Rapid stomatal closure is driven by the activation of S-type anion channels in the plasma membrane of guard cells. This response has been linked to Ca2+ signalling, but the impact of transient Ca2+ signals on S-type anion channel activity remains unknown. In this study, transient elevation of the cytosolic Ca2+ level was provoked by voltage steps in guard cells of intact Nicotiana tabacum plants. Changes in the activity of S-type anion channels were monitored using intracellular triple-barrelled micro-electrodes. In cells kept at a holding potential of −100 mV, voltage steps to −180 mV triggered elevation of the cytosolic free Ca2+ concentration. The increase in the cytosolic Ca2+ level was accompanied by activation of S-type anion channels. Guard cell anion channels were activated by Ca2+ with a half maximum concentration of 515 nm (SE = 235) and a mean saturation value of −349 pA (SE = 107) at −100 mV. Ca2+ signals could also be evoked by prolonged (100 sec) depolarization of the plasma membrane to 0 mV. Upon returning to −100 mV, a transient increase in the cytosolic Ca2+ level was observed, activating S-type channels without measurable delay. These data show that cytosolic Ca2+ elevation can activate S-type anion channels in intact guard cells through a fast signalling pathway. Furthermore, prolonged depolarization to 0 mV alters the activity of Ca2+ transport proteins, resulting in an overshoot of the cytosolic Ca2+ level after returning the membrane potential to −100 mV.
The role of cytosolic Ca2+ signals in regulation of anion channels was also challenged by studies of single guard cells in intact V. faba plants. In this species, ABA was unable to trigger cytosolic free Ca2+ signals, but nevertheless triggered a transient activation of anion channels (Roelfsema et al., 2004; Levchenko et al., 2005). Similar experiments with N. tabacum revealed that ABA triggered a transient elevation of the cytosolic Ca2+ level in approximately two of every three guard cells tested (Marten et al., 2007). For ABA responses, the degree of anion channel activation was not affected by the absence or presence of Ca2+ signals. Likewise, switching off photosynthetic active radiation also caused an increase in the cytosolic Ca2+ concentration in N. tabacum guard cells, but the activity of anion channels in response to this stimulus was enhanced in the presence of Ca2+ signals (Marten et al., 2008). This suggests that an increase in the cytosolic free Ca2+ concentration can stimulate anion channel activity in guard cells, but this response depends on the stimulus or species studied.
Even though stimulation of plasma membrane anion channels by cytosolic Ca2+ has been established in patch-clamp experiments with guard cells of V. faba, A. thaliana and N. tabacum (Schroeder and Hagiwara, 1989; Hedrich et al., 1990; Allen et al., 1999; Marten et al., 2007), detailed information on this response is still lacking. Patch-clamp experiments revealed that the Ca2+ concentration dependence of S-type anion channel activation in guard cells is modulated by ABA (Siegel et al., 2009) as well as ATP (Marten et al., 2007). However, in contrast to that of inward-rectifying K+ channels (Grabov and Blatt, 1999), the concentration dependence of plasma membrane anion channel activation has not been studied with intact guard cells. Even less is known about the time course linking elevation of the cytosolic free Ca2+ concentration to activation of anion channels in guard cells. For this reason, simultaneous measurements of the cytosolic free Ca2+ concentration and anion channel activity were performed with guard cells in intact N. tabacum plants. Cytosolic Ca2+ signals were triggered by voltage clamp pulses to hyperpolarized potentials (Grabov and Blatt, 1998; Levchenko et al., 2008). This revealed that S-type anion channels can be rapidly activated by cytosolic Ca2+, with a half maximum saturation at 515 nm cytosolic free Ca2+ (SE 235). Furthermore, we found that the ability of hyperpolarized voltage pulses to trigger elevation of the cytosolic free Ca2+ level is enhanced by prolonged depolarization.
Ca2+-dependent activation of S-type anion channels with hyperpolarizing voltage steps
In previous papers, we described an experimental approach in which guard cells are studied using multi-barrelled electrodes in intact plants (Roelfsema et al., 2001; Levchenko et al., 2005). This method not only allows recording of the free running membrane potential or plasma membrane current, but also enables current injection of the fluorescent Ca2+ reporter dye FURA2. Furthermore, the plasma membrane potential can be manipulated, which in turn can provoke changes in the cytosolic Ca2+ concentration by altering the activity of voltage-dependent Ca2+-permeable channels (Grabov and Blatt, 1998; Hamilton et al., 2000; Pei et al., 2000). The ability of hyperpolarizing voltage steps to trigger elevation of the cytosolic free Ca2+ concentration was tested using 5 sec voltage pulses from a holding potential of −100 mV (Figure 1). In all cells tested (n =11), the stimulus strength of the first test potential of −140 mV was too low to evoke an increase in the cytosolic free Ca2+ (Figure 1a–c). However, lowering the test potential to −160 mV caused a small transient increase in the cytosolic free Ca2+ concentration in six out of 11 cells. In cells already responding to −160 mV, large transient Ca2+ elevations were observed at −180 mV (Figure 1a). In three of the 11 cells, voltage-induced elevation of the cytosolic Ca2+ concentration only occurred at −180 mV (Figure 1b), and the test potential had to be lowered to −200 mV to provoke a cytosolic Ca2+ change in the remaining two cells (Figure 1c). Thus guard cells apparently differ with respect to the activation threshold required to induce an increase in the cytosolic Ca2+ concentration.
In addition to provoking elevation of the cytosolic free Ca2+ concentration, the negative test potentials induced inward currents (Figure 1a–c, lower traces). The fast transient currents at the onset and termination of the voltage pulses are due to capacity compensation of the electrode. Inward-rectifying K+ channels are likely to contribute to slowly activating currents, as these channels are activated at potentials negative of −120 mV (Roelfsema et al., 2001; Marten et al., 2007). However, after returning to the holding potential of −100 mV, K+ channels are rapidly deactivated, and the remaining inward current will be predominantly carried by plasma membrane anion channels. A transient increase in inward current was measured after termination of test potentials that provoked a Ca2+ signal. This indicates that Ca2+ was able to activate plasma membrane anion channels, which conduct an inward current.
The nature of the channels facilitating the Ca2+-induced inward current was tested using short (2 sec) voltage pulses (Figure 2a,b). S-type anion channels are characterized by a slow voltage-dependent activation and deactivation (Linder and Raschke, 1992). This characteristic is usually tested by activating S-type channels at 0 mV or more positive potentials, followed by test pulses to negative potentials (Pei et al., 1997; Roelfsema et al., 2002). Here smaller voltage steps were used in order to prevent additional voltage-induced changes in the cytoplasmic Ca2+ concentration. Short (2 sec) voltage pulses from −100 mV elicited a slowly activating inward current at −60 mV. These currents were slowly deactivated again after returning to −100 mV, a characteristic of S-type anion channels (Figure 2b). During the increase in the cytosolic free Ca2+ concentration, S-type anion currents were enhanced. This shows that elevated cytosolic Ca2+ activities stimulate S-type anion channels in intact N. tabacum guard cells (Figure 2b).
Ca2+-dependent activation of S-type anion channels with depolarizing voltage steps
The very negative potentials required to trigger a transient elevation of the cytosolic Ca2+ level raises questions about the physiological role of this response. Guard cells in intact plants seldom reach membrane potentials as negative as −160 mV (Roelfsema et al., 2001; Marten et al., 2008), and no Ca2+-dependent activation of plasma membrane anion channels was observed at more positive potentials. We therefore explored the possibility that cytosolic Ca2+ signals can be elicited by alternative voltage protocols.
Clamping the plasma membrane from −100 mV to 0 mV decreased the cytosolic free Ca2+ concentration in N. tabacum guard cells (Figures 3a and Movie S1), as previously found for V. faba (Levchenko et al., 2008). This relative small decrease was followed by a large transient elevation of the cytosolic Ca2+ level after repolarization to the holding potential of −100 mV (Figures 3a,b and Movie S1). The increase in the intracellular Ca2+ level was accompanied by a transient increase of inward current (Figure 3a), indicating activation of S-type anion channels. The ability of depolarizing voltage pulses to stimulate S-type channels was tested using short (2 sec) voltage pulses to −60 mV (Figure 4a). These pulses caused slow activation of S-type channels after stepping the membrane potential from the holding potential of −100 mV to −60 mV, and slow deactivation of these channels was associated with a return to −100 mV (Figure 4b) (Linder and Raschke, 1992). The activity of these S-type channels was enhanced during elevation of the cytosolic Ca2+ level, but returned to the pre-stimulus values after termination of the Ca2+ increase (Figure 4a,b).
The Ca2+ signal triggered by prolonged depolarization to 0 mV is only initiated after returning to −100 mV. This suggests that this response involves hyperpolarization-activated Ca2+-permeable channels, which are activated after stepping from 0 to −100 mV. The kinetics of this process were studied using voltage steps to 0 mV that varied in length. Pulses of 1 sec were too short to evoke pronounced changes in the cytosolic free Ca2+ level. During pulses of 10 sec, only small transient changes in the intracellular free Ca2+ concentration were observed (Figure 5a). However, voltage pulses to 0 mV lasting for 100 sec lowered the cytosolic free Ca2+ concentration in all cells tested (Figure 5a,b). Furthermore, a large transient overshoot of the cytosolic Ca2+ level occurred after returning to the holding potential of −100 mV in seven of eight cells (Figure 5a,b). Likewise, only the long depolarizing pulses of 100 sec were able to activate S-type anion channels (Figure 5a), which shows that activation of S-type anion channels is dependent on Ca2+ signals.
Ca2+ concentration dependence of S-type anion channel activation
The concentration dependence of the Ca2+-dependent activation of S-type anion channels was analysed by plotting S-type anion channel currents against the cytosolic free Ca2+ concentration (Figure 6). In the majority of guard cells, an increase in the cytosolic Ca2+ level was linked to the activation of S-type anion channels (Figure 6). The voltage-induced increase in the cytosolic free Ca2+ concentration exceeded 3 μm in five cells. These data were excluded from further analysis, because FURA2 does not enable accurate measurements at such high Ca2+ concentrations (asterisk in Figure 6). The other data points were fitted with a sigmoidal dose–response function, revealing a half maximal Ca2+ concentration of 515 nm (SE = 235) and a mean saturation current of −349 pA (SE = 107) at the clamp potential of −100 mV (Figure 6). The Hill coefficient of the fitted curve was 1.8 (SE = 1.2).
Nicotiana tabacum guard cells varied considerably in Ca2+ and voltage responsiveness. In 19 of 69 cells, elevation of the cytosolic free cytosolic Ca2+ concentration was not accompanied by activation of S-type anion channels (Figure 6). No response to voltage stimulation was observed in 12 of 69 cells. In none of these cells without a Ca2+ response was activation of S-type anion channels observed, confirming the hypothesis that activation of S-type channels is Ca2+-dependent (Figure 6).
Timing of Ca2+ signals
An increase of the cytosolic free Ca2+ concentration of guard cells can be induced either through hyperpolarization of the plasma membrane (Figures 1 and 2) or by prolonged depolarization (Figures 3–5). However, elevation of the cytosolic Ca2+ level directly follows the hyperpolarization, whereas depolarization initially induces lowering of the cytosolic Ca2+ concentration, followed by a transient Ca2+ increase only after returning to the holding potential. This suggests that the Ca2+ responses for both voltage protocols are induced by an influx of Ca2+ through hyperpolarization-activated cation channels.
Guard cells stimulated with 10 sec voltage steps to −180 mV showed a fast increase in the cytosolic Ca2+ level, followed by a slow relaxation to the resting level (Figure 7 and Table 1). The responses of guard cells kept for 100 sec at 0 mV were split into two groups, based on the velocity of the Ca2+ increase (Figure 8). The first group comprised cells with Ca2+ responses that were very similar to those observed with hyperpolarizing voltage steps, although the peak Ca2+ concentration was smaller (compare Figures 7 and 8a). The second group of cells showed a delayed increase in the cytosolic free Ca2+ concentration and a slow recovery to the pre-stimulus level (Figure 8b). The latter behaviour indicates that at least two mechanisms contribute to the transient cytosolic Ca2+ rise in depolarization-stimulated cells.
Table 1. Velocity of voltage-induced changes in the cytosolic free Ca2+ concentration of Nicotiana tabacum guard cells and the delay in increase of the inward current
t½ [Ca2+] increase (sec)
t½ [Ca2+] decrease (sec)
Delay in Im peak (sec)
The half times for the increase or decrease in the Ca2+ concentration were determined from traces of the cytosolic free Ca2+ concentration based on the FURA2 F345:F390 excitation ratio, as shown in Figures 2(a) and 3(a). The delay in the inward current was determined from individual measurements as shown in Figures 2(a) and 3(a). The delay is given as the time difference between the peak of the increase in cytosolic Ca2+ level and the peak in inward current. The cells were either stimulated with a 10 sec hyperpolarization from −100 mV to −180 mV or with a 100 sec depolarization from −100 mV to 0 mV. For the latter stimulus, the Ca2+ increase and concomitant increase in inward current occur after returning to −100 mV (Figure 8). The responses of cells stimulated with depolarizing voltage steps were split into two groups, with either fast or slow changes in the cytosolic free Ca2+ concentration.
Hyperpolarization (n =10)
5 ± 1
10 ± 3
9 ± 2
Depolarization (fast, n =9)
8 ± 2
18 ± 5
1 ± 3
Depolarization (slow, n =7)
29 ± 4
44 ± 9
7 ± 6
Timing of S-type anion channel activation
Activation of S-type anion channels by high cytosolic Ca2+ concentrations implies a strong correlation between the increase in the cytosolic Ca2+ concentration and activation of S-type anion channels. Indeed, the peak concentration in cytosolic Ca2+ triggered by hyperpolarizing voltage steps was followed by activation of S-type anion channels after a short delay (Figure 7 and Table 1). However, in depolarization-stimulated cells (Figure 8a,b), S-type anion channels activate without significant delay (Table 1). In addition to the Ca2+-dependent activation, the voltage steps are thus likely to affect S-type anion channels through an additional mechanism. Because of voltage-dependent gating, S-type channels are activated at 0 mV, but deactivate at −180 mV (Linder and Raschke, 1992). The short delay of 9 sec (Table 1) in hyperpolarization-stimulated cells is therefore most likely due to slow voltage-dependent activation of S-type channels at −100 mV.
Ca2+-dependent activation of S-type anion channels
Voltage-induced changes in the cytosolic free Ca2+ concentration trigger rapid activation of S-type anion channels in guard cells of N. tabacum. The lack of a measurable delay between the Ca2+ increase and S-type anion channel activation after a prolonged depolarization indicates a short signaling pathway linking both events. The dose–response curve for this response revealed a half maximal cytosolic free Ca2+ concentration of 515 nm (SE = 235). This value is in the same range as the value of 329 ± 31 nm found for inhibition of inward-rectifying K+ channels in V. faba guard cells (Schroeder and Hagiwara, 1989; Grabov and Blatt, 1999). However, Ca2+-dependent stimulation of S-type anion channels differs from inhibition of inward K+ channels with respect to the steepness of their dose–response curves. The Hill coefficient for inhibition of inward-rectifying K+ channels was 4.1 ± 0.5, compared to a value of 1.8 (SE = 1.2) for activation of S-type anion channels. This suggests that the proteins involved in activation of S-type anion channels might bind fewer than the four Ca2+ ions suggested for inhibition of inward-rectifying K+ channels (Grabov and Blatt, 1999).
S-type anion channels are unlikely to be directly regulated by Ca2+, as no Ca2+ binding motifs have been identified in the SLAC1 gene that encodes S-type anion channels in Arabidopsis guard cells (Vahisalu et al., 2008; Geiger et al., 2009). It is more likely that Ca2+-dependent protein kinases within the guard cell closely interact with the S-type anion channels. Such indirect regulation by Ca2+-sensitive proteins also explains why guard cells of N. tabacum display considerable variation in responsiveness to Ca2+ signals. In a subset of cells, S-type anion channel activity was unaffected by an increase in the cytosolic Ca2+ level. Apparently, the Ca2+ sensitivity of S-type anion channels is modulated by other regulatory mechanisms in guard cells. This type of modulation was observed for Arabidopsis guard cells (Siegel et al., 2009), in which ABA enhanced the ability of cytosolic Ca2+ to activate S-type anion channels.
Within the genome of A. thaliana, two families of Ca2+-regulated protein kinases have been identified, representing good candidates for regulating S-type anion channels. The calcium-dependent protein kinases (CDPK) belong to the CDPK–SnRK super family of serine/threonine protein kinases (Hrabak et al., 2003). Two members of this family (CPK3 and CPK6) were found to be involved in ABA-dependent regulation of S-type anion channels (Mori et al., 2006), and two further CPKs also affect stomatal movement (Zhu et al., 2007). In addition, the SnRK3/CIPK proteins (CBL-interacting protein kinases) may link Ca2+ signals to changes in transport activity, as these protein kinases interact with calcineurin B-like (CBL) proteins that in turn bind calcium (Albrecht et al., 2003; Gong et al., 2004; Hedrich and Kudla, 2006; Waadt et al., 2008). Future experiments may elucidate whether one or more members of these protein kinase families represent Ca2+-regulated interaction partners for S-type anion channels.
Voltage-induced Ca2+ signals
Voltage steps to hyperpolarized potentials represent a strong stimulus for elevation of the cytosolic free Ca2+ concentration in guard cells (Grabov and Blatt, 1998; Levchenko et al., 2008). In most guard cells of N. tabacum, stimulated from a holding potential of −100 mV, hyperpolarization to −180 mV induced a robust Ca2+ signal. However, in a subset of cells, stronger hyperpolarizing voltage steps were required to provoke elevation of the cytosolic Ca2+ level. This indicates that the voltage sensitivity of cells is subject to modulation. Indeed, cells clamped to 0 mV showed Ca2+ responses upon stepping the membrane potential to values as low as −100 mV. Apparently, the threshold potential for triggering Ca2+ elevation shifts to more positive values if cells are clamped to depolarized membrane potentials. The cytosolic free Ca2+ level is thus not elevated at a certain membrane potential, but instead upon a defined change in the membrane potential. This suggests that guard cells are especially capable of sensing fast membrane potential changes and responding with Ca2+-dependent activation of S-type anion channels.
Modulation of the voltage sensitivity required a time period exceeding 10 sec for which guard cells were clamped to 0 mV. This indicates that depolarization has a rather slow effect on one or more transporters involved in Ca2+ homeostasis. The prolonged depolarization may inhibit P-type 2B Ca2+ ATPases in the plasma and vacuolar membrane (Geisler et al., 2000; Sze et al., 2000) by lowering the cytosolic free Ca2+ concentration. These Ca2+ ATPases are regulated by an N-terminal auto-inhibitory domain that becomes active after release of Ca2+–calmodulin (Fuglsang et al., 2003). Likewise, the CAX1-encoded Ca2+ transporter is activated by CIPK24/SOS2 (salt overly sensitive) (Cheng et al., 2004), which in turn is regulated by Ca2+-binding CBL proteins (Hrabak et al., 2003; Batistic and Kudla, 2004). The ability of low cytosolic free Ca2+ levels to activate Ca2+ channels in the plasma or vacuolar membrane has not been documented. However, hyperpolarization-activated Ca2+-permeable channels in guard cells are activated by H2O2 and ABA (Grabov and Blatt, 1998; Pei et al., 2000), and this ABA response depends on protein phosphorylation in V. faba (Köhler and Blatt, 2002) and CPK3 and CPK6 in Arabidopsis (Mori et al., 2006).
Based on the considerations stated above, a model explaining our observations is presented in Figure 9. Activation of Ca2+-permeable channels in the plasma membrane through hyperpolarization is the central element in this scheme. During prolonged depolarization, low cytosolic free Ca2+ concentrations are likely to feed back on the activity of Ca2+ transporters. The subsequent inhibition of Ca2+ ATPases and CAX transporters, or stimulation of Ca2+-permeable channels, will promote the effect of voltage stimulation. In turn, high cytosolic Ca2+ levels stimulate S-type anion channels in the plasma membrane (Figure 9).
Role of Ca2+ signals in guard cell responses
The data obtained with N. tabacum guard cells support the role of cytosolic Ca2+ as a second messenger regulating S-type anion channels, as well as Ca2+ transporters, in guard cells. In this species, ABA and dark periods can trigger transient changes in the cytosolic free Ca2+ concentration (Marten et al., 2007, 2008). The mean changes in the cytosolic Ca2+ level are approximately 200 nm upon application of ABA (Marten et al., 2007) and 100 nm after switching off red light (Marten et al., 2008). Based on the relationship shown in Figure 6 and a resting cytosolic free Ca2+ concentration of 150 nm, these Ca2+ signals correlate with S-type anion currents of approximately −116 and −74 pA. However, ABA and dark stimuli triggered larger S-type anion currents of approximately −250 and −140 pA, respectively (unpublished results and Marten et al., 2008). The larger increase in S-type anion channel activity as expected from the strength of the Ca2+ signal suggests that both stimuli also act through Ca2+-independent signaling pathways.
The presence of a Ca2+-independent ABA signaling pathway is also supported by studies of V. faba guard cells. ABA was shown to inhibit inward K+ channels in guard cell protoplasts of this species in the absence of Ca2+ signals (Romano et al., 2000). Likewise, guard cells of V. faba in intact plants did not display ABA-induced changes in the cytosolic free Ca2+ concentration, but ABA nevertheless triggered activation of S-type anion channels (Levchenko et al., 2008). In N. tabacum, ABA seems to act through Ca2+-dependent as well as -independent signaling pathways to activate S-type anion channels (Marten et al., 2007). The recent identification of an intracellular ABA receptor that inhibits the ABI1 and ABI2 protein phosphatases (Ma et al., 2009; Park et al., 2009) suggests that indeed the initial steps of the ABA signaling pathway are Ca2+-independent. Downstream of the receptor, ABI1 interacts with the Ca2+-independent protein kinase SnRK2.6 (OST1, open stomata 1) (Mustilli et al., 2002; Yoshida et al., 2006), as well as two other SnRK proteins (Park et al., 2009). The activity of these SnRK protein kinases is essential for ABA responses in Arabidopsis (Fujii and Zhu, 2009; Nakashima et al., 2009) as well as those in guard cells of V. faba (Li et al., 2000). The guard cell-specific OST1 (SnRK2.6) protein kinase was found to activate SLAC1-encoded S-type anion channels in an ABI1- and ABI2-dependent manner (Geiger et al., 2009; Lee et al., 2009).
Even though Ca2+ signals do not seem to be essential for ABA activation of S-type anion channels in V. faba (Levchenko et al., 2005), these signals enhance the degree of channel activity during light–dark transitions in N. tabacum (Marten et al., 2008). In line with the latter function, we found that voltage-induced elevation of cytosolic free Ca2+ concentration evokes rapid activation of S-type anion channels. This response could be evoked through hyperpolarization of the guard cells, and was promoted by prolonged depolarized potentials. The change in voltage sensitivity suggests Ca2+-dependent feedback on Ca2+ transport proteins in guard cells. Future research will focus on the mechanisms through which Ca2+ regulates its own concentration within guard cells.
Nicotiana tabacum L. cv. SR1 plants were grown in a greenhouse under Powerstar HQI-E pressure lamps (400 W) (Philips, http://www.lighting.philips.com) with a day/night cycle of 12/12 h. All measurements were performed using the 2nd or 3rd pair of leaves of 4–6-week-old plants.
Experimental set-up for impalement, micro-electrodes and electrical configuration
Intact plants were placed next to the table of an upright microscope (Axioskop 2FS; Zeiss, http://www.zeiss.com). The adaxial side of a leaf was mounted on a acrylicglass holder in the focal plane of the microscope using double-sided adhesive tape and medical adhesive (Medical Adhesive B, Aromando, http://www.amt-med.de). In this experimental configuration, guard cells of the abaxial epidermis were accessible for impalement with triple-barrelled micro-electrodes. Micro-electrode impalement of guard cells was performed using a piezo-driven micro-manipulator (MM3A; Kleindiek Nanotechnik, http://www.nanotechnik.com). Positioning of the electrode was monitored using a water immersion objective (Achroplan 40×/0.8 W, Zeiss), operated with a drop of bath solution (5 mm KCl, 0.1 mm CaCl2, 5 mm potassium citrate, pH 6) between the objective and the leaf surface. The triple-barrelled micro-electrodes were pulled from borosilicate glass capillaries (inner diameter 0.58 mm, outer diameter 1.0 mm; Hilgenberg, http://www.hilgenberg-gmbh.com), as described previously (Roelfsema et al., 2001). Three capillaries were aligned, heated and twisted 360°, and pre-pulled using a customized vertical electrode puller (L/M-3P-A; Heka, http://www.heka.com). The final pull was executed using a horizontal laser puller (P 2000, Sutter Instruments Co., http://www.sutter.com). Two barrels of the micro-electrodes were filled with 300 mm CsCl, and the 3rd barrel was filled with 2 mm of the FURA2 Ca2+ reporter dye (Fluka/Sigma-Aldrich, http://www.sigmaaldrich.com). The tip resistance of each barrel ranged from 90 to 200 MΩ. The reference electrode consisted of a 300 mm KCl, 2% agarose salt bridge, connected to an Ag/AgCl half cell, and was placed in the bath solution between the objective and the cuticle. All barrels of the electrode were connected via Ag/AgCl half cells to head stages (HS180; BioLogic, http://www.bio-logic.info) with an input resistance of approximately 1011Ω. Head stages were coupled to micro-electrode amplifiers (VF-102; BioLogic) and the membrane potential was clamped using a differential amplifier (CA-100; BioLogic). Voltage steps were controlled using Pulse software (Heka) with an LIH-1600 interface (Heka). Data were low-pass-filtered at 10 Hz using an eight-pole Bessel filter (type 902; Frequency Devices, http://www.frequencydevices.com), and sampled at 33 Hz.
Ratiometric fluorescence microscopy
The ratiometric fluorescent calcium indicator dye FURA2 was micro-injected iontophoretically into guard cells using an injection current of −300 pA. During FURA2 injection, the membrane potential was constantly clamped to −100 mV, and thus the injection current from the 3rd barrel was automatically compensated for by current from the 2nd barrel. Ratiometric fluorescence spectroscopy measurements were performed using Metafluor software (Universal Imaging, http://www.moleculardevices.com). FURA2 was excited using 100 msec flashes of UV light at 345 and 390 nm and a time interval of 1 sec (VisiChrome high-speed polychromator system, Visitron Systems, http://www.visitron.de). The fluorescence emission signal was passed through a beam splitter (FT 395; Zeiss), filtered using a 510 nm bandpass filter (D510/40 M, AHF Analysentechnik, http://www.ahf.de), and captured using a cooled charge-coupled device camera (CoolSNAP HQ, Roper Scientific, http://www.roperscientific.com). Background fluorescence levels for both wavelengths were taken from a reference region placed within part of a neighbouring epidermal cell. The intracellular free Ca2+ concentration was calculated from the FURA2 ratio (Grynkiewicz et al., 1985) using the equation:
where Kd represents the binding constant of FURA2 for Ca2+, R represents the 345:390 nm excitation ratio, Rmin and Rmax correspond to Ca2+-free and Ca2+-saturated FURA2, respectively, and Fmin and Fmax indicate the fluorescence intensity measured at 390 nm with Ca2+-free and Ca2+-saturated FURA2, respectively. Here we used a Kd of 270 nm, determined in vitro by Levchenko et al. (2008). Rmin and Fmin were defined as the values obtained after simultaneously injecting 2 mm FURA2 and 50 mm BAPTA (1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid) at 0 mV into guard cells of intact plants. The values for Rmax and Fmax were obtained by clamping the plasma membrane to −250 mV at the end of each experiment, inducing a massive and saturating Ca2+ influx. In cells with large Ca2+ increases, R approaches Rmax, leading to unrealistically high values for the free Ca2+ concentration. In these cells, the maximum Ca2+ concentration was arbitrarily set to 3 μm (approximately 10 times the Kd value).
The dose–response curve of Ca2+-dependent S-type anion channel activation was fitted using Sigmaplot 2000 software (SPPS Inc, http://www.spss.com) to the following three-parameter Hill function:
where [Ca2+cyt] is the cytosolic free Ca2+ concentration, [Ca2+cyt]EC50 is the cytosolic free Ca2+ concentration eliciting a half maximal response, ΔIm is the increase in S-type current, ΔIm,max is the increase in S-type anion current at saturating Ca2+ levels, and n is the Hill coefficient.
This work was supported by grants from the Deutsche Forschungsgemeinschaft to R.H. and M.R.G.R (GRK 1342 and SFB 567).