Download Receptor-like activity evoked by extracellular ADP in Arabidopsis

Plant Physiology Preview. Published on May 11, 2011, as DOI:10.1104/pp.111.174722
Running Head: Receptor-like activity evoked by extracellular ADP
Corresponding author: Julia M. Davies, Department of Plant Sciences, University of
Cambridge, Downing Street, Cambridge, CB2 3EA, United Kingdom.
Tel. 44 1223 333 939
Email: [email protected]
Research area: Cell biology and signal transduction
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Copyright 2011 by the American Society of Plant Biologists
Receptor-like activity evoked by extracellular ADP in Arabidopsis thaliana root epidermal
plasma membrane
Vadim Demidchik1, Zhonglin Shang2, Ryoung Shin3, Renato Colaço4, Anuphon Laohavisit4,
Sergey Shabala5 and Julia M. Davies4*
1
Department of Biology, University of Essex, Colchester, CO4 3SQ, United Kingdom.
2
College of Life Science, Hebei Normal University, Yuhua East Road, Shijiazhang 050016,
Hebei, China.
3
Riken Plant Science Center, Yokohama City, Kanagawa 230-0045, Japan.
4
Department of Plant Sciences, University of Cambridge, Downing Street, Cambridge CB2
3EA, United Kingdom.
5
School of Agricultural Sciences, University of Tasmania, Hobart, Tasmania 7001, Australia
Financial sources: Funding was received from the Leverhulme Trust (F/09 741/C), the
Biotechnology and Biological Sciences Research Council, the University of Cambridge Frank
Smart Trust and Brookes Fund, the Royal Society (China Incoming Fellowship), the Australian
Research Council and The Monsanto Company.
* Corresponding author. [email protected]
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ABSTRACT
Extracellular purine nucleotides are implicated in the control of plant development and stress
responses. While extracellular ATP is known to activate transcriptional pathways via plasma
membrane NADPH oxidase and calcium channel activation, very little is known about signal
transduction by extracellular ADP. Here, extracellular ADP was found to activate net Ca2+ influx
in roots of Arabidopsis thaliana and transiently elevate cytosolic free Ca2+ in root epidermal
protoplasts. An inward Ca2+-permeable conductance in root epidermal plasma membrane was
activated within 1 s of ADP application and repeated application evoked a smaller current. Such
response speed and densitisation are consistent with operation of equivalents to animal
ionotropic purine receptors, although to date no equivalent genes for such receptors have been
identified in higher plants. In contrast to ATP, extracellular ADP did not evoke accumulation of
intracellular reactive oxygen species. While high concentrations of ATP caused net Ca2+ efflux
from roots, equivalent concentrations of ADP caused net influx. Overall the results point to a
discrete ADP signalling pathway, reliant on receptor-like activity at the plasma membrane.
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INTRODUCTION
Extracellular purine nucleotides have been shown to be involved in the regulation of plant cell
viability, membrane permeability, immunity, symbiosis, stress responses and growth (Lew and
Dearnaley, 2000; Tang et al., 2003; Chivasa et al., 2005, 2009; Kim et al., 2006, 2009; Roux and
Steinebrunner, 2007; Wu et al., 2007; Riewe et al., 2008a; Wu and Wu, 2008; Yi et al., 2008;
Demidchik et al., 2009; Govindarajulu et al., 2009; Kim et al., 2009; Clark et al., 2010a,b;
Tanaka et al., 2010a,b; Terrile et al., 2010; Tonón et al., 2010). Purine nucleotide release from
plant cells may occur through wounding, exocytosis or through the activity of plasma membrane
ABC transporters (Thomas et al., 2000; Kim et al., 2006). Growth points are ‘hotspots’ of
extracellular ATP (Kim et al., 2006) while touch-stimulated ATP release is mediated by
heterotrimeric G protein activity (Weerasinghe et al., 2009). Levels of extracellular ATP and
ADP can be regulated by the concerted action of ecto-apyrases, phosphatases and adenosine
nucleosidases with retrieval of adenine (the ultimate hydrolytic product) by plasma membrane
purine permeases (Komoszynski, 1996; Steinebrunner et al., 2003; Wu et al., 2007; Riewe et al.,
2008a, b; Govindarajulu et al., 2009; Clark et al., 2010b; Liang et al., 2010). Manipulation of
ecto-apyrase levels results in abnormal growth in Arabidopsis (Arabidopsis thaliana), cotton
(Gossypium hirsutum) and potato (Solanum tuberosum) (Wu et al., 2007; Riewe et al, 2008a;
Clark et al., 2010a,b) while nodulation by Bradyrhizobium japonicum is impaired in ectoapyrase-deficient soybean (Govindarajulu et al., 2009). Current models suggest that poise of the
cell’s state between death, stress adaptation and growth involves signaling governed by the level
of extracellular ATP (Chivasa et al., 2003, 2009; Roux and Steinebrunner, 2007; Wu et al., 2007;
Wu and Wu, 2008; Kim et al., 2009; Clark et al., 2010a,b; Terrile et al., 2010; Tonón et al.,
2010).
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The mechanism of purine nucleotide perception is unclear as higher plant genomes do not
encode for homologues of animal plasma membrane purinoceptors (Kim et al., 2006; Fountain et
al., 2008). The latter are either coupled to heterotrimeric G proteins or are ionotropic receptors
(activated by nanomolar or even 0.1 mM ATP) that permit Ca2+ influx to transduce the purine
nucleotide signal (reviewed by Khakh and North, 2006). It remains possible that plants evolved
structurally different systems for purine sensing to animals but antagonists of animal
purinoceptors are effective against plant extracellular ATP responses. Despite the apparent
absence of classical purinoceptors in higher plants, extracellular purine nucleotides can cause
transient elevations of cytosolic free Ca2+ ([Ca2+]cyt), in common with animals (Demidchik et al.,
2003a, 2009; Jeter et al., 2004; Tanaka et al., 2010b). In Arabidopsis roots, extracellular ATP
most likely triggers intracellular Ca2+ release, resulting in stimulation of plasma membrane (PM)
NADPH oxidase activity (Demidchik et al., 2009). The resultant reactive oxygen species (ROS)
activate a PM Ca2+ influx pathway that governs a transcriptional response. In Arabidopsis root
hairs, extracellular ATP also causes intracellular ROS accumulation by activating NADPH
oxidases (Clark et al., 2010b). Leaves also support extracellular ATP-induced transcription via
PM NADPH oxidases (Song et al., 2006). Nitric oxide (NO) is now also established as a product
of extracellular purine nucleotide perception. NO evolution by plants has been observed on
application of extracellular ATP or extracellular ADP and linked to influx of Ca2+ or production
of phosphatidic acid (Foresi et al., 2007; Wu and Wu, 2008; Reichler et al., 2009; Clark et al.,
2010b; Sueldo et al., 2010; Tonón et al., 2010).
In this study, the effects of extracellular ADP on ion transport at the plant PM have been
examined. Extracellular ADP elevates [Ca2+]cyt in both roots and leaves of Arabidopsis and it can
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be equally or more effective than extracellular ATP (Demidchik et al., 2003a, 2009; Jeter et al.,
2004; Tanaka et al., 2010b). In common with extracellular ATP, extracellular ADP can elicit
oscillations of [Ca2+]cyt in Arabidopsis seedlings (Tanaka et al., 2010b). At the physiological
level, extracellular ADP stimulates nodule development (Govindarajulu et al., 2009) and was
shown to stimulate root hair elongation (Lew and Dearnaley, 2000). In common with
extracellular ATP, it is now known that extracellular ADP has a biphasic effect on cotton fibre
and root hair elongation with high concentrations being inhibitory (Clark et al., 2010a, b).
Arabidopsis root epidermal cells have been used here as these are well characterised
electrophysiologically and are amongst the first cell types to sense the pathogen attack and
environmental change which could involve purine signaling. By using a fast perfusion system in
patch clamp electrophysiology experiments, extracellular ADP has been delivered to the
extracellular PM face of protoplasts within 1 s, permitting the fastest resolution to date of rapid
activation of plant ion transport events by an extracellular purine nucleotide. These events appear
to be independent of heterotrimeric G proteins. Epidermal protoplasts expressing the [Ca2+]cyt
reporter aequorin have been used to explore further the ability of extracellular ADP to evoke
[Ca2+]cyt elevation. In further contrast to extracellular ATP, extracellular ADP does not elicit
intracellular ROS accumulation. Using whole roots, it has also been possible to resolve
extracellular ADP-induced net Ca2+ and K+ fluxes; high concentrations of ATP but not ADP
evoke net Ca2+ efflux. Overall the results show rapid activation of PM Ca2+- and K+conductances by extracellular ADP, with initial events supported by PM ionotropic receptor-like
activity. The differential responses of intracellular ROS and net Ca2+ flux to extracellular ADP
and extracellular ATP could provide the basis for distinct signaling outcomes.
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RESULTS
Extracellular ADP activates a PM Ca2+ influx conductance
To establish whether the PM could respond to extracellular ADP, protoplasts from mature
epidermal root cells were assayed using the whole cell patch clamp configuration to resolve
activity of channel populations as a function of applied membrane voltage and extracellular
agonist. Application of 20 μM extracellular ADP to protoplasts (using conventional peristaltic
pump delivery to the recording chamber) transiently activated a hyperpolarisation-activated (i.e.,
activated by increasingly negative voltage) Ca2+-permeable conductance (Figure 1A), very
similar to that characterized in response to 20 μM extracellular ATP in this cell type (Demidchik
et al., 2009). The concentration used is representative of wounding levels (Song et al., 2006).
The increase in inwardly-directed current at hyperpolarized (inside negative) voltage was
observed 1-3 min after extracellular ADP addition (a similar time course to the extracellular ATP
response reported by Demidchik et al., 2009 and Shang et al., 2009) and was evident over the
physiological voltage range for this cell type (Maathuis and Sanders, 1993). Mean ± SE currentvoltage (IV) relationships are shown in Figure 1B; negative voltages are physiologically relevant.
At -200 mV, extracellular ADP induced a peak mean ± SE current increase from -163 ± 36 pA to
-382 ± 63 pA at -200 mV (n = 4); current then declined over a 5-15 min period. Negative current
signifies influx of positive charge into the cytosol or efflux of negative charge. The peak current
response is not significantly different (Student’s t-test) to that found for 20 μM extracellular ATP
(Demidchik et al., 2009).
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Rapid, reversible and desensitising response to extracellular ADP at the PM
Having found that the PM was electrically responsive to extracellular ADP on slow perfusion
and long-term exposure, a fast perfusion system was used to deliver extracellular ADP to the PM
within 1s. In contrast to experiments using peristaltic pump delivery of the agonist to the
recording chamber (with a waiting time for flow from the chamber edge to the protoplast), the
fast perfusion system used a primed multi-channel delivery pipette placed near the protoplast
(described further in Materials and Methods). In these experiments, the PS (pipette solution,
equivalent to the cytosol) contained a non-hydrolysable GDP analog (0.5 mM GDPβS) to
minimise G-protein-mediated channel activation (Li and Assmann, 1993). In preliminary
experiments, this increased the K+ efflux current, as described by Li and Assmann (1993),
indicative of effective G-protein inhibition. It has been proposed that extracellular purine
nucleotides can relay a signal via heterotrimeric G proteins in plants (Song et al., 2006, Roux and
Steinebrunner, 2007; Wu and Wu, 2008), so their inhibition could delineate possible direct
extracellular ADP effects on a receptor-like channel. Experiments were performed within the
first 20-30 min of recording so that extracellular ADP-activated currents could be distinguished
from the time-dependent development of the constitutive hyperpolarisation-activated Ca2+permeable conductance. The latter becomes the dominant Ca2+ conductance in this cell type after
approximately 1 h (Demidchik et al., 2002).
Fast perfusion proved problematic as in approximately 80% of trials (65 out of 85), the recording
configuration was disturbed or lost by distortion of the protoplast. Given the high failure rate, a
higher concentration of extracellular ADP (0.3 mM) was used in these trials to ensure activation.
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In the remaining 15 trials in which seal resistance was unaffected by fast perfusion, extracellular
ADP evoked a rapid and reversible change in PM conductance (Figure 2A). With the PM
clamped at a holding potential (HP) of -90 mV, fast perfusion with control bathing solution (BS)
was without effect (Figure 2A, top trace). Addition of 0.3 mM extracellular ADP caused a
downward current deflection, consistent with entry of positive charge into the protoplast (Figure
2A, middle trace). Current activation occurred immediately after exposure to extracellular ADP.
Fast removal of extracellular ADP resulted in immediate recovery to control conditions. As can
be seen from the middle trace of Figure 2A, a repeated application of 0.3 mM extracellular ADP
to the same protoplast again caused rapid and reversible current activation. However in this case,
recovery to baseline current began in the presence of extracellular ADP. This was typical of all
protoplasts where extracellular ADP activation was observed.
Mean ± SE values of extracellular ADP-evoked currents at HP = -90 mV are given in Table 1.
In experimental Set A, the first application of 0.3 mM extracellular ADP (Event 1) caused a
mean 89% increase in inwardly-directed current. After removal of the stimulus and recovery to
baseline current, a second application of 0.3 mM extracellular ADP (Event 2) caused a mean
54% increase. All differences are statistically significant (Student’s t-test; P < 0.05). In
experimental Set B, Event 1 (0.3 mM extracellular ADP) evoked a mean 101% increase in
inwardly-directed current at HP = -90 mV; in Event 2, application of 0.3 mM extracellular ADP
and 0.3 mM Gd3+ to the same protoplasts only caused a mean 11% increase. Thus, the greater
part of the extracellular ADP-induced current was sensitive to the cation channel blocker, Gd3+.
To explore cation channel activity further, IV relationships were determined.
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Extracellular ADP increases PM Ca2+ influx and K+ efflux conductance
As Figure 2B shows, fast delivery of 0.3 mM extracellular ADP increased both inwardly- and
outwardly-directed currents across the PM of a single protoplast, resulting in a positive shift of
reversal potential (Erev) away from the equilibrium potentials for K+ and Cl- (EK = -158 mV; ECl
= -36 mV) and towards that for Ca2+ (ECa = 155 mV). The mean Erev of the extracellular ADPinduced current (estimated by subtracting the control IV from that determined in the presence of
extracellular ADP; Figure 3 inset) was 4.7 ± 8.4 mV (n = 4), indicating Ca2+ permeation.
Extracellular ADP approximately doubled the Gd3+-sensitive inward current (Figure 2B and
Figure 3) over the physiological voltage range for this cell type (-100 to -150 mV; Maathuis and
Sanders, 1993); thus, Ca2+ could be delivered to the cell to act as a signal in response to the
extracellular purine nucleotide. For example, from the mean response shown in Figure 3, at -120
mV inward Ca2+ current increased from -69 ± 23 mA m-2 to -159 ± 25 mA in response to 0.3 mM
extracellular ADP (n = 4; mean ± SE). This large Gd3+-sensitive Ca2+ influx current helps
explain an extracellular ADP-induced elevation in whole root and root epidermal protoplast
[Ca2+]cyt found in aequorin luminometry assays (Demidchik et al., 2003a, 2009). In our
experimental conditions only K+ efflux (50 mM K+ in the PS) could mediate the outward current
(Shabala et al., 2006). Both inward and outward currents were also sensitive to the cation
channel blocker Gd3+. It may be that both Ca2+ influx and K+ efflux conductances were caused
by the activation of one cation channel type or that Ca2+ influx activated K+ efflux.
Extracellular ADP elevates root epidermal [Ca2+]cyt
Epidermal protoplasts were isolated from the mature epidermis of plants constitutively
expressing cytosolic apoaequorin to examine extracellular ADP-induced [Ca2+]cyt elevation.
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Previously, such protoplasts were found to sustain transient [Ca2+]cyt elevations in response to 0.1
mM extracellular ATP, its poorly hydrolysable analog adenosine 5'-(α,ß-methylene)triphosphate
(αßATP) and ADP. All these were inhibited by Gd3+ and the purinoceptor antagonist, suramin
(Demidchik et al., 2009). AMP (as an ADP breakdown product) up to 1 mM has been shown to
be ineffective (Demidchik et al., 2009). A test concentration of 0.1 mM ADP was used in the
present study as it is a feasible concentration to simulate a wound response (Song et al., 2006)
and was found to cause Ca2+ current activation in 4 out of 9 whole cell patch clamp trials using
slow perfusion (data not shown). Application of 0.1 mM extracellular ADP at pH 6.0 caused a
transient increase in [Ca2+]cyt (Figure 4A). Overall the mean peak [Ca2+]cyt increase caused by
this extracellular ADP concentration was 0.43 ± 0.02 μM [Ca2+]cyt (n = 52), a value comparable
to that reported previously for protoplasts from this cell type in response to 0.1 mM ADP or ATP
(Demidchik et al., 2009). Extracellular ADP was added as its Na+ salt; NaCl as a control is not
effective as an elicitor in this assay (Demidchik et al., 2009). The extracellular ADP-induced
increase depended on Ca2+ influx as depletion of external Ca2+ from 1 mM to 5 μM in a paired
test reduced the extracellular ADP-induced increase by 84 % (0.1 mM extracellular ADP, 1 mM
Ca2+; 0.38 ± 0.04 μM [Ca2+]cyt,, n = 3: 0.1 mM extracellular ADP, 5 μM Ca2+; 0.06 ± 0.05 μM
[Ca2+]cyt, n = 4). An application of 0.1 mM extracellular ADP was used as the standard stimulus
in further trials.
The peak [Ca2+]cyt response to extracellular ADP was insensitive to the G protein inhibitor
pertussis toxin (PTX; 200 ng ml-1: Alloisio et al., 2004) (extracellular ADP; 0.34 ± 0.04 μM
[Ca2+]cyt , n = 6: extracellular ADP + PTX; 0.34 ± 0.02 μM [Ca2+]cyt, n = 11). The cation channel
blocker, Gd3+, is already known to inhibit the extracellular ADP-induced elevation of [Ca2+]cyt in
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these epidermal protoplasts (Demidchik et al., 2009). TEA+ (10 mM), a K+ channel blocker,
significantly inhibited the [Ca2+]cyt elevation by 25% (extracellular ADP; 0.63 ± 0.02 μM
[Ca2+]cyt , n = 6: ADP + TEA+; 0.47 ± 0.03 μM [Ca2+]cyt , n = 6; P<0.001, ANOVA). This
suggests that K+ channels may be involved in changing membrane voltage regulating driving
force for the Ca2+ influx. Depolarising the PM with 80 mM K+ (Demidchik et al., 2002)
significantly reduced the [Ca2+]cyt response by 55%, consistent with reduction of Ca2+ influx at
depolarised voltage in patch clamp trials (1 mM K+; 0.55 ± 0.03 μM [Ca2+]cyt , n = 5: 80 mM K+;
0.25 ± 0.01 μM [Ca2+]cyt , n = 4;P<0.001). The effect of pH was tested against the standard
application of 0.1 mM extracellular ADP, revealing a clear neutral pH optimum (0.57 ± 0.02 μM
[Ca2+]cyt , n = 4) and a significantly enhanced response at alkaline pH compared to physiological
apoplastic pH 5-6 (Figure 4B;P<0.001). Nevertheless, at pH 5, elevation of [Ca2+]cyt remained
significant (control [Ca2+]cyt baseline pH 5, 0.11 ± 0.02 μM; [Ca2+]cyt peak, 0.44 ± 0.03 μM, n =
5;P<0.001). In separate tests (using a plate reader rather than a luminometer), although absolute
values of peak [Ca2+]cyt were lower, the pattern of a greater response at neutral pH rather than pH
6 was retained for 0.1mM ADP and also 0.1 mM ATP (Figure 4C, n = 4;P<0.05). Response to
ATP was significantly greater than to ADP at both pH. The response to the poorly hydrolysable
ADP analog ADPßS (adenosine 5'-O-(2-thiodiphosphate) was not significantly different to the
ADP response while αßATP evoked significantly lower responses than ATP at both pH (n =
4;P<0.05). Thus part of the ATP response may involve hydrolysis products but the ADP
response does not.
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Extracellular ADP does not increase intracellular levels of ROS
In Arabidopsis roots, extracellular ATP causes accumulation of intracellular ROS via activation
of the PM NADPH oxidase AtRBOHC (Demidchik et al., 2009). At the whole root level,
extracellular ADP was applied under the same conditions used by Demidchik et al. (2009) for
intracellular
ROS
imaging
(using
ester-loaded
5-(and-6-)-chloromethyl-2’,7’-
dichlorohydrofluorscein diacetate) in response to extracellular ATP. Solutions were designed to
ensure that extracellular [Ca2+] remained constant at 0.1 mM throughout experimental treatments
as purine nucleotides are potent Ca2+ chelators. Chelation of extracellular Ca2+ can trigger
extracellular superoxide anion production by roots (Mortimer et al., 2008). Here, extracellular
ATP was found again to cause intracellular ROS accumulation (Figure 5A). In contrast,
extracellular ADP did not result in significant accumulation of intracellular ROS (Figure 5A,B).
Previously a response was seen by 15 s after extracellular ATP application (0.01 to 1 mM) with
0.1 mM extracellular ATP causing over a threefold increase in signal intensity (Demidchik et al.,
2009). Here, the same concentration range for extracellular ADP under identical conditions did
not evoke intracellular ROS production, even over an extended observation period. Under
control conditions, mean ± SE fluorescence intensity of single primary roots was 19 ± 0.9
fluorescence per pixel (n = 23) and there were no significant increases on application of
extracellular ADP (Tukey’s multiple comparison test; Figure 5B). Separate trials using a
different imaging system (Leica) and with extracellular Ca2+ maintained at 0.1 or 1 mM resulted
in higher control values than recorded previously, showing the variable nature of ROS
accumulation (Figure 5C,D). Extracellular ATP (1 mM) caused an increase in fluorescence
intensity with 1 mM Ca2+ but 1 mM ADP did not (Figure 5C, D; n = 11). Fluorescence intensity
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with ADP was significantly lower than with ATP (Student’s t-test;P<0.02). Overall the results
suggest that the extracellular ADP/[Ca2+]cyt pathway is independent of intracellular ROS.
Extracellular ADP regulates net Ca2+ and K+ fluxes from excised roots
To test whether extracellular ADP can activate cation fluxes in intact cells, we monitored net
fluxes at the epidermis of excised roots using the MIFE system for non-invasive microelectrode
Ca2+- and K+-flux measurements. This permits fine spatio-temporal mapping of net fluxes.
Constancy of extracellular Ca2+ (0.1 mM Ca2+) ensured that changes in flux were due to
extracellular ADP perception alone (rather than changes in [Ca2+]) and also aided electrode
function. Application of extracellular ADP (Na+ salt) to the elongation zone epidermis caused
transient increases in net Ca2+ influx (i.e., more positive flux values) in a dose-dependent manner
(Figure 6A and B, n = 3-6 for 1 μM to 1 mM extracellular ADP). A significant increase above
basal level was first detected at 3 μM (P<0.02; Student’s t-test). Duration of transients was up to
4 minutes. Control flux levels were recovered. In control experiments for the effect of Na+, 4
mM NaCl had no effect on net flux (n = 4).
Net Ca2+ influx in response to extracellular ADP was less pronounced in mature zone epidermis
and [ADP] below 10 μM did not evoked significant changes (Figure 6C, n = 3-7). The mature
zone has been found previously to be less receptive to exogenous ROS than the elongation zone
(Demidchik et al., 2003b, 2009). Differences between elongation zone and mature zone were
clearer for extracellular ADP-induced K+ efflux (Figure 6D), with elongation zone epidermis
(Figure 6E, n = 3-6) supporting greater net K+ efflux than the mature zone (Figure 6F, n = 3-7)
across the concentration range tested. Significant changes in K+ flux relative to basal were
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observed at 1 μM ADP in both zones. Extracellular AMP was found previously to be a very
weak inducer of root Ca2+ influx (Demidchik et al., 2009) and seedling [Ca2+]cyt elevation
(Tanaka et al., 2010b). Here, AMP did not elicit a K+ efflux response even at 1 mM, in either the
elongation zone or mature epidermis (elongation zone control; -37 ± 34 nmol.m-2.s-1, n = 3:
elongation zone + AMP, -31 ± 26 nmol.m-2.s-1, n = 3. Mature zone control; -19 ±13 nmol.m-2.s-1,
n = 3: AMP, -10 ± 18 nmol.m-2.s-1, n =3). This suggests that breakdown of extracellular ADP to
AMP would help terminate the cell’s response.
Extracellular ATP and its poorly hydrolysable analog adenosine 5'-(α,ß-methylene)triphosphate
(αßATP) were also tested. The pattern of greater net Ca2+ and K+ fluxes at the elongation zone
was retained for both agonists (Figures 7 and 8). In the elongation zone, both extracellular ATP
and αßATP up to 100 μM caused transient net Ca2+ influx comparable to that evoked by
extracellular ADP (Figure 7A-D). The threshold of the detectable ATP response lay between 0.3
and 1 μM (n = 6). However while at 300 μM and 1 mM extracellular ADP promoted net Ca2+
influx, ATP and αßATP caused net Ca2+ efflux (n = 5-17; Figures 7C,D). The mean peak net
Ca2+ efflux caused by 1mM αßATP was almost double that caused by 1mM ATP (Figure 7C,D).
At the mature epidermis, while 1mM ADP caused net Ca2+ influx, both 1 mM ATP and αßATP
again caused net Ca2+ efflux with αßATP eliciting the greater response (Figure 7E; 1 mM ATP,
-5 ± 21 nmol.m-2.s-1, n =8). Thus ATP hydrolysis appears necessary to limit the Ca2+ efflux
response. At both elongation zone and mature epidermis, both extracellular ATP and αßATP
caused dose-dependent net K+ efflux in common with ADP (Figure 8). Thus while at high
concentrations of ADP net Ca2+ influx and K+ efflux appear as a coupled response, net Ca2+ flux
is uncoupled from K+ efflux at high concentrations of ATP or αßATP.
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DISCUSSION
Evidence is now accumulating that extracellular ATP is a regulator of plant physiology and
development (Lew and Dearnaley, 2000; Tang et al., 2003; Chivasa et al., 2005,2009; Kim et al.,
2006, 2009; Roux and Steinebrunner, 2007; Wu et al., 2007; Riewe et al., 2008a, b; Wu and Wu,
2008; Yi et al., 2008; Demidchik et al., 2009; Reichler et al., 2009; Clark et al., 2010a,b; Tanaka
et al., 2010a,b; Terrile et al., 2010; Tonón et al., 2010). In common with ATP, it is likely that
ADP is released on wounding and could perhaps also be released with ATP by exocytosis or the
activity of ABC transporters (Thomas et al., 2000; Kim et al., 2006; Tanaka et al., 2010a).
Extracellular ATP “hotspots” have been found in elongating root cells (Kim et al., 2006) and
ADP may also be released there. This would be consistent with the finding here that the
Arabidopsis root elongation zone epidermis supported a greater flux response to extracellular
ADP than the mature zone. Such data support the premise that the sensitivity of different cells to
extracellular purine nucleotides differs (Tanaka et al., 2010b).
Suramin (a purinoceptor
antagonist) inhibits Arabidopsis root elongation and extracellular ADP-induced [Ca2+]cyt
elevation (Demidchik et al., 2009). Moreover, there is a dose-dependent effect of extracellular
ADP on Arabidopis root hair elongation (Clark et al., 2010b) further implicating extracellular
ADP in growth regulation. Plant extracellular apyrases can hydrolyse ADP to regulate its
extracellular levels and so terminate any signal it could invoke (Komoszynski, 1996;
Steinebrunner et al., 2003; Riewe et al., 2008a, b; Tanaka et al., 2010a,b).
Results here show that the epidermis is a contributory cell type to the whole root elevation of
[Ca2+]cyt by extracellular ADP observed by Demidchik et al. (2003a) and the whole seedling
response, thought to be generated largely by the root, observed by Tanaka et al., (2010b).
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Moreover, that protoplasts could sustain the [Ca2+]cyt elevation demonstrates that the wall is not
critical for the response of this cell type. Previously, it had been suggested that wall proteins
capable of phosphorylation intercept extracellular purine nucleotides to evoke a cellular response
(Chivasa et al., 2005). The results also show that extracellular ADP can generate a [Ca2+]cyt
signal in the mature epidermis seemingly independently of heterotrimeric G protein activity.
Further studies on the epidermis now need to focus on G protein mutants.
The response speed of the PM to extracellular ADP in the fast perfusion patch clamp trials here
is consistent with the receptor-like activity suggested by Lew and Dearnaley (2000) to explain
extracellular ADP-induced depolarization of Arabidopsis root hair PM voltage. Previously,
extracellular ADP-induced rapid elevation of [Ca2+]cyt in root epidermal protoplasts was inhibited
by 92% with the same concentration (0.3 mM) of Gd3+ used here to block PM influx currents
(Demidchik et al., 2009). The very rapid response of the PM inward Ca2+ conductance to
extracellular ADP (fast perfusion) and its block by 0.3 mM Gd3+ raises the possibility that Ca2+
influx is an initial event. In common with animal purinceptor responses (Roberts and Evans,
2007), extracellular ADP-induced current decayed in the continued presence of the agonist and
repeated application of extracellular ADP to the epidermal PM resulted in a lower level of
current activation (Figure 2A and Table 1). Thus, a [Ca2+]cyt signal evoked by extracellular ADP
release could be diminished by desensitisation and terminated by apyrase or phosphatase
activity. This helps explain the transient nature of the [Ca2+]cyt response and perhaps also that of
the epidermal Ca2+ and K+ fluxes. Efflux of K+ could readily counter the depolarising effect of
purine nucleotide-induced Ca2+ influx on PM voltage and help return this membrane to a
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hyperpolarised state. The greater magnitude of the K+ efflux compared to Ca2+ influx suggests
that K+ efflux may also serve other purposes.
Previously, extracellular ATP was found to stimulate production of extracellular superoxide
anion by Arabidopsis etiolated hypocotyls (Tonón et al., 2010), and leaves (Song et al., 2006). In
leaves, this was through activation of PM NADPH oxidases (Song et al., 2006). Extracellular
ADP stimulated leaf and root extracellular superoxide anion production by an unknown
mechanism (Song et al., 2006; Demidchik et al., 2009).
In roots of Arabidopsis, Medicago
truncatula and Phaseolus vulgaris extracellular ATP also resulted in accumulation of
intracellular ROS, essentially H2O2 (Kim et al., 2006; Cárdenas et al., 2008; Demidchik et al.,
2009; Clark et al., 2010b). In the present study, and in contrast to extracellular ATP, extracellular
ADP did not trigger accumulation of intracellular ROS. This is in agreement with the findings of
Kim et al. (2006) in which extracellular ATP but not extracellular ADP triggered intracellular
ROS accumulation in M. truncatula root hairs. Certainly, there is a precedent for triggers of Ca2+
influx resulting in distinct ROS responses. In Arabidopsis cell suspension cells, both hyper- and
hypo-osmotic stress trigger Ca2+ influx but only hypo-osmotic stress results in extracellular H2O2
accumulation (Beffagna et al., 2005). Extracellular superoxide anions (resulting from perception
of extracellular purine nucleotides) could generate H2O2 which could then be translocated to the
cytosol by PM aquaporins (Bienert et al., 2007; Dynowski et al., 2008). One possible explanation
for the disparity between the intracellular H2O2 accumulation response to extracellular ATP and
extracellular ADP is that in the extracellular ADP pathway, H2O2 translocation is blocked. This
could be achieved by closure and/or retrieval of H2O2-permeable aquaporins (Boursiac et al.,
2008). Alternatively, extracellular ADP perception could inhibit an intracellular ROS-generating
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system, stimulate an intracellular anti-oxidant activity or extracellular ADP perception in the
apoplast could result in suppression of extracellular ROS production downstream of superoxide
anions. Given the importance of ROS in plant cell signaling, these possibilities merit further
investigation particularly in the context of nodule development. Extracellular ADP has been
found to regulate this process which is known to involve complex spatio-temporal changes in
ROS (Chang et al., 2009; Govindarajulu et al., 2009). Whether ATP-dependent intracellular ROS
accumulation is the cause of net Ca2+ efflux from the root epidermis at high [ATP] now needs to
be determined as this may be relevant to the suppression of defense responses noted when
exogenous ATP is increased (Chivasa et al., 2009).
The pH dependence of animal ionotropic purinoreceptors varies with isoform (King et al., 1997;
Wildman et al., 1999; Wirkner et al., 2008). Here, the extracellular ADP-induced [Ca2+]cyt
response was operative over a wide pH range but activity at more neutral to alkaline pH suggests
that extracellular ADP (and ATP) signaling could operate during stress responses in which the
cytosol acidifies and apoplastic pH elevates above its normal acid value, such as hypo-osmotic
stress or elicitor challenge (Mathieu et al., 1996; Pugin et al., 1997; Beffagna et al., 2005).
Additionally, pH-dependence of ADP- or ATP-induced Ca2+ and K+ flux could be relevant at
growth points, such as the root hair apex, at which apoplastic pH oscillates and pH oscillations
are linked to nitrogen availability (Monshausen et al., 2007; Bloch et al., 2011). Overall,
extracellular ADP is competent to act as a trigger of [Ca2+]cyt-dependent signalling with the
potential to evoke responses distinct to those produced by ATP. Its initial perception at the PM
appears reliant on ionotropic receptor-like activity.
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CONCLUSION
Perception of extracellular ADP at the root epidermal plasma membrane is consistent with the
operation of functional equivalents of ionotropic purine nucleotide receptors. In contrast to
extracellular ATP, downstream events do not include accumulation of intracellular ROS thus
indicating disparate signalling pathways.
MATERIALS AND METHODS
Plant growth
Arabidopsis thaliana (Col-0, Col-0 expressing apoaequorin under a 35s promoter; Demidchik et
al., 2009) from laboratory stock was grown vertically on full-strength Murashige-Skoog (MS,
Duchefa; www.duchefa.com) medium with 1% (w/v) sucrose and 0.3% (w/v) Phytagel (Sigma;
www.sigmaaldrich.com). Growth conditions were 22 ºC and 16 h daylength, 100 μmol⋅m-2⋅s-1
irradiance.
Patch clamp electrophysiology
Protoplasts were isolated from root mature epidermis (10-15 day plants) using the protocol
described by Demidchik et al. (2003b). Protoplasts used for patch clamping were 20 ± 1.5 μm in
diameter. The pipette solution (PS) contained (mM): 40 K-gluconate, 10 KCl, 1 1,2-bis(2aminophenoxy)ethane-N,N,N’,N’-tetraacetic acid (free Ca2+ adjusted to 100 nM with 0.4 CaCl2),
pH 7.2 adjusted with Tris/MES. The G-protein antagonist GDPβS (0.5 mM) was incorporated
into the PS as indicated. Control bathing solution (BS) comprised (mM): 20 CaCl2, 0.1 KCl, 0.3
NaCl (varied to supply equimolar Na+ to that in purine-containing BS), pH 6 with 2 Tris/4 MES.
BS containing purine agonist was freshly prepared directly before application and the pH of the
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solution was adjusted where necessary (Tris). BS and PS were adjusted to 290 to 300 mOsM
with D-sorbitol. Delivery of BS +ADP to the recording chamber was either by gravity feed or by
a fast flow system comprising an AutoMate Perfusion System with 4 pinch valves, ValveBank®
controller and four-barrel delivery pipette (barrel diameter 0.1 mm; AutoMate Scientific Inc.,
CA, USA). This system allowed switching to test BS in 0.01 s. Average delivery time was less
than 1 s. The delivery pipette was placed at 30-40 μm from the protoplast surface and delivery
speed was 0.5-1 ml s-1 Two barrels of the delivery pipette were used – one for control BS and
another for purine-containing BS.
Solution changes were controlled by the programmable
ValveBank® controller. Recording equipment was as described in Demidchik et al., 2003b. Ion
activities were calculated using Geochem (Parker et al., 1995). Liquid junction potentials were
measured and corrected as described previously (Demidchik et al., 2003b).
[Ca2+]cyt measurement
The protocol described in Demidchik et al., 2007 was used to isolate protoplasts from root
mature epidermis of Arabidopsis (10-15 days old) expressing cytosolic (apo)aequorin (CAMV
35S promoter) with the modification that incubation with protoplasting enzymes was increased
from 50 to 60 min. Protoplasts were suspended for 3-4 hours (dark, 28 oC) in recording medium
(mM: 1 CaCl2, 1 KCl, 1 MgCl2, 10 glucose; pH 6.0 adjusted by 2 Tris/4 MES; 290-300 mOsM
with D-sorbitol) with 10 μM coelenterazine (NanoLight Technology; www.nanolight.com) to
match the protocol used by Demidchik et al. (2009) for ATP and ADP studies. After washing,
recording medium (without coelentrazine) and standard luminometry procedures were used. The
discharging solution enabling estimation of [Ca2+]cyt contained 2 M CaCl2 and 20 % (w/v)
ethanol. Luminescence was recorded using a luminometer or FLUOstar plate reader.
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Imaging of reactive oxygen species
Imaging of intracellular reactive oxygen species used 50 μM 5-(and-6-)-chloromethyl-2’,7’dichlorohydrofluorscein diacetate, acetyl ester (Molecular Probes; www.invitrogen.com) with
dye loading as described previously (Demidchik et al., 2009). Assay medium was as described
by Demidchik et al., 2009 and comprised (mM): 0.1 KCl, 0.1 or 1 CaCl2, 4 MES and 2 Tris base
(pH 6.0). Images of 5 day old roots were acquired either with a Nikon SMZ 1500 microscope
and a QImaging Retiga cooled 12-bit camera or a Leica M165 FC microscope and a Leica
DFC310 FX 1.4-megapixel camera. Ca2+ was maintained at 0.1 or 1 mM in the presence of
purines by appropriate additions of CaCl2 (calculated using MaxChelator and Geochem
programs; Patton et al., 2004; Parker et al., 1995).
MIFE recordings of net cation fluxes
Excised roots (5-15 days old) were fixed into a recording chamber by an agar drop and immersed
in 0.5 ml control solution comprising (in mM) 0.1 KCl, 0.1 CaCl2, 4 MES, 2 Tris base (pH 6.0)
(Demidchik et al., 2009). Chamber design permitted rapid mixing of test solution and resumption
of flux recording 20-30 s after addition of 0.5 ml test solution. Ca2+ was maintained at 0.1 mM in
the presence of purines by appropriate additions of CaCl2. Levels were confirmed experimentally
in a cell-free assay using a Ca2+-selective microelectrode. Purines were added as Tris, Na+ or Li+
salts. NaCl or LiCl at equimolar concentrations did not perturb K+ efflux (tested experimentally).
Measurements of mature epidermal flux were taken 1-1.5 mm from the root apex and for the
elongation zone, 100 -150 μm from the apex (Demidchik et al., 2007). Net Ca2+ influx and K+
efflux were determined using the non-invasive slowly-vibrating microelectrode (MIFE)
technique. K+- and Ca2+-selective microelectrodes (external tip diameter approximately 3 μm)
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Copyright © 2011 American Society of Plant Biologists. All rights reserved.
were made and calibrated as described previously (Shabala and Shabala, 2002). Electrodes were
calibrated before and after use. Agonists (tested up to 1 mM) had no effect on calibration or
electrode response time. The microelectrode was placed 20 μm above the root surface and
moved between two positions, 20 and 50 μm above the root surface in a square wave manner (5 s
half cycle).
ACKNOWLEGEMENTS
We thank Daniel Schachtmann for advice and Adeeba Dark for help with plant cultivation.
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Figure legends
Figure 1. Extracellular ADP activates a PM influx conductance in root epidermal
protoplasts. (A) Effect of 20 μM extracellular ADP on whole cell currents from a representative
protoplast, recording conditions were as used previously by Demidchik et al., 2009. (B) Mean ±
SE IV relationships of control and peak ADP peak response (n = 4). Control BS was as described
in Materials and Methods (20 mM CaCl2, 0.1 mM KCl and Na+ equimolar to test BS, pH 6). PS
comprised (mM): 40 K-gluconate, 10 KCl, 1 1,2-bis(2-aminophenoxy)ethane-N,N,N’,N’tetraacetic acid (free Ca2+ adjusted to 100 nM with 0.4 CaCl2), pH 7.2 adjusted with Tris/MES.
Figure 2. Rapid activation of epidermal PM cation conductances by extracellular ADP. The
results are from whole cell recordings on protoplasts from the root mature epidermis. (A) Top
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Trace: Two sequential applications of control bathing solution (BS; green bars) to a protoplast
from the mature epidermis failed to elicit a current; holding potential (HP) across the PM was 90 mV inside negative. Middle trace (same protoplast): Two sequential applications to the same
protoplast of 0.3 mM ADP (red bars) by fast perfusion evoked inwardly-directed current
(downward deflection). Baseline current was recovered on removal of the ADP stimulus. Bottom
trace (different protoplast); inhibiting effect of 0.3 mM Gd3+, added together with ADP in the
second application. BS was as in Figure 1. Test BS included 0.3 mM NaADP ± 0.3 mM GdCl3
as indicated. PS included 0.5 mM GDPβS. (B) Current in response to voltage ramp (40 s-long
started from +100 mV) recorded from a single protoplast, approximately 1 min after the addition
of control BS (green), 0.3 mM ADP (red) or 0.3 mM ADP with 0.3 mM Gd3+ (blue). Solutions
as in (A).
Figure 3. Extracellular ADP activates Ca2+ influx and K+ efflux in epidermal protoplasts.
Mean ± SE IV relationships for control conditions and in the presence of 0.3 mM extracellular
ADP, n = 4; solutions as in Figure 2. Current values were taken from voltage ramps applied 1-3
min after application of control BS or activation by ADP. Insert: difference IV relationship
(control currents were subtracted from the values obtained after ADP activation).
Figure 4. Transient [Ca2+]cyt elevation by extracellular ADP in epidermal protoplasts. (A)
Representative trace of [Ca2+]cyt response in mature epidermal protoplasts evoked by 0.1 mM
ADP, recorded using a luminometer. “Discharge” denotes the addition of 2 M CaCl2 and 20%
(v/v) ethanol to enable [Ca2+]cyt estimation. (B) pH-dependence of the epidermal protoplast
[Ca2+]cyt response. Mean ± SE peak [Ca2+]cyt response to 0.1 mM ADP with variable extracellular
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Copyright © 2011 American Society of Plant Biologists. All rights reserved.
pH. Results are from 4 to 5 independent trials. (C) pH-dependence of the epidermal protoplast
mean ± SE peak [Ca2+]cyt response in separate trials using a plate-reader. The effects of 0.1 mM
ADP, ATP, ADPßS or αßme-ATP were tested at pH 6 or 7 (n = 4).
Figure 5. Extracellular ADP does not cause intracellular ROS accumulation. Arabidopsis
roots were ester-loaded with 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein (CMH2DCF) and exposed to extracellular ADP or ATP. (A) Top panels, roots bathed in medium (0.1
mM KCl, 0.1 mM CaCl2, pH 6.0) with 0, 0.1 mM, 1 mM extracellular ADP or 0.1 mM
extracellular ATP with fluorescence false colour-mapping applied using the same scaling used
for the positive extracellular ATP response reported by Demidchik et al. (2009). Bottom panels,
corresponding bright field images. Scale bar = 2 mm. (B) Mean ± SE fluorescence per pixel for
control (n = 23), 0.01 mM extracellular ADP (n = 24), 0.1 mM extracellular ADP (n = 25), and 1
mM extracellular ADP (n = 23) exposure. (C) In separate trials, roots were exposed to 1 mM
ADP or ATP but with Ca2+ maintained at 1 mM. Scale bar = 0.2 mm, fluorescence false colourmapping scale is shown on the right. (D) Mean ± SE fluorescence per pixel for 0.1 mM Ca2+
buffer (n = 8), 1 mM Ca2+ buffer (n = 17), 1 mM ADP with 1 mM Ca2+ (n = 11), 1 mM ATP
with 1 mM Ca2+ (n = 11).
Figure 6. Extracellular ADP-induced elevation of net Ca2+ influx and K+ efflux at the root
epidermis. (A) Response of elongation zone epidermal net Ca2+ flux to 3 µM, 100 µM or 1 mM
ADP, representative traces superimposed. Positive values signify influx. (B) Mean ± SE net Ca2+
influx at the elongation zone epidermis (n = 3-6). (C) Mean ± SE net Ca2+ influx at the mature
epidermis (n = 3-7). (D) Response of mature and elongation zone epidermis to 300 µM ADP,
32
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representative traces superimposed. Negative values signify efflux. (E) Mean ± SE net K+ efflux
at the elongation zone epidermis (n = 3-6). (F) Mean ± SE net K+ efflux at the mature epidermis
(n = 3-7). Basal assay solution comprised 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0.
Figure 7. Extracellular ATP- and αßme-ATP-induced net Ca2+fluxes at the root epidermis.
(A) Response of elongation zone epidermis to 1-1000 µM ATP, representative traces
superimposed. (B) Mean ± SE net Ca2+ influx at the elongation zone epidermis in response to
0.3-100 µM ATP (n = 6-18). (C) Mean ± SE net Ca2+ flux at the elongation zone epidermis in
response to 100-1000 µM ATP (n = 7-18). (D) Mean ± SE net Ca2+ flux at the elongation zone
epidermis in response to αßme-ATP (n = 3-7). (E) Mean ± SE net Ca2+ flux at the mature
epidermis in response to αßme-ATP (n = 3-5). Recording conditions as Figure 6.
Figure 8. Extracellular ATP- and αßme-ATP-induced net K+ fluxes at the root epidermis.
(A) Response of elongation zone epidermis to 0.3-1000 µM ATP, representative traces
superimposed. (B) Mean ± SE net K+ efflux at the elongation zone epidermis in response to 0.31000 µM ATP (n = 6-37). (C) Mean ± SE net K+ efflux at the mature epidermis in response to
10-1000 µM ATP (n = 4-13). (D) Response of elongation zone and mature epidermis to 300 µM
αßme-ATP, representative traces superimposed. (E) Mean ± SE net K+ efflux at the elongation
zone in response to 3-1000 µM αßme-ATP (n = 3-7). (F) Mean ± SE net K+ efflux at the mature
epidermis in response to 3-1000 µM αßme-ATP (n = 3-6). Recording conditions as Figure 6.
33
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Table 1. The effect of sequential extracellular ADP addition on PM current. Control values
were taken 1-3 s immediately preceding the application of ADP. “Event 1” refers to the first
application of ligand and “Event 2” to the subsequent application to the same protoplast.
Recording conditions as in Figure 1A, HP = -90 mV.
Mean ± SE maximal currents, mA.m-2
Set A
Event 1 (n = 4)
Event 2 (n = 3)
Control
0.3 mM ADP
Control
0.3 mM ADP
-40.63 ± 3.45
-76.59 ± 4.95
-36.74 ± 2.00
-56.51 ± 5.3
Set B
Event 1 (n = 3)
Event 2 (n = 3)
Control
0.3 mM ADP
0.3 mM ADP
-48.36 ± 19.83
-97.01 ± 12.90
-43.4 ± 15.23
34
0.3 mM ADP +
0.3 mM Gd3
-48.33 ± 21.30
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Figure 1
A
control
20 μM ADP
100 pA
B -200
500 ms
-150 -100
V, mV
-50
0
0
-100
-200
-300
-400
control
ADP
-500
I, pA -600
Figure 1. Extracellular ADP activates a PM influx conductance in root epidermal
protoplasts. (A) Effect of 20 µM extracellular ADP on whole cell currents from a
representative protoplast, recording conditions were as used previously by Demidchik
et al., 2009. (B) Mean ± SE IV relationships of control and peak ADP peak response (n = 4).
Control BS was as described in Materials and Methods (20 mM CaCl2, 0.1 mM KCl and Na+
equimolar to test BS, pH 6). PS comprised (mM): 40 K-gluconate, 10 KCl, 1 1,2-bis(2-aminophenoxy)ethane-N,N,N’,N’-tetraacetic acid (free Ca2+ adjustedto 100 nM with 0.4 CaCl2),
pH 7.2 adjusted with Tris/MES.
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Figure 2. Rapid activation of epidermal PM cation conductances by extracellular ADP. The
results are from whole cell recordings on protoplasts from the root mature epidermis. (A) Top
Trace: Two sequential applications of control bathing solution (BS; green bars) to a protoplast from
the mature epidermis failed to elicit a current; holding potential (HP) across the PM was -90 mV
inside negative. Middle trace (same protoplast): Two sequential applications to the same protoplast
of 0.3 mM ADP (red bars) by fast perfusion evoked inwardly-directed current (downward
deflection). Baseline current was recovered on removal of the ADP stimulus. Bottom trace
(different protoplast); inhibiting effect of 0.3 mM Gd3+, added together with ADP in the second
application. BS was as in Figure 1. Test BS included 0.3 mM NaADP ± 0.3 mM GdCl3 as indicated.
PS included 0.5 mM GDPβS. (B) Current in response to voltage ramp (40 s-long started from +100
mV) recorded from a single protoplast, approximately 1 min after the addition of control BS
(green), 0.3 mM ADP (red) or 0.3 mM ADP with 0.3 mM Gd3+ (blue). Solutions as in (A).
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Figure 3. Extracellular ADP activates Ca2+ influx and K+ efflux in epidermal protoplasts.
Mean ± SE IV relationships for control conditions and in the presence of 0.3 mM extracellular ADP,
n = 4; solutions as in Figure 2. Current values were taken from voltage ramps applied 1-3 min after
application of control BS or activation by ADP. Insert: difference IV relationship (control currents
were subtracted from the values obtained after ADP activation).
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Figure 4. Transient [Ca2+]cyt elevation by extracellular ADP in epidermal protoplasts. (A)
Representative trace of [Ca2+]cyt response in mature epidermal protoplasts evoked by 0.1 mM ADP,
recorded using a luminometer. “Discharge” denotes the addition of 2 M CaCl2 and 20% (v/v)
ethanol to enable [Ca2+]cyt estimation. (B) pH-dependence of the epidermal protoplast [Ca2+]cyt
response. Mean ± SE peak [Ca2+]cyt response to 0.1 mM ADP with variable extracellular pH.
Results are from 4 to 5 independent trials. (C) pH-dependence of the epidermal protoplast mean ±
SE peak [Ca2+]cyt response in separate trials using a plate-reader. The effects of 0.1 mM ADP, ATP,
ADPßS or αßme-ATP were tested at pH 6 or 7 (n = 4).
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A
0 mM
ADP
0.1 mM
ADP
1 mM
ADP
0.1 mM
ATP
C
Transmitted
light
+ ATP
or ADP
Buffer
1 mM
ATP
1 mM
ADP
255
20
0
1 mM ATP
40
1 mM ADP
60
1 mM Ca2+
0
80
0.1 mM Ca2+
5
1 mM ADP
10
0.1 mM ADP
15
0.01 mM ADP
20
Fluorescence per pixel
D
25
0 mM ADP
Fluorescence per pixel
B
0
Figure 5. Extracellular ADP does not cause intracellular ROS accumulation. Arabidopsis roots
were ester-loaded with 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein (CM-H2DCF) and
exposed to extracellular ADP or ATP. (A) Top panels, roots bathed in medium (0.1 mM KCl, 0.1 mM
CaCl2, pH 6.0) with 0, 0.1 mM, 1 mM extracellular ADP or 0.1 mM extracellular ATP with fluorescence
false colour-mapping applied using the same scaling used for the positive extracellular ATP
response reported by Demidchik et al. (2009). Bottom panels, corresponding bright field images.
Scale bar = 2 mm. (B) Mean ± SE fluorescence per pixel for control (n = 23), 0.01 mM extracellular
ADP (n = 24), 0.1 mM extracellular ADP (n = 25), and 1 mM extracellular ADP (n = 23) exposure.
(C) In separate trials, roots were exposed to 1 mM ADP or ATP but with Ca2+ maintained at 1 mM.
Scale bar = 0. 2 mm, fluorescence false colour-mapping scale is shown on the right. (D) Mean ± SE
fluorescence per pixel for 0.1 mM Ca2+ buffer (n = 8), 1 mM Ca2+ buffer (n = 17), 1 mM ADP with
1 mM Ca2+ (n = 11), 1 mM ATP with 1 mM Ca2+ (n = 11).
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Figure 6. Extracellular ADP-induced elevation of net Ca2+ influx and K+ efflux at the root
epidermis. (A) Response of elongation zone epidermal net Ca2+ flux to 3 µM, 100 µM or 1 mM
ADP, representative traces superimposed. Positive values signify influx. (B) Mean ± SE net Ca2+
influx at the elongation zone epidermis (n = 3-6). (C) Mean ± SE net Ca2+ influx at the mature
epidermis (n = 3-7). (D) Response of mature and elongation zone epidermis to 300 µM ADP,
representative traces superimposed. Negative values signify efflux. (E) Mean ± SE net K+ efflux at
the elongation zone epidermis (n = 3-6). (F) Mean ± SE net K+ efflux at the mature epidermis (n =
3-7). Basal assay solution comprised 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0.
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Figure 7. Extracellular ATP- and αßATP-induced net Ca2+fluxes at the root epidermis. (A)
Response of elongation zone epidermis to 1-1000 µM ATP, representative traces superimposed. (B)
Mean ± SE net Ca2+ influx at the elongation zone epidermis in response to 0.3-100 µM ATP (n = 618). (C) Mean ± SE net Ca2+ flux at the elongation zone epidermis in response to 100-1000 µM
ATP (n = 7-18). (D) Mean ± SE net Ca2+ flux at the elongation zone epidermis in response to αßmeATP (n = 3-7). (E) Mean ± SE net Ca2+ flux at the mature epidermis in response to αßATP (n = 35). Recording conditions as Figure 6.
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Figure 8. Extracellular ATP- and αßATP-induced net K+ fluxes at the root epidermis. (A)
Response of elongation zone epidermis to 0.3-1000 µM ATP, representative traces superimposed.
(B) Mean ± SE net K+ efflux at the elongation zone epidermis in response to 0.3-1000 µM ATP (n =
6-37). (C) Mean ± SE net K+ efflux at the mature epidermis in response to 10-1000 µM ATP (n = 413). (D) Response of elongation zone and mature epidermis to 300 µM αßATP, representative traces
superimposed. (E) Mean ± SE net K+ efflux at the elongation zone in response to 3-1000 µM
αßATP (n = 3-7). (F) Mean ± SE net K + efflux at the mature epidermis in response to 3-1000 µM
αßATP (n = 3-6). Recording conditions as Figure 6.
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