Download Ways of Ion Channel Gating in Plant Cells

Annals of Botany 86: 449±469, 2000
doi:10.1006/anbo.2000.1226, available online at http://www.idealibrary.com on
R E V IE W
Ways of Ion Channel Gating in Plant Cells
E L Z B I E TA K RO L and K A Z I M I E R Z T R E B AC Z *
Department of Biophysics, Institute of Biology, Maria Curie-Skl/ odowska University, Akademicka 19, 20-033 Lublin,
Poland
Received: 12 April 2000 Returned for revision: 7 May 2000
Accepted: 12 June 2000
Published electronically: 21 July 2000
A precise control of ion channel opening is essential for the physiological functioning of plant cells. This process is
termed gating. Ion channel gating can be e€ected by ligand-binding, ¯uctuations in membrane potential, membrane
stretch and light quality. Modern electrophysiological and molecular-biological techniques have enabled the
characterization and classi®cation of many ion channels according to their gating phenomena. Indications are that
gating mechanisms are complex and that individual ion channels can be regulated by a number of factors. In this
paper, gating mechanisms are reviewed following a standard classi®cation of ion channels based on permeability. The
gating of K ‡ , Ca2‡ and anion channels in the plasma membrane, tonoplast and endomembranes of plant cells is
# 2000 Annals of Botany Company
described.
Key words: Review, ion channel, ligand-gating, voltage-gating, stretch-gating, light-gating, plasmalemma, tonoplast.
I N T RO D U C T I O N
Ion channels are integral components of all membranes and
they can be viewed as dynamic ion transport systems
coupled via membrane electrical activities (White et al.,
1999). Not only do they in¯uence membrane voltage
through the ionic currents they mediate, but their activities
can also be regulated by membrane voltage. Ion channels
can be divided into four `historically-based' groups according to gating mechanism: ligand-gated, voltage-gated,
stretch-activated and light-activated. Ligand-gated ion
channels bind intracellular second messengers which provide the essential links between external stimuli and speci®c
intracellular responses (Leckie et al., 1998). Moreover,
additional modulations by ATP or protons make the
channels capable of sensing changes in energy status or
acid metabolism, respectively (Schulz-Lessdorf et al., 1996).
Voltage-dependent channels appear optimally suited for
electrical signal transmission via membrane depolarization
(e.g. through action potentials) and/or for signal transduction in response to changes in membrane potential
(e.g. models investigating the coupling between membrane
potential and voltage-dependent Ca2‡ -channels suggest
that these are engaged in intracellular signalling). They
* For correspondence. E-mail [email protected]
Abbreviations: ABA, Abscisic acid; ABC, ATP binding cassette;
A-9-C, anthracene-9-carboxylic acid; AP, action potential; BL, blue
light; cADPR, cyclic ADP-ribose; CDPK, calmodulin-like domain
protein kinase; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethyl urea; E,
equilibrium potential; I, current intensity; IAA, indol-3-acetic acid; IP3,
inositol triphosphate; NPPB, (5-nitro-2-3-phenylpropylamino) benzoic
acid; OA, okadaic acid; PLC, phospholipase C; PKA, protein kinase
dependent on cyclic AMP; PKC, protein kinase dependent on [Ca2‡ ]cyt
and phospholipids; PKG, protein kinase dependent on cyclic GMP;
TMB-8, 8 (N,N diethylamino) octyl-3,4,5-trimethoxybenzoate.
0305-7364/00/090449+21 $35.00/00
are also involved in membrane voltage stabilization, which
is critical for maintaining ionic gradients and nutritional
ion ¯uxes. Stretch-activated ion channels serve as additional speci®c transmembrane `receptors' co-existing with
other cellular volume-sensing mechanisms. Light-activated
channels are in fact ligand-gated, although a precise
indication of the ligands is not yet possible because the
process of light signal transduction remains unclear. These
channels are distinguished particularly because of a special
importance of light stimuli in plant signalling processes.
Modern biomolecular techniques reveal how complicated
the processes controlling channel behaviour are. It becomes
increasingly apparent that the activity of a channel may
depend on the developmental and metabolic stage of the
cell. Moreover, regulation of ion channels relies not only on
the channel proteins themselves, but also to a great extent
on regulatory polypeptides, such as auxiliary b-subunits,
cytoskeletal components, 14-3-3 proteins, phosphates,
kinases, and G-proteins (Czempinski et al., 1999).
Jan and Jan (1997) recently reviewed receptor-regulated
ion channels in excitable and nonexcitable animal tissues
(G-protein-gated and cGMP-gated K ‡ channels; voltagegated K ‡ -, Na ‡ -, Cl ÿ -, Ca2‡ channels; voltage-insensitive
Ca2‡ channels; Ca2‡ -activated K ‡ channels; ligand-gated
Ca2‡ channels). The activities of these channels are sensitive to external and internal signals that are mediated by
receptors for hormones and transmitters. There are also
plant-derived elicitor-speci®c receptors, which are closely
coupled with plasma membrane ion channels important for
signal transduction in plant cells (Ward et al., 1995;
Blumwald et al., 1998). Studies on receptor-regulated ion
channels suggest that they too are gated via G-proteins,
either by direct protein-protein interaction or indirectly
by kinase (PKA, PKG, PKC)/phosphatase cascades or
# 2000 Annals of Botany Company
450
Krol and TrebaczÐIon Channel Gating in Plant Cells
T A B L E 1. Plant responses controlled by ion channel regulation
Plant response
Reference
Blue- and red-light induced phototropism
Cho and Spalding, 1996; Ermolayeva et al., 1996, 1997; Elzenga and Van Volkenburgh, 1997a;
Lewis et al., 1997; Parks et al., 1998; Suh et al., 1998
Leaf movement
Kim et al., 1992, 1996; Stoeckel and Takeda, 1993, 1995; Moran, 1996; Mayer et al., 1997
Plant excitability
Katsuhara and Tazawa, 1992; Thiel et al., 1993
Light-induced hypocotyl elongation
Sidler et al., 1998
Light-induced transient membrane
potential changes
Trebacz et al., 1994; Elzenga et al., 1995, 1997; Blom-Zandstra et al., 1997; SchoÈnknecht et al.,
1998; Szarek and Trebacz, 1999
Light-induced stomatal opening
Kinoshita and Shimazaki, 1997; Suh et al., 1998
ABA-induced stomatal closure
Armstrong et al., 1995; McAinsh et al., 1995, 1997; Schmidt et al., 1995; Ward et al., 1995;
Li and Assmann, 1996; Blatt and Grabov, 1997a,b; Esser et al., 1997; MacRobbie, 1997;
Mori and Muto, 1997; Pei et al., 1997, 1998; Grabov and Blatt, 1998a; Leckie et al., 1998;
Li et al., 1998; Schwarz and Schroeder, 1998; Barbier-Brygoo et al., 1999
Plant hormone-induced responses
Marten et al., 1991; Hedrich and Jeromin, 1992; Schumaker and Gizinski, 1993;
Blatt and Thiel, 1994; Zimmermann et al., 1994; Ward et al., 1995; Venis et al., 1996;
Claussen et al., 1997; Barbier-Brygoo et al., 1999
Ethylene-mediated responses
Berry et al., 1996
Cold-shock responses
Knight et al., 1996; Lewis et al., 1997
Nod- and pathogen-induced responses
Ward et al., 1995; Zimmermann et al., 1997; Blumwald et al., 1998
Pollination
Holdaway-Clarke et al., 1997; Brownlee et al., 1999
Water and solute transport
Johansson et al., 1996, 1998; Logan et al., 1997; Eckert et al., 1999
Salt tolerance and turgor regulation
Katsuhara and Tazawa, 1992; Taylor et al., 1996; Liu and Luan, 1998; Teodoro et al., 1998;
Brownlee et al., 1999
Cellular pH regulation
Johannes et al., 1998
Proton pump regulation
De Boer, 1997; Claussen et al., 1997; Logan et al., 1997
second messenger binding (Ca2‡ , IP3 , cGMP, cAMP). A
growing body of evidence indicates that G-proteins, second
messengers and phosphorylation/dephosphorylation processes mediate various plant responses through ion channel
and other transport system regulation (Table 1).
Moreover, plant transmembrane receptors resembling
receptor kinases of animal cells are involved in mediating a
variety of cellular processes and responses to diverse
extracellular signals (Braun and Walker, 1996; Trewavas
and Malho, 1997). PCR, advanced homology-based cloning and function-complementation techniques have already
led to identi®cation of more than 70 plant protein kinase
genes (Stone and Walker, 1995). However, the precise
function of speci®c protein kinases and phosphatases
during plant growth and development has been elucidated
in only a few cases (Stone and Walker, 1995).
POTA S S I U M C H A N N E L S
Ion transport across all biological membranes is highly
selective and thus electrochemical potentials can be
generated. The electrochemical potentials largely depend
on the potassium ion gradient, so most of the potassium
channels must remain active for long periods of time. Such
gradients are indispensable for long-term cell functions
such as nutrition, elongation, turgor and water regulation
or osmotically driven movements (Schroeder et al., 1984;
Schroeder, 1989; Roberts and Tester, 1995; Hedrich and
Dietrich, 1996; Logan et al., 1997; Maathuis et al., 1997;
Czempinski et al., 1999).
Ligand-gated potassium channels
Ligand binding causes conformational changes in
channel proteins. It is a process of great importance,
especially during signal transduction cascades when second
messengers synchronize the metabolism of the cell with
environmental conditions and enhance the input stimuli.
There are many K ‡ channels a€ected by calcium ion
‡
channels, KORC,
binding (namely: plasmalemma Kout
NORC, VK, FV, SVÐfor more information see below) in
plant cells (Katsuhara and Tazawa, 1992; Allen and
Sanders, 1996; Czempinski et al., 1997, 1999; Maathuis
et al., 1997; Muir et al., 1997; Allen et al., 1998a). Besides
Ca2‡ , H ‡ ions, nucleotides, proteins and plant hormones
can serve as potassium channel ligands (see below). Their
attachment corresponds accordingly to changes in voltage
sensitivity of voltage-gated K ‡ channels.
Voltage-gated potassium channels in the plasmalemma
Voltage-gated plasmalemma K ‡ channels are generally
‡
‡
‡
† and outward (Kout
) recti®ers. Kin
divided into inward …Kin
channels are activated by hyperpolarizing potentials while
Krol and TrebaczÐIon Channel Gating in Plant Cells
‡
‡
Kout
are activated by membrane depolarization. Both Kin
‡
and Kout channels serve as membrane safeguards preventing membrane voltage from becoming too negative or
positive, respectively. Such a role of voltage-gated K ‡
channels in stabilizing membrane voltages is universal
among all eukaryotes (Maathuis et al., 1997). Voltagedependent plasma membrane-bound outward potassium
recti®ers responsible for K ‡ e‚ux are involved in turgor
regulation (Liu and Luan, 1998), stomatal closure
(MacRobbie, 1997; Grabov and Blatt, 1998a), organ
movements (Iijima and Hagiwara, 1987; Stoeckel and
Takeda, 1993), cation release into xylem (Roberts and
Tester, 1995), light-induced potential changes of the
plasmalemma (Blom-Zandstra et al., 1997) or repolarization during action potentials (APs), and prevention of
excessive depolarization (Stoeckel and Takeda, 1993;
Trebacz et al., 1994; Maathuis et al., 1997). These roles
‡
channels are involved in:
are summarized in Table 2. Kin
potassium uptake into a cell during cell expansion, growth
processes, organ movements and stomatal openings; lowanity uptake pathway in root hair cells; xylem unloading
by conducting cations from xylem to symplast of growing
shoots; membrane voltage prevention against excessive
hyperpolarization (reviewed by Maathuis et al., 1997)
(summarized in Table 2).
Regulation of plasmalemma voltage-gated potassium
channels
In addition to membrane potential, e€ectors like H ‡ ,
Ca2‡ , nucleotides and K ‡ ions can either interact directly
(ligand binding) with both inward and outward plasmalemma K ‡ channels or act indirectly via membrane-bound,
attached or soluble regulators (Hedrich and Dietrich, 1996;
Kurosaki, 1997; Blatt, 1999; Czempinski et al., 1999).
Inwardly and outwardly rectifying K ‡ channels are controlled by cytosolic calcium, ATP and pH in very di€erent
ways (Grabov and Blatt, 1997). The action of pHcyt is most
pronounced on the depolarization-activated outwardrectifying K ‡ channels which are virtually insensitive to
increased [Ca2‡ ]cyt (Grabov and Blatt, 1997). They do not
show such pronounced sensitivity towards external pH but
require slightly alkaline cytosolic pH for activation (Blatt
and Grabov, 1997a). Alkaline pHcyt activates IKout in a
voltage-dependent manner through a co-operative binding
of two protons (Grabov and Blatt, 1997). Moreover,
their activation by depolarization depends critically on
phosphorylation (e.g. by a kinase tightly associated with the
channel protein in Samanea saman motor cellsÐMoran,
1996) or dephosphorylation events associated with [Ca2‡ ]cyt
increase (e.g. by calcium-dependent phosphatase in Arabidopsis thaliana guard cellsÐMacRobbie, 1997). In mesophyll and guard cells of Vicia faba there are outwardrectifying K ‡ channels regulated by calcium and G-protein
interaction as well (Li and Assmann, 1993). On the other
hand, there are potential Ca2‡ -binding sites (EF-hand
motifs) found at the C-terminus of a-subunits from putative
outward potassium recti®ers. These ion channels are very
likely to be directly regulated by Ca2‡ (Czempinski et al.,
1997, 1999). This also applies to KORC and NORC
451
channels which become active at depolarized membrane
potentials, but their respective activation depends on the
cytoplasmic Ca2‡ level (De Boer and Wegner, 1997).
KORC, NORC and SKOR are di€erent channels from
plasmalemma of root xylem parenchyma cells. They are
responsible for xylem loading (Roberts and Tester, 1995;
De Boer and Wegner, 1997; Maathuis et al., 1997; Gaymard
et al., 1998). KORC channels also show a considerable
conductance for Na ‡ but very low permeability for Li ‡ and
Cs ‡ . This indicates that KORC channels may also act as a
`®lter' protecting the shoot from harmful Cs ‡ or Li ‡ ions
(Maathuis et al., 1997). NORC channels discriminate only
slightly between cations and their role in solute release into
xylem is limited. However, they do provide a function in
resetting the membrane potential after excessive depolarization (Maathuis et al., 1997). Kout currents conducted by
SKOR are e€ectively inhibited by both cytosolic and
external acidi®cation (Lacombe et al., 2000). SKOR
channels have no Ca2‡ -binding sites, but they contain
ankyrin and cyclic nucleotide-binding domains (Gaymard
et al., 1998). Direct binding of nucleotides, calcium ions
(De Boer and Wegner, 1997; Czempinski et al., 1997, 1999)
or protons (Blatt and Grabov, 1997a) to the channel
proteins illustrates that voltage-gated outward-rectifying
plasmalemma potassium channels may be regarded as
ligand-gated in certain experimental conditions.
Recently Ca2‡ -gated outward rectifying potassium
channels have been described in the plasmalemma of the
alga Eremosphaera viridis (SchoÈnknecht et al., 1998). These
channels show very steep Ca2‡ -dependence and they can be
Ca2‡ -stimulated both directly and indirectly by interaction
with calmodulin (SchoÈnknecht et al., 1998). They are
involved in hyperpolarizing currents during darkeninginduced transient hyperpolarizations of the plasma
membrane (Table 2).
‡
current is e€ected by [K ‡ ]ext , so that
The gating of Kout
its voltage dependence shifts in parallel with EK (Blatt,
1999). K ‡ ions bind in a co-operative fashion to a set of
sites exposed on the extracellular face of the membrane to
‡
channels and they may be substituted with
inactivate Kout
‡
‡
Rb or Cs (Blatt, 1999). This inactivating binding of
monovalent ions to the channel protein is facilitated by
inside negative membrane voltage. Recent studies have
shown that IKout activation is also dependent on the
cooperative interaction of two K ‡ ions with the channel,
but at sites di€erent from the channel pore (Grabov and
Blatt, 1998a).
Voltage-dependent plant plasmalemma K ‡ -uptakechannels represent various types (KAT, AKT) of di€erent
spatial expression patterns (Bei and Luan, 1998), di€erent
functions (Bei and Luan, 1998; Tang et al., 1998) and
di€erent sensitivities to voltage, Cs ‡ , Ca2‡ and H ‡ (Dreyer
et al., 1997; Bei and Luan, 1998). This diversity partly
results from nonselective heteromerization of di€erent
a-subunits (Dreyer et al., 1997) as well as from the ability
of b-subunits to associate with more than one type of
a-subunit in vivo (Tang et al., 1996, 1998). All voltagedependent plant plasmalemma K ‡ -uptake-channels contain a conserved GYGD motif within a pore region, which
is responsible for K ‡ conductivity (Czempinski et al.,
Voltage-dependence (depolarization activated)
Activated at membrane voltages more positive
than ÿ50 mV
Ca2‡ -dependent activation
K ‡ Rb‡ Na ‡
Cs ‡ Li ‡
K‡
K‡
K‡
out from Samanea saman
motor cells
K‡
out from Mimosa pudica
motor cells
K‡
out from Dionaea muscipula
trap-lobe cells
‡
K‡
out from Conocephalum conicum K
KORC (K ‡ outward rectifying
conductance)
Active at membrane voltages more positive
than ‡30 mV
Ca2‡ -dependent activation
Non-selective
among cations
Non-selective
among cations
K ‡ Na‡
K‡
K‡
K‡
NORC (non-selective outward
rectifying conductance)
Maxi cation channel from
rye roots
K‡
out from Nitellopsis obtusa
K‡
out from Eremosphaera viridis
K‡
out from Nicotiana tabacum
L. mesophyll cells
K‡
out from guard cells of
Vicia faba L.
Stretch-activated
Light-activation
Voltage-dependence
Ca-dependent and stimulated both by direct
Ca2‡ -binding and indirectly by some
calmodulin interactions
Ligand-binding: ATP- and [Ca2‡ ]ext-dependent
regulation (inhibition)
Active at membrane voltages more positive
than EK
Voltage-dependent
Changes in both pHcyt and pHext regulate the
number of channels available for activation
Activation by depolarization
Depolarization-induced activation
Phosphorylation by a kinase tightly associated
with K ‡ out channel
Depolarization-dependent opening stimulated
by Ca-dependent phosphatase
Up-regulated by pHin increase
SKOR (Shaker-type K ‡ outward K ‡
rectifying channel)
K ‡ Na‡
Voltage-dependence (depolarization activated)
Outward recti®cation strongly depends on the
concentration of intracellular K ‡
K‡
K‡
out from Arabidopsis thaliana
guard cells
Depolarization-dependent opening
Up-regulated by pHin increase (strong
voltage-dependent stimulation)
Co-operative binding of two protons
‡
K gradient sensitive
Inhibited by external K ‡ -binding
Regulated by G-protein-induced Ca2‡ -increase
K‡
from Vicia faba guard cell
Potassium channels
K‡
out
Gating mechanism
Permeability
Channel
Volume and turgor regulation and
thereby control of leaf gas exchange
Membrane depolarization upon
light transition
Dark-induced hyperpolarization
of Vm and thereby divalent
cation uptake
Cosgrove and Hedrich, 1991
Blom-Zandstra et al., 1997
SchoÈnknecht et al., 1998
Katsuhara et al., 1990; Katsuhara and
Tazawa, 1992
White, 1998
Membrane voltage stabilization
Salt stress tolerance
Roberts and Tester, 1995; De Boer and
Wegner, 1997; Maathius et al., 1997;
White, 1998
Gaymard et al., 1998;
Lacombe et al., 2000
Roberts and Tester, 1995; De Boer and
Wegner, 1997; Maathius et al., 1997
Trebacz et al., 1994
Iijima and Hagiwara, 1987
Stoeckel and Takeda, 1993
Moran, 1996; Maathuis et al., 1997
MacRobbie, 1997
Li and Assmann, 1993; Blatt and
Grabov, 1997a; Maathuis et al., 1997;
MacRobbie, 1997; Grabov and
Blatt, 1998a; Leckie et al., 1998;
Pei et al., 1998; Blatt, 1999
References
Protection against high
depolarization
Xylem loading
Xylem loading
Xylem loading
Shoot protection from harmful
Cs ‡ and Li ‡ ions
Repolarization during AP
Closure of trap-lobes
Rapid movements in Mimosa
Repolarization during AP
Leaf movements
Involvement in circadian clock
Stomatal closure
Stomatal closure
Prevention from re¯ux of K ‡ into
the guard cell
Physiological role
T A B L E 2. Plasmalemma ion channels
452
Krol and TrebaczÐIon Channel Gating in Plant Cells
Open 60±80 % of the time at voltages more
positive than ÿ120 mV
Inhibited by divalent cations
‡
‡
NH‡
4 , Rb , K ,
Cs ‡ , Na‡ , Li ‡ ,
TEA ‡
K ‡ , Rb‡
K‡
K‡
K‡
K‡
VIC (voltage-insensitive cation
channel)
K‡
in from Zea mays coleoptile
K‡
in from Avena sativa mesophyll
cells
K‡
in from Samanea saman motor
cells
K‡
in from cultured carrot cells
Stretch activated K‡
in from
Vicia faba guard cells
Plasmalemma Vm stabilization
Stabilization of ionic and osmotic
conditions during cell expansion
Cell elongation
Low-anity NH‡
4 -uptake
Osmotic adjustment independent of
the membrane potentials
Compensatory cation ¯uxes
Xylem unloading
Osmoticum gradient-sensitive
Voltage-dependence
Regulated by actin ®laments
Controlled by cytoplasmic concentration of cAMP
Osmoregulation
Membrane changes and thus activation
of voltage-gated channels
Activation by H‡ pump-induced hyperpolarization Leaf movements
Inhibition by PLC-mediated IP3-induced
Ca2‡ increase
Direct response to light
Voltage-dependent
Hyperpolarization-dependent opening
Lowering pHext
Inhibited by Ca2‡
Modulated by auxin
Active at membrane voltages more negative
than ÿ110 mV
K ‡ , Rb‡ , Na ‡ ,
Cs ‡ , Li ‡
KIRC (K ‡ inward rectifying
conductance)
Regulation of membrane voltage
Low-anity K ‡ uptake
Hyperpolarization-dependent opening
Inward K ‡ gradient sensitive
Stomata opening
Regulation of stomatal aperture
Osmotic volume readjustment
AKT1 from Arabidopsis thaliana, K ‡ , Rb‡ , Na ‡ ,
SKT1ÐSolanum tuberosum root Cs ‡ , Li ‡
cells and channel analogue
from corn roots
Hyperpolarization-dependent opening
Lowering pHext promotes K ‡ current in voltagedependent manner
CDPK dependent phosphorylation of KAT1
protein in a Ca2‡ dependent manner
Inhibited by IP3-induced [Ca2‡ ]in elevation
Inhibited by polymerized actin ®laments
Modulated by auxin
Controlled by actin ®laments
Require external K ‡ ions for activation
Modulated by cAMP-dependent signalling system
and/or direct cyclic nucleotide binding
Voltage dependent (hyperpolarization activated)
Stomatal opening
ATP and cGMP activation
K ‡ uptake during other osmotic
movements
Ion permeation may feed back on gating
Competitively inhibited by Ca2‡ and Cs ‡ ions
pH regulated ( pHext acidi®cation shifts
voltage-dependence toward less negative voltages)
Regulation by cytoskeletal proteins
Modulated by cyclic nucleotide binding
K‡
‡
KAT1 from Arabidopsis thaliana K ‡ , NH‡
4 , Rb ,
and KST1 from guard cells and Na ‡ , Li ‡
¯owers of Solanum tuberosum
K‡
in (KAT1) from Vicia faba
guard cell
Table 2 continued on next page
Ramahaleo et al., 1996; Liu and
Luan, 1998
Kurosaki, 1997
Kim et al., 1992, 1996;
Maathuis et al., 1997
Kourie, 1996
Hedrich and Dietrich, 1996;
Thiel et al., 1996; Claussen et al., 1997
White, 1997, 1999
Maathius et al., 1997
Hedrich and Dietrich, 1996; Bertl et al.,
1997; Maathuis et al., 1997;
Czempinski et al., 1999
Armstrong et al., 1995; MuÈller-RoÈber
et al., 1995; Becker et al., 1996; Hedrich
and Dietrich, 1996; Hoth et al., 1997;
Maathuis et al., 1997;
Czempinski et al., 1999
Blatt et al., 1990; Fairley-Grenot
and Assmann, 1992; Blatt and Thiel,
1994; Wu and Assmann, 1995;
Ilan et al., 1996; Blatt and Grabov,
1997a; Claussen et al., 1997; Grabov and
Blatt, 1997, 1998a; Hwang et al., 1997;
Maathuis et al., 1997; MacRobbie, 1997;
McAinsh et al., 1997; Leckie et al., 1998;
Li et al., 1998; Liu and Luan, 1998;
Pei et al., 1998; Blatt, 1999;
Czempinski et al., 1999; Jin and
Wu, 1999
Krol and TrebaczÐIon Channel Gating in Plant Cells
453
Stretch-activated
Regulated by cytoskeletal proteins
Ca2‡
Mechanosensitive Ca-channels
from guard cells
Stretch-activated
Stretch-activated
Regulated by cytoskeletal proteins
Non-selective
Non-selective
Voltage-dependent
Stretch-activated
Ca2‡
VDCC from Fucus rhizoids
SAC from Fucus zygotes
Voltage-dependent
Stretch-activated
Ca2‡
VDCC from pollen tubes
Mechanosensitive Ca-channels
from root cells
Hyperpolarization-activated
Ca2‡
Depolarization activated
Voltage-dependent Ca-channels Ca2‡
from liver wort Conocephalum
conicum and moss
Physcomitrella patens
VDCC from Mimosa pudica
motor cells
Depolarization activated
Ca2‡
VDCC from Chara corallina
Cytokinin-induced depolarization activated
AP induction
Early events of turgor regulation and
salt tolerance
Depolarization activated
Ca2‡
VDCC from characean cells
Ca2‡
Cation uptake
Thion et al., 1996; White et al., 1998
Maintaining appropriate electrochemical
gradients important for the transport
of other ions and cell volume
regulation
Signalling mechanisms and priming
the cell for response
Ba2‡ , Sr2‡ , Ca2‡ , Depolarization activated
Mg2‡ , K ‡
Active under condition of microtubule
disorganization
Slow inactivation at negative voltages
VDCC from Arabidopsis roots
and Daucus carota suspension
protoplasts
Ca-channels from mosses
Divalent cation uptake into roots
Signalling mechanisms and priming
the cell for response
Ba2‡ , Sr2‡ , Ca2‡ , Depolarization activated
Mg2‡ , Mn2‡ , K ‡ , Strong voltage-dependence (depolarization
activated)
Na ‡ , Rb‡ , Li ‡
Cytosolic ATP shifts activation to more negative
potentials
Reid et al., 1997
Thion et al., 1996; White et al., 1998
Taylor et al., 1996
Holdaway-Clarke et al., 1997
Stoeckel and Takeda, 1995
Schumaker and Gizinski, 1993
Transmission of Ca-signals into
Cosgrove and Hedrich, 1991;
the cytoplasm
MacRobbie, 1997; McAinsh et al., 1997
Guard cell volume and turgor regulation
and thereby control of leaf gas
exchange
Control of other ion channels with
Ca-dependent activities
Regulation of turgor
Determination of the allometry of cell
expansion and morphogenesis
Growth processes
Growth processes
Activation of channels involved in
leaf movements
Early events of cytokinin-induced
responses
Light-induced membrane depolarization Trebacz et al., 1994; Ermolayeva et al.,
1996, 1997
During Ca-starvation channels might
open to scavenge available Ca2‡
Katsuhara and Tazawa, 1992;
Shimmen, 1997
White, 1998; Pineros and Tester, 1997;
White, 1998
McAinsh et al., 1995; Grabov and
Blatt, 1998b
VDCC from rye roots
VDCC from wheat roots
(rca channel)
Early events of plant hormone-induced
responses
Depolarization activated
Ca2‡
References
VDCCÐvoltage-dependent
Ca-channel from guard cells
Calcium channels
Physiological role
Gating mechanism
Permeability
Channel
T A B L E 2. Continued
454
Krol and TrebaczÐIon Channel Gating in Plant Cells
S-type shows weak voltage dependence
S-type serves as major pathway for
S-type requires hydrolysable ATP and activation of
anion e‚ux during stomatal closure
protein kinase
and as negative feedback during
OA-sensitive phosphatases are involved in
stomatal opening
down-regulation of S-type channel
R-type responsible for signal
S-type may be ABC protein or it is tightly controlled
transduction via membrane
by such protein
depolarization
R-type is activated by parallel voltage membrane
GCAC channels are capable of sensing
2‡
depolarization, pHcyt acidi®cation, [Ca ]cyt
changes in the energy status, acid
increase and nucleotide binding
metabolism and proton pump activity
Direct auxin binding shifts activation potential
in guard cells, because of time- and
towards resting potentials to favour channel
voltage dependent activity strongly
opening
modulated by ATP and H ‡
Cl ÿ , malate
GCAC1 from Vicia faba and
Commelina communis
Voltage-dependent (hyperpolarization activated)
Two mode kinetics di€erently controlled by ATP
(R- and S-type, S-type occurs in the presence
of ATP)
Ca-dependent activation
Voltage-dependent (hyperpolarization activated)
Voltage-dependent inactivation under large
hyperpolarization
Cl ÿ
Anion channels from mesophyll
cells of Pisum sativum
Anion channels from suspension- Cl ÿ
cultured carrot cells
Control of membrane potential
Regulation of osmotic balance
Light-induced transient depolarization
ATP-controlled voltage-dependence (depolarization Anion release during inhibition of
activated)
cell elongation
Modulated by auxin
Cl ÿ
S-type shows weak voltage dependence, requires
protein phosphatase activities and is downregulated by protein kinase
S-type is in¯uenced by pH gradient
R-type is activated by parallel voltage membrane
depolarization, pHcyt acidi®cation, [Ca2‡ ]cyt
increase and nucleotide binding
R-type is modulated by phosphorylation/
dephosphorylation processes
Blumwald et al., 1998
Zimmermann et al., 1997;
White et al., 1998
Taylor et al., 1996; McAinsh et al., 1997
Table 2 continued on next page
Barbara et al., 1994
Elzenga and Van Volkenburgh, 1997a,b
Zimmermann et al., 1994
Armstrong et al., 1995; Ward et al., 1995;
Schulz-Lessdorf et al., 1996; Elzenga and
Van Volkenburgh, 1997b;
Lewis et al., 1997; Grabov and
Blatt, 1998a; Pei et al., 1997, 1998
Keller et al., 1989; Marten et al., 1991;
Hedrich and Jeromin, 1992;
Linder and Raschke, 1992;
Schroeder and Keller, 1992;
Schroeder et al., 1993; Dietrich and
Hedrich, 1994; Schmidt et al., 1995;
Ward et al., 1995; Li and Assmann, 1996;
Esser et al., 1997; Mori and Muto, 1997;
Pei et al., 1997, 1998; Grabov and
Blatt, 1998a; Schwarz and
Schroeder, 1998; Leonhardt et al., 1999
Ca-in¯ux as an early response to various Gelli and Blumwald, 1997
signals including fungal elicitors
Early events of pathogen defence
system activation
TSACÐtobacco suspension-cell
anion channel
GCAC1 from Nicotiana
benthamiana and Arabidopsis
thaliana
Elicitor-activated
Hyperpolarization-activated
Ca2‡ , K ‡
Receptor-regulated Ca- from
tomato protoplasts
Anion channels
Elicitor-activated
Voltage-gated
Ca2‡
Receptor-regulated Ca-channels
Early events of pathogen defence
system activation
Elicitor-activated
Non-selective
Receptor-regulated Ca-channels
from parsley protoplasts and
root cells
Transmission of Ca-signals into
the cytoplasm
Cell volume regulation
Stretch-activated
Regulated by cytoskeleton proteins
Ca2‡
Mechanosensitive Ca-channels
from Fucus rhizoids
Krol and TrebaczÐIon Channel Gating in Plant Cells
455
Non-selective
SAC from guard cells of
Vicia faba L.
SAC from Arabidopsis thaliana
guard cells
Cl ÿ
Stretch-activated
Light-induced activation (increase in
open probability)
Ca-dependent activation
BL-activation (increase in open probability)
Anion channels from epidermal Cl ÿ
cells of Arabidopsis hypocotyls
Anion channels from mesophyll
cells of Pisum sativum
Strong and weak voltage-dependence of R- and
S-type unitary conductances, respectively
(activation by Vm depolarization)
R- and S-types have the same conductance but
di€erent open probabilities
The switch between R- and S-type is controlled by
ATP (R-type occurs in the presence of ATP)
Modulated by phosphorylation/dephosphorylation
processes
Cho and Spalding, 1996
Thomine et al., 1995, 1997; Cho and
Spalding, 1996; Elzenga and Van
Volkenburgh, 1997b; Lewis et al., 1997;
Parks et al., 1998
Johannes et al., 1998
Ermolayeva et al., 1996, 1997
Reduction of cell turgor
Activation of voltage-dependent ion
channels through membrane
depolarization
Control of leaf gas exchange
Teodoro et al., 1998
Cosgrove and Hedrich, 1991
Elzenga et al., 1995, 1997; Elzenga and
Light-induced transient membrane
Van Volkenburgh, 1997a,b
potential depolarization
Charge balance for light-induced H ‡
pump activation, thus control of
pHext , membrane voltage and osmotic
potential
Light-induced inhibition of cell
elongation
R-type may be involved in the
transduction of external signals and
transmission of AP
S-type may be involved in turgor
regulation and hypocotyl movements
H ‡ - and Ca2‡ -dependent activation (direct binding) Facilitation of enhanced proton e‚ux
Phosphorylation/dephosphorylation processes
under intracellular acidosis
Cl ÿ
Anion channels from
Charophyta cells
Anion channels from epidermal Cl ÿ
cells of Arabidopsis hypocotyls
Ca-dependent activation
Cl ÿ
Anion channels from
Physcomitrella patens
Phytochrome-mediated signalling
pathway
Okihara et al., 1991; Katsuhara and
Tazawa, 1992; Thiel et al., 1993;
Shimmen, 1997
Iijima and Sibaoka, 1985
Depolarizing current during AP
Anion channels from Aldrovanda
vesiculosa
Ca-dependent activation
Cl ÿ
Anion channels from
Charophyta cells
Limiting the amplitude of dark-induced SchoÈnknecht et al., 1998
transient hyperpolarization caused by
K ‡ -release
References
Trebacz et al., 1994
Hyperpolarization activated
Cl ÿ
Anion channels from
Eremosphaera viridis
Physiological role
Anion channels from liverwort
C. conicum
Gating mechanism
Permeability
Channel
T A B L E 2. Continued
456
Krol and TrebaczÐIon Channel Gating in Plant Cells
Krol and TrebaczÐIon Channel Gating in Plant Cells
1999). Kourie (1996) demonstrated that the relative number
of opened voltage-activated inward rectifying potassium
channels increased sigmoidally as a function of hyperpolarized membrane potential. The kinetics of inward
rectifying K ‡ currents in Avena sativa mesophyll cells
reported by Kourie was independent of [K ‡ ]ext and it lacked
time-dependent inactivation. Neither low [K ‡ ]ext nor
[Na ‡ ]ext caused inactivation of the above-mentioned
currents, while Cs ‡ -induced block was reversible and
strongly voltage-dependent. A role of preventing large
membrane hyperpolarization resulting from electrogenic
‡
channels by
proton pumping was proposed for these Kin
Kourie (1996). On the other hand, there are reports
‡
channels (AKT1) `sensing' external potasconcerning Kin
sium concentration (Bertl et al., 1997). AKT1 channels are
present in root cells. Extracellular K ‡ binds to a modulator
site thereby enhancing the rate of opening of AKT1 protein.
Blatt (1999) also noticed that IKin current in guard cells
requires external millimolar K ‡ concentrations for its
‡
channels appear to
activity. In submillimolar [K ‡ ]ext , Kin
enter a long-lived inactive state (Blatt, 1999).
‡
channels is modulated by
Control of plasmalemma Kin
2‡
increasing [Ca ]cyt (inactivation) and increasing external
proton concentration (voltage-dependent activation) or
decreasing pHcyt (voltage-independent activation; increase
in the pool of active channels through allosteric interaction)
(Ilan et al., 1996; Grabov and Blatt, 1997, 1998a; Hoth
et al., 1997; MacRobbie, 1997). Ca2‡ -dependent inactivation can proceed even when pH is bu€ered. Equally,
changes in pH and channel gating may occur without
measurable changes in calcium concentration (Allan et al.,
1994; Armstrong et al., 1995). Thus, the e€ects of cellular
pH and calcium are separable, although these two ionic
messengers do interact. In other words, pHcyt may act in
parallel with, but independently of, [Ca2‡ ]cyt in controlling
‡
channels (Grabov and Blatt, 1997). Kim et al. (1996)
Kin
reported that phosphoinositide turnover, phospholipase C
(PLC) activation or the presence of inositol triphosphate
‡
channel closure. Earlier, Blatt
(IP3) is correlated with Kin
‡
et al. (1990) demonstrated the possibility of controlling Kin
2‡
channel activity by IP3-mediated Ca release. Both the
above-mentioned results indicate that increase in [Ca2‡ ]cyt is
‡
channel inactivation and they support a
responsible for Kin
growing body of evidence that G-proteins function in
regulating IKin (reviewed by Blatt and Grabov, 1997a,b).
Recently, Li et al. (1998) identi®ed a Ca2‡ -dependent
protein kinase, with a calmodulin-like domain (CDPK),
‡
channels of Vicia faba guard cell
which phosphorylates Kin
protoplasts. Moreover, the cAMP-dependent signalling
‡
system `cross-talks' with Ca2‡ -dependent inhibition of Kin
channels from Vicia faba guard cells by reversing inhibitory
‡
calcium e€ects (Jin and Wu, 1999). In contrast to Kin
‡
channels from guard cells, Kin channels in the plasmalemma of rye root cells are insensitive to [Ca2‡ ]cyt (White,
1997).
‡
channels (KAT1ÐArabidopsis thaliana, KST1Ð
Kin
Solanum tuberosum) can also be inhibited by Ca2‡ and Cs ‡
via competition in binding to the pore forming region
exposed to the aqueous lumen of the channel (Becker et al.,
1996). Thiel et al. (1996) showed that Ca2‡ -binding to the
457
K ‡ channel protein is responsible for fast and reversible
inactivation of inward K ‡ currents in maize coleoptile
protoplasts.
In addition to their Ca2‡ and pH dependence, voltage‡
channels seem also to require ATP
gated plasmalemma Kin
(Hoshi, 1995; MuÈller-RoÈber et al., 1995; Wu and Assmann,
1995). Their structures contain ATP and cyclic nucleotidebinding cassettes in the C-terminal domains. The rundown
‡
recti®ers in the absence of ATP is explained in terms
of Kin
of a shift in the voltage-dependence (Hedrich and Dietrich,
1996). Kurosaki (1997) surveyed some of the inward K ‡
channels (located in the plasma membrane of cultured
carrot cells) whose gating was controlled by cytoplasmic
concentration of cAMP. Their activation resulted in
transient membrane potential changes, which in turn
activated voltage-gated Ca2‡ channels. Because plasmalemma voltage-gated inward K ‡ -channels described by
Hedrich and Dietrich (1996) and Kurosaki (1997) are
regulated via direct nucleotide binding to the channel
protein, they can be classi®ed as ligand-gated ones as well.
There is an obvious correlation between inward rectifying
K ‡ channels and cytoskeletal proteins (Table 2). As a
conserved structural feature, proteins of the AKT subfamily
contain so-called ankyrin repeats which are potential
domains for interaction with the cytoskeleton (Czempinski
et al., 1999). Because proteins from the KAT subfamily lack
such ankyrin sequences, but they are `sensitive' to
cytoskeletal drugs, there must be other channel domains
participating in the regulation by cytoskeletal compounds
(Hwang et al., 1997; Czempinski et al., 1999). Pharmacological studies on guard cells have shown that actin
‡
channels as well
®laments contribute to regulation of Kin
as of stomatal aperture (Hwang et al., 1997). Cytochalasin
D, which induces depolymerization of actin ®laments,
activates inward potassium currents, while phalloidinÐa
stabilizer of ®lamentous actinÐinhibits them (Hwang et al.,
1997). These authors demonstrated that polymerized actin
‡
channels in the closed state and thus makes
stabilizes Kin
them unresponsive to membrane hyperpolarization. As
‡
actin ®laments depolymerize, the closed state of Kin
channels becomes less stable and more channels become
ready to respond to the hyperpolarized membrane potential. Liu and Luan (1998) also correlated regulation of IKin
with the pattern of organization of actin ®laments. They
stated that actin structure may be a critical component in
‡
channels in
the osmosensing pathway conducted by Kin
plants.
There are also reports of auxin-induced modulation of
K ‡ -inward recti®ers at the plasma membrane in coleoptile
cells (Claussen et al., 1997) and guard cells (Blatt and Thiel,
1994) (Table 2).
Voltage-gated potassium channels in the tonoplast
In the tonoplast of higher plants there are three distinct
kinds of voltage-sensitive potassium channels (FV, fast
activating; SV, slow activating; and VK, strongly K ‡ selective) (Table 3). FV channels are instantaneously activated at
the resting levels of [Ca2‡ ]cyt and pHcyt by changes in
tonoplast voltage (Allen et al., 1998). They open at cytosol
Multi-cation
Cation-selective channel from
nuclear envelope from red beet
Ca-regulated voltage-dependence
Voltage-dependent
ATP-regulated
Modulated by Cs ‡ , Mg2‡
K‡
Cation-selective channel from
chloroplast envelope
Ca-regulated pathways for nuclear
processes
Compensation of light-driven
proton movements
Metabolite di€usion
Voltage-dependent
Osmotic volume and turgor
regulation
Multi-cation
Activated by micromolar [Ca2‡ ]cyt
Voltage dependence
Strong pH-dependence (inhibition by acidic pH)
K ‡ , Na ‡
Cation-selective channel from
tonoplast of algae
Lamprothamnium, Chara
buckellii, Chara australis and
Nitellopsis obtusa
Ca-dependent potassium uptake and
release during stomatal
movements (e.g. ABA-induced
stomatal closure)
Activation of voltage-gated
tonoplast channels
Cation-selective channel from
chloroplast envelope
Activated by micromolar [Ca2‡ ]cyt and acidic pHcyt
Voltage-independent, non-rectifying channel
K ‡ , Rb ‡ ,
NH4‡
Tonoplast VK (vacuolar K ‡ )
channels
Vacuolar receptor site for calcium
during stomatal closure
Possible participation in CICR
(Ca-induced Ca-release)
Turgor regulation
Vacuolar ion transport
Voltage-gated (activated by positive membrane potentials Compensation of light-induced
of stroma relative to lumen)
proton ¯uxes
Time-dependent activation at cytosol-positive potentials
Outward-rectifying
Strong voltage-dependence modulated by Ca2‡ , Mg2‡
and H ‡ ions (Ca- and Mg-activation and downregulation of SV channel activity by protons)
Ca2‡ induces lowering of the voltage threshold for
activation
Require alkaline pH at both sides of tonoplast
Regulated by protein phosphorylation and calmodulin
interaction
Single channel conductance dependent on [K ‡ ]cyt
Modulated by redox agents (increased open probability
in the presence of antioxidants)
Blocked by polyamines in a voltage-dependent manner
K ‡ , Na ‡ ,
Rb‡ , Li ‡ ,
NH4‡ , Ca2‡ ,
Mg2‡ ,
polyamines
Tonoplast SV (slow-activating)
cation channels
Control of the tonoplast electrical
potential di€erence around EK
A shunt conductance for the
vacuolar H ‡ pumps
Involvement in potassium release
during stomatal closure
Involvement in increase in cellular
osmolarity
Small monovalent cation uptake
Physiological role
Cation-selective channel from
K ‡ , Ca2‡ ,
thylakoids of Spinacea oleracea Mg2‡
and Pisum sativum cotyledons
Voltage-dependent open probability
Preferred outward recti®cation at positive potentials
(relative to the cytoplasm)
Active at the resting levels of [Ca2‡ ]cyt and pHcyt
Inhibited by vacuolar and cytosolic Ca-increases
FV currents are reduced at acidic pHcyt
ATP regulated
Blocked by Mg2‡ and polyamines
NH4‡ , K ‡ ,
Rb‡ , Cs ‡ ,
Na‡ , Li ‡
Tonoplast FV ( fast-activating)
cation channels
Potassium channels
Gating mechanism
Permeability
Channel
T A B L E 3. Ion channels in plant endomembranes
Grygorczyk and Grygorczyk, 1998
Heiber et al., 1995
Heiber et al., 1995
Pottosin and SchoÈnknecht, 1996;
Hinnah and Wagner, 1998
Katsuhara and Tazawa, 1992;
LuÈhring, 1999
Ward et al., 1995; Allen and Sanders, 1996,
1997; Maathuis et al., 1997;
MacRobbie, 1997; McAinsh et al., 1997;
Allen et al., 1998a; Grabov and Blatt, 1998a
Ward and Schroeder, 1994; Allen and
Sanders, 1995, 1996, 1997; Schulz-Lessdorf
and Hedrich, 1995; Ward et al., 1995;
Gambale et al., 1996; Bethke and
Jones, 1997; Maathuis et al., 1997;
MacRobbie, 1997; McAinsh et al., 1997;
Allen et al., 1998b; Grabov and Blatt,
1998a; Carpaneto et al., 1999; Cerana et al.,
1999; Dobrovinskaya et al., 1999
Linz and KoÈhler, 1994; Allen and Sanders,
1996, 1997; Maathuis et al., 1997;
Tikhonova et al., 1997; Allen et al., 1998a;
Grabov and Blatt, 1998a; BruÈggemann et al.,
1999a,b; Dobrovinskaya et al., 1999
References
458
Krol and TrebaczÐIon Channel Gating in Plant Cells
Voltage-dependent
Ca2‡ -gradient sensitive
Anion channels
Vacuolar malate uptake
Intracellular Ca2‡ -release during
responses to mechanical stimuli
Metabolite di€usion
Compensation of light-driven
proton movements
Non-selective Voltage-dependent
Voltage-dependent
Voltage-dependent
Cl ÿ
Cl ÿ
Anion channel from outer
envelope of chloroplasts
Anion channel from inner
envelope of chloroplasts
Anion channel from thylakoids
Compensation of light-driven
proton movements
Anion uniport
Non-selective pH-regulated (activated by low matrix pH)
Inner membrane anion channel
from mitochondria
Turgor regulation
Control of mitochondrial membrane
potential
Control of ATP di€usion
Control of signal transduction
Outward rectifying Ca-dependent regulation
Cl ÿ , NO3ÿ
Vacuolar anion channel from
Characean cells
Anion uptake during stomatal
opening
VDAC (voltage-dependent anion Non-selective Voltage-dependent
channels in outer membrane of
pH-dependent
mitochondria)
Second messenger-binding
Activation by tonoplast hyperpolarization
Channel activation depends on protein phosphorylation
Inward-recti®cation
Cl ÿ
Activation by tonoplast hyperpolarization (negative
Vacuolar anion uptake
potentials relative to the cytoplasm)Ðinward-recti®cation
Vacuolar VCl from Vicia faba
guard cells
Vacuolar chloride channelÐVCl Cl ÿ , NO3ÿ ;
SO2ÿ
4
Vacuolar malate channelÐVMal Malate
Activation by potentials more negative than EMal
from Arabidopsis thaliana
fumarate,
Strong inward recti®cation because of luminal Cl ÿ
ÿ
vacuoles
acetate NO3 ; blockade of malate re-entry
H2 PO4ÿ
Ca-channel from ER of Bryonia Ca2‡
diodica tendrils
Voltage-dependent (hyperpolarization activated)
Intracellular Ca2‡ -release
pH-sensitive
Require two Ca2‡ ions binding to open
Luminal Ca2‡ shifts the threshold for voltage activation to
less negative potentials
Inhibited by [Ca2‡ ]cyt increases
Ca2‡ , K ‡
VVCa (voltage-gated Cachannels from vacuoles of Vicia
faba guard cells and Beta
vulgaris roots)
Ca2‡ -release during signal
transduction
Activation by cADPR-binding
Ca2‡
Ligand-gated Ca-channel from
alga Eremosphaera viridis
Ca2‡ -release during signal
transduction
Activation by cADPR-binding
Ca2‡ -release during signal
transduction
Ca2‡ , K ‡
Activation by IP3-binding
Ca-current recti®cation over physiological tonoplast
potentials (cytosol negative with reference to lumen)
Ligand-gated Ca-channel in
vacuoles from red beets and
cauli¯ower ¯orets
Ligand-gated Ca-channel from
Ca2‡
vacuole and ER of cauli¯ower
and vacuoles of guard cells,
zucchini hypocotyls, oat roots,
carrot and red beet roots, mung
bean hypocotyls, maize cells
Calcium channels
Heiber et al., 1995; Pottosin and
SchoÈnknecht, 1995
Heiber et al., 1995; Fuks and Homble, 1999
Heiber et al., 1995; Pohlmeyer et al., 1998
Beavis and Vercesi, 1992
Elkeles et al., 1997; Mannella et al., 1997,
1998; Rostovtseva and Colombini, 1997;
Green and Reed, 1998; Song et al., 1998;
Shimizu et al., 1999
Katsuhara and Tazawa, 1992
Pei et al., 1996; Grabov and Blatt, 1998a
Allen and Sanders, 1997
Cerana et al., 1995; Allen and Sanders,
1997; Chengs et al., 1997
KluÈsener et al., 1995
Allen and Sanders, 1994, 1997; Johannes
and Sanders, 1995; McAinsh et al., 1997;
Pineros and Tester, 1997
Bauer et al., 1998
Allen et al., 1995; Muir and Sanders, 1996;
Allen and Sanders, 1997; McAinsh et al.,
1997; Muir et al., 1997; Leckie et al., 1998
Muir and Sanders, 1996, 1997; Allen and
Sanders, 1997; Muir et al., 1997; Leckie
et al., 1998; MacRobbie, 1997; McAinsh
et al., 1997
Krol and TrebaczÐIon Channel Gating in Plant Cells
459
460
Krol and TrebaczÐIon Channel Gating in Plant Cells
positive vacuolar membrane potential for longer times than
at negative potentials and hence they mainly allow K ‡ and
NH4‡ e‚ux from the cytoplasm into the vacuole ( preferred
outward recti®cation) (Tikhonova et al., 1997; BruÈggemann
et al., 1999b). Their function is to control the electrical
potential di€erence across the tonoplast (Tikhonova et al.,
1997). Vacuolar Ca2‡ suppresses FV channels in a voltagedependent manner while cytosolic Ca2‡ blocks them in a
voltage-independent manner (Allen and Sanders, 1996;
Tikhonova et al., 1997; Allen et al., 1998a). One of the most
pronounced features of FV channels is their blockade by
Mg2‡ . Increasing cytosolic free Mg2‡ decreases the open
probability of FV channels without a€ecting single current
amplitudes (BruÈggemann et al., 1999a). FV currents were
also shown to be reduced at acidic pHcyt (Linz and KoÈhler,
1994) or by cytosolic polyamines (Dobrovinskaya et al.,
1999). Recent studies on FV currents in red beet vacuoles
indicate that FV channels may be ATP regulated (Allen
et al., 1998a).
SV channels are strictly outward rectifying, cation
selective and they show characteristics typical of a multiion pore, i.e. more than one ion can occupy the channel
pore at the same time (Allen and Sanders, 1996). They
display time-dependent activation at cytosol-positive potentials and when [Ca2‡ ]cyt is higher than approx. 0.5 mM
(Schulz-Lessdorf and Hedrich, 1995; Allen and Sanders,
1996). Calcium and protons modulate the voltagedependence of SV channels (Schulz-Lessdorf and Hedrich,
1995). These two cations interact strongly with the voltage
sensor without changing the unitary conductance. The
open probability of the SV-type channel is a function of
[Ca2‡ ]cyt (Gambale et al., 1996). Schulz-Lessdorf and
Hedrich (1995) showed that there is a regulatory Ca2‡ binding site on the cytoplasmic face of the SV channel and
that calmodulin may be involved in the modulation of the
activation threshold of the SV-type channel. This is in
agreement with recent results of Bethke and Jones (1997)
who examined SV currents stimulated by both calmodulinlike domain protein kinase (CDPK) and okadaic acidsensitive phosphatases. On the other hand, Ca2‡ -dependent
protein phosphatase can induce the inhibition of SV
channels (Allen and Sanders, 1995). Bethke and Jones
(1997) proposed a model in which SV channel activity is
regulated by protein phosphorylation at two sites. In the
absence of calcium ions, Mg2‡ can activate SV currents
(Allen and Sanders, 1996; Cerana et al., 1999). Moreover,
the single channel conductance increases as a function of
the potassium concentration (Gambale et al., 1996). This
behaviour can be explained by a multi-ion occupancy
mechanism. However, at negative transtonoplast voltages,
the closure of SV channels is una€ected by either Ca2‡ or
Mg2‡ , indicating that the channel belongs to the voltagegated superfamily (Cerana et al., 1999). SV channels are
also reversibly activated by a variety of sulphydryl reducing
agents at the cytoplasmic side of the vacuole (Carpaneto
et al., 1999). Increase in the open probability in the presence
of antioxidants may correlate ion transport with other
crucial mechanisms that in plants control turgor regulation,
response to oxidative stresses, detoxi®cation and resistance
to heavy metals (Carpaneto et al., 1999).
Cytosolic polyamines are strong inhibitors of SV
channels, but in contrast to the inhibition of FV channels,
the blockage of SV channels displays a pronounced voltagedependence (Dobrovinskaya et al., 1999). Hence,
polyamine-blockage is relieved at a large depolarization
(because of the permeation of polyamines through the
channel pore) and in the presence of high concentrations of
polyamines the slow vacuolar channels are converted into
inward recti®ers (Dobrovinskaya et al., 1999).
VK channels are non-rectifying and are activated at
micromolar [Ca2‡ ]cyt and acidic pHcyt by tonoplast potentials ranging from ÿ100 to ‡60 mV (Allen and Sanders,
1996; Allen et al., 1998a). Therefore, they can be involved in
vacuolar potassium uptake and loss. So far their presence
has been proved only in guard cells.
Di€erent sensitivities of FV-, SV- and VK-channels to
[Ca2‡ ]cyt and pH may provide a mechanism whereby stimuli
activating various signalling pathways can generate
vacuolar ion uptake or loss. Muir et al. (1997) concluded
that this di€erential regulation of vacuolar channels by
Ca2‡ represents a downstream event in signal transduction
cascades induced by Ca2‡ -release. SV channels are thought
to participate in signalling processes because of their ability
to release Ca2‡ after Ca2‡ -dependent activation (CICRÐ
Ca2‡ -induced Ca2‡ -release) (Allen et al., 1998b). However,
Pottosin et al. (1997) demonstrated that the SV channel is
not suited for CICR from vacuoles, at least in the case of
barley mesophyll cells. Thus, the physiological role of SV
channels remains a matter for discussion.
‡
-channel
The most frequently observed voltage-gated Kin
in the tonoplast of Chara was examined by LuÈhring (1999)
(Table 3). Acidi®cation on both sides of the membrane
decreases open probability of the channel and changes its
voltage-dependence, most probably through protonation of
negatively charged residues in neighbouring voltage-sensing
transmembrane domains (LuÈhring, 1999). The channel
behaves like animal maxi-K channels and its gating kinetics
responds to cytosolic Ca2‡ . Under natural conditions,
pH changes contribute mainly to channel regulation at the
vacuolar membrane face (LuÈhring, 1999).
Voltage-gated potassium channels in other intracellular
membranes
Heiber et al. (1995) showed that the chloroplast envelope
contains voltage-dependent cation channels (Table 3) with
complex gating behaviour and subconductace states, as well
as cation-selective pores with high conductances. Voltagedependent cation channels favour potassium uptake and
their gating is a€ected by monovalent cations (Cs ‡ ),
divalent cations (Mg2‡ ) and millimolar concentrations of
ATP. Hinnah and Wagner (1998) observed potassium
selective pore-like channels in osmotically swollen thylakoids from pea protoplasts derived from cotyledons of
young Pisum sativum plants (Table 3). There is also a
nonselective (PK 4 PMg 4 PCa) cation channel in native
spinach thylakoid membranes (Table 3) found by Pottosin
and SchoÈnknecht (1996). This cation channel displays
bursting behaviour and its open probability increases at
positive membrane potentials (Pottosin and SchoÈnknecht,
Krol and TrebaczÐIon Channel Gating in Plant Cells
1996). It has only a moderate voltage-dependence compared to classical voltage-dependent recti®ers. It is postulated that its function is to compensate the light-driven
proton uptake into thylakoids (Pottosin and SchoÈnknecht,
1996).
A Ca2‡ - and voltage-dependent non-speci®c channel was
found in the nuclear envelope of red beet (Grygorczyk and
Grygorczyk, 1998) (Table 3). Micromolar [Ca2‡ ] on the
nucleoplasmic side of the envelope activates this cation
channel. The channel voltage-dependent activity changes
with the nucleoplasmic calcium concentration. Such a
channel may provide a Ca2‡ -regulated pathway for Ca2‡ dependent nuclear processes (e.g. gene transcription).
Plasmalemma voltage-insensitive cation channels (VIC)
The VIC channels are responsible for an in¯ux of a range
of monovalent cations into cereal root cells (Table 2). It has
been postulated that they could contribute to low-anity
NH4‡ uptake and rapid osmotic adjustment independent of
membrane potential. They may also compensate electrogenic cation ¯uxes (White, 1999). Under saline conditions
‡
channels play a major role in
VIC channels along with Kin
the toxic Na ‡ in¯ux across the plasma membrane (White,
1999). Inward currents through the VIC channels are
inhibited by Ca2‡ and Ba2‡ .
Stretch-activated potassium channels
Changes in turgor pressure induced by hyper- or hypoosmotic stress induce an early change in activities of stretchsensitive channels. Stretch-activated channels (SACs) also
respond when mechanical forces are exerted on the cell
(Ramahaleo et al., 1996). For the translation of membrane
stretch into channel gating it is generally argued that
attachment of membrane proteins to tension-transmitting
components is necessary, by linkage to cell wall proteins, or
cytoskeletal proteins, or both (MacRobbie, 1997). Anionic,
cationic, as well as non-selective SACs, have been reported
to occur in plasma membranes (Table 2). There is a
growing body of evidence for involvement of stretchactivated ion channels in regulation of the response of
guard cells to ABA through interactions with the cytoskeleton (MacRobbie, 1997; McAinsh et al., 1997). Liu and
Luan (1998) identi®ed two kinds of stretch-activated
potassium channels in Vicia faba guard cells: voltagegated and insensitive to membrane potential. This was the
®rst evidence that plants contain osmosensitive, voltagedependent channels, those previously described by Ramahaleo et al. (1996) being voltage-independent. Negative
pressure activates voltage-insensitive currents with conductance very di€erent from that of voltage-dependent K ‡ channels. Voltage-dependent currents (IKin and IKout) are in
turn sensitive to osmotic gradient rather than changes in
pressure, although actin ®laments are involved in IKin
regulation (Liu and Luan, 1998). Hypotonic conditions
activate IKin and inactivate IKout , while hypertonic conditions act in the opposite way. An alternation in channel
opening frequency is responsible for regulating IKin and
IKout under di€erent osmotic conditions. Hypertonic
461
inhibition of IKin can be prevented by disruption of actin
®laments. Actin ®lament disruption occurs in hypotonic
conditions providing a link between hypotonic stress and
hypotonic activation of the inward K ‡ channels. Also
cytochalasin D (a cytoskeleton disrupting drug) modulates
IKin in a similar way to hypotonic conditions (Liu and
Luan, 1998), which is consistent with the report of Hwang
et al. (1997). It seems reasonable that stretch-activated
channels in the plant plasma membrane, which is under
continuous compression resulting from turgor pressure and
the presence of the cell wall, interact with cytoskeletal
structures providing local stretch of the membrane. It is
postulated that during perception of gravitational stimuli,
statoliths exert local stretch on the membrane via cytoskeletal ®bres (Sievers et al., 1996).
Light-activated potassium channels
Blom-Zandstra et al. (1997) examined light e€ects on
‡
channels in mesophyll protoplasts of
voltage-gated Kout
Nicotiana tabacum (Table 2). Single channel data from
patch-clamp studies indicate that the activity of the channel
increases upon dark-light transition. The e€ect of light was
not observed in root cells or chlorophyll-de®cient cells,
suggesting that such a response requires photosynthetic
activity. These results are consistent with those of Kim et al.
(1992) who showed that K ‡ channels display responses to
light. The light activated ion channels and electrogenic
proton pump are regarded as important factors in the not
yet fully understood light stimulus transduction cascade
(discussed by Szarek and Trebacz, 1999).
CA L C I U M C H A N N E L S
Calcium ions are universal second messengers in plant and
animal cells. They mediate in various signalling pathways
(reviewed by Brownlee et al., 1999; Sanders et al., 1999)
from signal perception to gene expression, through the
activation of ion channels and enzyme cascades. Stimulusinduced increases in [Ca2‡ ]cyt encode information as speci®c
spatial and temporal changes in frequency of [Ca2‡ ]cyt
oscillationsÐthe `calcium signature' (McAinsh et al.,
1997; Leckie et al., 1998). After signal transition, excess
Ca2‡ must be sequestered into external and internal stores
to keep [Ca2‡ ]cyt at a low level ranging from tens to
hundreds nM. Thus, all Ca2‡ channels located in Ca2‡ sequestering membranes are strongly inward rectifying
( facilitating Ca2‡ in¯ux to the cytosol).
Ligand-gated calcium channels in plasma membrane
Recently, Zimmermann et al. (1997) reported a novel
Ca2‡ -permeable, La3‡ -sensitive plasma membrane ion
channel of large conductance (Table 2). The channel is
activated by elicitors and is essential in pathogen defence.
Receptor-mediated stimulation of these channels appears to
be involved in the signalling cascade triggering a pathogen
defence system. The activation of plasma membrane Ca2‡ channels by speci®c and non-speci®c elicitors provides a
direct demonstration of a pathway by which [Ca2‡ ]cyt
462
Krol and TrebaczÐIon Channel Gating in Plant Cells
increases to levels that can initiate the production of active
oxygen species, callose and phytoalexins via Ca2‡ ±
dependent gene expression (Blumwald et al., 1998).
Ligand-gated calcium channels in inner membranes
2‡
Ligand-gated Ca channels in plant cells reported to
date represent two classes: IP3 (inositol triphosphate)- or
cADPR (cyclic ADP-ribose)-gated (Table 3). Recently a
new signalling moleculeÐNAADP (nicotinic acid adenine
dinucleotide phosphate)Ðhas been found in animal cells
(Lee, 2000). Ligand-gated Ca2‡ channels are present only in
intracellular compartments, and thus their existence provides a convenient mechanism for linking perception of
stimuli (e.g. light, IAA, ABA, osmotic shock, pollination,
Nod-factors, cold shock) to intracellular calcium mobilization (Knight et al., 1996; McAinsh et al., 1997; Muir et al.,
1997; Trewavas and Malho, 1997). The IP3-induced Ca2‡ release originates mainly from vacuolar stores, although in
cauli¯ower, Muir and Sanders (1997) found at least two
distinct membrane populations sensitive to IP3. IP3-induced
Ca2‡ -currents are inwardly rectifying and highly selective
for calcium (Allen and Sanders, 1997). A speci®c IP3binding 400-kDa protein, which is competent to release
Ca2‡ when incorporated into proteoliposomes (Biswas
et al., 1995), was puri®ed from mung bean, though no
subsequent reports on this protein have appeared. There is
some indirect evidence for the presence of IP3-gated Ca2‡
channels in the tonoplast of the algae Chara and Nitella
(Katsuhara and Tazawa, 1992).
As well as IP3-gated channels, cADPR-gated Ca2‡
channels act as instantaneous strong inward recti®ers over
physiological membrane potentials and they are activated
by ligand binding only in the presence of calcium on the
luminal side of the membrane. Pharmacological studies
suggest that cADPR has the capacity to act as a Ca2‡ mobilizing intracellular messenger and an endogenous
modulator of Ca2‡ -induced Ca2‡ release (CICR) (Willmott
et al., 1996). Ryanodine and ca€eine (agonists of ryanodine
receptors in animal cells) are able to cause activation of
cADPR-gated channels in a dose-dependent manner (Allen
et al., 1995), while ruthenium red and procaine (antagonists
of ryanodine receptors in animal cells) block Ca2‡ release
(Allen et al., 1995; Muir and Sanders, 1996; Bauer et al.,
1998) in plant cells. Heparin of low molecular mass and
TMB-8, well known competitive inhibitors of IP3-receptors
in plant and animal cells, are without e€ect on cADPRgated Ca2‡ -channels (Muir and Sanders, 1996). Allen et al.
(1995) demonstrated that there is a relatively low density of
cADPR-gated channels in beet microsomes. cADPR-gated
channels could participate in calcium release only up to
25 % in comparison to the dominating IP3-induced Ca2‡ release. Similar results were obtained from cauli¯ower
microsomes (Muir et al., 1997) and the unicellular green
alga Eremosphaera viridis (Bauer et al., 1998). Preliminary
experiments on sea urchin egg homogenates indicate that
cADPR may bind to an accessory 100±140 kDa protein
(Galione and Summerhill, 1996).
The lack of modulation of plant ligand-gated Ca2‡ channels by cytosolic Ca2‡ is the most notable di€erence
recognized to date between these and animal channels
(Muir et al., 1997).
Voltage-gated calcium channels in the plasmalemma
Many voltage-gated Ca2‡ channels have been described
in a variety of plant tissues and species (reviewed by Pineros
and Tester, 1997) (Table 2). Most of these are activated
through membrane depolarization and stimuli causing
membrane depolarization such as increased [K ‡ ]ext
(Thuleau et al., 1994), Ca2‡ starvation (Reid et al., 1997),
cytokinins (Schumaker and Gizinski, 1993), light or
electrical pulses (Trebacz et al., 1994; Ermolayeva et al.,
1996, 1997) mechanical stimulation (Shimmen, 1997), ABA
(McAinsh et al., 1995; Grabov and Blatt, 1998b) and
microtubule inhibitors (Thion et al., 1996). White (1998),
focusing on Ca2‡ channels in the plasma membrane of root
cells, distinguished between them based on their di€erent
sensitivities to La3‡ , Gd3‡ and verapamil. He discussed
their roles in mineral nutrition, intracellular signalling and
polarized growth.
Kiegle et al. (2000), Gelli and Blumwald (1997) and
Stoeckel and Takeda (1995) described the hyperpolarization-activated in¯uxes of Ca2‡ through the plasmalemma.
The hyperpolarization-activated calcium current is postulated to allow nutritive Ca2‡ uptake. Hyperpolarizationactivated Ca2‡ channels described in the plasma membrane
of Vicia faba guard cells by Fairley-Grenot and Assmann
(1992) are in fact the inwardly rectifying K ‡ channels
mediating Ca2‡ in¯ux prior to their closure and they may
be involved in the regulatory mechanism of stomatal
aperture changes.
Voltage-gated calcium channels in inner membranes
Voltage-gated Ca2‡ -channels are also present in other
cell compartments such as the vacuole, thylakoids or ER
(Pineros and Tester, 1997) (Table 3). Vacuolar voltagegated Ca2‡ channels (VVCa), characterized by Allen and
Sanders (1994), behave as multi-ion pores inwardly rectifying over the voltage range between ÿ20 and ÿ50 mV
(hyperpolarization). Their activity is inhibited by lanthanides, verapamil, nifedipine and by [Ca2‡ ]cyt above 1 mM.
Luminal Ca2‡ shifts the threshold for VVCa activation to a
less negative potential, and therefore restricts the accumulation of calcium excess in the vacuole. Luminal pH of
about 5.5 prevents uncontrolled leakage of Ca2‡ , because at
this physiological pH value the channel openings are very
infrequent (the highest activation is around pH 7).
Johannes and Sanders (1995) showed that a binding of
two calcium ions is required to open the VVCa channel.
Voltage-gated vacuolar Ca2‡ channels, previously
described in tonoplasts of beet, Arabidopsis and tobacco,
are in fact manifestations of SV K ‡ channels (Ward and
Schroeder, 1994).
KluÈsener et al. (1995, 1997) have shown the voltage-gated
Ca2‡ channels derived from endoplasmic reticulum membranes of Bryonia dioica touch-sensitive tendrils. The range
of membrane potentials activating these channels was
a€ected by the Ca2‡ gradient across the membrane. Single
Krol and TrebaczÐIon Channel Gating in Plant Cells
channel currents were modulated by divalent cations,
protons and H2O2 . H2O2 is a strong inhibitor of these
channels. The channel conductance increases with cytosol
acidi®cation. These channels play an important role in the
modulation of [Ca2‡ ]cyt in response to changes in [H2O2]cyt
or pHcyt .
Stretch-activated calcium channels
Taylor et al. (1996) examined both stretch-activated and
voltage-gated mechanosensitive Ca2‡ -permeable cation
channels in subprotoplasts prepared from di€erent regions
of rhizoid and thallus cells of Fucus zygotes (Table 2). Their
results suggest that intercellular signal transduction is
patterned by interactions of the cell wall, plasma membrane
and intracellular Ca2‡ stores.
Thion et al. (1996) observed activation of voltage-gated
Ca2‡ channels by microtubule disruption. Their results are
consistent with a previous report of Davies (1993), who
postulated that variation potentials can be transduced via
mechano-sensitive Ca2‡ channels into gene expression
through Ca2‡ -dependent cytoskeleton-associated phosphorylation/dephosphorylation processes. In addition,
Ca2‡ in¯ux through `volume sensing' voltage-gated Ca2‡
channels is essential for an apical Ca2‡ gradient to be
maintained in a growing cell (Taylor et al., 1996; HoldawayClarke et al., 1997).
ANION CHANNELS
Plant anion channels regulate anion e‚ux from a cell
through plasmalemma (Table 2) and/or tonoplast
(Table 3). Anion e‚ux from the cytoplasm into the
extracellular space is driven by the anion gradient and the
negative membrane potential causing plasma membrane
depolarization, which in turn activates outward rectifying
voltage-gated K ‡ channels. Anion-induced depolarization
plays a crucial role in such processes as xylem loading,
generation and propagation of action potentials or lightinduced transient voltage changes of membrane potential.
In addition, anion and potassium losses promote osmoregulation, stomatal closure, tissue and organ movements.
Since plant cells experience low extracellular anion concentrations, anion uptake must be energetically coupled with
proton pumps.
Ligand-gated anion channels in the plasmalemma
There are many anion channels activated by cytoplasmic
calcium widespread in plant cells (Katsuhara and Tazawa,
1992). Ca2‡ -dependent anion channels are responsible for
the main depolarizing current during action potential in
Charophyta (Okihara et al., 1991; Katsuhara and Tazawa,
1992; Thiel et al., 1993; Shimmen, 1997), the liverwort
Conocephalum conicum (Trebacz et al., 1994), Aldrovanda
vesiculosa (Iijima and Sibaoka, 1985) and during phytochrome-mediated transient depolarization in the moss
Physcomitrella patens (Ermolayeva et al., 1996, 1997).
Johannes et al. (1998) showed a direct e€ect of cytoplasmic protons on Cl ÿ e‚ux in Chara corallina during
463
intracellular acidosis. H ‡ -activated anion channels responsible for Cl ÿ currents act to facilitate an enhanced proton
e‚ux under conditions of low pHcyt . Activity of these
channels is also indirectly pH- and Ca2‡ -dependent
through phosphorylation/dephosphorylation processes.
The above-mentioned ®ndings imply that plasma
membrane anion channels play a central role in pHcyt
regulation in plants, in addition to their established roles
in turgor/volume regulation and signal transduction.
Ligand-gated anion channels in the tonoplast
Katsuhara and Tazawa (1992) summarized calciumregulated channels and their bearing on physiological
activities in characean cells. They presented some evidence
for the presence of Ca2‡ -regulated anion channels in the
tonoplast of Chara, Nitellopsis and Lamprothamnium giant
internodal cells (Table 3). Activation of these channels by
[Ca2‡ ]cyt is assumed to occur during turgor regulation.
Voltage gated anion channels in the plasma membrane
In the plasma membrane, voltage-gated anion channels
are activated by depolarization and under an excess of
cytoplasmic Ca2‡ . They deactivate under hyperpolarizing
potentials (Keller et al., 1989; Hedrich et al., 1990; Hedrich
and Jeromin, 1992; Linder and Raschke, 1992; Schroeder
and Keller, 1992; Dietrich and Hedrich, 1994; Thomine
et al., 1995; Schultz-Lessdorf et al., 1996; Lewis et al., 1997;
Pei et al., 1998). Inverse voltage dependence (activation by
hyperpolarization) has been reported infrequently to date.
Barbara et al. (1994) reported hyperpolarization-activated
chloride currents contributing both to the control of
membrane potential and to osmotic balance regulation in
carrot cells. Neither calcium ions nor MgATP were
necessary for fast activation of these channels. Under
large hyperpolarization, Barbara et al. (1994) observed
rapid and voltage-dependent channel inactivation.
Recently, hyperpolarization-activated anion channels have
also been found in the plasmalemma of the unicellular
green alga Eremosphaera viridis (SchoÈnknecht et al., 1998).
They conduct an anion e‚ux and hence they are responsible for limiting the amplitude of dark-induced transient
hyperpolarization caused by K ‡ -release. The well-known
anion channel inhibitors such as A-9-C, NPPB and Zn2‡
block these channels. Elzenga and Van Volkenburgh
(1997b) reported that in pea mesophyll cells there are
Ca2‡ -dependent anion currents activated by hyperpolarizing pulses. These anion channels display ATP-dependent
bi-modular ( fast and slow) kinetics. R-mode ( fast activation and deactivation of the channel) occurs in the
absence of ATP. However when 3 mM MgATP is added to
the pipette solution facing the cytoplasmic side of the
membrane, the current shows slow but clear timeinactivation (S-mode).
Dietrich and Hedrich (1994) showed the bimodular
kinetics of the guard cell anion channel (GCAC1) in Vicia
faba protoplasts. Previously these two modes of one guard
cell anion channel were considered as two anion channels
contributing to di€erent depolarization-associated processes
464
Krol and TrebaczÐIon Channel Gating in Plant Cells
during regulation of stomatal movements (Schroeder and
Keller, 1992). Dietrich and Hedrich (1994) noted that the
mode of action of GCAC1 is under the control of
cytoplasmic factors. Later Thomine et al. (1995) also
identi®ed a voltage-dependent anion channel in epidermal
cells of Arabidopsis hypocotyls which showed two-mode
function: rapid and slow mode in the presence or absence of
intracellular ATP, respectively. R-type and S-type channels
are voltage-regulated in a quite di€erent way and they
display di€erent kinetics. Only R-type anion channels
display strong voltage-dependence, while weak voltagedependence of S-type channels leaves them partially active
even when the membrane is strongly hyperpolarized. Such
behaviour of S-type channels makes them responsible for
sustained e‚ux of anions (Keller et al., 1989; Linder and
Raschke, 1992; Schroeder and Keller, 1992; Schroeder et al.,
1993; Thomine et al., 1995), which serves as a negative
regulator during stomatal opening (Schroeder et al., 1993;
Pei et al., 1998) or hypocotyl movements (Cho and
Spalding, 1996). Transition between R- and S-mode of an
anion channel may correspond to ATP binding (SchulzLessdorf et al., 1996; Thomine et al., 1997) or alternatively
to ATP-dependent phosphorylation/dephosphorylation
processes (Schmidt et al., 1995; Thomine et al., 1995).
R-type guard cell anion channels (GCAC1) in Arabidopsis
were shown not to be directly regulated by phosphorylation
events (Schulz-Lessdorf et al., 1996). They require cytoplasmic ATP to undergo voltage- and Ca2‡ -dependent
activation, involving strongly cooperative binding of four
ATP molecules (Schulz-Lessdorf et al., 1996). On the other
hand, S-type GCAC1 channels are strongly activated by
phosphorylation (in Vicia faba and Commelina communis
guard cells) or dephosphorylation (in Arabidopsis and
Nicotiana cells) (Armstrong et al., 1995; Schmidt et al.,
1995; Cho and Spalding, 1996; Li and Assmann, 1996;
Schulz-Lessdorf et al., 1996; Esser et al., 1997; Mori and
Muto, 1997; Pei et al., 1997, 1998; Schwarz and Schroeder,
1998). Therefore, guard cell anion channels characterized in
Arabidopsis (GCAC1) can also be classi®ed as ligand-gated
channels, since Schulz-Lessdorf et al. (1996) showed direct
binding of ATP to the channel protein. Leonhardt et al.
(1999) in turn, suggest that the slow anion channel in guard
cells may belong to the class of ATP binding cassette (ABC)
proteins. The same situation applies in the case of voltagegated and nucleotide-regulated anion channels of Arabidopsis hypocotyls described by Thomine et al. (1997). They
con®rmed that nucleotide binding (ATP 4 ADP AMP)
regulates channel activity (alters the kinetics and voltagedependence, causing a shift toward depolarized potentials
and thus leading to a strong reduction of anion current
amplitude). This regulation may couple the electrical
properties of the membrane with the metabolic status of
the whole cell.
Rapid- and slow-modes of the Arabidopsis guard cell
anion channel (GCAC1) are also variously in¯uenced by
pHcyt (Schulz-Lessdorf et al., 1996). The kinetics of S-mode
is in¯uenced by the pH gradient across the plasmalemma
(the inactivation gate responds to pH gradient, which may
be converted into a change of a channel structure). Such
pH gradient-dependence of slow inactivation resembles a
carrier-mode action. In the case of R-mode, the proton
gradient does not seem to a€ect channel activation
following ATP-binding. The single channel activity of
R-type GCAC1 increases as a function of [H ‡ ]cyt ( protonation of the cytoplasmic site of the channel), while single
channel conductance is una€ected either by pHcyt or pHext .
Similar pH sensitivity was determined for anion-permeable
vacuolar channels (Schulz-Lessdorf and Hedrich, 1995).
Since the time- and voltage-dependent activity of guard cell
anion channels (GCAC1) was shown to be strongly
modulated by ATP and H ‡ (Schulz-Lessdorf et al., 1996),
these channels have been thought to be capable of sensing
changes in the energy status, the proton pump activity and
acid metabolism of the cell.
Patch-clamp studies revealed that growth hormones can
directly a€ect voltage-dependent activity of inwardly
rectifying anion channels in a dose-dependent manner
(Hedrich and Jeromin, 1992). Auxin binding is side- and
channel-speci®c, and results in a shift of the activation
potentials towards the resting potential favouring transient
channel opening (Marten et al., 1991). These authors
demonstrated that auxin can interact directly with the
extracellular face of the channel, eliciting stomatal opening.
Voltage-gated anion channels in the tonoplast
In the tonoplast (Table 3) there are two types of cytosolnegative-potential-activated (hyperpolarization-activated)
anion channels: VCl and VMal (Allen and Sanders,
1997). The ®rst is responsible for carrying Cl ÿ to the
vacuole (inward rectifying), while the second is mainly
permeable for malate, but also for succinate, fumarate,
acetate, oxaloacetate, NO3ÿ and H2 PO4ÿ : VMal is very
strongly inward rectifying over the physiological range of
negative potentials, but more negative than Emal (Cerana
et al., 1995). Cytosolic Ca2‡ and ATP do not a€ect VMal
channels (Cerana et al., 1995; Chengs et al., 1997). On the
other hand, Pei et al. (1996) reported that calmodulin-like
domain protein kinase (CDPK) activates vacuolar malate
and chloride conductances (VCl) in guard cell vacuoles of
Vicia faba. Activation of both currents depends on Ca2‡
and ATP, enabling anion uptake into the vacuole even at
physiological potentials. CDPK-activated VCl currents
were also observed in red beet vacuoles, suggesting that
these channels may provide a more general mechanism for
kinase dependent anion uptake (Pei et al., 1996).
Voltage-dependent anion channels in other endomembranes
Voltage-dependent anion channels (VDACs or mitochondrial porins) in the outer membrane of mitochondria
regulate the mitochondrial membrane potential, among
other things, during transduction of an apoptotic signal into
the cell (Green and Reed, 1998; Shimizu et al., 1999) or
metabolite di€usion (Elkeles et al., 1997; Rostovtseva and
Colombini, 1997; Mannella, 1998) (Table 3). According to
Rostovtseva and Colombini (1997) these channels are
ideally suited to controlling the ¯ow of ATP between the
cytosol and the mitochondrial spaces. VDAC pore is formed
by a single 30-kDa protein (Song et al., 1998) which
Krol and TrebaczÐIon Channel Gating in Plant Cells
undergoes a major conformational change at pH 5 5
(Mannella, 1997, 1998). However, functional VDAC is a
heterodimer including one pore protein and other modulating subunits (Elkeles et al., 1997). Apart from transmembrane voltage and pH, VDACs can be regulated by direct
binding of signalling proteins (Shimizu et al., 1999).
Anion uniport in plant mitochondria is mediated by a
pH-regulated inner membrane anion channel that is activated by matrix H ‡ (Beavis and Vercesi, 1992). Voltagedependent inner mitochondrial anion channels (IMACs),
which serve as a safeguard mechanism for recharging the
mitochondrial membrane potential, have been found in
animal tissues (Ballarin and Sorgato, 1996; Borecky et al.,
1997).
Voltage-dependent anion channels were characterized by
a patch-clamp study in osmotically swollen thylakoids from
Peperomia metallica (SchoÈnknecht et al., 1988) and the alga
Nitellopsis obtusa (Pottosin and SchoÈnknecht, 1995).
Voltage-gated anion channels found in thylakoids are most
probably responsible for the compensation of light-driven
H ‡ movements (SchoÈnknecht et al., 1988; Heiber et al.,
1995).
Recently, Pohlmeyer et al. (1998) discovered a new type
of voltage-dependent solute channel of high conductance in
the outer envelope of chloroplasts, etioplasts and non-green
root plastids (Table 3). The channels are permeable for
triosephosphate, ATP, Pi , dicarboxylic acids, amino acids,
and sugars. Their open probability is highest at 0 mV (which
is consistent with the absence of transmembrane potential
across the plastidic outer membranes). Previously, Heiber
et al. (1995) reported a voltage-dependent anion channel of
low conductance in the chloroplast envelope. There are also
anion channels found in the inner envelope membrane of
isolated intact chloroplasts (Fuks and Homble, 1999).
Stretch-activated anion channels
Falke et al. (1988) ®rst reported large conductance,
stretch-activated, anion-selective channels in protoplasts of
tobacco. Cosgrove and Hedrich (1991) then showed the
existence of stretch-activated Cl ÿ , Ca2‡ and K ‡ channels in
the plasma membrane of guard cells. Teodoro et al. (1998)
suggested that the changes in turgor pressure induced by
hyper-/hypo-osmotic stress may cause an early inactivation/
activation of stretch-sensitive anion channels, respectively.
Light-activated anion channels
By patch clamping hypocotyl cells isolated from darkgrown Arabidopsis thaliana seedlings, Cho and Spalding
(1996) revealed the existence of blue-light activated anion
channels responsible for light induced membrane depolarization (Table 2). Their results are consistent with previous
reports of Elzenga et al. (1995). Further studies on bluelight activated anion channels in Arabidopsis hypocotyl
conducted by Lewis et al. (1997) showed that the open
probability of the channel depends on [Ca2‡ ]cyt and that
within the calcium concentration range of 1±10 mM the
probability of channel activation increases. Their results
indicate that cytoplasmic calcium does not a€ect the anion
465
channel directly, but that it does so through intermediates
(e.g. Ca2‡ -dependent kinases or phosphatases). Activation
of blue light-induced anion channels plays a central role in
transducing light signals into hypocotyl growth inhibition
(Cho and Spalding, 1996; Parks et al., 1998).
Light-activated anion channels, resembling those above,
were also reported by Elzenga and Van Volkenburgh
(1997a) (Table 2). They examined light-induced transient
depolarization in Pisum sativum mesophyll cells due to
increased conductance for anions and concluded that:
(1) under illumination the anion current increases threefold because of an increase in the open probability of a 32pS anion channel; (2) this change in channel activity is not
due to light-induced changes in membrane potential; (3) the
anion current depends on light intensity and can be totally
blocked by the photosynthetic inhibitor DCMU; (4) the
anion current is strongly Ca2‡ -dependent; and (5) lightinduced anion e‚ux may balance light-induced proton
extrusion and therefore participate in a mechanism controlling cellular pH, transmembrane and osmotic potential.
CO N C L U S I O N S
From year to year the number of characterized ion channels
increases, which bene®ts our understanding of their roles in
numerous physiological processes. Modern electrophysiological and molecular biological techniques have enabled
the characterization and classi®cation of novel channel
types. On the other hand, some channels previously
described as di€erent types are in fact `synonyms'. These
mainly originate from multiple gating mechanisms that can
sense the energy status of the cell and thus make the cell
responsive to various stimuli in a very ecient way. The ®ne
tuning of channel activities depends on e€ectors available
in a certain cell type, i.e. it is plant and tissue speci®c
(Barbier-Brygoo et al., 1999). Further research concerning
regulation and gating of the ion channels described here will
help to unravel the intermediate signalling mechanisms used
by plants in dynamic responses to the environment during
growth and development.
AC K N OW L E D G E M E N T S
We thank Professor M. A. Venis and the reviewers for
helpful comments and critical reading of the manuscript.
The investigation was supported by the grant 6P04 C 04218
from the State Committee for Scienti®c Research.
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