Anion channels in plant cells

MINIREVIEW

Anion channels in plant cellsHannes Kollist1, Mathieu Jossier2, Kristiina Laanemets1 and Sebastien Thomine2

1 Institute of Technology, University of Tartu, Estonia

2 Institut des Sciences du Vegetal, CNRS, Gif-sur-Yvette, France

Keywords

ALMT1; ion channel; root; SLAC1; stomata

Correspondence

S. Thomine, Institut des Sciences du

Vegetal, CNRS, Avenue de la Terrasse,

91 198 Gif-sur-Yvette, France

Fax: +33 1 69 82 37 68

Tel: +33 1 69 82 37 93

E-mail: [email protected]

(Received 28 April 2011, revised 13 July

2011, accepted 25 July 2011)

doi:10.1111/j.1742-4658.2011.08370.x

Plant anion channels allow the efflux of anions from cells. They are

involved in turgor pressure control, changes in membrane potential,

organic acid excretion, tolerance to salinity and inorganic anion nutrition.

The recent molecular identification of anion channel genes in guard cells

and in roots allows a better understanding of their function and of the

mechanisms that control their activation.

Introduction

The role of anions in plant cells is clearly distinct from

that encountered in animal cells. First, chloride is by far

the most prevalent anion in animal cells whereas plant

cells contain a complex mixture of diverse anions includ-

ing, besides chloride, nitrate, sulfate, phosphate and

organic anions, such as citrate, malate or oxalate in

varying proportions [1]. Second, anions are accumulated

in plant cells, whereas in most animal cells the chloride

gradient favors the influx. Third, in mature plant cells,

the main reservoir of ions is not the cytosol but rather

the central vacuole which occupies about 80% of the cell

volume. These differences imply original functions for

plant anion channels and probably account for the

recent discovery of anion channel gene families unique

to plants and not found in animal genomes.

Ion channels switch between open and closed states

according to the factors that control their gating.

When an ion channel is open, massive ion fluxes occur

according to their electrochemical gradients. An

important step in studying and understanding the

function of ion channels was development of the patch

clamp technique by Neher and Sakmann in the late

1970s [2]. In contrast, the first anion channel structure

was determined in bacteria only in 2002 [3] and the

first plant genome sequence was released in 2000 [4].

Thus, the electrophysiological properties of plant cell

membranes were thoroughly studied before the genes

encoding anion channels were identified.

At the level of plant cell plasma membrane, both the

membrane potential, which is highly electronegative

(usually below )100 mV), and the intracellular anion

accumulation dictate anion efflux through anion chan-

nels when they open [5]. In addition, in the case of

organic anions such as malate and citrate, which are

carboxylic acids, the pH gradient (neutral inside, acidic

outside) further favours their efflux through channels,

as their protonation in the extracellular space main-

tains a steep gradient of the anionic species. The

slightly negative electrical gradient across the vacuolar

membrane (tonoplast) also drives anion ‘efflux’ from

Abbreviations

ABA, abscisic acid; ABC, ATP binding cassette; ALMT, aluminum activated malate transporter; CLC, chloride channel; CPK, calcium-

dependent protein kinase; MATE, multidrug and toxic efflux transporter; NAXT1, nitrate excretion transporter 1; NRT, nitrate transporter;

OST1, open stomata 1; PKC, protein kinase C; PP2C, protein phosphatase type 2C; PTR, peptide transporter; PYR, pyrabactin; QUAC,

quickly activating anion channel; R-type, rapid-type anion channel; SLAC1, slow anion channel associated 1; S-type, slow-type anion channel.

FEBS Journal 278 (2011) 4277–4292 ª 2011 The Authors Journal compilation ª 2011 FEBS 4277

the cytosol to the vacuolar lumen. However, active

transport, such as H+ ⁄ anion co-transport, is clearly

required for the high accumulation of certain anions in

this compartment [6,7]. Paradoxically, active transport

may also be necessary to release anions from the vacu-

ole when the cell undergoes important changes in tur-

gor pressure.

The diversity of anions in plant cells means that anion

channels serve a wide range of functions. Whatever its

anion selectivity, the opening of an anion channel in the

plasma membrane shifts the membrane electrical poten-

tial towards the equilibrium potential of anions, i.e. it

will lead to a depolarization [8,9]. Cell depolarization

can induce signalling events or lead to the activation of

voltage-gated ion channels. When anion efflux through

anion channels is coupled to potassium efflux, anion

channels act as major players in plant cell osmotic regu-

lation. Depending on their selectivity, anion channels

may play more specific roles. For example, chloride-

selective channels may be involved in salt tolerance

[10,11]; nitrate-selective channels in nitrogen homeosta-

sis and organic-acid-selective channels in carbon metab-

olism (e.g. malate channels in CAM (crassulacean acid

metabolism) plants) or pH regulation [12].

Besides their selectivity, another important feature

of anion channels is their gating. It is unlikely that a

plant cell can survive with constitutively open anion

channels because this would lead to massive loss of

ions and depolarization. This review will provide

examples of anion channels gated by diverse mecha-

nisms. Many of the anion channels described are volt-

age regulated, opening in response to membrane

depolarization or hyperpolarization [13]. Additionally,

some channels display strong regulation by intracellu-

lar signalling events, such as phosphorylation, and

intracellular or extracellular metabolites [14–17]. In

addition to canonical anion channels characterized by

open–closed transitions, the review will address trans-

porters that allow anion fluxes along their electrochem-

ical gradient (i.e. MATE or NAXT) but have not been

characterized as channels stricto sensu. Such transport-

ers obey similar biophysical constraints and lead to

similar consequences for the cell and their detailed

characterization may reveal in some cases that they

function as channels. Particular attention will be given

to the most recent progress on two systems in which

anion channels have been intensively studied: the

guard cells of stomata, which undergo fast reversible

anion channel dependent change in turgor, and roots,

which illustrate the functions of anion channels in

anion excretion to the rhizosphere or to the xylem.

For other aspects of plant anion channel biology, the

reader is referred to other review articles [1,9,18–21].

Guard cell anion channels: fromelectrophysiological characterizationto molecular structure

Stomata are small pores in the epidermis of plant

leaves and stems, which control plant gas exchange.

Each stomatal pore is surrounded by a pair of guard

cells which control the pore size by swelling or shrink-

ing as a response to changes in the surrounding envi-

ronment or to intrinsic signals. This depends on the

activation of ion channels, changes in the guard cell

osmotic pressure and movement of water in or out of

the guard cells. Adequate stomatal regulation ensures

sufficient uptake of carbon dioxide with minimal loss

of water. This is particularly important in situations

where water resources are limited. Rapid stomatal

closure also limits the entrance of pathogens [22] and

air pollutants such as ozone [23,24].

The activation of guard cell anion currents was recog-

nized already more than 20 years ago as one of the first

steps in the induction of plant stomatal closure [25,26];

however, genes encoding guard cell anion channels were

characterized only recently [24,27,28]. Activation of

guard cell anion channels and release of anions is the

critical step for induction of plant stomatal closure

[29,30]. This depolarizes guard cell plasma membrane

and triggers the activation of guard cell outwardly recti-

fying K+ channels [31]. Overall the osmotic pressure

inside the cell is reduced, water flows out, guard cells

become flaccid and the stomatal pore closes.

Guard cell plasma membranes exhibit fast- and

slow-type anion channel activities

Applying the patch clamp technique to the plasma

membrane of Vicia faba guard cell protoplasts led to

the characterization of two distinct, rapid-type (R-type)

and slow-type (S-type), anion channels [13,32,33]. Acti-

vation of R-type anion channels is voltage dependent

and they activate ⁄deactivate within milliseconds. In

V. faba guard cells, R-type currents also exhibit time-

dependent inactivation within tens of seconds. The

activation ⁄deactivation time constants of S-type anion

channels are in the range of 10 s, channel activity

shows weak voltage dependence and S-type currents

do not inactivate with time. Both channels participate

in stomatal closure but not equally in all responses.

Drought-induced plant hormone abscisic acid (ABA)

activates both S- and R-type anion currents [8]. In

contrast, increases in CO2 partial pressure trigger

consistent activation of S-type currents only, whereas

R-type channels are either activated or inactivated [34].

Plant anion channels H. Kollist et al.

4278 FEBS Journal 278 (2011) 4277–4292 ª 2011 The Authors Journal compilation ª 2011 FEBS

Identification, regulation of SLAC1

A gene encoding the guard cell plasma membrane

S-type anion channel was genetically isolated from

independent Arabidopsis mutant screens for ozone

sensitive rcd (radical induced cell death) and for car-

bon dioxide insensitive (cdi) mutants [35,36]. Both rcd3

and cdi3 turned out to carry mutations in the gene

At1g12480 and were thus renamed slac1-1 (S456F) [24]

and slac1-2 (G194D) [28], respectively. Guard cells

extracted from slac1 mutants do not have detectable

Ca2+- and ABA-induced S-type anion channel activity,

whereas activation of R-type channels remains intact

[24]. Heterologous expression of SLAC1 alone in

Xenopus oocytes does not generate clear anion

currents. Therefore, as it could not be excluded that

SLAC1 represented only a subunit of the S-type anion

channel, the protein was initially named slow anion

channel associated 1 [24]. SLAC1 expression is highly

specific for guard cells. Plants lacking functional

SLAC1 display impaired stomatal closure in response

to all major endogenous (ABA, Ca2+, NO) and envi-

ronmental (CO2, darkness, humidity, ozone) stimuli.

These results illustrate the crucial role of guard cell

plasma membrane S-type anion current activation for

the induction of stomatal closure [24,28,37].

The importance of regulatory processes such as

phosphorylation and dephosphorylation for activation

and inactivation of S-type anion channels was shown

already by Schmidt et al. [17]. On this basis, it was

suggested that lack of regulatory proteins in Xenopus

oocytes could be the reason why initial experiments

trying to produce SLAC1-type anion currents in heter-

ologous systems failed [24]. This possibility was inde-

pendently tested by two laboratories [15,38] which

showed that co-expression of SLAC1 with a protein

kinase consistently induced S-type anion channel cur-

rents in Xenopus oocytes confirming that SLAC1 forms

the guard cell plasma membrane S-type anion channel

per se. Functional expression of SLAC1 reconstituted

anion currents with slow kinetics, a higher permeabil-

ity for nitrate than for chloride and low permeability

to malate, bicarbonate and sulfate, as observed in

guard cells [15,39,40]. The large increases in malate

and fumarate content in slac1-2 and slac1-3 guard cells

are thus probably not due to lack of efflux of these

organic anions through the slow anion channels but is

associated instead with a profound alteration in guard

cell function in these mutants [28].

Both Geiger et al. [15] and Lee et al. [38] showed

that phosphorylation by the protein kinase OST1 (also

known as SnRK2.6 or SRK2E) is critical for full acti-

vation of SLAC1-dependent S-type anion currents in

Xenopus oocytes (Fig. 1). In addition, S-type anion

channel activities were clearly reduced in guard cells

isolated from the ost1-2 mutant illustrating the impor-

tance of OST1-dependent phosphorylation for SLAC1

activation in guard cells [15]. Additional evidence for

OST1-dependent activation of SLAC1 was provided

by Vahisalu et al. [41] who showed that OST1 phos-

phorylates multiple serines of SLAC1 hydrophilic

N-terminal fragment. Furthermore, plants carrying a

mutation in one of the phosphorylated serines (S120)

show impaired stomatal responses to ozone [41], simi-

lar to that of slac1 loss-of-function mutants [24], indi-

cating that OST1-dependent phosphorylation at this

site is physiologically relevant for induction of stoma-

tal closure. The C-terminal tail of SLAC1 is also

phosphorylated by OST1 in vitro [38]. Nevertheless,

SLAH1, a homologue of SLAC1 which lacks the

N- and C-terminal tails phosphorylated in SLAC1,

fully complements the stomatal phenotypes of slac1-2

when expressed in guard cells under control of the

SLAC1 promoter [28].

Elevation of cytosolic Ca2+ also triggers the activa-

tion of guard cell slow anion currents [26]. Analysis of

S-type anion channel activation by Ca2+ in the context

of ABA signaling showed that treatment by ABA or

protein phosphatase inhibitors facilitates Ca2+-medi-

ated activation of slow anion currents by lowering the

intracellular Ca2+ concentration required to trigger

their activation [42,43]. One way of translating Ca2+

signals goes through activation of Ca2+-dependent pro-

tein kinases (CPKs, Fig. 1). Guard cells from single and

double mutants of Arabidopsis CPK3 and CPK6 have

impaired ABA- and Ca2+-induced S-type anion cur-

rents [44]. Recently Geiger et al. [45] showed physical

interaction between CPK21, CPK23 and SLAC1. Fur-

thermore, co-expression of these proteins with SLAC1

activates S-type anion currents in heterologous systems.

However, only CPK21 kinase activity is Ca2+ sensitive.

Elevation of CO2 leads to the activation of S-type

anion currents. Recent data demonstrated that the

activation of SLAC1 currents is mediated by intracel-

lular bicarbonate generated from CO2 by b-carbonicanhydrase, rather than by intracellular pH changes

[46,47]. Bicarbonate-induced activation of S-type anion

currents is positively controlled by OST1 kinase and

negatively by HT1 kinase [47,48]. Interestingly, this

activation requires elevated intracellular calcium levels

suggesting the need for concomitant signals for guard

cell CO2 response.

One of the major recent breakthroughs in plant biol-

ogy was the discovery of the cytosolic ABA receptor

PYR ⁄PYL ⁄RCAR proteins [49,50], which inhibit pro-

tein phosphatase 2Cs (PP2Cs), such as ABI1 and

H. Kollist et al. Plant anion channels


ABI2, by the formation of a ternary complex between

ABA, PP2Cs and PYR ⁄PYL proteins (Fig. 1). This

mechanism also controls the OST1-dependent activa-

tion of SLAC1 as, in the absence of ABA, OST1 is

kept inactive by PP2Cs. Upon ABA binding to the

receptor complex, the activity of the PP2Cs is inhibited

and OST1 is activated either by autophosphorylation

[51] or by an upstream kinase [52,53]. In agreement

with these results, protein phosphatases ABI1, ABI2

[15] and PP2CA [38] abolish OST1-dependent activa-

tion of SLAC1-induced anion currents in Xenopus

oocytes. Interestingly ABI1 and ABI2 also control

CPK21- and CPK23-dependent activation of SLAC1

currents [45] suggesting that Ca2+-dependent SLAC1

activation is also controlled by ABA.

These findings provide an elegant signaling module

for Ca2+-dependent and Ca2+-independent regulation

of plant guard cell S-type anion channel SLAC1

(Fig. 1). However, the functional significance of

CPK21 and CPK23 for stomatal regulation in planta is

not yet fully understood. Lack of functional SLAC1 or

OST1 causes impaired stomatal response to ABA

[24,54], air humidity [24,55] CO2 [47] and ozone [41],

indicating that OST1-dependent activation of SLAC1

is required for stomatal closure in response to these

stimuli. In contrast, plants with impaired CPK23 gene

do not exhibit any stomatal phenotypes even though

their Ca2+-induced activation of SLAC1 is reduced by

70% [45]. Similarly, stomatal responses to major envi-

ronmental stimuli seem to be intact in plants with

impaired CPK21 gene (E Merilo & H Kollist, unpub-

lished). Collectively, current data suggest that,

although three different protein kinases can activate

SLAC1, OST1-dependent phosphorylation is the main

prerequisite for SLAC1 activation in guard cells.

Structure of SLAC1 and its homologues

Recently, the 3D structure of HiTehA, a bacterial

homologue of SLAC1, was resolved at high resolution

(Fig. 2). The bacterial TehA does not possess a soluble

C-terminal domain comparable with that found in

SLAC1 protein. Except for this domain, a model of the

SLAC1 3D structure could be constructed with high

confidence based on the 3D structure of TehA [39,56].

This revealed three critical features of this channel: its

trimeric structure, the properties of its pore and an

essential gating mechanism. First, TehA forms homo tri-

mers (Fig. 2C). This feature may account for the coop-

erative opening of this channel which was noted already

in early single channel recordings [40]. In addition, a

multimeric structure suggests a possible mode of activa-

Fig. 1. Regulation of guard cell slow-type (SLAC1 ⁄ SLAH3) and rapid-type (QUAC1) anion channels. SLAC1 ⁄ SLAH3: (1) In the absence of ABA

(grey box), 2C type protein phosphatases (PP2C) inactivate protein kinases OST1, CPK21 and CPK23 via dephosphorylation. (2) In the presence

of ABA, PP2Cs are inactivated by the formation of a ternary complex between ABA, cytosolic ABA receptors (PYR ⁄ PYL) and PP2Cs. (3) This in

turn leads to the activation of protein kinases which activate SLAC1 via phosphorylation and anions are released from the guard cell. (4) Activa-

tion of CPK21, but not CPK23, is dependent on Ca2+. (5) CPK21 also activates SLAH3, another S-type anion channel mainly permeable to NO�3 .

Activation of SLAH3 is enhanced by extracellular NO�3 through an effect on its gating by membrane potential. (6) Potassium uptake channel

KAT1 phosphorylation by OST1 negatively regulates Kþin activity, further supporting stomatal closure. (7) Mutants of calcium-dependent protein

kinases CPK3 and CPK6 and double mutant of MAP kinases MPK9 and MPK12 have also been shown to have impaired S-type anion channel

activity but the mechanism is not known. QUAC1: (8) Activation of QUAC1 ⁄ AtALMT12 is highly voltage dependent with peak activities near

)100 mV. Extracellular malate shifts QUAC1 activation to more negative values and enhances QUAC1 activity. QUAC1 is permeable to organic

anions such as malate and fumarate. Activation of SLAC1, SLAH3 and QUAC1 induces the release of anions and membrane depolarization,

which leads to the activation of voltage-gated Kþout channel GORK, guard cell turgor loss and stomatal closure.



tion by phosphorylation at the interface between the su-

bunits [57]. Second, the structure of TehA sheds light on

the structure of the anion-permeable pore of SLAC1.

Each subunit forms an independent pore (Fig. 2B, C,

E). The pore has a remarkably constant diameter of

about 5 A through the membrane and is lined by hydro-

phobic or hydroxyl residues (Fig. 2B, D, E); it does not

show any distinct binding site for anion in contrast to

the anion permeation pathway through chloride channel

(CLC) proteins [3]. Such weak interactions between the

pore and the permeating anions account for the

observed selectivity sequence of different anions, which

follows the energetic cost of their dehydration. Third,

the structure of TehA revealed the presence of a phenyl

ring blocking the anion permeation pathway (Fig. 2B,

D). This phenylalanine residue (F450 in SLAC1) is con-

served through the entire SLAC1 protein family. Muta-

tional analysis demonstrated that this phenyl ring gates

the pore of both TehA and SLAC1 [39]. The ability of

the channel to switch between closed and open states

thus relies on the ability to move the phenyl ring away

from the anion permeation pathway, probably by phos-

phorylation-induced conformational change of the pro-

tein. Thus, a central question will be to understand how

phosphorylation of SLAC1 is coupled to the positioning

of the phenyl ring in the permeating pore.

In addition to SLAC1, the Arabidopsis genome

encodes four SLAC1 homologues, SLAH1–4 [28,39]. At

the subcellular level, SLAH1–3 are located in the plasma

membrane [28]. Transformation with SLAH1 and

SLAH3 under control of the SLAC1 promoter rescued

slac1 mutant phenotypes, indicating that SLAH1 and

SLAH3 are also capable of forming S-type anion chan-

nels. SLAH1 and SLAH2 are expressed in roots and

SLAH3 is expressed in whole plant. Initial characteriza-

tion indicated that SLAH1–3 were not expressed in

guard cells [28]. However, a recent study revealed that,

in different growth conditions, SLAH3 is expressed in

guard cells where it functions as an S-type anion channel

[58]. Similar to SLAC1, SLAH3 activation is regulated

by phosphorylation by CPK21, which is controlled by

the ABA receptor–phosphatase complex (Fig. 1). In

contrast to SLAC1, SLAH3 voltage dependence is mod-

ulated by extracellular NO�3 which facilitates SLAH3

activation by shifting its activation threshold towards

guard cell resting membrane potential [58]. The function

of SLAH3 and its regulation by nitrate in guard cell

movements remains to be investigated.

Identification of AtALMT12 ⁄ QUAC1 –

a component of guard cell R-type anion channel

activation required for plant stomatal closure in

response to major endogenous and

environmental stimuli

Recently two parallel studies characterized Arabidopsis

AtALMT12 (aluminum activated malate trans-

porter12), which on the basis of sequence is similar to

ALMT1 (see below) and showed that this protein is

required for full stomatal closure induced by various

Fig. 2. Structure of bacterial TehA and homology model of plant SLAC1. Ribbon diagram of HiTehA viewed from within the membrane, from

the side (A) or from the top (B), and ribbon diagram of HiTehA trimer (C). (D) Cross-section through the homology model of AtSLAC1. Colors

show the electrostatic potential from electronegative (red) to electropositive (blue). Cylinder model of SLAC1 (E) and pore-lining residues in

the SLAC1 homology model (F). Reprinted by permission from Macmillan Publishers Ltd, Nature (Chen et al., 2010 [39]), copyright 2010.



stimuli [27,59]. AtALMT12 is preferentially localized

to the plasma membrane of guard cells; moreover

detailed electrophysiological studies of almt12 guard

cells and Xenopus oocytes expressing the protein

revealed that AtALMT12 forms the malate-sensitive

R-type anion channel (Fig. 1) [27]. AtALMT12 acti-

vates when plasma membrane is depolarized [27]. The

maximum current of R-type anion channels is around

)100 mV in Arabidopsis thaliana guard cells [25,27,60].

R-type anion currents generated upon expression of

AtALMT12 in Xenopus oocytes display a peak

around )100 mV which is shifted to more negative

values when extracellular malate concentration is

increased (Fig. 1) [27]. Contrary to what its name

implies, extracellular Al3+ treatment does not stimu-

late AtALMT12-dependent anion currents [27,59];

thus it was suggested to rename the protein quickly

activating anion channel 1, QUAC1 [27]. However, it

should be noted that R-type anion currents are only

reduced by 40% in almt12 mutant guard cells and

that, in the absence of extracellular malate, R-type

currents are indistinguishable from those observed in

wild-type [27,59]. This implies that other proteins also

play a role in the formation of guard cell R-type

anion channels. Of the 14 Arabidopsis ALMTs [61]

clear guard cell specific localization is only shown for

AtALMT12 ⁄QUAC1; however, AtALMT13 and A-

tALMT14 are good candidates based on their high

homology to AtALMT12 ⁄QUAC1 [18].

What is the relative contribution of S-type and

R-type anion channels in stomatal movements?

It is well established that the activation of S-type anion

currents requires phosphorylation by protein kinases

[15,38,41,45]. In contrast, no direct intracellular signal-

ing pathway is known for R-type anion channel activa-

tion or deactivation. R-type anion channel activity is

tightly regulated by membrane potential [27,59,60]. It

is therefore possible that R-type anion channel activa-

tion does not require any intracellular activation

pathway but is merely triggered by membrane depolar-

ization. The hypothesis that R-type channel does not

require any intracellular activator is indirectly sup-

ported by the recent finding that AtALMT12 ⁄QUAC1

is fully functional in Xenopus oocytes [27] without the

need of other plant proteins to activate it, as opposed

to SLAC1 [15,38,45]. In open stomata, which have a

strongly hyperpolarized membrane potential around

)150 mV, the guard cell R-type anion channels are

inactive (Fig. 1). Depolarization of plasma membrane

can be achieved by activation of S-type anion channel

[26,30,32], inhibition of H+-ATPases [62,63], activa-

tion of Ca2+ channel [60,64] or a combination of these

processes. In addition, increases in extracellular malate

concentration shift R-type outward anion current acti-

vation threshold to more negative voltages [16,27].

Thus the activation of R-type anion channels can be

triggered either by membrane depolarization or by an

increase in extracellular malate concentration. This

suggests the presence of a feed-forward regulation for

R-type anion channel activation where a slight mem-

brane depolarization would trigger a slight activation

and the release of malate which in turn would lead to

enhanced activation due to the shift of the activation

threshold to more hyperpolarized potentials.

Stomatal opening is initiated by plasma membrane

H+-ATPase driven proton efflux from guard cells.

This shifts the plasma membrane electrical potential

towards hyperpolarized values, which in turn triggers

activation of voltage-gated potassium uptake channels

[65]. Concomitant accumulation of positive charges

inside the guard cells has to be balanced by anions.

Classical studies from the 1970s by Raschke and Out-

law and their colleagues [66,67] showed that guard

cells are capable of anion uptake from the extracellular

space. In the absence of extracellular anions, stomatal

opening is achieved by biosynthesis of organic anions

inside the guard cells. Recently, it was shown that the

Arabidopsis ABC transporter AtABCB14 mediates

malate and possibly fumarate uptake from the apo-

plast to the guard cells during stomatal opening [68].

Conversely, anion efflux channels such as SLAC1 and

QUAC have to be closed to allow for guard cell turgor

build-up during stomatal opening. The membrane

hyperpolarization induced by proton pump activation

during stomatal opening is probably sufficient to close

R-type ⁄QUAC channels, due to their steep voltage

dependence. As described above, SLAC1 is activated

by phosphorylation. An intriguing question is whether

the same PP2Cs that inhibit OST1 activity also func-

tion in SLAC1 inactivation by dephosphorylation. It

was shown that PP2CA physically interacts with

SLAC1, and inhibits SLAC1 activity in Xenopus

oocytes [38]. In addition, other PP2Cs are able

to dephosphorylate the SLAC1 N-terminal domain

in vitro (Kollist, unpublished results) suggesting that

ABA-PYR ⁄PYL-PP2C signaling module might also

control SLAC1 inactivation during stomatal opening.

Anion fluxes through the vacuolar membrane of

guard cells

During stomatal movements, guard cells undergo great

dynamics changes in vacuole morphology. These

changes, associated with ion fluxes across the tono-



plast, are essential for stomatal movements, highlight-

ing the crucial function of the vacuole during this pro-

cess [69]. Vacuolar chloride channel currents regulated

by a calcium-dependent protein kinase were identified

in V. faba guard cells [70]. Apart from this report, lit-

tle is known about anion channels at the tonoplast of

guard cells.

AtMRP5 belongs to the ATP-binding cassette

(ABC) transporter family and is expressed in guard

cells [71,72]. Disruption of AtMRP5 leads to an ABA

insensitivity of stomatal closure. This is in agreement

with reports showing that ABC transporter modulators

affect guard cell anion currents and stomata aperture

[73,74]. It had been proposed that AtMRP5 is a sub-

unit or acts as a regulator of guard cell S-type anion

channels [75]. Recently, it was shown that AtMPR5 is

an inositol-hexakisphosphate transporter presumably

localized at the tonoplast [72]. The implication of ino-

sitol-hexakisphosphate in calcium mobilization and

inhibition of inward rectifying K+ conductance in

guard cells may account for the stomatal phenotype of

atmrp5 [72,76].

Recently, AtCLCc, a member of the CLC family was

shown to be targeted to the tonoplast and implicated in

stomatal movements. Plants lacking functional AtCLCc

display light and ABA insensitivity of stomatal move-

ments associated with a dramatic decrease in guard cell

chloride content [11]. The role of AtCLCc in chloride

homeostasis provided the first evidence of the impor-

tance of coordination between anion transport at the

plasma and vacuolar membranes during stomatal move-

ments. As the CLC family comprises channels and trans-

porters, electrophysiological studies will be necessary to

establish which of these two transport mechanisms is

used by AtCLCc. Three other AtCLC members are

located in the tonoplast (AtCLCa, AtCLCb and At-

CLCg); however, only one of them, AtCLCa, is

expressed in guard cells [11]. AtCLCa encodes an

NO�3 ⁄H+ antiporter involved in NO�3 homeostasis [6].

The high expression of AtCLCa in guard cells suggests

that it may also be implicated in stomata movements

[11]. This raises the possibility that two vacuolar CLC

family members, AtCLCc and AtCLCa, with distinct

preferences for chloride or nitrate, respectively, could

cooperate to mediate anion transport across guard cell

tonoplast depending on the availability of these two

anions. (See note added in proof.)

Root anion channels: anion excretionand loading into the xylem

Anion channels have been described in all root cell

types investigated [20]. Besides the ubiquitous rele-

vance of anion fluxes in any plant cell, anion channels

fulfill several root-specific functions. These functions

include anion loading to the xylem and anion excretion

to the rhizosphere. Xylem loading allows anion trans-

location to the shoots; it is especially relevant for

nitrate which is taken up by the root but mostly

reduced to be assimilated in amino acids in the leaves

[77]. Xylem loading of organic anions such as citrate is

important for the translocation of metal cations that

move from the root to the shoot as complexes with

organic acids [78,79]. Anion excretion to the rhizo-

sphere also serves diverse functions. It regulates the

uptake rate of some mineral nutrients through futile

cycles, or counterbalances the efflux of positive charges

[80]. The best documented anion efflux in root periph-

eral cells is the excretion of organic acids to the rhizo-

sphere. This is part of a process in which plants release

30% of the carbon fixed by photosynthesis [81].

Release of organic anions serves several functions. The

best established mechanism is the chelation of alumi-

num in acidic soils but it is also implicated in phos-

phate mobilization (see below). In addition, together

with many other compounds released in the rhizo-

sphere by plant roots, excreted organic anions may be

used as carbon sources by bacteria and fungi living in

the rhizophere and thus participate in the control of

the microorganism populations [82]. In the context of

symbiosis, organic acids are excreted to intracellular

symbiosomes to provide a carbon source to the

nitrogen fixing bacteria. An organic anion efflux

system belonging to the peptide transporter (PTR)

family putatively involved in this function has been

identified in alder nodules colonized by Frankia [83].

Anion efflux to the rhizosphere

Inorganic anion uptake is mediated by high and low

affinity root cell transporters specific for various nutri-

ents such as nitrate, phosphate and sulfate. Anion

efflux is also an important process in root peripheral

cells where it occurs along their electrochemical gradi-

ent and is probably mediated by anion channels or

other passive transport mechanisms [20]. Inorganic

anion efflux to the rhizosphere may be necessary to

regulate root cell pH by electrically counterbalancing

the efflux of protons or to regulate whole plant inor-

ganic anion uptake under stressful conditions. In addi-

tion, anion channels are important to control the

plasma membrane electrical potential, which is a key

parameter for nutrient acquisition [9]. The recent

molecular identification of the first nitrate efflux trans-

porter from root peripheral cells, NAXT1, allowed the



physiological function of root nitrate efflux to be

tested.

NAXT1 (nitrate excretion transporter 1) belongs to

the large NRT1 ⁄PTR family [84] and was identified by

a biochemical strategy performed on A. thaliana sus-

pension cells [80]. NAXT1 is targeted to the plasma

membrane and mainly expressed in cortical and epider-

mis cells of mature roots where it is responsible for

passive NO�3 excretion induced by medium acidificat-

ion. Root cell acidification occurs during anoxia in

flooded soils. Acid loading leads to a prolonged NO�3efflux associated with a decrease of root NO�3 content.

These responses are abolished in the naxt1 mutants

leading to the hypothesis that NAXT1 participates in

NO�3 excretion to counterbalance H+ excretion by

H+-ATPase required to attenuate cytosol acidification

[80,85]. The NAXT subfamily contains seven members

whose physiological functions mostly remain to be

explored. The discovery of NAXT1 suggests a diversity

of transport mechanisms within the PTR ⁄NRT family.

The best characterized member of this family,

NRT1.1, is a proton nitrate symporter [86]. Although

the transport mechanism used by NAXT1 has not yet

been fully characterized, it transports nitrate passively

along its electrochemical gradient, as a nitrate-perme-

able channel would.

Root peripheral cells also harbor R-type anion chan-

nels with high nitrate and sulfate permeability [87,88].

These channels, although not yet identified at the

molecular level, could serve functions similar to those

proposed for NAXT1: they could regulate the uptake

of nitrate and sulfate or counterbalance protons [80,89].

Aluminum toxicity is a serious problem on acidic

soils, which represent about 30% of arable land, world-

wide. In many species, Al3+ tolerance is associated with

increased excretion of organic acids, such as citrate,

malate or oxalate in Al3+-tolerant cultivars [90–92]. The

organic anion secretion occurs at the root tip, which is

most sensitive to Al3+ stress. Application of citrate or

oxalate can mitigate the toxic effect of Al3+ on root

growth. The importance of organic acid excretion for

Al3+ tolerance prompted several laboratories to con-

duct patch clamp studies on protoplasts isolated from

root tips of Al3+-sensitive or Al3+-tolerant cultivars of

wheat and maize [93–95]. Wheat and maize excrete

mostly malate or citrate, respectively. In studies on both

species, Al3+-activated anion conductances with chan-

nel properties were recorded in the plasma membrane of

root tip protoplasts. The channels are permeable to

organic acids with different selectivities. The Al3+-acti-

vated channels of wheat root tip cells are more perme-

able to malate than to chloride [96]. In maize, they are

also permeable to citrate [93,97]. The Al3+-activated

currents occur more frequently and are more strongly

activated in protoplasts from Al3+-tolerant cultivars of

wheat or maize compared with sensitive ones. The avail-

ability of cultivars with contrasting Al3+ tolerance

allowed the molecular identification of the chan-

nels ⁄ transporters responsible for the higher organic

anion efflux. In the case of wheat, cDNA library sub-

traction between two near-isogenic lines identified Ta-

ALMT1 [98]. TaALMT1 encodes a transmembrane

protein which defines a new protein family unique to

plants. Homologues of TaALMT1 have been character-

ized in Arabidopsis, barley and maize and are present in

all sequenced plant genomes [18]. ALMT1 is currently

one of the best characterized organic anion efflux chan-

nels, although other systems have been identified that

bring a major contribution to Al3+ tolerance in other

species (see below).

ALMT1 was shown to be an Al3+-activated malate

efflux protein able to confer Al3+ tolerance to plants

(Fig. 3) [98]. Subsequent studies performed on Xenopus

oocytes [99] and in tobacco cells [100] further eluci-

dated TaALMT1 transport mechanism, selectivity and

regulation. TaALMT1 is a malate-selective channel

generating low basal currents in the absence of Al3+,

whereas in the presence of Al3+ currents are strongly

enhanced. In tobacco cells, the permeability of

TaALMT1 is about 20-fold higher for malate than for

Cl) or NO�3 [100]. When expressed in Xenopus oocytes,

TaALMT1 is permeable not only to malate but also to

inorganic ions Cl), NO�3 or SO2�4 when external anion

concentrations are high [99]. These electrophysiological

results were confirmed in vivo in transgenic barley,

wheat and Arabidopsis where TaALMT1 expression

enhances malate efflux and thus Al3+ resistance

[92,98,101]. Nevertheless, not all ALMT1 homologues

are able to transport malate or are implicated in Al3+

tolerance. When ZmALMT1 is expressed in Xenopus

oocytes, the Al3+-activated currents are small and the

selectivity for organic acids (malate, citrate) over sev-

eral inorganic anions is poor [102]. This led to the

hypothesis that ZmALMT1 is rather involved in anion

homeostasis and mineral nutrition.

The analysis of TaALMT1 topology revealed that

the TaALMT1 polypeptide forms six transmembrane

a-helices with an N-terminal domain and a long C-ter-

minal domain both facing the extracellular side of the

plasma membrane (Fig. 3) [103]. Recent studies focus-

ing on the mechanism of Al3+-induced activation of

ALMT1 revealed a crucial importance for its C-termi-

nal domain. Al3+ enhances the activity of most

ALMT1 homologues identified in plants [98–100,104]

but the exact mechanism remains unclear. Al3+ activa-

tion is observed when ALMT1 is expressed in Xenopus



oocytes, suggesting that it is an intrinsic property of

ALMT1 protein [99]. Truncation of the C-terminal

domain of TaALMT1 leads to a loss of basal and

Al3+-activated transport activity, which is rescued by

grafting the Arabidopsis ALMT1 C-terminal domain

[105]. This domain is thus essential for the function of

TaALMT1 and its homologues. By mutating acidic

residues in the C-terminal domain, three residues

(E274, D275 and E284) were identified, which are spe-

cifically required for activation of ALMT1 transport

activity by Al3+ without affecting its basal activity.

This suggests that these residues participate in Al3+

binding domains (Fig. 3) [105].

Several lines of evidence suggest that TaALMT1

activity is regulated by phosphorylation. Application

of K252a, a protein kinase inhibitor, on wheat or

Arabidopsis roots reduces Al3+-activated malate efflux

[106,107]. The role of phosphorylation was also studied

by Ligaba et al. [108] on Xenopus oocytes expressing

TaALMT1. In this system, the application of protein

kinase antagonists inhibits basal and Al3+-activated

malate efflux. Moreover, the addition of a protein

kinase C (PKC) activator enhances TaALMT1 medi-

ated currents. In an attempt to identify phosphorylated

residues, the authors mutated six putative PKC phos-

phorylation sites located in the C-terminal domain of

TaALMT1. Mutation of threonine 323 to alanine

results in a significant increase of TaALMT1 activity;

in contrast, substitution of serine 384 to alanine greatly

reduces TaALMT1 activity [108]. This could indicate

that serine 384 needs to be phosphorylated before

Al3+ can activate AtALMT1 (Fig. 3). However, no

direct evidence that this site is phosphorylated in vivo

exists. Moreover, the phosphorylation sites were identi-

fied on the C-terminal domain localized outside of the

cell and extracellular protein kinases have not yet been

identified in plants. TaALMT1 topology is now subject

to controversy [12,92,103].

Whereas TaALMT1 locus is clearly important for

Al3+ tolerance in wheat, quantitative trait loci account-

ing for Al3+ tolerance in sorghum, barley and maize

identified genes coding organic anion transporters of the

multidrug and toxic compound exudation (MATE) fam-

ily [109–111]. In contrast to ALMT1 transporters, plant

MATE involved in Al3+ tolerance show a substrate

preference for citrate rather than malate. Although

canonic MATE involved in toxic compound efflux were

shown to function as a proton coupled anion efflux

Fig. 3. Structure and function of TaALMT1. TaALMT1 is an Al3+-activated malate transporter that confers Al3+ tolerance to wheat by excret-

ing malate to the rhizosphere to form non-toxic complex (malate–Al). TaALMT1 is composed of six transmembrane domains and a long

C-terminal domain. This domain displays three residues (E274, D275, E284) implicated in the Al3+-activated malate transport and suspected

to participate to Al3+ binding domains (represented by a question mark). Two other residues, T323, S384, were identified as putative phos-

phorylation sites regulating TaALMT1 activity.



pump, expression of ZmMATE1 from maize in Xenopus

oocytes triggers inward currents that are compatible

with organic anion efflux current through channels

[109,111]. More detailed characterization is required to

determine whether these MATE function as proton cou-

pled transporters, facilitators or constitute a new class

of anion channels in plants.

In addition to Al3+ tolerance, organic anion efflux

to the rhizosphere has also been implicated in phos-

phate nutrition [12]. Organic anion efflux is especially

relevant for plant families that do not form mycorrhiza

and grow on soils in which phosphate is poorly avail-

able. Organic acid secretion increases the availability

of phosphate tightly bound to soil particles through

ligand exchange. Specialized roots involved in organic

acid efflux were first described in Proteaceae. However,

the preferred experimental system to analyze this pro-

cess has been the cluster roots of white lupin [112].

Both the development of cluster roots and their

organic acid excretion are enhanced under phosphate

deficiency. Patch clamp analysis of protoplasts from

white lupin cluster roots revealed the presence of

citrate permeable channels [113]. However, these chan-

nels are not restricted to cells from cluster roots.

Another patch clamp study performed on A. thaliana

epidermal cells identified citrate permeable channels

that are found only in protoplasts isolated from phos-

phate starved roots [87]. This suggests that all root

epidermal cells may excrete citrate through citrate per-

meable channels and cluster roots may merely repre-

sent a way to increase the root–soil interface at which

efflux occurs. The molecular identity of the anion

channels mediating organic efflux under phosphate

starvation is not known yet. It will be interesting to

mine microarray or proteomic data from phosphate

starved roots for homologues of ALMT or MATE,

which are good candidates for this function.

Anion efflux to the xylem

Mineral nutrients are mostly translocated from roots

to shoots through the xylem. Translocation thus

requires the loading of mineral ions into the xylem.

Given their electrochemical gradient, many ions may

be excreted to the xylem sap through channel-mediated

mechanisms. In the case of potassium, a combination

of electrophysiological analyses and Arabidopsis molec-

ular genetics has demonstrated that SKOR, an out-

ward rectifying potassium channel of the Shaker

family, participates in potassium loading into the

xylem [114]. The electrochemical gradient for anions,

such as nitrate, is even more favorable to their loading

to the xylem through anion channels.

Several electrophysiological studies relying on the

ability to isolate and recognize protoplasts from stele

cells have identified anion channels in this cell type in

maize and barley [20,115–117]. These channels are

highly permeable to nitrate and are good candidates to

load this nutrient into the xylem. In addition, a pro-

ton-coupled nitrate transporter of the PTR family,

NRT1.5, was shown to participate in the loading of

nitrate to the xylem [118]. Nitrate transfer to the shoots

is reduced, but not completely abolished, in an nrt1-5

mutant. NRT1.5-mediated and anion-channel-mediated

nitrate loading systems may thus coexist. Interestingly,

one class of anion channel, similar to R-type currents,

present in stele cells is downregulated by the phytohor-

mone ABA [119]. This suggests that under drought

conditions, when the xylem flux is slowed down, coor-

dinated downregulation of anion loading to the xylem

occurs. The anion channels described by the patch

clamp technique in stele cells still await molecular iden-

tification. The discovery of the genes encoding these

channels will allow their importance for anion loading

in the xylem to be tested.

The major anions in the xylem sap are nitrate,

chloride, sulfate and phosphate. Citrate and malate,

however, are present at sub-millimolar concentrations

(0.1–0.5 mm). Based on physicochemical considerations

and the low pH of the xylem sap, it was proposed that

some metal cations, such as iron or aluminum, travel

through the xylem as complexes with citrate.

Mutations in FRD3 (ferric reductase deficient 3)

were recovered in screens for mutants that constitu-

tively activate iron-deficiency responses [120]. FRD3

encodes a MATE transporter showing high homology

to SbMATE, AtMATE, HvAACT1 and ZmMATE

that are involved in citrate excretion in sorghum, Ara-

bidopsis, barley and maize, respectively [121]. The

characterization of FRD3 both identified a pathway

for citrate excretion from the pericycle cells to the

xylem and supported the importance of iron–citrate

complexes for iron translocation from the roots to

the shoots [78] (see note added in proof). The closest

homologue of AtFRD3 in rice, OsFRDL1, is also

involved in iron translocation from roots to shoots

[79,122]. AtFRD3 and OsFRDL1 are expressed in

root pericycle cells. When expressed in Xenopus

oocytes, both AtFRD3 and OsFRDL1 facilitate cit-

rate efflux and both frdl1 and frd3 have decreased

xylem sap citrate concentrations. Interestingly, when

ectopically overexpressed in Arabidopsis, FRD3 con-

fers increased Al3+ tolerance [78]. These results indi-

cate that similar transport systems, the MATEs, are

involved in citrate excretion to the rhizosphere or to

the xylem sap. Depending on their expression pattern,



in epidermal or in pericycle cells, the citrate transport-

ing MATE are involved in Al3+ tolerance or iron

translocation, respectively. Although both processes

involve citrate fluxes along its electrochemical gradient

and the fluxes are further favored by the pH gradient

between the cytosol and the extracellular medium, it

remains to be determined whether the citrate trans-

porting MATE function as anion channels.

Conclusions

The knowledge in the field of plant anion channels has

grown very quickly in the last few years, with the iden-

tification of the genes underlying the rapid (R-type)

and the slow (S-type) anion channels of guard cells,

the analysis of their regulation and the determination

of the 3D structure of the slow anion channel, SLAC1.

It will be interesting to determine whether some plant

anion channels are encoded by other gene families. So

far, the best characterized member of the plant CLC

family, AtCLCa, works as a proton-coupled nitrate

transporter [6]. It is possible, however, that other plant

CLCs function as anion channels [18]. In addition, this

review describes membrane proteins of the MATE

family and of the PTR family (NAXT1) that mediate

passive anion fluxes and could also function as anion

channels [80,109,111].

The guard cell has proved to be a very efficient model

to identify plant ion channels [21]. Nevertheless, using

electrophysiological techniques, anion channel activity

can be recorded from virtually any cell type investigated.

These channel activities most often resemble S-type or

R-type guard cell anion currents. The identification of

two anion channel gene families, the SLAC1 family and

the ALMT1 family, should open the way to the identifi-

cation of the specific proteins or protein combinations

underlying most anion channel activities encountered in

plant cells. Mutations in the corresponding genes will

allow a more complete investigation of the physiological

role of anion channels in plant cells. Notably, it has been

proposed that anion channels participate in the control

of osmotic pressure and growth, in the massive anion

efflux triggered in response to pathogen attack or in the

generation of long-distance electrical signals [123,124].

It will soon be possible to test these hypotheses using

powerful genetic tools.

ALMT1, CLC and SLAC1 homologues are present

in genomes of all sequenced species [18]. With the

development of next generation high throughput

sequencing, homologous anion channel genes will be

identified in many more species. In particular, it should

allow the study of anion channels and their regulation

in plants where electrical signaling and fast movements

have been well characterized, such as Mimosa pudica

and carnivorous plants.

Finally, anion channels are clearly at the crossroad of

signaling, metabolism, nutrition and turgor regulation.

One of the future prospects will be to integrate their

function in the many networks in which they participate.

How do guard cell anion channels integrate with the

activity of other ion channels in this cell type to achieve

adequate turgor regulation under fluctuating condi-

tions? How do nitrate-permeable channels integrate with

nitrate transporters and nitrate assimilation pathways to

control nitrogen homeostasis? How do organic acid per-

meable channels integrate with organic acid transport-

ers, the primary carbon metabolism and the control of

cellular pH? Answering these questions will require a

detailed knowledge of anion channel genes and proteins

and their regulation and the building of quantitative

models integrating transmembrane and metabolic fluxes

together with cellular ion concentrations.

Acknowledgements

The work of M.J. and S.T. was supported by the Cen-

tre National de la Recherche Scientifique (CNRS) and

the Agence Nationale pour la Recherche (ANR-Nitra-

pool: grant number ANR-08-BLAN-0008-02). The

work of H.K. and K.L. was supported by ESF grant

7763, targeted funding theme SF0180071S07 and by

European Regional Fund (the Center of Excellence in

Environmental Adaptation).

Note added in proof

After this review was accepted, two important studies

were published. Meyer et al. characterized a malate

permeable channel belonging to the ALMT1 family in

the vacuolar membrane of guard cells. Roschzttardtz

et al. characterized the function of FRD3 malate efflux

transporter throughout plant development. [Meyer S,

Scholz-Starke J, De Angeli A, Kovermann P, Burla B,

Gambale F & Martinoia E (2011) Malate transport by

the vacuolar AtALMT6 channel in guard cells is

subject to multiple regulation. Plant J 67, 247–257.

Roschzttardtz H, Seguela-Arnaud M, Briat JF, Vert G

& Curie C (2011) The FRD3 citrate effluxer promotes

iron nutrition between symplastically disconnected tis-

sues throughout Arabidopsis development. Plant Cell

23, 2725–2737.]

References

1 Barbier-Brygoo H, Vinauger M, Colcombet J, Ephri-

tikhine G, Frachisse J & Maurel C (2000) Anion chan-



nels in higher plants: functional characterization,

molecular structure and physiological role. Biochim

Biophys Acta 1465, 199–218.

2 Neher E & Sakmann B (1976) Single-channel currents

recorded from membrane of denervated frog muscle

fibres. Nature 260, 799–802.

3 Dutzler R, Campbell EB, Cadene M, Chait BT &

MacKinnon R (2002) X-ray structure of a ClC chloride

channel at 3.0 A reveals the molecular basis of anion

selectivity. Nature 415, 287–294.

4 AGI (2000) Analysis of the genome sequence of the

flowering plant Arabidopsis thaliana. Nature 408,

796–815.

5 Hille B (1996) Ionic channels of excitable membranes,

second edition. Sinauer Associates Inc., Sunderland

MA, USA.

6 De Angeli A, Monachello D, Ephritikhine G, Frachisse

JM, Thomine S, Gambale F & Barbier-Brygoo H

(2006) The nitrate ⁄ proton antiporter AtCLCa mediates

nitrate accumulation in plant vacuoles. Nature 442,

939–942.

7 Emmerlich V, Linka N, Reinhold T, Hurth MA, Traub

M, Martinoia E & Neuhaus HE (2003) The plant

homolog to the human sodium ⁄ dicarboxylic cotrans-

porter is the vacuolar malate carrier. Proc Natl Acad

Sci USA 100, 11122–11126.

8 Roelfsema MR, Levchenko V & Hedrich R (2004)

ABA depolarizes guard cells in intact plants, through

a transient activation of R- and S-type anion channels.

Plant J 37, 578–588.

9 Schroeder JI (1995) Anion channels as central mecha-

nisms for signal transduction in guard cells and puta-

tive functions in roots for plant–soil interactions. Plant

Mol Biol 28, 353–361.

10 Garrill A, Tyerman SD & Findlay GP (1994) Ion

channels in the plasma membrane of protoplasts from

the halophytic angiosperm Zostera muelleri. J Membr

Biol 142, 381–393.

11 Jossier M, Kroniewicz L, Dalmas F, Le Thiec D,

Ephritikhine G, Thomine S, Barbier-Brygoo H, Filleur

S & Leonhardt N (2010) The Arabidopsis vacuolar

anion transporter, AtCLCc, is involved in the

regulation of stomatal movements and contributes to

salt tolerance. Plant J 64, 563–576.

12 Meyer S, De Angeli A, Fernie AR & Martinoia E

(2010) Intra- and extra-cellular excretion of carboxy-

lates. Trends Plant Sci 15, 40–47.

13 Hedrich R, Busch H & Raschke K (1990) Ca2+ and

nucleotide dependent regulation of voltage dependent

anion channels in the plasma membrane of guard cells.

EMBO J 9, 3889–3892.

14 Colcombet J, Thomine S, Guern J, Frachisse JM &

Barbier-Brygoo H (2001) Nucleotides provide a voltage-

sensitive gate for the rapid anion channel of Arabidopsis

hypocotyl cells. J Biol Chem 276, 36139–36145.

15 Geiger D, Scherzer S, Mumm P, Stange A, Marten I,

Bauer H, Ache P, Matschi S, Liese A, Al-Rasheid KA

et al. (2009) Activity of guard cell anion channel

SLAC1 is controlled by drought-stress signaling

kinase–phosphatase pair. Proc Natl Acad Sci USA 106,

21425–21430.

16 Hedrich R & Marten I (1993) Malate-induced feedback

regulation of plasma membrane anion channels could

provide a CO2 sensor to guard cells. EMBO J 12,

897–901.

17 Schmidt C, Schelle I, Liao YJ & Schroeder JI (1995)

Strong regulation of slow anion channels and abscisic

acid signaling in guard cells by phosphorylation and

dephosphorylation events. Proc Natl Acad Sci USA 92,

9535–9539.

18 Barbier-Brygoo H, De Angeli A, Filleur S, Frachisse

JM, Gambale F, Thomine S & Wege S (2011) Anion

channels ⁄ transporters in plants: from molecular bases

to regulatory networks. Annu Rev Plant Biol 62, 25–51.

19 De Angeli A, Thomine S, Frachisse JM, Ephritikhine

G, Gambale F & Barbier-Brygoo H (2007) Anion

channels and transporters in plant cell membranes.

FEBS Lett 581, 2367–2374.

20 Roberts SK (2006) Plasma membrane anion channels

in higher plants and their putative functions in roots.

New Phytol 169, 647–666.

21 Ward JM, Maser P & Schroeder JI (2009) Plant ion

channels: gene families, physiology, and functional

genomics analyses. Annu Rev Physiol 71, 59–82.

22 Melotto M, Underwood W, Koczan J, Nomura K &

He SY (2006) Plant stomata function in innate immu-

nity against bacterial invasion. Cell 126, 969–980.

23 Brosche M, Merilo E, Mayer F, Pechter P, Puzorjova

I, Brader G, Kangasjarvi J & Kollist H (2010) Natural

variation in ozone sensitivity among Arabidopsis

thaliana accessions and its relation to stomatal

conductance. Plant Cell Environ 33, 914–925.

24 Vahisalu T, Kollist H, Wang YF, Nishimura N, Chan

WY, Valerio G, Lamminmaki A, Brosche M, Moldau

H, Desikan R et al. (2008) SLAC1 is required for plant

guard cell S-type anion channel function in stomatal

signalling. Nature 452, 487–491.

25 Keller BU, Hedrich R & Raschke K (1989) Voltage-

dependent anion channels in the plasma membrane of

guard cells. Nature 341, 450–453.

26 Schroeder JI & Hagiwara S (1989) Cytosolic calcium

regulates ion channels in the plasma membrane of

Vicia faba guard cells. Nature 338, 427–430.

27 Meyer S, Mumm P, Imes D, Endler A, Weder B,

Al-Rasheid KA, Geiger D, Marten I, Martionia E &

Hedrich R (2010) AtALMT12 represents an R-type

anion channel required for stomatal movement in

Arabidopsis guard cells. Plant J 63, 1054–1062.

28 Negi J, Matsuda O, Nagasawa T, Oba Y, Takahashi

H, Kawai-Yamada M, Uchimiya H, Hashimoto M &



Iba K (2008) CO2 regulator SLAC1 and its homo-

logues are essential for anion homeostasis in plant

cells. Nature 452, 483–486.

29 Pei ZM, Kuchitsu K, Ward JM, Schwarz M & Schroe-

der JI (1997) Differential abscisic acid regulation of

guard cell slow anion channels in Arabidopsis wild-type

and abi1 and abi2 mutants. Plant Cell 9, 409–423.

30 Schroeder JI, Schmidt C & Sheaffer J (1993) Identifica-

tion of high-affinity slow anion channel blockers and

evidence for stomatal regulation by slow anion chan-

nels in guard cells. Plant Cell 5, 1831–1841.

31 Kim TH, Bohmer M, Hu H, Nishimura N & Schroe-

der JI (2010) Guard cell signal transduction network:

advances in understanding abscisic acid, CO2, and

Ca2+ signaling. Annu Rev Plant Biol 61, 561–591.

32 Linder B & Raschke K (1992) A slow anion channel

in guard cells, activating at large hyperpolarization,

may be principal for stomatal closing. FEBS Lett 313,

27–30.

33 Schroeder JI & Keller BU (1992) Two types of anion

channel currents in guard cells with distinct voltage

regulation. Proc Natl Acad Sci USA 89, 5025–5029.

34 Raschke K, Shabahang M & Wolf R (2003) The slow

and the quick anion conductance in whole guard cells:

their voltage-dependent alternation, and the modula-

tion of their activities by abscisic acid and CO2. Planta

217, 639–650.

35 Kangasjarvi J, Jaspers P & Kollist H (2005) Signalling

and cell death in ozone-exposed plants. Plant Cell

Environ 28, 1021–1036.

36 Negi J, Hashimoto M & Iba K (2005) Characterization

of CO2-insensitive Arabidopsis mutant cdi3. Plant Cell

Physiol 46, S176.

37 Saji S, Bathula S, Kubo A, Tamaoki M, Kanna M,

Aono M, Nakajima N, Nakaji T, Takeda T, Asayama

M et al. (2008) Disruption of a gene encoding

C4-dicarboxylate transporter-like protein increases

ozone sensitivity through deregulation of the stomatal

response in Arabidopsis thaliana. Plant Cell Physiol 49,

2–10.

38 Lee SC, Lan W, Buchanan BB & Luan S (2009)

A protein kinase–phosphatase pair interacts with an

ion channel to regulate ABA signaling in plant guard

cells. Proc Natl Acad Sci USA 106, 21419–21424.

39 Chen YH, Hu L, Punta M, Bruni R, Hillerich B, Kloss

B, Rost B, Love J, Siegelbaum SA & Hendrickson WA

(2010) Homologue structure of the SLAC1 anion

channel for closing stomata in leaves. Nature 467, 1074–

1080.

40 Schmidt C & Schroeder JI (1994) Anion selectivity of

slow anion channels in the plasma membrane of guard

cells (large nitrate permeability). Plant Physiol 106,

383–391.

41 Vahisalu T, Puzorjova I, Brosche M, Valk E, Lepiku

M, Moldau H, Pechter P, Wang YS, Lindgren O,

Salojarvi J et al. (2010) Ozone-triggered rapid stomatal

response involves the production of reactive oxygen

species, and is controlled by SLAC1 and OST1. Plant

J 62, 442–453.

42 Chen ZH, Hills A, Lim CK & Blatt MR (2010)

Dynamic regulation of guard cell anion channels by

cytosolic free Ca2+ concentration and protein phos-

phorylation. Plant J 61, 816–825.

43 Siegel RS, Xue S, Murata Y, Yang Y, Nishimura N,

Wang A & Schroeder JI (2009) Calcium elevation-

dependent and attenuated resting calcium-dependent

abscisic acid induction of stomatal closure and abscisic

acid-induced enhancement of calcium sensitivities of

S-type anion and inward-rectifying K channels in

Arabidopsis guard cells. Plant J 59, 207–220.

44 Mori IC, Murata Y, Yang Y, Munemasa S, Wang YF,

Andreoli S, Tiriac H, Alonso JM, Harper JF, Ecker

JR et al. (2006) CDPKs CPK6 and CPK3 function in

ABA regulation of guard cell S-type anion- and Ca2+-

permeable channels and stomatal closure. PLoS Biol 4,

e327.

45 Geiger D, Scherzer S, Mumm P, Marten I, Ache P,

Matschi S, Liese A, Wellmann C, Al-Rasheid KAS,

Grill E et al. (2010) Guard cell anion channel SLAC1 is

regulated by CDPK protein kinases with distinct

Ca2+ affinities. Proc Natl Acad Sci USA 107, 8023–

8028.

46 Hu H, Boisson-Dernier A, Israelsson-Nordstrom M,

Bohmer M, Xue S, Ries A, Godoski J, Kuhn JM &

Schroeder JI (2010) Carbonic anhydrases are upstream

regulators of CO2-controlled stomatal movements in

guard cells. Nat Cell Biol 12, 87–93.

47 Xue S, Hu H, Ries A, Merilo E, Kollist H & Schroe-

der JI (2011) Central functions of bicarbonate in S-type

anion channel activation and OST1 protein kinase in

CO2 signal transduction in guard cell. EMBO J 30,

1645–1658.

48 Hashimoto M, Negi J, Young J, Israelsson M, Schroe-

der JI & Iba K (2006) Arabidopsis HT1 kinase controls

stomatal movements in response to CO2. Nat Cell Biol

8, 391–397.

49 Ma Y, Szostkiewicz I, Korte A, Moes D, Yang Y,

Christmann A & Grill E (2009) Regulators of PP2C

phosphatase activity function as abscisic acid sensors.

Science 324, 1064–1068.

50 Park SY, Fung P, Nishimura N, Jensen DR, Fujii H,

Zhao Y, Lumba S, Santiago J, Rodrigues A, Chow TF

et al. (2009) Abscisic acid inhibits type 2C protein

phosphatases via the PYR ⁄PYL family of START

proteins. Science 324, 1068–1071.

51 Belin C, de Franco PO, Bourbousse C, Chaignepain S,

Schmitter JM, Vavasseur A, Giraudat J, Barbier-Bry-

goo H & Thomine S (2006) Identification of features

regulating OST1 kinase activity and OST1 function in

guard cells. Plant Physiol 141, 1316–1327.



52 Vlad F, Rubio S, Rodrigues A, Sirichandra C, Belin C,

Robert N, Leung J, Rodriguez PL, Lauriere C & Mer-

lot S (2009) Protein phosphatases 2C regulate the acti-

vation of the Snf1-related kinase OST1 by abscisic acid

in Arabidopsis. Plant Cell 21, 3170–3184.

53 Umezawa T, Sugiyama N, Mizoguchi M, Hayashi S,

Myouga F, Yamaguchi-Shinozaki K, Ishihama Y,

Hirayama T & Shinozaki K (2009) Type 2C protein

phosphatases directly regulate abscisic acid-activated

protein kinases in Arabidopsis. Proc Natl Acad Sci

USA 106, 17588–17593.

54 Mustilli AC, Merlot S, Vavasseur A, Fenzi F & Girau-

dat J (2002) Arabidopsis OST1 protein kinase mediates

the regulation of stomatal aperture by abscisic acid

and acts upstream of reactive oxygen species produc-

tion. Plant Cell 14, 3089–3099.

55 Xie X, Wang Y, Williamson L, Holroyd GH, Tagliavia

C, Murchie E, Theobald J, Knight MR, Davies WJ,

Leyser HM et al. (2006) The identification of genes

involved in the stomatal response to reduced atmo-

spheric relative humidity. Curr Biol 16, 882–887.

56 Thomine S & Barbier-Brygoo H (2010) Structural

biology: a peep through anion channels. Nature 467,

1058–1059.

57 Loque D, Lalonde S, Looger LL, von Wiren N &

Frommer WB (2007) A cytosolic trans-activation

domain essential for ammonium uptake. Nature 446,

195–198.

58 Geiger D, Maierhofer T, Al-Rasheid KA, Scherzer S,

Mumm P, Liese A, Ache P, Wellmann C, Marten I, Grill

E et al. (2011) Stomatal closure by fast abscisic acid sig-

naling is mediated by the guard cell anion channel

SLAH3 and the receptor RCAR1. Sci Signal 4, ra32.

59 Sasaki T, Mori IC, Furuichi T, Munemasa S, Toyooka

K, Matsuoka K, Murata Y & Yamamoto Y (2010)

Closing plant stomata requires a homolog of an alumi-

num-activated malate transporter. Plant Cell Physiol

51, 354–365.

60 Pei ZM, Murata Y, Benning G, Thomine S, Klusener B,

Allen GJ, Grill E & Schroeder JI (2000) Calcium chan-

nels activated by hydrogen peroxide mediate abscisic

acid signalling in guard cells. Nature 406, 731–734.

61 Hoekenga OA, Maron LG, Pineros MA, Cancado

GMA, Shaff J, Kobayashi Y, Ryan PR, Dong B,

Delhaize E, Sasaki T et al. (2006) AtALMT1, which

encodes a malate transporter, is identified as one of

several genes critical for aluminum tolerance in Arabid-

opsis. Proc Natl Acad Sci USA 103, 9738–9743.

62 Goh CH, Kinoshita T, Oku T & Shimazaki K (1996)

Inhibition of blue light-dependent H+ pumping by

abscisic acid in Vicia guard-cell protoplasts. Plant

Physiol 111, 433–440.

63 Merlot S, Leonhardt N, Fenzi F, Valon C, Costa M,

Piette L, Vavasseur A, Genty B, Boivin K, Muller A

et al. (2007) Constitutive activation of a plasma

membrane H+-ATPase prevents abscisic acid-mediated

stomatal closure. EMBO J 26, 3216–3226.

64 Hamilton DW, Hills A, Kohler B & Blatt MR (2000)

Ca2+ channels at the plasma membrane of stomatal

guard cells are activated by hyperpolarization and

abscisic acid. Proc Natl Acad Sci USA 97, 4967–

4972.

65 Kinoshita T & Shimazaki K (1999) Blue light activates

the plasma membrane H+-ATPase by phosphorylation

of the C-terminus in stomatal guard cells. EMBO J 18,

5548–5558.

66 Outlaw WH & Lowry OH (1977) Organic acid and

potassium accumulation in guard cells during stomatal

opening. Proc Natl Acad Sci USA 74, 4434–4438.

67 Raschke K & Schnabl H (1978) Availability of chloride

affects the balance between potassium chloride and

potassium malate in guard cells of Vicia faba L. Plant

Physiol 62, 84–87.

68 Lee M, Choi Y, Burla B, Kim YY, Jeon B, Maeshima

M, Yoo JY, Martinoia E & Lee Y (2008) The ABC trans-

porter AtABCB14 is a malate importer and modulates

stomatal response to CO2.Nat Cell Biol 10, 1217–1223.

69 Gao XQ, Li CG, Wei PC, Zhang XY, Chen J & Wang

XC (2005) The dynamic changes of tonoplasts in guard

cells are important for stomatal movement in Vicia

faba. Plant Physiol 139, 1207–1216.

70 Pei ZM, Ward JM, Harper JF & Schroeder JI (1996)

A novel chloride channel in Vicia faba guard cell

vacuoles activated by the serine ⁄ threonine kinase,

CDPK. EMBO J 15, 6564–6574.

71 Gaedeke N, Klein M, Kolukisaoglu U, Forestier C,

Muller A, Ansorge M, Becker D, Mamnun Y, Kuchler

K, Schulz B et al. (2001) The Arabidopsis thaliana

ABC transporter AtMRP5 controls root development

and stomata movement. EMBO J 20, 1875–1887.

72 Nagy R, Grob H, Weder B, Green P, Klein M, Frelet-

Barrand A, Schjoerring JK, Brearley C & Martinoia E

(2009) The Arabidopsis ATP-binding cassette protein

AtMRP5 ⁄AtABCC5 is a high affinity inositol hexakis-

phosphate transporter involved in guard cell signaling

and phytate storage. J Biol Chem 284, 33614–33622.

73 Leonhardt N, Marin E, Vavasseur A & Forestier C

(1997) Evidence for the existence of a sulfonylurea-

receptor-like protein in plants: modulation of stomatal

movements and guard cell potassium channels by

sulfonylureas and potassium channel openers. Proc

Natl Acad Sci USA 94, 14156–14161.

74 Leonhardt N, Vavasseur A & Forestier C (1999) ATP

binding cassette modulators control abscisic acid-regu-

lated slow anion channels in guard cells. Plant Cell 11,

1141–1152.

75 Suh SJ, Wang YF, Frelet A, Leonhardt N, Klein M,

Forestier C, Mueller-Roeber B, Cho MH, Martinoia E

& Schroeder JI (2007) The ATP binding cassette trans-

porter AtMRP5 modulates anion and calcium channel



activities in Arabidopsis guard cells. J Biol Chem 282,

1916–1924.

76 Lemtiri-Chlieh F, MacRobbie EA, Webb AA, Manison

NF, Brownlee C, Skepper JN, Chen J, Prestwich GD

& Brearley CA (2003) Inositol hexakisphosphate mobi-

lizes an endomembrane store of calcium in guard cells.

Proc Natl Acad Sci USA 100, 10091–10095.

77 Andrews M (1986) The partitioning of nitrate assimila-

tion between root and shoot of higher plants. Plant

Cell Environ 9, 511–519.

78 Durrett TP, Gassmann W & Rogers EE (2007) The

FRD3-mediated efflux of citrate into the root vascula-

ture is necessary for efficient iron translocation. Plant

Physiol 144, 197–205.

79 Yokosho K, Yamaji N, Ueno D, Mitani N & Ma JF

(2009) OsFRDL1 is a citrate transporter required for

efficient translocation of iron in rice. Plant Physiol 149,

297–305.

80 Segonzac C, Boyer JC, Ipotesi E, Szponarski W,

Tillard P, Touraine B, Sommerer N, Rossignol M &

Gibrat R (2007) Nitrate efflux at the root plasma

membrane: identification of an Arabidopsis excretion

transporter. Plant Cell 19, 3760–3777.

81 Badri DV & Vivanco JM (2009) Regulation and

function of root exudates. Plant Cell Environ 32,

666–681.

82 Rudrappa T, Czymmek KJ, Pare PW & Bais HP

(2008) Root-secreted malic acid recruits beneficial soil

bacteria. Plant Physiol 148, 1547–1556.

83 Jeong J, Suh S, Guan C, Tsay YF, Moran N, Oh CJ,

An CS, Demchenko KN, Pawlowski K & Lee Y (2004)

A nodule-specific dicarboxylate transporter from alder

is a member of the peptide transporter family. Plant

Physiol 134, 969–978.

84 Tsay YF, Chiu CC, Tsai CB, Ho CH & Hsu PK

(2007) Nitrate transporters and peptide transporters.

FEBS Lett 581, 2290–2300.

85 Felle HH (2005) pH regulation in anoxic plants. Ann

Bot 96, 519–532.

86 Tsay YF, Schroeder JI, Feldmann KA & Crawford

NM (1993) The herbicide sensitivity gene CHL1 of

Arabidopsis encodes a nitrate-inducible nitrate

transporter. Cell 72, 705–713.

87 Diatloff E, Roberts M, Sanders D & Roberts SK

(2004) Characterization of anion channels in the plasma

membrane of Arabidopsis epidermal root cells and the

identification of a citrate-permeable channel induced by

phosphate starvation. Plant Physiol 136, 4136–4149.

88 Kiegle E, Gilliham M, Haseloff J & Tester M (2000)

Hyperpolarisation-activated calcium currents found

only in cells from the elongation zone of Arabidopsis

thaliana roots. Plant J 21, 225–229.

89 Santi S & Schmidt W (2009) Dissecting iron deficiency-

induced proton extrusion in Arabidopsis roots. New

Phytol 183, 1072–1084.

90 Delhaize E, Craig S, Beaton CD, Bennet RJ, Jagadish

VC & Randall PJ (1993) Aluminum tolerance in wheat

(Triticum aestivum L.). 1. Uptake and distribution of

aluminum in root apices. Plant Physiol 103, 685–693.

91 Ma JF, Ryan PR & Delhaize E (2001) Aluminium

tolerance in plants and the complexing role of organic

acids. Trends Plant Sci 6, 273–278.

92 Ryan PR, Tyerman SD, Sasaki T, Furuichi T, Yamam-

oto Y, Zhang WH & Delhaize E (2011) The identifica-

tion of aluminium-resistance genes provides

opportunities for enhancing crop production on acid

soils. J Exp Bot 62, 9–20.

93 Kollmeier M, Dietrich P, Bauer CS, Horst WJ & Hed-

rich R (2001) Aluminum activates a citrate-permeable

anion channel in the aluminum-sensitive zone of the

maize root apex. A comparison between an aluminum-

sensitive and an aluminum-resistant cultivar. Plant

Physiol 126, 397–410.

94 Pineros MA & Kochian LV (2001) A patch-clamp

study on the physiology of aluminum toxicity and

aluminum tolerance in maize. Identification and char-

acterization of Al3+-induced anion channels. Plant

Physiol 125, 292–305.

95 Ryan PR, Skerrett M, Findlay GP, Delhaize E &

Tyerman SD (1997) Aluminum activates an anion

channel in the apical cells of wheat roots. Proc Natl

Acad Sci USA 94, 6547–6552.

96 Zhang WH, Ryan PR & Tyerman SD (2001) Malate-

permeable channels and cation channels activated by

aluminum in the apical cells of wheat roots. Plant

Physiol 125, 1459–1472.

97 Pineros MA, Magalhaes JV, Carvalho Alves VM &

Kochian LV (2002) The physiology and biophysics of

an aluminum tolerance mechanism based on root

citrate exudation in maize. Plant Physiol 129, 1194–

1206.

98 Sasaki T, Yamamoto Y, Ezaki B, Katsuhara M, Ahn

SJ, Ryan PR, Delhaize E & Matsumoto H (2004) A

wheat gene encoding an aluminum-activated malate

transporter. Plant J 37, 645–653.

99 Pineros MA, Cancado GM & Kochian LV (2008)

Novel properties of the wheat aluminum tolerance

organic acid transporter (TaALMT1) revealed by

electrophysiological characterization in Xenopus

oocytes: functional and structural implications.

Plant Physiol 147, 2131–2146.

100 Zhang WH, Ryan PR, Sasaki T, Yamamoto Y, Sulli-

van W & Tyerman SD (2008) Characterization of the

TaALMT1 protein as an Al3+-activated anion channel

in transformed tobacco (Nicotiana tabacum L.) cells.

Plant Cell Physiol 49, 1316–1330.

101 Pereira JF, Zhou G, Delhaize E, Richardson T, Zhou

M & Ryan PR (2010) Engineering greater aluminium

resistance in wheat by over-expressing TaALMT1. Ann

Bot 106, 205–214.



102 Pineros MA, Cancado GM, Maron LG, Lyi SM,

Menossi M & Kochian LV (2008) Not all ALMT1-

type transporters mediate aluminum-activated organic

acid responses: the case of ZmALMT1 – an anion-

selective transporter. Plant J 53, 352–367.

103 Motoda H, Sasaki T, Kano Y, Ryan PR, Delhaize E,

Matsumoto H & Yamamoto Y (2007) The membrane

topology of ALMT1, an aluminum-activated malate

transport protein in wheat (Triticum aestivum). Plant

Signal Behav 2, 467–472.

104 Yamaguchi M, Sasaki T, Sivaguru M, Yamamoto Y,

Osawa H, Ahn SJ & Matsumoto H (2005) Evidence

for the plasma membrane localization of Al-activated

malate transporter (ALMT1). Plant Cell Physiol 46,

812–816.

105 Furuichi T, Sasaki T, Tsuchiya Y, Ryan PR, Delhaize

E & Yamamoto Y (2010) An extracellular hydrophilic

carboxy-terminal domain regulates the activity of

TaALMT1, the aluminum-activated malate transport

protein of wheat. Plant J 64, 47–55.

106 Kobayashi Y, Hoekenga OA, Itoh H, Nakashima M,

Saito S, Shaff JE, Maron LG, Pineros MA, Kochian

LV & Koyama H (2007) Characterization of AtAL-

MT1 expression in aluminum-inducible malate release

and its role for rhizotoxic stress tolerance in

Arabidopsis. Plant Physiol 145, 843–852.

107 Osawa H & Matsumoto H (2001) Possible involvement

of protein phosphorylation in aluminum-responsive

malate efflux from wheat root apex. Plant Physiol 126,

411–420.

108 Ligaba A, Kochian L & Pineros M (2009) Phosphory-

lation at S384 regulates the activity of the TaALMT1

malate transporter that underlies aluminum resistance

in wheat. Plant J 60, 411–423.

109 Furukawa J, Yamaji N, Wang H, Mitani N, Murata

Y, Sato K, Katsuhara M, Takeda K & Ma JF (2007)

An aluminum-activated citrate transporter in barley.

Plant Cell Physiol 48, 1081–1091.

110 Magalhaes JV, Liu J, Guimaraes CT, Lana UG, Alves

VM, Wang YH, Schaffert RE, Hoekenga OA, Pineros

MA, Shaff JE et al. (2007) A gene in the multidrug

and toxic compound extrusion (MATE) family confers

aluminum tolerance in sorghum. Nat Genet 39,

1156–1161.

111 Maron LG, Pineros MA, Guimaraes CT, Magalhaes

JV, Pleiman JK, Mao C, Shaff J, Belicuas SN &

Kochian LV (2010) Two functionally distinct members

of the MATE (multi-drug and toxic compound extru-

sion) family of transporters potentially underlie two

major aluminum tolerance QTLs in maize. Plant J 61,

728–740.

112 Neumann G & Martinoia E (2002) Cluster roots – an

underground adaptation for survival in extreme

environments. Trends Plant Sci 7, 162–167.

113 Zhang WH, Ryan PR & Tyerman SD (2004) Citrate-

permeable channels in the plasma membrane of

cluster roots from white lupin. Plant Physiol 136,

3771–3783.

114 Gaymard F, Pilot G, Lacombe B, Bouchez D,

Bruneau D, Boucherez J, Michaux-Ferriere N,

Thibaud JB & Sentenac H (1998) Identification and

disruption of a plant shaker-like outward channel

involved in K+ release into the xylem sap. Cell 94,

647–655.

115 Kohler B & Raschke K (2000) The delivery of salts to

the xylem. Three types of anion conductance in the

plasmalemma of the xylem parenchyma of roots of

barley. Plant Physiol 122, 243–254.

116 Kohler B, Wegner LH, Osipov V & Raschke K (2002)

Loading of nitrate into the xylem: apoplastic nitrate

controls the voltage dependence of X-QUAC, the main

anion conductance in xylem-parenchyma cells of barley

roots. Plant J 30, 133–142.

117 Wegner LH & Raschke K (1994) Ion channels in the

xylem parenchyma of barley roots: a procedure to

isolate protoplasts from this tissue and a patch-clamp

exploration of salt passageways into xylem vessels.

Plant Physiol 105, 799–813.

118 Lin SH, Kuo HF, Canivenc G, Lin CS, Lepetit M,

Hsu PK, Tillard P, Lin HL, Wang YY, Tsai CB et al.

(2008) Mutation of the Arabidopsis NRT1.5 nitrate

transporter causes defective root-to-shoot nitrate

transport. Plant Cell 20, 2514–2528.

119 Gilliham M & Tester M (2005) The regulation of anion

loading to the maize root xylem. Plant Physiol 137,

819–828.

120 Yi Y & Guerinot ML (1996) Genetic evidence that

induction of root Fe(III) chelate reductase activity is

necessary for iron uptake under iron deficiency. Plant J

10, 835–844.

121 Rogers EE & Guerinot ML (2002) FRD3, a member

of the multidrug and toxin efflux family, controls iron

deficiency responses in Arabidopsis. Plant Cell 14,

1787–1799.

122 Inoue H, Mizuno D, Takahashi M, Nakanishi H,

Mori S & Nishizawa N (2004) A rice FRD3-like

(OsFRDL1) gene is expressed in the cells involved in

long-distance transport. Soil Sci Plant Nutr 50, 1133–

1140.

123 Diatloff E, Peyronnet R, Colcombet J, Thomine S,

Barbier-Brygoo H & Frachisse JM (2010) R type anion

channel: a multifunctional channel seeking its molecu-

lar identity. Plant Signal Behav 5, 1349–1359.

124 Thomine S, Lelievre F, Boufflet M, Guern J &

Barbier-Brygoo H (1997) Anion-channel blockers

interfere with auxin responses in dark-grown

Arabidopsis hypocotyls. Plant Physiol 115,

533–542.



Date post:	08-May-2023
Category:	Documents
Upload:	ut-ee
View:	1 times
Download:	0 times

Anion channels in plant cells

Documents