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Non-selective slow vacuolar (SV) channels mediate uptake of K+ and Na+ into vacuolar compartment. Under salt stress plant cells accumulate Na+ in the vacuole and release vacuolar K+ into the cytoplasm. It is, however, unclear how plants mediate transport of K+ from the vacuole without concomitant efflux of toxic Na+. Here we show by patch-clamp studies on isolated Arabidopsis thaliana cell culture vacuoles that SV channels do not mediate Na+ release from the vacuole as luminal Na+ blocks this channel. Gating of the SV channel is dependent on the K+ gradient across the vacuolar membrane. Under symmetrical K+ concentrations on both sides of the vacuolar membrane, SV channels mediate potassium uptake. When cytoplasmic K+ decreases, SV channels allow K+ release from the vacuole. In contrast to potassium, Na+ can be taken up by SV channels, but not released even in the presence of a 150-fold gradient (lumen to cytoplasm). Accumulation of Na+ in the vacuole shifts the activation potential of SV channels to more positive voltages and prevents gradient-driven efflux of K+. Similar to sodium, under physiological conditions, vacuolar Ca2+ is not released from vacuoles via SV channels. We suggest that a major Arabidopsis SV channel is equipped with a positively charged intrinsic gate located at the luminal side, which prevents release of Na+ and Ca2+, but permits efflux of K+. This property of the SV channel guarantees that K+ can shuttle across the vacuolar membrane while maintaining Na+ and Ca2+ stored in this organelle.
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Vacuoles assist the accumulation of solutes and water by plant cells. During the life cycle, plant cells face fluctuations in apoplastic and intracellular K+ concentrations, associated with nutrient supply, growth, stomatal movement and salt stress. Cytoplasmic K+ concentration in plant cells is maintained relatively constant, while vacuolar K+ pool responds to changes in growth conditions (Leigh, 2001; Walker et al., 1996). Under optimal potassium supply, K+ accumulates in plant cells at about 100 mm and K+ gradients across the vacuolar membrane remain relatively small. Under potassium starvation, however, K+ release from the vacuole provides for a K+ homeostasis in the cytoplasm and the maintenance of the membrane potential (Leigh, 2001; Leigh and Jones, 1984; Walker et al., 1996). Under salt stress plant vacuoles accumulate up to 80 mm Na+, while the vacuolar K+ concentration decreases below 30 mm (Carden et al., 2003). These observations raise a question about how vacuolar transport of K+ is regulated without concomitant leak of Na+ ?
The physiological role of SV channels is, however, still under debate. These channels were shown to activate at cytoplasmic Ca2+≥1 μm (Hedrich and Neher, 1987), that is higher than free Ca2+ concentrations in the cytoplasm of plant cells under ‘resting’ conditions (10–200 nm, see for references Gilroy and Trewavas, 2001). When Ca2+ oscillations in response to environmental and physiological stimuli take place in plant cell, cytosolic free Ca2+ can, however, rise to the micromolar range (reviewed by Evans et al., 2001; McAinsh and Hetherington, 1998; Ng and McAinsh, 2003; Trewavas, 1999). Therefore, Ca2+-activated SV channels (Hedrich and Neher, 1987) were proposed to mediate calcium-induced calcium release (CICR) in Vicia faba guard cell vacuoles (Ward and Schroeder, 1994). The proposed model predicts that ABA-induced increase in cytoplasmic Ca2+ activates Ca2+ release via SV channels and triggers stomatal closure (Allen and Sanders, 1996; Ward and Schroeder, 1994). The involvement of SV channels in CICR is, however, in conflict with several experimental data, demonstrating that under elevated cytoplasmic Ca2+ concentrations SV channels do not mediate Ca2+ release from barley mesophyll and sugar beet taproot vacuoles (Pottosin et al., 1997, 2004).
We applied the patch-clamp technique to vacuoles isolated from Arabidopsis cell culture and measured the uptake and release of K+, Ca2+ and Na+ via SV channels. Here we show that (i) SV channels mediate K+ gradient-driven bi-directional fluxes of potassium across the vacuolar membrane, and that (ii) SV channels allow Na+ and Ca2+ uptake, but not release. These findings are discussed on the basis of cation-dependent gating of the SV channel.
Arabidopsis cell culture vacuoles were released within the recording chamber upon perfusion of protoplasts with low osmolite solution. In the whole-vacuole configuration of the patch-clamp technique vacuoles were exposed to solutions, containing 10 mm EGTA and 30 mm K+ in the bath (cytoplasmic side) and 150 mm K+ in the pipette (luminal side). In the absence of Ca2+ from the cytoplasmic side and voltage range between −80 and +80 mV small voltage-independent currents were recorded (Figure 1a). To activate SV currents, vacuoles were perfused with solutions containing 0.1 mm Ca2+ (Figure 1b). In the presence of Ca2+ in the bath, slowly activating voltage-dependent currents were recorded at both positive and negative voltages (Figure 1c). When K+ concentration in the bath solution was changed from 30 to 3 mm, the activation potential of SV currents shifted from −60 to 0 mV and reversal potential shifted from −14 to −70 mV, indicating that this channel type conducts K+ (Figure 1d). According to the convention for electrical measurements on endomembranes (Bertl et al., 1992), outward vacuolar currents correspond to K+ uptake into and inward currents to K+ release from the vacuole. In cytoplasmic side-out macropatches excised from vacuoles dominated by SV currents, exposure to solutions containing 30 and 150 mm K+ at the cytoplasmic and vacuolar sides, respectively, evoked time-dependent K+ current fluctuations through open SV channels at both positive and negative voltages (Figure 1e). In smaller patches but at the same conditions we observed single channel openings in both positive and negative directions (Figure 1f), indicating that these channels account for bi-directional macroscopic K+ currents seen in the whole-vacuole configuration (Figure 1c). The open probability of single channels decreased significantly at −60 mV (Figure 1f), which corresponds to a strong reduction of whole-vacuole current at this voltage (Figure 1d, 30 mm K+). The unitary conductance of single channels in solutions, containing 30 and 150 mm K+ at cytoplasmic and luminal sides, respectively, was about 56 pS (Figure 1g).
SV channels mediate K+ uptake and release
Under symmetrical K+ concentrations (150 mm K+ on both sides of the vacuolar membrane) SV channels mediated outward currents only (Figure 2a), while at reduced cytoplasmic K+ concentrations (30 and 3 mm) SV channels conducted K+ fluxes in both directions (Figures 1c and 2b). In the nominal absence of K+ in the bath solution inward macroscopic K+ currents were recorded (Figure 2c). These data indicate that under conditions causing a drop in the cytosolic K+ pool, SV channels provide for K+ release from the vacuole and thus maintenance of cytosolic K+ homeostasis (Leigh, 2001; Leigh and Jones, 1984; Walker et al., 1996). Current-voltage plots of K+-dependent vacuolar currents demonstrate that (i) the voltage dependence of SV currents shifts to more negative voltages as cytoplasmic K+ increases and that (ii) the SV current amplitudes among the K+ concentrations tested are minimal at 3 mm K+ (Figure 2d). To analyse this behaviour in more detail, deactivation (tail) currents were plotted against voltage and fitted by a Boltzmann function (Figure 2e). Boltzmann analyses revealed similar gating charge (5.8 and 6.0) and V1/2 values (−40 and −41 mV) for 150 and 0 mm cytoplasmic K+ respectively. Lowering K+ concentration from 150 mm to 30 and 3 mm, however, caused V1/2 to shift from −40 mV to −21 and +12 mV, respectively, and gating charge to decrease from 6 to 2.0 (both 30 and 3 mm). Taking into account that Ca2+ can permeate SV channels (Ward and Schroeder, 1994) we plotted the relative conductance G/Gmax at zero voltage against the K+/Ca2+ ratio in bath solution (Figure 2f). This relation demonstrates that SV conductance goes through a minimum at 3 mm K+ and 1 mm Ca2+ at the cytoplasmic side as predicted by an anomalous mole-fraction effect (Figure 2f). An anomalous mole-fraction effect in reversal potential has been shown for V. faba guard cell SV channels by Allen and Sanders (1996) and used as evidence supporting the hypothesis that these channels conduct Ca2+. In this context it should be mentioned that high Ca2+ over K+ permeability (3:1) of Vicia guard cell vacuoles led to the prediction that SV channels mediate Ca2+-induced Ca2+ release (Allen and Sanders, 1996; Ward and Schroeder, 1994). In barley mesophyll vacuoles under conditions favouring Ca2+ release, SV channels, however, were not active (Pottosin et al., 1997).
SV channels do not mediate Ca2+ and Na+ release from vacuoles
To test whether Arabidopsis SV channels mediate vacuolar Ca2+ and Na+ uptake and release, we compared K+, Ca2+ and Na+ currents under conditions favouring either influx or efflux of these ions. In bath solutions, containing 30 mm K+, 30 mm Na+ or 15 mm Ca2+, slowly activating outward currents of comparable amplitudes were recorded, indicating that all three cations can enter the vacuole via SV channels (Figure 3a). When we applied either 100 mm Ca2+, 150 mm K+ or 150 mm Na+ to the luminal side, macroscopic inward currents were recorded in K+ solution only (Figure 3b). Ca2+ and Na+ were not released from the vacuole even when K+ was present at the cytoplasmic side (data not shown). Tail pulses to negative voltages demonstrated that deactivation of SV channels was faster in the presence of luminal Na+ and Ca2+ (τk = 108 ms, τNa = 8.7 ms, τCa = 1.25 ms, Figure 3c). To test the possibility that luminal K+ is required to activate Na+ efflux from the vacuole, we performed a series of experiments in the whole-vacuole configuration with different Na+ to K+ to ratios in the patch pipette (Figure 3d). Under all ratios tested we did not observe macroscopic inward currents, a measure for cation efflux from the vacuole. Boltzmann analyses of tail currents demonstrate that luminal K+ up to 10 mm does not significantly affect activation of SV channels (V1/2 approximately +80 mV, gating charge approximately 2). An increase in K+ to Na+ ratio (from 50K+/100Na+ to 100K+/50Na+) shifted activation potential to less positive voltages, +48 and +15 mV, respectively, without significant changes in gating charge (Figure 3d). It should be noted that under these conditions the macroscopic SV conductance, however, did not exhibit an anomalous mole-fraction effect, suggesting that the permeation pathway of SV channels is not accessible for vacuolar Na+. To test whether Na+ accumulation in the vacuole can prevent K+ release, we performed whole-vacuolar measurements with 150 mm K+ and 50 mm Na+ in the pipette in the presence of 30 mm Na+ in the bath (Figure 3e). Under these conditions we did not observe cation efflux currents as well. The activation potential of SV channel was shifted about 40 mV positive when compared with Na+-free solution. In solutions containing 150 mm Na+ at the luminal side the activation potential was even more positive (approximately +40 mV, Figure 3e), indicating that luminal Na+ shifts the voltage dependence of the SV channel to positive voltages, preventing cation efflux from the vacuole. It has been demonstrated that luminal Ca2+ and Mg2+ shift voltage-dependence of SV channels to more positive voltages too (Cerana et al., 1999; Pottosin et al., 2004). This raises the question as to how SV channels permit bi-directional K+ fluxes across the vacuolar membrane, but transport Na+ and Ca2+ into the vacuole only?
How do cations gate the SV channel?
In search for an answer to this question we concentrated on the gating of SV channels. We performed tail experiments in whole-vacuolar configuration with 150 mm K+ and 10 mm EGTA in the pipette to avoid block of SV channel by luminal Ca2+ (Pottosin et al., 2001, 2004) Voltage pulses to +80 mV followed by subsequent steps to more negative potentials were applied and channel deactivation was monitored (Figure 4a). At low negative voltages (−20 to −40 mV) SV channels deactivated only partially and therefore conducted macroscopic inward currents (Figure 4a,b). At voltages negative to −60 mV, deactivation was faster and complete, as a result SV channels did not mediate steady-state macroscopic currents under these conditions (Figure 4a,b). Similar gating behaviour was observed in solutions containing 5 mm EDTA and 5 mm EGTA at the luminal side (to remove both Ca2+ and Mg2+) as well as in the absence of Hepes/Tris buffer (data not shown). This indicates that the rectification of SV channels represents an intrinsic gating property rather than block by vacuolar cations. When tail pulses from −20 to −80 mV were applied in the whole-vacuole configuration, a transient increase in inward current was recorded before channels deactivated completely (Figure 4c, left). On the single channel level this deactivation corresponded to a decrease in the open probability of SV channels (Figure 4c right). When voltage pulses were applied in an inverse manner from −80 to −20 mV, time-dependent single channel openings were recorded (Figure 4d). Steps to more negative voltages resulted in decrease in open probability and increase in single channel amplitude (Figure 4d). Plotting the single channel amplitudes and instantaneous tail currents against voltage revealed an opposite relationship of these parameters (Figure 4e). These data provide the evidence that the SV channel is equipped with an intrinsic voltage-dependent gate, which prevents channel opening at voltages negative to −60 mV (Figure 4a–c). In the absence of luminal Na+ and voltages positive to −60 the open probability of SV channels increases (Figure 4d,e). Therefore, SV channels mediate K+ fluxes across the vacuolar membrane in both directions according to K+ electrochemical gradient (Figure 2a–c). However, the presence of Na+ at the luminal side promotes the closure of SV channels and shifts the channel activation to more positive voltages (Figure 3c,e). This gating property of the SV channel accounts for (i) Na+ storage in the vacuole and (ii) block of K+ currents at elevated luminal Na+ concentrations.
Slow vacuolar channels have been extensively characterized during the past 17 years, nevertheless, the physiological role of these transport proteins is still under discussion. SV channels were proposed to mediate K+ uptake into the vacuole (Bethke and Jones, 1994; Schulz-Lessdorf and Hedrich, 1995), CICR from the vacuole (Allen and Sanders, 1995; Bewell et al., 1999; Ward and Schroeder, 1994) and contribute to the regulation of the cell volume (de Boer, 2002). To play these roles, SV channels are supposed to operate at physiological cytosolic Ca2+ concentrations <1 μm (McAinsh and Hetherington, 1998; Trewavas, 1999) and physiological vacuolar membrane potentials, which are normally slightly negative (Gibrat et al., 1985; Rea and Sanders, 1987; Walker et al., 1995). Although in the majority of studies SV channels were activated by micromolar cytoplasmic Ca2+, millimolar Mg2+ concentrations were shown to shift the activation of SV channels to physiological (nanomolar) Ca2+ range (Carpaneto et al., 2001; Pei et al., 1999). Under conditions usually applied to patch-clamp experiments on vacuoles (symmetrical K+, millimolar luminal Ca2+) SV channels activate at positive potentials. Upon variations in K+ and Ca2+ gradients across the vacuolar membrane, activation of SV channels was observed at negative voltages (Hedrich et al., 1986; Pottosin et al., 1997). Here we studied cation release by the SV channel in more detail and demonstrate that SV channels from Arabidopsis cell culture vacuoles mediate both K+ uptake and release (Figure 2). Studies on the K+ compartmentation in barley root cells show that under low K+ supply (≤0.1 mm) the concentration of K+ in the cytoplasm is higher than in the vacuole (Walker et al., 1996), therefore K+ can passively enter the vacuole. Under optimal potassium supply (0.5–5.0 mm) the vacuolar K+ concentration is close to the cytoplasmic one, or higher, so that at negative vacuolar membrane potentials K+ can passively leave the vacuole (Walker et al., 1995, 1996). These data support the hypothesis that SV channels may play a role in regulation of K+ fluxes upon changes in K+ acquisition.
In contrast to K+, which is transported by SV channels in both directions, Na+ and Ca2+ can be taken up, but not released from the vacuole (Figure 3). Luminal Ca2+ has been recently shown to block SV channels in sugar beet vacuoles (Pottosin et al., 2004). Therefore, SV channels are unlikely to be involved in Ca2+-induced Ca2+ release (for discussion see Allen and Sanders, 1996; Carpaneto, 2003; Pei et al., 1999; Pottosin et al., 1997, 2004; Ward and Schroeder, 1994). Here we demonstrated that Na+ also blocks the SV channel by shifting its activation potential to more positive (non-physiological) voltages (Figure 3). Under salt stress Na+ accumulates in vacuoles due to the activity of Na+/H+ antiporters and H+-translocating pyrophosphatases (reviewed by Hasegawa et al., 2000; Serrano and Rodriguez-Navarro, 2001; Tester and Davenport, 2003; Zhu, 2003). High cytosolic Na+ to K+ ratios are toxic to plants, therefore Na+ has to be extruded or stored in the vacuole, while K+ should freely move across the vacuolar membrane. Therefore, the SV channel provides for a mechanism, which prevents Na+ release from the vacuole, but allows bi-directional transport of K+. High vacuolar Na+ concentrations, however, block K+ transport via the SV channel (Figure 3). This block may cause problems with osmoregulation, which plant cells face under salt stress (discussed by Tester and Davenport, 2003). Although no major differences in the properties of vacuolar channels in salt-tolerant and salt-sensitive species of Plantago as well as the extreme halophyte Suaeda maritime were observed (Maathuis and Prins, 1990; Maathuis et al., 1992), unique gating of SV channels by luminal Na+ could play an important role in ion homeostasis under salt stress.
This phenomenon can be explained by positively charged gating particle(s) located at the luminal side of the SV channel. At positive (cytoplasm) voltages the gate is open allowing K+, Na+ and Ca2+ to enter the vacuole (Figure 3a). Negative voltages move the gate into the channel pore and block the ion-conducting pathway. At voltages negative to −60 mV SV channels do not conduct macroscopic inward currents (Figures 1d and 2d). At less negative voltages the gating particle partially obstructs the pore, allowing efflux of K+, but not Na+ and Ca2+ (ions with bigger hydration shell) from the vacuole (Figure 3b–d). Moved by negative voltage towards the channel pore, luminal Na+ blocks the channel. This shifts the activation potential of the SV channel to more positive voltages (Figure 3d,e), which are required to push the blocking ion out of the pore and open the channel.
The gating mechanism of the SV channel is reminiscent of animal inward-rectifying Kir channels, which are gated by positively charged particles, blocking the ion-conducting pathway from the intracellular side of the membrane (reviewed by Nichols and Lopatin, 1997; Oliver et al., 2000). These particles are represented by polyamines, Mg2+ or C-terminal amino acid residues. The closest plant homologue to Kir channels, KCO3, containing one pore loop and two transmembrane domains, was identified in the Arabidopsis genome (reviewed by Czempinski et al., 1999; Mäser et al., 2001; Véry and Sentenac, 2002) and localized to the vacuolar membrane (D. Becker, Würzburg University, Würzburg, Germany, unpublished data). Another member of the KCO channel family, KCO1, containing two pore loops and four transmembrane domains, was shown to express in Arabidopsis mesophyll cells and localized to the vacuolar membrane as well (Schönknecht et al., 2002). However, as all KCO channels contain GYGD-related motives, they should represent K+-selective rather than non-selective channels. Therefore, the question about the molecular nature of SV channels still remains open. Given the differences in single channel conductances observed with different cell types, SV channels seem to comprise a multigene family.
A suspension cell culture from Arabidopsis thaliana ecotype Columbia-0 wildtype was grown on a solution containing 4.8 g l−1 Murashige and Skoog medium including vitamins and MES buffer (Duchefa), 20 g l−1 sucrose, 500 μg l−1 2,4-D and 28 mg l−1 Fe-EDTA. Cells were cultivated in 250-ml flasks in a shaker at 120 rpm and 21°C. The growth medium was changed twice a week.
Arabidopsis cell culture vacuoles were isolated from protoplasts, obtained by enzymatic treatment of suspension cells. The enzyme solution contained 0.8% (w/v) cellulase (Onozuka R-10), 0.1% pectolyase (Sigma-Aldrich, Taufkirchen, Germany), 0.5% bovine serum albumin, 0.5% polyvenylpyrrolidone, 1 mm CaCl2 and 8 mm Mes/Tris (pH 5.6). The osmolarity of the enzyme solution was adjusted to 300 mosmol kg−1 using d-sorbitol. Suspension cells were collected by centrifugation and incubated in enzyme solution for 1 h at 30°C and 70 rpm on a rotary shaker. Protoplasts released from cells were filtered through a 50 μm nylon mesh and washed twice in a 1 mm CaCl2 buffer (osmolarity 300 mosmol kg−1, pH 5.6). The protoplast suspension was stored on ice and aliquots were used for isolation of vacuoles. Vacuoles were osmotically isolated shortly before the measurements by perfusion of protoplasts with solution containing 10 mm EGTA, 30 mm K-gluconate and 10 mm Hepes/Tris pH 7.4, osmolarity 100 mosmol kg−1. Vacuole release was observed under the microscope.
Patch-clamp recordings were performed in the whole-vacuole and cytoplasmic side-out mode using an EPC-7 amplifier (List-Medical-Electronic, Darmstadt, Germany). Data were low-pass-filtered with an eight-pole Bessel filter (CompuMess Electronic GmbH, Garching, Germany) with a cut-off frequency of 2 kHz and sampled at 2.5 times of the filter frequency. Data were digitized by ITC-16 interface (Instrutech Corp., Elmont, NY, USA) and analysed using software Pulse and PulseFit (HEKA Elektronik, Lambrecht, Germany) and IGORPro (Wave Metrics Inc., Lake Oswego, OR, USA). Patch pipettes were prepared from Kimax-51 glass capillaries (Kimble products, Vineland, NY, USA) and coated with silicone (Sylgard 184 silicone elastomer kit; Dow Corning GmbH, Wiesbaden, Germany). External (cytoplasmic) and pipette (luminal) solute compositions are included in the figure legends. The osmolarity of solutions was adjusted to 300 mosmol kg−1 using d-sorbitol. Liquid junction potentials (see Supplementary Material) were determined as described by Neher (1992). Chemicals, unless indicated, were obtained from Sigma.
We thank A. Carpaneto, P. Dietrich, R. Roelfsema and D. Becker for the discussion and comments on the manuscript. This work was funded by DFG grants and Körber European Science Award to R.H.