Download The essential role of anionic transport in plant cells

Journal of Experimental Botany, Vol. 62, No. 7, pp. 2273–2298, 2011
doi:10.1093/jxb/err036
REVIEW PAPER
The essential role of anionic transport in plant cells: the
pollen tube as a case study
Bárbara Tavares1,2, Patrı́cia Domingos1,2, Pedro Nuno Dias1,2, José A. Feijó1,2,* and Ana Bicho1
1
2
Instituto Gulbenkian de Ciência, 2780-156 Oeiras, Portugal
Universidade de Lisboa, Faculdade de Ciências, Departamento de Biologia Vegetal, Campo Grande, C2, 1749-016 Lisboa, Portugal
* To whom correspondence should be addressed: Email: [email protected]
Received 19 November 2010; Revised 24 January 2011; Accepted 25 January 2011
Abstract
Plasma membrane anion transporters play fundamental roles in plant cell biology, especially in stomatal closure and
nutrition. Notwithstanding, a lot is still unknown about the specific function of these transporters, their specific
localization, or molecular nature. Here the fundamental roles of anionic transport in plant cells are reviewed. Special
attention will be paid to them in the control of pollen tube growth. Pollen tubes are extreme examples of cellular
polarity as they grow exclusively in their apical extremity. Their unique cell biology has been extensively exploited for
fundamental understanding of cellular growth and morphogenesis. Non-invasive methods have demonstrated that
tube growth is governed by different ion fluxes, with different properties and distribution. Not much is known about
the nature of the membrane transporters responsible for anionic transport and their regulation in the pollen tube.
Recent data indicate the importance of chloride (Cl–) transfer across the plasma membrane for pollen germination
and pollen tube growth. A general overview is presented of the well-known accumulated data in terms of biophysical
and functional characterization, transcriptomics, and genomic description of pollen ionic transport, and the various
controversies around the role of anionic fluxes during pollen tube germination, growth, and development. It is
concluded that, like all other plant cells so far analysed, pollen tubes depend on anion fluxes for a number of
fundamental homeostatic properties.
Key words: Anionic transport, apical growth, plants, pollen tube.
Introduction
Free anions are ubiquitously present in the cells of all
organisms and play a variety of chemical and regulatory
roles in cellular physiology. The cell content is essentially an
aqueous solution, and anions are always needed to maintain
the bulk electroneutrality, as many fundamental physiological processes require the presence of nanomolar to
milimolar concentrations of various cations. In natural
systems, the cationic cytosolic composition generally
exceeds the anionic and the excess positive charge is
balanced by the intracellular macromolecules (i.e. proteins),
as well as small organic anions such as citrate, phosphate
–
(PO3–
4 ), nitrate (NO3), and others, according to the type of
cell. Evolutionary processes, on the other hand, have led to
the development of specific proteins that interact with or
transport cations and anions leading to the regulation of
cellular osmolarity in most eukaryotic cells of higher
organisms. In many cells, the osmolarity is kept at a fairly
constant value. In others, such as the guard cells involved in
the opening and closure of stomata, osmolarity values vary
according to the physiological needs and the processes
involved are tightly regulated (e.g. Siegel et al., 2009).
Moreover, the common machinery involved includes the
transport of ions and other solutes across the plasma
membrane by means of membrane transporters, followed
Abbreviations: A-9-C, anthracene-9-carboxylic acid; ABA, abscisic acid; ARACs, Arabidopsis root anion channels, depolarization-activated; DIDS,
4,4-diisothiocyanostilbene-2,2-disulphonic acid; GCAC1, guard cell anion channel 1; MFS, major facilitator superfamily; NPPB, 5-nitro-2-(3-phenylpropylamino)benzoic acid; ORAC, outward-rectifying anion channel; VMAL, vacuole malate channel; X-IRAC, X-inward-rectifying anion conductance; X-QUAC, X-quicly activating
anion conductance.
ª The Author [2011]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
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2274 | Tavares et al.
by the passive flow of water according to the osmotic
gradient which, in turn, builds up the hydrostatic or turgor
pressure needed for cell volume change. Evolutionarily, that
has become a major fundamental homeostatic mechanism
in plant cells, this pressure being largely counteracted by the
cell wall. It is this control and tolerance of the hydrostatic
pressure that is essential for many physiological processes,
including stomatal opening, cell growth, reproduction, and
various transport processes across membranes. Further3–
–
more, anions such as sulphate (SO2–
4 ), PO4 , NO3, malate,
and citrate all have essential roles in plant physiology and
nutrition. The organic anions malate and citrate are
intermediates of the Calvin and Krebs cycles, respectively,
representing potential energy substrate molecules. These
also participate in the regulation of the uptake of other
ions, namely in the uptake of phosphorus (Lopez-Bucio
et al., 2000) and iron (Durrett et al., 2007), and in resistance
to aluminium (Al3+) in acidic soils (Pineros and Kochian,
2001; Ryan et al., 2009).
This review deals with the proposed roles of anions and
anionic transport in different aspects of plant cellular
physiology and regulation. Emphasis will be made on
passive transport through anionic channels, and on the
pollen tube system, which is progressively being established
as one of the most interesting plant models in terms of ionic
fluxes and intracellular gradients (Becker and Feijo, 2007;
Michard et al., 2009). A model encompassing the full
description of anionic transport in the pollen tube is
proposed and discussed.
Anionic transport: a comprehensive
overview
In spite of all the efforts put into molecular cloning, most of
the many Cl–-selective channels from which anionic currents
have been recorded in different cell types have yet to be
molecularly identified or linked with a cellular function.
Typically, for comparative purposes, physiologists opt to
classify
channels
according
to
their
functional
characteristics. Anionic currents are usually analysed by
voltage–clamp techniques, and the channels are
characterized on the basis of their ionic selectivity,
conductance, dependence of gating on membrane potential,
and the kinetics of their opening and closing states. Anion
currents that are indistinguishable by the above-mentioned
electrophysiological criteria can sometimes be identified
pharmacologically (e.g. use of more or less specific inhibitors), by activation through extracellular agonists (e.g. the
nicotinic acetylcholine receptor, which opens in response to
the binding of acetylcholine released from a presynaptic
nerve terminus), and by their modulation through intracellular regulatory molecules (e.g. regulation via intracellular
messengers
such
as
the
intracellular
calcium
concentration—[Ca2+]in—or intracellular cGMP). The functional criteria described for channels are not mutually
exclusive, and should not be taken as a direct indication of
the putative molecular families they may belong to, since
they may present several of these features. Because of the
diversity of channels anticipated on the basis of functional
criteria, one should additionally rely on a complementary
molecular biology approach to ultimately provide a definitive classification of channels based on structural characteristics, such as amino acid sequence homology and the kinds
of subunits of which they are composed.
What makes plant ion transport unique?
In higher plants, physiological processes such as mineral
nutrition, carbon and nitrogen metabolism, and more
specifically growth and development, strongly depend on
solute and water fluxes across the plasma membrane,
tonoplast and other endomembranes. Therefore, anionic
membrane transporters have been associated with various
processes in plants such as stomatal closure, hormone
signalling, membrane excitability, cellular osmoregulation,
growth regulation, and anionic nutrition. Because of their
importance, anion transporters are present in all plant cell
membranes, including the plasma membrane, tonoplast,
endoplasmic reticulum, mitochondria, and chloroplasts
(reviewed in Barbier-Brygoo et al., 2000; Roberts, 2006;
de Angeli et al., 2007).
The presence of the wall has hindered electrophysiological studies, which require free access to the plasma
membrane. The use of cell protoplasts, obtained by either
enzymatic digestion or laser microsurgery methods (e.g.
Fairley et al., 1991; Henriksen and Assmann, 1997;
Gehwolf et al., 2002), permitted the application of the
patch–clamp technique to several cellular types and
functional information on transport systems present in the
plasma membrane to be obtained. In general, the resting
potential of the majority of plant cells is more negative
than –100 mV. Therefore the command voltages required
for voltage activation of plant ion channels are usually
more negative than those used to study animal cells.
Patch–clamp methods were also extended to the vacuole,
and during the past two decades a variety of channels,
carriers, and pumps have been identified in both the
plasmalemma and the tonoplast of plant cells. The kinetics
of plant channels, although similar, tends to be slower
than those in excitable cells in animals (Hedrich, 2009;
Marten et al., 2010).
In plant cells, the vacuole occupies most of the cellular
volume and contains the bulk of the cell’s solutes. Ionic
concentrations in the cytosol and in the vacuole are
controlled by active and passive transport processes across
membrane proteins. Typically, transport across the plasma
membrane is energized by one primary active transport
system coupled to ATP hydrolysis—a P-type H+-ATPase.
A variety of carriers in the membrane are then able to
couple the transmembrane movement of one or two
protons back into the cell (down their electrochemical
potential) to the uphill movement of other ions or organic
substrates. In a similar way, the tonoplast regulates the
transport of ions from the cytosol and the vacuole. The
vacuole V-ATPase is an electrogenic pump that transports
Anionic transport in plant cells | 2275
protons from the cytosol to the vacuole, and generates
a proton-motive force across the tonoplast which also
drives secondary transport across this barrier. The movement of protons out of the cell and into the vacuole keeps
the cytosol pH at a value of ;7.0, while the vacuole sap
has typical values of 5.5. The bulk electroneutrality of the
vacuole sap is maintained by different anions, which are
transported from the cytosol through transport systems,
and in addition makes it possible for the V-pump to
generate a large concentration gradient of protons across
the tonoplast.
In addition to the secondary transporters, the electrochemical potential across these membranes is also
maintained by ionic transport through channels. A general
example is the passive anion efflux from the cytoplasm into
the extracellular space which is driven by the anion
gradient, triggering membrane depolarization, which in turn
may activate outward-rectifying voltage-gated K+ channels
(as reviewed in Krol and Trebacz, 2000). Anionic channels
are not selective to one type of anion, displaying different
permeability sequences. They are able to transport different
anions according to their localization in the plant and the
3–
role they play in the cell. NO–3, Cl–, SO2–
4 , and PO4 are the
major inorganic anions in plant cells and malate and citrate
are the major organic anions (Pineros et al., 2008, reviewed
in Barbier-Brygoo et al., 2000; Roberts, 2006).
Plant channel activity may be directly or indirectly
regulated by different effectors, calcium being the
major link between electrical signals and cellular activity
(e.g. Pei et al., 1996; for reviews, see Sanders et al., 1999;
Malho et al., 2006). Intracellular and extracellular pH, as
well as intercellular signalling coupled to heterotrimeric G
proteins, are also common in plants (e.g. Johannes et al.,
1998; Colcombet et al., 2005). Signal perception through
cell surface receptors, which are closely coupled with plasma
membrane ion channels, is also known in plants.
Anionic transport in plants
Plasma membrane
The application of electrophysiological techniques to different cell types revealed the activity of a variety of plasma
membrane anion channels and transporters, which are
either constitutively active or are activated by membrane
depolarization, hyperpolarization, light, or stretch. This
resulted in well-characterized anion currents through genetically unknown transporters and/or channels (for reviews
see Tyerman, 1992; Schroeder, 1995; Barbier-Brygoo et al.,
2000; Krol and Trebacz, 2000; White and Broadley, 2001;
Roberts, 2006; Teakle and Tyerman, 2010). The major
exceptions to this trend are the R-type channel activity
(AtALMT12) and the S-type channel activity (SLAC1, slow
anion channel 1) in guard cells from Arabidopsis thaliana.
The general characteristics of these channels, which include
electrophysiological and pharmacological properties, are
summarized in Table 1.
Anionic current characteristics
(i) Depolarization-activated anion channels. Rapidly activating anion efflux channels or R-type channels have been
found and thoroughly studied in the plasma membrane of
Vicia faba guard cells, A. thaliana hypocotyl epidermal cells
and guard cells, and Nicotiana tabacum suspension-cultured
cells. The molecular identity of the R-type channel activity
in the plasma membrane of guard cells from A. thaliana was
recently revealed to be a member of the aluminum-activated
malate transporter family (ALMT). AtALMT12 was found
to be highly expressed in guard cells and allocated to the
plasma membrane. Expression of this protein in Xenopus
oocytes rendered voltage-dependent anion currents, for
which the voltage activation threshold is shifted towards
more hyperpolarized potentials in the presence of extracellular malate. Plants deficient in this gene are impaired in
stomatal closure induced by dark, CO2, and abscisic acid
(ABA), and their guard cell protoplasts display reduced
R-type currents (Meyer et al., 2010).
Slowly activating anion efflux channels or S-type channels
have been described in the guard cells of several species such
as V. faba, A. thaliana, N. tabacum, and Xanthium strumarium, in the epidermal cells of A. thaliana hypocotyls, and in
Coffea arabica suspension-cultured cells. The molecular
identity of the S-type channel activity in the plasma
membrane of the guard cells from A. thaliana has been
recently attributed to a distant homologue of fungal and
bacterial organic acid transport proteins. The channel was
designated SLAC1 and was found to be preferentially
expressed in guard cells, and to be permeable to malate. This
protein is fundamental for stomatal closure in response to
CO2, ABA, ozone, light/dark transition, humidity variation,
elevation of [Ca2+]cyt, H2O2, and NO. Mutation in this
protein results in an ozone-sensitive plant, with impaired
S-type current activity, but with no change in R-type
currents or Ca2+ channel activity (Vahisalu et al., 2008).
(ii) Outward-rectifying anion channels (ORACs). Under
physiological conditions, the electrochemical gradient for
anions in plant cells favours anion efflux (Hedrich and
Becker, 1994), and yet outward-rectifying depolarizationactivated anion channels, which may mediate anion influx,
have been reported in Zea mays and A. thaliana suspensioncultured cells, root cells of Lupinus albus, and in Zostea
muelleri leaf cells. The physiological role in these last cells is
believed to be the stabilization of the plasma membrane
voltage and turgor pressure in the estuarine environment of
this species, which is known to contain high levels of Cl–
(Garrill et al., 1994). An ORAC was also reported in root
cells from Triticum aestivum, and is believed to allow the
entrance of either Cl– or NO–3 into the cell when the
extracellular concentrations of these ions are high (Skerrett
and Tyerman, 1994).
(iii) Hyperpolarization-activated anion channels
(HAACs). Inward-rectifying HAACs have been detected in
various species of plants and algae, such as in Amaranthus
tricolor cotyledons, in the mesophyll and epidermal leaf cells
of Pisum sativum, in the seed coats of Phaseolus vulgaris, in
DPC (diphenylamine–2-carboxylic acid); EA, ethacrynic acid; IAA-94, R(+)-2-[(2-cyclopentenyl-6,7-dichloro-2,3-dihydro-2-methyl-1-oxo-1H-inden-5-yl)oxy]acetic acid; SITS, 4-acetamido4#-isothiocyanatostilbene-2,2#-disulphonic acid; ARAC, Arabidopsis root anion channel; mal2–,malate; acet–, acetate; prop–, propionate; extracellular anion concentration ([anion]ext);
–
+
[Ca2+]cyt, cytosolic calcium concentration; [ATP]cyt, cytosolic ATP concentration; pHcyt, cytosolic pH;SO2–
4 cyt, cytosolic sulphate; [Cl ]ext, extracellular chloride concentration; [Na ]ext,
extracellular sodium concentration; [K+]ext, extracellular potassium concentration; [NO–3]ext, extracellular nitrate concentration.
Designation
Channel type and
gating
Kinetics
Unitary
conductance
Selectivity
Regulation
Inhibition
Cell type
References
R-type or GCAC1
Depolarization
activated;
hyperpolarization
deactivated Ushaped I/V curve;
activation current
peaks at Vm more
negative than EA-;
inward rectifying.
Rapid (ms), timedependent activation
and deactivation; slow
inactivation
Small, 30–40 pS (154
mM [Cl–]in:40 [Cl–]out);
dependent on
[anion]ext
SCN–>NO–3>
I–>Br–>Cl–> mal2–>>
acet–> pro–
[Ca2+]cyt, [ATP]cyt,
pHcyt. Auxin, mal2–,
acet–, pro–, Cl– and
SCN–.
NA¼IAA94¼NPPB>DIDS >>
EA>A-9C>probenecid
V. faba guard cells
NA >> NPPB>IAA-94
A. thaliana hypocotyl
epidermal cells
[ATP]cyt,
phosphorylation
NPPB>DIDS>A-9-C
Phosphorylation,
[Ca2+]cyt, ABA
NA >> NPPB>A-9C>IAA-94
A-9-C¼NA
N. tabacum
suspension-cultured
cells
V. faba guard cells
Keller et al., 1989;
Hedrich et al., 1990;
Marten et al., 1991;
Schroeder and Keller,
1992; Hedrich and
Marten, 1993; Hedrich
et al., 1994; Schmidt
and Schroeder, 1994;
Schmidt et al., 1995;
Schulz-Lessdorf et al.,
1996; Dietrich and
Hedrich, 1998
Thomine et al., 1995,
1997; Frachisse et al.,
1999; Diatloff et al.,
2004;
Zimmermann et al.,
1994, 1998
R-type
–
–
2–
Small, 21 pS (150 mM NO–3>SO2–
4 >Cl >HCO3 [ATP]cyt, SO4 cyt,
–
–
2–
>> mal
[anion]ext.
[Cl ]in : 100 [Cl ]out)
R-type or TSAC
Small, 15 pS (150 mM [Cl–]in:100 [Cl–]out)
S-type
Depolarization
activated; voltage
independent;
inward-rectifying;
less pronounced
U-shaped I/V
curve; activation
current peaks at Vm
more negatives
than the EA-
Slow (<1 min)
activation and
deactivation; never
inactivate
Moderate, 33–35 pS;
long open and closed
duration; not
dependent on
[anion]ext
High NO–3;
impermeable to SO2–
4
DPC¼glibenclamide
>> A-9-C
DIDS
Schroeder and Keller,
1992
Grabov et al., 1997
N. tabacum guard
cells,
A. thaliana guard cells, Forestier et al., 1998
A. thaliana hypocotyl
epidermal cells
X. strumarium guard
cells, and C. arabica
suspension-cultured
cells
Frachisse et al., 2000
Linder and Raschke,
1992; Dieudonne
et al., 1997
2276 | Tavares et al.
Table 1. Summary of plant anion channels electrophysiologically characterized
Table 1. Continued
Designation
Channel type and
gating
ORAC
Depolarization
activated, outward
rectifying.
Kinetics
Strong voltage
dependence, inward
rectification.
Selectivity
Regulation
Inhibition
Induced by high
[Cl–]ext
Rapid activation (ms)
HAAC
Unitary
conductance
Very small, 4 pS
Large, 300 pS
Time-dependent
activation (100–400
ms), slow inactivation
(1–10 s)
Rapid activation (20
Large, 100 pS
ms), rapid inactivation
(300 ms)
Rapid activation (<200
ms)
Rapid activation (<200
ms)
NO–3¼CI–>I–
Cl–>citrate
F–>I –> Cl –> Br
–
>malate2–
Unaffected by [Na+]ext DIDS>ClO–4
or [K+]ext; [Ca2+]cyt
A-9-C
[Ca2+]cyt, [ATP]cyt
SITS
Slight negative
pressure
Cl– >> other anions
Zn2+> EA
Z. mays and A.
thaliana suspensioncultured cells, and Z.
muelleri leaf cells
T. aestivum root cells
Fairley et al., 1991;
Cerana and Colombo,
1992; Garrill et al.,
1994
Skerrett and Tyerman,
1994
Zhang et al., 2004a
Elzenga and Van
Volkenburgh, 1997a,
b
L. albus root cells
P. sativum mesophyll
leaf cells
D. carota culture cells
Barbara et al., 1994
C. pelagicus
Taylor and Brownlee,
2003
Heidecker et al.,
1999; Binder et al.,
2003
Schauf and Wilson,
1987
Terry et al., 1991
Amtmann et al., 1997
A. tuberosa
Large, 200 pS
Large, 150 pS
[Ca2+]cyt, [ATP]cyt
Small, 7–44 pS
Small, 18 pS
[Ca2+]cyt
Large, 97 pS (220 mM
[Cl–]out:25 mM [Cl–]in)
Rapid flickering
Small, 27 pS, outward
between the open and current, 13 pS, inward
closed states
current (154 mM
[Cl–]out:85 mM [Cl–]in)
Small
coats of developing
seeds of P. vulgaris
Long open periods
Positive pressure
La3+
A. tricolor cotyledons
H. vulgare
suspension-cultured
cells
A. thaliana callus cells Lew, 1991
C. inflata
Kourie, 1994
Zhang et al., 2004b
N. tabacum cultured
stem cells
V. faba guard cells
Falke et al., 1988
A. thaliana mesophyll
lea cells
Qi et al., 2004
Cosgrove and
Hedrich, 1991
Anionic transport in plant cells | 2277
Anion-selective MS
References
V. utricularis
Slow activation (1.5 s)
Weak voltage
dependence, inward
rectification.
Cell type
Designation
Channel type and
gating
Kinetics
Unitary
conductance
Light-activated anion
channels
Voltage dependent.
Remains open after
the stimulus is
removed
Small, ranging from 23
pS to 46 pS
depending on Vm
Small, 32 pS
(symmetrical 100 mM
KCI solutions)
–
–
Small,12 pS
SO2–
4 >NO3>Cl >>
organic anions
ARAC
X-IRAC
X-QUAC
X-SLAC
Strong voltage
dependence R-type
like.
Rapid activation and
deactivation Partial
inactivation
Depolarization
activated S-type like.
Slow activation
Voltage-independent
inward rectifier
Hyperpolarization
activated Inward
rectifier.
Voltage-dependent,
inward–outward
rectifier.
Slow gating
Depolarization
activated
S-type like.
Selectivity
Regulation
Inhibition
Cell type
References
Blue light
NPPB (20 lM)
A. thaliana hypocotyl
cells
Cho and Spalding,
1996
P. sativum mesophyll
leaf cells
Elzenga and Van
Volkenburgh, 1997a
A. thaliana root cells
Diatloff et al., 2004
Blue light; [Ca2+]cyt;
[anion]ext, [SO2–
4 ]cyt
NA>NPPB>A-9-C
DIDS
Large, 70–85 pS
NO–3¼Cl–
[Ca2+]cyt, ABA
NO–3>Cl–>malate
[Ca2+]cyt, [NO–3]ext
DIDS¼IAA-94
H. vulgare root XPCs
NO–3¼Cl >I
>malate>SO2–
4 >citrate
[Ca2+]cyt, [ATP]cyt,
ABA
NA
Z. mays root stele
Cl–
[Ca2+]cyt
Large, 90 pS
Rapid activation, slow
inactivation
Instantaneous
activation,
slow inactivation
Slow
Cortical and epidermal Kiegle et al., 2000
root cells and mature
pericycle cells
A. thaliana root hairs, Dauphin et al., 2001
V. unguiculate, P.
vulgaris
H. vulgare root XPCs Kohler and Raschke,
2000
Z. mays root stele
Gilliham and Tester,
2005
H. vulgare root XPCs
Kohler and Raschke,
2000
Gilliham and Tester,
2005
Kohler and Raschke,
2000
2278 | Tavares et al.
Table 1. Continued
Anionic transport in plant cells | 2279
suspension-cultured cells from Daucus carota, in A. thaliana,
in Hordeum vulgare and Asclepias tuberose, in Chara inflata,
in the marine phytoplankton Coccolithus pelagicus, and the
marine alga Valonia utricularis. The only feature that these
channels have in common is a voltage-dependent inward
rectification, which might indicate different natures for
these channels.
(iv) Mechanosensitive anion-selective channels or MS.
Stretch-activated anion-selective channels were found in the
plasma membrane of cultured stem cells from N. tabacum,
guard cells from V. faba, and mesophyll cells from
A. thaliana leaves by means of the patch–clamp technique
(Falke et al., 1988; Cosgrove and Hedrich, 1991; Qi et al.,
2004). All these channels are believed to be involved in
osmoregulation during osmotic stress and cell expansion,
but the differences in their biophysical properties suggest
that they have distinct roles in this process. The first
channel is believed to be involved in mediating large (turgor
resetting) anion fluxes, while the second could have a role in
osmosensing.
Recently the molecular identities of two MSL (mechanosensitive channel of small conductance, MscS-like) channels
were reported in the plasma membrane of root cells from
A. thaliana. These were designated MSL9 and MSL10 and
had unitary conductances of 4562 pS and 13765 pS,
respectively (in 100 mM [Cl–]out:150 mM [Cl–]in). They were
activated by positive pressure and were suggested to rapidly
depolarize the membrane or to alter the turgor pressure of
plant cells in response to mechanical force (Haswell et al.,
2008; Peyronnet et al., 2008).
(v) Light-activated anion channels. Cho and Spalding
(1996) reported a blue light-activated anion channel in the
plasma membrane from A. thaliana hypocotyl cells, by means
of the patch–clamp technique, which activity is believed to
cause the transient depolarization of the plasma membrane
induced in hypocotyl cells by blue light. Elzenga and Van
Volkenburgh (1997a), reported a light-activated anion channel in the plasma membrane of mesophyll cells from P.
sativum leaves. The authors believed that this channel was
responsible for the light-induced transient depolarization
present in these cells (reviewed in Roberts, 2006).
Anion transport in roots. The anion channels present in
the plasma membrane of root cells are important in the
processes of osmoregulation, membrane stabilization, boron
tolerance, and in the regulation of passive salt loading into
the xylem vessels (Roberts, 2006). Generally, they are
considered to function according to their localization, anion
selectivity, and regulation, and have been divided into two
major groups: channels that transport inorganic anions and
channels that transport organic anions. Several inorganic
anionic channels have been well characterized in different
plant species.
ORACs found in root epidermal cells were shown to be
responsible for mediating anion influx (Table 1). On the other
hand, anion efflux from the root was attributed to Arabidopsis
root anion channels, depolarization-activated (ARACs),
found in A. thaliana root cells and hairs, with similar channels
being found in Vigna unguiculate, and P.vulgaris (Table 1).
Kiegle et al. (2000) in a preliminary study using cortical and
epidermal root cells and mature pericycle cells, identified
inward currents that were probably carried by SO2–
4 efflux and
exhibited similar voltage dependence to ARACs (Table 1).
Diatloff et al. (2004) also identified SO2–
efflux as the
4
responsible inorganic anion for the observed inward anionic
currents. These channels share similar properties with R-type
anion channels, which have been previously reported in
A. thaliana hypocotyl and V. faba guard cells (see Table 1
and section below). However, during prolonged application of
activating voltages ARACs are only partially inactivated (i.e.
by 50%) compared with 90% inactivation of R-type channels
in guard cells. They also display different selectivity, ARACs
showing no significant permeability to organic-acid anions,
malate and citrate (Table 1; Diatloff et al., 2004). In contrast,
Dauphin et al. (2001) observed S-type-like currents in
Arabidopsis root hair cells that were only induced by severe
drought stress, but did not detect R-type-like channels.
Finally, two types of channel conductance mediating
anion efflux from the root stele (XPC, xylem parenchyma
cells) were reported in H. vulgare and Z. mays. These are
the X-inward-rectifying anion conductance (X-IRAC), and
the X-quickly activating anion conductance (X-QUAC),
which is most frequently observed in barley and maize steles
(Table 1). A third channel, the X-slowly activating anion
conductance (X-SLAC), which is a depolarization activated
S-type-like channel, was also identified, but rarely observed,
in barley XPCs (Table 1). The three anion conductances
display distinct characteristics, indicating that they are
likely to play distinct roles in salt release (Table 1). The
X-IRAC activity in both maize and barley displays strong
inward rectification and increases with increasing Ca2+cyt,
but the gating mechanism is distinct. In maize, X-IRAC is
voltage dependent (i.e. the channel open probability
increases with negative voltages) whereas in barley it is
considered to be voltage independent, exhibiting an increase
in unitary conductance with membrane hyperpolarization
(reviewed in Barbier-Brygoo et al., 2000; Roberts, 2006).
The channels so far mentioned are responsible for maintaining anion homeostasis (e.g. in salt tolerance during the
repolarization of the membrane potential after Na+ uptake
by roots in saline soils, and in the regulation of osmotic stress
caused by drought or excess water), preventing excessive
plasma membrane depolarization or hyperpolarization, and
also in the regulation of anion loading into the xylem (Table
1; reviewed in Barbier-Brygoo et al., 2000; Roberts, 2006).
For example, anthracene-9-carboxylic acid (A-9-C) and
5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB) inhibits Cl– efflux (possibly through ARAC) during hypo-osmotic
stress, arresting membrane depolarization (Teodoro et al.,
1998). Other studies showed that the presence of 4,4diisothiocyanostilbene-2,2-disulphonic acid (DIDS), a potent
inhibitor of X-QUAC, as well as water stress and ABA
treatment, reduced significantly anion loading into the xylem
of intact excised barley roots (Kawachi et al., 2002) and the
accumulation of solutes in maize roots, respectively, probably via the down-regulation of X-QUAC (Cram and Pitman,
1972; Sharp and Davies, 1979; Gilliham and Tester, 2005).
2280 | Tavares et al.
An ABA-induced Ca2+-dependent signalling mechanism may
be involved (Gilliham and Tester, 2005).
Moreover, boron transport is crucial in root development
since it maintains cell wall integrity and protects shoots
from boron deficiency. A membrane protein transporter
BOR1, was shown to be responsible for the latter process
during xylem loading in A. thaliana root cells
(Takano et al., 2002). Recently an aquaporin was implicated in the regulation of boron efflux. The nodulin 26-like
intrinsic proteins (NIPs) (Chaumont et al., 2001; Johanson
et al., 2001; Johanson and Gustavsson, 2002; reviewed in
Chaumont et al., 2005) have a wide functional repertoire,
and are closely regulated by phosphorylation and/or pH
changes. In addition, it has become increasingly clear that
some of these proteins do not exhibit strict specificity for
water, but can transport a wide range of small neutral and
anionic molecules, such as CO2, NO, H2O2, and boron
(Miwa et al., 2010; reviewed in Miwa and Fujiwara, 2010).
For example, recent work demonstrated that the Arabidopsis major intrinsic protein, localized in the plasma membrane, NIP5;1, is essential for efficient boron uptake and
plant development under boron limitation (Takano et al.,
2006; reviewed in Miwa and Fujiwara, 2010).
Most notable are the organic anion-selective channels,
which are regulated by extracellular Al3+ or the PO3–
4 status
of the plant. In general, ALMT-type anion channels have
multiple functions in anion homeostasis, contributing to the
regulation of growth and responses to the environment,
specifically in acidic soils when Al3+ toxicity may be a major
limiting factor in plant growth (Kochian, 1995). In
particular, T. aestivum (wheat) has an Al3+-activated malatepermeable channel, ALMT1 (also known as TaALMT1) that
is localized in the plasma membrane, and confers resistance
to Al3+ (Sasaki et al., 2004; Yamaguchi et al., 2005; Pineros
et al., 2008; Zhang et al., 2008), through the secretion of
organic anions that chelate Al3+, such as citrate or malate
(Ma et al., 2001; Kochian et al., 2004; Delhaize et al., 2007;
Ryan et al., 2009). Similar transport systems involving
TaALMT1 homologues, conferring resistance to Al3+, were
also found in other species such as maize (Kollmeier et al.,
2001; Pineros and Kochian, 2001; Pineros et al., 2002), and
rape (Brassica napus) (Ligaba et al., 2006).
Additionally, a subgroup of transporters from the
multidrug and toxic compound extrusion (MATE) family
has been shown to mediate organic anion transport in roots
from several plant species. Alt(SB) encodes an Al3+
-activated citrate transporter, which is responsible for
sorghum (Sorghum bicolor) Al3+ tolerance (Magalhaes
et al., 2007). In barley (H. vulgare L.), the gene HvAACT1
was found to be responsible for Al3+-activated citrate
secretion and for Al3+ resistance (Furukawa et al., 2007).
Furthermore, two MATE family members, ZmMATE1 and
ZmMATE2, were found and characterized in maize.
ZmMATE1 is a citrate transporter that confers Al3+
tolerance, and a functional homologue of the Al3+ tolerance
genes described above, while ZmMATE2 could be involved
in a novel Al3+ tolerance mechanism, since it does not
encode a citrate transporter (Maron et al., 2010).
Anion transport is a key player in stomatal movement.
Stomatal aperture is tightly regulated according to environment fluctuations, limiting both carbon dioxide uptake and
water loss. As water osmotically enters and increases guard
cell volume, the cells swell and move apart, enlarging the
stomatal aperture. On the other hand, salt loss upon anion
channel activation causes the guard cells to shrink and the
stomata to close. Guard cells have highly specialized ion
transporters because, unlike in other plant tissues, ion
transport occurs in both directions to allow for stomatal
opening and closing (Roelfsema and Hedrich, 2005). Several
types of plasma membrane anion channels or transporters
have been positively implicated in stomatal movement. The
best known are the anion channels that mediate anion
efflux, which were first discovered in V. faba and were
classified electrophysiologically as rapid [R-type or guard
cell anion channel 1 (GCAC1), ORAC], or slow (S-type)
anion channels (reviewed in Roberts, 2006; Pandey et al.,
2007).
The R-type channel mediates a strongly voltagedependent activated anion current with rapid activation
and deactivation kinetics, and the S-type channel mediates
a weak voltage-dependent activated anion current, with
slow activation and deactivation kinetics. The two voltagedependent anion channels (R- and S-type) have been
described in the guard cells of several other species (Table
1), and were shown to co-exist at the plasma membrane of
V. faba guard cells (Schroeder and Keller, 1992) and
Arabidopsis hypocotyl cells (Frachisse et al., 2000). R- and
S-type channels exhibit divergent properties not only in
terms of voltage dependence, and activation and deactivation kinetics, but also in terms of selectivity, nucleotide
regulation, and pharmacological sensitivity (Table 1). This
last one, in particular, is dependent on both species and cell
type, which might indicate a variation in the nature of these
channels (Keller et al., 1989; Schroeder and Hagiwara,
1989; Hedrich et al., 1990; Schroeder et al., 1993; Schmidt
and Schroeder, 1994; Pei et al., 1996).
Nevertheless both channel types show a higher permeability to NO–3 than to Cl– and are characterized by a Ushaped I/V (current–voltage) curve, the current amplitude
of which peaks at Vm (membrane potentials) more negative
than EA- (anion equilibrium potential). S-type channels show
a peak that is substantially less pronounced and occurs at
more positive Vm than in R-type currents, allowing a wider
range of voltages in which this channel is open. Both have
small to moderate unitary conductances. R-type channels
are closed at hyperpolarizing Vm and open with the
depolarization of the membrane (Table 1).
Functionally, S-type channels are responsible for the
sustained anion efflux during stomata closure. It is highly
regulated by phosphorylation, [Ca2+]in, ABA, and CO2 or
by switching off photosynthetic active radiation in V. faba
(Schroeder and Hagiwara, 1989; Schmidt et al., 1995;
Brearley et al., 1997; Roelfsema et al., 2002), N. tabacum
(Marten et al., 2007, 2008), and A. thaliana guard cells
(Allen et al., 1999; Pei et al., 1997). Marten et al. (2008)
showed that Ca2+ signals enhance the degree of S-type
Anionic transport in plant cells | 2281
anion channel activity during light–dark transition. Additionally, Colcombet et al. (2005), using the patch–clamp
technique on Arabidopsis hypocotyl cells, reported that Rtype and S-type anion channels at the plasma membrane
were both regulated by pH via distinct mechanisms. This
regulation via pH might occur in several signalling networks
including anion fluxes and pH variations, such as those
observed in response to pathogens or hormones. Furthermore, it constitutes a powerful argument in favour of the
existence of two different channels, contradicting the
hypothesis of Dietrich and Hedrich (1994), which accounts
for an interconversion from rapid to slow mode of a single
channel. Recently, the SLAC1 protein has been shown to be
an essential component of the S-type anion channel
function in stomatal signalling in guard cells in Arabidopsis
(Negi et al., 2008; Vahisalu et al., 2008).
On the other hand, R-type anionic channels are involved
in membrane excitability and auxin sensing. These channels
are activated by nucleotide binding and [Ca2+]in (Hedrich
et al., 1990; Zimmermann et al., 1994; Colcombet et al.,
2001), and regulated by intracellular pH (Schulz-Lessdorf
et al., 1996; Colcombet et al., 2005) and extracellular
anions. Similarly, in Chara corallina, protons have a direct
effect on Cl– efflux during intracellular acidosis, suggesting
that the regulation of anion channels by cytosolic pH plays
a specific role in cytosolic pH regulation in plant cells by
providing an anionic shunt conductance (Johannes et al.,
1998). Particularly in the presence of extracellular anions,
including malate and Cl–, which would accumulate in the
apoplast during stomatal closure, the voltage regulation of
guard cell R-type channels shifts to more negative voltages,
contributing to membrane depolarization (Roelfsema et al.,
2004; Roelfsema and Hedrich, 2005; reviewed in Pandey
et al., 2007). Sasaki et al. (2010) using electrophysiological
and loss-of-function mutation studies, showed that
AtALMT12 (a homologue of the Al3+-activated malate
transporter of wheat, TaALMT, in Arabidopsis) is an anion
transporter, particularly permeable to Cl– and NO–3, and
a key regulator of stomatal closure, localized to both
endomembranes and plasma membrane of guard cells of
A. thaliana. Meyer et al. (2010) showed that AtALMT12
represented the guard cell R-type anion channel, and was
highly expressed in guard cells and allocated to the plasma
membrane.
The ABC transporters are known to be involved in a wide
range of processes, such as polar auxin transport, lipid
catabolism, xenobiotic detoxification, disease resistance,
and stomatal function (reviewed in Sanchez-Fernandez
et al., 2001; Rea, 2007; Verrier et al., 2008). AtMRP5 has
been reported to play a role in anion permeability regulation in Arabidopsis guard cells (Suh et al., 2007). Additionally, two genes, AtMRP1 and AtMRP2, were associated
with anionic transport. AtMRP1 promotes the accumulation of folic acid in the vacuole (Raichaudhuri et al., 2009),
while AtMRP2 was found to contribute to cell detoxification and chlorophyll degradation, and to transport organic
anions into the vacuole (Frelet-Barrand et al., 2008).
Furthermore, a malate transporter (AtABCB14, an ABC-
class transporter) and an NO–3 transporter (AtNRT1.1/
CHL1) were reported. The first modulates stomatal movement by increasing osmotic pressure in guard cells (Lee
et al., 2008), and the second functions in stomatal opening
in the presence of NO–3 (Guo et al., 2003).
Tonoplast
Vacuoles play key roles in many physiological processes of
plants (reviewed in de Angeli et al., 2007; Martinoia et al.,
2007). Some of the more prominent are turgor regulation,
pH homeostasis, storage of minerals and nutrients, cellular
signalling, and protein degradation. Anions, in particular
NO–3 (nitrogen source), malate, and Cl–, required for plant
growth, are stored in the central vacuole of cells from root
and shoot tissues, entering the cell via the plasma membrane
and translocated by vacuole transporters. Besides their
potential nutritional value, these anions play other important
roles in plant vacuoles, such as the maintenance of charge
balance and pH regulation, and as osmolytes involved in the
generation and maintenance of cell turgor. In particular,
stomatal opening requires high rates of uptake of Cl– and
malate into guard cell vacuoles. In addition, in plants
showing crassulacean acid metabolism (CAM), malate plays
a major role in plant carbon metabolism, being synthesized
at night from atmospheric CO2 and transferred to the
vacuole as malic acid, constituting a temporary storage of
carbon, and latter providing CO2 for assimilation through
the Calvin cycle during the day (reviewed in de Angeli et al.,
2007; Martinoia et al., 2007).
Functional studies on several species have confirmed the
presence of anionic channels in the tonoplast. The bestcharacterized vacuole anion channel in higher plants is the
vacuole malate channel (VMAL). Using the patch–clamp
technique, electrical currents carried by malate have been
described in the tonoplast of both C3 plants (Pantoja et al.,
1992; Plant et al., 1994; Cerana et al., 1995; Pei et al., 1996),
and CAM plants (Iwasaki et al., 1992; Smith and Bryce,
1992; Cheffings et al., 1997; Pantoja and Smith, 2002; Hafke
et al., 2003). Hafke et al. (2003), showed that malate is
efficiently transported by small conductance (3 pS) channels
in isolated vacuoles from leaf mesophyll cells of Kalanchoe
daigremontiana (a CAM plant), displaying very similar
macroscopic inward-rectifying currents as those observed in
vacuoles from both C3 and other CAM plants. According
to the authors, the kinetic characteristics of the macroscopic
current densities recorded at physiological voltages and the
estimated channel density of 0.2 lm2 are sufficient to
account for the observed rates of nocturnal malic acid
accumulation, thus representing the principal pathway for
malate influx into the vacuole. Furthermore, the CAM
malate channel was shown to be actively modulated by both
cytoplasmic pH and external pH, with the current decreasing with acidification of the cytoplasm (Pantoja and
Smith, 2002; Hafke et al., 2003). VMAL is responsible for
the transport of malate, but is also permeable to succinate,
fumarate, acetate, oxaloacetate, NO–3, and H2PO–4. Pronounced inward rectification, allowing anions to enter but
2282 | Tavares et al.
not to leave the vacuole, and slow activation kinetics are
general characteristic features of the whole-vacuole malate
currents described for various cell types, including vacuoles
from storage parenchyma cells of Beta vulgaris (Pantoja
et al., 1992), suspension culture cells of A. thaliana (Cerana
et al., 1995), stomatal guard cells of V. faba (Pei et al.,
1996), and leaf mesophyll cells of K. daigremontiana (Smith
and Bryce, 1992; Pantoja and Smith, 2002; Hafke et al.,
2003). Furthermore, VMAL was shown to be barely
affected by cytosolic Ca2+ or ATP (Cerana et al., 1995;
Cheffings et al., 1997). Cl– currents (VCL) have also been
found in plant vacuoles (Pantoja et al., 1992; Plant et al.,
1994; Cerana et al., 1995; Pei et al., 1996). Pei et al. (1996)
reported the presence of malate and Cl– (VCL) conductances in guard cell vacuoles of V. faba, which were activated
by a calmodulin-like domain protein kinase (CDPK) and
ATP. The activation of VCL at physiological potentials
enabled Cl– uptake into the vacuole, providing a pathway
for stomatal opening. Since CDPK-activated VCL currents
were also observed in red beet vacuoles, the authors
suggested that these channels may provide a more general
mechanism for kinase-dependent anion uptake, volume
regulation, and cell expansion in other types of plant cell
(Pei et al., 1996). Interestingly, a response of a Cl– channel
to acetylcholine was shown in the tonoplast of C. corallina
(Gong and Bisson, 2002). Acetylcholine has long been
suggested to play a role in controlling physiological processes in plants, but no mechanism has been shown for its
action. Gong and Bisson (2002) hypothesized that this
acetylcholine-gated channel has evolved separately from the
mammalian acetylcholine-gated channel, and suggest that
this represents a third form of acetylcholine signal transduction, after the nicotinic and muscarinic pathways in
animal systems.
At the molecular level, several anion transport systems
have been identified. One of these was AtTDT, which
encodes the vacuole malate transporter from Arabidopsis
leaves (A. thaliana tonoplast dicarboxylate transporter),
responsible for malate influx (Emmerlich et al., 2003).
Hurth et al. (2005), using entire leaf discs and mesophyll
protoplasts from A. thaliana tdt knock-outs, showed that
the activity of AtTDT catalyses malate import into isolated
vacuoles and is critical for the regulation of pH homeostasis. On the other hand, cytosolic acidification induced the
expression of the AtTDT gene and correlated with a dramatic decrease in total cellular dicarbonic acid levels,
indicating that AtTDT may also play a role in malate
export. The authors hypothesized that the decrease in
malate/fumarate and increase in citrate content observed in
mutant leaves in comparison with wild-type plants indicate
that AtTDT is not the sole vacuole malate exporter. These
alterations suggested that Arabidopsis vacuoles contain at
least two transporters and a channel (VMAL) for dicarboxylate and citrate (Hurth et al., 2005). Later, a malate
channel, AtALMT9, was molecularly identified by
Kovermann et al. (2007). This channel represents a key
point in the regulation of cytoplasmic pH and in the control
of cellular metabolism, particularly in plants showing
CAM, in which large fluxes of malate occur during the day/
night cycle (Pantoja and Smith, 2002; Kovermann et al.,
2007). Four of the seven homologue members of the Cl–
channel (CLC) family, AtCLCa–c and g, were localized in
the tonoplast (De Angeli et al., 2006; Lv et al., 2009; von
der Fecht-Bartenbach et al., 2010).
Anionic transporters in other endomembranes
There are reports and descriptions of the existence of
voltage-dependent anion channels in endomembranes, such
as the outer and inner membranes of mitochondria and
chloroplasts, and the Golgi system (reviewed in Krol and
Trebacz, 2000; Kusano et al., 2009). Voltage-dependent
anion channels, showing selectivity for Cl–, were characterized by a patch–clamp study in isolated membrane patches
in osmotically swollen thylakoids from Peperomia metallica
(Schönknecht et al., 1988). Physiologically, this Cl– channel
could provide a mechanism involved in charge balancing
during light-driven proton uptake by thylakoids
(Schönknecht et al., 1988). Different reports have already
demonstrated the presence of a voltage-dependent anion
channel and a voltage-dependent solute channel in the outer
envelope of chloroplasts. The latter is thought to function
as a general solute channel (Krol and Trebacz, 2000).
In the membranes of mitochondria, anion transporters
are thought to regulate membrane potential, e.g. during
transduction of an apoptotic signal into the cell or
metabolite diffusion. In contrast to the inner mitochondrial
membrane, the outer mitochondrial membrane is highly
permeable to small molecules. It contains a voltagedependent anion channel (VDAC) or mitochondrial porin,
which is thought to facilitate metabolite exchange between
the organelle and the cytosol, and is also involved in
mitochondria-mediated cell death and in defence against
pathogens. VDACs have been reported in pea, potato,
tobacco, and Arabidopsis (reviewed in Kusano et al., 2009).
It has long been established that the inner membrane of
plant mitochondria is permeable to Cl–. Beavis and Vercesi
(1992), applying a light-scattering technique in isolated
mitochondria from white potato tubers (Solanum tuberosum), provided evidence of a pH-regulated and Mg2+insensitive ‘plant inner membrane anion channel’ (PIMAC),
that transports a wide variety of anions including NO–3, Cl–,
ferrocyanide,
1,2,3-benzene-tricarboxylate,
malonate,
H2PO–4, a-ketoglutarate, malate, adipate, and glucuronate.
The PIMAC revealed similarities to the so-called inner
membrane anion channel (IMAC) already described in
animal mitochondria, which is inhibited by matrix Mg2+,
unlike PIMAC. It was suggested that PIMAC may play
a role in volume homeostasis and also in the malate–
oxaloacetate shuttle (Beavis and Vercesi, 1992). Later,
Lurin et al. (2000) showed that tobacco NtCLC-1 specifically co-localizes with the markers of the mitochondrial
inner membrane, cytochrome c oxidase, and NAD9 protein.
Their data suggest that NtCLC-1 is a plausible candidate
for the PIMAC, and its gene may be a unique molecular
tool to study the putative role of anion transport in
Anionic transport in plant cells | 2283
mitochondria (Lurin et al., 2000). More recently, Laus et al.
(2008), by means of swelling experiments in K+ and
ammonium salts, further characterized a PIMAC-like
anion-conducting pathway in mitochondria from durum
wheat. The authors suggested that PIMAC, normally
inactive in vivo due to ATP and high electrical membrane
potential, can suffer activation in mitochondria in order to
replace or integrate the operation of classical anion carriers
(Laus et al., 2008).
Additionally, Marmagne et al. (2007) showed that
AtCLCe and AtCLCd–f proteins are targeted to the
thylakoid membranes in chloroplasts and Golgi
membranes, respectively.
A family portrait of the anionic transport families
The chloride channel (CLC) family. CLC proteins constitute
anionic channels/transporters ubiquitously found in eukaryotes and prokaryotes (Mindell and Maduke, 2001). The
first member of this family to be characterized was the
CLC-0, a voltage-gated Cl– channel found in the electric
organ of Torpedo californica (White and Miller, 1979).
Mammalian CLCs are the best-known elements of this
family, and they encompass both Cl– channels, in the
plasma membrane, and Cl–/H+ antiporters in intracellular
compartments (Zifarelli and Pusch, 2007; Jentsch, 2008).
In A. thaliana seven homologues, AtCLCa–g, have been
identified (Hechenberger et al., 1996; Lv et al., 2009).
According to Lv et al. (2009) these homologues form two
distinct subclasses with a low similarity level (26–29%)
between them. The first subclass is constituted by
AtCLCa–d and AtCLCg and has greater similarity to
other eukaryotic CLC subclasses, while subclass II, composed of AtCLCe and AtCLCf, has low homology to the
other plant CLCs and is more closely related to prokaryote
CLCs. The expression of the AtCLC family was studied
and it was found to be ubiquitous both temporally and
spatially, but with distinct expression patterns between the
two subclasses. Lv et al. (2009) found that all members of
the AtCLC family were predominantly expressed in the
vascular tissues, especially those from subclass I, in both
roots and shoots, implying a possible role for this family in
long-distance ion transport within the plant. Strong
expression of all AtCLCs, particularly AtCLCc, was also
found in guard cells (Lv et al., 2009) where it is involved in
the regulation of stomatal movement and contributes to
salt tolerance (Jossier et al., 2010). The specificity of this
channel for NO–3 has been found recently to be associated
with a single amino acid residue, proline 160 (Wege et al.,
2010). The cellular localization of CLCs placed them all in
intracellular membrane systems: AtCLCa–c and g were
localized in the tonoplast (De Angeli et al., 2006; Lv et al.,
2009; von der Fecht-Bartenbach et al., 2010), while
AtCLCd and f were targeted to Golgi vesicles (Marmagne
et al., 2007; von der Fecht-Bartenbach et al., 2007; Lv
et al., 2009), and AtCLCe was localized in the chloroplasts
(Marmagne et al., 2007; Lv et al., 2009).
The best known of all AtCLC proteins in plants is
AtCLCa. De Angeli et al. (2006) localized this transporter
to the tonoplast, and demonstrated that this protein
behaves as a 2NO–3/1H+ exchanger, whose function is the
specific accumulation of NO–3 in the vacuole (De Angeli
et al., 2006). AtCLCb was also functionally characterized
as a NO–3/H+ exchanger, with strong outward rectification,
but its physiological context has yet to be investigated
(von der Fecht-Bartenbach et al., 2010). AtCLCc homozygous knockout mutants were found to present significantly
lower NO–3, Cl–, malate, and citrate concentrations than
their wild-type counterparts. This gene was also found to
be down-regulated in the presence of NO–3 (Harada et al.,
2004). AtCLCd was co-localized with the VHA-a1 subunit
of the V-type ATPase in the trans-Golgi network (TGN),
suggesting that AtCLCd mediates the transport of an
anion, Cl– or NO–3, into the TGN to counter the pumping
of H+ by the V-ATPase (von der Fecht-Bartenbach et al.,
2007). AtCLCe homozygous knockout mutants were
found to under-accumulate NO–3 and over-accumulate
nitrite (NO–2) (Monachello et al., 2009). The authors
believed that AtCLCe acted as an anion channel pumping
either Cl– or NO–2 into the thylakoid lumen to compensate
the excess positive charge in this subcellular compartment,
and stated that this protein was important in NO–3
assimilation through cytosolic nitrite. A transcriptional
link between AtCLCa and AtCLCe was also found in
AtCLCa homozygous knockout mutants, which showed
down-regulation of the AtCLCe gene (Monachello et al.,
2009). AtCLCf and AtCLCg functional properties are still
unknown, although some evidence has been presented
associating AtCLCf with the acidification of TGN vesicles
(Marmagne et al., 2007). Table 2 summarizes the
characteristics of these channels.
The ALMT channel family. ALMTs were found to
promote Al3+ tolerance in several cultivars, by excreting
into the root system Al3+-chelating organic anions, such as
malate and citrate, in response to Al3+ exposure. Al3+ was
shown, by means of electrophysiological essays, to activate
these channels in cortical cells in the root apex (Kollmeier
et al., 2001; Pineros and Kochian, 2001; reviewed in Ward
et al., 2009). Further evidence for the regulation of these
channels by Al3+ was provided by the expression of ALMTs
in Xenopus oocytes. This was enough to mediate Al3+
-induced malate currents, suggesting a direct Al3+ sensor
role for ALMT transporters (Hoekenga et al., 2006).
There are 14 predicted members of the ALMT channel
family in the genome of A. thaliana. Of these only three,
AtALMT1, AtALMT9, and AtALMT12, have had their
physiological role unravelled. AtALMT1 was found to be
a plasma membrane Al3+-activated malate transporter
expressed in the roots, and was associated with Al3+
tolerance in Arabidopsis (Hoekenga et al., 2006). AtALMT9
is targeted to the tonoplast and mediates malate uptake into
the vacuole (Kovermann et al., 2007). AtALMT12 was
recently associated with the R-type anion currents found in
the guard cell plasma membrane (Meyer et al., 2010). This
gene is transcribed in pollen (Pina et al., 2005).
2284 | Tavares et al.
Table 2. Summary of the characteristics of the A. thaliana CLC family (see text for specific references)
u.n., unknown.
Localization
Expression
pattern
AtCLCa
Tonoplast
Ubiquitous;
vascular tissues
AtCLCb
Tonoplast
Roots, stems,
and siliques;
vascular tissues
NO–3/H+
exchanger
u.n.
Transporter
type
Function
2NO–3/1H+
exchanger
Accumulation of
NO–3 in the vacuole
Voltage
dependence
Selectivity
Slight outward
Strong outward
rectification
rectification
NO
NO
3 ¼I >Br >Cl
3 >Br >Cl
2
>SO4 >glutamate
>malate¼I
AtCLCc
Tonoplast
Ubiquitous;
vascular tissues;
guard cells
u.n.
u.n.
u.n.
AtCLCe
Chloroplast
Leaves, flowers,
and siliques;
vascular tissues
Anion channel
(Cl– or NO–2)
NO–3 assimilation
through cytosolic
NO–2
u.n.
u.n.
u.n.
u.n.
u.n.
The NRT/PTR/POT family. NO–3 transport in plants is
mediated by members of the nitrate transporter/peptide
transporter/proton-dependent oligopeptide transporter
(NRT/PTR/POT) family. In higher plants there are at least
three subgroups of NO–3 transporters, the NRT1s, the
NRT2s, and the NAXTs (Segonzac et al., 2007; reviewed in
Chen et al., 2008). NRT1s are typically low-affinity nitrate
transporters while NRT2s are high-affinity nitrate transporters (Liu et al., 1999). Both families are members of the
major facilitator superfamily (MFS). NO–3 uptake requires
active transport even in the presence of high NO–3 concentration; this process is mediated by proton/NO–3 symporters
(Glass et al., 1992; Walker et al., 1995; Huang et al., 1999;
Zhou et al., 2000). One unique feature of NO–3 uptake,
different from uptake of other mineral nutrients, is that it
can be induced by the availability of its substrates, while
others are only induced by deficiency of their corresponding
substrates (Clarkson and Luttge, 1991; Crawford and Glass,
1998; Daniel-Vedele et al., 1998). In addition to transporting NO–3, NRTs can also uptake peptides and histidine
(Paulsen and Skurray, 1994; Steiner et al., 1995; Zhou et al.,
1998; Tsay et al., 2007).
The number of known genes from the NRT1 family in
plants is larger than in any other organism (Tsay et al.,
2007), which further supports the important role of NO–3 in
higher plants. AtNRT1.1 (or CHL1), a member of the
NRT1 family, was the first NRT1 gene identified in plants
and acts as a dual affinity proton-coupled NO–3 transporter
(Doddema et al., 1978; Tsay et al., 1993; Liu et al., 1999).
The affinity of CHL1 changes with phosphorylation and
dephosphorylation as was shown in Xenopus oocytes
expressing a mutation of the gene (T101D and T101A,
respectively, for the phosphorylation and the dephosphorylation mutations) (Liu and Tsay, 2003; Tsay et al., 2007).
Furthermore, the phosphorylation state of the gene is
regulated by external NO–3 concentration (Liu and Tsay,
2003). Recent reports also point out its dual function as
a NO–3 sensor and transporter (Hu et al., 2009; Wang et al.,
2009). This gene seems to be the exception for the NRT1
AtCLCd
TGN
Ubiquitous;
vascular tissues
Anion channel
(Cl– or NO–3)
Acidification
of TGN vesicles
AtCLCf
TGN
Roots, leaves,
and stems;
vascular tissues
u.n.
AtCLCg
Tonoplast
Ubiquitous;
vascular tissues
Acidification of
TGN vesicles
u.n.
u.n.
u.n.
u.n.
u.n.
u.n.
family, as all others only show low-affinity NO–3 transport
activity, e.g. AtNRT1.2 (epidermis, Liu et al., 1999),
OsNRT1 (root epidermis, Huang et al., 1999; Lin et al.,
2000), AtNRT1.4 (leaf petiole, Chiu et al., 2004),
AtNRT1.5 (root pericycle, Lin et al., 2008), AtNRT1.6
(silique and funiculus vascular tissue, Almagro et al., 2008),
AtNRT1.7 (leaf phloem, Fan et al., 2009), and AtNRT1.8
(root xylem, Li et al., 2010). Despite sequence similarities,
these genes are expressed in different tissues and may prove
to play significant differential roles in nitrogen transport
and uptake.
The NRT2 protein alone does not show any NO–3
transport activity unless in the presence of an additional
component, NAR2, as was demonstrated in Xenopus
oocytes (Zhou et al., 2000) and yeast split-ubiquitin system
(Orsel et al., 2006). AtNRT2.1 and AtNRT2.2 are both
involved in high-affinity NO–3 transport (Filleur et al., 2001)
and impairment of these genes causes reduced NO–3 uptake
(Li et al., 2007). Another gene, AtNRT2.7, has been
implicated in the loading of NO–3 to the vacuole during seed
maturation (Chopin et al., 2007). Meanwhile, Monachello
et al. (2009), suggested cross-talk between several transport
systems involving members of the CLC and NRT families.
There has also been a great effort to unveil the complex
regulatory network involved in NO–3 uptake (Guo et al.,
2001; Munos et al., 2004; Little et al., 2005; Remans et al.,
2006; Vidal et al., 2010).
In addition, an NO–3 excretion transporter that mediates
NO–3 efflux from the cortex of mature roots, NAXT1, which
is electrically linked to ATP-dependent proton pumping was
reported (Segonzac et al., 2007).
The Pht transporter families. Plants acquire PO3–
4 mainly
via Pi (H2PO–4) transporters, which are classified into three
different families: Pht1, Pht2, and Pht3 (Rausch and
Bucher, 2002). These transporters are typically driven by
a proton gradient generated by plasma membrane H+
-ATPases, since PO3–
4 concentration in the soil is generally
orders of magnitude lower than inside plant cells (UllrichEberius et al., 1981; Tu et al., 1990).
Anionic transport in plant cells | 2285
The plant Pht1 transporters belong to the Pi:H+ symporter (PHS) family, included in the MFS. The first plant
Pht1 gene was cloned from Arabidopsis (Muchhal et al.,
1996), many more having been identified since then. Results
from Bucher et al. (2001), suggest that members of the Pht1
family are high-affinity Pi cotransporters. Most of these
genes are expressed in roots, particularly in rhizoderm cells
and in the outer cortex (Daram et al., 1998; Liu et al., 1998;
Chiou et al., 2001; Mudge et al., 2002). Other reports also
indicate Pht1 genes in leaves and pollen, which suggest new
roles for these proteins in addition to Pi uptake at the roots
(Mudge et al., 2002; Rae et al., 2003; Nagy et al., 2006).
Recently, the characterization of the low-affinity barley
phosphate transporter PHT1.6 in Xenopus oocytes demonstrated that, apart from PO3–
4 , this protein is also able to
–
transport SO2–
4 , and to a lesser extent NO3 and chlorate
(Preuss et al., 2010).
Function analysis of Pht2;1 protein remains the most
comprehensive study of members of the Pht2 transporter
family so far, and it led to the assumption that the Pht2
family may be characterized by low-affinity Pi transporters
(Daram et al., 1999). Further studies indicated its localization in the chloroplast (Ferro et al., 2002; Versaw and
Harrison, 2002) while Pht3 family genes appear to be highly
conserved within the mitochondrial transporter family
(Rausch and Bucher, 2002), though little else is known
about their function so far.
The SULTR transporter family. Plants have developed
a wide range of tissue specific sulphate transporters (Smith
et al., 1997) due to their pivotal role as the main source of
sulphur-containing amino acids. Several genes, closely related to the animal and fungal suphate:H+ cotransporters of
the SuIP family, have been divided into five subgroups
(Hawkesford, 2003; Buchner et al., 2004) encoding SO2–
4
transporters. Typically, high-affinity transporters, such as
the SULTR1;1, SULTR2;1, and SULTR2;2, are preferentially expressed in roots (Leustek and Saito, 1999;
Takahashi et al., 2000; Yoshimoto et al., 2002), while the
low-affinity forms are present in all tissues, but mainly in
leaves (Leustek and Saito, 1999). Interestingly only a few of
these transporters have been involved in intracellular SO2–
4
transport, namely SULTR4;1 and SULTR4;2 (Takahashi
et al., 1999; Hawkesford, 2000; Kataoka et al., 2004).
The pollen tube: a case study
Pollen is the male gametophyte of higher plants. It is
usually produced in copious quantities in the anther and
transported either by air currents or by a variety of animal
pollinators. Part of its success lies in its remarkably tough
external wall and its dehydrated state upon release. When
the pollen grain lands on a compatible female sexual organ,
the vegetative cell of the pollen grain rehydrates and
germinates, originating the pollen tube.
The pollen tube is a highly specialized cell and its main
biological function is the delivery of the two sperm cells
contained in the pollen grain to the female gametophyte—
the embryo sac—in order for double fertilization to occur.
While growing, pollen tubes never divide or ramify, all their
resources being allocated to allow one of the fastest cellular
elongations in nature, reaching rates of up to 4 lm s1, and
lengths up to 40–50 cm. This growth implies an increase in
its volume of several orders of magnitude in just a few
hours (reviewed in Boavida et al., 2005a, b; Michard et al.,
2009). The peculiar cell biology, combined with the almost
complete description of the Arabidopsis pollen transcriptome (Pina et al., 2005; review in Becker and Feijo, 2007),
makes this system specially suited for the study of polarized
cell growth in plants.
Anion fluxes and oscillations
The association between pollen tube growth and ionic fluxes
was established by Lionel Jaffe’s group during the 1970s.
They used what was, at the time, a newly developed
method—the vibrating probe—which consisted of a platinum-black electrode that measured differences in voltage
between two points, thus inferring the total electrical
current flowing between them (Weisenseel et al., 1975;
Weisenseel and Jaffe, 1976). The introduction of ionic
selectivity (though the use of selective ionophores) and
further improvements to this basic method allowed a deeper
understanding of the ionic fluxes and the respective intervening ions. Recent advances in imaging techniques and
in fluorescent dyes have permitted the discovery of intracellular ion gradients that result from the extracellular
ion fluxes. Both phenomena were found to oscillate with
a similar period to that of growth, though with distinct
phase delays; they are also essential for pollen tube growth
(Holdaway-Clarke et al., 1997; Messerli et al., 1999; Feijo
et al., 2001; Zonia et al., 2002; Holdaway-Clarke and
Hepler, 2003).
Cl– fluxes in growing pollen tubes from Lilium longiflorum
and N. tabacum, were first described by Zonia et al. (2002) by
means of the ion-specific vibrating probe (Fig. 1). Strong
oscillatory Cl– efflux occurs specifically at the tube apex (50–
8000 pmol cm2 s1 in Lilium and 400–1200 pmol cm2 s1
in Nicotiana) with a period of 13.2 and 105 s, respectively.
The remainder of the tube shank showed a non-oscillatory
influx. In tobacco, this influx started at ;12 lm from the tip
and peaked at ;26 lm from the tip (4000 pmol cm2 s1;
Zonia et al., 2002). These Cl– fluxes were further confirmed
by the use of known Cl– channel blockers DIDS, niflumic
acid (NA), and NPPB. These compounds not only completely inhibited the tobacco pollen tube growth, but also
induced an increase in apical volume. Furthermore, DIDS
did not affect cytoplasmic streaming, but disrupted the Cl–
efflux. Inositol-3,4,5,6-tetrakisphosphate [Ins(3,4,5,6)P4],
a known Ca2+-activated Cl– conductance blocker (Carew
et al., 2000), inhibited pollen tube growth, induced cell
volume increase, and interrupted Cl– efflux. These effects
were specific for Ins(3,4,5,6)P4, the close analogues
Ins(1,3,4,5)P4 and Ins(1,3,4,5,6)P5 having no significant
effect on these parameters. The Cl– efflux oscillation was
coupled to and temporally in phase with the growth
2286 | Tavares et al.
Fig. 1. Topographical mapping of Cl– flux along the cell surface of the apical region of a tobacco pollen tube. (A) Profile of Cl– flux
starting at the apex and continuing to 26 lm distal to the tip. There is an inversion of oscillatory efflux to net influx at ;12 lm distal to the
tip. (B) Graphic representation of the vectorial flux of Cl– into the pollen tube and out from the apex. Bar 10 lm (adapted from Zonia
et al., 2002).
oscillations. All these data indicated a role for Cl– fluxes in
osmotic homeostasis and apical growth regulation in pollen
tubes (Zonia et al., 2002).
Further evidence supporting the existence of Cl– fluxes in
growing pollen tubes was provided by several reports from
different research groups. Matveyeva et al. (2003) studied
the effect of several blockers of Cl– channels and transporters
in germination and Cl– efflux in pollen grains from N.
tabacum. They observed that NPPB and NA completely
inhibited pollen germination and significantly reduced Cl–
efflux from the grain. DIDS prevented pollen germination,
while furosemide, bumetanide, and bis-(1,3-dibutyl-barbituric
acid)-pentamethine oxonol [DiBAC4(5)], only suppress germination by less than 50%. These authors believed that NPPBsensitive anion channels were involved in the activation of
pollen grains during germination (Matveyeva et al., 2003).
More recently Breygina et al. (2009b) studied variations in
membrane potential during pollen germination and pollen
tube growth, by means of quantitative fluorescent microscopy, and found that the plasma membranes of N. tabacum
and L. longiflorum become hyperpolarized during pollen
germination, and that they present an uneven potential
distribution on pollen grain and tubes. This group also
showed the involvement of the plasma membrane H+ATPase and anion channels in membrane potential regula-
tion (Breygina et al., 2009b). Further proof was provided by
means of the fluorescent dye 6-methoxy-N-ethylquinolinium
iodide (MEQ). These authors also studied the release of
anions from pollen grains and pollen tubes from N. tabacum,
during germination and growth. By applying the inhibitor
NPPB (40 lM) and by increasing the extracellular [Cl–]
([Cl–]ext) to 200 mM (pollen germination) or 100 mM (pollen
tube growth), this group succeeded in completely blocking
the efflux of anions and consequently stopped pollen grain
germination and pollen tube growth. Additional analysis
using the fluorescent dyes bis-(1,3-dibutyl-barbituric acid)trimethine oxonol [DiBAC4(3)], N-(3-tri-ethyl-ammoniumpropyl)-4-{6-[4-(diethylamino)phenyl]hexatrienyl}
pyridinium dibromide (FM4-64), and 10-N nonyl-acridine orange
(NAO), revealed that NPPB not only prevented the efflux of
Cl– and other anions, but also induced depolarization of the
plasma membrane and disruption of pollen tube apical
compartmentalization (Breygina et al. 2009b). Blocking
selective Cl– efflux in pollen tubes with 100 mM [Cl–]ext
caused a significant hyperpolarization of the plasma membrane. These authors believed that the selective block of Cl–
release induced the efflux of other anions, namely organic
anions, which caused the observed arrest of pollen germination and tube growth. This report showed the importance of
Cl– efflux through NPPB-sensitive channels for the normal
Anionic transport in plant cells | 2287
germination of pollen grains and pollen tube growth
(Breygina et al., 2009a). These observations are in accordance
with the predicted role of Cl– fluxes in pollen germination
and tube growth (Zonia et al., 2001, 2002).
Breygina et al. (2010) further reported the effect that the
anion channel blockers NPPB and DIDS had on
N. tabacum pollen tube growth and its mitochondrial state,
via fluorescence microscopy and flow cytometry. The
authors found that 40 lM NPPB completely blocked pollen
tube growth but caused no increase in its diameter, while
20–80 lM of DIDS induced pollen tube swelling and
bursting. Isolated pollen mitochondria treated with DIDS
showed hyperpolarized membranes and a variation in
reactive oxygen species content and excretion. This study
suggested that pollen tube growth is dependent on the
activity of different anion channels, namely in localization
and function (Breygina et al., 2010).
In line with what is known in guard cells and other wellstudied systems, there is a substantial amount of evidence
pointing to the importance of Cl– efflux in the germination of
pollen grains and in the growth of pollen tubes, specifically in
the regulation of cytoplasm compartmentalization, membrane potential regulation, and to a lesser extent mitochondrial regulation. But the main role that has been associated
with Cl– movements across the plasma membrane is that
of maintaining the osmotic pressure by driving the water
movement across the cell. There is abundant evidence of the
importance of the regulation of osmotic pressure during the
apical growth of pollen tubes, and it has been established
that osmotically induced variations in cell volume elicit rapid
variations in phospholipid membrane composition and
signalling (Zonia and Munnik, 2004). Even though the link
between the Cl– movements and the water movements in the
growing pollen tube system has not been thoroughly
recognized, the connection between these two phenomena in
other important regulatory plant systems (e.g. the guard cell)
has been well established. The movement of guard cells is
associated with variations in their turgor pressure, which is
regulated by the flow of K+, Cl–, and organic anions in and
out of the cells.
High Cl– levels selectively inhibit ‘kiss-and-run’ endocytosis–exocytosis or flicker fusion (Smith et al., 2008). The
endocytosis rate appears to be linked to the rate of
exocytosis, and these are sensitive to levels of Ca2+ and Cl–.
High Cl– levels blocked tobacco pollen tube growth, but did
not immediately block smooth vesicle endocytosis at the
apex (Breygina et al., 2009a), suggesting that an endosomal
pathway differs from the ‘kiss-and-run’ exocytosis–
endocytosis pathway.
Anions may thus play a vital role not only in maintenance of the membrane potential, by preserving the electroneutrality and cell osmotic potential, but also by directly
controlling important cellular events. The cellular processes
stated above have been proposed as the physiological
effectors of the Ca2+ gradient (Roy et al., 1999; Parton
et al., 2003; Becker et al., 2004; Hwang et al., 2005; Helling
et al., 2006). Thus one can speculate that anions may also
be responsible for the fine-tuning of the Ca2+ gradient,
which would account for a feedback system (since Ca2+ also
regulates anionic fluxes; e.g. Chen et al., 2010) during pollen
tube growth and development. The signalling cascades
downstream of Ca2+ are multiple (reviewed in Malho et al.,
2006), and may imply phosphorylation through Ca2+dependent protein kinases (Yoon et al., 2006), small
GTPases (Gu et al., 2005) or calmodulin (Rato et al., 2004;
Berkefeld et al., 2010), and could plausibly regulate anionic
currents as well, as revealed by the specificity of the IP4
inhibition response (Zonia et al., 2002).
Regardless of the overwhelming evidence supporting the
existence of Cl– fluxes, these have been involved in
controversy for a long time. Early descriptions of the total
electric currents in pollen tubes using the voltage vibrating
probe and ion substitution experiments concluded that
anions were not a part of the total electric current generated
(Weisenseel and Jaffe, 1976). Justifiable in those days, these
substitution studies are controversial in many senses, and
understandably these results have not been replicated or
reproduced ever since. Of notice, the same paper assigned
a non-existent role for Ca2+ fluxes, now widely accepted as
a major player in the global ion flux regulation by more
than a dozen studies using the Ca2+-specific vibrating probe
(review in Holdaway-Clarke and Hepler, 2003; Michard
et al., 2009).
This disagreement was further developed by Messerli
et al. (2004), in their report on the characterization of the
Cl–-selective microelectrodes used in the experiments described in Zonia et al. (2002). Of note, these authors
described exactly the same sort of fluxes as Zonia et al.
(2002) but developed experiments to claim that the electrodes were poorly selective for Cl– over other anions and that
the electrode could indirectly detect H+ gradients (Messerli
et al., 2004). Substantial rebuttal of the significance of these
arguments to downplay the conclusions of Zonia et al.
(2002) has been published elsewhere (Kunkel et al., 2006;
Moreno et al., 2007). Of relevance, the experimental
parameters of Zonia et al. (2002), namely the MES
concentration and the pH, were grossly altered. Since both
will condition the activity of the anionic form of MES, the
claim that the microelectrode was indirectly detecting
variations in pH falls short of the mark [see Fig. 2; see also
Kunkel et al. (2001) for discussion of the effect of buffers on
ion current measurement]. Furthermore Messerli et al.
(2004) were unable to dismiss the pharmacological evidence
presented by Zonia et al. (2002), namely the IP4 specificity.
The claims by Messerli et al. (2004) remain focused on
theoretical arguments and interpretation of indirect results.
Of essence, the non-specific nature of Cl– ionophores is well
known, but in conditions where the only anion present is
Cl–, as was the case in Zonia et al. (2002), and MES was
kept to a minimum (Kunkel et al., 2001), these arguments
cannot cope with the enormous fluxes measured, orders of
magnitude above what has been reported for protons
(Fig. 2; Feijo et al., 1999; Michard et al., 2008).
Further polemic was added by the same group (Dutta
and Robinson, 2004), by reporting the absence of Cl–
channels in the plasma membrane of pollen protoplasts
2288 | Tavares et al.
Fig. 2. Comparison of time course measurement of anion (A) and proton (B) fluxes in pollen tubes of lily. Oscillations of the apex are
conspicuous and proceed with almost invariable amplitude and period through long experimental periods (note the reference levels in
both traces for signal/noise reference). However, the magnitude of the Cl– fluxes is three or four orders of magnitude above those of
protons (B: adapted from Feijó et al., 1999).
from L. longiflorum. Taken literally, this observation
would make pollen tubes the only cell in nature so far
described without an anion transporting system. Among
other problems, water transport and volume regulation,
on the one hand, and membrane polarization on the
other, would be left unaccounted for, as the current
functional paradigms for these fundamental processes
seem to rely on Cl–/anions. In the same study (Dutta and
Robinson, 2004) the complete absence of hyperpolarization-activated Ca2+ channels was reported. These were
subsequently found by several other authors in the
plasma membrane of pollen protoplasts from Lilium
davidii D., A. thaliana, and Pyrus pyrifolia (Shang et al.,
2005; Qu et al., 2007; Wu et al., 2007). Taken together,
the easiest explanation would be that the cell-attached
configuration of the patch–clamp technique might not be
the most appropriate one to study the ion currents
present in such a poorly known system as the plasma
membrane of the pollen grain or pollen tube protoplasts.
In accordance with this, distinct anion currents have
recently been measured in lily by patch–clamp
(B. Tavares, P. Dias, P. Domingos, T. Moura, J. A. Feijó,
and A. Bicho, manuscript submitted).
Anion channels and transporters in pollen tubes
Plasma membrane ionic transporters and channels have
long since been associated with the regulation of intracellular ion gradients, with the ion fluxes across the plasma
membrane, with the maintenance of turgor pressure, and
with the furnishing of materials necessary for pollen tube
growth (reviewed in Feijo et al., 2001; Holdaway-Clarke
and Hepler, 2003; Song et al., 2009). Transcriptome analysis
in A. thaliana has shown that there are at least 459 possible
transporter genes expressed during pollen germination and
tube growth. Of these 459 genes at least 8 were found to be
pollen specific (Pina et al., 2005; Wang et al., 2008). Among
the putative transporters found in pollen transcriptome,
Anionic transport in plant cells | 2289
several were putative anion channels. These genes included
two CLC transporters (AtCLC-c, which was pollen
enriched, and AtCLC-d), two SLAC1 homologues (SLAH2
and SLAH3), an ALMT, known to be responsible for the
R-type currents found in guard cells (ALMT12), an
anion:cation symporter (CCC), an anion exchanger, and
a divalent anion:Na+ symporter. Transporters of the ABC
family have also been found, five of which were pollen
enriched and three pollen specific (Pina et al., 2005; Becker
and Feijo, 2007; Song et al., 2009). Even though the
functions of these putative transporters are still unknown,
one must keep in mind that the main Cl– transporter in
mammals, CFTR, is an ABC transporter. The cation:chloride cotransporter (CCC) showed a preferential expression in
the root and the shoot vasculature at the xylem–symplast
boundary, root tips, trichomes, leaf hydathodes, leaf
stipules, anthers, and pollen grains. Plants mutated for the
AtCCC transporter presented shorter organs, inflorescence
necrosis, reduced seed production, and defective Cl–
homeostasis under high-salinity conditions. In Xenopus
laevis oocytes, AtCCC proved to be a 1K+:1Na+:2Cl–
symporter, and its activity was inhibited by bumetanide
(Colmenero-Flores et al., 2007). The physiological profile of
this transporter makes it a good candidate to explain the
influx of Cl– in the pollen tube, although its localization has
yet to be determined. Since the efflux of anions from the
guard cells has been associated with the SLAC1 and
ALMT12 channels, it is possible that the efflux of Cl–
observed at the tip of growing pollen tubes could be due to
the activity of SLAC1 homologues and the ALMT12
channel.
Finally, anionic currents were found in pollen grain
protoplasts from A. thaliana and L. longiflorum. These
currents presented outward rectification, time-dependent
activation, and allowed the passage of current both inwardly and outwardly. They were found to be regulated by
[Ca2+]in. By varying this parameter and also the membrane
potential it is possible to mimic the oscillations that
characterize the Cl– effluxes found at the tip of growing
pollen tubes (B. Tavares, P. Dias, P. Domingos, T. Moura,
J. A. Feijó, and A. Bicho, manuscript submitted).
Final remarks
It is evident that anionic transport plays crucial roles in
different areas of plant biology, namely in signalling pathways, in the control of metabolism, and in the building and
maintenance of electrochemical gradients. In addition to
playing a role in growth itself, ion transporters and other
interacting proteins make certain ions pivotal components
of the cell signalling network by themselves.
The involvement of anionic fluxes in the formation and
growth of the pollen tube has remained controversial since
the first germination studies performed in the 1970s
(Weisenseel and Jaffe, 1976). Similarly, the nature of the
anionic fluxes observed with extracellular electrodes during
the past decade and the reliability of the measuring method
have also been a source of debate, which persists to date,
namely, the contradictory interpretations of Cl– fluxes and
relevance in pollen tubes, where they are thought to play
key roles during polar growth and osmoregulation, call for
a direct analysis of plasma membrane Cl– channel activity.
There is much information that suggests a role for Cl– as
important as that in animal cells, though shared with other
plant ubiquitous anions, like ammonia and NO–3. Despite
major steps in the recent past, most Cl– transport genetic
signatures, namely the CLC family, have proved to be NO–3
channels, Cl– transport in the plasma membrane remaining
unaccounted for. A key issue in the field thus remains to
connect orphan anion transport activities to the corresponding proteins and genes, for a better understanding of
their integrated function in the plant and more specifically
in the pollen tube. Several molecular candidates have been
reported in the pollen transcriptome, and a great effort
in the identification of possible channels underlying the
anionic fluxes is currently being made (Pina et al., 2005).
Future studies will have to take into account: (i) higher
levels of complexity, including the assessment of functional
complementarities of different transporters within a given
gene family; (ii) parallel transport activities between cellular
membrane, and compartments combined with functional
studies; and (iii) protein sequence homology and structure
similarities with orthologous proteins in other organisms.
These studies will increase the knowledge of the function
and regulation systems of plant anion transporters and of
their role in pollen tube cell signalling and metabolism.
Interestingly, selectivity studies indicate a general lack of
anion specificity through anionic transporters, meaning that the
major anion available will demonstrate larger fluxes. This
feature would be most advantageous during pollen tube
growth, since this cell displays continuous and strong ionic
fluctuations, and has to rapidly and efficiently transduce the
resulting signals into the basic features that allow the pollen
tube to correctly target the ovules and discharge the sperm cells.
Finally, in order to construct a complete spatial and
temporal model of signalling systems regulating pollen tube
growth, it has to be borne in mind that transport activity
depends on ion dynamics, e.g. in terms of polarization of
extracellular fluxes and intracellular ion gradients, on the
geometry of the cell, e.g. cell size, shape, cell wall thickness,
spatial distribution of biochemical cell wall components, and
ultimately nutritional demands. This information has to be
integrated with the apical volume, and hydrostatic oscillations, and with other relevant cell mechanical properties.
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