glutamate in plants - journal of experimental botany

Journal of Experimental Botany, Vol. 58, No. 9, pp. 2339–2358, 2007

Nitrogen Nutrition Special Issue

doi:10.1093/jxb/erm121 Advance Access publication 19 June, 2007

SPECIAL ISSUE PAPER

Glutamate in plants: metabolism, regulation, and signalling

Brian G. Forde* and Peter J. Lea

Lancaster Environment Centre, Department of Biological Sciences, Lancaster University, Lancaster LA1 4YQ, UK

Received 9 March 2007; Revised 30 April 2007; Accepted 2 May 2007

Abstract

Glutamate occupies a central position in amino acid

metabolism in plants. The acidic amino acid is formed

by the action of glutamate synthase, utilizing gluta-

mine and 2-oxoglutarate. However, glutamate is also

the substrate for the synthesis of glutamine from

ammonia, catalysed by glutamine synthetase. The

a-amino group of glutamate may be transferred to

other amino acids by the action of a wide range of

multispecific aminotransferases. In addition, both the

carbon skeleton and a-amino group of glutamate form

the basis for the synthesis of g-aminobutyric acid,

arginine, and proline. Finally, glutamate may be

deaminated by glutamate dehydrogenase to form

ammonia and 2-oxoglutarate. The possibility that the

cellular concentrations of glutamate within the plant

are homeostatically regulated by the combined action

of these pathways is examined. Evidence that the well-

known signalling properties of glutamate in animals

may also extend to the plant kingdom is reviewed. The

existence in plants of glutamate-activated ion channels

and their possible relationship to the GLR gene family

that is homologous to ionotropic glutamate receptors

(iGluRs) in animals are discussed. Glutamate signal-

ling is examined from an evolutionary perspective, and

the roles it might play in plants, both in endogenous

signalling pathways and in determining the capacity of

the root to respond to sources of organic N in the soil,

are considered.

Key words: Enzymes, GLR genes, glutamate, homeostasis,

ionotropic glutamate receptors, metabolism, root architecture,

signalling, synthesis.

Introduction

In their stimulating article ‘Glutamate: an amino acid ofparticular distinction’, Young and Ajami (2000) reflectthat perhaps God’s inordinate fondness may have been forglutamate, and not for beetles as the evolutionary biologist(and atheist) JBS Haldane is famously quoted as suggest-ing. The authors had in mind both the ubiquity of theamino acid in Nature and its remarkably diverse biologicalroles as a metabolite, an energy-yielding substrate, a nutri-ent, a structural determinant in proteins, and even a signal-ling molecule. They noted that glutamate has a number ofchemical properties, including its reactivity, that make itparticularly well suited to its multiplicity of functions.

Many aspects of the importance of glutamate in plantshave been appreciated for decades, but others are onlynow coming to light. The purpose of this focus paper is toreview key aspects of the place of glutamate in plantbiology, focusing on its metabolic roles, the question ofwhether its cellular concentration is homeostaticallyregulated, and the evidence surrounding its possible roleas a signalling molecule in plants.

Glutamate metabolism

As can be seen from Fig. 1, there is little doubt thatglutamate is a central molecule in amino acid metabolismin higher plants. The a-amino group of glutamate isdirectly involved in both the assimilation and dissimilationof ammonia and is transferred to all other amino acids. Inaddition, both the carbon skeleton and a-amino groupform the basis for the synthesis of c-aminobutyric acid(GABA), arginine, and proline. It should also be notedthat glutamate is the precursor for chlorophyll synthesis in

* To whom correspondence should be addressed. E-mail: [email protected]: AMPA, (S)-a-amino-3-hydroxy-5-methylisoxazole-4-propionic acid; BMAA, b-methylamino-L-alanine; CNS, central nervous system; DNQX,6,7-dinitroquinoxaline-2,3-dione; EAAT, excitatory amino acid transporter; Fd, ferredoxin; GABA, c-aminobutyric acid; GDH, glutamate dehydrogenase;GGAT, glutamate:glyoxylate aminotransferase; GLR, plant glutamate receptor gene; GPCR, G-protein-coupled receptor; GS, glutamine synthetase; iGluR,ionic glutamate receptor; LAOBP, lysine/arginine/ornithine-binding protein; LIVBP, leucine/isoleucine/valine-binding protein; MK-801, dizocilpine maleate;mGluR, metabotropic glutamate receptor; NMDA, N-methyl-D-aspartic acid; Pi, inorganic phosphate; PCA, principal component analysis; Rubisco, ribulosebisphosphate carboxylase oxygenase.

ª The Author [2007]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.For Permissions, please e-mail: [email protected]

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developing leaves (Yaronskaya et al., 2006). The bio-chemistry and molecular biology of glutamate metabolismhave been reviewed previously (Lea and Miflin, 2003;Suzuki and Knaff, 2005), and this paper provides anoverview of the enzymes involved in the metabolism ofglutamate as shown in Fig. 1, focusing on advances thathave been made in the past few years.

Glutamine synthetase–glutamate synthase

The key enzyme involved in the de novo synthesis ofglutamate is glutamate synthase, also known as gluta-mine:2-oxoglutarate amidotransferase (GOGAT). The re-action is a reductant-driven transfer of the amide aminogroup of glutamine to 2-oxoglutarate to yield twomolecules of glutamate. The enzyme in plants is presentin two distinct forms, one that uses reduced ferredoxin(Fd) as the electron donor (EC 1.4.7.1) and one that usesNADH as the electron donor (EC 1.4.1.14). The Fd-dependent enzyme is normally present in high activities inthe chloroplasts of photosynthetic tissues, where it is ableto utilize light energy directly as a supply of reductant.The NADH-dependent enzyme, which is also present inplastids, is located predominantly in non-photosynthesiz-ing cells, where reductant is supplied by the pentosephosphate pathway (Bowsher et al., 2007). The Fd- andNADH-dependent forms would appear to be expresseddifferently in separate plant tissues. However, the situationis now more complex, as evidence is accumulating tosuggest that in most plants there are two genes that encodeeach form of glutamate synthase. Two recent reviewshave covered the early history of the discovery of the twoenzymes, their structure, and gene regulation (Lea andMiflin, 2003; Suzuki and Knaff, 2005). In this volume,Tabuchi et al. (2007) and Canovas et al. (2007) discussthe role of glutamate synthase in rice and conifers. Clearevidence of perturbations in amino acid metabolism havebeen demonstrated in plants that have reduced amounts ofeither form of enzyme activity, caused by mutation or

gene knockout (Somerville and Ogren, 1980; Blackwellet al., 1988; Leegood et al., 1995; Ferrario-Mery et al.,2002a, b; Lancien et al., 2002).

The nitrogen-containing substrate for the glutamatesynthase reaction is provided in the form of glutamine.This itself is synthesized in the ATP-dependent combina-tion of glutamate and ammonia catalysed by glutaminesynthetase (GS; EC 6.3.1.2). The ammonia may have beengenerated by direct primary nitrate assimilation, orfrom secondary metabolism such as photorespiration(Leegood et al., 1995). GS activity is located in both thecytoplasm and chloroplasts/plastids in most higher plants,except conifers. The enzyme proteins can be readilyseparated by standard chromatographic localization andwestern blotting techniques into cytoplasmic (GS1) andplastidic (GS2) forms. However, this distinction is not assimple as was first thought, as although only one gene hasbeen shown to encode the plastidic form, a small family ofup to five genes is now known to encode the cytoplasmicform. Recent information on the regulation of the genesencoding GS in conifers (Canovas et al., 2007), maize(Hirel et al., 2007), and rice (Tabuchi et al., 2007) isincluded in this volume. Limited success in the improve-ment of growth rates has been demonstrated following theoverexpression of GS1 genes in Lotus corniculatus,poplar trees, tobacco, and maize (Martin et al., 2006;Tabuchi et al., 2007). Early mutants of barley lackingchloroplastic GS2 exhibited a severe phenotype and wereonly able to grow under non-photorespiratory conditions(Blackwell et al., 1988; Leegood et al., 1995). However,more recently, GS1 knockout lines have been isolated andcharacterized in both rice and maize. The growth rate,spikelet number, and weight were considerably reduced ina homozygous knockout line of OsGS1;1 in rice(Tabuchi et al., 2005). In contrast, maize mutants withknockouts of the gln1-3 and gln1-4 genes appeared togrow normally until grain filling (Martin et al., 2006).However, there was a reduction of kernel size in gln1-4

Glutamine

2-Oxoglutarate

Glutamine

Glutamate

GLUTAMATE 2-OxoglutarateGlutamate

NH3

AsparagineUreidesNucleic acids

Arginine

Carbonmetabolism

NH3

Glutamate Synthase

-Aminobutyrate

Proline

AminoAcids

2-OxoAcids

Amino Transferases

Respiration

Synthetase

Dehydrogenase

Fig. 1. Pathways of glutamate synthesis and metabolism in plants.

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and of kernel number in gln1-3 mutants. In the gln1-3/1-4double mutant, a cumulative effect of the two mutationswas observed (Martin et al., 2006). The data from thesetwo experiments clearly show that individual GS1proteins have a non-redundant role in plant metabolism.

Glutamate dehydrogenase

The two enzymes involved in glutamate synthesis dis-cussed previously catalyse irreversible reactions. A thirdenzyme, glutamate dehydrogenase (GDH; EC 1.4.1.2),catalyses a reversible amination/deamination reaction,which could lead to either the synthesis or the catabolismof glutamate. During the last 33 years, the role of GDH inglutamate metabolism in plants has been the subject ofcontinued controversy (Oaks and Hirel, 1985; Duboiset al., 2003; Terce-Laforgue et al., 2004b). However,following recent investigations into the regulation of thegenes encoding the enzyme protein, the presence ofoverexpressing and antisense lines and the use of nuclearmagnetic resonance (NMR) and gas chromatography–mass spectrometry (GC-MS) techniques, the role isbecoming clearer. Very recently, Masclaux-Daubresseet al. (2006) have reconfirmed that in both young and oldtobacco leaves, glutamate is synthesized via the combinedaction of GS and glutamate synthase, whilst GDH isresponsible for the deamination of glutamate.

GDH is located in the mitochondria, and on occasionthe cytoplasm, within the phloem companion cells ofshoots (Terce-Laforgue et al., 2004a; Fontaine et al.,2006). GDH extracted from most plant species can bereadily separated into seven isoenzymic forms followingnative gel electrophoresis (Thurman et al., 1965; Loulaka-kis and Roubelakis-Angelakis, 1996). The reason for thisis that GDH comprises two distinct subunits (a and b) thatare able to assemble, apparently at random, into enzymat-ically active hexamers. The relative proportion of the a-and b-subunits and hence the isoenzyme pattern observedvaries with plant organ and nitrogen source (Loulakakisand Roubelakis-Angelakis, 1991; Turano et al., 1997).The a-subunit is encoded by GDHA in Nicotianaplumbaginifolia and GDH2 in Arabidopsis, and theb-subunit by GDHB in N. plumbaginifolia and GDH1 inArabidopsis (Purnell et al., 2005).

Antisense and mutant lines of both genes in tobacco andArabidopsis have been constructed, but a full metabolicanalysis of the plants has not yet been carried out(Fontaine et al., 2006). Tobacco lines overexpressing theGDH b-subunit exhibited a greater capacity to catabolizeglutamate but were not able to assimilate ammonia, whenGS was inhibited (Purnell and Botella, 2007). GDHactivity has long been known to increase during senes-cence and following the application of a range of stresses.It has recently been shown that high NaCl induces theformation of reactive oxygen species, which in turninduces the synthesis of the a-subunit of GDH in tobacco

and grapevine. When GS was inhibited, there wasevidence of incorporation of ammonia via GDH into[15N]glutamate and [15N]proline in the presence of highsalt (Skopelitis et al., 2006).

Glutamate as a substrate for aminotransferasereactions

Following the formation of glutamate, the a-amino groupcan be transferred to a wide variety of 2-oxo acidacceptors to form amino acids, and similarly the a-aminogroup can be transferred back to form glutamate when2-oxoglutarate and other amino acids are available. Thereactions are carried out by reversible pyridoxal-5#-phosphate-containing enzymes termed aminotransferases(EC 2.6.1.x), also known as transaminases. It is probablythe reversibility of these enzyme reactions that accountsfor the relative stability of the glutamate concentrationfound in plants, which will be discussed in a later section.Studies on the substrate specificity of aminotransferases inthe past have been confused by the use of impure plantextracts; however, there is now substantial evidence thataminotransferases can exhibit wide preferences for boththe amino acid and oxo acid substrate. Following thepublication of the genome sequence of Arabidopsisthaliana, it was calculated that there were 44 genes thatencoded aminotransferases (Liepman and Olsen, 2004).Such a large number of enzymes is clearly beyond thescope of this article, and for this reason only a fewreviews and key papers will be discussed.

The a-amino group of glutamate may be transferred tooxaloacetate to form aspartate by aspartate aminotransfer-ase (EC 2.6.1.1). Aspartate is a precursor of asparagine(Lea et al., 2007) and the aspartate family of amino acids,lysine, threonine, methionine, and isoleucine (Azevedoet al., 2006). Aspartate aminotransferase also plays a keyrole in C4 photosynthesis (Edwards et al., 2004). InArabidopsis there are five distinct forms of aspartateaminotransferase that are located in the cytosol, mitochon-dria, plastids, and peroxisomes (Wadsworth, 1997;Liepman and Olsen, 2004). Very recently, a plastid-localized aspartate aminotransferase has been identifiedthat is unrelated to other plant forms of the enzyme, but issimilar to a prokaryotic enzyme (De la Torre et al., 2006).

The a-amino group of glutamate may also be trans-ferred to pyruvate to form alanine by the action of alanineaminotransferase (EC 2.6.1.2). Alanine synthesis has beenshown to play a key role in the response to hypoxia/anoxia (Ricoult et al., 2006) and in C4 photosynthesis(Edwards et al., 2004). Four genes encoding alanineaminotransferases have been identified in Arabidopsis(Liepman and Olsen, 2004) and Medicago truncatula(Ricoult et al., 2006), with the enzymes being located inthe cytosol, mitochondria, and peroxisomes. The transferof amino groups to glyoxylate to form glycine in theperoxisomes in the photorespiratory nitrogen cycle (Keys,

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2006; Reumann and Weber, 2006) is an unusual case, asthe reaction is considered to be unidirectional. As well asa serine:glyoxylate aminotransferase, two forms of gluta-mate:glyoxylate aminotransferase (GGAT) are present inthe peroxisome. Recombinant GGT1 was shown to haveglutamate:glyoxylate, alanine:glyoxylate, glutamate:pyru-vate, and alanine:2-oxoglutarate activities (Liepman andOlsen, 2004; Reumann and Weber, 2006). Aminotrans-ferases involved in the metabolism of GABA, proline, andarginine will be discussed in later sections. At least sixaminotransferases are involved in the synthesis andmetabolism of branched chain amino acids (Binder et al.,2007) and two in the formation of histidinol phosphate inthe synthesis of histidine (Mo et al., 2006). Very recently,a totally new aminotransferase capable of convertingtetrahydrodipicolinate to LL-diaminopimelate, bridgingthree enzyme reactions, has been identified in the lysinesynthetic pathway of Arabidopsis (Hudson et al., 2006).

GABA, arginine, and proline

Glutamate may be converted to GABA by the irreversibleaction of glutamate decarboxylase (GAD; EC 4.1.1.15) inthe cytoplasm. GAD was initially shown to have a low pHoptimum of <6.0. However, it is now known that theenzyme protein has a Ca2+/calmodulin-binding site at theC-terminus (Zik et al., 2006), which allows the enzyme tobe stimulated in the presence of Ca2+/calmodulin at pHvalues >7.0. GABA accumulates in higher plants follow-ing the onset of a variety of stresses such as acidification,oxygen deficiency, low temperature, heat shock, mechan-ical stimulation, pathogen attack, and drought (Shelpet al., 1999; Bouche and Fromm, 2004; Bown et al.,2006). Shelp et al. (2006) have presented evidence thatGABA is involved in communication between plants andanimals, fungi, bacteria, and other plants. Very recently,Lancien and Roberts (2006) have also shown that GABAcan down-regulate the expression of genes encoding 14-3-3proteins in a calcium-, ethylene-, and abscisic acid-dependent manner.

Glutamate is the precursor of arginine and is metabo-lized via acetylated derivatives to ornithine, citrulline, andarginosuccinate in a nine-step process (Slocum, 2005).Arginine has a high N:C ratio (4:6), and along withasparagine (2:4) acts as a major nitrogen storage com-pound in higher plants, where it occurs in both the proteinand soluble form. Arginine plays a key metabolic role inseed maturation and germination, phloem and xylemtransport, particularly in conifer trees, and accumulatesunder stress and deficiency conditions (Lea et al., 2007).Arginine and ornithine may also act as precursors ofpolyamines, which can play an important role in theresponse of plants to stress (Alcazar et al., 2006). Earlystudies indicated that the major control point of argininesynthesis was at N-acetylglutamate kinase (NAGK; EC2.7.2.8), the activity of which was inhibited by arginine

and activated by N-acetylglutamate. There is now strongevidence that this control is mediated through the bindingof PII, a 2-oxoglutarate- and amino acid-sensing protein(Lam et al., 2006), which is able to relieve the inhibitionof NAGK by the end-product arginine (Sugiyama et al.,2004; Chen et al., 2006; Ferrario-Mery et al., 2006).

Glutamate can act as the precursor of proline ina pathway that requires three enzyme-catalysed reactionsand a spontaneous chemical reaction. Proline is a cyclicamino acid that has been shown to accumulate followinga wide range of abiotic stresses such as that induced bydrought, salinity, low and high temperatures, and heavymetals (Munns, 2005; Sharma and Dietz, 2006). There isalso evidence that proline is a major component of thenitrogen transport stream in both the xylem and phloem(Brugiere et al., 1999). The bifunctional enzyme proteinD1-pyrroline-5-carboxylate synthetase (P5CS) catalysestwo reactions (EC 2.7.2.11 and EC 1.2.1.41), the firstof which, the phosphorylation of glutamate, is subjectto feedback inhibition by proline (Delauney andVerma, 1993; Strizhov et al., 1997; Hong et al., 2000;Armengaud et al., 2004).

Glutamate concentrations in plant tissues

Methods for identifying and quantifying amino acids haveimproved dramatically over the last 50 years, frompaper chromatography to ion exchange chromatography(Steward and Durzan, 1965), HPLC (Lindroth andMopper, 1979; Geigenberger et al., 1996), NMR (Mes-nard and Ratcliffe, 2005), GC-MS (Roessner et al., 2000),and LC-MS (Jander et al., 2004), with the recent additionof principal component analysis (PCA) (Fritz et al.,2006a). It should be remembered that the plant extractsused will have been derived from a range of intracellularorganelles as well as extracellular material (Lunn, 2007),although the differences in amino acid concentrationsbetween these organelles may not be very large (Rienset al., 1991). In addition, there are known to be >40different cell types in plants, each of which may havea distinct transcriptome, proteome, and metabolome(Ohtsu et al., 2007). A vast number of papers havedetermined the soluble amino acid contents of higherplants under various conditions, including altered nitrogennutrition, developmental changes, and, more recently, inmutants and transgenic lines. It would be impossible tocover even a fraction of the relevant publications, so onlya small number of recent key papers will be discussed.

Light/dark diurnal cycle

In their paper entitled ‘Steps towards an integrated view ofnitrogen metabolism’, Stitt et al. (2002) proposed that‘glutamate plays a pivotal role in a sensitive feedbackmechanism that regulates ammonium assimilation’. As

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supporting evidence for this statement they drew ourattention to the fact that in tobacco leaves, glutamateconcentrations remained remarkably constant between3 lmol g�1 FW and 4 lmol g�1 FW throughout the day,irrespective of the growth conditions, whilst glutamineconcentrations varied considerably between 5 lmol g�1

FW and 15 lmol g�1 FW (Geiger et al., 1998). Similarlyin tobacco plants grown continuously in 2 mM nitrate, theleaf glutamate concentration remained constant around4 lmol g�1 FW, whilst the glutamine concentration variedbetween 5 lmol g�1 FW and 15 lmol g�1 FW during thelight/dark cycle (Matt et al., 2001a). Masclaux-Daubresseet al. (2002) confirmed that there were only smalloscillations in the glutamate content of older source leavesof tobacco during the diurnal cycle (1100–1400 nmolmg�1 chlorophyll), but that in younger sink leaves theglutamate concentration increased during the light periodto 1600 mmol mg�1 chlorophyll and decreased to900 nmol mg�1 chlorophyll in the dark. Similarly, inpotato leaves, glutamate showed very little diurnalvariation (1.0–1.3 lmol g�1 FW), by comparison withalanine, aspartate, glutamine, and glycine (1–3 lmol g�1

FW) (Urbanczyk-Wochniak et al., 2005). In a later studyon Arabidopsis, Gibon et al. (2006) noted that a largeproportion of the 137 metabolites measured in the rosettesexhibited marked diurnal changes during a 12/12 h light/dark cycle. These smooth oscillations in metaboliteconcentration included all of the amino acids, with thenotable exception of glutamate, which showed only verysmall fluctuations.

Fritz et al. (2006a) re-examined data obtained pre-viously that detailed the amino acid concentrations duringa diurnal light/dark cycle in wild-type, nitrate reductase-deficient (Fritz et al., 2006b), and Rubisco-deficient(Matt et al., 2002) lines of tobacco, grown at twoconcentrations of nitrate and varying light intensities. Thedata set was analysed using PCA and regression analysis,and by normalizing the level of each individual aminoacid on the total amino acid pool. Whilst major variationsin some amino acids were detected, the concentration ofglutamate remained relatively constant, with amplitudes ofchange being only 1.39 during the diurnal cycle and 2.34(–) for nitrogen status, compared with 8.29 and 197.85 (–)for glutamine. Fritz et al. (2006a) did however notea reduction in glutamate concentration in Rubisco-deficient plants with an amplitude of change of 11.92 (–),which correlated with lower concentrations of 2-oxogluta-rate, possibly due to high ATP and an increased redoxstate. Novitskaya et al. (2002) highlighted the importanceof 2-oxoglutarate in the synthesis of glutamate, when theyexamined the effect of photosynthesis and photorespira-tion on the amino acid content of wheat and potato leaves.Although there was a major increase in glycine contentfollowing transfer to conditions of high photorespiratoryrates, this was at the expense of aspartate, alanine, and

serine, whilst the glutamate and glutamine concentrationsremained constant. In a mutant where the mitochondrial Icomplex was non-functional and NADH concentrationswere increased, reductions in the concentrations of both2-oxoglutarate and glutamate were detected (Dutilleulet al., 2005).

Developmental stage

The germination of seeds is a period when the storedprotein is rapidly hydrolysed and transported as solubleamino acids to the growing root and shoot. The majorforms of the transported amino acids are usually thosewith a high N:C ratio such as asparagine, glutamine, andarginine (Lea et al., 2007). During the first 12 d ofgermination of loblolly pine, the increases in the concen-tration of asparagine, glutamine, and arginine were 4487-,1587- and 542-fold, respectively, whilst that of glutamatewas only 44-fold (King and Gifford, 1997). During theearly phases of germination (20 h) of Medicago truncatula,the amount of glutamate increased in the cotyledons (180–260 nmol per organ), but then remained remarkablyconstant during post-germinative growth, whilst the totalamino acids increased from 1500 nmol to 5500 nmol perorgan; similar data were also obtained with the embryoaxis. Glevarec et al. (2004) proposed that GDH in theembryo axis of M. truncatula was involved in maintainingthe constant glutamate concentration.

The senescence of leaves is another period whenproteins are hydrolysed and amino acids are transported,again often as the amides and arginine (Hortensteiner andFeller, 2002; Lea et al., 2007). Masclaux et al. (2000)analysed a number of metabolites in a wide range ofdifferent aged leaves on a mature vegetative tobacco plant.The total amino acid content was highest in the youngestleaves, but was reduced by 70% in the oldest leaves. Theproportion of glutamate of the total (25%) remainedconstant, whilst that of GABA increased and that ofproline decreased. Diaz et al. (2005) analysed the solubleamino acids in the leaves of different recombinant inbredlines of Arabidopsis during senescence. For all the lines,glutamate increased just after the onset of senescence from20% to 40% of the total amino acid pool, but thendecreased to between 10% and 20%, the rate of decreaseof glutamate varying between the lines but correlatingwith senescence. In darkened Arabidopsis leaves, theasparagine concentration increased 20-fold over 6 d from1 nmol mg�1 FW to 10 nmol mg�1 FW, whilst glutamateremained constant at 2 nmol mg�1 FW (Lin and Wu,2004). Some of the largest increases in glutamateconcentration have been demonstrated during the ripeningof tomato fruits (Gallardo et al., 1993; Baxter et al.,2005). In the leaves of tomato, the glutamate concentra-tion was 2.32 lmol g�1 FW, whilst in green fruit it was40.07 lmol g�1 FW, rising to 282.1 lmol g�1 FW in redfruit (Roessner-Tunali et al., 2003). Carrari et al. (2006)


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speculated that as glutamate was a precursor of chloro-phyll, this accumulation towards the end of ripening couldbe due to the cessation of chlorophyll biosynthesis

Nitrogen nutrition

The availability and uptake of nitrogen is considered asthe major factor affecting growth (Lea and Azevedo,2006). Increased nitrate has a beneficial effect andstimulates the synthesis of amino acids and protein, whilstexcess ammonium ions can be toxic and can promote theformation of amides (Britto and Kronzucker, 2002).

Geiger et al. (1999) grew tobacco at three concentra-tions of ammonium nitrate and three of potassium nitrateat normal and elevated CO2. Small increases in glutamateconcentration in the leaves were determined at highernitrogen inputs, which were further enhanced at elevatedCO2. Far greater increases in alanine, arginine, aspartate,glutamine, and serine were detected. However, at 3 mMammonium nitrate, there was a 4-fold increase inglutamate in the leaves that was not enhanced by elevatedCO2. In the roots, increased ammonium nitrate had noeffect, whilst 6 mM and 20 mM potassium nitrate causeda 2-fold increase in glutamate. Scheible et al. (2000)compared the soluble amino concentrations in two nitratereductase-deficient lines of tobacco, with wild-type plantsgrown on high and low nitrate. Remarkably, at noon, atthe interface of the 12/12 h light/dark day cycle, theglutamate concentrations were the same in the leaves ofall four plants at 3 lmol g�1 FW, whilst the glutamineconcentration ranged from zero to 17 lmol g�1 FW.During the day there was a small variation in theglutamate concentration, and overall the low nitrogenplants ranged from 2 lmol g�1 FW to 3 lmol g�1 FWand the high nitrogen plants from 3 lmol g�1 FW to4 lmol g�1 FW. Matt et al. (2001b) also detected littledifference in glutamate concentrations, ranging between4 lmol g�1 FW and 5 lmol g�1 FW in tobacco plantsgrown in 2 mM potassium nitrate or 1 mM ammoniumnitrate at either normal or elevated CO2 throughout theday in source leaves, whilst glutamine ranged from5 lmol g�1 FW to 15 lmol g�1 FW and glycine from1 lmol g�1 FW to 8 lmol g�1 FW

Terce-Laforgue et al. (2004b) grew tobacco under threeconditions of nitrogen fertilization, nitrate, ammonium,and low nitrogen, for 6 weeks and examined the aminoacid content of leaves of different developmental stages.In the low-nitrogen-grown plants, the glutamate and totalamino acids increased <4-fold from the old to youngleaves (from 6.3 nmol mg�1 DW to 22.5 nmol mg�1

DW). In the nitrate-grown plants, the total amino acidsincreased 10-fold from the old to young leaves, whilst theglutamate concentration only increased 5-fold from3.0 nmol mg�1 DW to 15.2 nmol mg�1 DW, but overallthe values were similar to the low nitrogen concentrations.In the older leaves of ammonium-grown plants, whilst the

total amino acids were >30-fold higher than in the othertwo treatments, the glutamate concentration was verysimilar, ranging from 7.3 nmol mg�1 DW to 8.1 nmolmg�1 DW, and with little age-dependent change.

In a series of detailed metabolite profiling studies onhydroponically grown tomato, Urbanczyk-Wochniak andFernie (2005) showed that glutamate declined to 25%following nitrate starvation (0.4 mM) for 7 d at high lightintensity, with a lesser decrease in low light. The additionof 8 mM nitrate caused a small increase in glutamateirrespective of light. Overall the changes in glutamatewere less than those observed for the other amino acids.

Fritz et al. (2006a, b) grew wild-type tobacco on12 mM nitrate and compared the leaf amino acid contentwith plants grown on 0.2 mM nitrate and a nitratereductase-deficient mutant grown on 12 mM nitrate. Theconcentrations of alanine, asparagine, aspartate, gluta-mine, glycine, and serine in the nitrate-replete wild typeall showed considerable diurnal variation which reachedvalues of 10–20-fold higher than the two nitrogen-deficient lines. However, the glutamate concentration inthe 12 mM nitrate-grown wild type never exceeded twicethat of the two deficient lines and showed no diurnalvariation. In tobacco plants transformed with a nitratereductase enzyme that was not subject to either transcrip-tional or post-translational regulation, there were increasesin glutamine and asparagine by factors of 9 and 14,respectively, when compared with the wild type, whichwere maintained during both the light and dark period.However, the glutamate concentration in the deregulatedplant was only greater by a maximum of 2-fold comparedwith the wild type (32 lmol g�1 DW to 16 lmol g�1

DW) during the light period and was not different duringthe dark period (Lea et al., 2006).

Nitrate-deficient Arabidopsis seedlings grown in sterileculture contained 2–3-fold higher 2-oxoglutarate(250 nmol g�1 FW), 50-fold lower glutamine (0.25 lmolg�1 FW), and 6-fold lower glutamate concentrations(1.0 lmol g�1 FW) compared with seedlings grown infull nutrient medium. Three hours after the addition of3 mM nitrate, glutamine and glutamate increased by6-fold and 2-fold, respectively, whilst 2-oxoglutaratedecreased by approximately 40% (Scheible et al., 2004).In a similar series of experiments involving carbonstarvation of Arabidopsis seedlings, followed by theresupply of 15 mM sucrose, Osuna et al. (2007) demon-strated, using a false colour chart, that whilst alanine,arginine, aspartate, glycine, and serine exhibited majorchanges, glutamate remained constant.

Feeding of metabolites

One way of investigating how glutamate concentrationscan vary is to feed metabolites into plant tissues from theoutside. The feeding of high concentrations, 150 mMGABA and proline, or 100 mM glutamine and nitrate, to

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leaf discs of tobacco had little effect on the endogenousglutamate concentration, which remained around 30 nmolmg�1 DW. Only the feeding of 100 mM glutamate causedan increase of 3–4-fold in glutamate, whilst glutamineincreased up to 30-fold to 302 nmol mg�1 DW, and at thesame time there was an induction of GS1 and GDH(Masclaux-Daubresse et al., 2005). The feeding of 20 mMglutamate through the petioles to tobacco leaves causeda small increase in the glutamate concentration from2.6 lmol mg�1 FW to 3.7 lmol mg�1 FW and there wasa decrease in nitrate reductase activity, whilst 40 mM 2-oxoglutarate caused a 5-fold increase in glutamate from2.0 lmol g�1 FW to 9.8 lmol g�1 FW (Fritz et al.,2006a). Similar results obtained by a different researchgroup showed that the feeding of 40 mM glutamate totobacco had little effect on the glutamate pool, although2-oxoglutarate increased 6–9-fold; in contrast, the feedingof 40 mM 2-oxoglutarate increased glutamate by >3-foldfrom 2 lmol g�1 FW to 7 lmol g�1 FW (Schneidereitet al., 2006). Thus it would appear that whilst even thesupply of very high concentrations of amino acids has littleeffect on the glutamate concentrations in leaves, it ispossible to manipulate the glutamate pools by feeding2-oxoglutarate, indicating the important role that the supplyof 2-oxoglutarate, probably through the action of isocitratedehydrogenase, plays in glutamate synthesis (Hodges,2002; Abiko et al., 2005).

Use of inhibitors, mutants, and antisense/knockout lines

The pathways involved in glutamate metabolism in vivocan be identified by preventing the action of key enzymes.Early work on establishing fluctuations in the amino acidpools of plants was carried out using inhibitors of GS(Fentem et al., 1982; Rhodes et al., 1986), glutamatesynthase (Baron et al., 1994), and aminotransferases(Brunk and Rhodes, 1988). However, doubt was alwayscast on the specificity of such inhibitors and the possibilityraised that other enzymes may have been inadvertentlyaffected. The first isolation of a mutant of Arabidopsisdeficient in Fd-glutamate synthase (Somerville and Ogren,1980) led the way to the characterization of a range ofmutants which were impaired in the photorespiratorynitrogen cycle (Keys, 2006). The mutants of Arabidopsisand barley deficient in Fd-glutamate synthase were charac-terized by an accumulation of glutamine and a reduction inthe concentration of glutamate following transfer fromelevated CO2 to normal air (Blackwell et al., 1988;Leegood et al., 1995). Leaves of mutants of barley deficientin chloroplastic GS accumulated high concentrations ofammonia, whilst glutamate decreased from 3.5 lmol g�1

FW to 0.5 lmol g�1 FW in 2 h following transfer fromelevated CO2 to normal air (Blackwell et al., 1988).

When antisense lines of tobacco containing only 15%Fd-glutamate synthase activity were exposed to air, therewas a 20-fold increase in glutamine, ammonia, and

2-oxoglutarate, whilst the glutamate concentration onlyfell by a maximum of 40% to 10 nmol mg�1 DW towardsthe end of the light period (Ferrario-Mery et al., 2002a,b). The leaves of antisense lines of tobacco deficient ina chloroplast 2-oxoglutarate/malate translocator, whenexposed to air in the light contained considerably reducedglutamate, aspartate, serine, and proline, and elevatedglutamine and 2-oxoglutarate concentrations (Schneidereitet al., 2006). Interestingly, the glutamate concentration inNADH-glutamate synthase knockout lines of Arabidopsiswas unaffected when the plants were grown in air, butwas reduced from 3 pmol mg�1 FW to <1 pmol mg�1

FW in plants grown in high CO2 (Lancien et al., 2002).The glutamate content of the roots and leaves of rice is

low when compared with glutamine and asparagine, andonly small increases were detected in a homozygousOsGS1;1 knockout line deficient in GS1 (Tabuchi et al.,2005). In maize knockout lines of the gln1-3 and gln1-4GS1 genes, the concentration of glutamate was increasedin the mature leaves (where no phenotype was observed),but was slightly reduced in the phloem (79 lmol ll�1 to56 lmol ll�1) and xylem (5.6 lmol ll�1 to 4.6 lmolll�1) (Martin et al., 2006). In GGAT knockout lines ofArabidopsis, glutamate increased from 3.8 nmol mg�1

FW to a maximum of 5.8 nmol mg�1 FW, with similarincreases in glutamine and aspartate being detected.However, in GGAT-overexpressing lines, there was littlechange in glutamate, glutamine, and aspartate, but serineincreased from 1 nmol mg�1 FW to 20 nmol mg�1 FWand glycine from 0.2 nmol mg�1 FW to 2 nmol mg�1 FW(Igarashi et al., 2006). The glutamate concentration wasunaffected in GDH antisense lines of tobacco (Purnellet al., 2005) and GAD insertion lines of Arabidopsis(Bouche et al., 2004).

Is there glutamate homeostasis in plants?

The experiments described above, whilst admittedlyselective, do provide some evidence that the glutamateconcentration of plant leaves is maintained relativelyconstant during a diurnal cycle. Considering the flux ofnitrogen through the photorespiratory pathway, this is nomean feat (Reumann and Weber, 2006). At differentstages of development, there can be a huge variation insoluble amino acid content, but glutamate changes lessthan the other amino acids, in particular glutamine.Although the glutamate concentration can change inresponse to major alterations in nitrogen nutrition or whencertain key enzymes have been deactivated, this variationis again not normally as great as that measured for otheramino acids. Amino acids when supplied externally arerapidly metabolized, with only 2-oxoglutarate causinga significant accumulation of glutamate.

The next question is how are the glutamate concen-trations held within reasonable limits, whilst concurrentlyother amino acids can change dramatically? The answer


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may well lie in the large number of different enzymes thatcan use glutamate as a substrate or form it as a product. Aclose scrutiny of Fig. 1 shows that the two major routes ofsynthesis of glutamate are from glutamine via glutamatesynthase or from amino acids via aminotransferases. Boththese reactions require a supply of 2-oxoglutarate ifglutamate is to accumulate. If there is a supply of other2-oxo acids, then a wide range of amino acids can beformed by transamination from glutamate. In the absenceof 2-oxo acids, glutamate may be metabolized toglutamine, arginine, or proline, which may accumulate,often as the result of an external stress. A shortage of2-oxoglutarate will also lead to the deamination ofglutamate by GDH. Masclaux-Daubresse et al. (2006)argued that GS/glutamate synthase and GDH were themajor enzymes responsible for maintaining a constantconcentration of glutamate. However, it is also likely thatthe supply of 2-oxoglutarate is a key regulator. The workof Hodges and colleagues has suggested that isocitratedehydrogenase is the primary enzyme required for thesynthesis of this key intermediate (Hodges, 2002; Abikoet al., 2005; Lemaitre and Hodges, 2006).

Glutamate as a phylogenetically conserved signalmolecule

L-Glutamate is now well established as an importantsignalling molecule in the mammalian central nervoussystem (CNS), but for many years this was a matter ofcontroversy. Speculation about a possible signalling rolefor L-glutamate began in the early 1950s, but it was notuntil well into the 1970s that its true status as the majorexcitatory neurotransmitter in the CNS became widelyaccepted (Watkins and Jane, 2006). Central to this storywas the development by Watkins and colleagues ofspecific L-glutamate agonists and antagonists that couldbe used to identify and characterize the neuronalglutamate receptors. Indeed the broad classification ofmammalian ionotropic glutamate receptors (iGluRs) intoN-methyl-D-aspartate (NMDA) receptors and non-NMDA[(S)-a-amino-3-hydroxy-5-methylisoxazole-4-propionic acid(AMPA)-kainate and Delta] receptors, according to theresponses evoked by selective agonists, is still in use(Mayer, 2005).

The iGluRs are glutamate-gated ion channels that areinvolved in the movement of Na+ and Ca2+ across thepost-synaptic plasma membrane. The existence of a secondclass of glutamate receptor in the CNS, whose activity wascoupled to small GTP-binding proteins, was recognized inthe 1980s. These were termed metabotropic glutamatereceptors (mGluRs) to distinguish them from the iGluRs.The mGluRs in general seem to play a modulatory role insynaptic activity and to be involved in effects that areslower and longer lasting than those mediated by iGluRs

(Watkins, 2000). The mGluRs belong to Family C of theseven transmembrane family of G-protein-coupled recep-tors (GPCRs), along with the GABAB receptors.

In the past decade, evidence has unexpectedly emergedfrom different directions that glutamate signalling mayalso occur in higher plants (reviewed in Chiu et al., 1999;Davenport, 2002; White et al., 2002; Filleur et al., 2005;Lam et al., 2006). In the following sections, recentprogress towards establishing the role and the molecularbasis for glutamate signalling in plants is discussed.

Molecular evidence: existence of a family of iGluR-likegenes in plants

The first important clue to the occurrence of glutamatesignalling in plants was the discovery in 1998 thatArabidopsis possesses a family of AtGLR genes homolo-gous to mammalian iGluRs (Lam et al., 1998). Comple-tion of the Arabidopsis genome sequence revealed thatthere are 20 AtGLR genes (compared with only 11 inhumans), which can be grouped into three clades (Chiuet al., 2002). The regions of homology between the plantGLR sequences and their animal counterparts span all theimportant domains, including the four transmembranesegments (M1–M4) and the S1 and S2 regions that formthe agonist-binding domain (Chiu et al., 1999, 2002;Davenport, 2002) (Fig. 2). Phylogenetic analysis suggeststhat the divergence of animal iGluRs and the AtGLRsoccurred before the divergence of the different animaliGluR classes (NMDA, AMPA-kainate, and Delta)(Chiu et al., 1999). The majority of the AtGLR proteinsare predicted to be targeted to the secretory pathway(Davenport, 2002), and a recent report has confirmed thata radish GLR homologue is localized to the plasmamembrane (Kang et al., 2006).

It has been established that all 20 AtGLR genes aretranscribed, and in some cases there is evidence formRNA splice variants, adding to the total number ofpotential AtGLR gene products (Chiu et al., 2002). Studiesof the expression patterns of the AtGLR family haveshown that their transcripts are widely distributed through-out the plant and that all 20 are expressed in roots (Chiuet al., 2002). Clade II genes appear to show a preferencefor root expression, with five of the nine genes in thisclade being expressed specifically in the roots of 8-week-old plants (Chiu et al., 2002).

Despite the overall similarities in their domain organi-zation, the plant GLRs are sufficiently divergent insequence from the iGluRs that it is extremely difficult todeduce either their channel activity or their agonist-binding properties (Lacombe et al., 2001). As has beennoted (Davenport, 2002), the plant GLRs have an unusualsequence in their putative pore region, suggesting that ifthey do have channel activity it either has novel ionselectivity or a novel mechanism of ion selectivity isinvolved. Attempts to model the agonist-binding sites of

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the AtGLRs led to the suggestion that only AtGLR1.1would bind glutamate and that glycine would be thenatural ligand for the other members of the gene family(Dubos et al., 2003). Experimental support for thishypothesis came from evidence that glycine was able toelicit transient increases in [Ca2+]cyt in Arabidopsis seed-lings in a manner similar to glutamate (Dubos et al.,2003). However, as discussed below, the relationshipbetween these Ca2+ spikes and the AtGLRs has not yetbeen unequivocally established. Furthermore, there is theadditional possibility that glycine is not directly activatingCa2+ channels but is having an indirect effect bytriggering the efflux of glutamate (or a glutamate-likeligand) from the plant cells (Frauli et al., 2006; Qi et al.,2006; see also below).

Mammalian and plant members of the iGluR/GLRfamily have two domains that are homologous to bacterialperiplasmic binding proteins: one acts as the agonist-binding site and belongs to the lysine/arginine/ornithine-binding protein (LAOBP)-like superfamily, while theother is in the N-terminal proximal region of the extendedN-terminal domain (NTD) and belongs to the leucine/isoleucine/valine-binding protein (LIVBP)-like superfam-ily (Fig. 2). The LIVBP domain is not found in a pro-karyotic member of the iGluR family, the GluR0glutamate-gated K+ channel, nor in the kainate-binding

iGluRs from lower vertebrates (Oswald, 2004), but isrelated to the LIVBP domain that serves as the agonist-binding domain in members of Family C of the GPCRs(which includes mGluRs, GABAB receptors, and Ca2+

receptors). Functions ascribed to the LIVBP domain iniGluRs include roles in binding allosteric modulators(such as Zn2+, redox agents, and polyamines) and infacilitating the oligomeric assembly of subunits (Oswald,2004). The LIVBP domain in the plant GLRs isparticularly intriguing because it is more closely relatedto the LIVBP domain in the Family C GPCRs than to theLIVBP domain in mammalian GLRs (Turano et al., 2001;Acher and Bertrand, 2004). To explain this, it has beenproposed that, early in the evolutionary history of theiGluR/GLR family and prior to the divergence of plantsand animals, a recombination event occurred that fusedthe LIVBP domain of the ancestral iGluR/GLR to theseven transmembrane domains of a GPCR to create theancestral Family C GPCR (Turano et al., 2001).

Crystal structure analysis of the LIVBP domain of ratmGluR1 has identified a number of key residues (Ser165,Thr188, Asp208, Tyr236, and Asp318) that participate inthe binding of the a-carboxylic and a-amino groups ofglutamate (Kunishima et al., 2000). It has been noted thatthese five residues are conserved in the N-terminal regionof AtGLR3.5 (GLR6) (Taniura et al., 2006), and

Fig. 2. Schematic representation of the 3D structures of eukaryotic glutamate receptors and the bacterial proteins with which they share homologousdomains. Two types of Venus fly trap (VFT) domain occur in members of the iGluR/GLR family: the agonist-binding LAOBP domain (formed bythe S1 and S2 regions of the primary sequence) and the LIVBP domain in the extended N-terminal region, which can act as an allosteric modulatorydomain. The bacterial glutamate receptor (GluR0) lacks the C-terminal and LIVBP domains. M1–M4 refer to the transmembrane domains in theGluR0/iGluR/GLR family (modified from Acher and Bertrand, 2004).


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alignment of the LIVBP domains of all 20 AtGLRsconfirms that AtGLR3.5 is the only member of the familyin which this is the case (BG Forde unpublished data). Thisindicates that glutamate (or a structurally related ligand)may have a role in regulating the function of AtGLR3.5(and of any heteromeric complexes of which AtGLR3.5is a component). It is noteworthy that the five glutamate-binding residues are also conserved in the LIVBP domainof two rice GLR genes belonging to clade III, Os06g46670and Os04g49570 (OsGLR3.1). This suggests that theligand-binding properties of this domain are conservedbetween monocots and dicots, and that glutamate poten-tially also has an important role in modulating theactivity of these members of the rice GLR family.

Cytological and electrophysiological evidence for iGluR-like channel activity

While mammalian iGluRs have been characterized usingheterologous expression systems such as Xenopusoocytes, attempts to do the same with plant GLR geneshave so far been unsuccessful (Li et al., 2006; Lai-HuaLiu, personal communication). It is possible that the GLRsubunits only function as heteromers, as is the case withNMDA receptors (Davenport, 2002), or there might bea requirement for specific interactions with auxiliaryproteins, such as the transmembrane AMPA receptorregulatory proteins (TARPs) that modify the kinetics ofAMPA receptors (Mayer, 2005).

The lack of success with heterologous expressionsystems means that it has not yet been possible toestablish unequivocally the ligand-binding properties orthe ion channel specificity of any plant GLR. Neverthe-less, there is evidence for the existence of glutamate-gatedion channel activity in plants that has properties consistentwith those of iGluR-related proteins. Dennison andSpalding (2000) were able to show that a rapid membranedepolarization which is triggered in root tip cells byexternal glutamate is accompanied by a spike in cytosolicCa2+. Current evidence suggests that these glutamate-activated channels have similarities to excitatory aminoacid-activated channels in animal cells, being voltageinsensitive and permeable to cations, notably Na+ andCa2+ (Dubos et al., 2003; Demidchik et al., 2004).

Until recently, the only evidence tentatively linkingthese cytological and physiological observations to theGLR gene products was pharmacological, based on theuse of agonists and antagonists of mammalian iGluRs (seebelow). However, two recent papers have providedtantalizing evidence that at least some GLRs might havefunctional properties similar to their animal homologues(Kang et al., 2006; Qi et al., 2006). In one study, a radishGLR cDNA (RsGluR) was overexpressed in transgenicArabidopsis seedlings (Kang et al., 2006). Glutamatetreatment of the root cells of the transgenic lines was

found to trigger enhanced Ca2+ influx compared with thecontrols, as expected if RsGluR is a glutamate-gated Ca2+

channel. However, since it remains possible that theectopic expression of the RsGluR gene has somehowenhanced the activity of an endogenous glutamate-activatedchannel, these results fall short of providing formal proofthat RsGluR encodes a glutamate-gated Ca2+ channel.

In a second paper, two T-DNA insertion mutants in theAtGLR3.3 gene were identified and were shown to bedefective in both the fast electrical response to glutamateand the associated Ca2+ response (Qi et al., 2006). Whilethese results clearly link the AtGLR3.3 gene to theseresponses, there are some puzzling aspects to the mutantphenotype. First, since AtGLR3.3 is just one of 20 AtGLRgenes expressed in roots (Chiu et al., 2002), it is not clearwhy the phenotype of the mutants was so strong (e.g. theCa2+ spike triggered by 1 mM glutamate was completelyabsent). Secondly, all 20 proteinogenic amino acids weretested and five were indistinguishable from glutamate intheir effects on both the mutants and the wild type. Thefive that behaved like glutamate were glycine, alanine,asparagine, cysteine, and serine, a structurally diversegroup that, apart from glutamate and glycine, are notknown agonists of iGluRs. It is difficult to envisage howall five amino acids could be agonists of AtGLR3.3. Assuggested by the authors, a possible explanation is thattreating the roots with any of the six effective amino acidstriggered the efflux of the true AtGLR3.3 ligand, so thatwhat was observed was a secondary response. Signifi-cantly, a similar conclusion was reached in a recent studyof the effect of all classical L-amino acids on the eightdifferent mGluR receptors expressed in human embryonickidney 293 (HEK293) cells (Frauli et al., 2006). Initialobservations had indicated that some mGluR receptorsubtypes in these cells could be activated by cysteine,aspartate, and asparagine, as well as glutamate. Althoughthese effects could each be blocked by specific mGluRantagonists, the authors were able to show that glutamatewas the only true agonist and that the activating effects ofthe other three amino acids resulted from their ability totrigger the release of endogenous glutamate from the cells(Frauli et al., 2006). These results underline the difficul-ties inherent in basing the identification of agonists ofglutamate-activated channels solely on the results of invivo experiments where the response may entail un-suspected levels of complexity.

Pharmacological evidence for iGluR-like channels inplants

Several well-known agonists and antagonists of mamma-lian iGluRs have been used as tools to obtain supportingevidence for the existence of functional iGluR-likeproteins in plants. b-Methylamino-L-alanine (BMAA),a cycad-derived agonist of mammalian iGluRs, was

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shown to inhibit Arabidopsis root growth and cotyledonopening, and to stimulate elongation of light-grownhypocotyls, in a glutamate- (and glutamine-) reversiblemanner (Brenner et al., 2000). The antagonist mostcommonly used is 6,7-dinitroquinoxaline-2,3-dione(DNQX), a specific antagonist of mammalian AMPA-and kainate-type iGluRs. A role for plant iGluR-likeproteins in light signalling was surmised from experimentsshowing that DNQX inhibited light-induced hypocotylshortening and the reduction of chlorophyll accumulation(Lam et al., 1998). In other studies, DNQX was able toblock glutamate- (and glycine-) mediated increases incytosolic [Ca2+] in cotyledons (Dubos et al., 2003).However, DNQX was ineffective at inhibiting glutamate-elicited changes in cytosolic [Ca2+] in root tips, suggestingthat glutamate-gated channels in root tips are insensitiveto this compound (Dubos et al., 2003). It is important tonote that DNQX and other antagonists that have beenused in plants, such as 2-amino-5-phosphonopentanoate(Sivaguru et al., 2003), were developed and characterizedfor use with mammalian rather than plant iGluRs. Giventhe major structural differences that appear to existbetween mammalian and plant GLR proteins, as alreadynoted, it is necessary to interpret the effects of thesecompounds with caution until their physiological targetsin plant cells have been confirmed.

Forward and reverse genetics approaches toelucidating the function of GLR genes

A number of studies have adopted a reverse geneticsapproach to investigate the role of specific AtGLR genes.Phenotypic analysis of antisense AtGLR1.1 lines hasimplicated this gene in the regulation of C/N metabolism(Kang and Turano, 2003) and in the modulation ofabscisic acid biosynthesis and signalling (Kang andTurano, 2003; Kang et al., 2004). The phenotype oftransgenic plants overexpressing AtGLR3.2 (formerlyAtGluR2) suggested a role in Ca2+ allocation within theplant (Kim et al., 2001). Microarray analysis of Arabi-dopsis lines overexpressing a radish GLR gene (RsGluR)revealed the up-regulation of jasmonic acid-responsivegenes and jasmonic acid-biosynthetic genes (Kang et al.,2006). The transgenic lines also had enhanced resistanceto a fungal pathogen, suggesting a possible connectionbetween GLR function and plant defence responses.

The only GLR gene so far to emerge from a forwardgenetics screen was recently identified in rice by Li et al.(2006). In a collection of T-DNA insertion lines theyfound a short root mutant that was subsequently shown tohave an insertion in the OsGLR3.1 gene. The OsGLR3.1knockout was associated with disruption of root capdevelopment, a reduction in the size of the quiescentcentre, a decline in radial expansion, cessation of rootmeristematic activity, and, ultimately, cell death in the

root apex (Li et al., 2006). The authors suggested thatOsGLR3.1 may maintain and co-ordinate the normalfunction of the root apical meristem, drawing parallelswith the way in which iGluRs have been implicated inregulating the balance between cell proliferation and celldeath in early CNS development (Hardingham andBading, 2003).

Are there other candidates for the role ofglutamate sensor in plants?

Based on current data, and in the absence of homologuesof the mGluR family in plants, members of the GLRfamily must currently be considered the strongest candi-dates for the role of glutamate receptors. However, aminoacid-sensing mechanisms occur in many forms (Conigraveet al., 2000; Hyde et al., 2003; Kimball and Jefferson,2005), and one should keep an open mind to thepossibility that there are glutamate-sensing systems ofother kinds. In particular, amino acid sensing is a propertyassociated with some amino acid transport proteins, whichin many cases are well placed to act as receptors by virtueof their location in the plasma membrane. In yeast, forexample, the Ssy1p protein is a member of the AAP(amino acid permease family) with a role in sensingextracellular amino acid availability (Iraqui et al., 1999).Ssy1p has been reported to have the interesting propertyof enabling the yeast cell to respond to changes in therelative concentrations of intra- and extracellular aminoacids (Wu et al., 2006).

Members of the excitatory amino acid transporter(EAAT) family in animals function as glutamate-gatedanion channels, an activity that is distinct from theirability to transport glutamate (Fairman et al., 1995). Theinteraction of glutamate with the EAAT1 transporterappears to be involved in regulating the morphology ofglial cells, as well as in up-regulating glutamate transportitself (Hyde et al., 2003). Although homologues of theEAAT1 transporters appear to be absent in plants,Arabidopsis has at least 50 amino acid transporters ofvarious types, most of which are largely uncharacterized(Liu and Bush, 2006).

What is glutamate signalling doing in plants?

An evolutionary perspective

The existence of a large family of plant GLR genes,together with the presence of glutamate-activated channelactivity in roots and shoots, strongly suggests thatglutamate signalling in plants is a reality. When glutamatesignalling was thought of as something strictly associatedwith neurons and nervous systems, the notion of itsexistence in plants seemed paradoxical (Lam et al.,


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1998). However, in the past decade, it has emerged thatglutamate signalling in animals is not restricted to thenervous system, and indeed that it also occurs in primitivemetazoans that lack a nervous system.

Examples where functional glutamate receptors(mGluRs and iGluRs) have been detected in non-neuronalmammalian tissues include bone cells, the pineal gland,and the pancreas (Gill and Pulido, 2001; Hinoi et al.,2004; Moriyama and Yamamoto, 2004). The same tissuesalso express glutamate/aspartate transporters (GLASTs),which are required in synapses for signal termination,and vesicular glutamate transporters (VGLUTs), whichare responsible for the active transport and storage ofL-glutamate in synaptic vesicles. These findings have ledto the conclusion that glutamatergic signalling is a generaland ubiquitous system for intercellular communication inanimals (Hinoi et al., 2004; Moriyama and Yamamoto,2004). Amongst the diverse functions ascribed to gluta-mate signalling in non-neuronal cells are included theregulation of insulin secretion (Storto et al., 2006), thecontrol of cellular differentiation in osteoblasts (Hinoiet al., 2003), and the modulation of tumour cell pro-liferation (Kalariti et al., 2005).

That glutamate signalling has early evolutionary origins,before the divergence of plants and animals, was an ideafirst proposed by Chiu et al. (1999). Support for this ideahas come from studies on the marine sponge Geodiacydonium and the cellular slime mould Dictyostelium.Isolated Geodia cells were shown to be responsive toglutamate and to agonists and anatagonists of mGluRs,and a gene related to the mGluR/GABAB-type receptorswas identified (Perovic et al., 1999). Marine sponges areregarded as belonging to the most primitive metazoanphylum (Muller, 2001), indicating that sophisticatedglutamate signalling systems existed in the earliestmetazoans. The slime mould Dictyostelium is an evenmore primitive unicellular eukaryote that appears to havesplit from the animal–fungal lineage after the divergenceof plants and animals (Eichinger et al., 2005). Theformation of spores on the fruiting body of this socialamoeba is induced by a GABA signal, and glutamate actsas a competitive inhibitor of this process (Anjard andLoomis, 2006). Dictyostelium possesses a mGluR-likegene (DdmGluPR or GrlE) (Taniura et al., 2006), anddisrupting this gene abolishes the responses to GABA andglutamate, suggesting that both amino acids can bind tothe DdmGluPR receptor (Anjard and Loomis, 2006).

Homologues of iGluRs/GLRs even exist in cyanobac-teria (GluR0) where they function as glutamate-gated K+

channels (Chen et al., 1999). Although these receptorslack the extended C-terminal and N-terminal domainsthat characterize the eukaryotic members of the family(Fig. 2), it is likely that iGluRs suitable for intercellularcommunication evolved from an ancestral GluR0-typechannel (Oswald, 2004).

In summary, it appears that both iGluR/GLR-type andmGluR/GABAB-type receptors were already present in theprogenitors of the Viridiplantae and the Metazoa. Sincemulticellularity is thought to have arisen independentlyin the evolution of the major eukaryotic kingdoms(Viridiplantae, Fungi, and Metazoa) (Schopf, 1993), itseems that glutamate/GABA signalling mechanisms mayhave been a feature of the most primitive unicellular lifeforms. It is possible that the earliest function of glutamate/GABA signalling in single-celled organisms was intriggering chemotactic responses. Chemotaxis is recog-nized as an important adaptive mechanism which enhan-ces an organism’s ability to exploit nutrient patches(Fenchel, 2002), and glutamate has been identified asa chemotactic signal in a number of modern dayunicellular organisms, both prokaryotic (Brown and Berg,1974; Barbour et al., 1991) and eukaryotic (Lee et al.,1999; Van Houten et al., 2000). Glutamate also acts asa cue that initiates a range of feeding-related responses ina variety of marine metazoans (Bellis et al., 1991; Trottet al., 1997; Daniel et al., 2001; Kidawa, 2005). In someinstances there is preliminary evidence suggesting thatchemodetection of environmental glutamate may involveiGluR-type receptors (Bellis et al., 1991; Murphy andHadfield, 1997; Van Houten et al., 2000). In this context,it is intriguing that glutamate, acting through iGluR-typereceptors, has a role as a guidance cue for neuronalmigration during embryogenesis (Behar et al., 1999;Manent et al., 2005; Matsugami et al., 2006; McGowan,2006), perhaps echoing an evolutionarily ancient role asa chemoattractant.

If it is the case that glutamate signalling evolved ina primitive progenitor of plants and animals, clues to thefunction of glutamate signalling pathways in plants maycome from a greater understanding of non-neuronalglutamate signalling in animals, and vice versa.

Glutamate signalling and root apical meristem activity

An unusual and striking effect of external glutamate onArabidopsis root growth and branching has recently beenreported (Walch-Liu et al., 2006b). When Arabidopsisroots were exposed to low concentrations of L-glutamatethere was a marked inhibition of primary root growth andan increase in root branching near the root apex. Thiseffect on root architecture resulted from inhibition ofmeristematic activity at the primary root tip and an initialstimulation of lateral root outgrowth in the apical regionof the primary root. Lateral roots were also glutamatesensitive, but only after they had reached 5–10 mm inlength, indicating a developmentally delayed response.A similar response to glutamate was found in roots ofa number of other species, including Thlaspi caerulescens,Thellungiella halophila, wild poppy, and tomato (Walch-Liu et al., 2006a, b). Intriguingly, different Arabidopsis

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ecotypes displayed widely differing sensitivities to gluta-mate, the most sensitive being C24 and the least sensitiveRLD1.

Both positive and negative effects of amino acids onplant growth have been reported by other authors (Skinnerand Street, 1953; Rognes et al., 1986; Katonoguchi et al.,1994; Barazani and Friedman, 2000), and a rapid in-hibition of primary root growth by glutamate was reportedby Sivaguru et al. (2003). However, the effects observedby Walch-Liu et al. (2006b) are distinctive in a number ofrespects. Most notably, the effects were detected at muchlower concentrations than the millimolar concentrationscommonly used in previous studies and were stronglygenotype dependent. Furthermore, the effects were highlyspecific to L-glutamate, not being mimicked by relatedamino acids such as aspartate, glutamine, and GABA, oreven by the D-stereoisomer of glutamate. The authorswere able to rule out the possibility that the L-glutamatetreatment was causing any kind of general nutritional ormetabolic disturbance to the plant. Indeed, glutamate wasonly inhibitory to primary root growth if it was applieddirectly to the primary root tip.

The specificity of the glutamate effect, its localizednature, and its occurrence at low concentrations (>20 lM)led to the suggestion that the root tip is probablyresponding to changes in the apoplastic L-glutamate con-centration, and that some kind of signalling mechanism isinvolved (Walch-Liu et al., 2006b). Attempts to useagonists and antagonists of mammalian iGluRs to inves-tigate the possible involvement of AtGLR-encoded gluta-mate receptors proved unfruitful: the antagonists tested[DNQX, MK-801, and 2-amino-5-phosphonopentanoate(AP-5)] neither suppressed the L-glutamate effect norhad any direct effect on root growth (Walch-Liu andForde, 2007). Furthermore, although the agonist BMAAwas inhibitory to root growth, the root phenotype it pro-duced was very different from that seen with L-glutamate(Walch-Liu and Forde, 2007).

Glutamate signalling in plants: autocrine/paracrine, endocrine, or environmental?

What might be the functional relevance of an exogenousglutamate-sensing mechanism? It has been noted that therange of concentrations reported for apoplastic glutamatein a variety of tissues and species is 0.3–1.3 mM, whichmatches fairly closely the range of glutamate concentra-tions in which glutamate-elicited currents are half activated(0.2–0.5 mM) (Demidchik et al., 2004). Although verylittle is known about glutamate fluxes and the dynamics ofapoplastic glutamate in plant tissues, it seems possible thatglutamate could participate in intercellular signalling, andeven in long-distance signalling within the plant.

There is a striking precedent for an amino acid-likesignalling molecule in plants, in the form of auxin. The

major form of auxin in plant tissues, indole-3-acetic acid(IAA), is an analogue of tryptophan and its role as a planthormone is implicated in almost every aspect of plantbiology. Like glutamate, auxin added exogenously toplant cells elicits rapid membrane depolarization anda transient increase in [Ca2+]cyt (Zimmermann et al.,1999). It is clear that exogenous auxin is able to modifyroot growth at very much lower concentrations thanglutamate (10�11 M versus 10�5 M) (Evans et al., 1994),an observation that might be taken to indicate that theirsignalling properties are very different. However, thisdisparity may simply reflect the fact that glutamate, unlikeauxin, has to fulfil the dual role of being both a signalmolecule and a key primary metabolite whose endogenousconcentration must be maintained at levels appropriate forcatalytic activity (see above).

For glutamate to act as an autocrine or paracrine signalin plants, it would be necessary to have efficientmechanisms for its efflux and re-absorption, as there arein the synapse (Moriyama and Yamamoto, 2004). Auxinefflux in plant tissues appears to be mediated bya vesicular cycling mechanism (Blakeslee et al., 2005),and parallels have been drawn between this process andthe mechanism by which glutamate is secreted duringsynaptic transmission in mammals (Balusxka et al., 2005).It is therefore possible that glutamate effluxes into theapoplast in a similar way. Alternatively, glutamate couldbe released from plant cells directly via anion channelssuch as those that facilitate organic acid efflux in roots(Roberts, 2006).

Effective mechanisms for glutamate re-absorption alsoexist. Six members of the Arabidopsis AAP family(AAP1–AAP6) have been heterologously expressed andshown to catalyse the low affinity influx of a broad rangeof amino acids, including glutamate (Km

Glu¼0.36–5.0 mM)(Fischer et al., 2002), while the related LHT1 geneappears to encode a high-affinity glutamate influx system(Km

Glu ;14 lM) (Hirner et al., 2006).There is preliminary evidence that an autocrine/para-

crine signalling pathway involving glutamate might beoperating in Al3+-treated Arabidopsis root tips (Sivaguruet al., 2003). Rapid responses to Al3+, including mem-brane depolarization and microtubule depolymerization,could be mimicked by glutamate treatment, but wereevoked more rapidly by glutamate. The authors proposedthat Al3+ signalling in the root tip may be initiated byefflux of glutamate (or a glutamate-like ligand) via anAl3+-gated anion channel, with subsequent binding of thisligand to a glutamate receptor (Fig. 3). This may be anexample of the stress response mechanism originallypostulated by Dennison and Spalding (2000), in whichendogenous or environmental factors trigger glutamateefflux into the apoplast to activate glutamate-gated chan-nels, causing a transient change in [Ca2+]cyt and a cascadeof downstream responses (Dennison and Spalding, 2000).


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As an amino acid commonly found in the phloem sap,glutamate could potentially also perform a long-distancesignalling role. It was the observation that amino acidscycle between the shoot and root that first prompted thehypothesis that these amino acids could enable the shootto communicate its changing N/C status to the root andthereby regulate the nitrate uptake system appropriately(Cooper and Clarkson, 1989). Although evidence support-ing this hypothesis is still largely lacking (Forde, 2002), itis nevertheless an attractive concept that deserves furtherinvestigation. One possible way in which a long-distanceglutamate signal might operate in the context of rootgrowth is depicted in Fig. 3. In this model, shoot-derivedglutamate arriving at the root tip [via a partially apoplasticroute similar to the one taken by shoot-derived auxin(Marchant et al., 2002)] is sensed by plasma membraneglutamate receptors, potentially enabling meristematicactivity in the root tip to respond to changes in the N/Cstatus of the shoot.

A further possibility, and one that does not exclude thepotential for glutamate to act as an endogenous signal, isthat roots have evolved the capacity to respond tovariations in glutamate concentration in the externalenvironment (Walch-Liu et al., 2006a, b) (Fig. 3).Glutamate is one of the most abundant amino acids in thesoil, being a component of the pool of dissolved organicnitrogen (DON), as well as a constituent of root exudates(Paynel et al., 2001). In the bulk soil solution, glutamateconcentrations may be <10 lM (Jones et al., 2005), butconcentrations are expected to be much higher locally,particularly in the vicinity of decomposing animal orvegetable matter (Walch-Liu et al., 2006a). As the rootsystem develops and the growing root tips continuouslyexplore the soil volume, they can therefore expect toencounter significant variations in the external glutamateconcentration. In agar plate experiments, locally highglutamate concentrations of >50 lM were shown to slowprimary root growth and encourage root branching behindthe primary root tip (Walch-Liu et al., 2006b). It was

pointed out that this response, if replicated in the field,would lead to an increase in the density of the root systemwithin glutamate-rich patches of soil. Although differingfrom the commonly observed root foraging response tolocalized supplies of inorganic nutrients (NO3

–, Pi, etc.),which is based on the direct stimulation of lateral rootgrowth and branching (Robinson, 1994), this response toglutamate can similarly be interpreted as a means toenhance the precision of root placement within the soil.The outcome in both cases is to focus root growth in theregion of the soil where resources are most abundant andtherefore to enhance the plant’s ability to compete (withits neighbours and microorganisms) for those resources.

There is an increasing appreciation of the potentialimportance of organic N as a source of N for plants (Joneset al., 2005; Weigelt et al., 2005), and it has been arguedthat plants will be most effective at competing with micro-organisms in organic N-rich patches where amino acidconcentrations are highest (Jones et al., 2005). It will beinteresting to establish whether the variation in glutamatesensitivity observed between different Arabidopsis eco-types has an adaptive significance, related, for example, tothe importance of organic N as an N source in their nativesoils.

Concluding remarks

Glutamate plays a central role in plant nitrogen metabo-lism. It may be synthesized and metabolized by a numberof different pathways (Fig. 1). There is evidence thatunder most circumstances the plant is able to maintain thesoluble concentration within fairly narrow limits. Themechanism by which this occurs involves a number ofenzymes present within different cellular compartments, inparticular GS, glutamate synthase, and GDH. It is likelythat, ultimately, the supply of 2-oxoglutarate catalysed byisocitrate dehydrogenase will play a key role in regulatingthe accumulation of glutamate.

The mounting evidence in favour of a signalling role forglutamate has been reviewed here. However, untilglutamate can conclusively be shown to be a ligand ofa specific GLR-type receptor, or until a receptor forglutamate of a different type is identified, the case must beconsidered unproven. As has been discussed elsewhere(Lam et al., 2006), glutamate signalling may be part ofa much broader network of N signalling pathways thatenable the plant to monitor and adapt to changes in itsN status and the N supply.

In terms of the effect of external L-glutamate on rootarchitecture, several key questions need to be addressed.First, what is the molecular basis for L-glutamate sensingand signal transduction at the root tip? Secondly, whataccounts for the remarkable natural variation in L-gluta-mate sensitivity between Arabidopsis ecotypes? Thirdly,

Fig. 3. Possible modes of glutamate signalling in the root. The areaindicated in yellow represents the symplast at the root tip. See text forfurther details.

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does the L-glutamate-elicited change in root architectureconstitute an adaptive response, perhaps by enhancing theplant’s ability to compete for localized patches of organicN? The first two questions are currently being addressedusing forward and reverse genetics and quantitative traitloci analysis (P Walch-Liu, T Remans, and BG Fordeunpublished data). Furthermore, mutants and near-isogeniclines differing in their glutamate sensitivity that areemerging from these studies will make it possible to testexperimentally the adaptive significance of the rootarchitectural response to external L-glutamate.

Acknowledgements

Work in BGF’s laboratory has been supported by grants from theBiotechnology and Biological Sciences Research Council (BBSRC)and by European Commission Research Training Network grant no.HPRN-CT-2002-00247 (PLUSN).

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