Download Arabidopsis root K - Journal of Cell Science

1468
Research Article
Arabidopsis root K+-efflux conductance activated by
hydroxyl radicals: single-channel properties, genetic
basis and involvement in stress-induced cell death
Vadim Demidchik1,*, Tracey A. Cuin2, Dimitri Svistunenko1, Susan J. Smith3, Anthony J. Miller3,
Sergey Shabala2, Anatoliy Sokolik4 and Vladimir Yurin4
1
Department of Biological Sciences, University of Essex, Colchester, Essex CO4 5AP, UK
School of Agricultural Science, University of Tasmania, Private Bag 54, Hobart, Tasmania 7001, Australia
Crop Performance and Improvement Division, Rothamsted Research, Harpenden, Hertfordshire AL5 2JQ, UK
4
Department of Physiology and Biochemistry of Plants, Biological Faculty, Belarusian State University, Minsk 220030, Belarus
2
3
*Author for correspondence ([email protected])
Journal of Cell Science
Accepted 11 February 2010
Journal of Cell Science 123, 1468-1479
© 2010. Published by The Company of Biologists Ltd
doi:10.1242/jcs.064352
Summary
Reactive oxygen species (ROS) are central to plant stress response, signalling, development and a multitude of other processes. In this
study, the plasma-membrane hydroxyl radical (HR)-activated K+ channel responsible for K+ efflux from root cells during stress
accompanied by ROS generation is characterised. The channel showed 16-pS unitary conductance and was sensitive to Ca2+,
tetraethylammonium, Ba2+, Cs+ and free-radical scavengers. The channel was not found in the gork1-1 mutant, which lacks a major
plasma-membrane outwardly rectifying K+ channel. In intact Arabidopsis roots, both HRs and stress induced a dramatic K+ efflux that
was much smaller in gork1-1 plants. Tests with electron paramagnetic resonance spectroscopy showed that NaCl can stimulate HR
generation in roots and this might lead to K+-channel activation. In animals, activation of K+-efflux channels by HRs can trigger
programmed cell death (PCD). PCD symptoms in Arabidopsis roots developed much more slowly in gork1-1 and wild-type plants
treated with K+-channel blockers or HR scavengers. Therefore, similar to animal counterparts, plant HR-activated K+ channels are also
involved in PCD. Overall, this study provides new insight into the regulation of plant cation transport by ROS and demonstrates
possible physiological properties of plant HR-activated K+ channels.
Key words: Ion channels, Plant, Potassium, Programmed cell death, Reactive oxygen species, Stress
Introduction
Reactive oxygen species (ROS) are important toxic and regulatory
agents in plants (Apel and Hirt, 2004; Møller et al., 2007). They
are produced in response to a number of stimuli, including biotic
and abiotic stresses, hormones, developmental and gravitropic
signals, and mineral deficiency (Apel and Hirt, 2004; Kwak et al.,
2006). Respiratory and photosynthetic electron-transport chains, as
well as NADPH oxidases and peroxidases, are involved in their
generation in plants (Apel and Hirt, 2004). Several ‘ROS-sensing’
systems have been proposed, including two-component histidine
kinases, mitogen-activated protein kinases, and certain protein
phosphatases and transcription factors (Apel and Hirt, 2004; Kwak
et al., 2006). In addition, plasma-membrane ion channels are
probably also involved in ROS sensing and ROS-mediated
regulatory reactions in plants. In animals, ion-channel-‘based’ ROS
sensors play crucial roles in key physiological functions (Lahiri et
al., 2006), but their physiological roles in plants are much less well
understood.
There is now ample evidence that ROS can activate plasmamembrane cation influx channels in plant cells, resulting in the
elevation of cytosolic Ca2+ activity (Pei et al., 2000; Köhler et al.,
2003; Demidchik et al., 2003; Foreman et al., 2003). The biophysical
properties and physiological roles of such ROS-activated Ca2+permeable channels have been investigated in a number of plant
preparations (Pei et al., 2000; Coelho et al., 2002; Demidchik et al.,
2003; Demidchik et al., 2007; Demidchik et al., 2009; Foreman et
al., 2003; Köhler et al., 2003; Takeda et al., 2008). In contrast to
cation influx, the effect of ROS on cation efflux remains poorly
studied. Hydroxyl radical (HR)-activated K+-efflux conductance has
been observed alongside the activation of Ca2+ channels (Demidchik
et al., 2003), but has not been investigated in detail; the singlechannel characteristics, genetic basis and physiological roles of HRactivated K+ conductance remain unknown.
HR-activated K+-efflux channels can potentially play very
important roles in plants. Potassium is an essential nutrient, a
major osmoticum and regulator of enzymes, so decreasing the K+
activity will result in significant changes in metabolic reactions
and physiological processes (Marschner, 1995). Moreover, retaining
high cytosolic K+ concentration is crucial for plant salinity
tolerance, although the mechanisms by which this protects cells
from Na+ is still unclear (Maathuis and Amtmann, 1999; Munns
and Tester, 2008). We have recently demonstrated that K+ channels
are responsible for K+ loss from Arabidopsis roots and leaves
during salt stress, and that this triggers an oxidative burst in
Arabidopsis roots (Demidchik et al., 2003; Shabala et al., 2006).
However, it was not shown whether the ROS produced under salt
stress include HRs (capable of activating K+-efflux channels). As
well as salinity, both K+ efflux and ROS generation are induced by
pathogen attack, cold and osmotic shock, heavy metals, UV light,
paraquat and ozone (Murphy and Taiz, 1997; Apel and Hirt, 2004;
Shabala and Cuin, 2008). This suggests that K+ efflux and ROS
generation are a general plant-cell response to stress.
Arabidopsis ROS-activated channels
1469
The main aim of this study was to investigate in detail the
activation of plant K+-efflux channels by HRs and to test the
possible physiological roles of this phenomenon. The model plant
Arabidopsis thaliana was used, focusing on the root epidermis as
it is the most likely first target of stress. This study reports, for the
first time, the single-channel properties of a plant HR-activated K+
channel, reveals the gene encoding this channel and shows its
possible role in plant programmed cell death (PCD).
Results
Journal of Cell Science
HRs activate single K+ channels in A. thaliana root
protoplasts
The exposure of excised outside-out plasma-membrane patches
exhibiting no channel activity to a HR-generating mixture
containing 1 mM CuCl2 and 1 mM L-ascorbic acid (Cu/a) activated
a single-channel outwardly directed conductance (Fig. 1A), with
the open probability (Popen) increasing with depolarisation (Fig.
1B). The maximal activation was observed 15-20 minutes after the
addition of Cu/a [Fig. 1C; mean ± s.e. Popen0.55±0.09; n6;
holding potential (HP)80 mV; 20 minutes exposure to 1 mM
Cu/a]. This activation was found in 36% of protoplasts (n136).
Removal of Cu/a from the bathing solution decreased the HRactivated currents when washing out started after 20-30 minutes of
treatment (Fig. 1D). Longer-term treatment (30-45 minutes) with
Cu/a caused irreversible activation of the K+ current that, over
time, slowly ‘ran down’ (Fig. 1D).
To demonstrate that the observed activation of an outwardly
directed channel was caused by HRs, the HR scavenger thiourea
(10 mM) was applied together with Cu/a (Fig. 1C,E). This treatment
inhibited the Cu/a-induced activation of K+ conductance, causing
a tenfold decrease in its Popen (mean ± s.e. Popen0.051±0.019; n5;
HP80 mV; 20 minutes; 1 mM Cu/a) (Fig. 1C,E). Dimethyl
sulfoxide (DMSO), another free-radical scavenger, resulted in a
similar effect (0.3% DMSO; mean ± s.e. Popen0.021±0.004; n5;
HP80 mV; 20 minutes; 1 mM Cu/a) (Fig. 1F). In contrast to HRs,
exogenous H2O2 (10 mM) did not activate currents (n14; 20-30
minutes; data not shown).
Exogenous application of the K+-channel blockers Ba2+, Ca2+,
Cs+, tetraethylammonium+ (TEA+) and Na+ significantly decreased
the Popen of the HR-activated conductance channel (Fig. 1F). The
Ca2+ channel antagonists verapamil and nifedipine did not cause
significant changes in HR-activated single-channel conductance
(Fig. 1F; P0.624 for verapamil and P0.552 for nifedipine,
ANOVA test). Overall, the pharmacological analysis showed that
HR-activated currents were mediated by K+ channels, not by any
other group of cation channel.
The extrapolated reversal potential of the single-channel HRactivated conductance was –75.2±10.4 mV (n6), close to the
equilibrium potential of K+ (–107.6 mV) (Fig. 1G). With 80 mM
pipette K+ solution, the average unitary conductance derived from
the first open state was 16.1±1.9 pS (mean ± s.e.; n6; Fig. 1G).
Substitution of K+ in the pipette solution with NH4+ (Fig. 2A),
which has similar physical properties to K+ (Hille, 2001), did not
cause any significant change in the open probability (mean ± s.e.
Popen0.49±0.08; n4; HP80 mV; 20 minutes; 1 mM Cu/a; Popen
derived from the first open state; P0.643, ANOVA test, comparison
with K+) and only slightly decreased the unitary conductance
(mean ± s.e. 13.3±1.2 pS; n4; same conditions; P0.303, ANOVA
test, comparison with K+). Substitution of K+ for the K+-channel
blocker Cs+ (Fig. 2A) dramatically decreased the Popen (mean ± s.e.
Popen0.051±0.013; n5; same conditions; P0.001, ANOVA test,
Fig. 1. The activation of single K+ channels by HRs in excised outside-out
patches from Arabidopsis root epidermal protoplasts. (A)Depolarisationactivated currents before (control) and after (+Cu/a) addition of HR-generating
mixture (20 minutes exposure to Cu/a: 1 mM CuCl2 and 1 mM L-ascorbic
acid). The closed state (C) and open states (O1-O3) of the channel are shown
by dotted lines. (B)Voltage dependence of the K+-channel open probability
(Popen) (mean ± s.e.; n5-6). (C)The increase in Popen with duration of
exposure to Cu/a (open circles; HP80 mV) and the effect of thiourea (10
mM) on this process (closed circles; HP120 mV) (mean ± s.e.; n4-6).
(D)Change in Popen during wash out of Cu/a (indicated by arrow; HP80 mV;
mean ± s.e.; n4) after 20 minute (open circles) or 45 minute (closed circles)
treatment. (E)Typical currents measured at 120 mV in the presence of 10 mM
thiourea or 0.3% DMSO. (F)The effect of externally applied cation-channel
blockers (in mM: 10 TEA+, 10 Cs+, 10 Na+, 2 Ca2+, 2 Ba2+, 0.05 verapamil
and 0.05 nifedipine) on Popen (20 minutes Cu/a; HP80 mV; mean ± s.e.; n56). (G)Graphical determination of the unitary HR-activated conductance from
the current-voltage (I-V) relationship curve (mean ± s.e.; n5-6). In B,C,D,F,
the first open state was used to calculate Popen. In all cases (A-G), the bathing
solution contained (in mM): 1 K+, 1.6 Cl– and 0.3 Ca2+ (pH 5.8); pipette
solution was as follows (mM): 80 K+, 70 gluconate– and 10 Cl– (100 nM Ca2+,
pH 7.2).
comparison with K+) and single-channel conductance (mean ± s.e.
8.1±1.3 pS; n5; same conditions; P0.009, ANOVA test,
comparison with K+). These results show that, typical of K+
Journal of Cell Science
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Journal of Cell Science 123 (9)
Fig. 2. Properties of single HR-activated K+ channels in excised outsideout patches from Arabidopsis root epidermal protoplasts. (A)Typical
currents induced by 1 mM Cu/a when pipette solution contained 80 mM NH4+
or Cs+ instead of K+. The closed state (C) and open states (O1-O2) of the
channel are shown by dotted lines. (B)Typical currents induced by 5M or
0.3 mM Cu2+ applied with 0.1 mM L-ascorbic acid. (C)Dependence of Popen
on different Cu2+ concentrations applied with 0.1 mM ascorbic acid (mean ±
s.e.; n4-6). (D)Typical current in gork1-1 after 20 minutes exposure to 1 mM
Cu/a. In all cases (A-D), the bathing solution contained (in mM): 1 K+, 1.6 Cl–
and 0.3 Ca2+ (pH 5.8); pipette solution (apart from in A) was (in mM): 80 K+,
70 gluconate– and 10 Cl– (100 nM Ca2+, pH 7.2); exposure time to Cu/a was
20 minutes; HP80 mV.
channels, HR-activated channels are highly permeable to K+ and
NH4+, and poorly permeable to Cs+.
Fig. 3. The effect of HRs on whole-cell K+ currents and membrane
potential in Arabidopsis root epidermis. (A)Currents activated by 20
minutes exposure to 1 mM Cu/a in WT and gork1-1 plants. Currents were
smaller when 80 mM TEA+, instead of 80 mM K+, was added to the pipette
solution. The corresponding average current-voltage (I-V) curves (symbols are
the same for the time course and I-V curves; n5; mean ± s.e.) are shown in
the bottom panels. (B,C)Depolarisation caused by different concentrations of
(B) NaCl and (C) Cu2+ applied with 0.1 mM ascorbate (mean ± s.e.; n5).
(D)Currents activated by 1 mM Cu/a in WT cells in the presence of bathing
solution containing 0.1 mM K+ (instead 1 mM K+) (mean ± s.e.; n4). In all
cases, the bathing solution contained (in mM): 1 K+ (apart from D, where it
was 0.1 K+), 1.6 Cl– and 0.3 Ca2+ (pH 5.8); pipette solution was (in mM): 80
K+ (apart from 80 TEA panel), 70 gluconate– and 10 Cl– (100 nM Ca2+, pH
7.2); exposure time to Cu/a (or NaCl in B) was 20 minutes. Liquid junction
potentials for the 80 K and 80 TEA solutions were 29 mV and 2.7 mV,
respectively.
Physiological concentrations of Cu2+ and ascorbate
activate K+-efflux channels
Ascorbate reaches submillimolar levels in plant cell walls
(Pignocchi and Foyer, 2003), and plant cell catalytic activities of
transition metals (Cu2+ and Fe3+) are in the range 0.3 to 15 M
(Becana and Klucas, 1992). We tested several Cu2+ concentrations,
added together with 0.1 mM ascorbate (Fig. 2B), and found that
the minimal Cu2+ concentration that causes K+ current activation
(mean ± s.e. Popen0.043±0.009; HP80 mV; n6) was 5 M.
Maximal K+ efflux was achieved after the addition of 0.3 mM
Cu2+. These data show that physiological levels of Cu/a are capable
of activating K+ channels.
Gork1-1 lacks HR-activated K+-efflux channels
The outwardly directed K+ channels in Arabidopsis root epidermis
are likely to be GORK1 and/or AKT2/3 (Véry and Sentenac,
2002). However, AKT2/3 has been recently shown to conduct a
steeply inwardly rectifying current, when expressed in plants (Latz
et al., 2007). Some cyclic-nucleotide-gated channels could also be
involved, but this has not yet been proven experimentally
(Demidchik and Maathuis, 2007). Therefore, the main focus of this
study is GORK, which is known to conduct large outwardly
rectifying K+ currents (Ache et al., 2000; Ivashikina et al., 2001;
Hosy et al., 2003). We tested HR-activated K+-efflux currents in
root epidermal protoplasts isolated from gork1-1 plants lacking
functional GORK (Hosy et al., 2003) (Fig. 2D). The 16 pS channel
was not observed in gork1-1 plants, indicating that the HR-activated
K+ channel is encoded by GORK. Outwardly directed single
channels with smaller unitary conductance (mean ± s.e. 5.9±1.6
pS; n3; data not shown) were activated in 3 out of 18 gork1-1
protoplasts in response to 1 mM Cu/a (note that these channels
were not observed in the wild type). Because the probability of
observing these ‘smaller’ channels was very low, a detailed
investigation was not possible. Nonetheless, it might be speculated
that these channels were expressed ‘to fill a gap’ in K+-efflux
function in the absence of GORK and could be encoded by some
other cation channel gene.
Whole-cell HR-activated K+-efflux conductance in wildtype and gork1-1 plants
To confirm that the HR-activated K+-efflux channel is encoded by
GORK at the cellular level, we compared whole-cell currents in
wild-type (WT) and gork1-1 plants (Fig. 3A). HRs activated a
steeply rectifying, slowly activating (time-dependent), outward
current in WT protoplasts (Fig. 3A). After 20 minutes exposure to
HRs, the outward current recorded in response to an 18 second
depolarising pulse from –90 to 120 mV increased from 87.9±20.2
to 467.7±91.3 mA m–2 (mean ± s.e.; n5; P0.004, ANOVA test).
Arabidopsis ROS-activated channels
1471
The current did not increase in response to HRs when the K+channel blocker TEA+ replaced K+ in the pipette (Fig. 3A). No
activation of time-dependent K+ currents was found in response to
HRs in gork1-1 plants (n16; Fig. 3A), although the rapidly
activating (instantaneous) conductance increased by about 30-40%.
This current could be due to a channel with a ‘smaller’ unitary
conductance (see above).
In our previous study (Demidchik et al., 2003), the time course
of the voltage-activated current was measured over 2.4 seconds.
Here, long voltage pulses (18 seconds) were applied, allowing the
recording of a more ‘pronounced’ time-dependent current (Fig.
3A). Pulses longer than 18 seconds were not applied because they
destabilised the gigaseal contact. The HR-activated whole-cell K+
conductance revealed two component currents: instantaneous (time
independent) and slowly activating (time dependent). The timedependent component showed greater activation than the
instantaneous (Fig. 3A).
Journal of Cell Science
HR-activated K+ channels can mediate K+ efflux during
stress-induced depolarisation
An important question is whether HR-activated K+ channels can
conduct current (efflux) upon depolarisation caused by stress. To
explore this, we measured membrane potential changes induced by
salinity and oxidative stress (Cu/a) in mature root epidermis (Fig.
3B,C). Measurements were carried out in a bath solution containing
0.1 mM K+, a typical soil solution K+ concentration (Marschner,
1995; Cuin et al., 2003). Both stresses cause depolarisation (Fig.
3B,C). Maximal salt-induced depolarisation (from –127.5±12.7
mV to 7.5±17.1 mV; mean ± s.e.; n5; P<0.001, ANOVA test) was
found after 20 minutes root exposure to 250 mM NaCl (Fig. 3B).
Similar to the patch-clamp experiments described in Fig. 2B,C, we
recorded changes in the membrane potential resulting from the
addition of different Cu2+ concentrations, together with 0.1 mM
ascorbate (Fig. 3C). The addition of 1 M Cu2+ and 0.1 mM
ascorbate did not modify the membrane potential, but 10 M Cu2+
and 0.1 mM ascorbate caused significant depolarisation (Fig. 3C).
At 1 mM Cu2+ and 0.1 mM ascorbate, strong depolarisation was
measured (from –123.2±15.5 mV to –46.2±12.3 mV; mean ± s.e.;
n5; P0.005, ANOVA test) (Fig. 3C). The mixture that was used
in most patch-clamp experiments, 1 mM CuCl2 and 1 mM
ascorbate, depolarised the plasma membrane from –119.9±14.1
mV to –41.9±14.4 mV (mean ± s.e.; n5; P0.005, ANOVA test).
Whole-cell HR-activated K+-efflux currents were measured with
0.1 mM K+ in the bathing medium (Fig. 3D). Application of 0.1
mM K+ (as compared to 1 mM K+) shifted the reversal potential
to more negative voltages. This resulted in the activation of a
significant K+-efflux current at depolarised voltages (Fig. 3D); for
example, at –40 mV, the outward current was 96.4±33.2 mA m–2
(mean ± s.e.; n4). Overall, these results reveal that HR-activated
K+ channels are capable of mediating significant K+ efflux from
root cells during depolarisation caused by stress.
HR-activated K+ efflux from intact roots in wild-type and
gork1-1 plants
To relate the results obtained in protoplasts with K+ flux in intact
roots, we used non-invasive vibrating K+ electrodes [so-called
microelectrode ion flux estimation (MIFE)] (Shabala et al., 1997)
to record the time course of K+ efflux from mature root cells (Fig.
4). Before HR application (control), the K+ efflux was –47.7±24.7
nmol m–2s–1 (net K+ flux; WT, mean ± s.e.; n6) (Fig. 4A). Upon
application of HR, dramatic K+ efflux occurred, reaching a peak of
Fig. 4. Stress-induced changes in K+ efflux in WT and gork1-1 mature root
epidermis, as measured by K+ vibrating microelectrodes (MIFE).
(A,B)TEA+-sensitive responses of K+ flux to 1 mM Cu/a in (A) WT and (B)
gork1-1 cells (mean ± s.e.; n6). (C,D)Similar responses induced by (C) 100
mM NaCl and (D) 1% cellulysin (mean ± s.e.; n6). In all cases, the bathing
solution contained (in mM): 0.1 K+, 0.3 Cl– and 0.1 Ca2+. Addition of
substances is indicated by an arrow. In A,B, the concentration of TEA+ was 10
mM.
–234.4±29.4 nmol m–2s–1 (net K+ flux; mean ± s.e.; n6; P0.002,
ANOVA test, comparison with control) after 2-4 minutes, before
slowly decreasing over time. This effect of HRs was significantly
inhibited by TEA+ (10 mM) (Fig. 4A; P0.003, ANOVA test,
comparison of peaks) and was smaller in gork1-1 plants (Fig. 4B;
P0.025, ANOVA test, comparison of peaks), in agreement with
the patch-clamp data (Fig. 2D). HR-activated K+ efflux in gork11 plants could be related to K+ efflux from cortex cells that might
have different HR-activated channels. Hypothetically, it might also
be due to the ‘smaller’ K+ channel observed in the single-channel
analyses. Nonetheless, K+ efflux in gork1-1 plants is sensitive to
TEA+ (Fig. 4B), suggesting that efflux is the result of K+-channel
activity.
Stress-activated K+ efflux from intact roots
A crucial step of our research was to establish a link between the
activity of the HR-activated channel GORK and the K+ efflux
induced by stress. The generation of ROS in response to NaCl and
the Trichoderma viride elicitor cellulysin (Piel et al., 1997) has
already been demonstrated for Arabidopsis roots (Demidchik et
al., 2003). Here, we tested the possible involvement of GORK in
NaCl- and cellulysin-induced K+ efflux. The application of 100
mM NaCl to mature WT root cells activated K+ efflux (peak net
K+ flux: –200.9±45.3 nmol m–2s–1, mean ± s.e.; n6) (Fig. 4C). In
gork1-1, the NaCl effect was significantly smaller (peak net K+
flux: –41.9±6.4 nmol m–2s–1, mean ± s.e.; n6; P0.006, ANOVA
test) (Fig. 4C). Applying 1% cellulysin resulted in dramatic K+
efflux in the WT, with a peak net K+ flux of –295.9±13.5 nmol
m–2s–1 (mean ± s.e.; n6), but caused a much smaller effect in
gork1-1 (peak net K+ flux: –80.8±32.9 nmol m–2s–1; mean ± s.e.;
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Journal of Cell Science 123 (9)
n6; P<0.001, ANOVA test) (Fig. 4D). The stress-induced K+
efflux in both WT and gork1-1 plants was significantly inhibited
by 10 mM TEA+ (n6; data not shown; P<0.01, ANOVA test).
HR-activated K+ efflux in the root elongation zone
The growing root tip, which interacts first with a new soil
environment, could potentially have greater sensitivity to signalling
molecules such as ROS than other parts of the root. Our attempts
to investigate single K+ channels in outside-out patches from
elongation zone protoplasts were not successful because the
gigaseal contact was not stable in this preparation when depolarising
pulses were applied (n32). Therefore, we investigated HRactivated K+ efflux in the elongation zone using MIFE (Fig. 5A).
HR-activated K+ efflux from the WT elongation zone was about
eight times greater than that from the mature epidermis. This
shows that the elongation zone has an elevated sensitivity to ROS
and points towards their possible involvement in plant stress
perception and response In gork1-1 plants, HR-induced K+ efflux
was again much smaller than in the WT, demonstrating that GORK
encodes HR-activated K+ channels in the elongation zone as well
(Fig. 5A).
Journal of Cell Science
The effect of H2O2 on K+ efflux from root cells
The most abundant and stable ROS in plants is H2O2, which
accumulates in response to stress (Apel and Hirt, 2004). In root
protoplasts, H2O2 does not activate K+-efflux currents (Demidchik
et al., 2003). However, the application of H2O2 to intact roots
could potentially activate K+ efflux, because cell-wall transition
metals and ascorbate could convert H2O2 to HRs (Fry et al.,
2002; Fry, 2004). We tested this hypothesis by applying 10 mM
H2O2 to intact roots (mature cells) and measuring the K+-flux
responses using MIFE (Fig. 5B). We found a TEA+-sensitive
increase in K+ efflux that was smaller in gork1-1 (Fig. 5C). The
affect of H2O2 developed more slowly and was smaller than that
caused by HRs, NaCl and cellulysin (Fig. 4). The difference in
response between WT and gork1-1 (Fig. 5B,C) was also not as
pronounced as with HR-, NaCl- and cellulysin-induced K+ efflux
(Fig. 4B-D).
Fig. 5. ROS-induced K+ efflux in WT and gork1-1 roots, as measured by
K+ vibrating microelectrodes (MIFE). (A)Responses of K+ flux to 1 mM
Cu/a in the elongation zone (mean ± s.e.; n6). (B,C)Responses of K+ flux in
mature epidermis induced by 10 mM H2O2 (and effect of 10 mM TEA+). In all
cases, the bathing solution contained (in mM): 0.1 K+, 0.3 Cl– and 0.1 Ca2+.
Addition of substances is indicated by an arrow.
Salt-induced generation of HRs in intact roots
Although some stresses, such as pathogen elicitors, drought and
cadmium, have been shown to cause HR generation in plants
(Becana and Klucas, 1992; Moran et al., 1994; Shen et al., 1997;
Van Doorslaeder et al., 1999), the effect of salinity has not been
studied (Chung et al., 2008). Electron paramagnetic resonance
(EPR) spectroscopy is one of the most accurate and sensitive
techniques for the detection of radicals in living systems (Halliwell
and Gutteridge, 1999). We added the classical spin trap 5,5dimethyl-1-pyrroline-N-oxide (DMPO) to intact roots. Spectra with
four characteristic peaks, indicative of the formation of HR-DMPO
adducts (Yamazaki et al., 1990; Halliwell and Gutteridge, 1999;
Ikeda et al., 2002), were detected (Fig. 6). Intact roots demonstrated
a small basal level of HR production (Fig. 6A), consistent with the
literature on ‘constitutive’ HR generation in Arabidopsis roots
(Renew et al., 2005). The effect of two NaCl concentrations (100
and 250 mM) was examined (note that neither NaCl concentration,
when added to DMPO without plants, changed the EPR signal;
data not shown). Although it is not a lethal concentration, 100 mM
NaCl does affect germination and growth of Arabidopsis; 250 mM
NaCl does cause cell death (Demidchik and Tester, 2002; Munns
and Tester, 2008; Shabala and Cuin, 2008). The shape of the EPR
spectrum did not change after treatment with NaCl, whereas the
intensity significantly increased (Fig. 6B). A 2.5- to 3-fold increase
in HR generation was caused by the addition of 100 mM NaCl,
whereas 250 mM NaCl triggered a 4- to 4.5-fold increase compared
to the basal level of HR generation (Fig. 6B,F; P<0.01, ANOVA
test).
The mixture of Cu2+ and ascorbate can produce HRs in the
presence of H2O2 or O2 (Shtamm et al., 1977). Because some O2
is always dissolved in water, we used deoxygenated water; all
solutions were sealed with parafilm (Halliwell and Gutteridge,
1999). Solution containing 1 mM Cu/a prepared in this way caused
only a little HR generation (Fig. 6C; top panel). H2O2 can be
synthesised by living cells, so in the presence of roots, Cu/a is
expected to generate HR. Indeed, when applied to intact roots (Fig.
6C; bottom panel) or mixed with H2O2 (Fig. 6D), Cu/a generated
a large amount of HR. Fig. 6F demonstrates how the average
(mean ± s.e.) EPR signal intensities change in different solutions,
the dotted line indicating the level of the background DMPO
signal. Additionally, we also applied the HR scavenger thiourea,
commonly used for validating HR production in EPR assays
(Liszkay et al., 2004). Application of 10 mM thiourea to the
Journal of Cell Science
Arabidopsis ROS-activated channels
Fig. 6. Generation of HRs in Arabidopsis roots, as measured by EPR
spectroscopy and spin-trap DMPO. (A)Control signals in 40 mM DMPO
(this concentration was used throughout) before (top panel) and after (bottom
panel) addition of plants. (B)Effects of 100 (top) and 250 (bottom) mM NaCl.
(C)Effect of 1 mM Cu/a in the absence (top panel) and presence (bottom
panel) of plants. (D)Effect of 1 mM Cu/a in the presence of 1 mM H2O2 (no
plants added). (E)Effect of thiourea. (F)EPR signal intensities under different
conditions (n4-6; mean ± s.e.). Stresses were applied together with plants for
20 minutes. Bathing solution (in mM): 0.1 KCl, 0.1 CaCl2 (pH 6.0).
reaction media caused the disappearance of the four-peak signal,
confirming that NaCl induced HR generation (Fig. 6E).
Involvement of HR-activated K+ channels in programmed
cell death
One important affect of HR-activated K+ loss in animal cells is to
trigger PCD, a ‘self-destruction’ mechanism associated with the
induction of specific K+-inhibited proteases and endonucleases
(Hughes et al., 1997; Yu, 2003; Ajiro et al., 2008). Here, 250 mM
NaCl and oxidative stress (1 mM Cu/a) were applied to initiate cell
death. We found that practically all root cells (in both WT and
gork1-1) showed PCD symptoms (Lam, 2004) during 2-3 days
treatment with these stresses (Fig. 7). Treatment with HR (Cu/a)
caused membrane damage and protoplasm shrinkage (Fig. 7B),
whereas NaCl mainly induced the appearance of condensed dark
areas in place of nuclei (Fig. 7C). However, the observed
morphological changes varied significantly and the extent of
damage was not possible to determine quantitatively. Therefore,
we used biophysical and biochemical approaches to assess whether
HR-activated K+ channels are involved in root PCD.
Sustained depolarisation is perhaps the best marker of cell
damage and death (Bortner et al., 2001); consequently, we measured
changes in the plasma-membrane potential. Membrane potential
tests do not differentiate between PCD and necrosis, but distinguish,
1473
Fig. 7. Changes in Arabidopsis root-cell morphology induced by stress.
(A)No stresses applied. (B)40 hours treatment with 1 mM Cu/a. (C)40 hours
treatment with 250 mM NaCl. Arrows show areas with condensed bodies in
place of nuclei. Bathing solution (in mM): 0.1 KCl, 0.1 CaCl2 (pH 6.0).
very accurately compared with other techniques, dead cells from
those that are alive. Measurements were taken from mature root
epidermal cells in both WT and gork1-1 plants, and the stresses
(250 mM NaCl and 1 mM Cu/a) were washed out for 2 hours
before measurement. For WT cells, 36% and 24% demonstrated
sustained depolarisation (above –40 mV) after 15 hours treatment
with Cu/a and NaCl, respectively (Fig. 8). Forty hours exposure to
250 mM NaCl resulted in the collapse of the membrane potential
in all tested cells, whereas the same exposure to a HR-generating
mixture induced sustained depolarisation in 85% of cells. gork1-1
had about three times fewer cells with a collapsed membrane
potential than WT cells after 15 hours treatment with both stresses
(Fig. 8). The difference between gork1-1 and WT cells was also
evident after 40 hours treatment with stresses; however, it was not
observed for exposure times over 3 days (Fig. 8). Overall, these
measurements show that the absence of functional GORK delays
stress-induced cell death.
Fluorescently labelled protease inhibitor zVAD-fmk (FITCVAD-fmk) is a widely used tool for the observation of PCD
protease activation in intact tissues, because, at micromolar
concentrations, this substance does not significantly inhibit
proteases, although it binds to and fluorescently labels them (Elbaz
et al., 2002; Bonneau et al., 2008). Increased intracellular
fluorescence indicates a higher protease activity and tests on
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Journal of Cell Science 123 (9)
Journal of Cell Science
Fig. 8. Collapse of the membrane potential in Arabidopsis WT and gork1-1
mature root epidermal cells exposed to 1 mM Cu/a or 250 mM NaCl.
(A) 1 mM Cu/a; (B) 250 mM NaCl. Eight to twelve cells from same root were
tested. Measurements are the average for 9-11 roots (open and closed
triangles) and 5-6 roots (open and closed circles) (mean ± s.e.). Wash outs
(2 hours) were applied before measurement. Bathing solution (in mM): 0.1 K+,
0.3 Cl– and 0.1 Ca2+.
Arabidopsis roots showed only a little background staining under
control conditions (Fig. 9). However, fluorescence increased under
stress treatment, indicating protease activation. Fluorescence was
about three to four times lower in gork1-1 compared with the WT
under either HR or NaCl treatments. DMSO, TEA+ and Ba2+
inhibited PCD protease activation, as reported by FITC-VAD-fmk
(Fig. 9A,B,D), demonstrating that the HR-mediated K+-efflux
channel is essential for stress-induced stimulation of proteases.
Longer-term treatment of gork1-1 (>40 hours NaCl) caused a
strong increase in protease activity, indicating that K+-efflux
channels are mainly involved in the early PCD phase (Fig. 9B). A
total of 10 mM Cu/a and H2O2 (used as a positive control) induced
a very strong signal in both WT (fluorescence intensity 494±19.7;
n16; mean ± s.e.) and gork1-1 (fluorescence intensity 468±14.1;
n15; mean ± s.e.) cells (Fig. 9C).
TdT-mediated dUTP nick end labelling (TUNEL) is a standard
technique for visualising DNA cleavage in PCD (Elbaz et al.,
+2002; Reape et al., 2008). Gork1-1 plants revealed about four
times less TUNEL staining after 15 hours exposure to stress (Fig.
10A,B), implying that HR-activated K+-efflux channels are likely
to be involved in endonuclease activation caused by oxidative
stress and salinity. DNase controls confirmed that the measured
fluorescence was caused by DNA degradation (Fig. 10).
Discussion
The data presented here demonstrate that HRs activate an outwardly
directed K+-selective channel with unitary conductance of 16 pS
that is encoded by the Shaker channel GORK (Figs 1-3). Activation
can be induced by physiological concentrations of Cu/a and thus
can probably take place in nature (Fig. 2B,C). The HR-activated
K+ channel mediates K+ efflux caused by both biotic and abiotic
stress – stresses that are able to stimulate HR generation (Figs 46) and are likely to contribute to stress-induced PCD (Figs 7-10).
Overall, this study shows the key functional properties of HRactivated K+ channels and provides evidence for their role in
linking stress-induced ROS, K+ leakage and PCD (Fig. 11).
Fig. 9. The effect of HRs and NaCl on cell-death proteases, as measured
by the FITC-VAD-fmk system. An increase in FITC fluorescence (green
light) indicates protease activity elevation in (A) WT and (B) gork1-1 plants
treated for 15 hours (apart from one 40 hour treatment of gork1-1; right bottom
panel) with control buffer solution (0.1 mM KCl and 0.1 mM CaCl2; pH 6.0),
HR (Cu/a) and 250 mM NaCl. (C)15 hour treatment with solution containing
(in mM): 10 CuCl2, 10 L-ascorbic acid and 10 H2O2. (D)Average fluorescence
intensities under different conditions (n15-20; mean ± s.e.). DMSO (0.3%)
and K+-channel blockers (10 mM TEA+ and 2 mM Ba2+) were applied
together with the stress.
A newly observed mechanism of plant K+-channel
regulation
The regulation of K+ channels is an important issue because these
proteins control membrane potential, ion homeostasis and the
activity of signalling systems (Hille, 2001). Potassium channels
are the best-studied group of plant ion channels (Ward et al., 2009).
They were initially discovered in the late 1970s in algae (Sokolik
and Yurin, 1981), and a detailed analysis of their voltage control,
selectivity, single-channel properties, pharmacology and regulation
by exogenous and endogenous factors, G proteins and
phosphorylation network then followed (Tester, 1990; Maathuis
and Amtmann, 1999; Dreyer and Blatt, 2009; Ward et al., 2009).
The data presented here suggest a new mechanism of regulation of
plant K+ channels – activation by ROS. This mechanism has not
been characterised in plants before, although, in animals, it has
been shown to be important for regulation of blood pressure,
pacemaker activities in the heart and brain, phagocytosis, and
maintaining ionic and osmotic homeostasis (Lahiri et al., 2006).
The genes and structure of plant K+ channels (nine genes in
Arabidopsis) have been extensively investigated (Véry and
Sentenac, 2002; Véry and Sentenac, 2003). Here, we demonstrate
that one of these genes, GORK, encodes a root HR-activated K+
channel. To our knowledge, this is the first identification of the
gene encoding the ROS-activated ion channel in plants. BLAST
analyses (NCBI) show that genes similar to GORK exist in other
plants (such as Populus trichocarpa, Nicotiana tabacum, Samanea
Journal of Cell Science
Arabidopsis ROS-activated channels
1475
Fig. 11. Hypothetical scheme of stress-induced and ROS-mediated K+
efflux from the plant cell. Stress factors interact with receptors that induce
the ROS generation. ROS (such as superoxide and H2O2) can be converted to
HRs by transition metals (TM). This leads to the activation of K+-efflux
channels (GORK), resulting in K+ loss from the cell and stimulation of celldeath enzymes.
Fig. 10. The effect of HRs and NaCl on endonuclease activity, as measured
using fluorescent TUNEL staining. (A)An increase in the FITC fluorescence
(green light) indicates endonuclease activity elevation in WT and gork1-1
plants treated for 15 hours with control buffer solution (0.1 mM KCl and 0.1
mM CaCl2; pH 6.0), HR (Cu/a) and 250 mM NaCl. DNase I (Sigma; 10 units
ml–1) was used for control. (B)Average fluorescence intensities under different
conditions (n12; mean ± s.e.).
saman, Solanum tuberosum), suggesting that HR-activated K+
channels are widespread in the plant kingdom.
The ‘well-defined’ physiological functions of plant outwardly
directed K+ channels include K+ efflux from guard cells leading to
stomata closure (mediated by GORK) and K+ loading to the xylem
(catalysed by SKOR) (Gaymard et al., 1998; De Boer, 1999; Hosy
et al., 2003). K+-efflux channels probably also repolarise the plasma
membrane after depolarisation caused by activation of Ca2+ and
Cl– channels (Tester, 1990; Ward et al., 2009). Our results suggest
that these functions might be regulated by ROS (through HR
activation of GORK).
The activation of K+-efflux channels or an increase in their
expression level has also been found during stresses, such as
salinity, drought, cold, heavy metals and pathogen attack (Blatt et
al., 1999; Becker et al., 2003; Shabala et al., 2006). The data
presented here show that stress does activate K+-efflux channels
through the generation of HRs. This provides, for the first time, a
mechanistic explanation for how the dramatic K+-channel activation
during a number of stresses leads to K+ loss and the death of plant
cells (Fig. 11).
The stimulatory effect of HRs on plant K+ channels appears to
be HR specific; other oxygen-derived species (H2O2, O3 and NO),
which are weaker oxidants than HRs (Halliwell and Gutteridge,
1999), have been shown to inhibit K+ conductance in roots
(Demidchik et al., 2003) and leaves (Torsethaugen et al., 1999;
Köhler et al., 2003; Sokolovski and Blatt, 2004).
Properties of plant HR-activated K+ channels and plant
response to stress
HR-activated K+-efflux channels demonstrate electrophysiological
properties (selectivity, voltage dependence, kinetics of activation,
pharmacological profile and single-channel characteristics) very
similar to those of ‘constitutive’ outwardly rectifying K+ channels
that were previously identified in Arabidopsis (Demidchik et al.,
2006). A classical study carried out by Maathuis and Sanders
showed that Arabidopsis root K+-efflux channels have unitary
conductance of about 15 pS (Maathuis and Sanders, 1995). This is
similar to results reported here (16 pS; Fig. 1G). The unitary
conductance of heterologously expressed GORK is 13.5 pS (Ache
et al., 2000), also very close to the values measured for the HRactivated K+-efflux channel in this study. Taking into account the
absence of HR-induced activation in gork1-1, it can be concluded
that HRs activate a ‘constitutively’ expressed outwardly directed
K+ channel encoded by the GORK gene in Arabidopsis roots.
The presence of two components in the whole-cell HR-activated
K+-efflux current (Fig. 3A) can be explained by either the activation
of two channels encoded by different genes or the coexistence of
two populations of the same channel with different activation
kinetics (Hille, 2001). The latter is more likely for HR-activated
K+-efflux channels because only a 16 pS single channel has been
found in excised patches. Nonetheless, this requires further
investigation.
HRs caused an approximately fivefold increase in the outward
K+ current (Fig. 3A). This is two to three times greater than the
cytosolic alkalinisation-induced outward K+ current previously
measured (Miedema and Assmann, 1996). Locking G proteins in
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Journal of Cell Science 123 (9)
an inactive state using the G-protein blocker guanosine 5⬘-[thio]diphosphate and decreasing the cytosolic Ca2+ concentration
(Li and Assmann, 1993) also causes smaller effects than HRs.
However, the Cladosporium fulvum elicitor resulted in very similar
activation of outward K+ currents in Nicotiana tabacum guard
cells (3–4-fold increase) (Blatt et al., 1999). Interestingly, this
elicitor is known to induce the generation of HR (Veraestrella et
al., 1992).
Notably, both HR production and K+ leakage have been observed
in plants treated with other pathogenic elicitors, originating, for
example, from Botrytis cinerea (Govrin et al., 2006), Alternaria
alternate (Jennings et al., 2002) and Magnaporthe grisea
(Pasechnik et al., 1998). In these cases, however, K+-channel
activation was not characterised. The T. viride elicitor cellulysin,
which previously was shown to induce ROS production in
Arabidopsis roots (Demidchik et al., 2003), activated K+ efflux
and activation was smaller in gork1-1 (Fig. 4D). This shows that
the HR-activated channel GORK can probably mediate elicitorinduced K+ efflux, which could potentially be involved in pathogeninduced PCD.
Journal of Cell Science
Possible mechanisms of K+-channel activation by HRs
H2O2, which accumulates abundantly in plant cells during stress
(Apel and Hirt, 2004), probably requires conversion (catalysed by
transition metals) to the more reactive HR species to activate K+
channels and efflux. Intriguingly, transition metals can be bound
directly within the Shaker K+ channel. For example, animal hERG
channels, which are closely related to plant GORK channels
(BLAST analysis; NCBI), have extracellular transition-metalbinding centres that generate HRs in the presence of H2O2 and are
responsible for channel opening (Yu et al., 1997; Yu, 2003). The
search for and examination of similar metal-binding centres that
can convert H2O2 to HR in GORK (and other plant ion channels)
is currently in progress in our laboratories.
We should not exclude other possible modes of action of H2O2
and HRs on channel activity. Oxidation of membrane lipids and/or
generation of other signalling molecules, such as NO, could also
target the channel; such mechanisms can coexist in animal cells
(Yu, 2003).
The oxidising effect of HRs on plant cells is widely believed to
be irreversible and harmful, although experimental evidence for
this is patchy (Møller et al., 2007). Moderate HR production is
essential for cell growth and shaping, cell-wall loosening and
probably other reactions (Liszkay et al., 2004; Fry, 2004). Although
the role of catalytic copper and iron in plant signalling remains
virtually unexplored, some facts point towards their possible
involvement in plant stress responses. For example, the catalytic
activity of transition metals (Cu+/2+ and Fe2+/3+) increases many
times in plant cells during senescence, pathogen attack and drought
conditions. This leads to an increase in HR generation and cell
death (Becana and Klucas, 1992; Moran et al., 1994; Liu et al.,
2007). Thus, it can be hypothesised that the regulation of catalytic
activities of transition metals could play a crucial role in plant
ROS-mediated regulatory reactions.
Our results demonstrate that the effect of HRs on K+ conductance
is reversible after 20 minutes exposure to a HR-generating mixture.
This ‘regulatory’ phase is relatively short and is followed by
irreversible activation (after 30-45 minutes exposure), very similar
to the effect of HR on animal cation channels (Simon et al., 2004;
Tang et al., 2004). The reversibility of the HR effect could derive
from incomplete and reversible oxidation of amino acids,
particularly His, Cys and Met (Tang et al., 2004). For example,
HR-induced Cys oxidation leads to the sequential formation of
more oxidised derivatives, such as cystine, cysteine sulfenic acid
and cysteine sulfinic acid, all of which are enzymatically reversible
(Biteau et al., 2003).
Cation channels and plant PCD
PCD helps plants to survive under stress (Apel and Hirt, 2004;
Mur et al., 2008; Cutler and Somerville, 2005). Dead cells probably
provide a ‘shield’ from stress factors and signal stress to surviving
cells. Generation of ROS seems to be a key mechanism in plant
PCD (Apel and Hirt, 2004). Stress-induced PCD begins from the
effect of stress factors on as yet undetermined receptors, which
activate ROS-producing enzymes, such as NADPH oxidases,
peroxidases, and chloroplast and mitochondrial redox cascades
(Foyer and Noctor, 2009). It is widely accepted that ROS induces
the elevation of cytosolic free Ca2+; this is not only a stress signal,
but probably also a trigger of PCD reactions that result in damage
to endomembranes and collapse of vacuoles (Apel and Hirt, 2004;
Demidchik and Maathuis, 2007). The data reported here
demonstrate that, as well as Ca2+ elevation, ROS also activate K+
efflux, thereby inducing PCD hydrolase activation.
Our results show that K+-channel blockers and the lack of a
functional GORK both inhibit the stress-induced activation of PCD
proteases and endonucleases (Figs 9 and 10). In animals, these
hydrolytic enzymes are directly inhibited by their natural blocker,
K+, which is at high concentration in the cytosol (70-100 mM) (Yu
et al., 1997; Lam, 2004; Remillard and Yuan, 2004). Death factors
activate animal K+-efflux channels, leading to K+ loss, and this
stimulates protease and endonuclease activity (Yu, 2003; Remillard
and Yuan, 2004). Our data strongly suggest that a similar
mechanism of hydrolase activation exists in plants.
HR-activated K+-efflux channels cannot activate without
depolarisation, which, in plant cells, relies on Ca2+ influx and
anion efflux or can be caused by toxic Na+ influx (Tester, 1990;
Demidchik and Maathuis, 2007). Notably, the involvement of Ca2+
and Cl– channels in plant PCD has already been reported
(Wendehenne et al., 2002). Here, we show that NaCl and oxidative
stress cause depolarisation that is sufficient for K+-channel
activation (Fig. 3B,C).
Other examples of PCD involvement in the plant lifecycle
include self-incompatibility of pollen tubes (Bosch and FranklinTong, 2007), senescence and death of floral organs (Rogers, 2006),
xylogenesis (Fukuda, 2000) and embryogenesis (Reape et al.,
2008). Future experiments should explore a possible role for HRactivated K+-efflux channels in these physiological processes.
The role of HR-activated K+ efflux in NaCl-induced cell
death
Ion disequilibrium is a prime reason for plant damage under salt
stress, causing huge crop yield losses worldwide (Munns and
Tester, 2008). The ability of plants to retain a high K+/Na+ ratio is
crucial for salt tolerance (Chen et al., 2005; Volkov and Amtmann,
2006; Amtmann, 2009). Although some explanations for this
phenomenon exist, they are clearly speculative (Shabala and Cuin,
2008). It is still not established experimentally how K+ loss causes
cell death. It could be hypothesised that K+ efflux from root cells,
mediated by K+-efflux channels activated by salt-induced HR (Fig.
6), stimulates PCD hydrolases, inducing cell death (Fig. 11). In
animal cells, the cytoplasmic and extracellular K+/Na+ ratio is a
major parameter regulating PCD (Bortner et al., 1997; Orlov et al.,
Arabidopsis ROS-activated channels
1999); Na+ cannot substitute for K+ in its protease inhibition
reactions, so the replacement of K+ by Na+ results in PCD (Orlov
et al., 1999). It is obvious, then, that K+ nutrition and K+ fertilisation
are crucially important for preventing plant cell damage and death
mediated by hydrolases. Future work should show how individual
plant caspase-like activities and endonucleases are regulated by
K+.
Conclusions
In conclusion, HR-activated K+-efflux channels have been
characterised in detail. They are similar to constitutive plant K+conductance channels and are probably encoded by the GORK
gene. Although their functions might be very broad, they are clearly
involved in stress-induced K+ loss and stimulation of PCD proteases
and endonucleases.
Materials and Methods
Plant material
Journal of Cell Science
A. thaliana Wassilevskija (WS 0) ecotype was from the European Arabidopsis Stock
Centre (Nottingham). gork1-1 seeds were a generous gift from Hervé Sentenac
(ENSAM, Montpellier, France). Plants were grown vertically, in sterile conditions,
on the surface of 0.4% Phytogel with full-strength Murashige and Skoog media
(Duchefa, Netherlands), 1% sucrose (w/v), pH 6.0 adjusted with KOH (Demidchik
et al., 2003). Four- to nine-day-old root seedlings were used.
Electron paramagnetic resonance spectroscopy
To measure HR generation in intact roots, standard EPR techniques were used
(Halliwell and Gutteridge, 1999; Liszkay et al., 2004). Twenty intact seedlings were
removed from the medium and washed several times in a buffer solution containing
0.1 mM KCl and 0.1 mM CaCl2 (pH 6.0; KH2PO4/K2HPO4). Their 1.5 cm root-tip
parts were immersed in the buffer solution containing spin trap and stress treatments.
The HR-specific spin trap DMPO (Sigma; 40 mM) was purified using activated
charcoal to remove transition metals and decrease the background. It was applied
together with 20 minute stress treatments in a 0.4 ml vial; the vial was sealed from
the air to avoid DMPO oxidation (all solutions were first deoxygenated with N2).
Root cells remained viable (did not lose membrane potential or turgor) upon 20
minutes exposure to DMPO.
After treatment, the eluate from the vial was collected in a flat Bruker EPR
cuvette. The EPR spectra were recorded in the liquid phase at room temperature
using a Bruker EMX (X-band) spectrometer and analysed using the standard Bruker
software. The HR-specific radical scavenger thiourea (Liszkay et al., 2004) was used
to validate HR production. The line-shape of the EPR spectra of the DMPO adducts
(Fig. 6D) was consistent with trapping the HR, because the HR-DMPO and
superoxide-DMPO adduct spectra can be clearly distinguished by specific features
in the line-shape (Yamazaki et al., 1990; Ikeda et al., 2002). It cannot be excluded
that superoxide radicals were generated as precursors of HR, but their activity was
not detected using a DMPO-based system. The superoxide-adduct-specific spectrum
(Yamazaki et al., 1990) was also not observed after either NaCl or Cu/a treatment.
Patch-clamp experiments and membrane potential measurements
Conventional patch-clamp and protoplast-isolation techniques were used (Demidchik
and Tester, 2002; Demidchik et al., 2006; Demidchik et al., 2007, Demidchik et al.,
2009). The standard bathing solution contained (in mM): 1 KCl, 0.3 CaCl2, 2 Tris,
adjusted to pH 5.8 with MES and to 290-300 mOsm with sorbitol. A freshly prepared
mixture of the bathing solution with 1 mM CuCl2 and 1 mM L-ascorbic acid was
applied to generate HRs (Halliwell and Gutteridge, 1999). The standard pipette
solution contained 80 mM K+, 70 mM gluconate–, 10 mM Cl–, 100 nM Ca2+
[adjusted with 1 mM 1,2-bis(o-aminophenoxy)ethane-N,N,N⬘,N⬘-tetraacetic acid] at
pH 7.2 (2 mM Tris/MES). TEA+, Cs+, Na+, Ca2+ and Ba2+ were added as chlorides.
Liquid junction potentials were calculated (corrections were made by amplifier if it
exceeded ±5 mV).
A voltage step protocol was used to activate K+-efflux channels as described
elsewhere (Demidchik and Tester, 2002). Voltage was held at –90 mV, then square
18 second depolarising or hyperpolarising voltage pulses were applied. Whole-cell
or single-channel (outside-out mode) conductances were recorded and analysed
using conventional approaches (Demidchik and Tester, 2002; Demidchik et al.,
2007; Demidchik et al., 2009).
To measure the membrane potential, a single-barrelled borosilicate microelectrode
with a sharp tip and filled with 1 M KCl was inserted into young epidermal cells
(usually 1 to 1.5 cm from the tip – at the beginning of the mature epidermis zone).
Up to eight cells were tested by one electrode. The bathing solution contained (in
mM): 0.3 KCl, 0.1 CaCl2, pH 6.0 buffered with 2 Tris/4 MES. Roots were treated
with HR or NaCl for 15, 40 and 80 hours. The stress was then withdrawn (washed
out for 2 hours) and the resting membrane potential was measured using a FD223
1477
amplifier (WPI, USA). To measure short-term changes (20 minutes exposure) in the
membrane potential (Fig. 3B,C), Cu/a and NaCl were not withdrawn.
Recording of extracellular K+ flux
K+-selective MIFE microelectrodes were fabricated and calibrated as described
elsewhere (Shabala et al., 1997). Arabidopsis roots were immobilised as described
in Shabala et al. (Shabala et al., 2006). During measurement, a computer-controlled
stepper motor moved the electrodes between two positions (20 and 50 m,
respectively) from the root surface in a 10 second square-wave manner. The CHART
software recorded the potential difference between two positions and converted them
to electrochemical potential differences using the calibrated Nernst slope of the
electrodes (Shabala et al., 1997). Ion fluxes were then calculated using MIFEFLUX
software. The bathing solution contained (in mM): 0.1 KCl, 0.1 CaCl2, pH 6.0 (4
MES/2 Tris).
Imaging cell death
An Olympus DX41 fluorescent microscope with standard FITC filters (490/520 nm)
and Olympus analySIS Imaging System and software were used. The exposure time
was 0.5 seconds. The bathing solution contained (in mM): 0.1 KCl, 0.1 CaCl2, pH
6.0 (4 MES/2 Tris). To analyse protease activity, the CaspACE FITC-VAD-fmk in
situ marker kit (Promega, UK) was used. A total of 10 M FITC-VAD-fmk was
applied together with the stress. After 15 hours stress exposure, roots were washed
three times for 5 minutes and observed under the microscope. To determine
endonuclease activity, the DeadEnd Fluorimetric TUNEL System (Promega) was
used. The technique was adjusted for the study of the whole root and observation of
epidermal cells. After treatment, whole roots were fixed with paraformaldehyde and
pasted using inert silicon adhesive to the surface of a microscope slide (roots were
not sliced). Cell walls were permeabilised for 10 minutes using cellulolytic and
pectolytic enzymes (1.5% cellulase Onozuka RS, 1% cellulysin, 0.1% pectolyase Y23, 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0 adjusted by 4 mM MES/2 mM Tris).
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