mung bean plants. These experiments suggested that only a portion ofthe lipids (4 to 7%) was actually undergoing a phase change in the 0˚ to 12˚C temperaturerange.

Meanwhile, Murata and coworkers demonstrated a strong correlation across differentplant species between the degree of chilling sensitivity and the proportion of disaturatedphosphatidylglycerol (PG); molecules that contain only 16:0, 18:0 and 16:1-trans fattyacids.8 Chloroplast PG is invariably synthesized with 16:0 at the sn-2 position of the glycerolbackbone. Although this 16:0 may be converted to ∆3-16:1-trans, the geometry of thistrans-unsaturated fatty acid is very similar to that of saturated fatty acids. For this reason, thelevel of disaturated PG depends on the extent to which the glycerol-3-phosphateacyltransferase specifically selects 18:1-ACP to the exclusion of 16:0-ACP and 18:0-ACP thatare also available as substrates in the chloroplast stroma.4 Invoking disaturated molecularspecies of PG as the cause of chilling sensitivity was attractive because, in contrast toproposals based on less precise concepts of lipid unsaturation, it provided a mechanismunderpinned by a firm biophysical explanation. Thus, preparations of PG purified fromthree chilling-sensitive plants were observed to enter the Lα to Lβ phase transition at 29˚ to33˚C, whereas PG from chilling resistant plants did not enter the transition until thetemperature was below 15˚C.9 More recently, the molecular-species distribution of PG intobacco and Arabidopsis plants has been altered by molecular genetic techniques.10,11 Murataet al10 transformed tobacco plants with gene constructs encoding glycerol-3-phosphateacyltransferase from either squash or Arabidopsis. Transgenic plants containing the squashgene contained elevated levels of disaturated PG (76% of total PG) compared with controls(36%) and showed more damage after chilling. Conversely, transgenic plants expressingthe Arabidopsis gene contained 28% disaturated PG and showed less chilling damage thancontrol tobacco plants. One of the measures of chilling injury in this study was the extentof photoinhibition of photosynthesis. Subsequent studies by Moon et al12 revealed thatthere is no difference between the rate at which transgenic and wild type plants undergochilling-induced photoinhibition. Rather, the principal effect of the variation in the amountof disaturated PG seems to be on the rate at which damaged photosystems can be repaired.The main target for photoinhibition is thought to be the D1 polypeptide of the photosystem IIreaction center.13 When this protein is damaged, presumably by side products from thephotochemical reactions,14 newly synthesized D1 protein is inserted into the photosystemII complex to restore photochemical activity. Thus, Moon et al12 hypothesize that thealtered amount of disaturated PG has an effect on the rate at which damaged D1 protein isremoved from the photosystem II complex and replaced by synthesis and insertion of newprotein. An important unanswered question is whether the effect of disaturated-PG

63Plant Cold Tolerance

content on D1 turnover extends to other chloroplast membrane proteins. Conceivably, theD1 protein is simply an efficient reporter of a general defect in the assembly or removal ofmembrane proteins.

Transformation of tobacco plants with a suitably modified version of the des9 genefrom Anacystis nidulans, which encodes a broad-specificity glycerolipid ∆9 desaturase substantiallyreduced the level of saturated fatty acids in PG and other membrane lipids.15 Again, the transgenicplants were more tolerant to chilling treatments than control plants. Interestingly, modestincreases in overall membrane unsaturation also appear to reduce chilling damage intobacco. Kodama et al16 used the Arabidopsis FAD7 gene in transgenic tobacco to producea modest increase (~10%) in the conversion of dienoic to trienoic fatty acids in membranelipids with no apparent change in the level of disaturated PG. Compared with controlplants, the transgenic tobacco exhibited significantly higher leaf expansion rates followinga 7 day exposure to 1˚C. This result is somewhat surprising since the change in lipidcomposition would not be expected to influence the tendency of membrane lipids toundergo a phase transition.

In a similar approach, Wolter et al11 restructured the E. coli plsB gene, which encodesa membrane-bound glycerol-3-phosphate acyltransferase, so that the peptide was directedinto the chloroplasts of transgenic Arabidopsis that expressed the gene. The bacterialacyltransferase utilizes both 16:0- and 18:1-ACPs as substrates. The resulting transgenicArabidopsis plants contained 48-54% disaturated PG and were damaged by treatment at4˚C for 7 days. These findings suggest that disaturated PG species can induce low-temperaturesensitivity in a chilling-tolerant plant such as Arabidopsis, although it is not formallypossible to rule out the possibility that accumulation of the plsB protein might havecontributed to the phenotype observed.11

Notwithstanding this accumulated body of evidence, studies on the Arabidopsis fab1mutant demonstrated that the level of disaturated PG cannot be the sole determinant ofplant chilling sensitivity. fab1 plants contain an increased proportion of 16:0 fatty acidsbecause of a partial defect in 3-ketoacyl-ACP synthase II, the enzyme responsible forelongation of 16:0 to 18:0.17 As a consequence, PG from fab1 leaves contains 43%disaturated molecular species compared with only 9% in PG from the wild type. Theproportion of disaturated PG in fab1 falls close to the middle of the range found for chilling-sensitive plants and makes the fab1 mutant comparable to species such as castor bean,cucumber, maize and tobacco.18 However, fab1 plants were able to grow and completetheir life cycle normally at 10˚C. They were also unaffected (as compared with wild typecontrols) by more severe chilling treatments that quickly led to the death of cucumber andother chilling-sensitive plants. These treatments included 4˚C for 7 days in the dark, 2˚C for 7days in the light and freezing to -2˚C for 24 hours.17 Following each of these treatments,mutant plants returned to 22˚C remained indistinguishable from wild type controls andflowered and set seed normally.

fab1 mutant plants do eventually show damage when grown continuously at 2˚C withreduced photosynthesis, reduced growth and leaf chlorosis developing gradually from 10to 35 days of low-temperature treatment. At 2˚C, fab1 plants undergo a process of chloroplastautophagy.19 Therefore, although the fab1 mutant does not exhibit classic chilling sensitivity,the results of Wu et al19 confirm a deleterious effect of high levels of disaturated PG onlow-temperature fitness and provide a rationale for the relatively low content of disaturatedPG among chilling-tolerant species which may be exposed to low temperatures forextended periods of the life cycle. Indeed, the chilling-induced chlorosis and slow growthof the fab1 mutant could be due to a defect in chloroplast membrane protein turnover oraccumulation of the kind described by Moon et al12 However, the side-by-side comparisonof the fab1 mutant with naturally chilling-sensitive species that contain similar levels of


disaturated PG emphasizes the point that factors other than high levels of disaturated PG areresponsible for the injury sustained by these and other chilling-sensitive plants.

In summary, these results make it clear that reducing the proportion of disaturatedPG, and perhaps increasing overall lipid unsaturation, can measurably improve thelow-temperature performance of tobacco plants, possibly through facilitating turnoverof the D1 protein and thereby allowing faster recovery from photoinhibition. At the sametime, disaturated PG cannot be considered the sole cause of chilling sensitivity becausehigh levels of disaturated PG in Arabidopsis fab1 did not produce a typical chilling-sensitive phenotype. Nor is the manipulation of membrane lipids the only way to improvelow-temperature performance of tobacco plants. Gupta et al20 produced transgenictobacco plants that overexpress a chloroplastic Cu/Zn superoxide dismutase. Leaf disksfrom transgenic plants had higher rates of photosynthesis at 10˚C compared withuntransformed controls and also exhibited a greater capacity for recovery at 25˚C afterphotoinhibition at 3˚C for 4 hours. These findings imply that protecting tissues from theeffects of oxidative stress may also reduce chilling damage.

The work described here on higher plants is complemented by studies in cyanobacteria.In these prokaryotes, a high level of saturated fatty acids is correlated with an inability togrow at low temperatures, either because of reduced processing of D1 protein21 orreduced activity of nitrate uptake.22 A more complete review of studies in cyanobacteria isincluded in Nishida and Murata.23

Additional Mechanisms of Chilling InjuryAlthough the roles of membrane lipid unsaturation and disaturated PG in chilling

sensitivity have been well established, there are also well-documented examples where otherprocesses must be responsible. One of these involves the chilling of tomato plants in thedark, under which condition photoinhibition and considerations of D1 turnover are clearlynot relevant. Tomato plants that have been chilled in the dark show greatly reducedphotosynthesis rates during subsequent illumination. Understanding the possible causeof this chilling damage started with the observation that many proteins involved in photo-synthesis are products of genes whose transcriptional activities cycle under control of thecircadian clock. Martino-Catt and Ort24 used genes for the chlorophyll a/b bindingprotein of photosystem II (Cab) and for ribulose-1,5-bisphosphate carboxylase/oxygenaseactivase (rca) to demonstrate that chilling stops the circadian clock. They discoveredthat low temperature has two separate effects on the normal pattern of expression of Cab andrca proteins:

1. Progression of the timing of the circadian clock controlling gene transcription issuspended throughout the period of low-temperature exposure; and

2. Normal turnover of the existing transcripts is suspended.Upon rewarming, the circadian rhythm of transcriptional and translational activity is

reestablished, but is out of phase with the actual time of day by the amount of time that thetomato plant was at low temperature. In addition, after rewarming, the messages that werestabilized at low temperature can no longer be translated into protein. From these results, itis reasonable to suggest that a range of gene-expression and other functions controlled bythe circadian clock may be affected by chilling in tomato. This would clearly be expected toresult in considerable disruption of photosynthesis and other cellular functions.

A few plant species have been demonstrated to have a capacity to increase their chillingtolerance in response to treatment at modestly cool temperatures. Dark-grown seedlings of themaize inbred G50 were killed by exposure to 4˚C for 7 days but could be induced to survivethis treatment by prior exposure to 14˚C for 3 days.25 Differential screening techniquesallowed the isolation of cDNAs representing chilling acclimation responsive genes including


cat3, which encodes the mitochondrial catalase 3 isozyme. Hydrogen peroxide levels in theseedlings were increased during acclimation at 14˚C and treatment of seedlings grown at2˚C with H2O2 induced chilling tolerance and increased both cat3 transcript levels and theactivities of catalase 3 and guaiacol peroxidase. From these results, it appears that peroxidehas dual effects at low temperatures. During acclimation at 14˚C, its early accumulationsignals the production of antioxidant enzymes such as catalase 3 and guaiacol peroxidase.At 4˚C, in nonacclimated seedlings, it accumulates due to low levels of these, and perhapsother, antioxidant enzymes and may cause damage through oxidation of lipids and proteins.26

Genes Required for Chilling ToleranceMuch of the discussion of temperature adaptation of plants focuses on finding

defects in chilling-sensitive species that can explain why they are damaged by low tempera-tures. However, it is probably more useful, especially at the genetic level, to identify thetraits that are responsible for the chilling tolerance observed in temperate plants. Thus, itis possible to screen mutant populations of a chilling tolerant species such as Arabidopsisfor plants that are no longer fully tolerant to low temperature. Mutants with impairedchilling tolerance are defined as those which have a wild type appearance at normal growthtemperatures but which show damage when transferred to chilling temperatures. Thesemutants each contain a mutation that has no effect at normal temperatures but is disruptive atchilling temperatures. In such a screen, only mutational defects that sensitize a mutant tochilling will be identified; mutations associated exclusively with other processes such asfreezing tolerance are excluded. Somerville and coworkers initiated such a mutationalapproach.27 They screened a population of Arabidopsis mutated by ethyl methane sulfonate

Fig.4.1. The effects of chillingtemperature on the growth ratesof Arabidopsis lipid mutants. Theresults shown are for (A) fad5 andfad6 at 5˚C (B); fab1 at 2˚C; and(C) fad2 at 6˚C. Wild type controlsare included in each experiment.


(EMS). About 20 mutants were isolated that showed damage symptoms in response to a briefand mild chilling regime; they had a normal, wild type phenotype at 22˚C, but after a weekat 13˚C exhibited visual damage.28,29 An extensive characterization of one mutant, chs1,27-30

reinforces the validity of the mutational approach. The chs1 mutant showedlow-temperature-induced chlorosis indicating a lesion in chloroplast maintenance at lowtemperatures. Subsequent investigations revealed a loss of chloroplast integrity30 andreduced accumulation of proteins localized to the chloroplast.28,29 The detected changesindicate a sequence of chilling-induced damage caused by disrupted protein accumulation inthe chloroplast. Nevertheless, it has not been possible to identify the precise biochemicallesion responsible for initiating these changes.

A collection of Arabidopsis mutants with defects in membrane lipid unsaturation31

have offered useful perspectives on the role of membrane unsaturation. Five mutant lines-fab1 (see above), fad5, fad6, fad2 and the triple mutant—fad3-fad7-fad8—show damagesymptoms when grown at 2˚-6˚C.17,32,33 All of these mutant lines are similar to wild typewhen grown at 22˚C, but their growth rates are lower than wild type at chilling temperatures(Fig. 4.1). However, in all cases (as discussed above for fab1), the low-temperature damageis distinct from typical chilling sensitivity. For example, symptom development is moregradual and damage is not exacerbated following a return to 22˚C. These results make itclear that a suitable membrane lipid composition is required for chilling tolerance, butthat it is unlikely that membrane defects are the sole cause of chilling sensitivity.

A more extreme chilling screen was used by Tokuhisa et al34 who exposed Arabidopsisplants to 5˚C for up to 42 days and looked for mutants both during the chilling treatmentand after return of the plants to 22˚C. A screen of EMS mutagenized plants using this protocolidentified 3% of the plants as having chilling-induced phenotypes including chlorosis,reduced growth, necrosis and death. One drawback of EMS mutagenesis is that themutations are primarily single base pair substitutions. In many cases, these mutationsdestroy the function of the gene product. However, there are many examples wheremissense mutations result in an amino acid substitution in the mutated gene such thatthe altered polypeptide product functions adequately at normal (permissive) temperatures butloses function at low (nonpermissive) temperatures. If such a missense mutation is in anessential gene, the mutation will render a chilling-induced phenotype. Such alleles havebeen used extensively in yeast35 and E. coli36 to characterize essential housekeeping genes,and have been termed cold-sensitive or cs alleles. To circumvent this problem, Tokuhisa etal34 repeated the screen on a population in which mutations have been generated byT-DNA insertion.37 Insertion mutagenesis produces a high proportion of null allelesand will thus facilitate the identification, in our screen, of genes which are unnecessaryat 22˚C but which are essential for proper growth at 5˚C. Just as importantly, the T-DNAinsertion can act as a starting-point to clone and characterize the specific chilling-tolerancegene. Over 8,000 lines of mutants generated by T-DNA insertional mutagenesis were screenedand about 280 putative mutants were identified. To date, about 200 of these putativeshave been rescreened and 21 mutants have been shown to have heritable, chilling-impaired phenotypes. Two of these mutants, which exhibited chilling-induced chlorosiswere designated paleface1 (pfc1) and pfc2. A third mutant that was inhibited in leafexpansion at 5˚C was designated stop1 (sop1). By segregation analysis, each of thesemutants has been shown to have linkage, within 2-3 centiMorgans between the kanamycinresistance marker in the T-DNA and the chilling-induced phenotype. Therefore, it ishighly probable that the T-DNA in each of these lines is inserted in a gene which isrequired for chilling tolerance.

Molecular characterization of the pfc1 mutant has demonstrated a previouslyunrecognized requirement for ribosomal RNA processing and modification to provide chilling


tolerance.38 The wild type allele of the mutated gene and a near-full length (>93%) cDNAclone were isolated by using the T-DNA as a tag. The deduced polypeptide has a 50 aminoacid transit peptide for chloroplast targeting, an S-adenosylmethionine-binding motif and34% identity with genes from bacteria and yeast encoding ribosomal RNA methylases whichare required for ribosomal RNA processing or translation. The PFC1 transcript was absentfrom pfc1 plants and biochemical analyses indicated that the expected methylation ofadenosines 1518 and 1519 in the small subunit rRNA occurred in the wild type but not inpfc1. Finally, expression of an antisense PFC1 construct in wild type Arabidopsis producedplants which exhibited the same low-temperature chlorosis seen in the pfc1 mutant. These resultsdemonstrate that PFC1 function is specifically required for low-temperature tolerance ofArabidopsis.

Freezing Tolerance

Causes of Freezing InjuryAs temperatures drop below freezing, ice forms primarily in the intercellular spaces

(formation of intracellular ice is generally thought to be a fatal event).39 Ice formation isinitiated extracellularly largely due to the apoplastic fluid having a higher freezing pointthan the intracellular fluids, but may also involve the relative levels of ice nucleating agents.40

Accumulation of ice in the intercellular spaces can potentially result in physical disruptionof the tissues and cells.41 However, most of the injury is thought to result from the severecellular dehydration that occurs with freezing.39,42 The chemical potential of ice is lessthan that of unfrozen water at a given temperature. Thus, when ice forms extracellularly,there is a drop in water potential outside the cell. Consequently, there is movement ofunfrozen water from inside the cell to outside the cell. The net amount of water movementdepends on both the initial solute concentration of the intracellular fluid and the freezingtemperature, which directly determines the chemical potential of the ice. Freezing at -10˚Cresults in an osmotic potential of about five osmolar and typically, movement of greaterthan 90% of the osmotically active water out of the cell.

Freeze-induced cellular dehydration could have a number of deleterious effects,resulting in cellular damage such as the denaturation of proteins and precipitation ofsolutes.39,43 However, the best documented injury occurs at the membrane level.42,44

Detailed analyses by Steponkus and colleagues45,46 have demonstrated that multiple formsof membrane lesions occur in response to freezing. The specific type of membrane damagedepends on the freezing temperature and corresponding severity of cellular dehydration.At freezing temperatures between about -2˚C and -5˚C, the predominant form of injury innonacclimated plants is “expansion-induced lysis”. It results from the cycle of osmoticcontraction and expansion that occurs with freezing and thawing. Specifically, whenprotoplasts from leaves of nonacclimated plants are frozen to about -4˚C, they dehydrate, andas they shrink, endocytotic vesicles bud off from the plasma membrane. When the protoplastsare thawed and water moves back into the cells, the vesicular material is not reincorporatedinto the plasma membranes, resulting in a decrease in membrane surface area. Consequently,rehydration results in an intolerable osmotic pressure and the cells burst. Freezing ofnonacclimated cells to slightly lower temperatures, approximately -5˚ to -10˚C, results inanother form of membrane damage, lamellar-to-hexagonal-II phase transitions.45,46 In thiscase, cells do not burst upon thawing, but instead become osmotically unresponsive due to themembranes losing their semipermeable characteristics. Freezing cold-acclimated cells to evenlower temperatures, with consequent lower water potentials and more severe dehydration,results in additional forms of membrane damage, including “fracture jump lesions”.45,46


Mechanisms of Freezing ToleranceGiven the central role of membranes in freezing-injury, it is not surprising that multiple

mechanisms appear to be involved in increasing the cryostability of membranes during coldacclimation. Steponkus and colleagues45,46 have demonstrated that unlike plasma membranesfrom nonacclimated plants, plasma membranes from cold-acclimated plants do not sufferexpansion-induced lysis or formation of hexagonal II phase lipids. The elimination ofthese forms of membrane damage involve a number of changes in lipid composition,including increased levels of fatty acid desaturation in membrane phospholipids.45,46 Inaddition, the accumulation of sucrose and other simple sugars that typically occurs withcold acclimation seems likely to contribute to the stabilization of membranes, as thesemolecules can protect membranes against freeze-induced damage in vitro.47,48 Finally, asdiscussed below, there is emerging evidence that certain cold-induced hydrophilicpolypeptides help stabilize membranes against freeze-induced injury.

Additional mechanisms could also potentially contribute to freezing tolerance,including ones that help prevent or reverse freeze-induced denaturation of proteins orlessen the direct physical damage to cells caused by the accumulation of extracellular ice.Indeed, molecular chaperones have been shown to accumulate during cold acclimation,including a spinach Hsp70,49,50 a soybean Hsp7051 and a Brassica napus Hsp90.52 In addition,there is evidence suggesting that “freeze-inhibitor” sugars might lessen cellular damage bypreventing the formation of adhesions between extracellular ice and the cell walls.53 Also,recent studies indicate that many plants accumulate antifreeze proteins during coldacclimation, some of which are present in the apoplastic fluids.54-56 These proteinsprobably do not act by preventing ice formation as they are capable of imparting only afew tenths of a degree of thermal hysteresis (i.e., lower the freezing temperature by a fewtenths of a degree without affecting the melting point of the solution). However, theycould potentially contribute to freezing tolerance by modifying ice crystal structure and/orpreventing ice recrystallization. In all of these cases, however, further study is required toclearly establish whether the proteins or sugars contribute significantly to freezingtolerance.

Role of Cold-Responsive Genes in Freezing ToleranceIn 1985, Guy et al57 established that changes in gene expression occur during cold

acclimation. Since then, a fundamental question in cold acclimation research has been todetermine whether cold-responsive genes have roles in freezing tolerance. To address thisissue, researchers have engaged in the isolation and characterization of genes that areinduced during cold acclimation.58,59 Many of these cold-responsive genes encode proteinswith known activities that could potentially contribute to freezing tolerance. Forinstance, the Arabidopsis FAD8 gene60 and barley blt4 genes,61 which encode a fatty aciddesaturase and a putative lipid transfer protein, respectively, are induced in response to lowtemperature. These genes might contribute to freezing tolerance by altering lipid composition.As alluded to above, cold-responsive genes encoding molecular chaperones49-52 mightcontribute to freezing tolerance by stabilizing proteins against freeze-induced denaturation.Cold-responsive genes encoding various signal transduction and regulatory proteins havealso been identified, including MAP kinases,62,63 calcium-dependent protein kinases64,65

and 14-3-3 proteins.66 These proteins might contribute to freezing tolerance by controllingthe expression of cold-responsive genes or by regulating the activity of proteins involved infreezing tolerance. Whether any of these cold-responsive genes have important roles in freezingtolerance, however, remains to be determined.

While many of the cold-responsive genes that have been isolated from cold-acclimatedplants encode proteins with known activities, the majority do not. Indeed, most encode


Fig. 4.2. Genomic organization of COR gene families. Coding and intron regions are depicted asfilled and open boxes, respectively. Alternative gene designations are listed. Accession numbersfor the genes are: COR15a, X64138; COR15b, L24070; KIN1, X51474; COR6.6/KIN2, X55053/X62281; LTI65/RD29b, X67670/D13044; COR78/LTI78/RD29a, L22567/X67071/D13044; LTI29/ERD10, X90958/D17714; COR47/RD17, X90959/AB004872. The figure was drawn by KathyWilhelm.

extremely hydrophilic polypeptides that are either newly discovered or are homologs of LEA(late embryogenesis abundant) proteins.59,67 LEA genes are induced late in embryogenesis,just prior to seed desiccation, and like many of the cold-responsive genes, are induced inresponse to dehydration and ABA.68-70 Based largely on these expression characteristicsand the close relationship between freezing and dehydration injury, it has been widelyspeculated that the cold-responsive genes encoding the novel hydrophilic and LEA proteins mightcontribute to freezing tolerance. Indeed, recent results provide direct evidence that theArabidopsis COR (cold-regulated) genes contribute to the increase in freezing tolerancethat occurs with cold acclimation.71,72

Arabidopsis COR Genes The Arabidopsis COR genes—also designated LTI (low temperature induced), KIN (cold-

inducible), RD (responsive to desiccation) and ERD (early dehydration-inducible)—comprisefour gene families.67 Each family is composed of two genes that are physically linked in thegenome in tandem array (Fig. 4.2). The COR78, COR15, and COR6.6 gene pairs encode newlydiscovered hydrophilic polypeptides, while the COR47 gen pair encodes homologs of LEA groupII proteins (also known as dehydrins and LEA D11 proteins).68,69 At least one member ofeach gene pair is induced in response to low temperature, dehydration and exogenousapplication of ABA.58


To determine whether COR15a might have a role in freezing tolerance, Artus et al.71

constructed transgenic plants that constitutively express the gene and assessed the effectsthat this had on freezing tolerance. COR15a encodes a 15 kDa polypeptide that is targetedto the stromal compartment of chloroplasts.73,74 The mature 9.4 kDa polypeptide,COR15am, is extremely hydrophilic and, like the other COR polypeptides and many LEAproteins, has the unusual property of remaining soluble upon boiling in aqueous buffer. Inthe initial experiments, Artus et al71 compared the freezing tolerance of chloroplasts innonacclimated transgenic and wild type plants. The results indicated that theCOR15am-containing chloroplasts in transgenic plants were 1 to 2˚C more freezingtolerant than were the chloroplasts in wild type plants that did not contain COR15am(cold acclimation increased chloroplast freezing tolerance about 6˚C). In additionalexperiments, they found that the effects of COR15am were not limited to the chloroplasts.Protoplasts isolated from leaves of the nonacclimated transgenic plants that constitutivelyproduced COR15am were about 1˚C more freezing tolerant at freezing temperaturesbetween -5 and -8˚C than were those isolated from nonacclimated wild type plants.Significantly, protoplast survival was measured by vital staining with fluorescein diacetate,a method that reports on retention of the semipermeable characteristic of the plasmamembrane. Thus, it could be concluded from the protoplast survival experiments thatconstitutive expression of COR15a resulted in an increase in plasma membranecyrostability.

The results of Artus et al71 indicate a role for COR15a in freezing tolerance. However,unlike cold acclimation which increases protoplast survival over the range of -2˚ to -8˚C,expression of COR15a only increased survival over the temperature range of -5˚ to -8˚C (ifanything, COR15a expression resulted in a slight decrease in protoplast survival between-2˚ and -4˚C). A possible explanation for this finding is that COR15a expression mightprevent certain membrane lesions, but not others. As discussed earlier, the predominantform of membrane injury over the range of -2˚ to -4˚C appears to be expansion-inducedlysis, while over the range of -5˚ to -8˚C, the predominant form of injury is freeze-inducedlamellar-to-hexagonal II phase transitions. Thus, it is possible that constitutive expressionof COR15a might defer the incidence of freeze-induced formation of hexagonal II phase lipidsto a lower temperature, but have little or no effect on the incidence of expansion-induced lysis.Additional experimentation is required to test this hypothesis.

The mechanism by which COR15a stabilizes membranes against freeze-inducedinjury is not yet known. It seems unlikely that the COR15am protein has enzymatic activity,as it has a very simple amino acid composition and structure: It is rich in alanine (21%),lysine (18%),glutamic acid (15%) and aspartic acid (10%) residues (which comprise greaterthan 60% of the protein); is devoid of proline, methionine, tryptophan, cysteine, arginineand histidine residues; and is comprised largely of a 13 amino acid sequence that isrepeated (imperfectly) four times. This, however, leaves open many possibilities. COR15ammay act indirectly to stabilize membranes. For example, it could potentially regulate theactivity of proteins that have roles in freezing tolerance, such as enzymes involved in sugaror lipid metabolism. Alternatively, COR15am might interact directly with the chloroplastenvelope and increase membrane cryostability in some manner. The location of COR15amwithin the chloroplast is not necessarily inconsistent with protection of the plasma membrane,as formation of the hexagonal II phase is an interbilayer event that occurs largely between theplasmalemma and the chloroplast envelope. Decreasing the propensity of the chloroplastenvelope to fuse with the plasma membrane could result in less damage to the plasmamembrane. Experiments to detect a direct effect of COR15am on the stabilization ofmembranes, however, have yielded equivocal results.75,76


Although constitutive expression of COR15a enhances freezing tolerance at both theorganelle (chloroplast) and cellular (protoplast) level, the effects are modest.71 Moreover,unlike cold acclimation, COR15a expression alone does not result in a detectable increasein freezing survival of whole plants.72 These findings are not surprising given the results ofgenetic analyses indicating that freezing tolerance is a multigenic trait involving genes withadditive effects.77 Indeed, multiple genes are activated with cold acclimation in Arabidopsis,including at least one member of each of the four COR gene pairs.58

If multiple COR genes act in concert to increase freezing tolerance, then expression ofthe entire COR gene “regulon” would presumably increase freezing tolerance more than

Fig. 4.3. CBF1 and COR transcript levels in nonacclimated wild-type Arabidopsis plants (RLD)and nonacclimated transgenic Arabidopsis plant lines that overexpress either CBF1 (A6 and B16)72

or COR15a (T8).71 Overexpression of CBF1 and COR15a was accomplished by transformingwild type RLD plants with hybrid gene constructs having the coding sequences for either CBF1or COR15a under control of the cauliflower mosaic virus 35S promoter.71,72 Total RNA wasprepared from leaves of nonacclimated plants and analyzed for CBF1 and COR transcripts byRNA blot analysis using 32P-radiolabeled probes.71,72 Overexpression of CBF1 results in thestimulation of COR gene expression, but does not affect the transcript levels of eIF4A (eukaryoticinitiation factor 4A),92 a constitutively expressed gene that is not responsive to low temperature.


expressing COR15a alone. This hypothesis was recently tested by Jaglo-Ottosen et al.72

Expression of the entire battery of COR genes was accomplished by overexpressing theArabidopsis transcriptional activator CBF1 (CRT/DRE binding factor 1).78 CBF1 binds toa DNA regulatory element, the CRT (C-repeat)/DRE (drought responsive element), thatstimulates transcription in response to both low temperature and water deficit.79 The elementis present in the promoters of COR15a, COR78, COR6.6, COR47 and presumably other yet tobe identified COR genes. Jaglo-Ottosen et al72 found that constitutive overexpression of CBF1induce expression of the COR genes in nonacclimated Arabidopsis plants (Fig. 4.3) andincreased freezing tolerance at the whole plant level, an effect that was not observed byexpressing COR15a alone. Thus, it appears that additional members of the ArabidopsisCRT/DRE regulon are freezing tolerance genes that have roles in cold acclimation.Determining which CRT/DRE-regulated genes have roles in freezing tolerance andtheir functions are now important goals. In addition, a critical point to establish is whetherthe CRT/DRE-containing COR genes regulate the full array, or only a subset, of thebiochemical changes that occur with cold acclimation (alterations in lipid composition,accumulation of sugars, synthesis of anthocyanin, etc.).

Other Possible Freezing Tolerance ProteinsMore than 20 years ago, Volger and Heber80 reported that cold-acclimated spinach

and cabbage synthesize polypeptides that are highly effective in protecting isolated thylakoidmembranes against freeze-thaw damage in vitro. These putative cryoprotective polypeptideswere detected in cold-acclimated plants, but not nonacclimated plants, suggesting thatthey were encoded by cold-regulated genes. Subsequent studies by Hincha and colleagues81,82

indicated that the cryoprotective polypeptides act to protect membranes against freeze-induced damage by reducing membrane permeability during freezing and increasingmembrane expandability during thawing. A significant limitation in all of these studies,however, was that only partially purified protein preparations were used. Thus, it wasunclear whether the cryoprotective activity detected was due to a single protein ormultiple polypeptides. Interestingly, however, from the enrichment procedures used, itwas evident that the polypeptides, like the COR polypeptides, were very hydrophilic andremained soluble upon boiling.

A significant advance in the study of the spinach and cabbage cryoprotective proteinswas recently made by Sieg et al.83 These investigators purified a single cryoprotective proteinfrom cold-acclimated cabbage that is effective in protecting isolated thylakoids againstfreeze-thaw damage in vitro. This protein, which was designated “cryoprotectin,” has a mass of 7kDa, remains soluble upon boiling and appears to be encoded by a cold-inducible gene(the protein is present in cold-acclimated plants, but not in nonacclimated plants).Unfortunately, there is no information on the amino acid sequence of cryoprotectin,and thus, it is unknown whether it is related to any of the hydrophilic polypeptidesencoded by the cold-responsive genes described above. Additional investigation shouldreveal more about the nature of cryoprotectin, its mode of action in vitro, and providedirect evidence whether it has a role in protecting membranes against freezing-injuryin vivo.

There is evidence accumulating that suggests certain LEA proteins may also contribute tofreezing tolerance. The HVA1 gene of barley, which encodes a LEA group III protein (alsoknown as LEA D7 proteins), is expressed in aleurone layers late in embryogenesis and inseedlings in response to low temperature, ABA and water deficit.84 Although there is nodirect evidence that HVA1 expression contributes to increased freezing tolerance, recentresults indicate that the gene is able to confer tolerance to dehydration stress. Xu et al85

have reported that expression of HVA1 in transgenic rice results in increased tolerance to


both water deficit and high salinity stress. Given the relationship between dehydrationtolerance and freezing tolerance, HVA1 seems to be a strong candidate for having a role infreezing tolerance. Similarly, Iu et al86 have recently shown that expression of the tomatoLe25 gene, which encodes a LEA D113 protein, increases both the freezing and high salinitytolerance of yeast cells. Thus, homologs of Le25 would seem to be good candidates forbeing freezing tolerance genes. In tomato, which is a chilling sensitive plant that does notcold acclimate, Le25 is expressed at very low levels (if at all) in response to low temperature.87

Whether it is expressed at high levels at low temperature in plants that cold acclimateremains to be determined.

Isolation of Mutants Affected in Freezing ToleranceA powerful approach to identify freezing tolerance genes is to isolate and characterize

mutants that are altered in their ability to survive freezing. Warren and colleagues88,89 andXin and Browse90 have recently made exciting progress in this area. The specific approachtaken by Warren and colleagues88,89 has been to isolate Arabidopsis mutants that are lessfreezing tolerant than wild type plants after a period of cold acclimation. They screenedM3 seed pools from 1804 chemically mutagenized M2 Arabidopsis plants for mutants witha decreased capacity to cold acclimate. These efforts resulted in the identification of 13mutant lines that were defective in freezing tolerance. Seven of these lines displayed chillingsensitive phenotypes that might have indirectly resulted in a diminished capacity to coldacclimate. However, six of the lines were able to withstand long periods of cold acclimation(up to 56 days) without displaying any obvious adverse effects. Thus, in these cases, it wouldappear that the mutations have direct effects on the cold acclimation process itself. Geneticcomplementation analysis indicated that these lines had suffered mutations in fiveSFR (sensitivity to freezing) genes—SFR1, 2, 4, 5 and 6 (two sfr5 mutant alleles were isolated).

Wild type Arabidopsis plants that are cold-acclimated for 2 weeks at 4˚C suffer noobvious damage upon being frozen at -6˚C for 24 hours followed by transfer to normalgrowth temperature. In contrast, Arabidopsis plants carrying the sfr1, 2, 4, 5-1, 5-2 and 6mutations do suffer injury. The injury observed varies with the different mutations (Table

Table 4.1 Phenotypes associated with sfr mutations in cold-acclimated plants.88,89

Mutant Freezing Anthocyanin Glucose and Fatty AcidGene Sensitivity Level (% wt) Sucrose Levels Composition

sfr1 Young leaves 162 Normal No changes detected

sfr2 All leaves (severe) 87 Normal No changes detected

sfr4 All leaves (severe) 8 Reduced amounts of Reduced levels of 16:0,both glucose (<<10% wt) 18:1 and 18:2 fatty acidsand sucrose (<25% wt)

sfr5 All leaves 115 Normal No changes detected

sfr6 Worst in young leaves 43 Normal No changes detected


4.1). The sfr1 mutation affects the freezing tolerance of only young leaves; the sfr6 mutation hasits most severe effects on young leaves, but affects all leaves to some extent; and the sfr2, 4,sfr5-1 and sfr5-2 mutations affect all leaves equally. Significantly, all of the mutations affectthe cryostability of the plasma membrane, as indicated by the electrolyte leakage test. Inthis test, detached leaves are frozen to various temperatures below zero and after thawing,cellular damage is assessed by measuring electrolyte leakage. Leakage of ions from the cellsis an indication that the semipermeable nature of the plasma membrane has been lost, atleast transiently, in response to freezing. With the sfr1, 4, 5 and 6 mutations, the severity ofthe freezing damage observed in the whole plant freezing tests corresponded with theresults of the electrolyte leakage test. Thus, the freezing sensitivity caused by these mutationsappears to result largely from a decrease in membrane cryostability. In contrast, the sfr2mutation resulted in severe injury in the whole plant freeze test, but only minor damage inthe electrolyte leakage test. Thus, it was suggested89 that the freezing-sensitive lesion causedby this mutation might not have a primary effect on cellular membranes.

The identity of the sfr genes are not yet known, nor is the molecular basis for howmutations in the genes affect freezing tolerance. The sfr2, sfr5-1 and sfr5-2 mutations donot have any obvious effects on the alterations in fatty acid composition and increases insucrose and anthocyanin levels that normally occur with cold acclimation (Table 4.1). Thesfr4 mutation, however, results in reduced accumulation of sucrose, glucose and anthocyaninand lowered levels of 18:1 and 18:2 fatty acids (Table 4.1). Given the likely role of sugars ascryoprotectants and demonstrated roles of fatty acid composition in membranecryostability, it is reasonable to speculate that the effects that the sfr4 mutation has onsugar and fatty acid composition account, at least in part, for the freezing sensitive phenotypeof these mutants. A role for anthocyanins in freezing tolerance is less clear. Interestingly,however, the sfr1 and sfr6 freezing sensitive mutations also affect anthocyanin production;they cause increased and decreased accumulation, respectively (Table 4.1). This suggestssome link between anthocyanin biosynthesis and freezing tolerance, but the nature of thatlink is obscure. Finally, both the sfr4 and sfr6 mutations have phenotypes that are notdirectly associated with low temperature, namely slow growth of plants at the seedlingstage and racemes having a “bottle brush” appearance, respectively.88 The isolation andidentification of the products encoded by the sfr genes should provide significant newinsight into our understanding of the cold acclimation response.

Another gene that has a major effect on freezing tolerance, eskimo1 (esk1), hasrecently been identified by Xin and Browse.90 These investigators screened 800,000 chemicallymutangenized M2 seedlings of Arabidopsis for mutants that displayed “constitutive” freezingtolerance, i.e., mutants that were more freezing tolerant than wild type plants without coldacclimation. A number of such mutants were isolated, the best characterized being esk1.Whereas nonacclimated wild type plants were found to have an LT50 of -2.8˚C in a wholeplant freeze test, the esk1 mutant plants had an LT50 of -10.6˚C. Moreover, the esk1 mutationincreased the freezing tolerance of cold-acclimated plants. Wild type plants that had beencold-acclimated had an LT50 of -12.6˚C, while cold-acclimated esk1 plants had an LT50

of -14.8˚C.The molecular basis for the increase in freezing tolerance displayed by the esk1

mutation is not yet certain. However, the esk1 mutation has been shown to have a dramaticeffect on proline concentration; the proline levels in the esk1 mutant are about 30-foldhigher than they are in wild type plants.90 It seems likely that this contributes to theincreased freezing tolerance of the esk1 plants, as proline has been shown to be an effectivecryoprotectant in vitro.91 In addition, total sugars are elevated in the esk1 mutant abouttwo-fold and expression of the RAB18 cold-responsive LEA group II gene is elevated aboutthree-fold. These alterations may also contribute to the increase in freezing tolerance.


Perhaps the most important observation made in regard to the esk1 mutation, however, isthat it does not affect the expression of any of the COR genes; expression of COR15a,COR6.6, COR47 and COR78 remains at low levels under normal growth conditions and isgreatly induced in response to low temperature. These results suggest that there may bemultiple signaling pathways involved in activating different aspects of the cold acclimationresponse and that activation of one pathway can result in considerable freezing tolerancewithout activation of the other pathways. As discussed above, overexpression of the CBF1transcription factor induces expression of the CRT/DRE gene regulon and results in asignificant increase in freezing tolerance. The “CBF1 pathway” might therefore control oneset of cold acclimation responses. Similarly, the ESK1 gene may participate in the control ofanother set of freezing tolerance responses that includes synthesis of proline, and to alesser degree, synthesis of sugars and expression of RAB18. The mechanism of ESK1action is not known. However, the fact that the two available esk1 alleles are recessive suggeststhat ESK1 might act as a negative regulator. Further insight into the nature and function ofESK1 will come from isolation of the ESK1 gene and characterization of the encoded geneproduct.

Conclusions and PerspectivesIn a short review such as this, it is impossible to do justice to all of the significant

investigations that have been made regarding plant cold tolerance. We, therefore, chose tofocus primarily on recent advances in our understanding of chilling sensitivity and freezingtolerance at the molecular and genetic levels. From the studies presented on chilling sensitivity,we suggest the following hypotheses and framework for considering chilling responses inplants. For plants that evolved in consistently warm habitats, there has been no selectionagainst traits that compromise low-temperature growth. Thus, many such traits may haveevolved in tropical and subtropical species, especially if they confer even a small selectiveadvantage at higher temperatures. Some of these acquired characteristics might affect plantperformance only after extended cold treatment, whereas others might result in damage onthe much shorter time scale normally associated with chilling sensitivity. The progressivedispersal of angiosperms to temperate climates would have required the elimination of alltraits that affected plant performance in the new, periodically cooler environment. Animportant corollary of this proposal is that any chilling-sensitive species is likely to possessmultiple traits that restrict its geographical range. This suggestion is supported by theexamples we have described and by the results of plant breeding experiments indicatingthat improvements in chilling tolerance are typically multigenic and may be specificto particular stages in the plant life cycle. A second corollary is that any particular chilling-associated trait may not be found in all chilling-sensitive species because each trait mayhave arisen more than once during evolution.

In regard to freezing tolerance, it had been widely speculated for more than a decadethat the changes in gene expression that occur with cold acclimation might contribute tothe increased freezing tolerance displayed by cold-acclimated plants. The recent results of Artuset al71 and Jaglo-Ottosen et al72 indicate that this is indeed the case. Thus, the fundamentalissue of whether cold-responsive genes have roles in freezing tolerance now shifts to identifyingwhich cold-responsive genes have key roles in cold acclimation and determining their specificmodes of action. In Arabidopsis, it will be important to determine which members of theCRT/DRE regulon contribute to freezing tolerance and to establish their functions. It will alsobe of fundamental interest to determine whether the CRT/DRE regulon is highly conservedamong plants and if so, whether it is regulated by CBF1-related transcription factors.Finally, it will be important to determine the extent to which the cold acclimationresponse is conditioned by the CRT/DRE regulon, i.e., determine how much of the


increase in freezing tolerance that occurs with cold acclimation is due to action ofCRT/DRE-regulated genes. That the CRT/DRE regulon may account for only a portion ofthe increase in freezing tolerance is suggested by the study of Xin and Browse;90 analysis ofthe esk1 mutants suggests the possibility that cold acclimation involves multiple cold-activated processes that are controlled by different signal transduction pathways andthat each process/pathway contributes to freezing tolerance in an independentfashion.The investigations now underway detailing the function and regulation of cold-responsive genes and the isolation and characterization of mutants altered in freezing toleranceshould provide, in the near future, fundamental insight into the number and nature of“freezing tolerance regulons” and the signaling pathways that control their expression.

References1. Lyons JM. Chilling injury in plants. Annu Rev Plant Physiol 1973; 24:445-466.2. Wang CY. Chilling injury of Horticultural Crops. Boca Raton:CRC Press, 1990.3. Lyons JM, Graham D, and Raison JK. Low Temperature Stress in Plants. New York:Academic

Press, 1979.4. Murata N, Nishida I. Lipids in relation to chilling sensitivity of plants. In: Wang CY, ed.

Chilling injury of Horticultural Crops. Boca Raton:CRC Press, 1990:181-199.5. Raison JK. The influence of temperature-induced phase changes on the kinetics of respira-

tory and other membrane associated enzyme systems. Bioenergetics 1973; 4:285-309.6. Raison JK, Orr GR. Proposals for a better understanding of the molecular basis of chilling

injury. In: Wang CY, ed. Chilling injury of Horticultural Crops. Boca Raton:CRC Press,1990:145-164.

7. Raison, JK, Wright LC. Thermal phase transitions in the polar lipids of plant membranes:Their induction by disaturated phospholipids and their possible relation to chilling injury.Biochim Biophys Acta 1983; 731:69-74.

8. Murata N, Sato N, Takahashi N et al. Compositions and positional distributions of fattyacids in phospholipids from leaves of chilling-sensitive and chilling-resistant plants. PlantCell Physiol 1982; 23:1071-1079.

9. Murata N, Yamaya J. Temperature-dependent phase behavior of phosphatidylglycerols fromchilling-sensitive and chilling-resistant plants. Plant Physiol 1984; 74:1016-1024.

10. Murata N, Ishizaki-Nishizawa O, Higashi, S. et al. Genetically engineered alteration in thechilling sensitivity of plants. Nature 1992; 356:313-326.

11. Wolter FP, Schmidt R, Heinz E. Chilling sensitivity of Arabidopsis thaliana with geneticallyengineered membrane lipids. EMBO J 1992; 11:4685-4692.

12. Moon BY, Higashi S-I, Gombos Z, et al. Unsaturation of the membrane lipids of chloro-plasts stabilizes the photosynthetic machinery against low-temperature photoinhibitionin transgenic tobacco plants. Proc Natl Acad Sci USA 1995; 92:6219-6223.

13. Aro E-M, McCaffery S, Anderson JM. Recovery from photoinhibition in peas (Pisumsativum l.) acclimated to varying growth irradiances. Role of D1 protein turnover. PlantPhysiol 1994; 104:1033-1041.

14. Aro E-M, Virgin I, Andersson B. Photoinhibition of photosystem II-inactivation, proteindamage and turnover. Biochim Biophys Acta 1993; 1143:113-134.

15. Ishizaki-Nishizawa O, Fujii T, Azuma M, et al. Low-temperature resistance of higher plantsis significantly enhanced by a nonspecific cyanobacterial desaturase. Nature Biotech 1996;14:1003-1006.

16. Kodama H, Hamada T, Horiguchi G et al. Genetic enhancement of cold tolerance by expressionof a gene for chloroplast (∆−3 fatty acid desaturase in transgenic tobacco. Plant Physiol1994; 105:601-605.

17. Wu J, Browse J. Elevated levels of high-melting-point phosphatidylglycerols do not inducechilling sensitivity in a mutant of Arabidopsis. The Plant Cell 1995; 7:17-27.

18. Murata N. Molecular species composition of phosphatidylglycerols from chilling-sensitiveand chilling-resistant plants. Plant Cell Physiol 1983; 24:81-86.


19. Wu JW, Lightner J, Warwick N et al. Low-temperature damage and subsequent recoveryof fab1 mutant Arabidopsis exposed to 2˚C. Plant Physiol 1997; 113:347-356.

20. Gupta AS, Heinen JL, Holaday AS et al. Increased resistance to oxidative stress in transgenicplants that overexpress chloroplastic Cu/Zn superoxide dismutase. Proc Natl Acad Sci USA1993; 90:1629-1633.

21. Kanervo E, Aro E-M, Murata N. Low unsaturation level of thylakoid membrane lipidslimits turnover of the D1 protein of photosystem II at high irradiance. FEBS Lett 1995;364:239-242.

22. Sakamoto T, Bryant DA. Growth at low temperature causes nitrogen limitation in thecyanobacterium synechococcus sp. PCC 7002. Arch Microbiol 1998; 169:10-19.

23. Nishida I, Murata N. Chilling sensitivity in plants and cyanobacteria. The crucial contri-bution of membrane lipids. Annu Rev Plant Physiol Plant Mol Biol 1996; 47:541-568.

24. Martino-Catt S, Ort DR. Low temperature interrupts circadian regulation of transcriptionalactivity in chilling-sensitive plants. Proc Natl Acad Sci USA 1992; 89:3731-3735.

25. Prasad TK, Anderson MD, Martin BA et al. Evidence for chilling-induced oxidative stressin maize seedlings and a regulatory role for hydrogen peroxide. Plant Cell 1994; 6:65-74.

26. Prasad TK. Mechanisms of chilling-induced oxidative stress injury and tolerance in devel-oping maize seedlings: Changes in antioxidant system, oxidation of proteins and lipids,and protease activities. Plant J 1996; 10:1017-1026.

27. Hugly S, McCourt P, Browse J et al. A chilling sensitive mutant of Arabidopsis with alteredsteryl-ester metabolism. Plant Physiol 1990; 93:1053-1062.

28. Schneider JC, Hugly S, Somerville CR. Chilling sensitive mutants of Arabidopsis. PlantMol Biol Rep 1995; 13:11-17.

29. Schneider JC, Nielsen E, Somerville C. A chilling-sensitive mutant of Arabidopsis is deficientin chloroplast protein accumulation at low temperature. Plant Cell Environ 1995; 18:23-31.

30. Patterson GW, Hugly S, Harrison D. Sterols and phytyl esters of Arabidopsis thaliana undernormal and chilling temperatures. Phytochemistry 1993; 33:1381-1383.

31. Browse J, Somerville C. Glycerolipids. In: Meyerowitz E, Somerville C, eds. Arabidopsis.New York: Cold Spring Harbor Press, 1994:881-912.

32. Miquel M, James D, Dooner H et al. Arabidopsis requires polyunsaturated lipids for lowtemperature survival. Proc Natl Acad Sci USA 1993; 90:6208-6212.

33. Hugly S, Somerville C. A role for membrane lipid polyunsaturation in chloroplast biogenesisat low temperature. Plant Physiol 1992; 99:197-202.

34. Tokuhisa JG, Feldmann KA, LaBrie ST et al. Mutational analysis of chilling tolerance inplants. Plant Cell and Environment 1997; 20:1391-1400.

35. Moir D, Botstein D. Determination of the order of gene function in the yeast nucleardivision pathway using cs and ts mutants. Genetics 1982; 100:565-577.

36. Botstein D, Maurer R. Genetic approaches to the analysis of microbial development. AnnRev Genet 1982; 16:61-83.

37. Feldmann KA. T-DNA insertion mutagenesis in Arabidopsis: Mutational spectrum. Plant J1991; 1:71-82.

38. Tokuhisa JG, Vijayan P, Feldmann KA et al. A homolog of DIM1, a yeast gene encoding aribosomal RNA dimethylase is essential for chloroplast development at low temperatures.Plant Cell 1998; in press.

39. Levitt J. Responses of Plants to Environmental Stress. Chilling, Freezing, and High Tem-perature Stresses. 2nd ed. New York:Academic Press, 1980.

40. Brush AR, Griffith M, Mlynarz A. Characterization and quantification of intrinsic icenucleators in winter rye (Secale cereale) leaves. Plant Physiol 1994; 104:725-735.

41. Olien CR. Analysis of midwinter freezing stress. In: Olien CR, Smith MN eds. Analysis andImprovement of Plant Cold Hardiness: Boca Raton:CRC Press Inc, 1981:61-87.

42. Steponkus PL. Role of the plasma membrane in freezing injury and cold acclimation. AnnuRev Plant Physiol 1984; 35:543-584.

43. Guy CL. Cold acclimation and freezing stress tolerance: role of protein metabolism. AnnuRev Plant Physiol Plant Mol Biol 1990; 41:187-223.


45. Steponkus PL, Uemura M, Webb MS. A contrast of the cryostability of the plasmamembrane of winter rye and spring oat. Two species that widely differ in their freezingtolerance and plasma membrane lipid composition. In: Steponkus, PL ed. Advances inLow-Temperature Biology. Volume 2. London:JAI Press Ltd, 1993:211-312.

46. Uemura M, Steponkus PL. Effect of cold acclimation on membrane lipid compositionand freezing-induced membrane destabilization. In: Li PH, Chen THH, eds. Plant ColdHardiness. New York:Plenum Press, 1997:171-179.

47. Strauss G and Hauser H. Stabilization of lipid bilayer vesicles by sucrose during freezing.Proc Natl Acad Sci USA 1986; 83:2422-2426.

48. Anchordoguy TJ, Rudolph AS, Carpenter JF et al. Modes of interaction of cryoprotectantswith membrane phospholipids during freezing. Cryobiology 1987; 24:324-331.

49. Neven, LG, Haskell DW, Guy CL et al. Association of 70-kilodalton heat-shock cognateproteins with acclimation to cold. Plant Physiol 1992; 99:1362-1369.

50. Anderson JV, Li Q-B, Haskell DW et al. Structural organization of the spinach endoplasmicreticulum-luminal 70-kilodalton heat-shock cognate gene and expression of 70-kilodaltonheat-shock genes during cold acclimation. Plant Physiol 1994; 104:1359-1370.

51. Caban ÈM, Calver P, Vincens P et al. Characterization of chilling-acclimation-relatedproteins in soybean and identification of one as a member of the heat shock protein(HSP70) family. Planta 1993; 190:346-353.

52. Krishna P, Sacco M, Cherutti JF, Hill S. Cold-induced accumulation of hsp90 transcriptsin Brassica napus. Plant Physiol 1995; 107:915-923.

53. Olien CR, Clark JL. Freeze-induced changes in carbohydrates associated with hardiness ofbarley and rye. Crop Sci 1995; 35:496-502.

54. Duman JG. Purification and characterization of a thermal hysteresis protein from a plant,the bittersweet nightshade Solanum dulcamara. Biochim Biophys Acta 1994; 1206:129-135.

55. Griffith M, Antikainen M, Hon W-C et al. Antifreeze proteins in winter rye. PhysiolPlant 1997; 100:327-332.

56. Antikainen M, Griffith M. Antifreeze protein accumulation in freezing-tolerant cereals.Physiol Plant 1997; 99:423-432.

57. Guy CL, Niemi KJ, Brambl R. Altered gene expression during cold acclimation of spinach.Proc Natl Acad Sci USA 1985; 82:3673-3677.

58. Thomashow MF. Arabidopsis thaliana as a model for studying mechanisms of plant coldtolerance. In: Meyerowitz E, Somerville C, eds Arabidopsis. New York:Cold Spring HarborLaboratory Press, 1994:807-834.

59. Hughes MA, Dunn MA. The molecular biology of plant acclimation to low temperature. JExpt Bot 1996; 47:291-305.

60. Gibson S, Arondel V, Iba K et al. Cloning of a temperature-regulated gene encodinga chloroplast omega-3 desaturase from Arabidopsis thaliana. Plant Physiol 1994;106:1615-1621.

61. White AJ, Dunn MA, Brown K et al. Comparative analysis of genomic sequence andexpression of a lipid transfer protein gene family in winter barley. J Experimental Botany1994; 45:1885-1892.

62. Jonak C, Kiegerl S, Ligterink W et al. Stress signaling in plants: A mitogen-activatedprotein kinase pathway is activated by cold and drought. Proc Natl Acad Sci USA 1996;93:11274-11279.

63. Mizoguchi T, Irie K, Hirayama T et al. A gene encoding a mitogen-activated protein kinaseis induced simultaneously with genes for a mitogen-activated protein kinase and an S6ribosomal protein kinase by touch, cold, and water stress in Arabidopsis thaliana. ProcNatl Acad Sci USA 1996; 93:765-769.

64. Monroy AF, Dhindsa RS. Low-temperature signal transduction: Induction of cold accli-mation-specific genes of alfalfa by calcium at 25˚C. The Plant Cell 1995; 7:321-331.

65. Tahtiharju S, Sangwan V, Monroy AF et al. The induction of kin genes in cold-acclimatingArabidopsis thaliana. Evidence of a role for calcium. Planta 1997; 203:442-447.


66. Jarillo JA, Capel J, Leyva A et al. Two related low-temperature-inducible genes of Arabidopsisencode proteins showing high homology to 14-4-3 proteins, a family of putative kinase regu-lators. Plant Mol Biol 1994; 25:693-704.

67. Thomashow MF. Role of cold-responsive genes in plant freezing tolerance. Plant Physiol1998; in press.

68. Dure III, L. Structural motifs in Lea proteins. In: Close TL, Bray EA, eds. Plant Responsesto Cellular Dehydration During Environmental Stress. Rockville:American Society of PlantPhysiologists, 1993:91-103.

69. Close, TJ. Dehydrins: A commonality in the response of plants to dehydration and lowtemperature. Physiologia Plantarum 1997; 100:291-296.

70. Ingram J, Bartels D. The molecular basis of dehydration tolerance in plants. Annu RevPlant Physiol Plant Mol Biol 1996; 47:377-403.

71. Artus NN, Uemura M, Steponkus PL et al. Constitutive expression of the cold-regulatedArabidopsis thaliana COR15a gene affects both chloroplast and protoplast freezing tolerance.Proc Natl Acad Sci USA 1996; 93:13404-13409.

72. Jaglo-Ottosen KR, Gilmour SJ, Zarka DG et al. Arabidopsis CBF1 overexpression inducesCOR genes and enhances freezing tolerance. Science 1998; 280:104-106.

73. Lin C, Thomashow MF. DNA sequence analysis of a cDNA for cold-regulated Arabidopsisgene cor15 and characterization of the COR15 polypeptide. Plant Physiol 1992; 99:519-525.

74. Gilmour and Thomashow, unpublished data.75. Webb MS, Gilmour SJ, Thomashow MF et al. Effects of COR6.6 and COR15am polypep-

tides encoded by COR (cold-regulated) genes of Arabidopsis thaliana on dehydration-induced phase transitions of phospholipid membranes. Plant Physiol 1996; 111:301-312.

76. Uemura M, Gilmour SJ, Thomashow MF et al. Effects of COR6.6 and COR15am polypep-tides encoded by COR (cold-regulated) genes of Arabidopsis thaliana on the freeze-induced fusion and leakage of liposomes. Plant Physiol 1996; 111:313-327.

77. Thomashow MF. Molecular genetics of cold acclimation in higher plants. Adv Genet 1990;28:99-131.

78. Stockinger EJ, Gilmour SJ, Thomashow MF. Arabidopsis thaliana CBF1 encodes an AP2domain-containing transcriptional activator that binds to the C-repeat/DRE, a cis-actingDNA regulatory element that stimulates transcription in response to low temperature andwater deficit. Proc Natl Acad Sci USA 1997; 94:1035-1040.

79. Yamaguchi-Shinozaki K, Shinozaki K. A novel cis-acting element in an Arabidopsis gene isinvolved in responsiveness to drought, low-temperature, or high-salt stress. Plant Cell 1994;6:251-264.

80. Volger HG, Heber U. Cryoprotective leaf proteins. Biochim Biophys Acta 1975; 412:335-349.81. Hincha DK, Heber U, Schmitt JM. Proteins from frost-hardy leaves protect thylakoids

against mechanical freeze-thaw damage in vitro. Planta 1990; 180:416-419.82. Hincha DK, Schmitt JM. Freeze-thaw injury and cryoprotection of thylakoid membranes. In:

Somero G, Osmond B eds. Water and life: Comparative Analysis of Water Relationships atthe Organismic, Cellular and Molecular Level. Berlin:Springer Verlag, 1992:316-337.

83. Sieg F, Schroder W, Schmidt JM et al. Purification and characterization of a cryoprotectiveprotein (cryoprotectin) from the leaves of cold-acclimated cabbage. Plant Physiol 1996;111:215-221.

84. Hong B, Uknes SJ, Ho T-HD. Cloning and characterization of a cDNA encoding a mRNArapidly induced by ABA in barley aleurone layers. Plant Mol Biol 1988; 11:495-506

85. Xu D, Duan X, Wang B et al. Expression of a late embryogenesis abundant protein gene,HVA1, from barley confers tolerance to water deficit and salt stress in transgenic rice. PlantPhysiol 1996; 110:249-257.

86. Imai R, Chang L, Ohta A et al. A lea-class gene of tomato confers salt and freezing tolerancewhen expressed in Saccharomyces cerevisiae. Gene 1996; 170:243-248.

87. Cohen A, Plant AL, Moses MS et al. Organ-specific and environmentally regulatedexpression of two abscisic acid-induced genes of tomato. Plant Physiol 1991; 97:1367-1374.

88. Warren G, McKown R, Marin A et al. Isolation of mutations affecting the development offreezing tolerance in Arabidopsis thaliana (L.) Heynh. Plant Physiol 1996; 111:1011-1019.


89. McKown R, Kuroki G, Warren G. Cold responses of Arabidopsis mutants impaired in freezingtolerance. J Expt Bot 1996; 47:1919-1925.

90. Xin Z, Browse, J eskimo1 mutants of Arabidopsis are constitutively freezing-tolerant. Proc NatlAcad Sci USA 1998; 95:7799-7804.

91. Rudolph AS, and Crowe JH. Membrane stabilization during freezing: The role of two naturalcryoprotectants, trehalose and proline. Cryobiology 1985; 22:367-377.

92. Metz AM, Timmer RT, Allen ML, Browning KS. Sequence of a cDNA encoding thealpha-subunit of wheat translation elongation factor 1. Gene 1992; 120:315-316.

CHAPTER 5


Molecular Responses to Heat StressFritz Schöffl, Ralf Prändl and Andreas Reindl

The heat shock response, defined as a transient reprogramming of gene expression, is aconserved reaction of cells and organisms to elevated temperatures (heat stress). The

features of this response are the induction of heat shock protein (HSP) synthesis and,physiologically, the acquisition of a higher level of thermotolerance. In plant speciesincluding soybean the synthesis of HSP is transient and accompanied by a cessation of normalprotein synthesis.1 During heat stress HSP seem to accumulate in a dosage-dependent manner,sufficient to protect cells from severe damage and to allow resumption of normal cellularand physiological activites. The transient synthesis of HSP suggests that the signal which istriggering the reponse is either lost, inactivated or no longer recognized under conditions oflong term heat treatment.

The temperature for the induction of the heat shock response (hyperthermic treatment)in plants is related to the optimum growth temperature of species and varieties and is usually5˚-10˚C above normothermic conditions. Plants evolved in low temperature environmentsrespond to much lower heat shock temperatures than, for example, desert plants.2,3 Thedifferences in temperature set points for the heat shock response indicate that the cells arenot sensing the absolute temperature. Instead, an internal stress signal, generated at differenttemperatures in different species and varieties seems to trigger the response. As exemplifiedfor the soybean, the heat shock response can be induced by different temperature regimes:2,4

1. Treatment at the maximum heat shock temperature (endured by the unconditionedcell for several hours),

2. Short heat pulse (applied for several minutes only) at an otherwise lethal temperature, and3. Also, by slowly, stepwise increasing temperatures.Severe heat stress leads to cellular damage and cell death. The thermal death point or

killing temperature of plants is highly dependent on the time of exposure. At very hightemperatures, death occurs within minutes and can be attributed to a catastrophic collapse ofcellular organization. Attempts have been made to discriminate between rapid direct (proteindenaturation and aggregation, increased fluidity of lipids in membranes) and slower indirecttypes of heat injuries (inactivation of enzymes in chloroplasts and mitochondria, block inprotein synthesis, degradation of proteins and membranes) that eventually lead to starvation,inhibition of growth, generation of toxic compounds, Reactive oxygen species, ion efflux,etc.5 However, the crucial point always remaining is the sensitivity of proteins andenzymes to heat inactivation and denaturation. Hence, adaptive mechanisms that protectcells from the proteotoxic effects of heat stress should be the key in acquisition ofthermotolerance.

The term tolerance is defined as the ability of cells and organisms to endure aninternal stress induced by an externally applied stressor.6 Thermotolerance refers to the


ability to cope with extreme temperatures; however, it is almost exclusively used fortolerance to heat, not for cold stress tolerance. Basal tolerance refers to the ability tosurvive a certain dose of heat stress without being conditioned by prior heat treatments.Acquired thermotolerance refers to an enhanced level of tolerance (significantly higherthan basal tolerance) induced by the application of a relatively mild or graduallyincreasing heat treatment. All plants tested to date are capable of acclimating or hardeningtowards heat stress. It has been shown that heat hardening of plant cells is effective onlyunder conditions that also induce the synthesis of HSP.7 This acquisition of a higherlevel of thermotolerance protects the cells and organisms from a subsequent, otherwiselethal heat stress.8-10

The heat shock response is of great interest for studying the molecular mechanisms ofstress tolerance and regulation of gene expression. In this paper we summarize the workon the characterization of the heat shock response with respect to the molecular functionof HSP in thermotolerance and general stress tolerance, the expression of heat shock genes,and the regulation of the heat shock transcription factor HSF.

Heat Shock Proteins and ThermotoleranceSeveral classes of HSP have been defined in eukaryotes, including plants. Nomenclature

refers to the approximate molecular mass which is characteristic for each group (HSP100,HSP90, HSP70, HSP60, HSP20). Synonymous designations are according to the designationsof homologues in prokaryotes (see Table 5.1).

HSP are functionally linked to the large and diverse group of molecular chaperoneswhich are defined by their capacity to recognize and to bind substrate proteins that are inan unstable, inactive state. It becomes evident that probably all cellular proteins have tointeract with molecular chaperones at least once in their lifetime, either during synthesis,subcellular targeting, or degradation. Owing to heat denaturation, the fraction of potentialtargets for molecular chaperones seems to dramatically increase upon heat stress andconsequently the cellular chaperone pool has to be replenished. It is not surprising thatexcept for HSP20 and HSP100 each class of HSP is matched by one or several otherconstitutive or cognate proteins (HSC) expressed at normal temperatures or respectivelynon-stress conditions. Different HSP may have different functional properties, but common toalmost all of them is their capacity to interact with other proteins and to act as molecularchaperones (for review see ref. 11). General and plant-specific characteristics of the differentclasses of HSP are briefly described; a broader overview is given by Boston et al.12

HSP100HSP100 belongs to the larger class of Clp proteins and is most similar to the ClpB

subgroup, a family of chaperones with diverged structures. The proteins of the HSP100/Clp family are crucial for protein disassembly of aggregated and higher order proteinstructures. The chaperone activity of certain members of this group assists in proteolyticpathways, and the ATPase activity of HSP100 is essential for this function. Heat-inducibleplant HSP100 genes cloned from soybean and Arabidopsis were able to functionallyreplace the homologous yeast HSP100 in the development of thermotolerance.13,14 In yeast,HSP100 was shown to be important also for the restoration of heat-inactivated splicingcomplexes.15 Of great interest and perhaps also important to plants is the implementationof a regulatory role of HSP100 in controlling the extrachromosomal inheritance of theprion-like protein PSI in yeast.15

83Molecular Responses to Heat Stress

Table 5.1 General Functional Properties of HSP Families

HSP familya Properties

HSP100 Homologous to ClpB , ATP-dependent disassembly of higherorder protein structures, important for thermotolerance in yeast

HSP90 Chaperone with ATP binding site, association with proteins involvedin signal transduction

HSP70 Homologous to DnaK, ATP-dependent chaperone, negative regulatorof the heat shock response, isoforms involved in protein transport

HSP60 GroEL/GroES homologs, ATP-dependent chaperonin, not typicallyhs-induced, isoforms in mitochondria and chloroplasts(Cpn60/Cpn10), cytoplasmic TCP-1

HSP20b Prevalent in plants, aggregation into homooligomeric complexes inthe cytoplasm, formation of heat shock granules, ATP-independentmolecular chaperone in vitro

aclassified by molar mass in kDa, different sizes in many organisms, structural and functionalconservation within families

bHSP-families in plants, only one HSP in yeast and man, structural relationship with the lenscrystallines of the vertebrate eye

HSP90The members of the HSP90 family, like HSP70 proteins, are highly conserved.

Constitutive and heat-regulated expression has been reported for HSP90 proteins. In yeastat least one HSP90 is required for viability, and biochemical evidence supports a role ofHSP90 in preventing thermal denaturation and aggregation of protein substrates in vitro.16

Except for studies about the expression of Hsp90 genes, nothing is known about biochemical,functional, or genetic properties of proteins of this group in plants. The assembly ofglucocorticoid receptor/HSP90 complexes in wheat germ extracts17 suggests that plantHSP90 may be also involved in signal transduction as shown for animal and human cells,by forming complexes with, for example, steroid receptors and kinases.18

HSP70The HSP70 family is probably the most highly conserved protein/gene family in

nature, with about 50% identical amino acid residues between the E. coli DnaK and thehomologous HSP70 proteins in eukaryotes.19 There is genetic evidence for an essentialfunction of HSC70 in yeast. Members of the chaperone 70 family are involved in translocationcompetence of precursor proteins, uncoating of coated vesicles, and association withribosomes in the cytosol; others are the driving force for protein uptake in mitochondriaand the ER (for review see Rassow et al 20) and in translocation of proteins into chloroplasts(for review see Boston et al12). There is structural/biochemical evidence that plant HSP70/HSC70 genes share the conserved domains for an N-terminally located ATPase activity


and a C-terminally located peptide-binding site. Current models propose that HSP70/HSC70 bind to nascent or partially denatured polypeptides, thereby preventing improperfolding and keeping proteins in a translocation competent and/or refoldable conformation.ATP binding and hydrolysis seem to modulate substrate binding. Cochaperones homologousto DnaJ and GrpE of E. coli accelerate ATP-hydrolysis and stimulate ATP-exchange,respectively. Numerous genes of the HSP70/HSC70 family have been identified in plants andit is typical that single species encode multiple genes for cytosolic and organelle-targetedHSP70-chaperones.12 The multiplicity of HSC70 genes/proteins is partially explained by themultiple localizations of HSP70-like proteins in different compartments (cytoplasm, chloro-plasts, mitochondria, ER). The genetic redundancy of cytosolic variants of HSC70 is notyet understood; multiple forms of this conserved chaperone may reflect the requirementfor differences in biochemical function of these proteins. Identifying the in vivo targets ofHSP70/HSC70 chaperones and a mutant analysis will be a prerequisite to the elucidationof functional properties.

HSP60Chaperonins which are homologous to the bacterial GroEL/GroES proteins are not

typically heat-induced stress proteins. The chloroplast-localized Cpn60/Cpn10 are nuclearencoded proteins and are imported into chloroplasts (for review see Boston et al12). Bothsubunit proteins are expressed in the absence of stress and moderately increased uponheat shock. Ch-Cpn60 forms a cylindrically shaped double-stacked ring of two heptamerswith intrinsic ATPase activity. In contrast to the bacterial GroEL complex, ch-Cpn60appears to be a hetero-oligomer consisting of stoichiometric amounts of α- and β-subunits.The chloroplast co-chaperonin Cpn10, compared to its bacterial counterpart GroES, is anobviously head-to-tail duplicate form of approximately 21 kDa. GroEL/GroES complexesfacilitate folding, assembly, and translocation of numerous other proteins, as exemplifiedby the Rubisco complex in chloroplasts. However, in plants the exact mechanism is stillobscure. It is known that GroEL oscillates between states of high and low affinity fornon-native proteins, release requires binding of adenine nucleotides, and that theGroEL-assisted folding reaction is strictly dependent on ATP-hydrolysis and participationof co-chaperone GroES. GroEL and GroES-like proteins are also found in plant mitochondria.The TCP-1 protein of the cytosol is only distantly related to GroEL but probably performsanalogous functions.

HSP20HSP20 proteins, also termed small (sHSP) or low molecular weight (lmw) HSP, are a

complex group of prevalent HSP in plants. Characteristic to plants is also the occurrenceof members of the HSP20 group in chloroplasts, mitochondria, and in the ER (for reviewsee Boston et al12). Recently a tomato chloroplast sHSP was found to be imported intochromoplasts, indicating a function for this HSP in fruit ripening.21 Meanwhile, sHSPcomprise 6 gene families for strictly heat-inducible HSP ranging between 17 and 30 kDa. HSPof two families (class I and II) are targeted to the cytoplasm. It should be noted that to dateonly in higher plants were sHSP also found in the endomembrane system. The conservation ofamino acid residues within classes is 80-90%, but only 30% between different classes.However, all sHSP share a common structural domain, a feature for all sHSP and theα-crystalline, in the C-terminal part of the molecule. Small HSP tend to form homooligomericcomplexes of 200 to 800 kDa in vitro.22,23 Studies using recombinant proteins in vitrosuggest that sHSP act as chaperones preventing heat-induced aggregation and promoterenaturation of model substrates.24 Interestingly, this chaperone activity acts insubstoichiometric amounts and, in contrast to HSP70 and HSP60 chaperones, does not


require stimulation by nucleotides. Although the in vivo targets of sHSP chaperonecomplexes are not known, a model suggests that sHSP capture unfolded polypeptides byhydrophobic interactions and maintain them in a state competent for refolding the nativestate by other chaperones, or likewise for degradation by the proteolytic systems.

At the cellular level, one consequence of the heat shock response is the appearance ofgranular structures containing class I-sHSP located throughout the cytoplasm and thenucleus in soybean.25 Heat shock granules may be formed from different HSP; their exactcomposition in vivo is unknown. Dependent on the heat stress regimes applied, granulesmay form higher order structures of aggregation.26

Mutants Affecting Expression of HSP and ThermotoleranceThere is a striking correlation between the occurrence of HSP and acquisition of

thermotolerance, but there is only little direct evidence proving a causal relationship.Mutagenesis and genetic engineering strategies have been proposed for the analysis ofregulation and for the manipulation of the heat shock response.27,28 Mutations in regulatorygenes resulting in a coordinate change in expression of HSP would be required for studying:

1. the signal pathway from stress to gene,2. the mechanism of transcriptional regulation and3. the role of HSP in thermotolerance.Using genetic engineering of Arabidopsis as a model for higher plants, dominant

mutations were generated showing a deregulated synthesis of HSP at normal temperature.41,42

Such transgenic plants exhibit a significantly higher level of basic thermotolerance. Thesedata demonstrate the fundamental role of HSP in thermotolerance, however, it is not yetclear whether, apart from HSP, other genes are also affected and involved in thermotolerancein transgenic lines expressing a derepressed HSF.

The effects of mutations in individual heat shock genes have been investigated indifferent organisms. Analyses in yeast provided evidence for an important role ofHSP10429 and a minor, accessory role of HSP7030 concerning thermotolerance. Mutationsin Hsp26, the sole gene for sHSP in yeast,31 or overexpression of sHSP and antisenseapproaches in transgenic plants32,33 had no obvious effect on the phenotype. It is possiblethat the protective effect of HSP is sometimes dependent on the physiological conditionsof the cell, as shown by the disruption of a mitochondrial HSP30 gene in Neurosporaresulting in strains that were less thermotolerant under certain carbohydrate limitations.34

In other eukaryotes, other groups of HSP seem to play important roles in thermotolerance,as for example shown by HSP70 overexpression in mammalian cells and in Drosophila.35-40

Other Heat-Induced ProteinsBesides the described groups of major HSP chaperones there is a number of plant

proteins/genes including ubiquitin,43,44 cytosolic Cu/Zn-superoxide dismutase45 andmanganese peroxidase,46 whose expression is also stimulated upon heat stress. The function ofthese proteins is related to protein degradation pathway and oxidative stress response, twoobviously very important processes to be maintained during heat stress or restored duringrecovery.

Links to Other Abiotic StressesExpression of HSP is also linked to several other environmental stresses and an

increasing number of studies show cross protection in plants. The results imply that HSPare important constituents of the molecular mechanism of common stress tolerance. Themost obvious links are between HSP and dehydration/drought, cold/chilling/freezing, heavymetal, and oxidative stress.


Dehydration/DroughtIn sunflower47 and the resurrection plant Craterostigma plantagineum48 certain sHSP

are expressed upon water stress/dehydration in vegetative tissue. Expression of sHSP wasidentified in a screen for early dehydration responsive genes in Arabidopis thaliana.49 HSPare also expressed in the absence of heat stress during late seed maturation/desiccationperiod in the Arabidopsis embryo.50

Cold/ChillingChilling injury is a physiological disorder that develops in some plants that are indigenous

to tropic and subtropic regions. As shown for tomato fruit, heat stress can protect againstchilling injury and the accumulation of HSP correlated with the persistence of chillingtolerance.51 A similar correlation was found also for chilling tolerance induced by heatstress in the mung bean hypocotyl52 and in germinating Cucumis sativus seeds.53 Cold-inducedtranscripts of HSP90 in Brassica napus, HSC70 in spinach and soybean, and sHSP inpotato, suggest that HSP seem to play a role in plant responses and acclimation to lowtemperatures.54-57

Heavy MetalIn soybean hypocotyl the most efficient non-heat shock inducers of heat shock gene

expression are arsenite and cadmium.58-60 Arsenite-treated cells synthesize HSP andacquire a certain level of thermotolerance.8 Heat shock protects against metal toxicity inwheat leaves, and cultured cells of Lycopsersicon peruvianum can be protected by heat stressfrom injuries inflicted by cadmium treatment.61,62 On the other hand, expression of HSPis also induced by heavy metal ions, which is in accordance with the concept of cross protectionvia HSP.63

Oxidative StressCell death in plants caused by abiotic stresses is frequently associated with symptoms

of oxidative stress. Mutant analyses of SOD, catalase, and cytochrome C oxidase genes inyeast provide evidence for an involvement of oxidative stress in heat-induced cell death;overexpression of these enzymes improved thermotolerance under anaerobic conditions.64

In human cells mitochondria were found to be selective targets for the effect of heat shockagainst oxidative stress, probably mediated by HSP70.65 In plants, heat shock protects PSIIfrom photoinhibition and it is speculated that a chloroplast-targeted sHSP is involved inthe protection of the D1 protein.63

On the Molecular Mechanism of Cross ProtectionHeat shock-induced cross tolerance and the occurrence of HSP induced by other abiotic

stresses suggest that HSP are important determinants of a common stress response in plants.If HSP have a generally protective function in abiotic stress response, what is the commontheme in the cellular damage by abiotic stresses? This is most likely the denaturation ofcellular proteins and/or the generation of Reactive oxygen species. Higher levels ofenzymes and enzymatic activities of the antioxidant pathway including superoxidedismutases (SOD), catalyses, ascobate peroxidase, glutathione reductase and others werefound in response to a number of environmental stresses (for overview see Foyer et al 67).Overexpression of SOD in tobacco,68,69 alfalfa,70 potato,71 and cotton72 has been found toenhance tolerance to oxidative stress and to some extent also to freezing, chilling injuryand water deficit.67,73

With the availability of mutants of the heat shock response, showing a synthesis ofHSP at lower non-heat shock temperatures,41,42 it will be possible to test whether under


environmental stress the chaperone function of HSP is involved in stabilization of enzymes(or enzymatic activities) of the antioxidant pathway.

Transcriptional RegulationThe expression of heat shock genes in plants is, similarly to the situation in other

eukaryotes, primarily regulated at the transcriptional level.74 The thermoinducibility isattributed to conserved cis-regulatory promoter elements (HSE) which are located in theTATA box proximal 5'-flanking regions of heat shock genes. The occurrence of multipleHSEs within a few hundred base pairs is a signature of most eukaryotic heat shock genes.The importance of HSE for heat-dependent transcriptional regulation in plants has beenverified by promoter deletions of a soybean heat shock gene75,76 and by the capacity ofsynthetic HSE sequences, integrated in a truncated CaMV-35S promoter, to stimulateheat-inducible gene expression in transgenic tobacco.76 The requirement of the TATA boxwas demonstrated by deletion analysis of a soybean heat shock gene in sunflower.77 Thesedata are in accordance with the protection pattern of HSE and TATA box-containingregions in footprinting experiments using a Drosophila HSP70 promoter.78

HSE-HSF InteractionThe eukaryotic HSE consensus sequence has been ultimately defined by Amin et al79

and Xiao and Lis80 as alternating units of 5'-nGAAn-3'. HSE are the binding sites for thetransactive heat shock transcription factor HSF; efficient binding requires at least threeunits resulting in 5'-nGAAnnTTCnnGAAn-3'.81,82 Chemical footprinting and interferenceexperiments show that the binding of HSF to HSE occurs in the major groove. The nGAAnbox is considered as the fundamental unit of HSF-binding, with each subunit of a HSFtrimer interacting with one nGAAn-unit.81 The stoichiometry for binding was directlydemonstrated by analytical ultracentrifugation.83 There is evidence for trimer formationof recombinant plant HSF following expression in E. coli and, for binding to both consensusHSE sequences84,85 and to the HSE-containing regions of an authentic Drosophila HSP70promoter.85 Transgenic expression provides evidence that HSF-HSE interaction andtranscriptional activation is highly conserved in nature, as demonstrated by the recognitionand proper regulation of a Drosophila HSP70 promoter in plants86 and by the activation ofthe same promoter in Drosophila cells via the transiently expressed HSF of Arabidopsis,AtHSF1.85

The promoter strength seems to depend on several different criteria, including thedegree of conservation and spacing of HSE sequences.87 Mismatches in a naturally occurringHSE2 consensus sequence of a Drosophila HSP70 gene result in only weak binding of HSF;however, in the presence of an adjacent full matching HSE1 the binding affinity to HSE2 isgreatly increased, indicating cooperative binding of trimeric HSF.88 The cooperativity inHSF binding may act as a supplementary mechanism for increasing the range of heat-inducible binding to DNA.89

Other Sequences Affecting Heat Shock Gene ExpressionA number of additional sequence motifs have been identified that have quantitative

effects on expression of certain heat shock genes. In plants, there is evidence for aninvolvement of CCAAT box90 and AT-rich sequences.75,91 One AT-rich sequence located inthe flanking region of a soybean heat shock gene exerted an enhancing effect on theexpression of heat shock promoter-reporter gene constructs in tobacco and was able tobind to the nuclear scaffolds.92 These data suggest that sequences affecting the chromatinstructure may be important for efficient access of transcription factors (e.g., TATA-boxbinding protein TBP) and/or the transcriptional activator proteins (e.g., HSF). The following


model integrates the current knowledge about the activation of heat shock geneexpression: The binding of a GAGA sequence binding factor93,94 or, likewise, scaffoldattachment affects chromatin structure in a way that provides TBP access to the TATA box,which is prerequisite for subsequent assembly of the basal transcription complex. In this“standby mode” heat shock genes are primed for transcriptional activation upon heat stress,mediated by the trimerization and binding of HSF to the HSE sequences.

Developmental ExpressionIn many organisms including plants, the expression of heat shock genes is not only

triggered by a number of environmental stresses but also by developmental cues. In plantsthe regulation of developmental expression of HSP, indicated by the occurrence of mRNAsand HSP in dry seeds, has not yet been investigated in greater detail. The analysis of adevelopmentally regulated soybean heat shock promoter in transgenic tobacco providesevidence for participation of HSE sequences and consequently suggests binding andinvolvement of HSF.95 It cannot be excluded that other sequences and transactive factorsare involved in seed specific expression of HSP. The control of this expression by thedevelopmental program rather than by dehydration is indicated by the negative effect ofthe abi3 mutation in Arabidopsis on seed-specific expression of sHSP.96 ABI3, originallyidentified as an abscisic acid-insensitive mutant allele in Arabidopsis, appears to have adominant regulatory effect on the developmental expression of heat shock genes in theembryo. Recent models for the action of Vp1,97 the structural/functional homologue ofABI3 in maize,98 suggest that Vp1 acts in stabilization/activation of regulatory complexesinvolved in the transcription of target genes. Further investigation of the activation of heatshock promoters during seed maturation will be required to test the hypothesis that ABI3,directly or via the action of secondary factors, is a regulator of HSF activity. It should benoted that in Drosophila, developmental regulation of certain heat shock genes, such as theexpression of HSP82 and HSP26 in oocytes and early larval stages, seems to be regulatedby steroid hormones and does not involve HSE-HSF interaction. However, the sole HSF ofDrosophila plays an essential role at this stage of development and this function appears tobe not directly related to the expression of HSP.99

The Regulation of HSFActivation of HSF of higher eukaryotes is a multi-step process. In response to heat

stress, HSF is converted from a monomeric to a trimeric form. Trimeric HSF is localizedpredominantly in the nucleus, binding to HSE with high affinity. The acquisition oftranscriptional competence is separately regulated; it can be uncoupled from HSE binding, asshown for human HSF1 and Drosophila HSF. HSF of Saccharomyces cerevisiae andKluyveromyces lactis is bound to heat shock promoters in the absence of stress and is thereforebelieved to be regulated primarily at the level of transactivating ability.89,100

HSF DomainsThe domains for DNA binding (DBD) and oligomerization (HR-A/B) are located in

the N-terminal region of HSF (Fig. 5.1). Both domains are conserved in primary structurethroughout the HSF protein family. Other regions show significant homology onlybetween closely related HSF. Nuclear localization signals (NLS), hydrophobic heptadrepeats localized in the C-terminal region (HR-C), and activation domains (AD) havebeen identified by functional studies in several HSF.89,100

Similar to vertebrates, all plant species investigated so far contain multiple HSF incontrast to single HSF genes reported for yeast and Drosophila. To date, three HSF havebeen described from Arabidopsis thaliana, six from soybean (Glycine max.), three from


tomato (Lycopersicon peruvianum), and three from maize (Zea maize).42,101-105 Molecularmasses of plant HSF are in the range from 31.2 to 57.5 kD. Based on sequence homologyand domain structure, plant HSF can be subdivided in the two classes A and B.101

DNA-Binding DomainHSF carry a conserved DBD consisting of an antiparallel four stranded β-sheet packed

against a bundle of three α-helices, as determined for HSF from Kluyveromyces lactis,Drosophila, and tomato.106-108 The second and the third helix form a typical helix-turn-helixmotif. The third helix is responsible for establishing the specific nucleic acid contacts withthe HSEs. A distinguishing feature between non-plant and plant HSF is an 11 amino aciddeletion between two β-sheets in plant HSF. The significance of the lack of this probablysolvent-exposed loop is unknown.101

Oligomerization DomainThe HR-A/B is separated from the DBD by a linker of variable length and sequence

and comprises the two regions A and B. Region A is based on heptad repeats of hydrophobicamino acids, whereas region B is composed of two overlapping heptad repeats. In class Aplant HSF, these arrays are separated by additional heptad repeats brought about by theinsertion of 21 amino acids. Plant HSF of class B lack this insertion. It is assumed that thefunction of the HR-A/B region is to allow homotrimer formation through a triple-stranded,α-helical coiled-coil structure.

Obviously, the formation of trimeric HSF in higher eukaryotes requires heat stress,but how is the suppression of HSF trimerization achieved under non-stress conditions?The C-terminal heptad repeats of hydrophobic residues (HR-C), which are well conservedin animal HSF but only poorly in plant and yeast HSF, is involved in the regulation oftrimerization. Mutations in the HR-C region lead to HSF with constitutive trimerizationand DNA-binding competence, as shown for Drosophila HSF, chicken HSF1 and HSF3,and human HSF1.110-112 A model proposes that intramolecular coiled-coil interactionsbetween the HR-A/B and HR-C hydrophobic heptad repeats suppress trimer formationunder normal growth conditions. However, deletion mapping of Drosophila HSF revealedlarger portions of HSF involved in the negative control of trimer formation.113

Nuclear Localization SignalHSF carry two clusters of basic amino acids that have been proposed to function as

NLS. A highly conserved cluster of basic amino acids is located at the C-terminus of theDBD, and a second cluster resides C-terminal of the HR-A/B.89,100,101 In functional studieswith two class A tomato HSF, the more C-terminal NLS has been found to be exclusivelyrequired for nuclear import.104 Again, this is a discrepancy with vertebrate HSF, which

DBD HR-AB NLS HR-C AD

Fig. 5.1. Schematic domain structure of a prototypic HSF. The DNA-binding domain (DBD), theoligomerization domain (HR-A/B), a nuclear localization signal (NLS), the C-terminal hydro-phobic heptad repeat (HR-C), and an activation domain (AD) are marked. The sizes of theindividual domains as well as the lengths of the linker sequences are not drawn to scale.


require either both or solely the more N-terminal NLS for translocation.115,116 By usingfusion proteins between the NLS of the Drosophila HSF and a β-galactosidase reporter, ithas been shown recently that the NLS is sufficient for stress-induced nuclear entry, supportingthe view that nuclear import is one layer of HSF regulation by stress.117

Activation DomainThe activation domains (AD) of HSF of higher eukaryotes are localized C-terminally,

whereas the HSF of Saccharomyces cerevisieae and Kluyveromyces lactis carry AD at the C- andthe N-terminus of the protein.89,100 The AD of human HSF1 and Drosophila HSF showlimited sequence identity and are rich in hydrophobic and acidic amino acids.118,119 Studiescarried out to characterize the AD of tomato HSF indicate the involvement of motifs whichconsist of aromatic, hydrophobic, and acidic amino acid residues.120,121

Animal HSF acquire transactivating competence as a final step during the activationprocess. Regulatory sequences repress the AD under non-stress conditions and render itheat-inducible. A central regulatory domain has been mapped between the HR-A/B andthe AD of human HSF1. The regulatory domain confers heat responsiveness even to theheterologous AD of VP16.118 In contrast to higher eukaryotes, the Saccharomyces cerevisiaeHSF is associated with the high affinity binding sites of heat shock promoters in theabsence of stress.122 HSF in yeast is assumed to be regulated primarily at the level oftransactivating competence. An amino acid in the DBD, the HR-B, and a yeast-specificcontrol element (CE2) have been shown to be involved in the repression of the AD undernon-stress conditions.123,124

Regulators of HSF Activity

Negative Regulation of HSF by HSP70There is genetic evidence for an autoregulation of the heat shock response in Escherichia

coli, yeast, and higher eukaryotes.89,100 In Saccharomyces cerevisae mutations in twoconstitutively expressed HSC70/HSP70 genes activate a β-galactosidase reporter gene in aHSE-dependent manner in the absence of heat stress.125 The derived model of HSF regulationby chaperone titration proposes that the pool of free HSC70/HSP70 is deplenished duringheat shock due to binding of HSC70/HSP70 to unfolded proteins, thereby relieving therepression of HSC70/HSP70 on HSF. Derepression of the AD domain and increasedtrimer formation in higher eukaryotes activate HSF and consequently heat shock gene tran-scription. In a negative feedback loop, the synthesis of appropriate levels of HSP70 shutsoff HSF. With respect to trimer formation, HSC70/HSP70 may maintain HSF in monomericstate or may participate in the disassembly of trimeric HSF. Stoichiometric complexesbetween non-activated HSF1 and HSP70 have been described, as well as an inhibition of heatactivation of HSF1 in mammalian cells which transiently overexpress HSP70.126

Recently, genetic evidence for a negative regulation of Arabidopsis HSF1 under non-stressconditions has been obtained. Arabidopsis HSF1 is repressed under non-stress conditionsand trimerizes upon heat shock. A heat stress-independent derepression of ArabidopsisHSF1 was obtained by constitutive overexpression of HSF1 fusion proteins with aβ-glucuronidase reporter.41 The molecular mechanism of derepression is still unknownbut seems not only restricted to glucuronidase fusions of HSF. The conformation of thefusion protein may be either inaccessible to a negative regulatory molecule or overexpressionof this protein titrates a transacting negative regulator. Interestingly, overexpression ofanother Arabidopsis HSF, HSF3, appears to be sufficient for derepression of the heat shockresponse in transgenic Arabidopsis.


Arabidopsis HSF1 shows also a constitutive DNA binding upon heterologous expressionin Drosophila and human cells. Furthermore, a derepressed transactivating competencehas been demonstrated for Arabidopsis HSF1 in Drosophila cells.102 Thus, the negativecontrol seems to depend on a factor which is absent in the cultured animal cells. Directevidence for HSP70 as a negative regulator of HSF in Arabidopsis comes from the analysisof transgenic Arabidopsis plants carrying an HSP70 antisense gene. As a result of expression ofthe antisense RNA, endogenous HSC70/HSP70 levels are reduced, and during therecovery from heat shock HSF1 trimers are significantly longer present than in controlplants.128

Negative Regulation of HSF by PhosphorylationPhosphorylation has been proposed to play a role in activation and inactivation of

HSF.89,100 However, recent functional studies suggest that phosphorylation is primarilyinvolved in repression of HSF. In yeast, phosphorylation of CE2-adjacent serine residueshas been shown to enhance deactivation of HSF after heat shock.129 The major phos-phorylation sites of human HSF1 in cell culture at control temperature have been localized totwo serines in the regulatory domain, which modulates the AD. Mutational conversion ofthese serines to alanine residues leads to constitutively active HSF, whereas conversion toglutamic acid, mimicking a phosphorylated serine, represses HSF in the absence of stress.Phosphorylation of the serine residues is increased upon stimulation of the Raf/ERK pathway,which is a mitogen-activated protein kinase pathway responsive to growth factors.130,131

Recently, HSF phosphorylation has been reported for plants. A kinase activity has beendetected in extracts of Arabidopsis suspension culture cells which phosphorylates HSF1 atserine residues and consequently decreases the HSE-binding of HSF1. Immunologicalcharacterization has identified the kinase as CDC2a, a cyclin-dependent kinase regulatingthe cell cycle.132

Thus, in human cells as well as in Arabidopsis, phosphorylation of HSF through variouskinases may integrate growth signals. As yet it is unknown whether cyclin-dependentkinases are involved in HSF phosphorylation in animals or whether mitogen-activatedprotein kinases play a role in HSF regulation in plants. It is conceivable that in growingcells, subjected to moderate stress conditions, phosphorylation of HSF may be requiredfor a repression of the heat shock response, which might otherwise interfere with proliferation.Such a phosphorylation-dependent repression can be overridden under conditions of heatstress.129 On the other hand, HSF may have also essential functions during development.Drosophila HSF is indispensable for oogenesis and early larval development.99 Thus it willbe interesting to investigate whether kinases modulate the developmental activities of HSFand which pathway is involved in signaling that results in developmental control of HSPexpression in plants.

Conclusions and PerspectivesSome of the plant responses to heat stress show certain characteristics that are unique

to plants, originally discovered in plants or more important to plants than to other organisms.One such area is the biological role and molecular mechanism of HSP in thermotoleranceand common stress tolerance. Based on the results of genetic engineering of the entireheat shock response and thermotolerance via HSF,41,42 future research will focus on theroles of HSP100, HSP90, HSP70 and sHSP as the primary targets for identifying specificdeterminants involved in protection from deleterious effects of heat, cold, heavy metal,desiccation, reactive oxygen, and other stresses. In order to understand the underlyingmolecular mechanisms, it will be necessary to identify the cellular targets, enzymes, orstructural proteins to become associated with these HSP in their native form in protein


complexes. The problem of functional specificity of molecular chaperones is exemplifiedby the proteins of the chaperone 70 family, comprising the heat-induced cytosolic HSP70and several constitutive HSC70 proteins localized in different subcellular compartments.The important questions are: What is the specific target and functional role of HSP70? IsHSP70 and/or another protein of the 70 kDa stress protein family involved in the negativeregulation of HSF and consequently of the heat shock response?

The regulation of HSF activity and the multiplicity of HSF in plants are other problems ofgreat scientific interest. The mechanism of repression of HSF activity is still not understood.HSF1 protein fusions41 and HSF342 of Arabidopsis are active upon transgenic overexpression,suggesting that negative regulation and/or conformational changes are involved in themechanism of activation. Three HSF-like genes were identified in tomato,104 Arabidopsis,42,84

and maize105 and six in soybean.103 The question about the biological role of the geneticredundancy of HSF has to be addressed and answered by future research. It seems possiblethat some putative HSF, classified by the criterion of structural features in the DNAbinding and multimerization domains, may in fact act as DNA-binding-proteins, lacking,however, the capacity of transcriptional activation. Such proteins could act through DNAbinding, either as repressors or through protein-protein interaction as modulators of HSFactivity.

Is there a signal pathway that senses stress from external sources and triggers the heatshock response via HSF? Components in the pathway upstream from HSF are not yetknown. It is conceivable that HSF itself or its interaction with HSC70 and other proteins isthe sensor of heat stress and results in an activation of HSF via conformational changesinvolving monomer to trimer transition and nuclear targeting. An alternative model fortemperature sensing and regulation of the heat shock response integrates observed membranealterations. The lipid perturbation model is based on the effects resulting from changes inthe ratio of saturated: unsaturated fatty acids on the set point of temperature for the heatshock response in yeast and proposes that these alter HSF activity.133 Developmental signalingseems to be responsible for the expression of HSP during seed maturation. The involvement ofHSF is indicated by the dependence on HSE promoter sequences; signaling through ABApathways is suggested by the negative effect of an abi3 mutation in Arabidopsis.96 However,neither the responsible HSF nor the level of control by ABI3 have been identified.

Other areas of important research in the field of heat stress responses, not covered inthis review, concern changes in the translation machinery that result in a shut down ofnormal protein synthesis and preferential synthesis of HSP during heat stress. In a numberof new approaches, interesting results were gained about translational control for the firsttime in this field in plants.134-136

References1. Key JL, Lin CY, Chen YM. Heat shock proteins of higher plants. Proc Natl Acad Sci USA

1981; 78:3526-3530.2. Key JL, Kimpel J, Vierling E et al. Physiological and molecular analyses of heat shock

response in plants. In: Atkinson BG, Walden DB, eds. Changes in eukaryotic geneexpression in response to environmental stress. Orlando: Academic Press, 1985:327-348.

3. Kee SC, Nobel PS. Concomitant changes in high temperature tolerance and heat shockproteins in desert succulents. Plant Physiol 1986; 80:596-598.

4. Key JL Nagao RT, Czarnecka E et al. Heat stress: Expression and structure of heat shockproteins. In: Von Wettstein D, Chua NH ed. Plant Molecular Biology. New York:PlenumPubl Corp, 1987:385-397.

5. Levitt J. Responses of plants to envronmental stresses. Vol I: Chilling freezing and hightemperature stresses. 2nd ed. New York:Academic Press, 1980:27-64.


6. Street HE, Oepik H. Physiology of flowering plants: Their growth and development. 3rded., London: Edward Arnold, 1984:110-134

7. Wu MT, Wallner SJ. Heat stress responses in cultured plant cells. Plant Physiol 1984;75:778-780.

8. Lin CY, Roberts JR, Key JL. Acquisition of thermotolerance in soybean seedlings. PlantPhysiol 1984; 74:152-160.

9. Yarwood CE. Acquired tolerance in leaves to heat. Science 1961; 134:941-942.10. Yarwood CE. Adaptions of plants and plant pathogens to heat. In: Posser CL, ed.

Molecular mechanisms of temperature adaption, Publ. No. 84 Washington DC: AmerAssoc Advance Sci 1967:75-89.

11. Vierling E. The roles of heat shock proteins in plants. Ann Rev Plant Physiol Plant MolBiol 1991; 42:579-620.

12. Boston RS, Viitanen PV, Vierling E. Molecular chaperones and protein folding in plants.Plant Mol Biol 1996; 32:191-222.

13. Lee YR, Nagao RT, Key JL. A soybean 101-kD heat shock protein complements a yeastHSP104 deletion mutant in acquiring thermotolerance. Plant Cell 1994; 6:1889-1897.

14. Schirmer EC, Lindquist S, Vierling E. An Arabidopsis heat shock protein complements athermotolerance defect in yeast. Plant Cell 1994; 6:1899-1909.

15. Vogel JL, Parsell DA, Lindquist S. Heat shock proteins HSP104 and HSP70 reactivatemRNA splicing after heat inactivation. Curr Biol 1995; 306-317.

16. Jakob U, Buchner J. Assisting spontaniety: The role of HSP90 and small HSPS as molecularchaperones. Trends Biochem Sci 1994; 19:205-211.

17. Stancato LF, Hutchinson KA, Krishna P, Pratt WB. Animal and plant cell lysates share aconserved chaperone system that assembles the glucocorticoid receptor into a functionalheterocomplex with HSP90. Biochemistry 1996; 35:554-561.

18. Rutherford SL, Zucker CS. Protein folding and the regulation of signaling pathway. Cell1994; 79:1129-1132.

19. Boorstein WR, Ziegelfoffer T, Craig EA. Molecular evolution of the HSP70 gene family.J Mol Evol 1994; 38:1-17.

20. Rassow J, von Ahsen O, Böhmer U, Pfanner N. Molecular chaperones: Toward a character-ization of the heat shock protein 70 family. Trends Cell Biol 1997; 7:129-133.

21. Lawrence SD, Cline K, Moore GA. Chloroplast development on ripening tomato fruit:Identification of cDNAs for chromoplast targeted proteins and characterization of a cDNAencoding a plastid localized low molecular weight heat shock protein. Plant Mol Biol1997; 33:483-492.

22. Waters ER, Lee GJ, Vierling E. Evolution, structure and function of small heat shockproteins in plants. J Exptl Bot 1995; 47:325-338.

23. Lee GJ, Pokala N, Vierling E. Structure and in vitro molecular chaperone activity of cytosolicsmall heat shock proteins from pea. J Biol Chem 1995; 270:19432-10438.

24. Lee GJ, Roseman AM, Saibil HR et al. A small heat shock protein stably binds heat-denatured model substrates and can maintain a substrate in a folding-competent state.EMBO J 1997; 16:659-671.

25. Jinn T-L, Chang PFL, Chen YM et al. Tissue-type-specific heat-shock response andimmunolocalization of class I low molecular weight heat-shock proteins in soybean. PlantPhysiol 1997; 114:429-438.

26. Nover L, Scharf KD, Neumann D. Cytoplasmic heat shock granules are formed fromprecursor particles and are associated with a specific set of mRNAs. Molec Cell Biol 1989;9:1298-1308.

27. Schöffl F, Baumann G, Raschke E. The expression of heat shock genes—A model forenvironmental stress response. In: Verma DP, Goldberg RB ed. Temporal and spatialregulation of plant genes, Plant gene research V, New York: Springer Verlag, 1988:253-273.

28. Schöffl F. Molecular genetics of the heat shock response. In: Maybry HT, Nguyen HT,Dixon RA, Bonness MS ed. Biotechnology for arid plants. Austin: IC2 Institute. University ofTexas at Austin, 1994:83-92.


29. Sanchez Y, Lindquist S. HSP104 is required for induced thermotolerance. Science 1990;248:1112-1115.

30. Sanchez Y, Taulien J, Borkowich KA et al. HSP104 is required for tolerance to manyforms of stress. EMBO J 1992; 11:2357-2364.

31. Petko L, Lindquist S. HSP26 is not required for growth at high temperatures, nor forthermotolerance, spore development, or germination. Cell 1986; 45:885-894.

32. Schöffl F, Rieping M, Baumann G. Constitutive transcription of a soybean heat shockgene by a cauliflower mosaic virus promoter in transgenic tobacco plants. Dev Genetics1987; 8:365-374.

33. Schöffl F, Diedring V, Kliem M et al. The heat shock response in transgenic plants: Theuse of chimeric heat shock genes. In: Wray JL, ed. Inducible plant proteins: Theirbiochemistry and molecular biology. Cambridge: Cambridge University Press, 1992:247-265.

34. Plesovsky-Vig N, Brambl R. Disruption of the gene for HSP30, an α-cristallin-related heatshock protein of Neurosposra crassa, causes defects in thermotolerance. Proc Natl Acad SciUSA 1995; 92:5032-5036.

35. Morimoto RI, Tissieres A, Georgopoulos C. Stress proteins in biology and medicine. NewYork: Cold Spring Harbor Laboratory, 1990.

36. Li GC, Li L, Liu Y-K et al. Thermal response of rat fibroblast stably transfected with thehuman 70-Kda heat shock protein-encoding gene. Proc Natl Acad Sci USA 1991;88:1681-1685.

37. Li GC, Li L, Liu RY et al. Heat shock protein HSP70 protects cells from thermal stresseven after deletion of its ATP-binding domain. Proc Natl Acad Sci USA 1992; 89:2036-2040.

38. Suzuki K, Watanabe M. Modulation of cell growth and mutation induction by introductionof the expresssion vector of human HSP70 gene. Experimental Cell Res 1994; 215:75-81.

39. Welte MA, Tetrault JM, Dellavalle RP et al. A new method for manipulating transgenes:Engineering heat tolerance in a complex, multicellular organism. Current Biol 1993;3:842-853.

40. Kim D, Ouyang H, Li G. Heat shock protein HSP70 accelerates the recovery of heat-shockedmammalien cells through its modulation of heat shock transcription factor HSF1. ProcNatl Acad Sci USA 1995; 92:2126-2130.

41. Lee JH, Hübel A, Schöffl F. Derepression of the activity of genetically engineered heatshock factor causes constitutive synthesis of heat shock proteins and increasedthermotolerance in transgenic Arabidopsis. Plant J 1998; 258:269-278.

42. Prändl R, Hinderhofer K, Eggers-Schumacher G, Schoeffl F. HSF3, a new heat shockfactor from Arabidopsis thaliana, derepresses the heat shock response and confersthermotolerance when overexpressed in transgenic plants. Mol Gen Genet 1998;258:269-278.

43. Burke TJ, Callis J, Vierstra RD. Characterization of a polyubiquititn gene from Arabidopsisthaliana. Mol Gen Genet 1988; 213:435-443.

44. Sun CW, Callis J. Independent modulation of Arabidopsis thaliana polyubiquitin mRNAsin different organs of and in response to environmental changes. Plant J 1997; 11:1017-1027.

45. Hérouart D, Van Montagu M, Inzé D. Developmental and environmental regulation ofthe Nicotiana plumbaginifolia cytosolic Cu/Zn-superoxide dismutase promoter intransgenic tobacco. Plant Physiol 1994; 104:873-880.

46. Brown JA, Li D, Ic M et al. Heat shock induction of manganese peroxidase genetranscription in Phanerochaete chryosporium. Appl Environ Microbiol 1993; 59:4295-4299.

47. Almoguera C, Coca MA, Jordano J. Tissue-specific expression of sunflower heat shockproteins in response to water stress. Plant J 1993; 4:947-958.

48. Alamillo J, Almoguera C, Bartels D, Jordano J. Constitutive expression of small heat shockproteins in vegetative tissues of the resurrection plant Craterostigma plantagineum. PlantMol Biol 1995; 29:1093-1095.

49. Kiyosue T, Yamaguchi-Shinozaki K, Shinozaki K. Cloning of cDNAs for genes that areearly responsive to dehydration stress in Arabidopsis thaliana L.: Identification of threeERDs as HSP cognate genes. Plant Mol Biol 1994; 25:791-798.


50. Prändl R, Kloske E, Schöffl R. Developmental regulation and tissue specific differencesin the expression of a chimeric heat shock gene in transgenic tobacco and Arabidopsisplants. Plant Mol Biol 1995; 28:73-82.

51. Sabehat A, Weiss D, Lurie S. The correlation between heat shock protein accumulationand persistence and chilling tolerance in tomato fruit. Plant Physiol 1996; 110:531-537.

52. Collins GG, Nie XL, Saltveit E. Heat shock proteins and chilling sensitivity in mung bean.J Exptl Botany 1995; 46:795-802.

53. Jennings P, Salteveit ME. Temperature and chemical shocks chilling tolerance in germinatingCucumis sativus (cv. Poinsett 76) seeds. Physiol Plant 1994; 91:703-707.

54. Krishna P, Sacco M, Cherutti JF et al. Cold-induced accumulation of HSP90 transcriptsin Brassica napus. Plant Physiol 1995; 107:915-923.

55. Anderson JV, Li QB Haskell DW et al. Structural organization of the spinach endoplasmicreticulum-luminal 70-kilodalton heat shock cognate gene and expression of 70-kilodaltonheat shock genes during cold acclimation. Plant Physiol 1994; 104:1359-1370.

56. Cabane M, Calvet P, Vincens P, Boudet AM. Characterization of chilling-acclimation-relatedproteins in soybean and identification of one as a member of the heat shock protein(HSP70) family. Planta 1993; 190:346-353.

57. Van Berkel J, Salamini F, Gebhardt C. Transcripts accumulating during cold storage ofpotato (Solanum tuberosum L) tubers are sequence related to stress responsive genes.Palnt Physiol 1994; 104:445-452.

58. Czarnecka E, Edelman L, Schöffl F et al. Comparative analysis of physical stress responsesin soybean seedlings using cloned heat shock cDNAs. Plant Mol Biol 1984; 3:45-58.

59. Edelman L, Czarnecka E, Key JL. Induction and accumulation of heat-shock-specificpoly(A+) RNAs and proteins in soybean seedlings during arsenite and cadmium treatments.Plant Physiol 1983; 86:1048-1056.

60. Howarth CJ. Heat shock proteins in sorghum and pearl millet; ethanol, sodium arsenite,sodium malonate and the development of thermotolerance. J Eptl Botany 1990; 41:877-884.

61. Orzech KA, Burke JJ. Heat shock and the protection against metal toxicity in wheat leaves.Plant Cell Envir 1988; 11:711-714.

62. Neumann D, Lichtenberger O, Günther D et al. Heat-shock proteins induce heavy-metaltolerance in higher plants. Planta 1994; 194:360-367.

63. Wollgiehn R, Neumann, D. (1995) Stress Response of tomato cell cultures to toxic metalsand heat shock: Differences and similarities. J Plant Physiol 1990; 146:736-742.

64. Davidson JF, Whyte B, Bissinger PH et al. Oxidative stress is involved in heat-inducedcell death in Saccharomyces cerevisiae. Proc Natl Acad Sci USA 1996; 93:5116-5121.

65. Polla BS, Kantengwa S, Francois D et al. Mitochondria are selective targets foe the protectiveeffects of heat shock against oxidative injury. Proc Natl Acad Sci USA 1996; 93:6458-6463.

66. Schuster G, Even D, Kloppstech K et al. Evidence for protection by heat shock proteinsagainst photoinhibition during heat shock. EMBO J 1988; 7:1-6.

67. Foyer CH, Descourviers P, Kunert KJ. Protection against oxygen radicals: An importantdefense mechanism studied in transgenic plants. Plant Cell Environ 1994; 17:507-523.

68. Bowler C, Slooten L, Vandenbranden S et al. Manganese superoxide dismutase canreduce cellular damage mediated by oxygen radicals in transgenic plants. EMBO J 1991;10:1723-1732:

69. Van Camp W, Capiau K, Van Montagu M et al. Enhancement of oxidative stress tolerancein transgenic tobacco plants overproducing Fe-superoxide dismutase in chloroplasts. PlantPhysiol 1996; 112:1703-1714.

70. McKersie BD, Chen Y, de Beus M et al. Superoxide dismutase enhances tolerance offreezing stress in transgenic alfalfa (Medicago sativa L.). Plant Physiol 1993; 103:1155-1163.

71. Perl A, Perl-Treves R, Galili S et al. Enhanced oxidative stress defence in transgenic potatooverexpressing tomato CuZn superoxide dismutase. Theor Appl Genet 1993; 85:586-576.

72. Allen RD. Dissection of oxidative stress tolerance using transgenic plants. Plant Physiol1995; 107:1049-1054.

73. McKersie B, Bowley S, Harjanto E et al.Water-deficit tolerance and field performance oftransgenic alfalfa overexpressing superoxide dismutase. Plant Physiol 1996; 111:1177-1181.


74. Schöffl F, Rossol I, Angermüller S. Regulation of the transcription of heat shock genes innuclei from soybean (Glycine max.) seedlings. Plant Cell Environ 1987; 10:113-119.

75. Baumann G, Raschke E, Bevan M et al. Functional analysis of sequences required fortranscriptional activation of a soybean heat shock gene in transgenic tobacco. EMBO J1987; 6:1161-1166.

76. Schöffl F, Rieping M. Baumann G et al. The function of heat shock promoter elementsin the regulated expression of chimaeric genes in transgenic tobacco. Mol Gen Genet1989; 217:246-253.

77. Czarnecka E, Key JL, Gurley WB. Regulatory domains of the Gmhsp17.5-E heat shockpromoter of soybean: A mutational ananlysis. Mol Cell Biol 1989; 9:3457-3463.

78. Wu C; Activating protein factors bind in vitro to upstream control sequences in the heatshock gene chromatin. Nature 1984; 311:81-84.

79. Amin J, Anathan J, Voellmy R. Key features of heat shock regulatory elements. Mol CellBiol 1988; 8:3761-3769.

80. Xiao H, Lis JT. Germ line transformation used to define key features of heat shockresponse elements. Science 1988; 239:1139-1142.

81. Perisic O, Xiao H, Lis JT. Stable binding of Drosophila heat shock factor to head to headand tail to tail repeats of a conserved 5 bp recognition unit. Cell 1989; 59:797-806.

82. Sorger PK, Nelson HCM. Trimerization of a yeast transcriptional activator via coiled-coilmotif. Cell 1989; 59:807-813.

83. Kim SJ, Tsukiyama T, Lewis MS, Wu C. Interaction of the DNA binding domain ofDrosophila heat shock factor with its cognate DNA site: A thermodynamic analysis usingultracentrifugation. Protein Sci 1994; 3:1040-1051.

84. Hübel A, Schöffl F. Arabidopsis heat shock factor: Characterization of the gene and therecombinant protein. Plant Mol Biol 1994; 26:353-362.

85. Hübel A, Lee JH, Wu C et al. Arabidopsis heat shock factor is constitutively active inDrosophila and human cells. Mol Gen Genet 1995; 248:136-141.

86. Spena A, Hain R, Ziervogel U et al. Construction of a heat-inducible gene for plants.Demonstration of heat-inducible activity of the Drosophila HSP70 promoter in plants.EMBO J 1985; 4:2739-2743.

87. Fernandez M, O’Brien T, Lis JT. Structure and regulation of heat shock gene promoters.In: Morimoto RI, Tissières A, Gorogopoulus C ed. The biology of heat shock proteinsand chaperones. New Work: Cold Spring Harbor Press, 1994:375-394.

88. Topol J, Ruden DM, Parker CS. Sequences required for in vitro transcriptional activation ofa Drosophila heat shock gene. Cell 1985; 42:527-537.

89. Wu C. Heat schock transcription factors: Structure and regulation. Annu Rev Cell DevBiol 1995; 11:441-469.

90. Rieping M, Schöffl F. Synergistic effect of upstream sequences, CCAAT box elements,and HSE sequences for enhanced expression of chimaeric genes in transgenic tobacco.Mol Gen Genet 1992; 231,226-232.

91. Czarnecka E, Key JL, Gurley WB. Regulatory domains of the Gmhsp17.5-E heat shockpromoter of soybean: a mutational ananlysis. Mol Cell Biol 1989; 9:3457-3463.

92. Schöffl F, Schröder G, Kliem M. et al. A SAR sequence containing 395 bp fragmentmediates enhanced, gene-dosage-correlated expression of a chimaeric heat shock genenin transgenic tobacco plants. Transgenic Res 1993; 2:93-100.

93. Giardina C, Perez-Riba M, Lis JT. Promoter melting and TFIID complexes on Drosophilagenes in vivo. Genes Dev 1992; 15:2737-2744.

94. Tsukijama T, Becker PB, Wu C. ATP-dependent nuclesome disruption at a heat shockpromoter mediated by binding of GAGA transcription factor. Nature 1994; 367:525-532.

95. Prändl R, Schöffl F. Heat shock elements are involved in heat shock promoter activationduring tobacco seed maturation. Plant Mol Biol 1996; 31,157-162.

96. Wehmeyer N, Hernandez LD, Finkelstein RR et al. Synthesis of small heat shock proteins ispart of the developmental program of late seed maturation. Plant Physiol 1996;112:747-757.


97. Quatrano RS, Bartels D, Ho TH et al. New insights into ABA-mediated processes. PlantCell 1997; 470-475.

98. Hill A, Nantel A, Rock CD et al. A conserved domain of the viviparous-1 gene productenhances the DNA binding activity of the bZIP protein EmBP-1 and other transcriptionfactors. J Biol Chem 1996; 271:3366-3374.

99. Jedlicka P, Mortin MA, Wu C. Multiple functions of Drosophila heat shock transcriptionfactor in vivo. EMBO J 1997; 16:2452-2462.

100. Mager WH, De Krujiff AJJ. Stress-induced transcriptional activation. Microbiol Rev 1995;59:506-532.

101. Nover L, Scharf KD, Gagliardi D et al. The HSF world: Classification of plant heat stresstranscription factors. Cell Stress Chaperones 1996; 1:215-223.

102. Hübel A, Lee JH, Wu C et al. Arabidopsis heat shock factor is constitutively active inDrosophila and human cells. Mol Gen Genet 1995; 248:136-141.

103. Czarnecka-Verner E, Yuan CH, Fox PC et al. Isolation and characterization of six heatshock transcription factor cDNA clones from soybean. Plant Mol Biol 1995; 29:37-51.

104. Scharf KD, Rose S, Zott W et al. Three tomato genes code for heat stress transcriptionfactors with a region of remarkable homology to the DNA-binding domain of yeast HSF.EMBO J 1990; 9:4495-4501.

105. Gagliardi D, Breton C, Chaboud A et al. Expression of heat shock factor and heat shockprotein 70 genes during maize pollen development. Plant Mol Biol 1995; 29:841-856.

106. Harrison CJ, Bohm AA, Nelson HC. Crystal structure of the DNA-binding domain ofthe heat shock transcription factor. Science 1994; 263:224-227.

107. Vuister GW, Kim SJ, Orosz A et al. Solution structure of the DNA-binding domain ofDrosophila heat shock transcription factor. Struct Biol 1994; 1:605-614.

108. Schultheiss J, Kunert O, Gase U et al. Solution structure of the DNA-binding domain ofthe tomato heat stress transcription factor HSF24. Eur J Biochem 1996; 236:911-921.

109. Peteranderl R, Nelson HCM. Trimerization of the heat shock transcription factor by atriple-stranded alpha-helical coiled-coil. Biochem 1992; 31:12272-12276.

110. Rabindran SK, Haroun RI, Clos J et al. Regulation of heat shock factor trimer formation:Role of a conserved leucine zipper. Science 1993; 259:230-234.

111. Nakai A, Morimoto RI. Characterization of a novel chicken heat shock transcription factor,heat shock factor 3, suggests a new regulatory pathway. Mol Cell Biol 1993; 13:1983-1997.

112. Zuo J, Baler R, Dahl G, Voellmy R. Activation of the DNA-binding ability of humanheat shock transcription factor 1 may involve the transition from an intramolecular toan intermolecular triple-stranded coiled-coil structure. Mol Cell Biol 1994; 14:7557-7568.

113. Orosz A Wisniewski J, Wu C. Regulation of Drosophila heat shock factor trimerization:Global sequence requirements and independence of nuclear localization. Mol Cell Biol1996; 16:7018-7030.

114. Lyck R, Harmening U, Höhfeld I et al. Intracellular distribution and identification of thenuclear localization signals of two plant heat-stress transcription factors. Planta 1997;202:117-125.

115. Sheldon LA, Kingston RE. Hydrophobic coiled-coil domains regulate the subcellularlocalization of human heat shock factor 2. Genes Dev 1993; 7:1549-1558.

116. Zuo J, Rungger D, Voellmy R. Multiple layers of regulation of human heat shocktranscription factor 1. Mol Cell Biol 1995; 15:4319-4330.

117. Zandi E, Tran TN, Chamberlain W et al. Nuclear entry, oligomerization, and DNA-bindingof the Drosophila heat shock transcription factor are regulated by a unique nuclearlocalization sequence. Genes Dev 1997; 11:1299-1314.

118. Newton EM, Knauf U, Green M et al. The regulatory domain of human heat shock factor 1is sufficient to sense heat stress. Mol Cell Biol 1996; 16:839-846.

119. Wisniewski J, Orosz A, Allada R et al. The C-terminal region of Drosophila heat shockfactor (HSF) contains a constitutively functional transactivation domain. Nucleic AcidsRes 1996; 24:367-374.

120. Treuter E, Nover L, Ohme K et al. Promotor specificity and deletion analysis of threeheat shock transcription factors of tomato. Mol Gen Genet 1993; 240:113-125.


121. Scharf KD, Materna T, Treuter E et al. Heat stress promoters and transcription factors.In: Nover L, ed. Plant Promoters and Transcription Factors. Springer-Verlag, Berlin, 1994;125-162.

122. Giardina C, Lis JT. Dynamic protein-DNA architecture of a yeast heat shock promoter.Mol Cell Biol 1995; 15:2737-2744.

123. Bonner JJ, Heyward S, Fackenthal DL. Temperature-dependent regulation of a heterologoustranscriptional activation domain fused to heat shock transcription factor. Mol Cell Biol1992; 12:1021-1030.

124. Chen Y, Barlev NA, Westergaard O et al. Identification of the C-terminal activatordomain in yeast heat shock factor: independent control of transient and sustainedtranscriptional activity. EMBO J 1993; 12:5007-5018.

125. Boorstein WR, Craig EA. Transcriptional regulation of SSA3, an HSP70 gene fromSaccharomyces cerevisiae. Mol Cell Biol 1990; 10:3262-3267.

126. Baler R, Zou J, Voellmy R. Evidence for a role of HSP70 in the regulation of the heatshock response in mammalian cells. Cell Stress Chaperones 1996; 1:33-39.

127. Rabindran SK, WisniewskiJ, Li G et al. Interaction between heat shock factor and HSP70is insufficient to suppress induction of DNA-binding activity in vivo. Mol Cell Biol 1994;14:6552-6560.

128. Lee JH, Schöffl F. A HSP70 antisense gene affects the expression of HSP70/HSC70, theregulation of HSF, and the acquisition of thermotolerance in transgenic Arabidopsisthaliana. Mol Gen Genet 1996; 252:11-19.

129. Høj A, Jakobsen BK. A short element required for turning off heat shock transcriptionfactor: Evidence that phosphorylation enhances deactivation. EMBO J 1994; 3:2614-2624.

130. Knauf U, Newton EM, Kyriakis J et al. Repression of human heat shock factor 1 activityat control temperature by phosphorylation. Genes Dev 1996; 10:2782-2793.

131. Chu B, Soncin F, Price BD et al. Sequential phosphorylation by mitogen-activated proteinkinase and glycogen synthase kinase 3 represses transcriptional activation by heat shockfactor 1. J Biol Chem 1996; 271:30847-30857.

132. Reindl A, Schöffl F, Schell J et al. Phosphorylation by a cyclin-dependent kinase modulatesDNA-binding of the Arabidopsis heat shock transcription factor HSF1 in vitro. PlantPhysiol 1997; 115:93-100.

133. Carratu L, Franceschelli S, Pardini CL et al. Membrane lipid perturbation modifies theset point of the temperature of heat shock response in yeast. Proc Natl Acad Sci USA1996; 93:3870-3875.

134. Gallie DR, Caldwell C, Pitto L. Heat shock disrupts cap and poly(A) tail function duringtranslation and increases mRNA stability of introduced reporter mRNA. Plant Physiol1995; 108:1703-1713.

135. Gallie DR, Le H, Caldwell C et al. The posphorylation state of translation initiation factors isregulated developmentally and following heat shock in wheat. J Biol Chem 1997;272:1046-1053.

136. Pitto L, Gallie DR, Walbot V. Role of the leader sequence during thermal repression oftranslation in maize, tobacco, and carrot protoplasts. Plant Physiol 1992; 100:1827-1833.

CHAPTER 6

Molecular Responses to Cold, Drought, Heat and Salt Stess in Higher Plants, edited by KazuoShinozaki and Kazuko Yamaguchi-Shinozaki. ©1999 R.G. Landes Company.

Cellular Responses to Water StressMichael R. Blatt, Barbara Leyman and Alexander Grabov

Plant biomass is inextricably tied to water transpiration: increases in crop productionmust appear inevitably in an increased water use, regardless of improvements in water

management or farming practice. Thus, on a global scale water use and the physiologicalconsequences of stress, when water becomes limiting, remain among the most importantfactors that influence vegetative plant growth and yield.1 Although water limitation is anobvious feature of arid environments, conditions of water stress develop in association withsaline soils and, hence, may also become evident where irrigation leads to a buildup of salt.2

Paradoxically flooding of soils, too, can lead to water shortage of the aerial parts of the plantas the root environment becomes deprived of oxygen.3,4

The primary defense of the plant against water loss and dehydration, especially overrelatively short time scales, is to reduce transpirational water loss. It is now generallyrecognized that plants respond to water stress by synthesizing the hormone abscisic acid(ABA) which is transported via the xylem to the leaf tissues. Most important to this pro-cess, ABA effects the closure of stomatal pores in the leaf epidermis and dramaticallyreduces foliar transpiration, in many cases by two orders of magnitude or more. Foliarconductance to water vapor of mesophytes and crop plants often lie in the range of 10-20mm s-1 under conditions in which stomata are largely open, and these figures fall to valuesnear 0.1 mm s-1 or lower—equivalent to the cuticular conductance—when stomata close.6,7

In xerophytes and many trees, conductances under water stress can fall still lower to valuesapproaching 0.01 mm s-1.7 Clearly, understanding the factors that control stomatal aperturewill be crucial to future developments toward improving vegetative yields in the face ofincreasing pressure on water resources and arable land usage.

At the same time, the guard cells that surround the stomatal pore have become a focusof attention in fundamental research as well. The ability of these cells to integrate bothenvironmental and internal signals—and their unique situation within the leaf tissue—has provided a wealth of experimental access points to signal cascades that link membranetransport to stomatal control.8 The past 15 years of research has raised the status of theguard cell to that of an undisputed higher-plant cell “model”, and some of the mostexciting new findings in plant cell signaling have come from work with guard cells, takingadvantage of the “stomatal interface” between molecular genetics and biophysics. In thischapter we examine some of these recent developments and their background that formthe center of much debate about second messengers in plant cell biology. We also exploretheir implications for plant response to water stress.


The Stomatal SituationStomata are pores formed by pairs of specialized cells, the guard cells, and are found

on the epidermis of all aerial parts of most higher plants. Stomata open and close to control gasexchange between the intercellular spaces within the plant tissue and the surroundingenvironment. Thus, stomata have a fundamental role in controlling two of the mostimportant processes in vegetative plant physiology, photosynthesis and transpiration: theyopen to allow sufficient CO2 to enter the leaf for photosynthetic carbon fixation, and theyclose to reduce transpiration under conditions of water stress.

Stomata (sing. stoma, Greek for mouth) were some of the first microscopic structuresto be identified in higher plant tissues,9 coinciding with Hooke’s discovery of cells in thesecond half of the 17th century, and early on were associated with "plant breathing".10 DeCandolle11 demonstrated that stomatal apertures were variable. However, recognition thatstomatal movements were driven by turgor came only later with von Mohl’s anatomicaland physiological studies.12 The relationship between the accumulation of inorganic K+

and stomatal opening was identified by Immamura13 and Yamashita14 in the first half ofthis century, but it was not until Fujino15 published his work in English in 1967 thatscientists outside Japan were made fully aware of there significance. Fischer’s work16,17

independently confirmed these findings in epidermal peels of Vicia, concluding that theamount of K+ together with an accompanying anion would be sufficient to account for theincrease in osmotic pressure during stomatal opening.

The mechanics of stomatal function is intimately connected with their morphology.From an anatomical point of view there are two basic types of stomata, although intermediateforms also exist in gymnosperms and sedges.7 The first type has kidney-shaped guard cellsthat surround an elliptical pore in the epidermal surface. These stomata are typical ofdicotyledonous plants including Vicia, Nicotiana and Arabidopsis, but are also found inCommelina. They remain the best-studied form in terms of guard cell membrane transportand cell signaling. The second type of stomata are restricted to the monocotyledons andhave dumbell-shaped guard cells and an almost rectangular-shaped pore.18 In each case,the swelling of the guard cells in opposition to one another leads to an opening of thepore. However the mechanics differ according to the shape of the guard cells. Swellingof the monocotyledonous guard cells is restricted by cell wall depositions primarily to thedumbell-like ends, and opposition between the two cells thus opens the pore slitwise. Inkidney-shaped guard cells the cell-wall thickenings around each guard cell are mostpronounced on the side facing the pore. Cell expansion thus leads to an outward “bowing”which opens the pore in an oval.

As may be expected, estimates of the change in guard cell volume between the closedand open states of stomata vary between species. Volume changes of 0.2-0.35 pl changeper µm of pore aperture have been reported for Vicia,19,20 while values for Commelina maybe as high as 0.4 pl/µm.19 These values are significantly larger than our estimates forArabidopsis that fall between 0.03 and 0.08 pl/µm. Obviously such estimates depend onthe relative size of the guard cells and provide a rough guide only, because even in onespecies guard cell size can vary, dependent on growth conditions and the age of the plant.7

However, the proportional changes in volume they imply are substantial, considering thesmall size of guard cells compared with most other plant cell types. Dimensions of Viciaguard cells, for example, are typically about 40 µm in length and 8-10 µm in diameter attheir widest point, giving volumes near 4 pl when apertures are between 6-8 µm.21 Thusthe volume changes between the closed state and fully-open state (typically 10-14 µm)entail increases of 2- to 3-fold (200-300%).

101Cellular Responses to Water Stress

Transport MechanicsAt maturity guard cells lack functional plasmodesmata,22 so all of the osmotic solute

flux that drives cell volume changes for stomatal movement must take place across theplasma membrane. The transport mechanics that underpin these solute fluxes are nowwell-established as a result of the electrophysiological and radiotracer analyses of the pastdecade. Until recently, much of this work was carried out on Vicia, primarily because oftheir ease of handling. However, comparable data are now to hand for guard cells ofNicotiana23,24 and of Arabidopsis.25,26 These studies so far indicate only minor differencesbetween species in transport characteristics, although more subtle differences in their controlhave yet to be examined systematically.

Plasma MembraneThe fluxes entailed—notably K+, Cl- and in some instances also organic acids such as

malate—are considerable: between the open and closed states of the stomata, guard cellsof Vicia take up and release, respectively, 2-4 pmol of KCl which on a cell volume basis isequivalent to changes of approximately 300 mOsM in osmotically active solutes.7 Theseevents commonly take place over periods of 20 min to 2 h,27 implying a high level oftransport activity by comparison with many other higher-plant cells. In fact, there isevidence that Vicia guard cells express unusually high levels of H+-ATPase at the plasmamembrane,28-30 consistent with the higher demand expected for membrane energization.Likewise, measurements of H+-ATPase current under intracellular voltage clamp havegenerally yielded values at 0 mV around 3-10 µA cm-2 (= 50-200 pA per cell).31-33 Forcomparison a current of 35 pA can be estimated as the minimum required for stomatalopening. This estimate assumes (probably unrealistically, in view of the energetic needs ofother homeostatic transport processes) that the H+-ATPase current is used entirely to driveK+ uptake into the cell in exchange for H+ over a 2 h opening period. [It is worth notingthat initial values of 1.5-6 pA, based on patch electrode measurements34,35 were probablycompromised by cytosolic exchange with the patch electrode filling solution and theconsequent loss of regulatory and other cofactors.36 More recent “perforated patch”techniques have gone some way to overcoming these difficulties and have yielded currentsof 10-20 pA.37,38]

Quantitative differences apart, primary energisation of the guard cell plasma membrane,like the plasma membranes of most plant and fungal cells, is achieved by ATP-dependent H+

extrusion. At least two distinct H+-ATPase genes are expressed in Vicia guard cells,28

although the physiological characteristics of these isoforms remain to be examined.Analysis of H+-ATPase function has shown that H+ extrusion is coupled to ATP hydrolysis in a1:1 ratio33 and shares kinetic features in common with H+ pumps of Neurospora39 andChara.40 In theory the H+-ATPase will support stable membrane voltages near -360 mVwith an external pH of 6.33 However, this electrical driving force is utilized to power themovement of other solutes (notably K+ and Cl- uptake) across the membrane, so thetheoretical limit is never achieved in practice. Membrane voltages near -300 mV havebeen observed in low external K+,31 thus ruling out coupling ratios of 2:1 (H+:ATP) orhigher which would limit the H+- ATPase output to a maximum near -200 mV.33

Our knowledge of the mechanisms for K+ across the plasma membrane is dominatedby two classes of K+ channels that give rise to current rectifying inward (IK,in) and outward(IK,out), respectively. These two pathways are major contributors to K+ flux during stomatalopening and closing, respectively, and are clearly separable on bases of their biophysicaland pharmacological properties (below; see also ref. 41). However, guard cells probably alsopossess secondary coupled transporters for K+ 42 similar to those known in fungi43

and other plant cells.44 Ion flux through channels is inherently passive and thus critically


dependent on the membrane voltage and equilibrium voltage for the permeant ions(Ex). There are circumstances in which uptake of K+ occurs against the prevailing electro-chemical driving force for K+ (V>EK), for example in low extracellular [K+] and in thepresence of fusicoccin.42

The major inward-rectifying K+ channel in guard cells of Arabidopsis has beenidentified with the KAT1 K+ channel45 and its homologues that show single-channelconductances of 5-8 pS at near-physiological [K+] as well as similar physiological andbiophysical properties when expressed in Xenopus oocytes46,47 and in yeast.48 Uptake of K+

through IK,in is demonstrable electrically in millimolar external [K+].31,49,50 These channels areactivated normally by membrane voltages more negative than -100 mV,37,51 they show anapparent gating charge of about 1.4-1.5,37,51 and give rise to an inward-directed (byconvention, negative) current and corresponding K+ flux. The current appears to requiremillimolar external [K+] for activity,51 consistent with a putative extracellular K+-bindingsite with a Kd near 0.4 mM.52 However, K+ otherwise has no effect on IK,in gating or itsvoltage sensitivity.

By contrast, the gating of IK,out is affected by extracellular [K+].53 This channel has asomewhat higher single-channel conductance (=10-25 pS)54,55 and shows an apparentgating charge near 2,53,56 implying a steeper dependence on membrane voltage. With 10mM [K+]outside, activation of IK,out normally occurs at voltages near -70 mV and thecurrent is fully activated at voltages near 0 mV and more positive. Furthermore unlikeIK,in, the IK,out gate is affected by extracellular [K+]so that its voltage dependence shifts inparallel with EK.26,53,57 This current has been indicated as the major pathway for K+ effluxduring stomatal closure,42,58 so its dependence on extracellular [K+] makes good “physiologicalsense” in an environment in which [K+] can vary over more than an order of magnitude: itensures that K+ passage can only occur when the driving force on the ion will lead to itsefflux from the cell.57 Blatt and Gradmann53 carried out a detailed analysis of the currentas a function of [K+] and found that its activation depended on theco-operative interaction of 2 K+ ions with the channel, but at a site (or sites) distinct fromthe channel pore. They also observed that the apparent K1/2 for interaction was stronglyvoltage-dependent, accounting for the equivalence of (negative) membrane voltage and[K+] in regulating channel activity. Thus, their data indicate that extracellular K+ acts as aligand and negative regulator of IK,out, and point to a pair of K+-binding sites associatedwith the channel deep within the membrane, but accessible only from the outside ofthe cell.

To an extent, the fluxes of K+ carried by these channels—and hence the accompanyingelectrical charge—are balanced by parallel transport of Cl. Anion efflux is particularlyimportant for stomatal closure, as it is essential that the voltage across the guard cell plasmamembrane is brought positive of EK to effect a net loss of K+ through IK,out. Becausecytosolic [Cl-] under most conditions is probably close to 1 mM,59 ECl typically will besituated close to and positive of 0 mV, that is well positive of EK and sufficient to drive themembrane voltage positive of EK if the membrane conductance to Cl- is elevated.

Two anion channels have been identified with different macroscopic currentkinetics, although both exhibit roughly equivalent single-channel conductances (≅35pS). One of these anion currents activates rapidly with halftimes of less than 50 ms ondepolarization to voltages positive of about -100 mV and deactivates with halftimes of2-5 s on repolarization.60 Once activated, these channels also inactivate with a halftime ofabout 10 s, thus disabling the current even when the voltage is held positive of -100 mV.61

The second anion current (hereafter identified as ICl) has been reported to activate anddeactivate slowly (halftimes, 5-30 s), shows no inactivation with time,62,63 and exhibits asignificant residual conductance at voltages negative of -100 mV.23,62,63


Dietrich and Hedrich64 have suggested an interconversion between these two channeltypes, proposing a single Cl- channel exists in one of two different gating “modes.”65-67

However, the relationship between the two channel forms remains to be examined in anydetail. Regardless of origin, the fast-activating anion current can be ruled out as amajor pathway for anion efflux during stomatal closure. Its narrow voltage range for acti-vation and the fact that the current inactivates within seconds cannot be reconciledwith the requirement for sustained channel functioning over periods of 20-30 min ormore during closure.8,68 In fact, recent evidence has confirmed that ICl responds rapidly andwith prolonged activation to stimuli that evoke stomatal closure.23,25,69

TonoplastThe greater proportion of solutes that pass across the plasma membrane during

stomatal movements must also find their way across the tonoplast. As is the case forvirtually all mature, higher-plant cells the vacuole in guard cells makes up 80-90% of thetotal cell volume. Thus most of the volume changes as the guard cells swell and shrink aretaken up in the vacuole7 and the fact implies a degree of transport coordination at thetonoplast and between the two membranes.70,71 By contrast with events at the plasmamembrane, our understanding of transport across the tonoplast remains very poorindeed. There has been remarkably little work on the pumps that might energize thetonoplast in guard cells. All evidence again indicates that significantly higher vacuole-typeATPase activities are found in these cells compared with leaf mesophyll.72,73 However,Willmer, et al73 could not find evidence for its control by extrinsic stimuli. Furthermore,nothing is known of the mechanisms that are responsible for solute accumulation,although it is likely that the cell must expend energy at least for cation transport into thevacuole.7

The guard cell tonoplast is known to harbor at least two different cation channelsthat are capable of carrying K+ and Ca2+ 74-76 and an anion channel with high selectivity for Cl-

over K+ and dependent on protein phosphorylation for activity.77 Pei, et al77 have suggestedthat the anion channel could be a pathway for Cl- accumulation. The cation channelsare more likely to be important to events of solute loss and stomatal closure. Each mayprove important targets for hormonal and/or environmental control, but their respectivecontributions still remain to be demonstrated. Equally, the vacuole constitutes an importantsource and sink for Ca2+ and H+ generally and, thus, is likely to be a contributor tosignaling events that lead to changes in the free concentration of these ions in thecytosol. Indeed, the vacuole clearly contributes to hormone-mediated changes in pHi

as well as its homeostatic control78 and it is also thought to comprise a key reservoirfor Ca2+ release and stimulus-evoked changes in [Ca2+]i,

79 but much detail is still missing.

Transport Coordination in the Face of StressAbscisic acid acts as a universal signal in plants to prevent stomatal opening and

promote stomatal closure. The potency of ABA to inhibit transpiration was first discovered inthe late 1960s80,81 and these observations were soon linked with early studies on ABAaccumulation under water stress conditions.82,83 It is now recognized that during thevegetative phase of the plant life cycle ABA mediates generally to signal adverse environ-mental conditions including cold, drought and high salinity (see especially chapters 4 and5, this volume). During water stress conditions ABA accumulates in leaf tissues around theguard cells and promotes stomatal closure to reduce transpirational water loss84 (forextensive reviews of the literature see refs. 5,7).


Abscisic acid will initiate stomatal closure within minutes of its application toepidermal strips, to the surface of leaves and via the transpiration stream to intactleaves. However, stomatal responses can vary even between stomata on one leaf (so-called“patchy” or mosaic behavior85,86). Stomatal response to ABA is subject to environmentalfactors such as external [K+]17 and [Ca2+],87 CO2,18,88 circadian period89 and prior stressor exposure to ABA.90 There is also a more prolonged effect of ABA that is observed whenplants are first relieved from a period of water stress. Stomata then often remain partiallyclosed, a characteristic that can extend over periods of hours or even days,91 possibly throughan increased sensitivity to residual ABA levels90 that may reflect an enhanced expression ofsignaling components.92 These observations imply a high degree of coordination incoupling ABA to stomatal movements; they also underscore the integration of ABA-mediated events with other signal cascades within the cells.

How is such a degree of regulation achieved? To this question a very large body ofdata can now be brought to bear. In fact, for guard cells we know more about thedownstream events of evoked transport control than for any other higher-plant cell. Inthe first instance, concerted regulation of sets of transporters, including ion channels andpumps at the plasma membrane and endomembranes, is essential to achieve the requisitesolute flux. For ABA the events leading to stomatal closure requires the modulation of allof the major ion currents at the plasma membrane to effect K+ and Cl- loss from the guardcells. Additional effects at the tonoplast as well as other endomembranes are anticipated,both in relation to bulk K+ salt efflux and to signals dependent on Ca2+ and pH. In thesimplest of schemes, the events that ensue on ABA exposure can be ordered as follows(Fig. 6.1):

1. ABA evokes the development of an inward-directed current, mediated at least inpart by slow-activating anion channels23,25 to depolarize the membrane and generate adriving force for K+ efflux;31

2. It inactivates IK,in that normally mediates in K+ uptake;93 and3. It activates current through IK,out that, together with the anion channels, facilitates

the net loss of salt from the cells.8,93

At least two parallel signaling pathways are known to underpin K+ and anion channelbehavior alone, in addition to alterations in H+-ATPase activity.31,94 Current evidencesupports the idea of [Ca2+]i- and pHi-mediated signaling in guard cells, and the influenceof these signal cascades and their downstream targets overlap significantly. However,whether both cascades originate with the same receptor-binding event and, more still,where the site (or sites?) of ABA perception occurs remains speculative. Furthermore, ithas become increasingly clear that signaling pathways evoked by other stimuli such asCO2

69 converge with ABA-mediated control of plasma membrane transport independent ofthese known second messengers. Thus, as many questions are still unanswered about ABA-evoked stomatal closure. In the remainder of this chapter we review several of these issuesand their background that form the center of current debate about guard cell signaltransduction and transport control.

The Ca2+ Second MessengerEvidence of a role for cytosolic-free [Ca2+] ([Ca2+]i) in regulating K+ channels of

guard cells was uncovered early on and has remained a focus of interest, as it has subsequentlyfor anion channel control in relation to ABA. Elevating [Ca2+]i to micromolar concentrationsgreatly reduces IK,in

95 and alters its voltage dependence while promoting bothanion currents (see also refs.60,63). These characteristics mimic the channel responses toABA.31,93,96 Because [Ca2+]i has been observed to rise during ABA exposures from restnear 100 nM to values on some occasions exceeding 1 µM,97-101 the idea that [Ca2+]i


Fig. 6.1. ABA leads to a concerted modulation (+)=increase or activation, (-)=decrease orinactivation of at least three subsets of plasma membrane ion channels—including thetwo dominant K+ currents IK,in and IK,out, and the slow-activating anion current ICl—and,probably, the H+-ATPase. A scheme for ABA signal perception including hypothical andknown steps entails binding to an integral plasma membrane receptor (1), possibly alsointernal ABA binding sites not shown. Binding may activate a G-protein (G) and triggeractivation of phospholipase C (PLC)-mediated hydrolysis of phosphoinositol bisphosphate(PIP2) to inositol 1,4,5-trisphosphate (IP3) releasing Ca2+ from intracellular stores (2).ABA also affects Ca2+ entry across the plasma membrane and its interaction with intracellular Ca2+

release pathways (shading) leads to high-gain Ca2+-induced Ca2+ release (3). The consequent risein Ca2+i affects IK,in, ICl and the H+-ATPase. A concurrent, but Ca2+-independent rise inpHi (4) acts on IK,in, IK,out and ICl, as wel l as depleting substrate for theH+-ATPase. The abi1 protein phosphatase, and by inference also protein kinases (PK/PP),gate pHi signal transmission (5) to IK,in and IK,out but do not influence ICl. A 2B-type proteinphosphatase may also gate the Ca2+- sensitivity of IK,in as well as vacuolar Ca2+ release notshown. Alterations in phosphorylation state (P*) may mediate in activating ICl directly aswell as modulating the characteristics of other plasma membrane and tonoplast ion channels/transporters.


increases are crucial to ABA stimulus transduction remains a potent concept. Indeed,simply raising [Ca2+]i can be sufficient to bias the membrane for solute efflux by controllingtwo key ion currents, IK,in and ICl.

Attaching a physiological interpretation to [Ca2+]i changes has not been straightforwardeven so. A major problem with the [Ca2+]i hypothesis for guard cell signaling has rested inthe fact that the rise in[Ca2+]i is not universally associated with ABA and related stimuli, oreven stomatal closure.98-101 Fricker, et al99 have pointed out the technical difficulties inherentto fluorescent measurements of [Ca2+]i in higher-plant cells:

1. That the intrinsic cytosolic volume is small, either sandwiched between plasmamembrane and tonoplast or dominated by perinuclear cytoplasm, making it difficultto obtain reliable measurements with high spatio-temporal resolution; and

2. That changes in [Ca2+]i relevant to ion channel control almost certainly occurlocally, adjacent to the plasma membrane.

However, environmental factors including growth temperature102 that are "secondary" tothe ABA stimulus can play a critical role in determining the scope and magnitude of the[Ca2+]i signal. One likely explanation for the influence of secondary factors relates to thetransport “poise” of the cell itself. Recent studies from this laboratory103 have linked changesof [Ca2+]i in ABA to the voltage across the plasma membrane. These observations suggestthat the [Ca2+]i response may be part of an adaptive feedback loop adjusting the ABAresponse of the guard cells to the prevailing requirements for solute flux (see [Ca2+]i

Oscillations... below). So it is all the more remarkable that no further kinetic detail hasbeen forthcoming relating the activities of either IK,in or ICl to [Ca2+]i. It will be importantto know now whether small increases of [Ca2+]i, for example to values near 0.3-0.5 µMsuch are often observed in ABA,97-101 could account for the characteristics of these currents inthe presence of ABA. It is interesting that IK,in was largely unaffected by high [Ca2+]i whenEGTA was used as the Ca2+ buffer in patch-clamp experiments. Kelly, et al104 havesuggested that this dependence on the buffer is related to the dynamics of its binding withCa2+ as, unlike BAPTA, EGTA shows a relatively slow Ca2+-binding rate.105-108 Nonetheless, asthese measurements were carried out under steady-state conditions in which [Ca2+]i washeld constant, it is not clear how the relaxation kinetics of Ca2+-buffer binding canaccount for such a difference. Whatever the explanation for their observations, the datamake it clear that estimating the Ca2+-sensitivity of IK,in in vivo is not straightforward inpatch clamp experiments.

[Ca2]i Oscillations and the Origins of [Ca2]i IncreaseFrom where does the Ca2+ originate that contributes [Ca2+]i increases? Clearly, the

apoplast is the ultimate source for all cytosolic Ca2+. Entry of Ca2+ across the plasmamembrane also contributes to the [Ca2+]i rise evoked by ABA. In an elegant study of K+ fluxcontrol by ABA, MacRobbie109 demonstrated that the K+ (86Rb+) efflux evoked by ABA inCommelina guard cells displays two peaks. The initial, rapid stimulation in efflux in the first1-2 min was probably associated with membrane depolarization and the shift in electro-chemical driving force for K+ across the plasma membrane. A second, and much slowerstimulation—with a maximum some 10 min after ABA addition—was thought to representK+ (86Rb+) release from the vacuole. The slow rise in efflux could also have reflectedlonger-term regulatory effects at the plasma membrane. But regardless of its origin, thissecond peak in efflux required the presence of extracellular Ca2+. In accord with theseobservations, Ca2+ channel blockers have been found to affect ABA-induced stomatalclosure.110 Thus, a simple interpretation is that Ca2+ entry from outside is important tomaintaining osmotic solute flux over the time periods of tens of minutes required forstomatal closure.


A large body of data also supports a role for Ca2+ release from internal stores duringABA stimulation—and in guard cell signaling generally—although there remains someuncertainty about the relative contributions of various pathways. Blatt, et al111 and Gilroy,et al112 showed that cytosolic inositol-1,4,5-trisphosphate was capable of stimulating a risein [Ca2+]i, even in the presence of extracellular Ca2+ channel blocker La3+ to blockCa2+ entry from outside, and of inactivating IK,in. Subsequent studies have added furthersupport to the idea of a signaling sequence that passes through inositol-1,4,5-trisphosphate-113-115

and, more recently, also through cADPR- (ryanodine receptor) mediated Ca2+

release.116,117 These studies have indicated parallels with animal cell models, although themonophasic dependencies on cADPR reported for Ca2+ release116 are at odds with thebiphasic cADPR characteristics that have been observed in this case.118

Interest in the [Ca2+]i hypothesis in guard cells has been still further heightened byrecent observations that [Ca2+]i may oscillate in response to external stimuli.119,120 It isplausible that these characteristics constitute “Ca2+ signatures” to encode specific responses,much as has been argued for animal cells,121,122 and the analogy raises questions about theorigins of such oscillations in guard cells. In many animal cells, [Ca2+]i signals depend onentry of Ca2+ across the plasma membrane, its release from intracellular stores and itssubsequent elimination from the cell or sequestration within organelles.118,122 Coordination ofthese processes gives rise to short-lived and repetitive [Ca2+]i spikes, with durations of afew seconds, that may serve to frequency-encode cellular responses, including geneexpression.123 However, the oscillations reported in guard cells to date are at least an orderof magnitude slower, with individual cycle durations of minutes and occurring with periods of5-10 min or more.

It now seems that the situation in guard cells may differ significantly from the animalmodels, and that these slow [Ca2+]i fluctuations are driven by oscillations in membranevoltage. Guard cells commonly show two states of membrane voltage,31,124 one state situatedclose to and positive of the K+ equilibrium voltage (EK), and the second characterized byvoltages well negative of EK. Transitions between these two states occur spontaneously andare potentiated by various stimuli, including ABA;31,49 they are rapid, often exceeding 150mV in amplitude and can occur in cyclic oscillations similar to cardiac action potentials,but with periods of tens of seconds to minutes. Our recent work103 has shown that mem-brane hyperpolarization during such oscillations or under voltage clamp evokes a Ca2+

influx across the plasma membrane and initiates a wave of high [Ca2+]i that propagatescentripetally from the cell periphery. Hyperpolarization beyond about -120 mV is essential toevoke the response and evidence from pharmacological analyses indicates that the processdepends also on intracellular release from Ca2+ stores.

The observations imply a capacity for Ca2+-induced Ca2+ release (CICR) in the guardcells that parallels established patterns of CICR coupling in animal cells.122,125 Nonetheless, thedata underscore some important differences to the animal models, notably the dependence onvoltages more negative than -120 mV. By contrast, voltage-evoked CICR in neuromusculartissues is normally triggered by membrane depolarization that activates Ca2+ influx throughpharmacologically-distinct, L-type Ca2+ channels.125-127 Both the acute voltage-sensitivityand [Ca2+]i relaxation kinetics clearly distinguish the present data from one previousreport of [Ca2+]i transients in Vicia guard cell protoplasts. Schroeder and Hagiwara128

proposed that Ca2+ release might be triggered after Ca2+ entry via nonselective,Ca2+-permeable channels. However these experiments do not convince, because Ca2+-medi-ated activation of Cl- channels was not ruled out. Furthermore, exchange of the cytosolwith the patch electrode filling solution in these experiments meant that dynamic controlof [Ca2+]i was lost. By contrast, voltage clamp using intracellular microelectrodes leavesthe cytosol largely intact.68,129


Most remarkable, we found that voltage and ABA acted in concert to potentate the[Ca2+]i signal, with ABA affecting both the voltage threshold for the [Ca2+]i response andits kinetics. These observations suggest that targets of ABA action may include the voltage-dependence of gating for the Ca2+ influx pathway itself and activation of intracellular Ca2+

release elements. One explanation may be that Ca2+ channel activation occurs via proteinphosphorylation,79 although these actions of ABA are by no means the only possibilities.Furthermore, voltage appeared to be a determining factor for [Ca2+]i increases evoked byABA. Indeed, with membrane voltage under experimental control, ABA elicited anappreciable [Ca2+]i increase only once the membrane was driven to voltages near andnegative of -100 mV.

Perhaps it is not surprising, then, that increases in [Ca2+]i in (non-voltage-clamped)guard cells have never fully correlated with stomatal closure in ABA.98,100,128 Membranevoltage differs between cells and its distribution between voltage states in a population ofguard cells shows seasonal variability.21,31,38 With 1-10 mM K+ outside the voltage can besituated positive of -60 mV.31,58 Yet [Ca2+]i increases require that the membrane besituated at a voltage near or negative of -100 mV. So, the variability in [Ca2+]i

increases evoked by ABA seems very likely to be a direct consequence of the voltagestate of the plasma membrane.

The H+ Second MessengerIncreases in [Ca2+]i do not mimic the effects of ABA in controlling the K+ channels that

mediate IK,out at the guard cell plasma membrane. Thus, additional control elements distinctfrom any [Ca2+]i-related events must take part in ABA signal transduction. In fact, this classof K+ channels has proven singularly insensitive to [Ca2+]i

54,95,130 and to factors affectingupsteam intermediates of [Ca2+]i-related signal cascades, although in each case an influenceon IK,in was reported complementary to a rise in [Ca2+]i. The current has been reported to beinsensitive to G-protein agonists and antagonists such as GTPγS, GDPβS, cholera andpertussis toxins,131 to mas7, a serpentine receptor mimetic,132 and to inositol-1,4,5,-trisphosphate.111 Yet, IK,out is activated in a roughly scalar manner in the presence of ABA31,93

which can evoke as much as a 14-fold (1400%) increase in the current at any one voltage.96

Despite early indications of its potential importance in signaling,133-135 a role forcytosolic-free [H+] (pHi) has surfaced only slowly. At first stimulus-evoked changes in pHi

were linked to control of the H+-ATPase by auxin and fusicoccin136-139 and, hence, to H+ asa transported substrate rather than as a second messenger. The proposal of pHi as a majorintermediate in ABA signaling rests on two key experimental developments:

1. ABA was found to alkalinize the cytosol in guard cells58,97 as well as in themesophyll of Zea and Petroselenium;140 and

2. Experimentally-imposed changes in pHi were found to alter guard cell K+ channelsin a manner distinct from that of changes in extracellular pH.51,58

The first point is important, because the findings implicitly separated the changes inpHi from the H+- ATPase. Common dogma maintained that ABA reduces H+-ATPaseactivity141,142 and it was expected therefore that decreasing H+ extrusion via the pumpshould decrease pHi (raise [H+]i). Here was a clue that hormonal control of pHi mightdepend on more than just changes in the pool of substrate for primary transport. Thesecond point, furthermore, ascribed to pHi an action that could not be result directly fromH+ flux across the plasma membrane and, instead, called into question an alternativefunction as an internal signal to control ion channels.

Consistent with the idea of a pHi signal intermediate, “pH clamp” experiments demonstratedthat an increase in pHi is both necessary and sufficient to account for the activation of IK,out inABA.58 In every case, the current response was fully accommodated as a simple function


of [H+]i without recourse to additional controls, whether guard cells were loaded with pHbuffers or with the weak acid butyrate to suppress changes in pHi with ABA and to raise andlower pHi in the absence of ABA. Subsequent work also pointed to pHi more generally inchannel control, for example that initiated by auxin,49,143 suggesting a more central role incellular stimulus-response coupling in guard cells. Indeed, the evoked changes in pHi

recorded by Thiel, et al143—roughly 0.5 pH units within 20 s, at a conservative estimate—rivals evoked changes of [Ca2+]i in many plant cells, both in amplitude and kinetics, andremains difficult to explain in terms of H+ transport across either plasma membrane ortonoplast.

The action of pHi on IK,out shows an unusual “fingerprint”. Blatt and Armstrong58

found that increasing pHi activated IK,out in a voltage-independent manner, that is thecurrent rose roughly in proportion at all voltages, and titration of the current as a functionof pHi suggested a cooperative binding of two H+ to effect the response. These resultshave since been extended in combined voltage clamp and BCECF fluorescence ratiomeasurements of pHi

130 to demonstrate that pHi has little effect on the kinetics of IK,out

activation. Instead, it appears to mobilize K+ channels into a pool that are activated onmembrane depolarization. Miedema and Assmann144 recently confirmed theseobservations, showing that the action of pHi results from H+ interaction with bindingsites closely associated with the membrane, possibly with the channel or with closelyassociated protein(s).

The action of pHi extends beyond control of IK,out. Along with its sensitivity to [Ca2+]i

(above), IK,in—or one of the major signaling elements controlling these channels—respondsto pHi and, significantly, with a fingerprint that is fundamentally different to that of IK,out.Initial studies pointed to a pHi dependence opposing that of IK,out in this case. Thus,reducing pHi was found to activate IK,in and the current was suppressed by alkaline pHi

loads.51,58 Titration of IK,in130 has since demonstrated that the current behaves as a simple

function of [H+]i, again in contrast to the cooperativity between H+ observed for IK,out.The data indicate a H+ binding at a site with a pKa near 6.4, consistent with the titration ofa single histidine residue and consonant with several recent studies of cloned K+ channelsfrom yeast145 and mammalian neuromuscular tissues.146

One immediate question is whether pHi action in this latter instance could bemediated by H+ titration of Ca2+ binding sites, in other words by interference with Ca2+

signal transmission. The answer is negative, but this reply belies the complexity of thesituation (see pHi Interaction with [Ca2+]i below). The Ca2+ and H+ messengers can beshown to function independently, converging finally on the K+ channels to control theiractivity via two kinetically-distinct mechanisms.130 Two lines of evidence support thisconclusion:

1. pHi- mediated control of IK,in is observed in the absence of any measurable changesin [Ca2+]i; and

2. The effect of pHi is evident predominantly as a voltage-independent change in IK,in

conductance, whereas the action of [Ca2+]i is profoundly voltage-sensitive.Thus, in a very straightforward sense, the actions of pHi on IK,in can be distinguished

from those of [Ca2+]i. Not surprising then, additional studies — including work with ABA24,58

demonstrating IK,in inactivation concomitant with a rise in pHi when measurements failedto show a rise in [Ca2+]i — have indicated that changes in pHi can predominate in regulatingIK,in in leu of any changes in [Ca2+]i.

Generality of the pHi SignalThe preceding evidence aside, there occurs at least one circumstance for which pHi

does not appear to play a major role in ion channel control. Stomata normally close in


response to a rise in the partial pressure of CO2 (pCO2), much as they do in the presence ofABA. Like ABA, elevated pCO2 promotes solute efflux from the guard cells7. Indeed, CO2

and ABA interact closely in control of stomatal aperture.18,88 So, CO2 might be anticipatedto draw on a similar sequence of changes in K+ and anion channel activities to facilitatethese ion fluxes. The expectation has been borne out. Recent voltage clamp studies withVicia guard cells69 demonstrated that raising pCO2 from ambient (350 µ11-1) to 1000 µ l-1

evokes a rapid inactivation of IK,in and activation of IK,out as well as enhancing the anionchannel current and altering its voltage-dependent kinetics. Significantly, parallel measure-ments with the H+- sensitive fluorescent dye BCECF failed to uncover any measurablechange in pHi and, in 10,000 µ11-1 pCO2 a small 0.1-0.2 unit decrease in pHi was evenobserved.

The observations make sense from the physiological standpoint. A rise in pCO2

increases the bicarbonate dissolved in aqueous solution which, as a weak acid, will favorcytosolic acidification.147 Yet the effect must inevitably suppress K+ salt loss from the guardcells and stomatal closure, because decreasing pHi inactivates IK,out. So, the existence of analternative control pathway circumvents the potential dilemma otherwise inherent whenthe response to a weak acid must be to facilitate K+ efflux for stomatal closure. Becauseelevating pCO2 actually promoted the activity of IK,out even in the face of mild decreases inpHi, Brearley, et al69 concluded that the CO2 signal must be transmitted via another, as yetunidentified, signal cascade. For the present, however, the identify of this third signalingpathway is a mystery.

Origin of Changes in pHiWhat are the possible origins for these pHi changes? Because of the high buffer

capacity of guard cells [≅100 mM/(pH unit)]58,185 a pHi rise of 0.2-0.3 units, such asoccurs in ABA, requires either that approximately 30 mM H+ are eliminated from thecytosol or that a comparable increase in H+ binding and sequestration occurs within thecell. Of the possible sinks for H+, transport by the H+-ATPase across the plasma membrane canbe ruled out, because the large flux of H+ would entail substantial currents (approx. 10 µAcm-2 over 5 min). Such H+ currents have not been observed in response to ABA.23,31,93 Twoother potential sinks for H+ are vacuolar H+ transport, and metabolic H+ consumptionduring organic acid breakdown and CO2 release. Neither of these options excludes theother and both are attractive. The second possibility would tie the pHi signal to themetabolic turnover of osmolytes that is known to take place during stomatal closure. Guardcells of many higher plants, including Vicia, use malate as well as Cl- to balance K+ uptakeduring stomatal opening.7 Much of this malate is decarboxylated during closure, therebyconsuming cellular H+. Movement of H+ into the vacuole is an equally likely explanationfor the pHi rise, and could contribute to charge balance during K+ efflux from the vacuole.Roughly 50-70% of the vacuolar K+ salt content passes across the tonoplast before exitingthe guard cell during stomatal closure.7,148,149 A cursory estimate shows that to account fora 0.3 pHi unit rise, that is approximately 30 mM H+, only 2-4% of the K+ efflux need bebalanced by H+ entry into the vacuole. In fact, vacuolar acidification on this scale is knownto take place during closure.150 Recent studies also lend support to this argument:Frohnmeyer, et al78 observed that evacuated guard cells and mesophyll protoplasts lose theability to dynamically buffer pHi and show no change in pHi in response to hormonetreatments.

Protein PhosphorylationChannel regulation by phosphorylation/dephosphorylation is indisputably a third

major element coupling ABA, and probably other stimuli, to solute flux during stomatal


movements. Even before a link was established to physiologically-related events,pharmacological evidence implicated protein kinase and protein phosphatase activities inmaintaining a homeostatic balance of channel function. Regulation of IK,in by a 2B-type(Ca2+-dependent) protein phosphatase was identified through its sensitivity to FK506,cyclophilin-cyclosporin A and related compounds in patch electrode studies151 and time-dependent block of both IK,in and IK,out by low concentrations of okadaic acid implicated adependence on protein phosphatase 1/2A activity.152,153 More recent studies have indi-cated that both the tonoplast SV channel79 and the tonoplast anion channel77 are alsosubject to control by phosphorylation. Because ABA, among other stimuli, activates Ca2+-dependent protein kinases,154,155 it is conceivable that ABA (and Ca2+) action could bemediated in part through a cascade of protein (de-)phosphorylation.

In the event, direct evidence for protein phosphatase (and by inference, proteinkinase) activity in ABA signaling has come from combined molecular genetic and electro-physiological studies of plants carrying the mutant abi1 gene of Arabidopsis. The abi1 genewas originally identified as one of a family of abscisic acid-insensitive mutants that germi-nated and grew on high concentrations of ABA.156 It is linked to a number of ABA-relatedphenotypes, including an aberrant control of stomatal aperture and consequent wiltycharacteristic. The ABI1 amino-acid sequence indicated that the protein is type 2Cprotein (serine/threonine) phosphatase (PP-2C),157,158 and subsequent work demonstratedthat the ABI1 gene product displays Mg2+-dependent protein phosphatase activity but isCa2+-insensitive.159 In vivo, the wild-type gene will partially complement a yeast PP-2Cmutant, and the mutant abi1 (dominant negative) gene confers a subset of phenotypes intransgenic Nicotiana similar to those of the Arabidopsis mutant, including the strongtendency to wilt.24 Thus, by association, the abi1 phenotype implicates protein phos-phorylation in ABA-mediated control of guard-cell ion transport.

Armstrong, et al24 have since linked the mutant phenotype with aberrant control ofthe guard cell K+ channels using abi1-transgenic tobacco. These studies showed that theabi1 gene not only reduced IK,out in the absence of ABA, but eliminated the response ofthe current in its presence. The transgene was also found to interfere with ABA-evokedcontrol of IK,in and to block stomatal closure. Furthermore, the background of IK,out

activity, as well as IK,in and IK,out responses to ABA and stomatal closure could be “rescued” bytreatments with protein kinase antagonists. So, how can abi1 gene action be understood?The simplest explanation is that the (dominant) mutant protein phosphatase interfereswith a wild type homologue in tobacco preventing protein dephosphorylation. Thekinase antagonists, then, redress the consequent imbalance in phosphorylation of thetarget protein(s) to reinstate K+ channel sensitivity to ABA. This interpretation isentirely consistent with the phosphatase activities shown by the wild type and mutantgene products.159

Protein (de-)phosphorylation also affects anion channel function. Evidence frompharmacological studies have shown that protein phosphatase antagonists such as okadaicacid and calyculin A (both protein phosphatase 1/2A antagonists) alter anion channelcharacteristics152 and rundown in patch electrode recordings of guard cell protoplasts.160

Recently, ICl was found to be activated by ABA in guard cells of N. benthamiana,23 andArabidopsis25 in a manner that depended on the phosphorylation status of the cell. Pei, etal25 recorded anion currents from protoplasts after obtaining patch seals either in the pres-ence or absence of ABA and/or with additions of okadaic acid. Statistical comparisonsindicated that the current was activated by ABA and that this activation could be suppressed byokadaic acid. The effect appeared to be a purely scalar increase in the number of channelsin the active pool. Furthermore, the ABA-mediated increase in ICl was notobserved in abi1 mutant plants or in Arabidopsis carrying the mutant abi2 gene that


encodes an abi1 homologue,161 although only in the former case was the impairmentrescued in the presence of the kinase antagonist K252A. Grabov et al,23 by contrast, madeuse of intracellular microelectrode methods, recording anion currents from individualguard cells before and throughout treatments with ABA and calyculin A. They found thatABA reversibly stimulated the anion current and that this stimulation was associated withalterations in the kinetics of voltage-dependent activation that favored the active state ofthe channels. Grabov, et al also observed that calyculin A acted synergistically with ABA inpromoting the anion current, and that the effects in every case were independent of theabi1 transgene.

How can these two sets of data be reconciled? Pei et al based their assessment on amistaken comparison of the Arabidopsis anion current in the presence of ABA with theN. benthamiana current recorded in the absence of ABA. In fact, resting activities ofthe anion channels were roughly equivalent, despite the obvious difference ofspecies and very different methods used to record these currents. It is likely that subtledifferences in kinase/phosphatase cascades mean that Arabidopsis and N. benthamianarespond differently to protein phosphatase antagonists (compare also ref. 160), andthis factor might well explain the discrepancy in action of the abi1 gene product. Equallyimportant, technical differences—including exchange of the cytosol with patch electrodefilling solutions36—raises the possibility that different parts of the same kinase/phosphatase cascade could be favored in each case. Regardless of these complications,the fact that both sets of experiments demonstrate an action of ABA on the anioncurrent is salutary.

Interaction of Signaling ElementsEven when evoked by a single stimulus such as ABA, signaling pathways interact in a

manner that can be envisaged to spread its perturbation like a ripple in a pond through anetwork of interrelated connections.162 Sufficient evidence is now to hand to demonstratesuch interactions between signaling elements of protein (de-)phosphorylation, [Ca2+]i andpHi. At present, the precise points of interaction are minimally defined, as is the extent towhich the various signals are interdependent. These issues are currently a major focus ofour attention. Their resolution will be particularly relevant both to determining thehierarchy of events behind stimulus-response coupling per se, and to establishing the role(s)of each element, whether primary to signal transmission or secondary in adaptiveconditioning of the response.

pHi Interaction with [Ca2+]iA case in point can be drawn from work on pHi and [Ca2+]i signals evoked by ABA.

From analysis of the current kinetics and voltage-dependence, it is evident that pHi and[Ca2+]i controls of IK,in are fundamentally different (see The H+ Second Messenger above).Nonetheless, these two signaling elements almost certainly interact. Grabov and Blatt130

observed that experimentally lowering pHi to values below about 7.0 resulted in a rise of[Ca2+]i in roughly 50% of the cells examined. It is significant, too, that these [Ca2+]i

increases did not correlate with the timecourse of pHi loads as might be expected for simple,bulk titration of Ca2+ binding sites. Instead it appears that pHi may “prime” signalingelements that mediate in [Ca2+]i control and indirectly trigger a rise in [Ca2+]i

8. Theprecise mechanism of this pH-induced [Ca2+]i rise is not currently known, however severalexperimentally testable scenarios are now worth examining. One possible explanationlies in the pH-sensitivity of Ca2+ release mechanisms within the cell, for example thebinding of IP3 to its receptor that leads to cytosolic Ca2+ release,163 although in animals itis normally alkaline rather than acid pHi that favors IP3 mediated Ca2+ release.164 In the


guard cells pHi is known to affect the gating of vacuolar ion channels that may contributeto [Ca2+]i changes either directly or indirectly.74,75 The SV channels, especially, aresensitive to pHi

74 with acid pHi favoring vacuolar Ca2+ release. It is also possible that pHi

could affect Ca2+ influx across the plasma membrane (see Fig. 6.1). The action of pHi ineach of these instances affects events upstream of the [Ca2+]i messenger itself, althoughthe associated mechanisms are different.

Equally, cytosolic-free Ca2+ may influence pHi. Recent studies (Fricker M, Wood J,personal communication) have suggested that increasing [Ca2+]i, by promoting Ca2+

influx across the plasma membrane, can trigger increases and fluctuations in pHi thatpersist over many minutes. These measurements were carried out using confocalratio fluorescence imaging techniques and also indicate a spatial heterogeneity to thepHi changes. Because local [Ca2+]i “hot spots” are though to underpin Ca2+-inducedCa2+ release in many cell types,122 we may speculate that complementary pHi domainsalso occur within cells, either superimposed on or associated with local changes in[Ca2+]i. The validity of such a premise certainly deserves attention. But whether or notcorrect, it is already evident that the interaction between [Ca2+]i and pHi is not simplybilateral: whereas decreasing pHi may promote a rise in [Ca2+]i, the converse ofincreasing [Ca2+]i appears to favor pHi increases.

[Ca2+]i, pHi and Protein PhosphorylationElevation of [Ca2+]i is probably closely linked to protein phosphorylation in plants as

it is in animals. Principal points of convergence are protein kinases and phosphatasesthat dependent on [Ca2+]i. In plant cells there is now considerable evidence for Ca2+-,Ca2+/calmodulin- and Ca2+/phospholipid- dependent kinases and phophatases.79,151,165-168

Calcium-dependent protein kinase activity is known to activate an ABA-responsivepromoter in Arabidopsis leaf protoplasts169 and a Cl- channel in the guard cell tonoplast.77

By contrast, the ABI1 gene product has proven to be insensitive to [Ca2+]i,159 although it

contains a putative EF-hand Ca2+-binding motif and was originally suspected to beCa2+-regulated.157,158

Protein phosphorylation has now been shown to affect pHi signal transmission inguard cells. Blatt and Grabov8 have reported that the abi1 transgene in N. benthamianadrastically reduces K+ channel sensitivity to experimentally-imposed changes in pHi. Theyfound that lowering pHi with 3 and 10 mM butyrate, equivalent to decreases of 0.3-0.5 pHi

units, had only marginal effects on the K+ currents in the transgenic plants compared withthe wild type. The observations are significant, because previous work with these plants24

had demonstrated that the transgene affected K+ channel response to ABA but had noinfluence on the rise in pHi evoked by the hormone. So, together, these two sets of dataconfirm that events downstream of pHi are dependent on the phosphorylation state ofone or more target proteins.

Initial Events in ABA Stimulus PerceptionBy contrast with our current knowledge of downstream signaling events and their

targets that mediate solute transport in guard cells, no substantive information existsrelating to the ABA receptor itself. The debate surrounding the localization of the ABAreceptor has itself failed to resolve the most basic question—whether the receptor is cyto-solic or situated in the plasma membrane and exposed to the external environment—andhas further fueled controversy. It is possible that different ABA receptors situated at thecell periphery and within the cell all contribute to ABA perception.170 However resolvingthis, and related issues will now require that the receptor gene(s) are cloned andcharacterized.92


Localization of ABA ReceptorsThe results of early attempts to identify the nature of the ABA receptor suggested that

the hormone was perceived outside at the cell surface.109 These arguments centered aroundthe fact that ABA should partition as a weak acid across the plasma membrane in apH-dependent manner171-173 and therefore should be more effective at acid than at alkalineexternal pH if the receptor were located inside the cell. Hornberg and Weiler174 reporteddirect ABA binding to a plasma membrane proteins using photo affinity methods, but thefailure of other groups to duplicate these results thereafter was a setback. Only recentlyhave more conventional biochemical approaches yielded indications of specific ABA-bindingproteins.175

Other studies exploring the subcellular localization of ABA perception site(s) in guardcells have used advanced in vivo techniques. Schwartz, et al176 reported a close correlationbetween 3H-ABA uptake and stomatal closure in Commelina guard cells. They also foundthat microinjecting ABA into individual guard cells promoted stomatal closure whileinclusion of ABA in patch electrodes suppressed IK,in of Vicia guard cell protoplasts. Parallelstudies,102 using a biologically-inert “caged”-ABA, also implicated an internal perceptionsite for ABA. In this case, microinjection and UV photolysis of the caged compound torelease active ABA promoted stomatal closure even when the epidermal strip was continuouslyperfused with ABA-free solution. Other work has indicated that ABA may be perceived atthe cell surface. Anderson, et al177 reported that intracellular microinjections of ABA alonewas insufficient to prevent stomatal opening in Commelina, although stomatal openingwas suppressed when ABA was provided externally. (Note that in a different tissuepreparation, in this case from barley aleurone, Gilroy and Jones178 found that -amylasesecretion that is normally stimulated by gibberelin and antagonized by ABA could not besuppressed when ABA was first microinjected into the cells.)

The more trivial explanations of differences in plant material aside, the apparentcontradiction between these two data sets might be explained by differences in experimentaldesign. Evidence of an internal site for ABA action was supported by experiments thatcentered on stomatal closure, whereas arguments for an external site of action wereindicated in experiments designed to show the ability of ABA to suppress other activities,either stomatal opening or amylase secretion. Promotion of stomatal closure and inhibition ofstomatal opening involve partially-separable mechanisms.179 So, it is possible that morethan one site of perception exists, even for short-term events that (presumably) do notrequire transcription/translation activities. In fact, this latter idea has found support inrecent radiotracer flux studies. MacRobbie70 observed differences in the efficacy and tim-ing for ABA flux response in Commelina guard cells. She found that under suboptimalconditions a delay in vacuolar efflux could be introduced into 86Rb+(K+) efflux measurementsthat were separable from the initial flux transient across the plasma membrane. It ispossible that intervening signaling steps—and differences in their kinetics—couldaccount for such differences in response time and ABA dependence. However, at least oneexplanation is that ABA binds with different affinities and at separate sites associated withthe plasma membrane and tonoplast.

Is the ABA Signal G-Protein Coupled?In lieu of direct evidence for ABA receptor binding, downstream events immediately

following ABA-receptor interaction have been sought that couple to late signalingintermediates such as [Ca2+]i increases and could yield some information about thenature of the receptor protein. Widespread interest has focused on the possible role for aserpentine (7-transmembrane-segment) receptor protein that could couple through theactivation of GTP-binding proteins (G-proteins).180-182 There is certainly some evidence


to support a role for heterotrimeric G-proteins in regulating IK,in. However, none of thesedata can be tied to pHi regulation of the current, nor are they wholly consistent withcoupling even through the [Ca2+]i intermediate. In their original studies, Fairley andAssmann131 showed that IK,in was sensitive to the G-protein antagonists cholera andpertussis toxins in Vicia guard cell protoplasts; the current was inactivated in the presenceof the non-hydrolyzable GTP analogue GTP-γ-S and this inactivation was overcome bybuffering changes in [Ca2+]i. These studies indicated that control of IK,in could passfrom Gα activation and, plausibly, stimulation of inositol-1,4,5- trisphosphate productionby phospholipase C to a rise in [Ca2+]i. Subsequently, Wu and Assmann183 added evidencefor coupling via a membrane-delimited Gα, and Kelly, et al104 have suggested that [Ca2+]i

may exert a dual effect with G-proteins on IK,in.G-proteins may also control IK,in activity independent of [Ca2+]i. Armstrong and

Blatt132 found that mas7, that mimics serpentine receptors and promotes G release,inactivated IK,in. Mas7 action was blocked by GDP-β-S, consistent with Gα control of IK,in.However, the effect was not linked to [Ca2+]i, as mas7 action could not be overcome bycytosolic Ca2+ buffering or by neomycin sulfate, an antagonist of phospholipase C andinositol- 1,4,5-trisphosphate-mediated Ca2+ release. Nor could evidence be found for mas7action passing through the pHi intermediate. Unlike the effects on auxin- and ABA-evokedresponses of the K+ channels, treatments with weak acid to clamp pHi near 7.0 failedto rescue IK,in from inactivation by mas7. Furthermore, mas7 had no influence on IK,out

which is profoundly pHi sensitive (see The H+ second messenger above). In fact, thereremains an overwhelming lack of evidence to support a function for G-proteins inhormonal responses of guard cells per se, whether coupled through [Ca2+]i or pHi.Thus, it appears ever more probable that relevant signaling events upstream in thesecells could differ from the classic G-protein-receptor model.

Perspectives and ConclusionThe complexity of signaling events that underlie stomatal response to ABA should

come as little surprise as guard cells integrate both environmental and internal signalsto achieve stomatal control. As the primary defense of the plant against water loss anddehydration, it is imperative that transport functions in guard cells are finely tuned tothese needs along with those of competing demands for CO2. However, our awareness ofthe “stomatal situation” arises from a physiological perspective and, most recently, fromthe advantages afforded by the interface between molecular genetics and biophysics. It islikely that many of these control networks, like the transporters that they regulate, arecommon among higher-plant cells. Indeed, of the pumps and ion channels that effectsolute movement for stomatal control—and these are, with several notable exceptions,now well documented at the plasma membrane—counterparts are to be found in arange of higher-plant cells in every case. Admittedly, our understanding of transportacross the tonoplast remains very poor and much more work is needed here to identify themajor pathways for solute movements and their respective functions. Nonetheless, at presentthere is little basis to set vacuolar transport in guard cells apart from other higher-plantcell types.

Our knowledge of the mechanisms controlling ion channels and, even more still, theH+-ATPase at the plasma membrane signal integration is unquestionably in the early stagesof development. Still less is known of transport regulation at the tonoplast, although thebulk of osmotic solutes must pass across this latter membrane. The vacuole is also likely tocomprise an important reservoir and sink for Ca2+ and H+, so separating transportand signaling functions will be especially important. A role for [Ca2+]i in associationwith ABA is certain but, remarkably, major questions still hang over its situation within


the ABA-signaling network, its origin(s), downstream targets and mechanism(s) ofaction. The emergence of pHi as a second messenger, and its interaction with the abi1protein phosphatase, further underscores our relative ignorance about the scope andnature of events both downstream and upstream within each respective pathway. Significantly,here is a second messenger for which we may find no direct counterpart in animal cellsand no preconceived model to frame our ideas.

Unquestionably, major advances in resolving each of these issues will now be aided bya combination of molecular genetic and biophysical tools. Once the identity of a first ABAreceptor has been established, relationships between many signaling elements should fallin place while still others may only then become evident. But almost certainly somecurrent perspective on the issues and their possible solutions will appear naive by theclose of this decade. We may echo JBS Haldane’s suspicion “that the universe is not onlyqueerer than we suppose, but queerer than we can suppose.”184

AcknowledgmentsWe are grateful to Mark Fricker (Oxford) for sharing unpublished results with us.

This work was supported by the Gatsby Charitable Foundation, Human Frontiers ScienceProgram grant RG303/95M and EC Biotech grant CT960062. AG is a Senior ResearchAssociate funded on BBSRC grant 32/C098-1. BL was a Sainsbury Ph.D. Student and iscurrently funded on BBSRC grant 32/C08406.

References1. Loveys BR. How useful is a knowledge of ABA physiology for crop improvement? In:

Davies WJ, Jones HG, eds. Abscisic Acid Physiology and Biochemistry. Oxford, BiosScientific. 1991:245-260.

2. Flowers T, Yeo M. Salt Tolerant Plants. In: Baker DA, Hall JL, eds. Solute Transport inPlant Cells and Tissues. Harlow, Longman Press. 1988:326-355.

3. Bradford KJ, Hsiao TC. Stomatal behavior and water relations of waterlogged tomatoplants. Plant Physiology 1982; 70:1508-1513.

4. Jackson MB. Regulation of water relationships in flooded plants by ABA from leaves,roots and xylem sap. In: Davies WJ, Jones HG, eds. Abscisic Acid Physiology andBiochemistry. Oxford, Bios Scientific. 1991:217-226

5. Davies WJ, Jones HG. Abscisic Acid: Physiology and Biochemistry. Bios Scientific, Oxford.1991:1-266.

6. Nobel PS. Physicochemical and Environmental Plant Physiology. Academic Press, SanDiego. 1991:1-413.

7. Willmer C, Fricker MD. Stomata. Chapman and Long, London. 1996:1-375.8. Blatt MR, Grabov, A. Signaling gates in abscisic acid-mediated control of guard cell ion

channels. Physiologia Plantarum 1997; 100:481-490.9. Malpighi M. Anatome Plantarum. Royal Society, London.1675:1-130.

10. Grew N. Anatomy of Plants. London. 1682:1-312.11. de Candolle AP. Organographie Végétale ou description raisone des organes des plantes.

Deterville, Paris. 1827:1-213.12. von Mohl H. Welche Ursachen bewirken die Erweiterung und Verengung der

Spaltoeffnungen? Bot Zeit. 1856; 14:697-720.13. Immamura S. Untersuchungen uber den mechanismus der turgorschwankung der

Spaltoeffnungschlieszellen. Jap J Bot. 1943; 12:251-346.14. Yamashita T. Influences of potassium supply upon various properties and movement of

guard cell. Sieboldia Acta Biol. 1952; 1:51-70.15. Fujino M. Role of ATP and ATPase in stomatal movement. Sci Bull Fac Ed Nagasaki

Univ. 1967; 18:1-47.


16. Fischer RA. Stomatal opening: Role of potassium uptake by guard cells. Science 1968;160:784-785.

17. Fischer R, Hsiao T. Stomatal opening in isolated epidermal strips of Vicia faba: II.Response to KCl concentration and role of potassium absorption. Plant Physiol. 1968;43:1953-1958.

18. Raschke K. The stomatal turgor mechanism and its response to CO2 and abscisic acid:observations and hypothesis. In: Marre E, Cifferrri O, eds. Regulation of cell membraneactivities in plants. Amsterdam, Elsevier. 1977:173-183.

19. Raschke K. Movements of stomata. In: Haupt W, Feinleib ME, eds. Encyclopedia of PlantPhysiology, N.S. Vol. 7. Berlin, Springer. 1979:373-441.

20. Brindley HM. Fluxes of 86Rb+ in “isolated” guard cells of Vicia faba L. Planta 1990;181:432-439.

21. Blatt MR. Electrical characteristics of stomatal guard cells: The ionic basis of the mem-brane potential and the consequence of potassium chloride leakage from microelectrodes.Planta 1987; 170:272-287.

22. Wille A, Lucas W. Ultrastructural and histochemical studies on guard cells. Planta 1984;160:129-142.

23. Grabov A, Leung J, Giraudat J, Blatt MR. Alteration of anion channel kinetics in wildtype and abi1-1 transgenic Nicotiana benthamiana guard cells by abscisic acid. Plant Journal1997; 12:203-213.

24. Armstrong F, Leung J, Grabov A, Brearley J, Giraudat J, Blatt MR. Sensitivity to abscisicacid of guard cell K+ channels is suppressed by abi1-1, a mutant Arabidopsis gene encoding aputative protein phosphatase. Proc Natl Acad Sci USA. 1995; 92:9520-9524.

25. Pei ZM, Kuchitsu K, Ward JM, Schwarz M, Schroeder JI. Differential abscisic acidregulation of guard cell slow anion channels in Arabidopsis wild type and abi1 and abi2mutants. Plant Cell. 1997; 9:409-423.

26. Roelfsema MRG, Prins HBA. Ion channels in guard cells of Arabidopsis thaliana (L.)Heynh. Planta. 1997; 202:18-27.

27. Raschke K, Firn RD, Pierce M. Stomatal closure in response to xanthoxin and abscisicacid. Planta. 1975; 125:149-160.

28. Hentzen AE, Smart LB, Wimmers LE, Fang HH, Schroeder JI, Bennett AB. 2 plasma-membrane H+-ATPase genes expressed in guard-cells of Vicia faba are also expressedthroughout the plant. Plant And Cell Physiology 1996; 37:650-659.

29. Becker D, Zeilinger C, Lohse G, Depta H, Hedrich R. Identification and biochemicalcharacterization of the plasma membrane H+-ATPase in guard cells of Vicia faba L.Planta. 1993; 190:44-50.

30. Villalba JM, Lutzelschwab M, Serrano R. Immunocytolocalization of plasma membraneH+-ATPase in maize coleoptiles and enclosed leaves. Planta. 1991; 185:458-461.

31. Thiel G, MacRobbie EAC, Blatt MR. Membrane transport in stomatal guard cells: Theimportance of voltage control. J Membr Biol. 1992; 126:1-18.

32. Blatt MR, Clint GM. Mechanisms of fusicoccin action kinetic modification and inactivationof potassium channels in guard cells. Planta. 1989; 178:509-23.

33. Blatt MR. Electrical characteristics of stomatal guard cells: The contribution of ATP-dependent, “electrogenic” transport revealed by current-voltage and difference-cur-rent-voltage analysis. J Membr Biol. 1987; 98:257-274.

34. Serrano EE, Zeiger E, Hagiwara S. Red light stimulates an electrogenic proton pump inVicia guard cell protoplasts. Proc Natl Acad Sci USA 1988; 85:436-440.

35. Assmann SM, Simoncini L, Schroeder JI. Blue light activates electrogenic ion pumpingin guard cell protoplasts of Vicia faba. Nature 1985; 318:285-287.

36. Pusch M, Neher E. Rates of diffusional exchange between small cells and a measuringpatch pipette. Pfluegers Archiv, Euro J Physiol 1988; 411:204-214.

37. Schroeder, J.I. 1988. K+ transport properties of K+ channels in the plasma membrane ofVicia faba guard cells. J Gen Physiol. 1988; 92:667-683.

38. Lohse G, Hedrich R. Characterization of the plasma-membrane H+- ATPase from Viciafaba guard cells. Planta. 1992; 188:206-214.


39. Slayman CL, Bertl A, Blatt MR. Partial reaction chemistry and charge displacement bythe fungal plasma-membrane H+-ATPase. In: Hirst BH, ed. Molecular and Cellularmechanisms of H+ Transport. Berlin, Springer Verlag. 1994:237-244.

40. Blatt MR, Beilby MJ, Tester M. Voltage dependence of the Chara proton pump revealedby current-voltage measurement during rapid metabolic blockade with cyanide. J MembrBiol. 1990; 114:205-223.

41. Thiel G, Wolf AH. Operation of K+ channels in stomatal movement. Trends In PlantScience 1997; 2:339-345.

42. Clint GM, Blatt MR. Mechanisms of fusicoccin action: evidence for concerted modulationsof secondary K+ transport in a higher-plant cell. Planta. 1989; 178:495-508.

43. Rodriguez-Navarro A, Blatt MR, Slayman CL. A potassium-proton symport in Neurosporacrassa. J Gen Physiol. 1986; 87:649-674.

44. Maathuis FJM, Sanders D. Mechanisms of potassium absorption by higher-plant roots.Physiologia Plantarum 1996; 96:158-168.

45. Nakamura RL, Mckendree WL, Hirsch RE, Sedbrook JC, Gaber RF, Sussman MR.Expression of an Arabidopsis potassium channel gene in guard cells. Plant Physiology1995; 109:371-374.

46. Very A-A, Gaymard F, Bosseux C, Sentenac H, Thibaud J-P. Expression of a cloned plantK+ channel in Xenopus oocytes: analysis of macroscopic currents. Plant J. 1995; 7:321-332.

47. Schroeder JI, Schachtman DP, Lucas WJ, Anderson JA, Gaber RF. The Arabidopsis KAT1cDNA confers functional expression of an inward-rectifying potassium channel inXenopus oocytes. Biophysical Journal 1993; 64:196-196.

48. Bertl A, Anderson JA, Slayman CL, Gaber RF. Use of Saccharomyces-cerevisiae forpatch-clamp analysis of heterologous membrane proteins—characterization of KAT1, aninward-rectifying K+ channel from Arabidopsis thaliana, and comparison with endogeneousyeast channels and carriers. Proc Natl Acad Sci USA. 1995; 92:2701-2705.

49. Blatt MR, Thiel G. K+ channels of stomatal guard cells: Bimodal control of the K+

inward-rectifier evoked by auxin. Plant J. 1994; 5:55-68.50. Schroeder JI, Fang HH. Inward-rectifying K+ channels in guard cells provide a mechanism

for low-affinity K+ uptake. Proc Natl Acad Sci USA 1991; 88:11583-11587.51. Blatt MR. K+ channels of stomatal guard cells: characteristics of the inward rectifier and

its control by pH. J Gen Physiol. 1992; 99:615-644.52. Obermeyer G, Armstrong F, Blatt MR. Selective block by α-dendrotoxin of the K+

inward rectifier at the Vicia guard cell plasma membrane. J Membr Biol. 1994;137:249-259.

53. Blatt MR, Gradmann D. K+-sensitive gating of the K+ outward rectifier in Vicia guardcells. J Membr Biol 1997; 158:241-256.

54. Hosoi S, Iino M, Shimazaki K. Outward-rectifying K+ channels in stomatal guard cellprotoplasts. Plant Cell Physiol. 1988; 29:907-911.

55. Schroeder JI, Raschke K, Neher E. Voltage dependence of K+ channels in guard-cellprotoplasts. Proc Natl Acad Sci USA 1987; 84:4108-4112.

56. Schroeder JI. Quantitative analysis of outward rectifying K+ channel currents in guardcell protoplasts from Vicia faba. J Membr Biol. 1989; 107:229-235.

57. Blatt MR. Potassium-dependent bipolar gating of potassium channels in guard cells. JMembr Biol. 1988; 102:235-246.

58. Blatt MR, Armstrong F. K+ channels of stomatal guard cells: abscisic acid-evoked controlof the outward rectifier mediated by cytoplasmic pH. Planta. 1993; 191:330-341.

59. Allen GJ, Sanders D. Vacuolar ion channels of higher plants. Advances In BotanicalResearch Incorporating Advances In Plant Pathology 1997; 25:217-252.

60. Hedrich R, Busch H, Raschke K. Ca2+ and nucleotide-dependent regulation of voltagedependent anion channels in the plasma membrane of guard cells. EMBO J. 1990;9:3889-3892.

61. Schulz-Lessdorf B, Lohse G, Hedrich R. GCAC1 recognizes the pH gradient across theplasma membrane: A pH-sensitive and ATP-dependent anion channel links guard cellmembrane potential to acid and energy metabolism. Plant Journal 1996; 10:993-1004.


62. Linder B, Raschke K. A slow anion channel in guard cells, activating at large hyper-polarization, may be principal for stomatal closing. FEBS Lett. 1992; 313:27-30.

63. Schroeder JI, Keller BU. Two types of anion channel currents in guard cells with distinctvoltage regulation. Proc Natl Acad Sci USA 1992; 89:5025-5029.

64. Dietrich P, Hedrich R. Interconversion of fast and slow gating modes of GCAC1, a guardcell anion channel. Planta 1994; 195:301-304.

65. Thomine S, Zimmermann S, Guern J, Barbier-Brygoo H. ATP-dependent regulation ofan anion channel at the plasma membrane of protoplasts from epidermal cells ofArabidopsis hypocotyls. Plant Cell. 1995; 7:2091-2100.

66. Artalejo CR, Ariano MA, Periman RL, Fox AP. Activation of facilitation calcium channels inchromaffin cells by D1 dopamine receptors through a cAMP/protein kinase A- dependentmechanism. Nature. 1990; 348:239-242.

67. Song Y, Huang L-YM. Modulation of glycine receptor chloride channels by cAMP-dependent protein kinase in spinal trigeminal neurons. Nature 1990; 348:242-245.

68. Blatt MR, Thiel G. Hormonal control of ion channel gating. Ann Rev Plant Physiol MolBiol. 1993; 44:543-567.

69. Brearley J, Venis MA, Blatt MR. CO2-mediated control of K+ and anion channels in guardcells of Vicia. Planta. 1997; 203:145-154.

70. MacRobbie EAC. Effects of ABA on 86Rb+ fluxes at plasmalemma and tonoplast ofstomatal guard cells. Plant Journal. 1995; 7:835-843.

71. MacRobbie EAC. ABA-induced ion efflux in stomatal guard-cells—multiple actions ofaba inside and outside the cell. Plant Journal. 1995; 7:565-576.

72. Fricker MD, Willmer CM. Nitrate-sensitive ATPase activity and proton pumping in guadcell protoplasts of Commelina. J Exp Botany 1990; 41:193-198.

73. Willmer CM, Grammatikopoulos G, Lasceve , Vavasseur A. Characterization of thevacuolar-type H+-ATPase from guard cell protoplasts of Commelina. J Exp Botany 1995;46:383-389.

74. Schulz-Lessdorf B, Hedrich R. Protons and calcium modulate SV-type channels in thevacuolar lysosomal compartment—channel interaction with calmodulin inhibitors. Planta.1995; 197:655-671.

75. Ward JM, Schroeder JI. Calcium-activated K+ channels and calcium- induced calciumrelease by slow vacuolar ion channels in guard-cell vacuoles implicated in the control ofstomatal closure. Plant Cell. 1994; 6:669-683.

76. Tikhonova LI, Pottosin II, Dietz KJ, Schonknecht G. Fast-activating cation channel inbarley mesophyll vacuoles. Inhibition by calcium. Plant Journal. 1997; 11:1059-1070.

77. Pei ZM, Ward JM, Harper JF, Schroeder JI. A novel chloride channel in Vicia faba guardcell vacuoles activated by the serine/threonine kinase, CDPK. Embo Journal 1996;15:6564-6574.

78. Frohnmeyer H, Grabov A, Blatt MR. A role for the vacuole in auxin-mediated control ofcytoplasmic pH by Vicia mesophyll and guard cell protoplasts. Plant J. 1998; 13:1098-116.

79. Allen GJ, Sanders D. Calcineurin, a type 2B protein phosphatase, modulates the Ca2+-permeable slow vacuolar ion channel of stomatal guard cells. Plant Cell 1995; 7:1473-1483.

80. Little CHA, Eidt DC Effect of abscisic acid on bud break and transpiration in woodyspecies. Nature. 1968; 220:498-499.

81. Mittelheuser CJ, van Steveninck RFM. Stomatal closure and inhibition of transpirationinduced by (RS)-abscisic acid. Nature. 1969; 221:281-282.

82. Wright STC. An increase in the “inhibitor-B” content of detached wheat leaves followinga period of wilting. Planta. 1969; 86:10-20.

83. Wright STC, Hiron, RWP. (+)-Abscisic acid, the growth inhibitor induced in wheat leavesby a period of wilting. Nature. 1969; 224:719-720.

84. Harris MJ, Outlaw WHJ, Mertens R, Weiler E. Water-stress-induced changes in theabscisic acid content of guard cells and other cells of Vicia faba L. leaves as determinedby enzyme-amplified immunoassay. Proc Natl Acad Sci USA 1988; 85:2584-2588.


85. Cheeseman JM. Patchy: Simulating and visualizing the effects of stomatal patchiness onphotosynthetic CO2 exchange studies. Plant Cell Environ. 1991; 14:593-599.

86. Daley PF, Raschke K, Ball JT, Berry JA. Topography of photosynthetic activity of leavesobtained from video images of chlorophyll flourescence. Plant Physiol. 1989; 90:1233-1238.

87. Atkinson CJ, Mansfield TA, Kean AM, Davies WJ. Control of stomatal aperture bycalcium in isolated epidermal tissue and whole leaves of Commelina communis L. NewPhytol 1989; 111:9-17.

88. Snaith PJ, Mansfield TA. Control of the CO2 response of stomata by indol-3ylacetic acidand abscisic acid. J Exptl Bot. 1982; 33:360-365.

89. Gorton HL, Williams WE, Assmann SM. Circadian rhythms in stomatal responsivenessto red and blue light. Plant Physiol. 1993; 103:399-406.

90. Peng ZY, Weyers JDB. Stomatal sensitivity to abscisic acid following water-deficit stress.J Exp Botany 1994; 45:835-845.

91. Harris MJ, Outlaw WH, Jr. Rapid adjustment of guard-cell abscisic acid levels to currentleaf-water status. Plant Physiol. 1991; 95:171-173.

92. Leyman B, Blatt MR. Towards cloning of an abscisic acid receptor. Institute Juan MarchCRISB 1997; 60:74.

93. Blatt MR. Potassium channel currents in intact stomatal guard cells: Rapid enhancementby abscisic acid. Planta. 1990; 180:445-455.

94. Goh CH, Oku T, Shimazaki K. Properties of proton-pumping in response to blue-lightand fusicoccin in guard-cell protoplasts isolated from adaxial epidermis of Vicia leaves.Plant Physiology 1995; 109:187-194.

95. Schroeder JI, Hagiwara S. Cytosolic calcium regulates ion channels in the plasmamembrane of Vicia faba guard cells. Nature. 1989; 338:427-430.

96. Lemtiri-Chlieh F, MacRobbie EAC. Role of calcium in the modulation of Vicia guardcell potassium channels by abscisic acid: a patch-clamp study. J Membr Biol. 1994;137:99-107.

97. Irving HR, Gehring CA, Parish RW. Changes in cytosolic pH and calcium of guard cellsprecede stomatal movements. Proc Natl Acad Sci USA 1992; 89:1790-1794.

98. McAinsh MR, Brownlee C, Hetherington AM. Visualizing changes in cytosolic-free Ca2+

during the response of stomatal guard cells to abscisic acid. Plant Cell. 1992; 4:1113-1122.99. Fricker MD, Gilroy S, Read ND, Trewavas AJ. Visualisation and measurement of the

calcium message in guard cells. In: Schuch W, Jenkins G, eds. Molecular Biology of Plantdevelopment. Cambridge, Cambridge Univ. Press. 1991:177-190.

100. Gilroy S, Fricker MD, Read ND, Trewavas AJ. Role of calcium in signal transduction ofCommelina guard cells. Plant Cell. 1991; 3:333-344.

101. McAinsh MR, Brownlee C, Hetherington AM. Abscisic acid-induced elevation of guardcell cytosolic Ca2+ precedes stomatal closure. Nature 1990; 343:186-188.

102. Allan AC, Fricker MD, Ward JL, Beale MH, Trewavas AJ. Two transduction pathwaysmediate rapid effects of abscisic acid in Commelina guard cells. Plant Cell. 1994;6:1319-1328.

103. Grabov, A. and Blatt, M.R. Ca2+-induced Ca2+ release triggered by membrane voltageand abscisic acid in guard cells of Vicia faba. Proc Natl Acad Sci USA. 1998; 95:4778-4783.

104. Kelly WB, Esser JE, Schroeder JI. Effects of cytosolic calcium and limited, possible dual,effects of G-protein modulators on guard cell inward potassium channels. Plant Journal1995; 8:479-489.

105. Cobbold PH, Rink TJ. Flourescence and bioluminescence measurement of cytoplasmicfree calcium. Biochem J. 1987; 248:313-328.

106. Fabiato A, Fabiato F. Calculator programs for computing the composition of solutionscontaining multiple metals and ligands used for experiments in skinned muscle cells. JPhysiol. 1979; 75:463-505.

107. Pethig R, Kuhn M, Payne R, Adler E, Chen T-H, Jaffe LF. On the dissociation constantsof BAPTA-type calcium buffers. Cell Calcium. 1989; 10:491-498.

108. Foehr G, Warchol W, Gratzl G. Calculation and control of free divalent cations insolutions used for membrane fusion studies. Methods Enzymol. 1993; 221:149-157.


109. MacRobbie EAC. Calcium-dependent and calcium-independent events in the initiationof stomatal closure by abscisic acid. Proc Roy Soc Lond B Biol Sci. 1990; 241:214-219.

110. McAinsh MR, Brownlee C, Hetherington AM. Partial inhibition of ABA-induced stomatalclosure by calcium—channel blockers. Proc R Soc Lond Ser B Biol Sci. 1991; 243:195-202.

111. Blatt MR, Thiel G, Trentham DR. Reversible inactivation of K+ channels of Vicia sto-matal guard cells following the photolysis of caged inositol 1,4,5- trisphosphate.Nature. 1990; 346:766-769.

112. Gilroy S, Read ND, Trewavas AJ. levation of cytoplasmic calcium by caged calcium orcaged inositol trisphosphate initiates stomatal closure. Nature. 1990; 346:769-771.

113. Lee YS, Choi YB, Suh S, Lee J, Assmann SM, Joe CO, Kelleher JF, Crain RC. Abscisicacid-induced phosphoinositide turnover in guard-cell protoplasts of Vicia faba. PlantPhysiology. 1992; 110:987-996.

114. Parmar PN, Brearley CA. Metabolism of 3-phosphorylated and 4-phosphorylatedphosphatidylinositols in stomatal guard cells of Commelina communis l. Plant Journal.1995; 8:425-433.

115. Parmar PN, Brearley CA. Identification of 3-phosphorylated and 4-phosphorylatedphosphoinositides and inositol phosphates in stomatal guard cells. Plant Journal. 1993;4:255-263.

116. Muir SR, Sanders D. Pharmacology of Ca2+ release from red beet microsomes suggeststhe presence of ryanodine receptor homologs in higher-plants. Febs Letters. 1996;395:39-42.

117. Allen GJ, Muir SR, Sanders D. Release of Ca2+ from individual plant vacuoles by bothinsp(3) and cyclic ADP-ribose. Science 1995; 268:735-737.

118. Clapham DE. Calcium signaling. Cell. 1995; 80:259-268.119. Webb AAR, McAinsh MR, Mansfield TA, Hetherington AM. Carbon dioxide induces

increases in guard cell cytosolic free calcium. Plant Journal. 1996; 9:297-304.120. McAinsh MR, Webb AAR, Taylor JE, Hetherington AM. Stimulus- induced oscillations

in guard cell cytosolic-free calcium. Plant Cell. 1995; 7:1207-1219.121. Meyer T, Stryer L. Calcium spiking. Ann Rev Biophys Biophys Chem. 1991;20:153-174.122. Berridge MJ. Microdomains and elemental events in calcium signaling. Cell Calcium. 1996;

20:95-96.123. Dolmetsch RE, Lewis RS, Goodnow CC, Healy JI. Differential activation of transcription

factors induced by Ca2+ response amplitude and duration. Nature. 1997; 386:855-858.124. Gradmann D, Blatt MR, Thiel G. Electrocoupling of ion transporters in plants. J Membrane

Biol 1993; 136:327-332.125. Bootman MD, Berridge MJ. Subcellular Ca2+ signals underlying waves and graded

responses in hela cells. Curr Biol. 1996; 6:855-865.126. Chavis P, Fagni L, Lansman JB, Bockaert J. Functional coupling between ryanodine

receptors and L-type calcium channels in neurons. Nature. 1996; 382:719-722.127. Schweitz H, Heurteaux C, Bois P, Moinier D, Romey G, Lazdunski M. Calcicludine, a

venom peptide of the Kunitz-type protease inhibitor family, is a potent blocker of high-threshold Ca2+ channels with a high-affinity for L-type channels in cerebellar granuleneurons. Proc Natl Acad Sci USA. 1994; 91:878-882.

128. Schroeder JI, Hagiwara S. Repetitive increases in cytosolic calcium of guard cells byabscisic acid: Activation of nonselective calcium permeable channels. Proc Natl Acad SciUSA. 1990; 87:9305-9309.

129. Tester M. Plant ion channels: Whole-cell and single-channel studies. New Phytol. 1990;114:305-340.

130. Grabov A, Blatt MR. Parallel control of the inward-rectifier K+ channel by cytosolic-freeCa2+ and pH in Vicia guard cells. Planta. 1997; 201:84-95.

131. Fairley GK, Assmann SM. Evidence for G-protein regulation of inward potassium ionchannel current in guard cells of fava bean. Plant Cell. 1991; 3:1037-1044.

132. Armstrong F, Blatt MR. Evidence for K+ channel control in Vicia guard cells coupled byG-proteins to a 7TMS receptor. Plant J. 1995; 8:187-198.


133. Guern J, Felle H, Mathieu Y, Kurkdjian A. Regulation of intracellular pH in plant-cells.International Review of Cytology. 1991; 127:111-173.

134. Felle H. pH as a second messenger in plants. In: Boss WF, Morre DJ, eds. Secondmessengers in plant growth and development. New York, Alan R. Liss, Inc. 1989:145-166.

135. Morison JIL. Sensitivity of stomata and water use efficiency to high CO2. Plant CellEnviron. 1985; 8:467-474.

136. Senn AP, Goldsmith MHM. Regulation of electrogenic proton pumping by auxin andfusicoccin as related to the growth of Avena coleoptiles. Plant Physiol. 1988; 88:131-138.

137. Brummer B, Bertl A, Potrykus I, Felle H, Parish RW. Evidence that fusicoccin andindole-3-acetic acid induce cytosolic acidifcation of Zea mays cells. FEBS Lett. 1985;189:109-114.

138. Hager A, Moser I. Acetic acid esters and permeable weak acids induce active protonextrusion and extension growth of coleoptile segments by lowering the cytoplasmic pH.Planta. 1985; 163:391-400.

139. Marre E. Fusicoccin: A tool in plant physiology. Ann Rev Plant Physiol Mol Biol. 1979;30:273-288.

140. Gehring CA, Irving HR, Parish RW. Effects of auxin and abscisic acid on cytosoliccalcium and pH in plant cells. Proc Natl Acad Sci USA 1990; 87:9645-9649.

141. Kasamo K. Effect of abscisic acid on membrane-bound epidermal ATPase from tobaccoleaves. Plant Cell Physiol. 1979; 20:293-300.

142. Lurie S, Hendrix D. Differential ion stimulation of plasmalemma adenosine triphosphatasefrom leaf epidermis and mesophyll of Nicotiana rustica L. Plant Physiol. 1979; 63:936-939.

143. Thiel G, Blatt MR, Fricker MD, White IR, Millner PA. Modulation of K+ channels inVicia stomatal guard cells by peptide homologs to the auxin-binding protein C-terminus.Proc Natl Acad Sci USA 1993; 90:11493-11497.

144. Miedema H, Assmann SM. A membrane-delimited effect of internal pH on the K+

outward rectifier of Vicia faba guard cells. J Membr Biol 1996; 154:227-237.145. Lesage F, Guillemare E, Fink M, Duprat F, Lazdunski M, Romey G, Barhanin J. A

pH-sensitive yeast outward rectifier K+ channel with 2 pore domains and novel gatingproperties. J Biol Chem 1996; 271:4183-4187.

146. Coulter KL, Perier F, Radeke CM, Vandenberg CA. Identification and molecular localizationof a pH-sensing domain for the inward rectifier potassium channel HIR. Neuron 1995;15:1157-1168.

147. Bown AW. CO2 and intracellular pH. Plant Cell Environ. 1985; 8:459-465.148. MacRobbie EAC. Effect of ABA on ion transport and stomatal regulation. In: Davies WJ,

Jones HG, eds. Abscisic Acid Physiology and Biochemistry. Oxford: Bios Scientific.1991:153-168.

149. MacRobbie EAC. Ionic relations of guard cells. In: Zeiger E, Farquhar GD, CowanIR, eds. Stomatal Function. Stanford: Stanford University Press. 1987:125-162.

150. Penny MG, Bowing DJF. Direct determination of pH in the stomatal complex ofCommelina. Planta. 1975; 122:209-212.

151. Luan S, Li W, Rusnak F, Assmann SM, Schreiber SL. Immunosuppressants implicateprotein phosphatase regulation of K+ channels in guard cells. Proc Natl Acad Sci USA1993; 90:2202-2206.

152. Thiel G, Blatt MR. Phosphate antagonist okadaic acid inhibits steady-state K+ currentsin guard cells of Vicia faba. Plant J. 1994; 5:727-733.

153. Li WW, Luan S, Schreiber SL, Assmann SM. Evidence for protein phosphatase 1 andphosphatase 2A regulation of K+ channels in 2 types of leaf cells. Plant Physiology. 1994;106:963-970.

154. Mori IC, Muto S. Abscisic acid activates a 48-kilodalton protein kinase in guard cellprotoplasts. Plant Physiology. 1997; 113:833-839.

155. Li JX, Assmann SM. An abscisic acid-activated and calcium-independent protein kinasefrom guard cells of fava bean. Plant Cell. 1996; 8:2359-2368.

156. Koornneef M, Reuling G, Karssen CM. The isolation and characterization of abscisic acid-insensitive mutants of Arabidopsis thaliana. Physiol.Plant. 1984; 61:377-383.


157. Meyer K, Leube MP, Grill E. A protein phosphatase 2C involved in ABA signal transductioninArabidopsis thaliana. Science. 1994; 264:1452-1455.

158. Leung J, Bouvier-Durand M, Morris P-C, Guerrier D, Chefdor F, Giraudat J. ArabidopsisABA response gene ABI1: Features of a calcium-modulated protein phosphatase. Science.1994; 264:1448-1452.

159. Bertauche N, Leung J, Giraudat J. Protein phosphatase activiy of the ABI1 (Abscisic acid-Insensitive 1) protein fromArabidopsis thaliana. European Journal Of Biochemistry. 1996;241:193-200.

160. Schmidt C, Schelle I, Liao YJ, chroeder JI. Strong regulation of slow anion channels andabscisic acid signaling in guard cells by phosphorylation and dephosphorylation events.Proc Natl Acad Sci USA. 1995; 92:9535-9539.

161. Leung J, Merlot S, Giraudat J. The Arabidopsis abscisic acid-insensitive2 (abi2) and abi1genes encode homologous protein phosphatases 2C involved in abscisic acid signaltransduction. Plant Cell. 1997; 9:759-771.

162. Trewavas AJ. Growth substances in context: a decade of sensitivity. Biochem Soc Trans.1992; 20:102-108.

163. Busa WB. Mechanisms and consequences of pH-mediated cell regulation. Annual Reviewof Physiology. 1986; 48:389-402.

164. Taylor CW, Richardson A. Structure and function of inositol trisphosphate receptors.Pharmac Ther. 1991; 51:97-137.

165. Stone JM, Walker JC. Plant protein kinase families and signal transduction. Plant Physiology.1995; 108:451-457.

166. Ma H. Protein phosphorylation in plants—enzymes, substrates and regulators. Trends InGenetics. 1993; 9:228-230.

167. Roberts DM, Harmon AC. Calcium-modulated proteins—targets of intracellular calciumsignals in higher-plants. Annual Review Of Plant Physiology And Plant Molecular Biology.1992; 43:375-414.

168. Urao T, Katagiri T, Mizoguchi T, Yamaguchishinozaki K, Hayashida N, Shinozaki K. 2genes that encode Ca2+-dependent protein-kinases are induced by drought and high-saltstresses in Arabidopsis thaliana. Molecular & General Genetics. 1994; 244:331-340.

169. Sheen J. Ca2+-dependent protein kinases and stress signal transduction in plants. Science.1997; 274:1900-1902.

170. MacRobbie EAC. Signaling in guard cells and regulation of ion channel activity. JournalOf Experimental Botany. 1997; 48:515-528.

171. Hartung W. The site of action of abscisic acid at the guard cell plasmalemma ovValerianella locusta. Plant Cell Environ. 1983; 6:427-428.

172. Baier M, Hartung W. Movement of abscisic acid across the plasmalemma and thetonoplast of guard cells of Valerianella locusta. Bot Acta. 1988; 101:332-337.

173. Paterson NW, Weyers JDB, A'Brook R. The effect of pH on stomatal sensitivity toabscisic acid. Plant Cell And Environment. 1988; 11:83-89.

174. Hornberg C, Weiler EW. High-affinity binding sites for abscisic acid on the plasmalemma ofVicia faba guard cells. Nature. 1984; 310:321-325.

175. Wan YS, Hasenstein KH. Purification and identification of ABA-binding proteins andantibody preparation. Journal Of Molecular Recognition. 1996; 9:722-727.

176. Schwartz A, Wu WH, Tucker EB, Assmann SM. Inhibition of inward K+ channels andstomatal response by abscisic acid-—an intracellular locus of phytohormone action. ProcNatl Acad Sci USA. 1994; 91:4019-4023.

177. Anderson BE, Ward JM, Schroder JI. Evidence for an extracellular reception site forabscisic-acid in commelina guard-cells. Plant Physiology. 1994; 104:1177-1183.

178. Gilroy S. Jones RL. Perception of gibberellin and abscisic-acid at the external face of theplasma-membrane of barley (Hordeum-vulgare l) aleurone protoplasts. Plant Physiology.1994; 104:1185-1192.

179. Assmann SM. Ins and outs of guard cell ABA receptors. Plant Cell. 1994; 6:1187-1190.180. Wickman KD, Clapham DE. G-protein regulation of ion channels. Current Opinion In

Neurobiology. 1995; 5:278-285.


181. Clapham DE. Direct G-protein activation of ion channels. Annual Review Of Neuro-science. 1995; 17:441-464.

182. Berridge MJ. Inositol trisphosphate and calcium signalling. Natur. 1993; 361:315-325.183. Wu WH, Assmann SM. A membrane-delimited pathway of G protein regulation of the

guard cell inward K+ channel. Proc Natl Acad Sci USA. 1994; 91:6310-6314.184. Haldane JBS. Possible Worlds and Other Papers. London, Chatto and Windus. 1927:1-149.185. Grabov A, Blatt MR. Parallel control of the inward-rectifier K+ channel by cytosolic-free

Ca2+ and pH in Vicia guard cells. Planta. 1997; 201:84-95.

125Role of Glycine Betaine and Dimethylsulfoniopropionate in Water-Stress Tolerance

Role of Glycine Betaine andDimethylsulfoniopropionatein Water-Stress ToleranceDouglas A. Gage and Bala Rathinasabapathi

Drought, cold and salinity are three major abiotic conditions limiting the biologicalproductivity of crops. As a metabolic response to the osmotic stress caused by these

environmental factors, many adapted higher plants, marine algae and bacteria accumulateorganic solutes to equalize external osmotic pressure. The most common osmolytes arezwitterionic quaternary ammonium compounds such as glycine betaine, the analogoustertiary sulfonium compound dimethylsulfoniopropionate (DMSP), the amino acidproline, and polyols like mannitol and glycerol. Together these solutes are known as“compatible solutes” or “compatible osmolytes” because they are nontoxic compounds thatdo not inhibit cellular structure and function even at high concentrations.1 In contrast, highconcentrations of perturbing (incompatible) solutes, such as inorganic ions, can causeprotein denaturation. The exclusion of compatible osmolytes from the hydration sphere ofproteins tends to stabilize the tertiary structure of proteins.2 These compounds also reversethe disruption of the tertiary structure caused by perturbing solutes, including inorganicions.2 In addition, compatible solutes may stabilize proteins during freeze-thaw cycles andact as cryoprotectants.3-4 To exert a protective metabolic function, it is important thatcompatible osmolytes be located primarily in the cytosol and chloroplastic compartments,but not the vacuole.5

Among the quaternary ammonium compounds, glycine betaine is the most well-knownand widely distributed.6-8 The tertiary sulfonium compound DMSP has an equally effectiveosmoprotective role in marine algae and certain higher plants.9-13 Aspects of the occurrence,compartmentation, synthesis and roles of glycine betaine and DMSP as osmolytes havebeen reviewed earlier.5,7,8,14-19 The functional equivalence of these and other quaternaryammonium and tertiary sulfonium (“onium”) compounds suggests each might be anappropriate target for metabolic engineering to improve crop stress tolerance.20 However,the introduction of genes coding for biosynthetic enzymes into new plants requires carefulconsideration of the different pathways by which these compounds are formed and theconsequent effects on related primary metabolic processes. Here, we present recentadvances in understanding the synthetic pathways for glycine betaine and DMSP emphasizingbiochemical and molecular genetic results relevant to the development of metabolicengineering strategies. Comparison of the metabolism of glycine betaine and DMSP isillustrative of the potential and challenges for engineering stress resistance.

CHAPTER 7



Stress Protection by Glycine Betaine and DMSP In Vivo and In VitroBoth glycine betaine and DMSP have been shown to afford stress protection for isolated

enzymes, membranes and cultured cells.2,12 For example, glycine betaine has been foundto have unusually strong stabilizing effects on the structure and function of the oxygen-evolvingPhotosystem II complex.21 Using bacterial growth bioassays for osmoprotection againsthigh external salt concentrations,22 glycine betaine and DMSP were found to be equallyeffective (Fig. 7.1).12 Many compatible osmolytes also have cryoprotectant activity,3 butboth comparative physiology23-25 and assays on isolated enzymes indicate that DMSP maybe particularly effective in this regard.4,26 Because DMSP does not contain nitrogen, theaccumulation of DMSP could potentially offer advantages over accumulation of glycinebetaine for osmotic adjustment in N-limited environments.

Occurrence of Glycine Betaine in Angiosperms and Other PlantsIn certain higher plant taxa, glycine betaine accumulates in response to stress, typically to

50-250 µmol g-1 dry weight. It is known to occur in osmotically significant concentrationsin at least thirteen higher plant families: Amaranthaceae, Asteraceae, Avicenniaceae,Caryophyllaceae, Chenopodiaceae, Convulvulaceae, Cuscutaceae, Fabaceae, Malvaceae,Plumbaginaceae, Scrophulariaceae and Solanaceae among the dicots and Poaceae amongthe monocots.6,18 A number of other families produce glycine betaine in trace amounts.6,18

Glycine betaine has also been found in a number of marine algae14 and bacteria.27,28 Massspectrometry and NMR methods have been most useful for identification and accuratequantification of glycine betaine. 8,29-31

Fig. 7.1. Osmoprotection of Salmonella typhimurium LT2 by glycine betaine and DMSP. Cellswere cultured in minimal medium (MOPS-Glucose) with 0.7 M NaCl (o) or 0.7 M NaCl supplementedwith 1 mM of either glycine betaine (�) or DMSP (∆). (Data replotted from reference 12).


Occurrence of DMSPDMSP (also known as dimethyl-β-propiothetin) accumulation to osmotically

significant levels is widespread in marine micro- and macroalgae; it is found in theevolutionary diverse classes Chlorophyceae, Rhodophyceae, Dinophyceae,Chrysophyceae, Bacillariophyceae and Prymnesiophyceae.14,23-25,32 The distribution ofDMSP in marine algae has received considerable attention because this compound is theprimary precursor of biogenic dimethylsulfide (DMS) in the environment. The DMSformed by specific bacterial or algal enzymatic degradation of DMSP33-35 released intothe water column is a significant component of the global sulfur cycle.36 This gas mayalso play a role in climate regulation, because DMS enters the atmosphere from the air-seainterface (approximately 40 million metric tons/year) and is oxidized to sulfate and othercompounds that form tropospheric aerosols and cloud condensation nuclei.36 Aerosolsand clouds in turn affect the earth's radiative balance and potentially influence climate.The decomposition of DMSP may also have a defensive function for some phytoplanktonspecies, because acrylate, the other product in the lyase reaction, deters herbivores.37

Among higher plants, DMSP accumulators are much more restricted, being confirmedin only two unrelated families, the Asteraceae (Wollastonia biflora13,38) and Poaceae (Spartinaspp39-41 and Saccharum spp- and related taxa12,42). Other members of these two familiescontain lower levels of this compound, while some do not produce detectable levels ofDMSP.42 Most of the DMSP-accumulating species are halophytes, although Saccharumofficianale (sugarcane) is a moderately salt-sensitive crop. The marine angiosperms Posidoniasp7 and Zostera sp43 have also been reported to accumulate DMSP, but these studies did notexclude potential contributions from marine algal epiphytes. A number of taxa in otherangiosperm families have been shown to make detectable quantities of DMSP, but thelevels are below those likely to contribute to osmotic stress protection. Cyanobacteria mayalso produce trace amounts of DMSP.44

Surveys of DMSP in different organisms have often employed assays involving thedetection of dimethylsulfide (DMS) released after the base-catalyzed decomposition ofDMSP44 or thin layer chromatography. While these data must be interpreted cautiously,many of the reports above have been subsequently confirmed by other methods, primarilymass spectrometry45 or NMR.14

Genetic Variation for Glycine Betaine SynthesisNatural variation for glycine betaine content is known in higher plants. Glycine

betaine accumulation has evolved in distantly related families, but non-accumulators docontain trace amounts. Therefore, it was suggested that glycine betaine accumulation is anarchetypal angiosperm characteristic, strongly expressed by some but weakly by others.8,20

Accumulators and non-accumulators could be found within a family, genus or species. Forexample, most members of Poaceae accumulate glycine betaine, but cultivated and the wildrice species46,47 Eremochloa ophiuroides,48 Echinochloa utilis49 and some cultivars of corn29

and sorghum50 do not. Among dicots, glycine betaine accumulators and non-accumulatorsare known within the genus Limonium in the Plumbaginaceae20 and Wollastonia in theAsteraceae.13,38 In both instances, glycine betaine was replaced with another compatiblesolute. 13,17,38

Glycine betaine deficiency in corn has been extensively characterized.8,29 Homozygouslines deficient in glycine betaine (bet1/bet1) did not oxidize choline to betaine aldehyde,51

suggesting that Bet1 may encode or regulate the choline oxidizing enzyme. To geneticallytest the role of glycine betaine in osmotic stress tolerance, near-isogenic F8 pairs of glycinebetaine-containing (Bet1/Bet1) and glycine betaine-deficient (bet1/bet1) lines of corn weredeveloped.52 When growth parameters were compared between these two genotypes


under control and salinity stress, both the lines were equal under control conditions, butthe glycine betaine, containing line grew significantly better under salinity stress53 (Table7.1). Glycine betaine-deficient lines were also more markedly impaired by high temperaturestress as evaluated by membrane integrity and in vivo photochemical activity of PSII.54

Genetic and Environmental Variation in DMSPThere is significant inter- and intraspecific variation in DMSP content among marine

algae.14,25,32,55 Even in the classes, such as the Dinophyceae and Prymnesiophyceae, whichcontain a significant number of DMSP accumulating taxa, content can vary widely fromover 2000 µmol/cm3 cell volume (Amphidenium carterae) to below detectable values (someProrocentrum accessions). developmental and physiological conditions may account forsome of this variation.55 Environmental conditions can also influence DMSP content inmarine algae. Phytoplankton do not experience significant changes in salinity, but they dohave to maintain high constitutive levels of osmolytes; for some groups DMSP is the dominantcompatible solute, often in concentrations between 50 and 400 mM. In some organisms(e.g., Tetraselmis subcordiformis) DMSP can occur together with other osmolytes such asglycine betaine and polyols.4 In contrast to pelagic phytoplankton, estuarine micro- andmacroalgae experience frequent major changes in external salinity. DMSP accumulatorsin both constant and variable salinity environments apparently maintain an ability toadjust to increases in external salt concentrations by increasing DMSP content.55 Reducedtemperatures can also promote increases in DMSP in some algal species.24 Nitrogen nutritionis another factor that affects DMSP concentrations in a number of algae. Generally, N-limitationincreases DMSP content.56 Internal DMSP concentrations usually increase slowly over aperiod of hours in response to environmental conditions, including salinity stress.4 However,DMSP concentrations can be lowered rapidly by excretion to the outside environment.4

Some algae produce a DMSP lyase to catalyze the breakdown of DMSP to dimethyl sulfide(DMS) and acrylate, and evidence suggests this enzyme is extracellular.35

Table 7.1 Role of Glycine Betaine In Stress Tolerance In Corn (see ref. 53)

Genotype, Treatment LAER LA PH DW

BET/bet1, Control 331 8867 102 68.3

Bet/Bet1, Control 340 9031 107 67.4

bet1/bet1, Salinized 206 5796 53 34.9

Bet/Bet1, Salinized 228a 6364a 61a 40.7a

LSD (0.05) 17 452 6 4.1

Leaf area expansion rate (LAER) (cm2 d-1) for the stress period, total leaf area at harvest (LA)(cm2), plant height (PH) (cm), and total shoot dry weight (g) at harvest for two F8 families ofcorn grown under nonsalinized (control) or salinized conditions (127.5 mM NaCl+ 22.5 mMCaCl2). aMean (n = 4) of Bet1/Bet1 plants significantly differant at p = 0.05 level from themean of sister line bet1/bet1 plants in the same treatment.


DMSP accumulation in higher plants is also variable. In addition to interspecificvariation,12,42 intraspecific variation in a few taxa has been reported. Different strains ofW. biflora contain different concentrations of DMSP13 and some accessions also accumulateglycine betaine.13,38 Similar variation in DMSP concentrations was found within Saccharumspecies and related taxa,12 as well as in Spartina alterniflora41 Increasing salinity causesDMSP content to increase in W. biflora,13 sugarcane (Saccharum sp) and S. alterniflora.40-41

Other data suggest that DMSP does not function in osmotic adjustment, but rather as aconstitutive osmolyte.57 As in marine algae, DMSP levels were found to be inversely related toN-nutrition in S. alterniflora.40,57

Glycine Betaine Biosynthesis in Animal and Microbial SystemsIn bacteria, animals and higher plants, glycine betaine is synthesized by a two-step

oxidation of choline via betaine aldehyde; however, the enzyme that catalyzes the cholineoxidation step is different in different organisms.

The enteric bacteria Escherichia coli and Salmonella typhimurium, rhizosphere bacteriasuch as Rhizobium meliloti, Azospirillum and Pseudomonas, certain marine algae andcyanobacteria use glycine betaine as a compatible solute. E. coli synthesizes glycine betainefrom exogenous supplies of choline or betaine aldehyde. Choline is oxidized by the actionof a membrane-bound choline dehydrogenase (CDH) to betaine aldehyde, which in turnis oxidized to glycine betaine by a soluble betaine aldehyde dehydrogenase (BADH). Cellsin which CDH alone was expressed can convert radiolabelled choline into glycine betaine,leading to the notion that CDH could catalyze both steps of choline oxidation. However,enzyme activity data suggest that if CDH also catalyzes betaine aldehyde oxidation it shouldbe at least 40-fold less efficient than its activity with choline.27 A regulatory gene betI controlsthe structural genes for choline porter, CDH (betA) and BADH (betB) and choline acts asan inducer.58 E. coli and S. typhimurium use glycine betaine for osmotic adaptation only,but R. meliloti demethylates it to glycine through dimethylglycine and sarcosine. Thedemethylation of glycine betaine is inhibited in a medium of high osmolality, which thuspermits cells to accumulate glycine betaine as a compatible solute.59 As in E. coli and R.meliloti, in animals, choline oxidation is catalyzed by CDH and BADH. CDH in liver andkidney is localized in the inner mitochondrial membrane and the soluble BADH is presentboth in the mitochondria and in the cytosol.60-62 Nucleotide sequences coding for CDHand BADH have been cloned from both animal63-64 and microbial sources.28,65-67 BADHfrom bacteria resembles its higher plant counterpart.65

In certain soil bacteria, choline is oxidized by choline oxidase (COX), a soluble flavoenzyme. In Alcaligenes sp, COX catalyzes the oxidation of both choline and betaine aldehyde,but the enzyme has about seven-fold lower affinity for betaine aldehyde than for choline.68

Genes encoding COX were cloned from Arthrobacter pascens and A. globiformis.69-70 In Arthrobacterglycine betaine does not function as an osmoprotectant,70,71 but is utilized as a carbonsource; accordingly COX expression is repressed by the end-product glycine betaine andregulated by the carbon source in the medium.69

Glycine Betaine Biosynthesis in Higher PlantsSynthesis and accumulation of glycine betaine in higher plants has been studied in

detail in the Chenopodiaceae and the Poaceae. In the leaf cells of halophytic chenopods,glycine betaine is predominantly localized in the cytoplasm and inorganic ions predominantlyin the vacuole.5,8 Radiotracer studies mainly in chenopods and grasses confirmed thatglycine betaine is synthesized by a two-step oxidation of choline via betaine aldehyde.51,71

Glycine betaine is not catabolized in higher plants tested so far.72-74 In chenopods, the firststep of choline oxidation to betaine aldehyde is catalyzed by choline monooxygenase (CMO)


and the oxidation of betaine aldehyde is catalyzed by betaine aldehyde dehydrogenase(BADH)75-76 (Fig. 7.2).

Choline MonooxygenaseCMO was purified and partially characterized from salinized spinach.78 Recently,

cDNAs were isolated from spinach79 and sugar beet.80 Unlike CDH and COX from microbialand animal systems, CMO is a soluble iron sulfur enzyme that requires reduced ferredoxinfor its activity. It is an oligomer of identical subunits of Mr 43,864 with one 2Fe-2S clusterper subunit. The iron-sulfur cluster is coordinated by two cysteine and two histidine ligandsas in Rieske type iron-sulfur proteins.79 The deduced amino acid sequence for CMO alsohas a consensus motif for a mononuclear Fe-binding.80 Upon drought and salinity stress,CMO mRNA, protein and enzyme activity rose three to seven-fold and returned touninduced levels when the stress was removed.80

Betaine Aldehyde DehydrogenaseIn spinach, BADH is a homodimer of nuclear-encoded subunits of Mr 60,000.81-82

Spinach BADH is localized mainly (90%) in the chloroplast stroma.76 Amino acid sequencesdeduced from spinach and sugar beet BADH cDNAs83-84 indicates that BADH lacks a typi-cal stromal targeting peptide. However, expression of these cDNAs in transgenictobacco indicated that spinach and sugar beet BADHs were correctly targeted to the chloroplasts intransgenic tobacco.73 BADH cDNAs or genomic clones were also isolated from spinach,83,85

Atriplex hortensis,86 barley,87 sorghum,88 grain amaranth,89 and rice.90

Several-fold induction of BADH mRNA and enzyme by salinity and drought havebeen studied in sugar beet, barley and sorghum.84,87-88 Recent work on purified BADHfrom amaranth and spinach BADH expressed in transgenic tobacco indicated that BADHis not a substrate-specific enzyme.91,92 BADH efficiently catalyzed dimethyl-sulfoniopropionaldehyde as well as 4-aminobutyraldehyde and 3-aminopropionaldehyde,which are intermediates in putrescine and polyamine degradation.92 There were also someendogenous activities in wild type tobacco with the amino aldehydes tested.92 This suggestedthat plants have a family of aldehyde dehydrogenases with distinct but overlapping substratespecificities. Nakamura et al90 cloned a BADH from rice, a glycine betaine non-accumulatorwhere it is only weakly expressed. Barley, rice and one of the two sorghum BADHs have aC-terminal tripeptide, SKL, known to be a signal for targeting preproteins to microbodies;barley BADH expressed in transgenic tobacco, a dicot, was shown to be localized in theperoxisomes.90 It was proposed that monocot BADHs function in the peroxisomes.90 Thissuggests that glycine betaine synthetic steps are compartmentalized differently between

Fig. 7.2. Glycine betaine biosynthetic pathway, as characterized in spinach. Both of theseenzymatic steps are localized in the chloroplast stroma.77


monocots and dicots. Alternatively, monocot BADHs, all isolated by their homology todicot BADHs, may represent one of many aldehyde dehydrogenases with overlapping substratespecifications and different subcellular location. At present CMO expression is known onlyin Amaranthaceae outside of Chenopodiaceae80 and nothing is known about CMOexpression in monocots. Further work on characterization and subcellular localization ofBADH and CMO (or other choline oxidizing enzymes) in a monocot species are requiredto confirm any differences in glycine betaine synthesis between monocots and dicots.

Choline BiosynthesisCholine, the precursor for glycine betaine, is also the precursor for phosphatidylcholine, a

dominant constituent of membrane phospholipids in eukaryotes. In higher plants, choline issynthesized from serine via ethanolamine. There could be several biosynthetic routes tofree choline from ethanolamine, the routes essentially varying by whether N-methylationof ethanolamine occurs at the free base, phospho-base or phosphatidyl-base levels (see thereview by Rhodes and Hanson8).

The major steps and enzymes involved in choline synthesis from ethanolamine, mostlyas deciphered in spinach leaf tissue, are shown in Figure 7.3. The metabolic basis for theincreased diversion of choline into glycine betaine was investigated in spinach.93 Ethanolaminekinase and the three S-adenosylmethionine:phospho-base N-methyl transferases catalyzingthe methylation of phosphoethanolamine are induced by salinity.93 The regulatory stepfor choline synthesis appears to be at the enzyme catalyzing the first N-methylation ofphosphoethanolamine. This enzyme's activity is highest at the end of the light period, andlight is required for the salt-responsive upregulation of choline synthesis.94 Comparable

Fig. 7.3. Choline biosynthesis in spinach. Reversibility of enzyme-catalyzed steps may not beindicated. Only the major route to phosphatidylcholine is shown. The enzymes are (1) ethanola-mine kinase (2) to (4) S-Adenosyl methionine-dependent N-methyltransferases, (5) cholinekinase, (6) P-choline phosphatase, (7) CTP:P-choline cytidylyl transferase, (8) CDP-cholinediacylglycerol choline phosphotransferase and (9) phospholipase D.


data on regulation of choline synthesis from a glycine betaine non-accumulator or amonocot are not available. Contrasting the chenopods, where free choline is derived fromphosphocholine (Fig. 7.3), in the Poaceae phosphocholine is incorporated to phosphati-dylcholine prior to its release as free choline95 and phosphatidylcholine turnover increasesupon salinity treatment.96

Biosynthesis of DMSPIn 1962 Greene97 conducted an initial series of radiotracer experiments with the

chlorophyte alga Ulva lactuca that established that the carbon skeleton, sulfur atom andmethyl groups of DMSP were derived from methionine. It was later demonstrated thatmethionine was also the precursor of DMSP in other algal groups98-100 and in higherplants.37,42,101 The conversion of methionine to DMSP requires four steps: decarboxylation,deamination, oxidation of the α-carbon and S-methylation. Because intermediates in thepathway were not characterized in the first biosynthetic studies, the order of these stepsremained unknown until recently, although some evidence from the earlier algal studies97,99

suggested that the S-methylation of methionine to form the sulfonium compoundS-methylmethionine (SMM) was not the first step in the pathway. Maw102 originallyproposed that methionine was converted to DMSP via 4-methylthio-2-oxobutyrate(MTOB), then to methylthiopropionate (MTP) and finally by methylation to DMSP. However,there are a number of biochemically plausible alternative routes compatible with the initialradiolabeling data. Recent work in both higher plants and several algal groups has shedlight on the DMSP pathway and led to the surprising finding that at least three, and perhapsfour, biosynthetic routes are found in nature. The results leading to this conclusion arereviewed in detail in the following sections. Progress in the isolation of genes coding forthe DMSP biosynthetic enzymes in both marine algae and higher plants lags behind thatfor glycine betaine, but some of the enzymes have been characterized, and in a few casespurified.103-104

DMSP Biosynthesis in Marine AlgaeBiosynthetic studies have now been carried out in four classes: the Chlorophyceae,

Bacillariophyceae, Prymnesiophyceae (Haptophyceae) and Dinophyceae. Using radioisotopefeeding, Ushida and coworkers105 showed that methionine was incorporated into DMSPin the heterotrophic dinoflagellate Crypthecodinium cohnii. As in the chlorophyte alga Ulvalactuca,97 the C-1 carboxyl group of methionine was lost, confirming that the biosynthesismust involve decarboxylation, deamination, oxidation of the α-carbon and S-methylation. Theorder of these steps in C. cohnii was not completely clear, but several lines of evidenceindicated that the DMSP pathway in this organism did not involve an initial S-methylationstep. First, feeding cold SMM did not inhibit the labeling of DMSP from radiolabeledmethionine, suggesting that SMM was not an intermediate in the pathway.104-105 Alternatively,the methionine transamination product, methylthio-2-oxobutryate (MTOB) did not blockthe incorporation of label from methionine to DMSP. This result could be interpreted tomean that a pathway, involving transamination of methionine to MTOB, is not the firststep in DMSP biosynthesis. In contrast, methylthiopropionate (MTP) did block uptake ofradiolabel into DMSP, supporting its assignment as a biosynthetic intermediate in thisdinoflagellate and inferring that S-methylation of MTP is the last step to DMSP (Fig. 7.4).104

However, as the authors point out, these data may be explained alternatively by the lack ofuptake of externally-supplied SMM or MTOB to the proper intracellular compartment orthe inhibition of methionine uptake by MTP. It should also be noted that MTOB is somewhatunstable, so that decomposition may have prevented supplied MTOB from trapping labelin feeding studies.


If MTP is an intermediate, then the formation of MTP from methionine requires C-1decarboxylation, deamination and oxidation of the C-2 carbon. The isolation of a pyridoxal5'-phosphate-dependent methionine decarboxylase from C. cohnii104 provided some supportfor the notion that decarboxylation was likely the first step in converting methionine toMTP. However, the labeling patterns of the methionine decarboxylation product,methylthiopropylamine (MTPA) or the other putative intermediate, MTP, were not investigated, sothe case for this pathway remains circumstantial. The presence of a methionine decarboxylase,while consistent with the hypothetical pathway, is not conclusive, since similar enzymesare likely widespread in non-DMSP producers (e.g., the fern, Dryopteris felix-mas and Streptomycessp106-107). A different mechanism to form MTP from methionine via the intermediatemethylthiopropylamide by peroxide-dependent oxidative decarboxylation108-109 might alsobe possible.

DMSP biosynthesis was recently investigated in several different classes of algae.100 Inthe chlorophyte alga Enteromorpha intestinalis, a species not too distantly related to Ulvalactuca, feeding studies with 35S-labeled methionine established that this amino acid was

Fig. 7.4. Biosynthetic pathway to DMSP in the dinoflagellate Crypthecodinium cohnii.104,105 A PLP-dependent methionine decarboxylase has been isolated and partially characterized.


Fig. 7.5. Biosynthesis of DMSP in the chlorophyte alga Enteromorpha intestinalis.100 The enzymaticmechanisms catalyzing the first three steps have been elucidated, and the intermediates MTHBand DMSHB have been shown to be the D isomers.118


the precursor of DMSP, as in all other organisms investigated to date. In this study theintermediates in the pathway were identified by following the incorporation of 35S-labelinto other metabolites. Two compounds, methylthio-2-hydroxybutyrate (MTHB) anddimethylsulfonium-2-hydroxybutyrate (DMSHB), were identified as metabolites that rapidlyacquired label and lost it as the 35S-methionine was depleted. Time course experiments andkinetic analysis showed that the MTHB and DMSHB labeling patterns fit those predictedfor biosynthetic intermediates. With the observed labeling kinetics, the most likely routefrom methionine to DMSP via these intermediates would be through the unstable compoundMTOB, which is then reduced to MTHB and subsequently methylated to DMSHB (Fig. 7.5).Because of its instability, the role of MTOB in the pathway was determined by two methods:

(1) The use of a gentle extraction procedure; and(2) The conversion of MTOB to a stable product, MTHB, by reduction with sodium

borohydride and correcting for endogenous MTHB levels. This study conclusivelydemonstrated that MTOB acquired and lost label consistent with its role as an earlyintermediate. Kinetic analysis of the labeling of MTOB, MTHB, DMSHB, confirmedthat flux through the pathway was quantitatively sufficient for these three compoundsto be intermediates in E. intestinalis (Fig. 7.5).

Other compounds monitored in this study, including SMM, dimethylsulfonio-propionaldehyde, acquired little or no 35S. The putative precursor in the C. cohnii DMSPpathway, methylthiopropylamine (MTPA), picked up label slowly, as would be expectedfor a minor end product, not an intermediate. MTP, another proposed precursor, waslabeled to widely varying extents in different E. intestinalis batches. Gage et al100 concludedthat the labeled MTP detected was formed from oxidative decarboxylation of MTOB, acatabolic reaction that is known in other organisms that do not accumulate DMSP.98,110-112

Feeding experiments with the synthetically prepared 35S-labeled metabolites SMM,MTHB and DMSHB supported the proposed route to DMSP (Fig. 7.5). SMM was takenup, but not metabolized to any appreciable extent, indicating that it was not an intermediate inthe pathway. MTHB was primarily converted back to methionine (and ultimately protein),which was not an unexpected fate for this metabolite,113,114 but some MTHB was converted toDMSP. These data do not exclude an indirect route through methionine. However, DMSHB,the key intermediate in the proposed pathway, was efficiently converted to DMSP, but notto other products. Stable isotope labeling and mass spectrometry were used to confirm theidentification of DMSHB as an intermediate. These data also shed light on the enzymaticreactions involved in the pathway in E. intestinalis (see the section below).

The possibility of two different pathways to DMSP in chlorophyte algae and heterotrophicdinoflagellates (see Figs. 7.4, 7.5), raises questions about the biosynthesis in other classesof algae that are known to accumulate this osmolyte. The pathway was also investigated inthe prymnesiophyte Emiliania huxleyi, the diatom Melosira nummuloides and Tetraselmissp, a prasinophyte.100 Each of these species was found to contain small pools of DMSHBwhich accumulated label from 35S-labeled methionine and lost it as the supplied methioninewas depleted. In addition, all three taxa were able to metabolize supplied 35S-labeled DMSHBto DMSP. These data suggest that at least two other algal classes, the Prymnesiophyceaeand Bacillariophyceae (diatoms), have the same pathway as the Chlorophyceae, whiledinoflagellates may use different means to synthesize this compound. As will be shownbelow, DMSP biosynthesis in higher plants proceeds by yet another pathway.

Enzymes in Algal DMSP BiosynthesisAs mentioned above, the first enzyme in the proposed DMSP pathway in the dinoflagellate

C. cohnii was characterized.104,105 A pyridoxal 5'-phosphate-dependent methionine


decarboxylase from this organism was isolated that is a homodimer of two 103 kDasubunits. This contrasts with the methionine decarboxylases previously isolated from Dryopterisfelix-mas and Streptomyces sp, which are homodimers of 57 and 59 kDa subunits, respectively.No kinetic data were reported for the dinoflagellate decarboxylase. The enzymes involved in thesubsequent conversions in the C. cohnii pathway (Fig. 7.4) have not been investigated yet.

In E. intestinalis stable isotope labeling studies provided initial information aboutseveral of the enzymatic steps in the DMSP pathway in this chlorophyte. The conversion ofDMSHB to DMSP was shown to be mediated by an oxidase mechanism analogous to thatof lactate oxidase.115 When [U-13C5]methionine was supplied to E. intestinalis in an atmospherecontaining either 16O2 or 18O2, a labeling pattern was observed that was consistent with anoxygenase-mediated oxidative decarboxylation in the last step in the pathway (DMSHB toDMSP, Fig. 7.5).100

Stable isotope labeling was also used to investigate the first step in the pathway, thedeamination of methionine to MTOB. There are two common mechanisms that couldeffect this conversion: transamination and oxidative deamination. To distinguish the tworeactions, 15N-labeled methionine was supplied to E. intestinalis. Glutamate, alanine andaspartate all rapidly acquired label, but the amide N of glutamine did not, supporting atransamination mechanism in the deamination of methionine, rather than oxidative deamination.In the latter case, free 15NH3 released in the reaction would be predicted to be incorporated inamide N of glutamine by glutamine synthase.116 Control experiments where 15NH4

+ wassupplied showed that the glutamine amide N could be labeled and by inference thatglutamine synthase was operational in E. intestinalis. Additional evidence for the involvementof a transaminase was provided by the data showing that MTHB was converted back tomethionine; transaminase reactions are reversible.117 By inference, the reductase catalyzingthe conversion of MTOB to MTHB must also be reversible.

Summers et al118 recently provided additional evidence about the enzymes catalyzingthe first three steps in the DMSP pathway in E. intestinalis and other chlorophyte algae.Employing in vitro assays with partially purified E. intestinalis extracts, these authors wereable to confirm that a transaminase was responsible for the conversion of methionine toMTOB. Of potential amino group acceptors, 2-oxoglutarate was found to be preferred.Kinetic analyses showed that the observed enzyme activity was approximately 30-fold higherthan that observed in comparable extracts of three non-DMSP accumulating chlorophytealgae (Halimeda discoides, Caulerpa ashmedii and Utodea conglutinata). Similar methioninetransaminase activity was found in other DMSP-accumulating chlorophytes (E. fasiculataand Ulva spp).

The second step, the conversion of MTOB to MTHB (Fig. 7.5) was catalyzed by anNADPH-dependent reductase. The MTHB produced in this reaction was exclusively theD-isomer. The reverse reaction, MTHB to MTOB, was NADP-dependent and proceededat only 0.4% of the forward rate. Comparisons with other algal extracts demonstrated thatthe activity of MTOB reductase was again significantly higher in E. intestinalis and theother DMSP-producing algae than in three taxa studied that do not accumulate DMSP. Thein vitro results were confirmed in vivo by feeding D-[35S]MTHB to intact E. intestinalisfronds.

The stereoselectivity for the production of D-MTHB was consistent with the substratepreference for the next enzyme in the pathway, MTHB S-methyltransferase. E. intestinalisextracts catalyzed the methylation of D-MTHB, but not L-MTHB, with S-adenosylmethionine acting as the methyl donor. In accord with the proposed biosynthetic route(Fig. 7.5) and previous radiolabeling experiments,100 the methyltransferase activity wasselective for MTHB; no activity was detected for MTP, or other thioethers (e.g.,methylthiopropylamine, D- or L-methionine). As with the previous two enzyme activities,


extracts of the other DMSP-accumulating algae were similar to the preparation fromE. intestinalis, while those of the non-accumulators did not contain significant catalyticactivity. These results suggest that the methylation product, D-DMSHB, might be the preferredsubstrate for the oxidase catalyzing the last step in the formation of DMSP. In vivo labelingexperiments with both L- and D-[35S]DMSHB demonstrated that the D isomer was converted9 times more efficiently to DMSP than the L isomer.

Biosynthesis of DMSP in Higher PlantsThe biosynthesis of DMSP in higher plants was first investigated in the Indo-Pacific

strand plant, Wollastonia biflora (synonyms Melanthera biflora and Wedelia biflora), amember of the Asteraceae.38 DMSP accumulation in this species is variable and someaccessions also accumulate glycine betaine.13 The DMSP biosynthetic studies wereperformed with a genotype rich in DMSP, but lacking glycine betaine. Initialradiolabeling studies with W. biflora leaf discs confirmed that methionine was theprecursor of DMSP.38

When [U-14C]methionine was fed to leaf discs and then chased by a supply of coldmethionine (pulse-chase labeling), one metabolite, SMM, acquired and lost label in a mannerquantitatively consistent with it being an intermediate. This is in contrast to Greene's 97

results in U. lactuca, where only small amounts of labeled SMM were detected. Anotherputative intermediate in the DMSP pathway, MTP, accumulated only a small amount oflabel slowly, suggesting that it is a minor end product in methionine metabolism in thisspecies, and not a precursor to DMSP. Feeding [14C]SMM and [14C]MTP showed thatonly SMM was efficiently converted to DMSP. Further, cold SMM, but not MTP, stronglyreduced the incorporation of label into DMSP from [U-14C]methionine.

While these data supported the role of SMM as an intermediate in DMSP biosynthesis inW. biflora, interpretation of the results is complicated by the facile interconversion of SMMand methionine via the SMM cycle.119 In this futile cycle that is likely ubiquitous inangiosperms, methionine is first S-methylated to form SMM by the following reaction:methionine + S-adenosylmethionine → SMM + S-adenosylhomocysteine. SMM can thenreact with homocysteine enzymatically released from S-adenosylhomocysteine byS-adenosylhomocysteine hydrolase119 to give two molecules of methionine. The functionof this cycle is still unclear, but it has been proposed that SMM may be a way to storemethionine in a less metabolically active form.119 Hanson et al38 showed that SMM was infact a direct precursor of DMSP, and not involved in the pathway only by conversion backto methionine, by labeling one of the methyl groups of SMM with 13C and the other with2H3. Both methyl groups of DMSP retained the labels and there was no evidence of methylgroup scrambling. If the SMM was first converted to methionine before going on to DMSP,then one of the two methyl groups in SMM would be lost and a much more complicatedlabeling pattern in DMSP would be expected. Thus, it appears that the widespreadmetabolite SMM119,120 has been diverted for DMSP synthesis in this species.

In W. biflora, radiolabeling pulse-chase and trapping experiments demonstratedthat the next intermediate in the pathway from SMM to DMSP was dimethylsulfonio-propionaldehyde (DMSP-ald).121 Although the conversion of SMM to DMSP-ald requirestwo steps, decarboxylation and deamination, no free intermediates such as dimethyl-sulfoniopropylamine (DMSP-amine) or dimethylsulfonio-2-oxobutanoic acid (DMSOB)were detected in radiolabeling studies. The latter compound has never been synthesizedand is predicted to be extremely unstable,122 so it is unlikely to be a free intermediate in thepathway. DMSP-ald itself is rather unstable, and degradation to DMS is the main byproductwhen the radiolabeled compound is supplied.122 Because no precursors of DMSP-ald otherthan SMM were detected in feeding studies with a number of labeled, biochemically-plausible


intermediates (prepared synthetically), it was proposed that intermediates between SMMand DMSP-ald might be tightly bound to a multifunctional enzyme complex catalyzingboth the oxidative deamination and decarboxylation steps.122 Evidence that in fact thisenzymatic step proceeds by a combined transamination-decarboxylation reaction123 will bediscussed below. The biosynthetic pathway to DMSP in W. biflora is summarized in Fig. 7.6.

Recent studies with another DMSP accumulator, Spartina alterniflora, has shown aninteresting variation in the higher plant pathway. In contrast to the pathway in W. biflora,feeding studies with [35S]methionine have shown that there is a free intermediate,dimethylsulfoniopropylamine (DMSP-amine), between SMM and DMSP-ald in this monocot.101

Both labeling kinetics and direct feeding studies showed that this novel compound was afree intermediate in the pathway. Modeling of the pathway flux with data from radiolabelingexperiments, and DMSP-amine’s limited ability to act as a cold trap, indicated that thereare metabolically active and “storage” pools of DMSP-amine in S. alterniflora. This analy-sis also indicated that exogenously-supplied DMSP-amine first enters the storage pool before

Fig. 7.6. Biosynthesis of DMSP in Wollastonia biflora38,121 and Spartina alterniflora.101 In W. biflorathe 2-step conversion of SMM to DMSP-ald is catalyzed by a transamination/decarboxylasecomplex125with no free intermediate (Step1). A different pathway is found in S. alterniflorainvolving decarboxylation of SMM to DMSP-amine (Step 2). The mechanism by whichDMSP-amine is converted to DMSP-ald (Step 3) is not clear, but it could be catalyzed by anamine oxidase, dehydrogenase or transaminase.


being metabolized. In control experiments, non-DMSP accumulating grasses could notconvert [35S]DMSP-amine to DMSP.101 Together these data indicate that DMSP biosynthesisin the monocot S. alterniflora is distinct, but related to that in the dicot W. biflora. SMM isa biosynthetic intermediate in several other taxa that are minor producers of DMSP,42 butsubsequent steps in the pathway remain unknown in these groups.

The occurrence of at least three, and likely four, distinct biosynthetic pathways indifferent groups of organisms (the dinoflagellate C. cohnii, the chlorophyte E. intestinalisand other algae, the dicot W. biflora and the monocot S. alterniflora) makes DMSP uniqueamong natural products. This diversity presents several alternative approaches for thegenetic engineering of this compound, each with possible advantages. The enzymesinvolved in higher plant DMSP biosynthesis are only beginning to be isolated and characterized, butthe mechanisms involved in the individual steps are becoming clear.

Enzymes in Higher Plant DMSP BiosynthesisIn both W. biflora and S. alterniflora, the first step in the DMSP pathway is the

methylation of methionine by the enzyme S-adenosylmethionine:methionineS-methyltransferase (MMT).103 S-adenosylmethionine is the methyl donor. As part of theubiquitous SMM cycle,119 this enzyme is likely found in all angiosperms. In DMSP-producing higher plants, MMT has been recruited as the first committed step in thepathway. This enzyme has now been purified and characterized from W. biflora. MMT is ahomotetramer of an approximately 115 kDa polypeptide. Antibodies to MMT cross-reactwith a similar sized polypeptide in non-DMSP accumulators, such as cabbage, clover andmaize. This structure is significantly different from that of other methyltransferases, whichare usually monomers or dimers of 20-45 kDa polypeptides.124 Because only partialsequence of MMT has been obtained, and the cloning of the corresponding gene is not yetcomplete, the significance of the size of MMT is not yet clear.

The enzyme involved in the direct conversion of SMM to DMSP-ald in W. biflora hasalso been investigated.125 As discussed earlier, this conversion requires a two-step reactioninvolving removal of the amino group and decarboxylation. Stable isotope labeling experiments with[15N]SMM showed that a transamination, rather than a deamination, step was likely.125

However, simple transamination of SMM produces the extremely unstable compounddimethylsulfonio-2-oxobutyrate (DMSOB), which would be expected to rapidly undergob,g-elimination to yield DMS and vinylglyoxylate.122 Because the kinetics of thedecomposition reaction indicates that free DMSOB is not likely a product, a coupled tran-saminase/decarboxylase enzyme complex was proposed, perhaps involving pyridoxal5'-phosphate (PLP) as a cofactor or cosubstrate. Given the nature of the substrate andintermediates in this reaction, this complex would be biochemically unusual, if notunprecedented.

In S. alterniflora, little is known about the enzymatic steps catalyzing the conversionof SMM to DMSP-ald via the intermediate DMSP-amine.101 However, an SMM decarboxylasemust be present. Subsequent conversion of DMSP-amine to DMSP-ald could proceed viaone of several alternative mechanisms involving an amine oxidase, dehydrogenase ortransaminase. It is possible that the two novel enzymatic steps in the S. alterniflora DMSPpathway might have originated from more widespread enzymes. For example, SMM isstructurally analogous to S-adenosylmethionine, and S-adenosylmethionine decarboxylasesare widespread in plants.126 Whether the similarity of DMSP-amine to other diaminesindicates that a diamine oxidase might be involved is intriguing, but speculative at thispoint.

The last step in higher plant DMSP biosynthesis is common to both the S. alternifloraand W. biflora pathways; in monocots and dicots DMSP-ald is oxidized to DMSP. The


enzyme responsible for catalyzing this reaction has not yet been isolated, but some of itsfeatures have been characterized. A pyridine nucleotide-dependent DMSP-ald dehydrogenase(DDH) activity in W. biflora has been identified.127 This enzyme utilized either NAD orNADP, though NAD was preferred, and the effect of these cofactors was not additive. Themeasured activity and Km was sufficient to account for the in vivo rate of DMSP synthesis.There is an interesting parallel between this enzymatic step in DMSP synthesis and that ofBADH in glycine betaine. Antisera against BADH neutralized DDH activity in W. bifloraextracts and immunoblot analysis showed a single polypeptide band at 63 kDa,127 similarto that of BADH subunits in other plants.76,82 Betaine aldehyde was also found to be aweak competitive inhibitor of DDH.127 These data suggest that DDH and BADH may beclosely related enzymes. BADH isolated from the non-DMSP producer Amaranthushypochondriacus efficiently catalyzes the oxidation of DMSP-ald to DMSP.91 In fact, theVmax/Km value indicates that DMSP-ald is a better substrate for BADH than betaine aldehyde.91

Further, as was discussed above, tobacco plants engineered to express sugar beet BADHwere able to oxidize DMSP-ald and several other aldehyde substrates.92 Thus, BADH and,by inference, DDH, are clearly not as substrate-specific as previously supposed and mayhave originally evolved for another role, perhaps in polyamine metabolism. In any case,this enzymatic step in DMSP biosynthesis may have evolved by the recruitment of preexistingenzymes.

Fig. 7.7. Subcellular localization of the DMSP biosynthetic steps in W. biflora. A schematic repre-sentative of the subcellular localization of the biosynthetic enzymes in the DMSP pathway (afterrefs. 125, 127). MMT is cytosolic, but the enzyme(s) involved in the conversion of SMM to DSMP-ald andDDH are stromal. There is recent evidence for salt stress-activated SMM import into the chloroplast, butlittle is known about the export mechanism for DMSP.128 Although it is not shown, the intermediateidentified in S. alterniflora between SMM and DMSP-ald, DMSP-amine,101 is also likely chloroplastic.


Localization of DMSP BiosynthesisLittle is known of the intracellular localization of the DMSP biosynthesis in marine

algae, but recent studies have provided insight into the compartmentation of DMSP biosyntheticenzymes in higher plants. Studies with W. biflora mesophyll protoplasts employing subcellularfractionation and immunological techniques demonstrated that MMT is a cytosolicenzyme.127 This observation is in accord with earlier investigations of SMM metabolism.119

DDH, in contrast, was localized in the chloroplast stroma (Fig. 7.7).127 The relative instabilityof the substrate DSMP-ald suggests that it should be synthesized in close proximity toDDH and labeling experiments showed this is the case. Feeding [35S]SMM to intact chloroplasts,incubated in the light with HCO3

- and 3-phosphoglyceric acid to promote photosynthesis,efficiently produced labeled DMSP. (The enhancement of DMSP synthesis under photosyntheticconditions implies there may be some link to the Calvin cycle, but at present this relationship isunclear.) The synthesis of DMSP indicates that there is a mechanism to import SMM intothe chloroplast and that the next enzyme in the pathway, the transaminase/decarboxylasecomplex that converts SMM to DMSP-ald, is in the same compartment. A similarcompartmentation of DMSP biosynthetic enzymes may be occurring in S. alterniflora,where DMSP-amine is an intermediate between SMM and DMSP-ald. Although analogoussubcellular fractionation experiments have not been conducted, the limited ability ofexogenously-supplied DMSP-amine to act as a cold trap in feeding studies may reflect thefact that the metabolically active DMSP-amine is chloroplastic and that the “storage pool”is cytosolic.101

Follow-up studies have shown that SMM import into the chloroplast may play a keyregulatory role in DMSP biosynthesis in W. biflora.128 When salinized with 30% seawater,intact protoplasts from W. biflora were found to produce increased levels of DMSP.Quantitative analysis of SMM and DMSP in the intact protoplasts and in chloroplastsderived from the protoplasts demonstrated that there was a significant intracellular redis-tribution of SMM. SMM levels decreased in salinized leaves, with the drop primarily inextrachloroplastic SMM, while levels inside the chloroplast were similar. Thus, 40% of theSMM was chloroplastic in unsalinized protoplasts, while 80% was found in the chloro-plasts following salinization. These results suggest that in W. biflora the SMM transporteris activated under salt stress in order to increase DMSP biosynthesis. As expected, DMSPcontent also increased in the chloroplasts upon salinization.128 Estimates indicate thatstromal concentrations go from approximately 60 mM in control chloroplasts (44% ofthe total) to 130 mM in salinized chloroplasts (69% of the total). While there must be ameans to translocate DMSP from the chloroplasts into the cytosol, this compound’sintracellular distribution in other compartments has not been thoroughly investigated.Leakage of DMSP from chloroplasts during their isolation was observed, however.128

Whether active or passive transport of DMSP out of the chloroplast occurs in vivo isunknown.

Metabolic Engineering of Glycine Betaine SynthesisIn recent years glycine betaine has been an active target for genetic engineering into

nonproducing organisms to provide improved osmotic stress resistance. Genes encodingglycine betaine biosynthetic enzymes from microbes and higher plants have been used toengineer this pathway in heterologous organisms. For example, the bacteriumS. typhimurium lacks natural ability to oxidize choline; expression of E. coli genes forcholine transport, CDH and BADH in S. typhimurium conferred increased osmotolerancein the presence of choline.27 Similarly, introduction of a gene for COX from A. pascens intoan E. coli mutant defective in betaine synthesis resulted in glycine betaine synthesis andosmotolerance upon exogenous choline supply.69 Nomura et al129 transformed the fresh


water cyanobacterium Synechococcus sp. with E. coli bet genes. The transformed cells tookup exogenous choline, accumulated glycine betaine up to a concentration of 45 mM andtolerated salt stress.129 Similarly, the gene for choline oxidase (codA) from A. globiformiswas introduced to Synechococcus. The transformed cells synthesized glycine betaine fromexogenous choline and were tolerant to salt stress70 and low temperature stress.71 In thesemicrobial models, the role of glycine betaine in stress protection has been well demonstratedand in each case the host organism depended on exogenous choline to synthesize glycinebetaine.

Engineering glycine betaine synthesis in higher plants is more complex, especiallywith reference to the availability and regulation of choline. Some of the considerations inthis context were presented by McCue and Hanson.5 Many important crops such as rice,legumes, canola, tomato and cucurbits lack the ability to accumulate glycine betaine. Hence,engineering its synthesis is an attractive route to improve their stress tolerance. The naturalvariation for glycine betaine synthesis in higher plants (see above) suggests that engineeringthis pathway should be a biological feasibility. Both glycine betaine non-accumulators andaccumulators have comparable free choline pools and there is evidence that choline isfeedback regulated.8,52 However, at present it is not clear how much plasticity there is incholine production if significant flux to a new sink is introduced. It is quite possible thatthe large methyl group demand required to accumulate osmotically significant quantitiesof glycine betaine would exceed the normal capacity of choline synthesis to respond. Inorder to engineer glycine betaine (or DMSP) accumulation in higher plants, manipulationof folate-mediated methyl group metabolism130 may be required.

Some initial experiments have been done to address this question. Coding sequencesfor BADH from spinach, sugar beet, barley and E. coli were successfully expressed intransgenic tobacco under constitutive promoters.73,87,131 When supplied with betainealdehyde, the transgenic tobacco expressing spinach or sugar beet BADH accumulatedglycine betaine to levels comparable to glycine betaine accumulators.73 However, endogenouspools of choline were not diverted to glycine betaine in these transformants because thecholine derived precursor, betaine aldehyde, was provided. Engineering of the cholineoxidation (CMO) step into any of these BADH-expressing transgenic tobacco has not yetbeen done.

Lilius et al132 introduced the E. coli betA gene encoding CDH into tobacco. Twotransgenic lines of tobacco were shown to grow better than one wild type control undersalt stress, but glycine betaine synthesis was not confirmed or quantified.132 When betAwas expressed in potato, one transgenic line tested produced glycine betaine up to 108nanomoles g-1 fwt compared to 44 nanomoles g-1 fwt in the wild type control, though theplants were supplied with 15 mM choline in the medium.133 Low levels of glycine betaineaccumulation could be due to poor expression of the transgene or limited access by CDHto supplied choline with resulting poor betaine aldehyde oxidation.

Recently a chimeric construct containing the codA gene (from A. globiformis) for COXunder the control of a constitutive promoter was introduced into Arabidopsis thaliana.134

COX was targeted to the chloroplasts by using the transit peptide of the small subunit ofRubisco.134 Three lines of transformants were shown to accumulate about 1 µmole g-1 fwt(equalling about 50 mM internal concentration) and had improved tolerance for salt andcold stress.134 This is the most unequivocal and direct proof for the role of glycine betainein stress tolerance in higher plants. Similar results have been achieved by transforming thecodA gene into rice.134 Yet, the glycine betaine concentrations in these transformants arestill below those found in some glycine betaine producers. Further study is needed todetermine if choline metabolism has reached an upper limit.


When spinach CMO cDNA was expressed under a constitutive promoter in transgenictobacco (without spinach or beet BADH), the transgenic lines synthesized about two-foldmore glycine betaine than wild type tobacco or vector alone control (Hanson, personalcommunication). Presumably, in this case an endogenous aldehyde dehydrogenase catalyzedthe conversion of the betaine aldehyde product to glycine betaine. The limited increase inglycine betaine may have been the result of compartmentation and restricted access oravailability of the substrate to the dehydrogenase. Tobacco expressing both CMO and BADHfrom heterologous sources should provide valuable insights as to whether the transgenicplants synthesize osmotically significant quantities of glycine betaine. The results fromthese experiments will help to address whether or not choline metabolism will be a limitingfactor for glycine betaine synthesis.

Despite considerable success in engineering glycine betaine synthesis in plants usinggenes for choline oxidizing enzymes from microbial sources,134 genes from plants wouldappear to have several physiological advantages. The plant enzyme CMO requires reducedferredoxin for its activity. This links glycine betaine synthesis with the light reactions ofphotosynthesis and could help match the supply of glycine betaine with the demand forosmotic adjustment and osmoprotection. This demand for osmotic adjustment climbsrapidly after sunrise as the water potential and water content of salt- or drought-stressedleaves start falling.135 Similarly, the light control of the N-methyltransferase that catalyzesphosphoethanolamine methylation in choline synthesis is physiologically linked to thehigher demand for choline flux to glycine betaine under light. Another consideration isthat in higher plants the glycine betaine synthetic pathway has evolved for osmoprotection,while in microbial systems it may play either a nutritional role alone or a combined rolewith osmoprotection. The regulatory controls on CMO, BADH and choline biosyntheticenzymes from higher plants might therefore be superior to the microbial enzymes. Forexample, cis-regulatory elements for these genes should be ideal for engineering higherplants. Further insight into some of the issues concerning regulation might be provided byexperiments with antisense CMO to downregulate glycine betaine synthesis in plants thatnaturally accumulate glycine betaine.

Prospects for Engineering DMSP SynthesisBecause DMSP is a non-nitrogen-containing compatible osmolyte, the genetic

engineering of its biosynthesis into crop plants is attractive. DMSP's effectivecryoprotectant activity would be another potential benefit for crops.4,23-25 However,because DMSP’s initial precursor is methionine, accumulation of osmotically significantamounts of DMSP might triple the requirement for reduced sulfur and also increase thedemand for methyl groups.136 While the nitrogen from methionine can be recycled, thedemands on sulfur amino acid and C-1 metabolism may make metabolic compensation diffi-cult, without additional manipulations of these primary metabolic pathways (see ref. 136 forfurther discussion of DMSP in the context of primary metabolism). At present little is knownof the potential metabolic consequences on sulfur and methyl group metabolism ofengineering the DMSP pathway. Investigating primary metabolic adaptations in naturalDMSP accumulators might provide insight into this problem.

Another potential complication is that DMSP accumulation in crops couldintroduce a new source of DMS emissions into the environment. At physiological pH,DMSP is relatively stable. The significant release of DMS from marine systems is primarilymediated by the enzymatic breakdown of DMSP by specific bacterial and algal lyases.Therefore, DMS release could be negligible in the absence of these lyases. On the otherhand, in Spartina alterniflora and Wollastonia biflora it has been estimated thatapproximately 1% of the DMSP pools turn over by degradation to DMS every day.40,121


However, it is possible that the DMS emissions from these plants do not originate fromendogenous DMSP-lyase activity, but rather from the breakdown of the unstable biosyntheticintermediate DMSP-ald or degradation of SMM by a specific hydrolase.121 With additionalmanipulations, DMS emissions may be avoidable in plants engineered to accumulate DMSP.

The practical opportunities for genetically engineering the DMSP pathway into otherplants are currently limited, compared to those for the glycine betaine pathway, simplybecause the first genes for the biosynthetic enzymes are only now being cloned. However,as these genes become available, the alternative routes to DMSP in different organisms willprovide several options with alternative benefits and disadvantages. While it may be prematureto speculate on particular engineering strategies, it is interesting to consider the alternatives.Targeting of the gene product or intermediate to the proper compartment (e.g., DDH orSMM to the chloroplast stroma) is another important issue. Engineering enzymes fortranslocation to a specific compartment may be much easier than targeting metabolites.Specific transporters may be required,128 about which almost nothing is currently known.

There are four potential DMSP biosynthetic pathways to consider (see Figs. 7.4-7.6).The number of steps in each of these pathways may seem daunting from the perspective ofengineering, especially given the simplicity of the glycine betaine pathway. However, aswas illustrated above, some of the enzymes involved are common to most plants, so thatintroduction of new genes may not be required. For example, the dinoflagellate pathway104

is initiated by decarboxylation of methionine by a PLP-dependent decarboxylase. Althoughthe C. cohnii enzyme appears to be unusual,104-105 functionally equivalent enzymes areknown from other organisms106-107 and may be widespread. The mechanism(s) involvedin the next step in the pathway (MTPA→MTP) is not clear at this time and might requirean amine oxidase, dehydrogenase or transaminase. The last reaction would also requirethe engineering of a specific methyltransferase.

The E. intestinalis pathway100 (Fig. 7.5) again appears to have evolved by utilizingsome of the enzymes involved in primary amino acid metabolism.118 Methionine transaminases(methionine→MTOB) are likely ubiquitous, although the enzyme in the DMSP producerappears to be 30- to 100-fold more active, with a much lower Km (30 µM vs mM range formost amino transferases) and is therefore likely a novel enzyme rather than over-expression ofa standard amino transferase.118 To successfully engineer DMSP accumulation, the genefor the specialized methionine amino transfersase would likely have to be introduced. Theposition of a transaminase at the head of the DMSP pathway in E. intestinalis may explainthe increased production of DMSP under N-deficit conditions. Reduction of cellular aminoacid content would increase transamination reactions and thus activate the DMSP pathway.For some environments this regulatory point in an engineered crop might be valuable. Incontrast, under low N conditions, glycine betaine production is diminished.

Like the first enzyme in the pathway, many plants have a low level of MTOB reductaseactivity (MTOB→MTHB).137 The stereochemical configuration of the MTHB producedhas not been determined for plants other than E. intestinalis, however. Because the MTOBreductase is probably also specialized, introduction of the reductase gene would also berequired to engineer the pathway. Finally, a unique methyltransferase converting D-MTHBto D-DMSHB is only found in DMSP accumulators and would by necessity be requiredfor the pathway. However, it is possible that engineering just the first three steps would besufficient to impart osmotic stress resistance. The last step in the pathway, conversion ofDMSHB to DMSP, might not be necessary. DMSHB is an effective compatible osmolyte, 118

although it is subject to enzymatic degradation to DMS in vivo in the algae that use thiscompound in the DMSP pathway.100 However, it is a chemically stable compound in theabsence of the degradative enzymes, so in principle it would be possible to accumulateDMSHB without DMS emissions.


The W. biflora and S. alterniflora pathways differ only in the nature of the conversionof SMM to DMSP-ald (Fig. 7.6). As with the other DMSP pathways, higher plant DMSPbiosynthesis has recruited steps from primary metabolism, in this case the SMM cycle (toslightly stretch the definition of “primary metabolism”). In W. biflora the first step in thepathway occurs in the cytosol (methionine→SMM). MMT is abundant in many non-DMSPaccumulators, so that this first step may not have to be engineered. However, as the nextsteps in the pathway occur in the chloroplast, cytosolic SMM must be transported into thechloroplast stroma.127 The transporter(s) that perform this function are unknown. SMMimport to the chloroplast is quite active in the DMSP accumulator W. biflora, particularlyunder salinized conditions, while non accumulators have only residual SMM trans-port function under any conditions.128 Thus, even if the next step in the pathway, thetransamination/decarboxylation of SMM to DMSP-ald,125 could be engineered to beexpressed in the chloroplast of non-DMSP accumultors, it is probable that insufficientSMM substrate would be available. Further investigation of the SMM transporter inDMSP and non-DMSP accumulating plants is definitely warranted. If this limitationis overcome, the last step in the pathway may not require the introduction of DDH.Both BADH and DDH appear to have less substrate specificity than expected. Theseenzymes may both be related to the constitutive aldehyde dehydrogenases involved inpolyamine metabolism.

The relative advantages and disadvantages of engineering a particular DMSP pathway (orgenes from different pathways) will become apparent as we learn more about the biosyntheticenzymes involved. These studies are only in the early stages.

ConclusionThe engineering of compatible osmolyte biosynthetic pathways into crop plants to

impart improved stress tolerance has long been an objective. There has been recent successin the introduction of bacterial glycine betaine genes to higher plants to impart improvedstress resistance. Now that the plant genes are available, it will be interesting to compareglycine betaine sythesis and the stress response of other plants transformed with theseplant genes. The engineering of DMSP accumuation will remain a challenging target forthe future. A great deal of fundamental biochemical investigation must precede anyapplications in genetic engineering. The remarkable biosynthetic diverstiy for the productionof this compound is unprecedented and will offer many opportunities for genetic engineering.That DMSP production has evolved so many times, suggests that it may be a very usefulmolecule. Still, attempts to engineer DMSP accumulation will face the same hurdles thatconfront glycine betaine engineering. How can substrate limitations be overome withoutdisrupting primary metabolism? It is clear that the engineering of compatible osmolytescannot be viewed in isolation from primary metabolism, particularly that involved inmethyl group metablolism, and in the case of DMSP, sulfur metabolism also. Theintroduction of the genes for the biosythesis of compounds like glycine betaine and intro-duction of the genese for the biosythesis of compounds like glycine betaines and DMSPmay not immediately produce stress-tolerant crops, but they will be useful tools forunderstanding how some key areas in primary metabolism interact and are regulated.

AcknowledgmentsThis publication is Florida Agricultural Experiment Station Journal Series Number

R-06603 and support to B.R. from the College of Agriculture, University of Florida isgratefully acknowledged.


References1. Timasheff SN. A physiochemical basis for the selection of osmolytes by nature. In: Somero

GN, Osmond CB, Bolis CL, eds. Water and Life. Berlin:Springer Verlag, 1992:70-84.2. Yancey PH. Compatible and counteracting solutes. In: Strange K, ed. Cellular and Molecular

Physiology of Cell Volume Regulation. Boca Raton: CRC Press, 1994:81-109.3. Carpenter JF, Crowe JH. The mechanism of cryoprotection of proteins by solutes. Cryobiol

1988; 25:244-255.4. Kirst GO. Osmotic adjustment in phytoplankton and macroalgae. The use of dimethyl-

sulfoniopropionate. In:Kiene RP, Visscher PT, Keller MD, Kirst GO, eds. Biological andEnvironmental Chemistry of DMSP and Related Sulfonium Compounds. New York: PlenumPress, 1996:121-129.

5. McCue KF, Hanson AD. Drought and salt tolerance: Towards understanding and applica-tion. Trends Biotechnol 1990; 8:358-362.

6. Weretilnyk EA, Bednarek S, McCue KF, Rhodes D, Hanson AD. Comparative biochemicaland immunological studies of the glycine betaine synthesis pathway in diverse families ofdicotyledons. Planta 1989; 178:342-352.

7. Wyn Jones RG, Storey R. Betaines. In: Paleg LG, Aspinall D, eds. The Physiology andBiochemistry of Drought Resistance in Plants. Sydney: Academic Press, 1981:171-204.

8. Rhodes D, Hanson AD. Quaternary ammonium and tertiary sulfonium compounds inhigher-plants. Ann Rev of Plant Physiol Mol Biol 1993; 44:357-384.

9. Dickson DJM, Kirst GO. The role of β-dimethylsulfoniopropionate, glycine betaine andhomarine in the osmoacclimation of Platymonas subcordiformis. Planta 1986; 167:536-543.

10. Dickson DJM, Kirst GO. Osmotic adjustment in marine eukaryotic algae: The role ofinorganic ions, quaternary ammonium, tertiary sulfonium and carbohydrate solutes. NewPhytol 1987; 106:657-666.

11. Reed RH. Measurement and osmotic significance of (-dimethylsulfoniopropionate in marinemacroalgae. Mar Biol Lett 1983; 4:173-181.

12. Paquet L, Rathinasabapathi B, Saini H, Zamir, L, Gage DA, Huang Z-H, Hanson, AD.Accumulation of the compatible solute 3-dimethylsulfoniopropionate in sugarcane andits relatives, but not other gramineous crops. Aust J Plant Physiol 1994; 21:37-48.

13. Storey R, Gorham J, Pitman MG, Hanson AD, Gage DA. Responses of Melanthera biflorato salinity and water stress. J Exp bot 1993; 44:1551-1560.

14. Blunden G, Gordon SM. Betaines and their sulphonio analogues in marine algae. ProgPhycol Res 1986; 4:39-80.

15. Csonka LN, Hanson AD. Prokaryotic osmoregulation: Genetics and physiology. Ann RevMicrobiol 1991; 45:569-606.

16. Hanson AD. Compatible solute synthesis and compartmentation in higher plants. In: SomeroGN, Osmond CB, Bolis CL, eds. Water and Life. Berlin: Springer-Verlag,1992:52-60.

17. Hanson AD, Rivoal J, Burnet M, Rathinasabapathi B. Biosynthesis of quaternary ammoniumand tertiary sulphonium compounds in response to water deficit. In: Smirnoff N, ed.Environment & Plant Metabolism. Oxford: Bios Scientific Publishers,1994:89-198.

18. Gorham J. Betaines in higher plants—biosynthesis and role in stress metabolism. In:Wallsgrove RM, ed. Amino acids and their derivatives in higher plants. Cambridge University Press,1995:173-203.

19. Miller KJ, Wood JM. Osmoadaptation by rhizosphere bacteria. Ann Rev Microbiol1996; 50:101-136.

20. Hanson AD, Rathinasabapathi B, Rivoal J, Burnet M, Dillon MO, Gage DA. Osmo-protectantcompounds in the Plumbaginaceae: A natural experiment in metabolic engineering. ProcNat Acad Sci USA 1994; 91:306-310.

21. Papageorgiou GC, Murata N. The unusually strong stabilizing effects of glycine betaineon the structure and function of the oxygen-evolving Photosystem II complex. Photo-synthesis Res 1995; 44:243-252.

22. Hanson AD, Rathinasabapathi B, Chamberlin B, Gage DA. Comparative physiologicalevidence that β-alanine betaine and choline-O-sulfate act as compatible osmolytes in halophyticLimonium species. Plant Physiol 1991; 97:1199-1205.


23. Kirst GO, Thiel C, Wolff H, Nothnagel J, Wanzec M, Ulmke R. Dimethylsulfonio-propionate(DMSP) in ice-algae and its possible biological role. Mar Chem 1991; 35:381-388.

24. Karsten U, Wienke C, Kirst GO. The β-dimethylsulfoniopropionate (DMSP) content ofmacroalgae from Antarctica and southern Chile. Bot Mar 1990; 33:143-146.

25. Karsten U, Wienke C, Kirst GO. Dimethylsulfoniopropionate (DMSP) accumulation ingreen macroalgae from polar to temperate regions: interactive effects of light versus salinityand light versus temperature. Polar Biol 1992; 12:603-607.

26. Nishiguchi MK, Somero GN. Temperature and concentration dependence of compatibilityof the organic osmolyte dimethylsulfoniopropionate. Cryobiol 1992; 29:118-124.

27. Andersen PA, Kaasen I, Styrvold OB, Boulnois G, Strom AR. Molecular cloning, physicalmapping and expression of the bet genes governing the osmoregulatory choline-glycinebetaine pathway of Escherichia coli. J Gen Microbiol 1988; 134:737-1746.

28. Boch J, Kempf B, Bremer E. Synthesis of the osmoprotectant glycine betaine in Bacillussubtilis: Characterization of the gbsAB genes. J Bacteriol 1996; 178:5121-5129.

29. Rhodes D, Rich PJ, Myers AC, Reuter CR, Jamieson GC. Determination of betaines byfast atom bombardment mass spectrometry: Identification of glycine betaine deficientgenotypes of Zea mays. Plant Physiol 1987; 84:781-788.

30. Selvaraj G, Jain RK, Olson DJ, Hirji R, Jana S, Hogge LR. Glycine betaine in oilseed rapeand flax leaves: Detection by liquid chromatography/continuous flow secondary ion-massspectrometry. Phytochemistry 1995; 38:1143-1146.

31. Gorham J. Glycine betaine is a major nitrogen-containing solute in the Malvacceae.Phytochemistry 1996; 43:367-369.

32. Keller MD, Bellows WK, Guillard RRL. Dimethylsulfide production in marine phy-toplankton. In: Salzman ES, Cooper WJ, eds. Biogenic Sulfur in the Environment.Washington, DC: American Chemical Society,1989:167-182.

33. Kiene RP. DMS production from DMSP in coastal seawater samples and bacterial cultures.Appl Environ Microbiol 1990; 56:3292-3297.

34. de Souza MP, Yoch DC. N-terminal amino acid sequences and comparisons of DMSPlyases from Pseudomonas doudorffii and Alcaligens strain M3A. In: Kiene RP, VisscherPT, Keller MD, Kirst GO, eds. Biological and Environmental Chemistry of DMSP andRelated Sulfonium Compounds. New York: Plenum Press,1996:293-304.

35. Stefels J, Gieskes WWC, Dijkhuizen L. Intriguing functionality of the production andconversion of DMSP in Phaeocystis sp. In: Kiene RP, Visscher PT, Keller MD, Kirst, GO,eds. Biological and Environmental Chemistry of DMSP and Related Sulfonium Compounds.New York: Plenum Press,1996:305-315.

36. Malin G. The role of DMSP and DMS in the global sulfur cycle and in climate regulation. In:Kiene RP, Visscher PT, Keller MD, Kirst GO, eds. Biological and Environmental Chemistry ofDMSP and Related Sulfonium Compounds. New York: Plenum Press, 1996:77-189.

37. Wolfe GV, Steinke M, Kirst GO. Grazing-activated chemical defence in a unicellular marinealga. Nature 1997; 387:894-896.

38. Hanson AD, Rivoal J, Paquet L, Gage DA. Biosynthesis of 3-dimethylsulfoniopropionatein Wollastonia biflora (L.)DC. Evidence that S-methylmethionine is an intermediate. PlantPhysiol 1994; 105:103-110.

39. Larher F, Hamelin J, Stewart GR. L’acide dimethylsulfonium-3-propanoique de Spartinaangelica. Phytochemistry 1977; 16:2019-2020.

40. Dacey JWH, King GM, Wakeman SG. Factors controlling the emission of dimethylsulfidefrom salt marshes. Nature 1987; 330:643-645.

41. Otte ML, Morris JT. Dimethylsulfoniopropionate (DMSP) in Spartina alterniflora Loisel.Aquatic Bot 1994; 48:239-259.

42. Paquet L, Lafontaine PJ, Saini HS, James F, Hanson AD. Evidence en faveur de la presencedu 3-dimethylsulfoniopropionate chez une large gamme d’Angiospermes. Can J Bot 1995;73:1889-1896.

43. White H. Analysis of dimethylsulfonium compounds in marine algae. J Marine Res 1982;40:529-536.


44. Vogt C, Rabenstein A, Rethmeier J, Fischer U. Alkali-labile precursors of dimethyl sulfide inmarine benthic cyanobacteria. Arch Microbiol 1998; 169:263-266.

45. Gage DA, Hanson AD. Characterization of 3-dimethylsulfoniopropionate (DMSP) andits analogs with mass spectrometry. In: Kiene R, Visscher PT, Keller MD, Kirst GO, ed.Biological and Environmental Chemistry of DMSP and Related Sulfonium Compounds.New York: Plenum, 1996:29-44.

46. Hitz WD, Hanson AD. Determination of glycine betaine by pyrolysis-gas chromatography incereals and grasses. Phytochemistry 1980; 19:2371-2374.

47. Rathinasabapathi B, Gage DA, Mackill DJ, Hanson AD. Cultivated and wild rices do notaccumulate glycine betaine due to deficiencies in two biosynthetic steps. Crop Sci 1993;33:534-538.

48. Marcum KB, Murdoch CL. Salinity tolerance mechanisms of six C4 turfgrasses. J AmSoc Hort Sci 1994; 119:779-784.

49. Ishitani M, Arakawa K, Mizuno K, Kishitani S, Takabe T. Betaine aldehyde dehydrogenase inthe Gramineae: Levels in leaves of both betaine-accumulating and nonaccumulating cerealplants. Plant Cell Physiol 1993; 34:493-495.

50. Grote EM, Ejeta G, Rhodes D. Inheritance of glycine betaine deficiency in sorghum. CropSci 1994; 34:1217-1220.

51. Lerma C, Rich PJ, Ju GC, Yang W-J, Hanson AD, Rhodes D. Betaine deficiency in maize:Complementation tests and metabolic basis. Plant Physiol 1991; 95:1113-1119.

52. Yang W, Nadolska-Orczyk A, Wood KV, Hahn DT, Rich PJ, Wood AJ, Saneoka H,Premachandra GS, Bonham CC, Rhodes JC, Joly RJ, Samaras Y, Goldsbrough PB,Rhodes D. Near-isogenic lines of maize differing for glycine betaine. Plant Physiol1995; 107:621-630.

53. Saneoka H, Nagasaka C, Hahn DT, Yang W, Premachandra GS, Joly RJ, Rhodes D. Salttolerance of glycine betaine-deficient and -containing maize lines. Plant Physiol 1995; 107:631-638.

54. Yang G, Rhodes D, Joly RJ. Effects of high temperature on membrane stability and chlorophyllfluorescence in glycine betaine-deficient and glycine betaine-containing maize lines. AustJ Plant Physiol 1996; 23:437-443.

55. Keller MD, Korjeff-Belows W. Physiological aspects of the production of dimethyl-sulfoniopropionate (DMSP) by marine phytoplankton. In: Kiene RP, Visscher PT, KellerMD, Kirst GO, eds. Biological and Environmental Chemistry of DMSP and Related SulfoniumCompounds. New York: Plenum Press,1996:131-142.

56. Gröne T, Kirst GO. The effect of nitrogen deficiency, methionine, and inhibitors of methioninemetabolism on the DMSP content of Tetraselmis subcordiformis. Mar Biol 1992; 112:97-503.

57. Colmer TD, Fan TW-M, Lauchi A, Higashi RM. Interactive effects of salinity, nitrogen andsulfur on the organic solutes in Spartina alterniflora leaf blades. J Exp Bot 1996; 47:369-375.

58. Lamark T, Rokenes TP, McDougall J, Strom AR. The complex bet promoters of Escherichiacoli: Regulation by oxygen (ArcA), choline (BetI), and osmotic stress. J Bacteriol 1996;178:1655-1662.

59. Bernard T, Pocard JA, Perroud B, Le Rudulier D. Variations in the response of salt-stressed Rhizobium strains to betaines. Arch Microbiol 1986; 143:359-364.

60. Tsuge H, Nakano Y, Onishi H, Futamura Y, Ohashi K. A novel purification and some propertiesof rat liver mitochondrial choline dehydrogenase. Biochim Biophys Acta 1980; 614:274-284.

61. Haubrich DR, Gerber GH. Choline dehydrogenase: Assay, properties and inhibitors.Biochem Pharmacol 1981; 30:2993-3000.

62. Zhang J, Blusztajn JK, Zeisel SH. Measurement of the formation of betaine aldehyde andbetaine in rat liver mitochondria by a high pressure liquid chromatography-radioenzymaticassay. Biochim Biophys Acta 1992; 1117:333-339.

63. Loeffler M. NCBI sequence Accession number 1154950. 1994.64. Wilson R. NCBI Accession number Z66494. 1995.65. Boyd LA, Adam L, Pelcher LE, McHughen A, Hirji R, Selvaraj, G. Characterization of an

Escherichia coli gene encoding betaine aldehyde dehydrogenase (BADH): Structural similarityto mammalian ALDHs and plant BADH. Gene 1991; 103:45-52.


66. Lamark T, Kaasen I, Eshoo MW, Falkenberg P, McDougall J, Strom AR. DNA sequenceand analysis of the bet genes encoding the osmoregulatory choline-glycine betaine pathwayof Escherichia coli. Mol Microbiol 1991; 5:1049-1064.

67. Pocard JA, Vincent N, Boncompagni E, Smith LT, Poggi MC, Le Rudulier D. Molecularcharacterization of the bet genes encoding glycine betaine synthesis in Sinorhizobiummeliloti 102F34. Microbiol 1997; 43:1369-1379.

68. Ohta-Fukuyama M, Miyake Y, Emi S, Yamano T. Identification and properties of theprosthetic group of choline oxidase from Alcaligenes sp. J Biochem 1980; 88:197-203.

69. Rozwadowski KL, Khachatourians GG, Selvaraj G. Choline oxidase, a catabolic enzymein Arthrobacter pascens, facilitates adaptation to osmotic stress in Escherichia coli. JBacteriol 1991; 173:472-478.

70. Deshinium P, Los DA, Hayashi H, Mustardy L, Murata N. Transformation ofSynechococcus with a gene for choline oxidase enhances tolerance to salt stress. Plant MolBiol 1995; 29:897-907.

71. Deshinium P, Gombos Z. Nishiyama, Y, Murata N. The action in vivo of glycine betainein enhancement of tolerance of Synechococcus sp. Strain PCC7942 to low temperature. JBacteriol 1997; 179:339-334.

72. Ladyman JAR, Hitz WD, Hanson AD. Translocation and metabolism of glycine betaine.Planta 1980; 150:191-196.

73. Rathinasabapathi B, McCue KF, Gage DA, Hanson AD. Metabolic engineering of glycinebetaine synthesis: Plant betaine aldehyde dehydrogenases lacking typical transit peptides aretargeted to tobacco chloroplasts where they confer betaine aldehyde resistance. Planta1994; 193:155-162.

74. Makela P, Sainio P, Jokinen K, Pehu E, Setala H, Hinkkanen R, Somersalo S. Uptake andtranslocation of foliar-applied glycine betaine in crop plants. Plant Sci. 1996; 121:221-230.

75. Brouquisse R, Weigel P, Rhodes D, Yocum CF, Hanson AD. Evidence for a ferredoxin-dependentcholine monooxygenase from spinach chloroplast stroma. Plant Physiol 1989; 90:322-329.

76. Weigel P, Weretilnyk EA, Hanson AD. Betaine aldehyde oxidation by spinach chloroplasts.Plant Physiol 1986; 82:753-759.

77. Hanson AD, May AM, Grumet R, Bode J, Jamieson GC, Rhodes D. Betaine synthesis inchenopods: Localization in chloroplasts. Proc Nat Acad Sci USA 1985; 82:3678-3682.

78. Burnet M, Lafontaine PJ, Hanson AD. Assay, purification, and partial characterization ofcholine monooxygenase from spinach chloroplast stroma. Plant Physiol 1995; 90:581-588.

79. Rathinasabapathi B, Burnet M, Russell BL, Gage DA, Liao P-C, Nye GJ, Scott P, GolbeckJH, Hanson AD. Choline monooxygenase, an unusual iron-sulfur enzyme catalyzing thefirst step of glycine betaine synthesis in plants: Prosthetic group characterization andcDNA cloning. Proc Natl Acad Sci USA 1997; 94:3454-3458.

80. Russell BL, Rathinasabapathi B, Hanson AD. Osmotic stress induces expression of cholinemonooxygenase in sugar beet and amaranth. Plant Physiol 1998; 116:859-865.

81. Arakawa K, Takabe T, Sugiyama T, Akazawa, T. Purification of betaine-aldehyde dehydrogenasefrom spinach leaves and preparation of its antibody. J Biochem 1987; 101:1485-1488.

82. Weretilnyk EA, Hanson, AD. Betaine aldehyde dehydrogenase from spinach leaves: Purification, invitro translation of the mRNA, and regulation by salinity. Arch Biochem Biophys 1989;271:56-63.

83. Weretilnyk EA, Hanson AD. Molecular cloning of a plant betaine-aldehyde dehydrogenase,an enzyme implicated in adaptation to salinity and drought. Proc Natl Acad Sci USA1990; 87:2745-2749.

84. McCue KF, Hanson AD. Effects of soil salinity on the expression of betaine aldehydedehydrogenase in leaves: Investigation on hydraulic, ionic and biochemical signals. AustJ Plant Physiol 1992; 19:555-564.

85. Shu W, Ai W, Chen S. NCBI Accession number 1813538. 1995.86. Xiao G, Zhang G, Liu F, Chen S. Study on BADH gene from Atriplex hortensis. Chin Sci

Bull 1995; 40:741-745.


87. Ishitani M, Nakamura T, Han SY, Takabe T. Expression of the betaine aldehyde dehy-drogenase gene in barley in response to osmotic stress and abscisic acid. Plant Mol Biol1995; 27:307-315.

88. Wood AJ, Saneoka H, Rhodes D, Joly RJ, Goldsbrough PB. Betaine aldehyde dehydrogenasein sorghum: Molecular cloning and expression of two related genes. Plant Physiol 1996;110:1301-1308.

89. Legaria, J, Iturriaga, G. NCBI Accession number 2388710. 1997.90. Nakamura T, Yokota S, Muramoto Y, Tsutsui K, Oguri Y, Fukui K, Takabe T. Expression of

a betaine aldehyde dehydrogenase gene in rice, a glycine betaine nonaccumulator, andpossible localization of its protein in peroxisomes. Plant J 1997; 11:1115-1120.

91. Vojtechova M, Hanson AD, Munoz-Clares RA. Betaine aldehyde dehydrogenase from amaranthleaves efficiently catalyzes the NAD-dependent oxidation of dimethyl-sulfoniopropionaldehydeto dimethylsulfoniopionate. Arch Biochem Biophys 1997; 337:81-88.

92. Trossat C, Rathinasabapathi B, Hanson AD. Transgenically expressed betaine aldehydedehydrogenase efficiently catalyzes oxidation of dimethylsulfoniopropionaldehyde andomega-aminoaldehydes. Plant Physiol 1997; 113:1457-1461.

93. Summers PS, Weretilyn EA. Choline synthesis in spinach in relation to salt stress. PlantPhysiol 1993; 103:1269-1276.

94. Weretilnyk EA, Smith DD, Wilch GA, Summers PS. Enzymes of choline synthesis in spinach.Response of phospho-base N-methyltransferase activities to light and salinity. Plant Physiol1995; 109:1085-1091.

95. Hitz WD, Rhodes D, Hanson AD. Radiotracer evidence implicating phosphoryl and phosphatidylbases as intermediates in betaine synthesis by water-stressed barley leaves. Plant Physiol1981; 68:814-822.

96. Giddings TH Jr., Hanson AD. Water stress provokes a generalized increase in phosphatidylcholineturnover in barley leaves. Planta 1982; 155:493-501.

97. Greene RC. Biosynthesis of dimethyl-β-propiothetin. J Biol Chem 1962; 237:2251-2254.98. Pokorny M, Marcenko E, Keglevic D. Comparative studies of L- and D-methionine

metabolism in lower and higher plants. Phytochemistry 1970; 9:2175-2188.99. Chillemi R, Patti A, Morrone R, Piatelli M, Sciutto S. The role of methylsulfonium

compounds in the biosynthesis of N-methylated metabolites in Chondria coerulescens. JNat Prod 1990; 53:87-93.

100. Gage DA, Rhodes D, Nolte KD, Hicks WA, Leustek T, Cooper AJL, Hanson, AD. A newroute for synthesis of dimethylsulphoniopropionate in marine algae. Nature 1997; 387:891-894.

101. Kocsis MG, Nolte KD, Rhodes D, Shen T-L, Gage DA, Hanson AD. Dimethyl-sulfoniopropionate biosynthesis in Spartina alterniflora. Plant Physiol 1998; 117:273-281.

102. Maw GA. The biochemistry of sulfonium salts. In: Stirling CJM, Patai S, eds. The Chemistryof the Sulfonium Group. Part 2. Chichester, UK: Wiley, 1981:703-770.

103. James F, Nolte KD, Hanson AD. Purification and properties of S-adenosyl-L-methionine:L-methionine S-methyltransferase from Wollastonia biflora leaves. J BiolChem 1995; 270:22344-22350.

104. Uchida A, Ooguri, T, Ishida, T, Kitaguchi, H, Ishida, Y. Biosynthesis of dimethyl-sulfoniopropionate in Crypthecodinium cohnii (Dinophyceae). In: Kiene RP, Visscher PT,Keller MD, Kirst GO, eds. Biological and Environmental Chemistry of DMSP andRelated Sulfonium Compounds. New York: Plenum Press, 1996:97-107.

105. Uchida A, Ooguri, T, Ishida, T, Ishida, Y. Incorporation of methionine into dimethyl-thiopropionic acid in the dinoflagellate Crypthecodinium cohnii. Nippon Suisan Gakkaishi1993; 59:851-855.

106. Stevenson, DE, Akhtar, M, Gani, D. Streptomyces L-methionine decarboxylase: Purifica-tion and properties of the enzyme and stereochemical course of substrate decarboxylation.Biochemistry 1990; 29:7660-7666.

107. Stevenson DE, Akhtar M, Gani D. L-methionine decarboxylase from Dryopteris felix-mas: Purification, characterization, substrate specificity, abortive transamination of thecoenzyme, and stereochemical courses of substrate decarboxylation and coenzymetransamination. Biochem 1990; 29:7631-7647.


108. Mazelis M. The pyridoxal phosphate-dependent oxidative decarboxylation of methionineby peroxide. I. Characteristics and properties of the reaction. J Biol Chem 1962; 237:104-108.

109. Mazelis M, Ingraham LL. The pyridoxal phosphate-dependent oxidative decarboxylationof methionine by peroxide. II. Identification of 3-methylthiopropylamide as the product.J Biol Chem 1962; 237:109-112.

110. Scislowski WD, Hokland BM, Davis van-Thienen WIA, Bremer J, Davis EJ. Methioninemetabolism by rat muscle and other organisms. Biochem J 1987; 247:35-40.

111. Walters DS, Steffens JC. Branched chain amino acid metabolism in the biosynthesis ofLycopersicon pennellii glucose esters. Plant Physiol 1990; 93:1544-1551.

112. Livesy G. Methionine degradation: Anabolic and catabolic. Trends Biochem Sci 1984; 9: 27-29.113. Dibner JJ, Knight CD. Conversion of 2-hydroxy-4-(methylthio)butanoic acid to L-methionine in

the chick: a stereospecific pathway. J Nutr 1984; 114:1716-1723.114. Mizayaki HH, Yang SF. Metabolism of 5-methylthioribose to methionine. Plant Physiol

1987; 84:277-281.115. Walsh C. Enzymatic Reaction Mechanisms. New York:Freeman. 1979.116. Miflin BJ, Lea PJ. Amino acid metabolism. Ann Rev Plant Physiol 1977; 28:299-329.117. Christen P, Metzler DE. Transaminases. New York:Wiley. 1985.118. Summers PS, Nolte KD, Cooper AJL, Borgeas H, Leustek T, Rhodes D, Hanson AD.

Identification and stereospecificty of the first three enzymes of 3-dimethylsulfoniopro-pionate biosynthesis in a chlorophyte alga. Plant Physiol 1998; 116:369-378.

119. Mudd SH, Datko AH. The S-methylmethionine cycle in Lemna paucicostata. Plant Physiol1990; 93:623-630.

120. Giovanelli J, Mudd SH, Datko A. Sulfur amino acids in plants. In: Stumpf PK Conn EE,eds. The Biochemistry of Plants, Vol. 5, Amino Acids and Derivatives. New York: AcademicPress, 1980:453-505.

121. James F, Paquet L, Sparace A, Gage DA, Hanson AD. Evidence implicating dimethyl-sulfoniopropionaldehyde as an intermediate in dimethylsulfoniopropionate biosynthesis.Plant Physiol 1995; 108:1439-1448.

122. Cooper ALJ, Hollander MM, Anders MW. Formation of highly reactive vinylglyoxylate(2-oxo-3-butenoate) from amino acids with good leaving groups in the γ-position.Toxicological implications and therapeutic potential. Biochem Pharmacol 1989;38:3895-3901.

123. Toney MD, Hohenester E, Cowan SW, Jansonius JN. Dialkylglycine decarboxylase structure:Bifunctional active site and alkali metal sites. Science 1993; 261:756-759.

124. Walton NJ, Peerless ACJ, Robins RJ, Rhodes MJC, Boswell HD, Robins DJ. Purification andproperties of putrescine N-methyltransferase from transformed roots of Datura stramoniumL. Planta 1994; 193:9-15.

125. Rhodes D, Gage DA, Cooper AJL, Hanson AD. S-methylmethionine conversion todimethylsulfoniopropionate: Evidence for an unusual transamination reaction. PlantPhysiol 1997; 115:1541-1548.

126. Kumar A, Altabella T, Taylor MA, Tiburico, AF. Recent advance in polyamine research.Trends Plant Sci 1997; 2:124-130.

127. Trossat C, Nolte KD, Hanson AD. Evidence that the pathway of dimethylsulfoniopro-pionate biosynthesis begins in the cytosol and ends in the chloroplast. Plant Physiol 1996;111:965-973.

128. Trossat C, Rathinasabapathi B, Weretylnik EA, Shen T-L, Huan Z-H, Gage DA, HansonAD. Salinity promotes accumulation of dimethylsulfoniopropionate and its precursorS-methylmethionine in chloroplasts. Plant Physiol 1998; 116:165-171.

129. Nomura M, Ishitani M, Takabe T, Rai AK, Takabe T. Synechococcus sp. PCC7942 transformedwith Escherichia coli bet genes produces glycine betaine from choline and acquiresresistance to salt stress. Plant Physiol 1995; 107:703-708.

130. Cossins, EA, Chen, L. Folates and one carbon metabolism in plants and fungi. Phytochemistry1997; 45:437-452.


131. Holmstrom KO, Welin B, Mandal A, Kristiansdottir I, Teeri T, Lamark T, Strom, AR,Palva ET. Production of the Escherichia coli betaine-aldehyde dehydrogenase, an enzymerequired for the synthesis of the osmoprotectant glycine betaine in transgenic plants.Plant J 1994; 6:749-758.

132. Lilius G, Holmberg N, Bulow L. Enhanced NaCl stress tolerance in transgenic tobaccoexpressing bacterial choline dehydrogenase. Biotechnol 1996; 14:177-180.

133. Holmberg N. Metabolic engineering: Approaches towards improved stress tolerance inmicroorganisms and plants. Doctoral Dissertation. Lund, Sweden: University of Lund. 1996:77-92.

134. Hayashi H, Mustardy L, Deshnium P, Ida M, Murata N. Transformation of Arabidopsisthaliana with the codA gene for choline oxidase; accumulation of glycinebetaine and enhancedtolerance to salt and cold stress. Plant J 1997; 12:133-142.

135. Hitz WD, Ladyman JAR, Hanson AD. Betaine synthesis and accumulation in barley (Hordeumvulgare) during field water-stress. Crop Sci 1982; 22:47-54.

136. Hanson AD, Gage DA. 3-Dimethylsulfoniopropionate biosynthesis and use by flowering plants. In:Kiene RP, Visscher PT, Keller MD, Kirst GO, eds. Biological and Environmental Chemistry ofDMSP and Related Sulfonium Compounds. New York: Plenum Press, 1996:75-86.

137. Pokorny M, Marcenko E, Keglevic D. Comparative studies of L- and D-methioninemetabolism in lower and higher plants. Phytochemistry 1970; 9:2175-2188.

Osmotic Stress Tolerance in Plants:Role of Proline and SulfurMetabolismsDesh Pal S. Verma

Osmotic stress, caused either due to the loss of water or increase in soil salinity, reducesgrowth and productivity of plants. The responses of plants to both drought and salinity

have many steps in common and some of them overlap with cold stress. Anincrease in external osmolarity results in an efflux of water from the interior, which bringsabout a reduction in the turgor pressure in the cell and a reduction in the cytoplasmicvolume. A decrease in cell volume elevates the concentration of various intracellular ions,which are toxic to the cell. To prevent volume change and loss of water, organisms generallyincrease the concentration of compatible solutes in response to osmotic stress. The majorcompatible solutes are K+, proline, glutamate, and quaternary ammonium compounds. Someother molecules that act as osmolytes include trehalose, glycerol, choline, s,s-dimethyl-sulfoniumacetate, stachydrine (N,N-dimethylproline, proline betaine), β-butyrobetaine,L-pipecolate, 5-hydroxyl-1-pipecolate, N,N-dimethylglycine, N-methylproline, glutamatebetaine and γ-aminobutyrate.1-3 Different organisms accumulate one or more of thesecompounds in response to drought or salinity.

Accumulation of proline in response to osmotic stress is very common in many plants.4

Our recent data suggests that the primary role of proline in osmoprotection may not besolely as an osmoregulatory osmolyte, but it also helps the cell to overcome oxidative stress.Other known attributes of proline, such as protecting enzymes from denaturation,5 interactingwith membrane systems,6 regulating cytosolic acidity,7 scavenging free radicals,8 balancing theratio of NADH/NAD+,9 and acting as a energy source10 may be more important for theoverall health of the plant under osmotic stress. We have demonstrated that high levels ofendogenous proline help reduce free radicals generated during oxidative stress induced bythe osmotic stress.11 Furthermore, free radicals produced during the oxidative stress causeserious damage to SH groups, oxidize cystine and methionine, which may impair the functionof proteins.12 We have shown that the regulation of sulfur metabolism can reduce oxidativestress and thus help alleviate osmotic stress.

Osmoregulation in Microorganisms

Proline as an Osmoprotectant Although potassium is the most prevalent cation that acts as a major osmolyte in

bacteria,13,14 accumulation of proline has been found to benefit many microorganisms in

CHAPTER 8



sustaining osmotic stress. The role of proline in osmotic stress tolerance was deduced fromthe observation that exogenously applied proline could alleviate the growth inhibition ofbacteria imposed by osmotic stress. Proline is taken up from the medium by an activetransport system, and often proline levels are proportional to the osmotic strength of themedium.15

Csonka demonstrated that proline-overproducing mutants of Salmonella typhimuriumexhibit enhanced tolerance to osmotic stress.16 In bacteria, proline synthesis is controlledby feedback regulation of the first enzyme of the proline biosynthesis pathway, γ-glutamylkinase. Mutations resulting in proline overproduction and enhanced osmotic tolerancewere shown to be located in the proB gene, encoding γ-glutamyl kinase. One of the mostpronounced osmotic stress tolerant mutants (proB74) had a single base pair change whichresulted in a 100-fold loss of sensitivity of g-glutamyl kinase to feedback inhibition byproline.16,17 When this mutated gene was transferred from S. typhimurium to otherenteric bacteria, it showed an enhanced osmotic stress tolerance.18,19 These studies confirmedthat proline plays a crucial role in osmotolerance in bacteria.

Other Osmolytes Betaines and glycinebetaine (N,N,N-trimethlglycine) are widely used as osmolytes

by bacteria.20 Most bacteria are unable to synthesize betaines and depend on the transportof these compounds from their environment.1 The transport of betaines is mediated bythe proP and proU transport system, as used for proline.21- 23 It appears that other systemsfor transport also exist1,24 because proP and proU double mutants still grow well onglycine betaine though with a long lag in growth.

In yeast (Saccharomyces cerevisiae), glycerol seems to be the primary compatiblesolute produced under osmotic stress.25,26 Glycerol is synthesized in cytosol fromdihydroxyacetonephosphate, an intermediate of the glycolytic pathway catalyzed by anNADH-dependent glycerol-3-phosphate dehydrogenase (GPDH) and a phosphatase,respectively. The GPDH activity is enhanced several fold under osmotic stress.27 Somehigh salt tolerant algae, e.g., Dunaliella, also accumulate glycerol as an osmoprotectant.Trehalose is accumulated in many organisms and may serve as a non-osmoregulatoryprotector in stress conditions.28

Osmosensing and Signal Transduction Machinery The osmosensing machinery is well studied in E. coli and yeast. In E. coli, two proteins,

OmpF and OmpC, are involved in passive diffusion of small hydrophilic molecules acrossthe membrane.29 The expression of their genes, ompF and ompC, is affected in a reciprocalmanner by the osmolarity of the medium. As the osmolarity increases, the ompC gene ispreferentially activated, whereas a decrease in osmolarity results in the activation of theompF gene. The expression of these two genes is controlled at the transcriptional level byOmpR and EvnZ proteins. The OmpR is a DNA binding protein and specifically interactswith the promoter regions of the ompF and ompC genes.30,31 The EnvZ is a membraneprotein containing two membrane-spanning regions at its amino-terminus32 and functions asan environmental sensor.33 It has been demonstrated that the EnvZ is autophosphorylatedat the histidine residue (His243) in the presence of ATP, and the phosphate group is thentransferred to the OmpR protein.29 Thus, EnvZ acts as a protein kinase. The phosphorylatedOmpR binds with DNA and controls the transcription of the cognate genes. The EnvZ andOmpR phosphotransfer system has a resemblance to the family of two-component regulatorysystems involved in response to environmental stimuli.34 The osmosensing system must becoupled with the induction of genes involved in the synthesis of specific osmolyte. Theactual mechanism for this, however, is not yet understood.

155Osmotic Stress Tolerance in Plants: Role of Proline and Sulfur Metabolisms

In yeast (S. cerevisiae), a more complex signal transduction pathway exists that sensesthe osmotic condition of the cell and induces physiological changes that lead to the synthesis ofglycerol, which acts as an osmolyte.35-37 Yeast cells can accumulate glycerol in cytoplasm up toa concentration of 1 M.35 Several genes of the signal transduction cascade have recentlybeen isolated from yeast. The mechanism involved in osmosensing is briefly summarizedbelow.

The primary sensor of osmotic stress, Sln1, contains a classical “input-transmitter-receiver”structure and shares sequence similarities to both the histidine kinase and the responseregulator protein of the prokaryotic two-component systems.38 The transmitter domain(histine kinase) of Sln1 is similar to the histidine kinase modules of the bacterial regulators.Another similarity between Sln1 and bacterial sensor proteins is the presence of twohydrophobic transmembrane domains in the N-terminal region. This region also containsa membrane-bound signal-sensing domain which may be exposed on the cell surface. Severalsuppressor mutants have been isolated from either sln1∆ or slnts strains. One of themutants, ssk1∆, suppresses sln1∆ lethality. The deduced amino acid sequence of Ssk1 showshomology to the bacterial response regulators. Mutagenesis studies have shown that theunphosphorylated Ssk1 is functional.36 A new protein, Ypd1p, was recently isolated andshown to be a constituent of the two component system.37 The mutant of ypd1 is lethal butit can be suppressed by the overexpression of Ptp2 (tyrosine phosphatase). The ypd1 geneencodes a protein of 167 amino acids, which has similarity with the chemotactic CheAprotein of bacteria.

Early events of osmosensing start from the autophosphorylation of Sln1 at a His residuefollowed by a cascade of phosphorylation steps. The downstream osmotic signal transductionpathway is composed of three tiers of protein kinases, namely SSK2 and MAPKKKs (MAPkinase kinase kinases), MAPKK (MAP kinase kinase) and high osmolarity glycerol (HOG)response MAP (mitogen-activated protein) kinases39,40 (see chapter 2). SSK2 was found toact as an extragenic suppressor of the sln1D mutant.40 It has high homology with MAPKKKat the -COOH terminal region. Hog1 is not phosphorylated in pbs2-∆1 cells, suggestingthat phosphorylation of Hog1 requires Pbs2.

Both Ssk2p and Ssk22p interact with Ssk1p and HOG1 as shown by two hybrid analysis.The phosphorylated Ssk1p is non-functional. HOG1 and HOG4 genes were cloned bycomplementing yeast osmoregulation-defective mutants Osms. These mutants grow wellon YEPD medium but not on high-osmolarity medium and show reduced accumulationof glycerol. The HOG1 sequence contains a single large open reading frame encoding a proteinof 416 amino aids with a molecular weight of 47 kDa. Near the NH2-terminus of the predictedamino acid sequence of Hog1, a stretch contains the most conserved amino acids found inthis family of protein kinases. Two residues, corresponding to Thr174 and Tyr176, incomparable positions in the MAP kinases encoded by ERK2 and FUS3 are phosphory-lated in response to extracellular signals. Pbs2 is also a member of the MAP kinase kinasegene family. The mutant pbs2 is unable to grow in medium of high osmolarity.

Glycerol synthesis in yeast is limited by the activity of glycerol-3-phosphate dehydroge-nase (GPD1).27 gpd1∆ mutants produce little glycerol and are sensitive to osmotic stress. Asexpected, hog1∆ mutant fails to increase glycerol-3-phosphate dehydrogenase activity. Thus,the expression of GPD1 appears to be regulated through the HOG pathway. However,there may be Hog1-independent mechanisms mediating osmotic stress-induced glycerolaccumulation in yeast, since a hog1∆ mutant still shows glycerol accumulation duringosmotic stress, although at a reduced level. The gpd1∆ and hog1∆ double mutants are moresensitive to osmotic stress than gpd1∆ mutant.

By screening high osmolarity sensitive mutants, an SHO1 gene was identified,37 whichcan be rescued by transformation either SSK2 or SSK22. SHO1 encodes a protein of 367


amino acids with four hydrophobic transmembrane domains at the N-terminal region.The COOH terminal of the SHO1 contains an SH3 domain. The triple mutations of ssk2∆,ssk22∆, and sho1∆ completely abolish tyrosine phosphorylation of Hog1p and osmotic stresssensitivity. Sho1p appears to be a component of an alternate pathway that activates Pbs2pin response to high osmolarity. The HOG pathway thus controls the expression of genesinvolved in glycerol biosynthesis.

Two mammalian genes involved in osmoregulation have recently been identified. Thesegenes, encoding p38 and Jnkl, have high homology with HOG1 and belong to the MAPkinase gene family. Both genes can complement hog1∆ mutants.41,42 These results suggestthat the signal transduction machinery may be conserved in eukaryotes. We have recentlyisolated a plant homolog of the HOG1 gene (unpublished data). However, this gene isunable to complement the hog1∆ mutant of yeast. The plants have a large MAP kinase genefamily.43 Recently, Jonak et al (ref. 43a). reported that a specific MAP kinase p44MMK4 isfound to be activated by drought and cold stress but not by salt stress. Furthermore,the transcription of this gene is not induced by ABA. Recent studies from Zhu’s laboratory,44

have demonstrated the presence of ABA-dependent and ABA-independent pathways thatoverlap in the transduction of cold and osmotic stress signals in plants.

Osmotic Stress Tolerance in PlantsPlants have evolved a variety of adaptations to water deficit and high salinity. These

adaptations include45 developmental and structural traits, time of flowering, rooting patterns,leaf waxiness, and physiological mechanisms such as the ability to exclude salt or thecompartmentalization of ions within the cell.46 The biochemical traits, synthesis andaccumulation of compatible osmolytes and changes in patterns of carbon and nitrogenmetabolism are most important. Among the most prevalent osmolytes are proline andbetaines.47 These, however, appear to be non-osmoregulatory osmolytes since the con-centration of these compounds in most plants is not very high for them to significantlyaffect the osmoticum of the cell.

Regulation of Proline Biosynthesis in Plants The biosynthesis pathways for proline and betaine have been well studied.48,49 In

plants, proline is synthesized from glutamate and ornithine.4 We have demonstrated thatproline biosynthesis from glutamate is accomplished in two major steps and have isolatedboth gene encoding ∆1-pyrroline-5-caroxylate (P5C) reductase (P5CR) and P5C synthetase(P5CS). The latter is a bifunctional enzyme which catalyzes the first two steps in prolinebiosynthesis from glutamate; in E. coli these steps are catalyzed by two different enzymesencoded by two separate genes (proB and proA). In addition, we have demonstrated thatnitrogen flux to proline is tightly regulated.50 Under normal conditions, ornithine is theprimary source of nitrogen for proline synthesis while glutamate takes precedent overornithine under stress conditions. Accordingly, the genes encoding ornithine aminotransferase (OAT) and P5CS are reciprocally regulated.50 Furthermore, we have demonstratedthat the accumulation of proline is tightly controlled by the end product of the pathwaywhen it is synthesized from glutamate and degraded by proline dehydrogenase (PDH).11,51, 52 Aproline cycle is thus established (Fig. 8.1).48

In order to determine the relationship between proline concentration and the levelsof P5CS and PDH gene expression, we first isolated a PDH gene from Arabidopsis.52 ThemRNA levels of P5CS and PDH were measured along with the free proline contents.Under osmotic stress, the P5CS transcription is significantly induced and results in highlevels of proline synthesis. When osmotic stress is released, P5CS transcript declines to thenormal levels, but at the same time the PDH transcript significantly increases. Consequently,


the free proline level declines following the induction of the PDH gene. The inhibition ofPDH by osmotic stress, both in bacteria and plants,53,54 may be a key factor in theaccumulation of proline under stress conditions.

Proline Induces Proline-Degradation, but Under Osmotic Stress Conditionsthis Degradation is Inhibited

In yeast and bacteria, the proline oxidase is induced by adding proline to the medium.55

The bacterial PutA protein not only has proline dehydrogenase and P5C dehydrogenaseactivities, it also functions as a repressor of the put operon.55 When proline is provided, thePutA protein binds with proline. The transcription of the put operon is induced and prolinedegradation starts.56,57 In plants, our results indicate that PDH transcription is significantlyinduced by exogenously applied proline. However, the level of PDH mRNA induced byproline was reduced during saline stress. During stress, as the free proline concentrationincreases, the PDH expression remains at a low level.52 Conversely, the transcription ofPDH is significantly induced within two hours upon removal of the osmotic stress. Theproline concentration has been shown to reach over 125 mM in NaCl-adapted tobaccocells.58 The above results suggest that a transcriptional mechanism may be involved inorder to prevent proline-induced proline degradation during osmotic stress. This mechanismmay avoid the futile cycling and would precisely control the level of enzymes needed fordegradation of proline in response to osmotic stress to ensure proline accumulation.

Fig. 8.1. Proline cycle in plants. Increase in the synthesis of proline during stress conditions andits degradation to glutamate during recovery from stress maintains a contenuous flux of nitrogenand energy to the plant. The enzymes involved in the synthesis are P5CS, catalysing step 4; P5CR,catalysing step 6. For the degradation of proline, the enzymes involved are proline dehydrogenasefor step 1 and P5C dehydrogenase for step 3. Reactions 2 and 5 are the same and are autocatalytic.


Proline Cycle and the Role of Proline Degradation During Recovery of Plantsfrom Stress

Proline is one of the few amino acids that can be used as a sole source of carbon andnitrogen, and as shown above, the synthesis and degradation of proline is tightly regulated. InE. coli, PDH couples proline oxidation to the reduction of FAD cofactor, which is bound tothe PutA protein and thus delivers electrons to the membrane-associated electron transportchain.56,57 Removal of FAD retains P5C dehydrogenase activity but not the PDH activityof the PutA protein. Plant PDH is also a flavoprotein located in the inner membrane ofmitochondria,54 suggesting that proline oxidation donates electrons to the respiratory electrontransport chain and thus provides energy during recovery from stress. This was also suggestedfrom the studies on root nodules.9 In soybean root nodules, the proline concentration isvery high and it has been proposed that the NADP+ generated during proline synthesisprovides cofactor for the synthesis of the purine precursor, ribose 5-phosphate, and thusregulating the synthesis of purines.9 Proline is oxidized by bacteroids which provide muchrequired energy for symbiotic nitrogen fixation.

In yeast, it has been reported that the proline degradation is regulated by nitrogensources and both PUT1 and PUT2 genes have been shown to be regulated by nitrogen. 55,59,60 Inmaize, proline degradation was shown to be inhibited to 25% of the control levels inwater-stressed mitochondria. However, only PDH was inhibited, while P5C dehydrogenasewas not affected.54 Our results indicate that exogenously applied proline significantlyinduces PDH expression. Removal of osmotic stress also induced PDH expression, resulting inthe decline of proline levels. These data indicate that the PDH expression level is directlycorrelated with the increase in proline degradation, suggesting that proline dehydrogenaseis the rate limiting enzyme in the proline degradation pathway. Since accumulation ofproline is not deleterious to the cell, it acts as a reserve for nitrogen and energy and mayhelp plant recover rapidly from stress.52 The proline cycle (Fig. 8.1) thus helps maintain anoptimum flow of nitrogen and energy in the cell.

Accumulation of Other Osmolytes

Accumulation of Sugar Alcohol Sugar alcohols, also called polyols or polyalcohol, have many hydroxyl groups, and

could possibly take the place of water in biopolymers of cell cytoplasm and help maintainthe function of enzymes and membranes at a time when water is limited due to osmoticstress.61,62 Furthermore, sugar alcohols are inert and harmless to enzymes even at veryhigh concentrations.63 In bacteria and animals, the function of sugar alcohols in osmoregulation iswell studied. In plants, sugar alcohols are not as common as proline, glycine betaine, andcholine-0-sulfate. Mannitol-1- phosphate deyhdrogenase is an enzyme that regulates mannitolsynthesis in bacteria but plants do not normally produce and accumulate mannitol. Theexpression of the bacterial mt1D gene results in the synthesis and accumulation of mannitol intransgenic plants.64 The mannitol concentration in leaves and roots of the transformedplants was estimated to be at a level of 100 mM. When the control and transgenic plantswere subjected to salinity stress, the plants producing mannitol were found to havean increased ability to tolerate high salinity. These results clearly demonstrated thatoverproducing sugar alcohols also helps plants deal with high salinity stress.

Betaines Many organic molecules such as β-alanine betaine, proline betaine, and hydroxyproline

betaine, act as effective non-osmoregulatory osmolytes in plants.3,65 Different osmolytesappear to have different selective advantages in a particular stress environment and in a


given species. Proline betaine and hydroxyproline betaine are found in plants adapted tovery dry or saline environments such as species of Citrus.66 It has been demonstrated thatproline betaine and hydroxyproline betaine are much better osmoprotectants than proline(our unpublished data), particularly in chronically dry environments.3 The pathwayinvolved in the synthesis of proline betaine has, however, not yet been worked out and nogene involved in this pathway has so far been identified. Once proline methyltransferasesare isolated and characterized, their genes can be used for plant transformation. Expression ofthese genes in plants producing high levels of proline may convert some of this excessproline into dimethyl proline and thus provide excellent protection from osmotic stress.

Sulfur Osmolytes Choline-0-sulfate is another important osmolyte in plants.65,66 Choline-0-sulfate is

formed from choline. The choline sulfotransferase has not yet been identified in plants,but this enzyme has been identified in both fungi and bacteria. The sulfate salinity leads tohigher choline-0-sulfate accumulation.3 With an increase in choline-0-sulfate, a proportionaldecrease in glycine betaine level is observed.2 In some green alga, red algae and brownalgae and in the higher plants, e.g.,Wedelia biflora,65 β-dimethyl sulfoniopropioate (DMSP),a sulfur osmolyte, accumulates. DMSP originates from methionine via methylation. Theenzyme S-adenosylmethionine:methionone S-methyltransferrase, catalyzing the first stepof DMSP synthesis, has recently been purified.67 It appears that the regulation ofnitrogen and sulfur metabolisms may influence the type of osmolytes which a givenspecies accumulates.

Accumulation of Proline in Transgenic Plants Expressing ElevatedLevels of P5CS

We have earlier demonstrated that P5CS is the rate limiting enzyme in the prolinebiosynthesis pathway and the level of P5CR has no effect on proline synthesis.68,69 Weintroduced a Vigna P5CS cDNA under the control of 35S promoter in tobacco plants andassessed the level of proline produced. Transgenic lines that produced high levels of VignaP5CS mRNA and proteins were analyzed and the effect of proline accumulation on planttolerance to water stress was assessed. Plants expressing high levels of P5CS accumulatedhigh levels of proline.69 Transgenic lines that did not produce Vigna P5CS mRNA norP5CS protein were found to have as low levels of proline as the control plants. Prolinelevels in leaves of 10 transgenic lines ranged from 830 to 1590 µg/g fresh weight of leaves(on average:1100 µg/g), compared to 80 to 89 µg/g in control plants. Corresponding to thelevel of expression of P5CS, proline contents were increased in transgenic lines. A directcorrelation between P5CS expression level and proline accumulation in the transgenicplants confirms that P5C synthetase is rate limiting in proline biosynthesis.

Data on amino acid analysis showed that accumulation of proline occurs at theexpense of glutamate.69 This indicates that availability of glutamate may act as a factor forproline overproduction under water or salt stress. This conclusion is consistent with ourobservation that transgenic plants expressing high levels of P5CS and supplied with20 m M NH4NO3 produced nearly 25 times more proline than did the pBI121 controls.69

A recent study from B. Hirel’s group showed (unpublished data) that antisense of GS inphloem cells reduces the level of proline, rendering plants sensitive to osmotic stress. Thus,proline synthesis is tightly coupled with nitrogen assimilation. Furthermore, the source ofnitrogen for proline is also regulated, as was observed by our studies on OAT expression.4


Fig. 8.2. Effect of free radicals-induced damage due to osmotic stress as measured by malondialdehyde(MDA) production, a measure of lipid peroxidation reaction. (A) Effect of externally added prolineon the level of MDA produced in transgenic tobacco cell lines after treatment with 250 mM NaCl for8 h. (B) Effect of endogenously accumulated proline on MDA content in 14 day old seedlings of wildtype, P5CS and P5CS129A seedlings.

Feedback Inhibition of Proline Biosynthesis Earlier experiments suggested that proline accumulation in plants under stress may

involve the loss of feedback regulation due to a conformational change in the P5CSprotein.70,71 In bacteria, proline biosynthesis has been shown to be regulated by the endproduct inhibition of γ-GK activity.72 A Salmonella typhimurium mutant resistant to a toxicproline analog accumulated proline and showed enhanced tolerance to osmotic stress.73

The mutation was due to the change of an aspartate (at position 107) to asparagine in theγ-GK, resulting in a mutant γ-GK which was much less sensitive to proline inhibition.16,74

Our experiments revealed that the conserved aspartate residue (at position 128) in theVigna P5CS is not involved in the feedback inhibition. Using site-directed mutagenesis, areplacement of phenylalanine at position 129 by alanine in Vigna P5CS (P5CSF129A) wascreated. This mutant enzyme was shown to retain similar kinetic characteristics as thewild type P5CS, but its feedback inhibition was virtually eliminated.51

We compared plants overexpressing a wild type form of Vigna P5CS and themutant enzyme P5CSF129A, whose feedback inhibition was eliminated. These two groupsof transgenic plants expressed comparable levels of Vigna P5CS mRNA and proteins asrevealed by Northern and Western blot analyses. Under normal conditions, P5CSF129A


plants accumulated about two-fold more proline over the plants expressing Vigna wild typeP5CS.10 This difference was further increased in plants under salinity stress,11 demonstratingthat feedback regulation of P5CS does play a role in controlling the level of proline inplants in both normal and stress conditions.

Proline Accumulation Confers Osmoprotection Proline levels are increased in both control and transgenic plants after drought treatment.

These values increased from about 80 µg proline/g fresh leaf (before stress) to about 3000µg/g (after stress) in control (wild type and pBI121) plants, and from 1000 µg/g to anaverage of 6500 µg/g in transgenic lines.69 While the proline contents were approximately14-fold greater in transgenic lines than in control plants before stress, only about 2-foldgreater after stress. Control plants started wilting in 5-6 days after drought treatment, whilewilting was delayed by at least two to three days in transgenic plants and the wilting wasmore severe in controls than in transgenic plants. High constitutive levels of proline inP5CS transgenic lines may be responsible for the observed effect on wilting rather than theinduced level of proline. Shinozaki’s group has recently constructed P5CS antisenseAribidopsis plants and has demonstrated (personal communication) that the transgenicplants with reduced P5CS are more sensitive to osmotic stress suggesting a direct role ofproline in osmoprotective machinery in plants.

The elevated levels of proline, either from exogenous addition in the medium orproduced endogenously effectively reduces free radical levels caused by salinity stress(Fig. 8.2). This was determined by measuring the levels of melondialdehyde (MDA), amarker for lipid peroxidation and membrane damage due to free radicals. The resultsof externally added proline to suspension culture cells and the transgenic seedlingsoverproducing proline due to P5CS were very similar.10 These data clearly show thatproline confers a significantly increased ability for the transgenic seedlings to grow in mediacontaining NaCl (Fig. 8.3). These findings shed new light on the regulation of prolinebiosynthesis in plants and its role in reducing oxidative damage conferred by the osmoticstress. Figure 8.4 summarizes different roles of proline in plan metabolism and opens thepossibility of improving crops for stress tolerance through genetic engineering for prolinesynthesis.

Fig. 8.3. Overexpression of Vigna P5CS and its mutagenized derivative devoid of feedbackinhibition by proline (F129A), and the effect of overproduction of proline on the ability oftrangennic tobacco seeds to germinate on 200 mM NaCl. Pictures are taken 20 days aftergermination. WT, wild type control seeds. For details, see ref. 10.

A B C


Role of Sulfur Metabolism in Osmotic Stress Tolerance Proteins, sulfated polysaccharides, sulfolipids, coenzymes and other sulfur-containing

secondary compounds are actively involved in cellular metabolism. Sulfate must be activatedin order to be used in cellular metabolism. This activation is achieved by the enzymes ATP-sulfurylase forming adenosine 5'-phosphosulfate (APS) and the APS-kinase formingphosphoadenosine 5'-phosphosulfate (PAPS).75 The PAPS is further reduced to sulfite andsulfide, which contribute sulfur to cysteine and methionine. Some of the intermediatesand their derivatives in the sulfur assimilation pathway, such as sulfite,75 are toxic to thecell when accumulated beyond certain levels. Under oxidative stress imposed by osmoticstress, the SH groups of proteins are likely to be oxidized, which may impair the functionof many proteins.12 To avoid accumulation of the sulfur compounds to toxic levels, a sensitiveflux (rate of flow) of the intermediates in the sulfur assimilation pathway is necessary. A3'(2'), 5'- diphosphonucleoside 3'(2')-phosphohydrolase (DPNPase) catalyzes the conversionof PAPS to APS in vitro. This enzyme is widely distributed from algae to plants and suggests thepresence of a futile cycle (substrate cycle). The futile cycle is one of the most sensitive fluxcontrol systems in metabolic pathways.

In yeast, a halotolerance gene (HAL2) was identified by functional assay of supporting thegrowth of cells under high salinity stress.76 The yeast HAL2 gene is identical to MET22.The yeast met22 mutant is a methionine auxotroph and can not use sulfate, sulfite or sulfide assulfur sources. However, the mutant exhibits wild type activities of the enzymes necessary toassimilate sulfate and has a normal sulfur uptake system. HAL2 also shows high homologywith the E. coli cysQ gene.77 The cysQ mutant is a cysteine auxotroph but mutations whichresulted in sulfate transport defects compensated for cysQ mutation. Over expression ofHAL2 in yeast improved salt tolerance.74

The protein encoded by yeast HAL2 gene is shown to have the activity of 3'(2'),5'-bisphosphatenucleotidase (DPNPase) with both PAPS and PAP as substrates.78 We have isolated a planthomolog of HAL2 gene from rice (RHL) and enzymatic studies on the expressed protein

Fig. 8.4. Possible roles of proline in protecting plants from osmotic stress.The other roles, besides being an osmolyte, appear to be more importantfor reducing osmotic stress-induced oxidative stress.


confirmed that it encodes a DPNPase. The RHL cDNA complemented both cysQ and met22mutations. Thus, we demonstrated that the proteins encoded by cysQ, HAL2 and RHL geneshave the same function in sulfur assimilation pathways in E. coli, yeast and plants. Thisenzyme, together with APS kinase, appears to catalyze a futile cycle in sulfur assimilationpathway (Fig. 8.5).

Similar to the Chlorella enzyme, the rice DPNPase is inhibited by Ca2+ and has optimal at9. The optimum Mg2+ concentration of DPNPase is about 2.5 mM. Since the RHL enzymeand the Chlorella DPNPase have the same substrate specificity, similar kinetics and bothdepend on Mg2+ and, inhibited by Ca2+, we believe the two are the same enzyme. DPNPase,together with APS sulfotransferase, transfers sulfur from PAPS to a thio carrier which isfurther reduced. The isolation of the RHL gene and the complementation of yeast HAL2and E. coli cysQ mutants provided direct evidence that DPNPase is involved in sulfurreduction in plants.

The assimilation of sulfur starts with sulfur activation. The first enzyme, ATP-sulfurylase, catalyzes APS synthesis. Since the equilibrium for APS formation is far to theleft, the second reaction catalyzed by APS kinase, to phosphorylate APS at the 3' position,plays an important role in pulling the first reaction forward. [Consequently, the PAPS accumulatesand an enzyme that controls the PAPS pool and removes unnecessary PAPS]. DPNPaseperforms this function. Murguía et al78 suggested that PAPS is the substrate for the yeastHAL2 encoded enzyme and the function of the HAL2 gene product is to remove PAP. Thishypothesis explains some phenotypes of the HAL2 mutant, met22; however, it does notexplain several facts.52

The phenotypes of cysQ and met22 can be explained if PAPS is the native substrate.The cysQ gene product converts PAPS to APS and APS kinase catalyzes APS to PAPS. Thetwo enzymes run a futile cycle in the sulfur activation pathway (Fig. 8.5). In the cysQ mutant,the PAPS accumulates immediately which is toxic to the cell.76 When cysteine is provided to E coli, thesulfate uptake system is inhibited by feedback regulation and the cell stops synthesizing theenzymes involved in the sulfur assimilatory pathway. Therefore, the toxic PAPS is not accumulatedand the cysQ mutant grows normally. When sulfite is provided, it can be easily converted tocysteine without producing PAPS. The same principles may apply in yeast.

It has been demonstrated that fructose 1,6-biphosphatase (FBPase) and DPNPasebelong to the same structural protein family.80 FBPase is an allosterically regulatedenzyme which controls the flux of glycolysis by forming a “futile cycle” with phosphofructokinase(PFK). Furthermore, FBPase is also regulated by cellular redox and the oxidized form isinactive.81 If DPNPase is also regulated by cellular redox, the free radicals generated by

Fig. 8.5. The futile sulfur cycle and the role of DPNPase in the control of this cycle, leading to theregulation of thr flux of active sulfur. For details, see ref. 52.


osmotic stress will render the enzyme inactive and thus disturb the balance of the sulfurassimilation pathway. Consequently, overexpression of the HAL2 gene may help the cellrestore the sulfur flux under stress conditions and confer salt tolerance. In addition, thecations accumulated during osmotic stress may also inhibit the DPNPase enzyme activity,and overexpression of the HAL2 gene may overcome this problem.

The product of RHL gene (DPNPase) converts PAPS to APS controlling the sulfurflux and thus may reduce the accumulation of the toxic compounds. Gläser et al (1993)reported that supplying methionine improved salt tolerance in yeast. This phenotype couldbe due to the fact that availability of methionine inhibits sulfate uptake.82 Sequesteringtoxic sulfur compounds may be a common phenomenon under salt stress. In plants,choline-O-sulfate (an osmolyte) may sequester extra sulfate under stress conditions.3

Although the transcription of the DPNPase gene is not increased by salt, the activityof the DPNPase enzyme and the yeast HAL2 enzyme are increased by K+. Osmotic stressincreases the K+ uptake.82 This indicates that the RHL enzyme may response to salt stressat the protein level. Overexpression of RHL in plants produces more glutathione, renderingplant cells more resistant to heavy metal ions. These cells also produce less free radicals.The latter may be the primary reason why HAL2 overexpression shows osmotic protection bylowering oxidative damage, as does the proline.

A Possible Role of DPNPase in Salt ToleranceSince our results have demonstrated that salt stress causes free oxygen radical production

and oxidative damage, it is likely that salt stress causes some of the HAL2 enzyme to loseactivity due to the oxidation of protein. The decrease in DPNPase activity disturbs thesulfur metabolism which is required to remove oxidative stress. Because the DPNPase issensitive to Li+ and Na+, Murguia et al,78 proposed that Ha12 loses enzyme activity duringsalt stress due to the increase of cellular Li+ or Na+ and overexpression of DPNPase overcomesthe problem of sulfur reduction.

Considering that the sulfur-rich compounds play significant roles in antioxidationstress and that overexpression of the HAL2 gene in yeast conferred osmotolerance, the roleof DPNPase in controlling the “futile cycle” in sulfur assimilation pathways is important.

Overexpression of Plant HAL2 Gene Confers Reduction in FreeRadical Production and in Heavy Metal Toxicity

Plants exposed to various oxidative stresses including heavy metal stress, exhibit anincrease in lipid peroxidation due to excessive free radical generation.83 Interaction of heavymetals with functional -SH groups has been proposed as the mechanism of inhibition ofseveral physiological reactions.84,85 A rapid decline in cellular glutathione (GSH) levelswas shown in plant cells exposed to cadmium.86,87 It has also been proposed that glutathionemay be of significance in the protection of plants against the harmful effects of activeoxygen species and free radicals.88 Exposure of plants to excess Cd particularly enhancesthe demand for organic sulfur and even sulfide.89 The metal chelating phytochelatins aresynthesized from glutathione by plants exposed to metals such as Cd2+, Cu2+ and Zn2+.90

The studies on the overexpression of P5CS, mutated P5CS and HAL2 have clearlydemonstrated that metabolic engineering to produce specific compounds is now a reality.For ensuring a continuous supply of nitrogen for proline synthesis a multiple gene cassettecontaining constitutive GS, and stress-inducible P5CS may be necessary for producingoptimum levels of proline. Coexpression of HAL2 and superoxide dismutase may becomplementory in reducing drought or salt stress. A significant reduction in free radicalformation may, however, be deleterious from the point of pathogen attack.


References1. Csonka LN. Physiological and genetic responses of bacteria to osmotic stress. Microbiological

Reviews. 1989; 51:121-147.2. Hanson AD, Rathinasabapathi B, Chamberlin B, Gage DA. Comparative physiological

evidence that β-alanine betaine and choline-O-sulfate act as compatible osmolytes inhalophytic Limonium species. Plant Physiol. 1991; 97,1199-1205.

3. Hanson AD, Rathinasabapathi B, Rivoal J, Burnet M, Dillon MO, Gage DA.Osmoprotective compounds in the plumbaginaceae: A natural experiment in metabolicengineering of stress tolerance. Proc Natl Acad Sci USA 1994; 29,306-310.

4. Delauney AJ, Verma DPS. Proline biosynthesis and osmoregulation in plants. The PlantJournal. 1993; 4,215-223

5. Rajendrakumar CS, Reddy BV, Reddy AR. Proline-protein interaction: Protection ofstructural and functional integrity of M4 lactate dehydrogenase. Biochem Biophys ResCommun. 1994; 201,957-963.

6. Rudolph AS, Crowe, JH, Crowe LM. Effects of three stablizing agents-proline, betaine,and trehalose on membrane phospholipids. Arch Bioche Biophys. 1986; 245,134-143.

7. Venekamp JH. Regulation of cytosol acidity in plants under conditions of drought. PhysiolPlantarum. 1989; 76,112-117.

8. Smirnof N, Cumbes QJ. Hydroxyl radical scavenging activity of compatible solutes.Phytochemistry. 1989; 28,1057-1060.

9. Alia, Saradhi PP. Suppression in mitochondrial electron transport is the primary causebehind stress induced proline accumulation. Biochem Biophys Res Commun, 1991;193:54-58.

10. Kohl DH, Schubert KR, Carter MB, Hagedorn CH, Shearer G. Proline metabolism inN2-fixing root nodules: Energy transfer and regulation of synthesis. Proc Natl Acad SciUSA 1998; 85,2036-2040.

11. Hong Z, Lakinini K, Zhang Z, Verma DPS. Removal of feedback inhibition of ∆1-pyrroline-5-carboxylate synthetase enhances proline accumulation and removes oxidative stress1999; submitted.

12. Smirnof N. The role of active oxygen in the response of plants to water deficit and desiccation.New Phytol 1993; 125:27-58.

13. Christian JBH, Waltho J. The sodium and potassium content of non-halophilic bacteriain relation to salt tolerance. J Gen Microbiol. 1961; 25:292-335.

14. Walderhaug MO, Dosch DC, Epstein W. Potassium transport in bacteria. In: Ion transportin prokaryotes. Rosen BP, Silver S, eds. San Diego: Acadmic Press, 1987; 85-130.

15. Britten RJ, McClure FT. The amino acid pool in Escherichia coli. Bacteriol Rev 1962;26:292-335.

16. Csonka LN. Regulation of cytopalsmic proline levels in Salmonella typhimurium: Effectof osmotic stress on synthesis, degradation, and cellular retention of proline. J Bacteriol1988; 170:2374-2378.

17. Smith LT. Characterization of g-glutamyl kinase from Escerichia coli that confers prolineoverproduction and osmotic tolerance. J Bacteriol 1985; 164:1088-1093.

18. Jakowec MW, Smith LT, Dandekar AM. Recombinant plasmid conferring proline overproductionand osmotic tolerance. Appl Environ Microbiol 1985; 50:441-446.

19. LeRudulier D, Yang SS, Csonka LN. Nitrogen fixation in Klebsiella pneumoniae duringosmotic stress: Effect of exogenous proline or a proline overproducing plasmid. BiophysActa 1982; 719:273-283.

20. Hanson AD, Hitz WD. Metabolic responses of mesophytes to plant water deficits. AnnRev Plant Physiol. 1982; 33:163-203.

21. Cairney J, Booth IR, Higgins CF. Salmonella typhimurium proP gene encodes a transportsystem for the osmoprotectant betaine. J Bacteriol 1985a; 164:1218-1223.

22. Cairney J, Booth IR, Higgins CF. Osmoregulation of gene expression in Salmonellatyphimurium: proU gene encodes an osmotically induced betaine transport system.J Bacteriol,1985b; 164:1224-1232.


23. May, G, Faatz E, Villarejo M, Bremer E. Binding protein dependent transport of glycinebetaine and its osmotic regulation in Escerichia coli K12. Mol Gen Genet 1983; 205:225-233.

24. LeRudulier D, Bouillard L. Glycine betaine, an osmotic effector in Klebsiella pneumoniaeand other members of the Enterobacteriaceae. Appl Environ Microbiol,1986; 46:152-159.

25. Bloomberg A, Adler, L. Physiology of osmotolerance in fungi. Adv Microbiol Physiol1992; 33:145-212.

26. Mager WH, Varel JCS. Osmostress response of the yeast Saccharomyces. Mol Microbiol,1993; 10:253-258.

27. Albertyn JS, Hohman JM, Thevelein, and Prior (1994). GPD1, which encodes glycerol-3-phosphate dehydrogenase, is essential for growth under osmotic stress in Saccaromyces cerevisiae,and its expression is regulated by the high-osmolarity glycerol response pathway. MolCell Biol 1994; 14:4135-4144.

28. Streeter J. Accumulation of a,a-trehalose by Rhizobium bacteria and bacteroids. J Bact1985; 164:78-84.

29. Aiba H, Mizuno T, Mizushima S. Transfer of phosphoryl group between two regulatoryproteins involved in osmoregulatory expression of the ompF and ompC genes in Escherichia coli.J Biol Chem 1989; 264:8563-8567.

30. JoY-L, Nara F, Ichihara S, Mizuno T, Mizushima S. Purification and characterization ofthe OmpR protein, a positive regulator involved in osmoregulatory expression of theompF and ompC genes in Escerichia coli. J Biol Chem 1986; 261:15252-15256.

31. Norioka S, Ramakrisshnan G, Ikenaka K, Inouye M. Interaction of a transcriptional activator,OmpR, with reciprocally osmoregulated genes, ompF and ompC, of Escerichia coli.J Biol Chem 1986; 261:17113-17119.

32. Forst S, Comeau D, Norioka S, Inouye M. Localization and membrane topology of EnvZ,a protein involved in osmoregulation of OmpF and OmpC on Escherchia coli. J Biol Chem,1987; 262:16433-16438.

33. Hall MN, Silhavy TJ. Genetic analysis of the major outer membrane proteins ofEscherchia coli K-12. Annu Rev Genet 1981b; 15:91-142.

34. Albright LM, Huala E, Ausubel FM. Prokaryotic signal transduction mediated by sensorand regulator protein pairs. Annu Rev Genet 1989; 23:311.

35. Brewster JL, DeValoir T, Dwyer ND, Winter E, Gustin MC. An osmosensing signal transductionpathway in yeast. Science. 1993; 259:1760-1763.

36. Maeda T, Wurgler-Murphy SM, Saito H. A two-component system that regulates anosmosensing MAP kinase cascade in yeast. Nature. 1994; 369:242-245.

37. Posas F, Wurgler-Murpy SM, Maeda T, Witten EA, Thai TC, Saito H. Yeast HOG1 MAPkinase cascade is regulated by a multistep phosphorelay mechanism in the SLN1-YPD1-SSK1“Two-Component” osmosensor. Cell 1996; 86:865-875.

38. Ota IM, Varshaavsky A. A yeast protein similar to bacterial two-compound regulators.Science. 1993; 262:566-569.

39. Boguslawski G. PBS2, a yeast gene encoding a putative protein kinase, interacts with theRAS2 pathway and afects osmotic sensitivity of Saccharomyces cerevisiae. J Gen Microbiol1992; 138:2425-2432.

40. Maeda T, Takekawa M, Saito H. Activation of yeast PBS2 MAPKK by MAPKKKs or bybinding of an SH3-containing osmosensor. Science 1995; 269:554-558.

41. Han J, Lee JD, Bibbs L, Ulevitch RJ. A MAP kinase targeted by endotoxin andhyperosmolarity in mammalian cells. 1994; 265:808-811.

42. Galcheva-Gargova Z, Derijard B, Wu I-H, Davis RJ. An osmosensing signal transductionpathway in mammalian cells. Science 1994; 265:806-808.

43. Mizoguchi T, Hayashida N, Yamaguchi-Shinozaki K, Kamada H, Shinozaki K. ATMPKs:A gene family of MAP kinases in Arabidopsis thaliana. FEBS Letters. 1993; 336:440-444.43a. Jonak C, Kiegerl S, Ligterink W, Barker PJ, Huskisson NS. Stress signaling in plants:A mitogen-activated protein kinasee pathway is activated by cold and drought. Proc NatlAcad Sci USA 1996; 93:11274-11279.


44. Ishitani M, Xiong L, Stevenson B, Zhu J-K. Genetic Analysis of Osmotic and cold stresssignal transduction in Arabidopsis: Interactions and convergence of abscisic acid-dependentand abscisic acid-independent pathways. Plant Cell 1997; 9:1935-1949.

45. McCue KF, Hanson AD. Salt-inducible betaine aldehyde dehydrogenase from sugar beet:cDNA cloning and expression. Plant Mol Biol. 1990; 8:358-362.

46. Binzel ML, Hess FD, Bressan RA, Hasegawa PM. Intracellular compartmentation of ionsin salt-adapted tobacco cells. Plant Physiol. 1988; 86:607-614.

47. Yancey PH, Clark ME, Hand SC, Bowlus RD, Somero G. Living with water stress: Evolutionof osmolyte systemes. Science 1982; 217:1214-1222.

48. Verma DPS, Zhang C-S. Regulation of proline and arginine biosynthesis in plants. In:Singh BK, ed. Amino acid biosynthesis. Marcell and Decker Pub 1998:249-265.

49. Hanson AD, May AM, Grumet R, Bode J, Jamieson GC, Rhodes D. Betaine synthesis inchenopods: Localization in chloroplasts. Proc Natl Acad Sci USA 1985; 82:3678-3682.

50. Delauney AJ, Hu CA, Kishore PBK, Verma DPS. Cloning of ornithine δ-aminotransferasecDNA from Vigna aconitifolia by transcomplementation in Escherichia coli and regulation ofproline biosynthesis. J Biol Chem. 1993; 268:18673-18678.

51. Zhang C, Lu Q, Verma DPS. Removal of feedback inhibition of D1-pyrroline-5-carboxylatesynthetase, a bifunctional enzyme catalysing the first two steps of proline biosynthesis inplants. J Biol Chem. 1995; 270:20491-20496.

52. Peng Z, Verma DPS. Reciprocal regulation of ∆1-Pyrroline-5-carboxylate synthetase andproline dehydrogenase genes control probline levels during and after osmotic stress inplants. Mol Gen Gent. 1996; 253:334-341.

53. Ekena K, Maloy S. Regulation of proline utilization in Salmonella typhimurium: How docells avoid futile cycle. Mol Gen Genet. 1990; 220:492-494.

54. Rayapati PJ, Stewart CR. Solublization of proline dehydrogenase from maize mitochondria.Plant Physiol. 1991; 95:787-791.

55. Wang S, Brandriss MC. Proline utilization in Saccharomyces cervisiae: Analysis of clonedPUT1 gene. Mol and Cell Biol 1986; 6:2638-2645.

56. Wood JM. Membrane association of proline dehydrogenase in Escherichia coli is redoxdependent. Proc Nat Acad Sci USA 1987; 84:373-377.

57. De Spicer PO, Maloy S. PutA protein, a membrane-associated flavin dehydrogenase, actsas a redox-dependent transcriptional regulator. Proc Natl Acad Sci, USA 1993; 90:4295-4298.

58. Binzel ML, Hasegawa PM, Rhodes D, Handa S, Handa AK, Bressan RA. Soluteacuumulation in tobacco cells adapted to NaCl. Plant Physiol 1987; 84:1408-1415.

59. Wang S, Brandriss MC. Proline utilization in Saccharomyces cerevisiae: Sequence, regula-tion, and mitochondrial localization of the PUTI gene product. Mol Cell Biol 1987;7:4431-4440.

60. Brandris MC. Proline utilization in Saccharomyces cerevisiae: Analysis of cloned PUT2gene. Mol Cell Biol 1983; 3:1846-1856.

61. Schobert B. Is there an osmotic regulatory mechanism in algae and higher plants? J TheorBiol 1977; 68:17-26.

62. Ahmad I, Lahrher F, Stewart GR. Sorbitol, a compatible osmotic solute in Plantago maritima,New Phytol 1979; 82:671-678.

63. Bieleski RL, Pedgewell RJ. Sorbitol metabolism in nectarines from flowers of Rosaeae.Aust J Plant Physiol 1980; 7:15-25.

64. Tarczynski MM, Jensen RG, Bohnert HJ. Stress protection of transgenic tobacco by production ofthe osmolyte mannitol. Science 1993; 259:508-510.

65. Rhodes D, Hanson AD. Quaternary ammonium and tertiary sulfonium compounds inhigher plants. Annu Rev Plant Physiol Plant Mol Biol 1993; 44:357-384.

66. Hanson AD, Gage DA. Identification and determination by fast atom bombardment massspectometry of the compatible solute choline-o-sulfate in Limonium species and otherhalophytes. Aust J Plant Physiol 1991; 18:317-327.

67. James F, Nolte KD, Hanson AD. Purification and properties of S-adenosyl-l-methionine:L-methionine s-methyltransferase from Wollastonia biflora leaves. J Biol Chem 1995; 270:22344-22350.


68. Szoke A, Miao GH, Hong Z, Verma DPS. Subcellular location of D1-pyrroline-5-carboxylatereductase in root/nodules and leaf of soybean. Plant Physiol 1992; 99:1642-1649.

69. Kavi Kishore PB, Hong Z, Miao G, Hu C, Verma DPS. Overexpression of ∆1-pyrroline-5-carboxylate synthetase increases proline production and confers osmotol-erance in transgenicplants. Plant Physiol 1995; 108:1387-1394.

70. Boggess SF, Aspinall D, Paleg L. Stress metabolism IX. The significance of end-productinhibtion of proline biosynthesis and of compartmentation in relation to stress inducedproline accumulation. Aust J Plant Physiol 1976; 3:513-525

71. Boggess SF, Stewart CR, Aspinall D, Paleg L. Effect of water stress on proline synthesisfrom radioactive precursors. Plant Physiol 1976; 58:398-401.

72. SmithJC, Deutch AH, Rushlow KE. Purification and characteristics of a γ-glutamyl kinaseinvolved in E. coli proline biosynthesis. J Bacteriol 1984; 157:545-551.

73. Csonka LN. Proline overproduction results in enhanced osmoregulation in Salmonellatyphimurium. Mol Gen Genet 1981; 182:82-86.

74. Dandekar AM, Uratsu SL. A single base pair change in proline biosynthesis genes causesosotic stress tolerance. J Bacteriol 1988; 170:5943-5945.

75. Schmidt A, Jager K. Open questions about sulfur metabolism in plants. Ann Rev PlantPhysiol Plant Mol Biol 1992; 43:325-349.

76. Glaser HU, Thomas D, Gaxiola R, Montrichard F, Surdin-Kerjan Y, Serrano R. Salt toleranceand methionine biosynthesis in Saccharomyces cerevisiae involve a putative phosphatasegene. EMBOJ 1993; 3105-3110.

77. Neuwald AF, Krishnan BR, Brikun I, Kulakauskas S, Suziedelis K, Tomcsanyi T, LeyhTS, Berg De. cysQ: A gene needed for cysteine synthesis in Escherichia coli K-12 onlyduring aeobic growth. J Bacteriol 1992; 174:415-425.

78. Murguia JR, Belles JM, Serrano R. A salt-sensitive 3'(2'), 5'-bisphosphate nucleotidaseinvolved in sulfate activation. Science 1995; 267:232-243.

79. York JD, Ponder JW, Majerus PW. Definition of a metal-dependant/Li+-inhibitedphosphomonoesterase protein family based upon a conserved three-dimensional corestructure. Proc Natl Acad Sci USA 1995; 92:5149-5153.

80. Chardot T, Meunier JC. Properties of oxidized and reduced spinach (Spinacia oleracea) chlo-roplast fructros-1,6-bisphosphatase activated by various agents. J Biochem 1991; 278:787-791.

81. Markovich D, Forgo J, Strange G, Biber J, Murer H. Expression cloning of rat renalNa+/So4(2-) cotransport. Proc Natl Acad Sci USA 1993; 90:8073-8077.

82. Watad A-E, A, Reuveni M, Bressan RA, Hasegawa PM. Enchanced net K+ uptake capacity ofNaCl-adapted cells. Plant Physiol 1991; 95:1265-1269.

83. Chaoui A, Mazhoudi S, Ghorbal MH, Ferjani EE. Cadmium and zince induction of lipidperoxidation and effects on antioxidant enzyme activities in bean (Phaseolus vulgaris L.). PlantSci 1997; 127:139-147.

84. Shio Y, Tamai H, Sasa T. Effects of copper on photosynthetic electron transport systemsin spinach chloroplasts. Plant Cell Physiol 1978; 19:203-209.

85. Sandmann G, Boger P. Copper mediated lipid peroxidation processes in photosyntheticmembranes. Plant Physiol 1980; 66:797-800.

86. Goldsbrough PB, Mendum ML, Gupta SC. Effect of glutathione on phytochelation synthesisand cadmium tolerance in tomato cells. In: Rennenber et al, eds. Sulfur and Sulfur Assimilation inHigher Plants. The Hague, The Netherlands: SPB Academic Publishing 1990:185-189.

87. Scheller HV, Huang B, Hatch E, Goldsbrough PB. Phytochelation synthesis and glutathionelevels in response to heavy metals in tomato cells. Plant Physiol 1987; 85:1031-1035.

88. De Kok LJ, Stulen I. Role of glutathione in plants under oxidative stress. In: De Kok LJet al, eds. Sulfur Nutrition and Assimilation in Higher Plants. The Netherlands: SPBAcademic Publishing, 1993; 125-138.

89. Rauser WE. Metal binding peptides in plants. In: De Kok LJ et al, eds. Sulfur Nutrition andAssimilation in Higher Plants. The Hague, The Netherlands: SPB Academic Publishing 1993;239-251.

90. Meuwly P, Thibault P, Schwan AL, Rauser WE. Three families of thiol peptides are inducedby cadmium in maize. Plant J 1995; 7:391-400.

Index

A

ABA 3-7, 11, 14-20, 22-25, 41, 71, 74, 101,105-118, 158

ABA signaling 7, 19, 24, 110, 113ABA-dependent pathway 14, 15, 17, 18Abiotic stress 1-4, 7, 11, 87, 88ABRE 7, 15-18Arabidopsis 102-104, 113-115, 144, 158Arabidopsis thaliana 29, 38, 45, 49, 90, 144

B

Betaines 1, 4, 147, 156, 158, 160Biosynthesis 3, 5, 6, 12, 14, 15, 17-20, 24, 25,

33, 34, 38, 47, 49, 76, 131, 133-137,139-143, 145, 147, 156, 158, 161-163

BZIP transcription factor 17

C

Ca2+ 105-111, 113-117, 165CBF1 16, 17, 73, 74, 77, 81Chilling injury 63, 64, 66, 78, 88Chilling tolerance 63, 66-68, 77, 79, 88Cl- channel 105, 109, 115Cold acclimation 63, 70-78Compatible solute 29-32, 37-39, 47, 49, 127,

129-131, 155, 156COR genes 71, 73, 74, 77Cross protection 87, 88CRT/DRE 74

D

Development 1, 3, 5, 6, 14, 23, 24, 29, 30, 41,44, 48, 49, 63, 68, 84, 90, 93, 94, 106, 117,158

DRE/CRT 15, 16Drought 1, 2, 6, 7, 11-25, 29, 38, 41, 43, 48,

49, 63, 74, 83, 87, 88, 101, 105, 127, 132,145, 155, 158, 163, 166

F

Freezing tolerance 4, 63, 67, 69-78Frost sensitive mutants 5

G

Guard cell 5, 24, 36, 37, 101-106, 108-118

H

H+-ATPase 103, 106, 107, 110, 112, 117Heat shock protein 83, 84, 94HSF phosphorylation 93HSF regulation 92, 93

I

Ion homeostasis 31, 36, 48

K

K+ channel 103, 104, 106, 110, 111, 113, 115,117

M

MAP kinase cascade 19, 21, 22, 33, 34Marine algae 38, 127-131, 134, 143Metabolic engineering 39, 47, 48, 127, 143,

166Metabolite accumulation 29, 37MYB 13, 18, 19, 27MYC 13, 18, 19

N

Nicotiana 102, 103, 113

O

Osmolytes 12, 30, 32-34, 45, 51, 112, 127,128, 130, 147, 155, 156, 158, 160, 161

Osmoprotectants 1, 14, 30, 161Osmoregulation 36, 49, 155, 157, 158, 160Osmotic adjustment 7, 30-32, 46, 47, 128,

131, 145Osmotic stress 2, 5, 11, 18-22, 29, 31-35,

37-39, 44, 127, 129, 143, 146, 155-164,166


Oxidative stress 42, 43, 47, 66, 87, 88, 155,164, 166

P

Plasma membrane 13, 31, 34, 35, 40, 41, 44,45, 69, 70, 72, 76, 103-112, 115-117

Proline 1, 12-14, 29, 34, 37, 38, 43, 48, 49, 72,76, 77, 127, 155, 156, 158-164, 166

Protein dephosphorylation 19, 113Protein phosphorylation 24, 105, 110, 112,

113, 115

Q

QTLs 3, 7Quantitative trait loci 3Quaternary ammonium compounds 1, 127,

155

R

Reactive oxygen species (ROS) 2, 3, 42, 43, 48,83, 88

Reverse genetics 3, 25

S

Salinity 11, 14, 17, 21, 22, 29, 30, 34, 38, 39,44-49, 75, 105, 127, 130-134, 155, 158,160, 161, 163, 164

Salt tolerance 4, 31-38, 45, 46, 48, 49, 164,166

Seed dormancy 4, 6SFR 75Sulfur metabolism 147, 155, 161, 164, 166

T

Thermotolerance 83-85, 87, 88, 93Tonoplast 13, 35, 40, 44, 46, 105-108,

111-113, 115-117Two component system 157

V

Vicia 102, 103, 109, 112, 116, 117

W

Water channels 39, 42Water stress 5, 20, 88, 101, 102, 105, 106, 161

Documents

Molecular Responses