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Novel roles of ascorbate in plants: induction of cytosolic Ca2+ signals and efflux from cells via anion channels

Novel roles of ascorbate in plants: induction of cytosolic Ca2+ signals and efflux from cells via... Abstract Ascorbate is not often considered as a signalling molecule in plants. This study demonstrates that, in Arabidopsis roots, exogenous l-ascorbic acid triggers a transient increase of the cytosolic free calcium activity ([Ca2+]cyt.) that is central to plant signalling. Exogenous copper and iron stimulate the ascorbate-induced [Ca2+]cyt. elevation, while cation channel blockers, free radical scavengers, low extracellular [Ca2+], transition metal chelators, and removal of the cell wall inhibit this reaction. These data show that apoplastic redox-active transition metals are involved in the ascorbate-induced [Ca2+]cyt. elevation. Exogenous ascorbate also induces a moderate increase in programmed cell death symptoms in intact roots, but it does not activate Ca2+ influx currents in patch-clamped root protoplasts. Intriguingly, the replacement of gluconate with ascorbate in the patch-clamp pipette reveals a large ascorbate efflux current, which shows sensitivity to the anion channel blocker, anthracene-9-carboxylic acid (A9C), indicative of the ascorbate release via anion channels. EPR spectroscopy measurements demonstrate that salinity (NaCl) triggers the accumulation of root apoplastic ascorbyl radicals in an A9C-dependent manner, confirming that l-ascorbate leaks through anion channels under depolarization. This mechanism may underlie ascorbate release, signalling phenomena, apoplastic redox reactions, iron acquisition, and control the ionic and electrical equilibrium (together with K+ efflux via GORK channels). Anion channels, Arabidopsis, ascorbate transport, ascorbic acid, ascorbyl radicals, calcium signalling, hydroxyl radicals, plasma membrane, reactive oxygen species, salinity Introduction l-Ascorbic acid (AA) is a key antioxidant, cofactor of redox enzymes, and precursor for some biosyntheses. In water solutions, this substance is present as a monovalent anion, ascorbate– (Asc–), which easily donates electrons to oxidized molecules and scavenges reactive oxygen species (ROS). For example, AA directly reacts with the most important ROS, such as superoxide (O2·–), hydroxyl radicals (HO·), and singlet oxygen (Zhang, 2013; Halliwell and Gutteridge, 2015). Moreover, AA is a major reducing agent for iron and copper [Cu2+/Fe3++electron from AA→Cu+/Fe2++dehydroascorbate (DHA)] and potentially for other transition metal ions, such as Mn2+/3+, inside and, hypothetically, outside the cell. This makes AA a pro-oxidant because Cu+ and Fe2+ are catalysts of HO· production, while HO· is the most powerful oxidizing species in plants (Demidchik, 2015; Halliwell and Gutteridge, 2015; Demidchik and Shabala, 2018). The pro-oxidative properties of AA in plants have not been studied in detail, while its role as an antioxidant has attracted enormous interest over the last few decades (Foyer and Noctor, 2011; Halliwell and Gutteridge, 2015; Wertz et al., 2017). In plant cells, the antioxidant function of AA is related to a number of processes, but the most important one is the donation of electrons in the Foyer–Halliwell–Asada cycle (FHA cycle), also known as the ‘ascorbate–gluthatione cycle’ (Foyer and Halliwell, 1976, 1977; Foyer and Noctor, 2011). The FHA cycle is active inside the cell, detoxifying superoxide anion radicals (O2·–), which are mainly generated as a result of so-called ‘electron leakage’ to triplet oxygen (O2) in photosynthetic, mitochondrial, peroxisomal, and probably other electron transport chains (Ozyigit et al., 2016). AA also takes part in the regeneration of α-tocopherol (vitamin E) in photosynthetic membranes and participates in chloroplast metabolism as a cofactor for violaxanthin de-epoxidase, an enzyme involved in xanthophyll cycle-mediated photoprotection (Smirnoff and Wheeler, 2000). Overall, these are very important physiological roles of AA relating this molecule exclusively to antioxidant functions. Nevertheless, this might not be the only function of AA in plants. In some plant species, AA is the substrate for the biosynthesis of oxalate and tartrate (Smirnoff and Wheeler, 2000). AA also controls plant development via regulation of phytohormone synthesis (Pastori et al., 2003). An exogenous AA is capable of stimulating an expansive growth, and the high ascorbate oxidase activity in the cell wall is correlated with areas of rapid cell expansion (Arrigoni, 1994; Gallie, 2013). The mechanistic explanation of these phenomena is largely missing. Hypothetically, AA can act as a cofactor for prolyl hydroxylase that hydroxylates proline residues in the cell wall hydroxyproline-rich glycoproteins required for cell expansion (Smirnoff and Wheeler, 2000). Another possibility is that AA stimulates HO· production via reduction of cell wall-bound transition metals, such as copper and iron (Demidchik, 2015). This can potentially result in the activation of Ca2+ influx, induction of the ROS–Ca2+ hub (via NADPH oxidase) required for sustained ROS production and Ca2+ influx stimulating exocytosis and cell elongation (Foreman et al., 2003; Demidchik and Shabala, 2018), and/or cleavage of cell wall polymers inducing cell wall loosening (Müller et al., 2009). Plants synthesize AA by at least by two pathways. The first one is an intracellular Smirnoff–Wheeler pathway (cytosol/mitochondria), when plants produce AA from l-galactose, which in turn is synthesized from GDP-d-mannose (Wheeler et al., 1998). A second AA biosynthetic pathway is located in the cell wall and relies on the substrate d-galacturonic acid, generated from the breakdown of pectin (Loewus and Kelly, 1961; Gallie, 2013). It is believed that the first pathway is the more important and capable of producing the most AA in plants (Smirnoff and Wheeler, 2000; Gallie, 2013). The physiological role of the second pathway of AA production is poorly understood. The levels of apoplastic AA and its reversibly oxidized forms (Asc·– and DHA) can control the activities of apoplastic transition metals, and, vice versa, metals, such as copper and iron, can affect AA stability and AA-mediated reactions in the cell wall. Transition metals such as copper and iron are abundant in the cell wall (Thornton and Macklon, 1989; Zhang et al., 1991; Fry et al., 2002). They can catalyse HO· generation from O2·–/H2O2 in the presence of AA in this compartment (Fry, 1998; Demidchik et al., 2010; Demidchik, 2015). The induction of Ca2+ influx for signalling purposes via AA-mediated HO· generation in the apoplast can also be proposed. In this context, a new signalling role for apoplastic AA is emerging. This possibility has not been addressed in higher plants. It can be hypothesized that apoplastic AA functions as a signalling molecule in plants. This can be due to redox properties of this molecule or potentially via some receptor-like mechanisms. In this context, the testing of potential elevations in cytosolic free Ca2+ ([Ca2+]cyt.) using Ca2+ aequorin luminometry is crucial; it can directly demonstrate AA involvement in central reactions of plant signalling (via generation of Ca2+ signals). If AA is a signalling molecule, it can have a system for release from plant cells and breakdown in the extracellular space. The breakdown can be catalysed by ascorbate peroxidase (De Gara, 2004) and ascorbate oxidase (De Tullio et al., 2013), while its release can hypothetically be mediated by anion channels. The negative electric potential difference across the plasma membrane of typical plant cell is –120 mV to –60 mV (Demidchik et al., 2002). This provides a driving force for passive ascorbate efflux from cells, if the plasma membrane is permeable to this anion. In order to answer this question, the plasma membrane conductance mediated by the AA efflux should be measured. The permeability of plant plasma membranes to Asc– has not previously been examined by electrophysiological techniques, although this question is definitely of fundamental importance for plant biology. Endogenous AA represents as much as 10% of soluble carbohydrates in some plant tissues (Noctor and Foyer, 1998); therefore, a tight control over transport of AA is crucial for the organism. In animal cells, anion channels dominate ascorbate efflux, although other mechanisms are also involved (Corti et al., 2010). Thus, it can be hypothesized that anion channels may also be involved in plant AA efflux. The aim of this study was to examine AA effects on Ca2+ signalling and explore the potential role of transition metals, Cu+/2+ and Fe2+/3+, in the development of these effects. The AA efflux currents were measured by conventional patch-clamp techniques using protoplasts isolated from roots. The ascorbate release was assessed by examining apoplastic ascorbyl radicals using EPR spectroscopy. The model plant, Arabidopsis thaliana, was used as a convenient system for electrophysiological, cell death, luminometry, and EPR spectroscopy assays. Materials and methods Plant material Wild-type (WT) A. thaliana, ecotype Columbia-0 (Col-0), was used to study programmed cell death (PCD) symptoms (microscopy), ionic currents (patch-clamp), and formation of Asc·– (EPR spectroscopy). Transgenic Col-0 were used to measure changes in [Ca2+]cyt. by aequorin luminometry. Col-0 seeds were obtained from the European Arabidopsis Stock Centre (arabidopsis.info). For all experiments, the plants were grown vertically on a surface of sterilized gel (in Petri dishes), which contained full-strength Murashige and Skoog (MS) medium (Murashige and Skoog, 1962), 1% sucrose (w/v), and 0.35% Phytagel (w/v, Sigma-Aldrich), pH 6.0/KOH. Constant temperature and irradiance photosynthetically active radiation (PAR) conditions were maintained in growth cabinets (22 ± 1 °C, 16 h daylight, LED lamps, with PAR wavelength, 200 μmol m−2 s−1 irradiance). For patch-clamp and luminometry measurements, root protoplasts were isolated using techniques developed by Demidchik and Tester (2002) and Demidchik et al. (2002, 2010). Briefly, roots were excised from 50–100 A. thaliana seedlings (7–12 d old) and cut into 0.5–2 mm long pieces using a razor blade in a plastic dish, which contained 4 ml of enzymatic solution comprising 1.5% (w/v) Cellulase Onozuka RS (Yakult Honsha, Tokyo, Japan), 1% (w/v) cellulysin (CalBiochem, Nottingham, UK), 0.1% (w/v) pectolyase Y-23 (Yakult Honsha), 0.1% (w/v) BSA (Sigma), 10 mM KCl, 10 mM CaCl2, 2 mM MgCl2, 2 mM MES, pH 6.0 with Tris; 290–300 mOsM adjusted with 330 mM sorbitol. After gentle shaking (45 rpm) in the enzyme solution for 30–50 min at 28 °C, protoplasts were filtered (30 µm pore mesh) and rinsed with holding solution (HS; 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM sucrose, 10 mM glucose, 2 mM MES, pH 6.0 with Tris; 290–300 mOsM with sorbitol). Protoplasts were collected by 5 min centrifugation at 200 g and ~0.5 ml was diluted with 8–10 ml of HS. Luminometric measurements of [Ca2+]cyt. Seven- to 12-day-old seedlings of A. thaliana plants constitutively expressing apoaequorin, under the control of the 35S Cauliflower mosaic virus (CaMV) promoter, were used (Knight et al., 1996, 1997; Polisensky and Braam, 1996). Seeds were kindly provided by Professor Marc Knight (Durham University). Luminometric measurements were carried out directly on these seedlings or on protoplasts. Ca2+ activity was recorded continuously up to 15 min with a single-tube Turner BioSystems Luminometer Model TD-20/20 controlled by Spreadsheet Interface Software (Promega). Standard aequorin chemiluminometry techniques were used to examine changes in [Ca2+]cyt. (Knight et al., 1996, 1997; Demidchik et al., 2011; Sosan et al., 2016). Coelenterazine, native (4 µM; Prolume Ltd, USA) was used in all measurements. The remaining aequorin was discharged for 5 min using a mixture containing 2 M CaCl2, 20% ethanol (v/v), and 0.01% Triton X-100 (v/v). The amount of free Ca2+ entering the cells was calculated using the calibration equation derived empirically: pCa=0.332588(−log k)+5.5593, where k=(luminescence counts s−1)/(total luminescence counts remaining) (Knight et al., 1996, 1997). Curves showing dose dependence of [Ca2+]cyt. in response to AA were fitted and analysed using SigmaPlot 10.0 (Systat Software Inc.). Patch-clamp measurements Conventional patch-clamp and protoplast isolation techniques were used (Demidchik et al., 2010). The standard bathing solution contained (in mM): 20 CaCl2, 2 Tris, 0.1 NaCl, adjusted to pH 6.0 with MES, and 290–300 mOsM, with sorbitol. A freshly prepared mixture of this solution with AA was applied in whole-cell outside-out patches. All pipette solutions (PSs) contained at least 10 mM Cl– and 100 nM Ca2+ adjusted with 2 mM Tris/MES, pH 7.2. Other saline solutions are indicated in the figure legends. Liquid junction potentials were calculated by JPCalc which is included in the Axon Clampex 10.6 software (Molecular Devices, USA) and corrected. The voltage was held at –90 mV, then square 7.6 seconds-long depolarizing or hyperpolarizing voltage pulses were applied. Currents were measured using PC-ONE Patch/Whole Cell Clamp (CORNERSTONE Series) amplifier (Dagan Corporation, USA) controlled by Digidata 1320/Clampex 10.6 (Molecular Devices, USA). Imaging symptoms of programmed cell death Standard bright-field and epi-fluorescent microscopy were used for the analysis of morphological (irreversible plasmolysis, cytoplasm shrinkage, plasma membrane rupture, appearance of dark bodies, etc.) and biochemical (caspase-like proteolytic activity; CaspACE FITC-VAD-fmk in situ marker kit; Promega, UK) cell death symptoms in root cells as described by Demidchik et al. (2010) and Hogg et al., (2011). An inverted Nikon TS100F microscope (Nikon, Japan) powered by NIS-Elements AR software and a standard fluorescein isothiocyanate (FITC) filter were used. The exposure time was 0.5 s. The bathing solution contained (in mM): 0.1 KCl, 0.1 CaCl2, pH 6.0 (4 MES/2 Tris). Five minutes staining with 0.5% Evans blue (Roth; w/v) was applied to AA-treated roots; then the dye was washed five times. To analyse protease activity, 10 μM FITC-VAD-fmk was applied together with the AA. After treatment with AA, the roots were washed three times for 5 min and observed under the microscope. Detection of Asc·– by EPR spectroscopy Standard EPR spectroscopy in liquid phase (20 °C) was used for the examination of Asc·– in intact roots, in control conditions, and after treatment by 100 mM NaCl (Liszkay et al., 2004; Demidchik et al., 2010). The cell-free tests were also carried out as required. Preliminary cell-free tests and analysis of the literature (Halliwell and Gutteridge, 2015) showed that the ascorbate radical signal is stable at pH 8.0; therefore, this pH level was used throughout the EPR tests. The basic solution for EPR spectroscopy tests included 2 mM Tris/MES (pH 8.0), 0.1 mM CaCl2, 0.1 mM KCl. For all tests, reagents were mixed 60 s before recording the EPR spectrum in oxygen-depleted conditions. ESR Grade Water (Noxygen) was used, and all solutions were additionally purified using ultrafine activated charcoal powder (Sigma-Aldrich) to remove any residues of transition metals. The EPR spectra were recorded on a Bruker EMX (X-band) spectrometer equipped with an ER 4103TM cylindrical mode resonator and analysed using WinEPR (Bruker). For in vitro tests, the EPR Spectrometer SPINSCAN (ADANI) was used with a standard cuvette set. Asc·– spectra were detected without the spin-trap as described by Buettner and Jurkiewicz (1993). In cell-free conditions, AA was directly mixed with buffer solution containing the substances being tested. To measure Asc·– generation in intact roots, 150 intact 1-week-old seedlings were removed from the medium, carefully washed several times, and adapted for 1 h in a 1 ml vial filled with buffer solution containing 0.1 mm KCl and 0.1 mM CaCl2 (pH 8.0; Tris/MES), then 2 cm lengths of the root tips were immersed for 5 min in the fresh buffer solution. After treatment, the eluate from the vial was directly collected in a Bruker AquaX system (the multiple bore design of closely spaced capillaries allows for much higher sensitivity compared with the capillary or flat-cell designs) for measurements of liquid samples. Results Exogenous l-ascorbate induces free radical- and transition metal-dependent elevation of cytosolic free Ca2+ To assess whether AA can trigger [Ca2+]cyt. elevations, 0.01–30 mM AA was added to excised roots of intact A. thaliana plants constitutively expressing aequorin (Fig. 1). Starting from 100 µM, AA induced a significant transient increase in [Ca2+]cyt., with a peak during 4–7 min (ANOVA test, P<0.001; Fig. 1A). The addition of buffer solution without AA or application of low AA concentrations (0.1–10 µM) did not cause significant changes in the cytosolic Ca2+ (Fig. 1B). Fig. 1. View largeDownload slide The effect of exogenously applied l-ascorbic acid (AA) on the activity of cytosolic Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 10 µM to 30 mM AA (I.–VII.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA are indicated in the panels. (IX.) [Ca2+]cyt. elevation in response to combined application of 1 mM Cu2+ and 1 µM AA. ‘DC’: time, when the discharge solution was added to roots (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different AA concentrations (mean ±SE; n=10–11). (C) Dependence of peak [Ca2+]cyt. on the concentration of Cu2+ applied with (open circles) or without (filled circles) 1 mM AA (mean ±SE; n=5–10). (D) Effect of iron ions applied as Fe(II)EDTA on peak AA-induced [Ca2+]cyt. transients (mean ±SE; n=4). The bathing solution in all tests contained 10 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). Fig. 1. View largeDownload slide The effect of exogenously applied l-ascorbic acid (AA) on the activity of cytosolic Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 10 µM to 30 mM AA (I.–VII.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA are indicated in the panels. (IX.) [Ca2+]cyt. elevation in response to combined application of 1 mM Cu2+ and 1 µM AA. ‘DC’: time, when the discharge solution was added to roots (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different AA concentrations (mean ±SE; n=10–11). (C) Dependence of peak [Ca2+]cyt. on the concentration of Cu2+ applied with (open circles) or without (filled circles) 1 mM AA (mean ±SE; n=5–10). (D) Effect of iron ions applied as Fe(II)EDTA on peak AA-induced [Ca2+]cyt. transients (mean ±SE; n=4). The bathing solution in all tests contained 10 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). The basal [Ca2+]cyt. level was 70.8 ± 3.8 nM (±SE; n=15). Peak values were calculated by subtracting this level (tested in each individual trial) from the obtained maximal (peak) increase of [Ca2+]cyt.. The relationship between the mean peak [Ca2+]cyt. increase and the concentration of applied AA is demonstrated in Fig. 1B. This ‘dose–response curve’ did not show saturation even at very high levels of AA in the bathing solution. The maximal [Ca2+]cyt. elevation was induced by 30 mM AA (321 ± 40.1 nM; ±SE; n=10). At the same time, physiological levels of AA, which in roots reach 5–10 mM (Smirnoff and Wheeler, 2000; Kawa et al., 2016; Alscher and Hess, 2017), triggered effects that were 2–5 times smaller (Fig. 1). Hypothetically, exogenous AA can undergo transition metal-catalysed oxidation leading to formation of HO·, activation of Ca2+-permeable cation channels, and elevation of [Ca2+]cyt. (Demidchik et al., 2003). Copper ions are the most powerful catalysts of Fenton-like reactions (generating HO·) compared with iron and other transition metals (Fry, 1998; Fry et al., 2002; Halliwell and Gutteridge, 2015). Addition of Cu2+ together with AA resulted in [Ca2+]cyt. elevation, which demonstrated parameters similar to AA-induced [Ca2+]cyt. but had much higher peak values (Fig. 1). Very low Cu2+ concentrations (10–7 M and 10–6 M) caused significant stimulation of AA-induced [Ca2+]cyt. transients. This is aligned with plant cell catalytic activities of transition metals (Cu2+ and Fe3+), which can reach 0.3–15 µM (Becana and Klucas, 1992). The minimal Cu2+ concentration which caused statistically significant stimulation of the AA effect was 0.1 µM (n=7; P<0.01; ANOVA; Fig. 1C). Iron [Fe(II)-EDTA] also stimulated AA-induced [Ca2+]cyt. elevation, but it was less effective than copper, causing significant stimulation at millimolar levels (Fig. 1D). To examine the involvement of free radicals, such as HO·, in the AA-induced [Ca2+]cyt. increase, AA was added together with a free radical scavenger, thiourea (Fig. 2). Thiourea effects on HO· generation and HO·-dependent processes have previously been observed in different preparations, including EPR studies on A. thaliana roots (Demidchik et al., 2003, 2010; Liszkay et al., 2004; Halliwell and Gutteridge, 2015). Addition of 3 mM thiourea to aequorin-expressing roots without AA caused a small increase in [Ca2+]cyt. (peak: 15 ± 6.2 nM; mean ±SE; n=6). When 3 mM thiourea was added together with 1 mM or 3 mM AA, the elevation of [Ca2+]cyt caused by AA decreased dramatically (up to eight times; Fig. 2). This test with thiourea, which is considered by some authors as a hydroxyl radical-specific scavenger (Liszkay et al., 2004), strongly suggests that AA acts on cytosolic free Ca2+ via a free radical-dependent pathway, such as generation of HO·, causing activation of Ca2+-permeable cation channels (Demidchik, 2015). Similar results (inhibition of AA-induced [Ca2+]cyt. transients; n=5; data not shown) were obtained here for DMSO (0.03%, v/v), which is a non-specific scavenger of free radicals (Demidchik et al., 2010; Halliwell and Gutteridge, 2015). Fig. 2. View largeDownload slide Inhibitory action of thiourea (free radical scavenger) on the l-ascorbate-induced elevation of the cytosolic free Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 1 mM and 3 mM AA (I. and III.) and combined application of these concentrations with 3 mM thiourea (II. and IV.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA and thiourea are indicated in the panels. DС: time of applying the discharge solution (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different treatments by AA and combined application of AA and thiourea (mean ±SE; n=6–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a) and (b)). The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Fig. 2. View largeDownload slide Inhibitory action of thiourea (free radical scavenger) on the l-ascorbate-induced elevation of the cytosolic free Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 1 mM and 3 mM AA (I. and III.) and combined application of these concentrations with 3 mM thiourea (II. and IV.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA and thiourea are indicated in the panels. DС: time of applying the discharge solution (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different treatments by AA and combined application of AA and thiourea (mean ±SE; n=6–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a) and (b)). The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Two classical blockers of plasma membrane Ca2+ influx channels, Gd3+ and La3+ (Demidchik et al., 2002), were used to define the involvement of cation channels in AA-induced [Ca2+]cyt. elevation (Fig. 3). The results showed that Gd3+ and La3+ were very effective inhibitors of cytosolic calcium burst induced by both 1 mM and 3 mM AA. This confirms that the effect on cytosolic Ca2+ was related to the activation of Ca2+-permeable cation channels. Moreover, the decrease of extracellular Ca2+ from 10 mM to 50 µM caused a decrease of peak [Ca2+]cyt. elevations induced by 1 mM AA from 44.4 ± 7.2 nM (mean ±SE; n=9) to 9.5 ± 3.6 nM; mean ±SE; n=6). Fig. 3. View largeDownload slide Peak transient [Ca2+]cyt. increase induced by 3 mM AA in the presence of lanthanide cations (La3+ and Gd3+), transition metal chelators (bathocuproine and deferoxamine), or after removal of the cell wall by isolation of protoplasts [mean ±SE; n=7–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a)]. Protoplast isolation techniques were the same as in patch-clamp analyses (see the Materials and methods). Concentrations of salines are indicated in the figure. The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Fig. 3. View largeDownload slide Peak transient [Ca2+]cyt. increase induced by 3 mM AA in the presence of lanthanide cations (La3+ and Gd3+), transition metal chelators (bathocuproine and deferoxamine), or after removal of the cell wall by isolation of protoplasts [mean ±SE; n=7–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a)]. Protoplast isolation techniques were the same as in patch-clamp analyses (see the Materials and methods). Concentrations of salines are indicated in the figure. The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Based on data presented in Fig. 2, the AA-induced [Ca2+]cyt. elevation is related to HO· generation in the cell wall, which can be catalysed by transition metals (copper and iron) residing in the cell wall. To examine this hypothesis, plants were pre-treated by copper (0.3 mM bathocuproine) and iron (0.3 mM defereoxamine) chelators (Fig. 3). This reduced the AA-induced [Ca2+]cyt. elevations by 40–60%. Combined application of AA with both 0.3 mM bathocuproine and 0.3 mM defereoxamine resulted in complete inhibition of AA-induced [Ca2+]cyt transients (Fig. 3), demonstrating that copper and iron (and HO· generation mediated by these metals) are central to the AA-induced Ca2+ signalling. Apart from the application of chelators, cell wall metals were depleted by the removal of the cell wall using enzymatic treatment (protoplast were isolated by the same protocols as were used for patch-clamping; Fig. 3). The protoplasts lacked AA-induced [Ca2+]cyt elevation. This additionally confirmed that the cell wall was crucial for AA effects. Cell death symptoms in roots treated by l-ascorbic acid PCD induced by a mixture of AA and Cu2+ and stressors associated with oxidative burst (NaCl, pathogens, heavy metals, etc.) has recently been studied in detail (Demidchik et al., 2010, 2017; Petrov et al., 2015). If AA added to the cell wall is capable of producing HO· and stimulating Ca2+ influx, then the induction of PCD in the AA-treated roots is a possible scenario. Stresses can induce a number of detectable morphological symptoms of PCD, such as irreversible plasmolysis, cytoplasm shrinkage, plasma membrane rupture, and appearance of dark spots at the site of the nucleus (Demidchik et al., 2010, 2017; Hogg et al., 2011). Plasma membrane damage can be monitored by staining with membrane-impermeant dyes, such as Evans blue. Hydrolytic ‘self-digestion’ reactions accompanying PCD can be assessed by staining the cell death proteases using the CaspACE FITC-VAD-fmk in situ marker kit (Promega, UK; Demidchik et al., 2010). Fluorescently labelled protease inhibitor zVAD-fmk (FITC-VAD-fmk) is a widely used tool for the observation of PCD protease activation in intact tissues, because, at micromolar concentrations, this substance does not significantly inhibit proteases, although it binds to and fluorescently labels them (Elbaz et al., 2002; Bonneau et al., 2008). In this case, an increased intracellular fluorescence indicates a higher protease activity. For AA-treated roots, morphology, Evans blue, and FITC-VAD-fmk tests demonstrated that long-term treatment with AA (15 h and 40 h) triggered a statistically significant increase in the number of cells with PCD symptoms (Fig. 4). Trichoblasts (root hairs) were more sensitive to both 15 h and 40 h treatment by AA than mature epidermal cells (atrichoblasts). The addition of 0.3 mM bathocuproine and 0.3 mM defereoxamine together with AA prevented the development of morphological PCD symptoms in all epidermal cell types. Treatment by 1 mM AA caused an increase of FITC-VAD-fmk fluorescence by 30–40%, which was abolished by the combined addition of 0.3 mM bathocuproine and 0.3 mM defereoxamine (Fig. 4C). Overall, these tests showed that physiological levels of exogenous AA induced a very moderate increase in the level of PCD symptoms in roots. AA was mainly targeting root hairs, where the cell wall is thinner, and metabolic activities, including those that are catalysed by copper and iron, are higher. Fig. 4. View largeDownload slide Symptoms of programmed cell death (PCD) in Arabidopsis thaliana L. roots treated with l-ascorbic acid. (A) Changes in Arabidopsis root cell morphology (‘Morphological symptoms’) and activity of cell death proteases (‘FITC-VAD-fmk’) induced by 1 mM l-ascorbic acid. Arrows show areas with condensed bodies within irreversibly plasmolysed cells treated over 15 h by 1 mM AA or 1 mM AA with a mixture of copper and iron chelators (0.3 mM bathocuproine, 0.3 mM deferoxamine). (B) Mean ± SE numbers of cells with morphological PCD symptoms in root hairs (trichoblasts) and atrichloblasts (data of 10 independent trials). (C) Mean ±SE FITC-VAD-fmk fluorescence intensities (15 independent trials) in the control and after 15 h treatment by 1 mM l-ascorbic acid or 1 mM l-ascorbic acid with a mixture of copper and iron chelators. In all tests, the bathing solution contained 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). **P<0.001 and ***P<0.0001, ANOVA, respectively [data which were compared are indicated as (a)]. Treatment by 1 mM l-ascorbate with a mixture of transition metal chelators did not cause a statistically significant increase in the number of cells with morphological PCD symptoms and FITC-VAD-fmk fluorescence intensities. Fig. 4. View largeDownload slide Symptoms of programmed cell death (PCD) in Arabidopsis thaliana L. roots treated with l-ascorbic acid. (A) Changes in Arabidopsis root cell morphology (‘Morphological symptoms’) and activity of cell death proteases (‘FITC-VAD-fmk’) induced by 1 mM l-ascorbic acid. Arrows show areas with condensed bodies within irreversibly plasmolysed cells treated over 15 h by 1 mM AA or 1 mM AA with a mixture of copper and iron chelators (0.3 mM bathocuproine, 0.3 mM deferoxamine). (B) Mean ± SE numbers of cells with morphological PCD symptoms in root hairs (trichoblasts) and atrichloblasts (data of 10 independent trials). (C) Mean ±SE FITC-VAD-fmk fluorescence intensities (15 independent trials) in the control and after 15 h treatment by 1 mM l-ascorbic acid or 1 mM l-ascorbic acid with a mixture of copper and iron chelators. In all tests, the bathing solution contained 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). **P<0.001 and ***P<0.0001, ANOVA, respectively [data which were compared are indicated as (a)]. Treatment by 1 mM l-ascorbate with a mixture of transition metal chelators did not cause a statistically significant increase in the number of cells with morphological PCD symptoms and FITC-VAD-fmk fluorescence intensities. Measurement of Asc–-mediated currents by the patch-clamp technique Demidchik et al. (2010) have shown that exogenously applied AA (1 mM) does not activate Ca2+ influx currents in Arabidopsis root epidermal protoplasts in conditions of high [K+] in the pipette solution (PS: 80 mM K+). Here, using the same experimental system, the effect of 1 mM AA was examined in conditions of high Na+ concentration in PS (50 mM Na+; Fig. 5). AA treatment during 15–30 min did not cause statistically significant modifications of Ca2+ influx currents (P>0.01; ANOVA; n=7). This confirmed that an intact cell wall is required for AA-induced activation of Ca2+ influx. Fig. 5. View largeDownload slide Changes in plasma membrane currents induced by intracellular and extracellular l-ascorbate in Arabidopsis thaliana root cell protoplasts. (A) Typical whole-cell currents obtained in different conditions. ‘40 gluconate in PS’: pipette solution (PS) contained 40 mM Na-gluconate, 10 mM NaCl, 0.75 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; bathing solution (BS) contained 20 mM CaCl2, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1 mM Tris. ‘40 ascorbate in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 gluconate in PS’. ‘40 gluconate in PS+1 mM exogenous AA’: PS salines were the same as in ‘40 gluconate in PS’; BS contained 20 mM CaCl2, 1 mM l-ascorbic acid, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1.5 mM Tris. ‘40 ascorbate, 1 mM A9C in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 1 mM anthracene-9-carboxylic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 ascorbate in PS’. Holding voltages were corrected by the JPCalc command in Clampex 10.6. (B) Mean ±SE current–voltage relationships (I–V curves) measured in protoplasts patch-clamped using different PSs (n=6–10). PS and BS compositions are shown in (A). Fig. 5. View largeDownload slide Changes in plasma membrane currents induced by intracellular and extracellular l-ascorbate in Arabidopsis thaliana root cell protoplasts. (A) Typical whole-cell currents obtained in different conditions. ‘40 gluconate in PS’: pipette solution (PS) contained 40 mM Na-gluconate, 10 mM NaCl, 0.75 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; bathing solution (BS) contained 20 mM CaCl2, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1 mM Tris. ‘40 ascorbate in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 gluconate in PS’. ‘40 gluconate in PS+1 mM exogenous AA’: PS salines were the same as in ‘40 gluconate in PS’; BS contained 20 mM CaCl2, 1 mM l-ascorbic acid, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1.5 mM Tris. ‘40 ascorbate, 1 mM A9C in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 1 mM anthracene-9-carboxylic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 ascorbate in PS’. Holding voltages were corrected by the JPCalc command in Clampex 10.6. (B) Mean ±SE current–voltage relationships (I–V curves) measured in protoplasts patch-clamped using different PSs (n=6–10). PS and BS compositions are shown in (A). Organic anions, including malate–, oxalate–, and citrate–, can move through the plant plasma membranes, including those of Arabidopsis roots, but the Asc– permeability has not been tested so far (Diatloff et al., 2004; Gruber et al., 2010; Hedrich, 2012). To assess Asc– efflux currents, gluconate– was substituted with Asc– in PS, and the test with an anion channel blocker, anthracene-9-carboxylic acid (A9C), was carried out (Fig. 5). Negative inwardly directed currents, which are responsible for anion efflux, were recorded and analysed. When a pipette was filled with 50 mM gluconate– (‘Gluconate pipette’ in Fig. 5A), a small inwardly directed current was observed, which corresponded to Ca2+ influx through constitutive Ca2+-permeable non-selective cation channels (Demidchik et al., 2002). Patch-clamp studies have demonstrated that gluconate appears to be ‘virtually impermeant’ to anion channels (Alvarez-Leefmans and Russell, 1990). In this respect, addition of Asc– instead of gluconate– to PS should reveal the Asc– conductance. Indeed, the addition of 40 mM Asc– in PS resulted in the appearance of a large inwardly directed current, which was three times larger than the current measured using the ‘Gluconate pipette’ (Fig. 5). The Asc– efflux current was completely blocked by the addition of 1 mM A9C to the PS (Fig. 5). A9C also caused a shift of the current–voltage curve in the direction of the reversal potential for Ca2+ (calculated reversal potentials were as follows: ECl= –33.5; ECa=44.2; ENa= –156.5). Thus, these data provided evidence for a relatively good Asc– permeability through the plasma membrane of A. thaliana root cells. It should be noted that the focus of this study was not on the analysis of ascorbate-permeable ion channels. Therefore, further research is required to investigate the biophysical properties and molecular nature of root Asc– efflux conductances. Testing ascorbate release: EPR spectroscopy of root Asc·– in normal and NaCl-stressed plants EPR spectroscopy techniques were applied to assess Asc– efflux qualitatively (Fig. 6). This approach does not show AA release per se but it provides information about the occurrence of its oxidized form, Asc·–, in the apoplast of intact roots. In this case, NaCl (250 mM) was used as an inducer of AA release, while A9C (1 mM) was applied as an inhibitor of AA efflux. Fig. 6. View largeDownload slide The effect of salt stress on the generation of ascorbyl radicals (Asc·–) in intact Arabidopsis thaliana roots studied with EPR spectroscopy. (A) Typical Asc·– spectra in the solution containing 1 mM l-ascorbic acid (‘1 mM AA’), the same solution with 10 mM H2O2 (‘1 mM AA, 10 mM H2O2’), and in root exudates from untreated roots [‘Roots’, corresponding to ‘Control’ in (B)], roots treated by 250 mM NaCl (‘Roots, 250 mM NaCl’), and roots treated with the same NaCl concentration in the presence of 1 mM anthracene-9-carboxylic acid (‘Roots, 250 mM NaCl, 1 mM A9C’) or reduced glutathione (‘Roots, 250 mM NaCl, 1 mM GSH’). (B, C) Mean ±SD EPR signal intensities (Asc·– spectra; n=6–9) obtained in the experimental conditions indicated in (A). Note the different scale on the x-axis in (C) and (B). All experiments were carried out using buffer solution (BS) containing 10 mM Tris (titrated by MES to pH 8.0; Sigma Ultra). Statistical analyses: ANOVA, comparison of control and NaCl-treated roots (B; ***P<0.0001), NaCl-treated roots and NaCl-treated roots with addition of A9C (B; ***P<0.001), cell-free 1 mM AA and 1 mM AA with 10 mM H2O2 (C; ***P<0.0001). Data which were compared are indicated as (a). Fig. 6. View largeDownload slide The effect of salt stress on the generation of ascorbyl radicals (Asc·–) in intact Arabidopsis thaliana roots studied with EPR spectroscopy. (A) Typical Asc·– spectra in the solution containing 1 mM l-ascorbic acid (‘1 mM AA’), the same solution with 10 mM H2O2 (‘1 mM AA, 10 mM H2O2’), and in root exudates from untreated roots [‘Roots’, corresponding to ‘Control’ in (B)], roots treated by 250 mM NaCl (‘Roots, 250 mM NaCl’), and roots treated with the same NaCl concentration in the presence of 1 mM anthracene-9-carboxylic acid (‘Roots, 250 mM NaCl, 1 mM A9C’) or reduced glutathione (‘Roots, 250 mM NaCl, 1 mM GSH’). (B, C) Mean ±SD EPR signal intensities (Asc·– spectra; n=6–9) obtained in the experimental conditions indicated in (A). Note the different scale on the x-axis in (C) and (B). All experiments were carried out using buffer solution (BS) containing 10 mM Tris (titrated by MES to pH 8.0; Sigma Ultra). Statistical analyses: ANOVA, comparison of control and NaCl-treated roots (B; ***P<0.0001), NaCl-treated roots and NaCl-treated roots with addition of A9C (B; ***P<0.001), cell-free 1 mM AA and 1 mM AA with 10 mM H2O2 (C; ***P<0.0001). Data which were compared are indicated as (a). EPR spectroscopy-based measurements of Asc·– in intact A. thaliana roots have recently been developed in our laboratories (Sosan et al., 2016). We have also analysed the potential of other sensitive techniques, such as measuring the radioactively labelled Asc– efflux (Parsons and Fry, 2010) and electrospray ionization MS (Grillet et al., 2014); however, these techniques had significant technical limitations in studying intact roots. In normal conditions, a constitutive Asc·– level was detected in A. thaliana roots (Fig. 6), suggesting that some Asc· is present in the apoplast. The characteristic ascorbyl radical EPR doublet (Laroff et al., 1972; Halliwell and Gutteridge, 2015) was comparable with those that were generated in buffer solution containing 1 mM AA (pH 8.0). Previously, Parsons and Fry (2010) have demonstrated that AA leaks from plant cells stressed by H2O2 for a few seconds. Here, treatment of roots during 3 min by 250 mM NaCl increased Asc·– peak intensity up to 3–4 times. Longer treatment times did not increase Asc·– signal intensity, although, in some cases, it started to decline after treatment for 3 min. The Asc·– signal was sensitive to reduced glutathione in both control and NaCl-treated roots. Reduced glutathione was also effective in vitro, quenching the Asc·– signal measured in solution containing 1 mM AA and 10 mM H2O2 (Fig. 6). Addition of the anion channel blocker, A9C (1 mM), which inhibited Asc– efflux currents in root protoplasts (Fig. 5), together with NaCl, caused a decrease of the NaCl-induced Asc·– signal. Overall, these data demonstrated that the amount of Asc·– increases in the apoplastic space after treatment by NaCl in an A9C-dependent manner. This is indicative of Asc·– release in response to the depolarizing action caused by NaCl (as the counterion for K+). Discussion The data presented here demonstrate that AA, starting from 30 µM, induces elevation of [Ca2+]cyt. in intact Arabidopsis roots. The shape of AA-induced [Ca2+]cyt. transients was similar to that of a mixture AA and Cu2+, which is capable of generating HO·, although the magnitudes of AA-induced [Ca2+]cyt. peaks were smaller (Fig. 1). The overall reaction of [Ca2+]cyt. to AA closely resembles the reaction to ADP (Demidchik et al., 2011) and some phytohormones (Straltsova et al., 2015), which potentially act via stimulation of ROS/HO· production by NADPH oxidase (Demidchik and Shabala, 2018). The peak [Ca2+]cyt. induced by physiological apoplastic AA concentrations (10 µM to 3 mM; Smirnoff and Wheeler, 2000) was reproducible and statistically significant, although it did not exceed 100–150 nM. Such a moderate [Ca2+]cyt. transient is supposedly a signal for finely tuned adjustment of cell metabolism and gene expression. Interestingly, long-term exposure to 1 mM AA (15 h and 40 h; 1 mM AA) moderately increased PCD symptoms in root cells but did not cause complete death of the root system even after 40 h exposure (Fig. 4). Potentially, this can also have a regulatory function. Here the application of AA alongside the mixture of copper and iron chelators prevented AA-induced PCD symptoms, clearly demonstrating the role of these metals in AA-induced PCD. Intriguingly, PCD symptoms evoked by AA were more often observed in trichoblasts than in atrichoblasts, coinciding with the reported higher density of ROS-induced Ca2+ influx current in root hair protoplasts (Demidchik et al., 2003). Hypothetically, AA levels >3 mM, which caused high [Ca2+]cyt. peaks (Fig. 1), can occur in the apoplast, when cells collapse and an intracellular medium and organelles containing high AA levels are excreted. Plant leaf intracellular AA levels vary from 10 mM to 75 mM, while roots accumulate up to 5–10 mM AA; therefore, a lot of ascorbate can potentially be released locally (Smirnoff and Wheeler, 2000; Kawa et al., 2016; Alscher and Hess, 2017). AA levels >10 mM triggered very strong Ca2+ influx according to the dose–response curve presented in Fig. 1. Release of AA can be caused by mechanical injury (wounding stress) and/or PCD. Inducing PCD itself (Fig. 4), AA may be a trigger of self-release, with a further induction of Ca2+ signals. This mechanism can hypothetically play a role in systemic signal transduction stimulating propagation of Ca2+ signals in the case of pathogen attack or wounding stress. Removal of the cell wall abolished the AA-induced activation of Ca2+ signals (Fig. 3), demonstrating that factors needed for this activation reside in the cell wall. Addition of copper or iron chelators halved peak [Ca2+]cyt. transients, while their combined application fully suppressed them (Fig. 3). Taken together, these data indicate that AA effects on [Ca2+]cyt. critically depend on the cell wall Cu+/2+ and Fe2+/3+. These metals are abundant in the apoplast, where they form one of the biggest pools in plants (Sattelmacher, 2001; Printz et al., 2016). In the cell wall, Cu+/2+ and Fe2+/3+ can bind to negatively charged residues of polysaccharides and other organic substances, such as pectin, hemicellulose, and lignin (Fry et al., 2002; Fry, 2004; Wertz et al., 2017). Normally, the apoplastic activities of ionic Cu+/2+ and Fe2+/3+ are very low, while their catalytic activities are maintained at a relatively high level and increase under stress conditions, such as drought (Becana and Klucas, 1992; Moran et al., 1994, 1997). Grillet et al. (2014) have recently shown that AA efflux from root cells is critically important for reduction and mobilization of apoplastic iron and that it is a necessary step for iron acquisition in Pisum sativum. Moreover, Guo et al. (2017) have demonstrated that AA efflux alleviated iron deficiency in an abscisic acid (ABA)-dependent manner. These authors have speculated that AA can stimulate the ABA biosynthesis regulating plant adaptation to iron deficiency. The data presented here demonstrate that AA in the presence of cell wall-bound transition metals can induce Ca2+ signals, which are also known as major modulators of ABA-mediated processes. Here, a very wide range of copper and iron concentrations was tested individually and in combination with AA. Application of CuCl2 at a level higher than 1 µM caused significant stimulation of the AA-induced [Ca2+]cyt. elevation, while addition of Fe(II)-EDTA showed similar effects only at levels >1 mM. This difference can be explained by much higher copper reactivity in catalysis of Fenton-like reactions compared with iron (Halliwell and Gutteridge, 2015) as well as better solubility of copper salts as compared with Fe(II/III)-EDTA complexes. Nevertheless, iron chelators were effective in aequorin luminometry, suggesting that catalytically active ‘bio-available’ iron, similar to copper, is important for induction of Ca2+ signals by exogenous AA. In animal cells, AA efflux is dominated by volume-sensitive and Ca2+-dependent anion channels, but some other mechanisms can also be involved, including gap junction hemichannels, exocytosis of secretory vesicles, and potentially homo- and hetero-exchange systems (Corti et al., 2010). In this study, addition of AA to a patch-clamp pipette instead of gluconate evoked large inward currents, which were sensitive to 1 mM A9C (a blocker of anion channels; Fig. 5). Ascorbate efflux currents showed rapid activation kinetics in response to voltage steps, similar to R-type anionic conductances reported in a number of plant preparations (Diatloff et al., 2004; Roberts, 2006; Kollist et al., 2011; Hedrich, 2012). Ascorbate currents were not previously measured in plant protoplasts, but they can be compared with currents mediated by citrate, malate, and other organic anions (Gruber et al., 2010). Among the major classes of plant anion channels (encoded by ALMT, CLC, and SLAC), members of the ‘aluminium-activated malate transporter’ (ALMT) gene family are the best candidates for mediating ascorbate efflux. These channels are responsible for transport of large organic anions, such as malate, oxalate, and citrate. They have been discovered as A9C-sensitive systems releasing malate and other organic acids to chelate soil Al3+ (Ryan et al., 1995, 1997). Here, we provided the evidence that these channels can also be involved in another important process, which is release of AA to the extracellular space. AA does not form complexes with Al3+; therefore, this reaction should have a different evolutionary route from Al tolerance. Current–voltage curves of ascorbate efflux conductance (Fig. 5) were very similar to those that were reported in Al-free conditions (control) for malate efflux channels from different preparations, including Arabidopsis thaliana ALMT1 expressed in Xenopus oocytes (Hoekenga et al., 2006; Piñeros et al., 2008; Gruber et al., 2010) or Triticum aestivum ALMT1 expressed in tobacco culture cells (Zhang et al., 2008). Using EPR spectroscopy allowed us to resolve monodehydroascorbate reductase (MDHAR) spectra in control plants and the 9AC-sensitive burst of the MDHAR spectrum intensity induced by the addition of NaCl (3 min, 250 mM NaCl; Fig. 6). After 10 min and 15 min exposure to NaCl, a 2- to 3-fold decrease in MDHAR spectrum intensity was noted. We may speculate that by that time released AA was oxidized by mobilized iron and copper. AA release from plant cells has been reported several times in a variety of circumstances in the plant literature (Luwe et al., 1993; Luwe and Heber, 1995; Parsons and Fry, 2010; Grillet et al., 2014). Luwe and Heber (1995) measured AA content in control and O3-treated leaves of spinach. They found that leaf AA content increased after O3 treatment from 1–2 mM to 2–3 mM. Grillet et al. (2014) found that embryos of pea and A. thaliana excrete AA for iron reduction and uptake. Parsons and Fry (2010) tested the effect of 1–10 mM H2O2 on AA efflux in rose and A. thaliana suspension cultures in relation to mechanisms of antioxidant defence in cell cultures. They showed that H2O2 induces loss of 20% of cell AA in a few minutes. Peak AA efflux occurred in the first 100 s of H2O2 application. These authors also measured H2O2-induсed electrolyte leakage, which accompanied AA release, but was not related to the loss of cell integrity and viability. According to the literature and data shown here, plant plasma membranes have good permeability to AA. Therefore, ascorbate can function as a counterion during ROS-induced K+ efflux under oxidative stress conditions (Demidchik et al., 2014; Demidchik, 2015). This partially explains the very fast kinetics of AA release found by Parsons and Fry (2010). Potassium is the most abundant ion in the cytosol and a major species that leaves a plant cell in response to key stresses, such as NaCl, drought, pathogens, hypoxia, O3, UV, xenobiotics, heavy metals, etc. (Shabala et al., 2006; Demidchik et al., 2014). The potassium efflux channel GORK is activated by ROS (H2O2 and HO·), which are generated in response to stresses (Demidchik et al., 2010, 2018). This activation normally results in stress-induced leakage of K+ accompanied by leakage of anions. Salt stress (50 mM NaCl) causes the decrease of cytosolic K+ from 70–80 mM to 20–30 mM (Shabala et al., 2006). An equal amount of the anions should leave the cell to equilibrate changes in electric charge and ionic balance caused by K+ loss. A number of anions can potentially function as counterions, and ascorbate could be one of them. If GORK-mediated K+ efflux and ascorbate release are related processes, this can have important consequences for plants, including (i) additional stimulation of the ROS-induced K+ efflux via HO· production by released AA; (ii) involvement of GORK in systemic ROS-dependent responses and signal transduction; (iii) loss of cytosol antioxidant capacity in response to stresses that are accompanied by K+ leakage; and (iv) ascorbate can participate in the adjustment of plant metabolism under stress. These opportunities and other implications of AA release are summarized in Fig. 7. We hypothesize that ascorbate efflux can be part of the plant cell response to stresses initiating redox-dependent Ca2+ signalling and K+ efflux at the plant plasma membrane (Fig. 7). Ascorbate release can also be a reason for a moderate increase in PCD, which can have an adaptive role or originate the long-distance signalling. Additionally, AA release can be an iron-mobilizing reaction, which is necessary for iron uptake by root cells, and AA, when released, may stimulate cell growth via the ROS–Ca2+ hub (Fig. 7; Demidchik and Shabala, 2018). Fig. 7. View largeDownload slide Hypothetical scheme of plant plasma membrane signalling and transport reactions involving l-ascorbate. Stresses or developmental signals can trigger NADPH oxidase activity, leading to production of ROS, such as O2·–, H2O2, and HO·, which activate Ca2+ influx and K+ efflux channels (see Demidchik and Shabala, 2018; Demidchik et al., 2003, 2010, 2018, for details). Stress-induced depolarization results in K+ efflux, which can potentially be accompanied by Asc– (ascorbate-) efflux. In this case, ascorbate– is transported as the counterion to maintain cell ionic and charge balance. Ascorbate accumulation in the apoplast results in reduction of transition metals, such as copper and iron, leading to generation of HO·, which can cause activation of Ca2+ influx channels in the same cell and neighbouring cells. Overall, ascorbate efflux stimulates the self-amplifying ROS–Ca2+ hub, which is based on activation of NADPH oxidase by cytosolic Ca2+ and stimulation of Ca2+ influx by transition metal/ascorbate-catalysed conversion of O2·–/H2O2 to HO·, which, in turn, activates Ca2+ influx channels. Moreover, ascorbate efflux may cause iron and copper reduction for nutritional needs. Please note that some other transporters, such as HO·-activated non-specific ion channels, which are permeable to both cation and anions (Zepeda-Jazo et al., 2011) and hypothetically mediate Ca2+ influx and AA efflux, can also be involved in AA signalling reactions. Fig. 7. View largeDownload slide Hypothetical scheme of plant plasma membrane signalling and transport reactions involving l-ascorbate. Stresses or developmental signals can trigger NADPH oxidase activity, leading to production of ROS, such as O2·–, H2O2, and HO·, which activate Ca2+ influx and K+ efflux channels (see Demidchik and Shabala, 2018; Demidchik et al., 2003, 2010, 2018, for details). Stress-induced depolarization results in K+ efflux, which can potentially be accompanied by Asc– (ascorbate-) efflux. In this case, ascorbate– is transported as the counterion to maintain cell ionic and charge balance. Ascorbate accumulation in the apoplast results in reduction of transition metals, such as copper and iron, leading to generation of HO·, which can cause activation of Ca2+ influx channels in the same cell and neighbouring cells. Overall, ascorbate efflux stimulates the self-amplifying ROS–Ca2+ hub, which is based on activation of NADPH oxidase by cytosolic Ca2+ and stimulation of Ca2+ influx by transition metal/ascorbate-catalysed conversion of O2·–/H2O2 to HO·, which, in turn, activates Ca2+ influx channels. Moreover, ascorbate efflux may cause iron and copper reduction for nutritional needs. Please note that some other transporters, such as HO·-activated non-specific ion channels, which are permeable to both cation and anions (Zepeda-Jazo et al., 2011) and hypothetically mediate Ca2+ influx and AA efflux, can also be involved in AA signalling reactions. Acknowledgements Financial support from the Russian Science Foundation (grant #15-14-30008 to VD) is gratefully acknowledged. 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All rights reserved. For permissions, please email: [email protected] This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Journal of Experimental Botany Oxford University Press

Novel roles of ascorbate in plants: induction of cytosolic Ca2+ signals and efflux from cells via anion channels

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Oxford University Press
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© The Author(s) 2018. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected]
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0022-0957
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10.1093/jxb/ery056
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Abstract

Abstract Ascorbate is not often considered as a signalling molecule in plants. This study demonstrates that, in Arabidopsis roots, exogenous l-ascorbic acid triggers a transient increase of the cytosolic free calcium activity ([Ca2+]cyt.) that is central to plant signalling. Exogenous copper and iron stimulate the ascorbate-induced [Ca2+]cyt. elevation, while cation channel blockers, free radical scavengers, low extracellular [Ca2+], transition metal chelators, and removal of the cell wall inhibit this reaction. These data show that apoplastic redox-active transition metals are involved in the ascorbate-induced [Ca2+]cyt. elevation. Exogenous ascorbate also induces a moderate increase in programmed cell death symptoms in intact roots, but it does not activate Ca2+ influx currents in patch-clamped root protoplasts. Intriguingly, the replacement of gluconate with ascorbate in the patch-clamp pipette reveals a large ascorbate efflux current, which shows sensitivity to the anion channel blocker, anthracene-9-carboxylic acid (A9C), indicative of the ascorbate release via anion channels. EPR spectroscopy measurements demonstrate that salinity (NaCl) triggers the accumulation of root apoplastic ascorbyl radicals in an A9C-dependent manner, confirming that l-ascorbate leaks through anion channels under depolarization. This mechanism may underlie ascorbate release, signalling phenomena, apoplastic redox reactions, iron acquisition, and control the ionic and electrical equilibrium (together with K+ efflux via GORK channels). Anion channels, Arabidopsis, ascorbate transport, ascorbic acid, ascorbyl radicals, calcium signalling, hydroxyl radicals, plasma membrane, reactive oxygen species, salinity Introduction l-Ascorbic acid (AA) is a key antioxidant, cofactor of redox enzymes, and precursor for some biosyntheses. In water solutions, this substance is present as a monovalent anion, ascorbate– (Asc–), which easily donates electrons to oxidized molecules and scavenges reactive oxygen species (ROS). For example, AA directly reacts with the most important ROS, such as superoxide (O2·–), hydroxyl radicals (HO·), and singlet oxygen (Zhang, 2013; Halliwell and Gutteridge, 2015). Moreover, AA is a major reducing agent for iron and copper [Cu2+/Fe3++electron from AA→Cu+/Fe2++dehydroascorbate (DHA)] and potentially for other transition metal ions, such as Mn2+/3+, inside and, hypothetically, outside the cell. This makes AA a pro-oxidant because Cu+ and Fe2+ are catalysts of HO· production, while HO· is the most powerful oxidizing species in plants (Demidchik, 2015; Halliwell and Gutteridge, 2015; Demidchik and Shabala, 2018). The pro-oxidative properties of AA in plants have not been studied in detail, while its role as an antioxidant has attracted enormous interest over the last few decades (Foyer and Noctor, 2011; Halliwell and Gutteridge, 2015; Wertz et al., 2017). In plant cells, the antioxidant function of AA is related to a number of processes, but the most important one is the donation of electrons in the Foyer–Halliwell–Asada cycle (FHA cycle), also known as the ‘ascorbate–gluthatione cycle’ (Foyer and Halliwell, 1976, 1977; Foyer and Noctor, 2011). The FHA cycle is active inside the cell, detoxifying superoxide anion radicals (O2·–), which are mainly generated as a result of so-called ‘electron leakage’ to triplet oxygen (O2) in photosynthetic, mitochondrial, peroxisomal, and probably other electron transport chains (Ozyigit et al., 2016). AA also takes part in the regeneration of α-tocopherol (vitamin E) in photosynthetic membranes and participates in chloroplast metabolism as a cofactor for violaxanthin de-epoxidase, an enzyme involved in xanthophyll cycle-mediated photoprotection (Smirnoff and Wheeler, 2000). Overall, these are very important physiological roles of AA relating this molecule exclusively to antioxidant functions. Nevertheless, this might not be the only function of AA in plants. In some plant species, AA is the substrate for the biosynthesis of oxalate and tartrate (Smirnoff and Wheeler, 2000). AA also controls plant development via regulation of phytohormone synthesis (Pastori et al., 2003). An exogenous AA is capable of stimulating an expansive growth, and the high ascorbate oxidase activity in the cell wall is correlated with areas of rapid cell expansion (Arrigoni, 1994; Gallie, 2013). The mechanistic explanation of these phenomena is largely missing. Hypothetically, AA can act as a cofactor for prolyl hydroxylase that hydroxylates proline residues in the cell wall hydroxyproline-rich glycoproteins required for cell expansion (Smirnoff and Wheeler, 2000). Another possibility is that AA stimulates HO· production via reduction of cell wall-bound transition metals, such as copper and iron (Demidchik, 2015). This can potentially result in the activation of Ca2+ influx, induction of the ROS–Ca2+ hub (via NADPH oxidase) required for sustained ROS production and Ca2+ influx stimulating exocytosis and cell elongation (Foreman et al., 2003; Demidchik and Shabala, 2018), and/or cleavage of cell wall polymers inducing cell wall loosening (Müller et al., 2009). Plants synthesize AA by at least by two pathways. The first one is an intracellular Smirnoff–Wheeler pathway (cytosol/mitochondria), when plants produce AA from l-galactose, which in turn is synthesized from GDP-d-mannose (Wheeler et al., 1998). A second AA biosynthetic pathway is located in the cell wall and relies on the substrate d-galacturonic acid, generated from the breakdown of pectin (Loewus and Kelly, 1961; Gallie, 2013). It is believed that the first pathway is the more important and capable of producing the most AA in plants (Smirnoff and Wheeler, 2000; Gallie, 2013). The physiological role of the second pathway of AA production is poorly understood. The levels of apoplastic AA and its reversibly oxidized forms (Asc·– and DHA) can control the activities of apoplastic transition metals, and, vice versa, metals, such as copper and iron, can affect AA stability and AA-mediated reactions in the cell wall. Transition metals such as copper and iron are abundant in the cell wall (Thornton and Macklon, 1989; Zhang et al., 1991; Fry et al., 2002). They can catalyse HO· generation from O2·–/H2O2 in the presence of AA in this compartment (Fry, 1998; Demidchik et al., 2010; Demidchik, 2015). The induction of Ca2+ influx for signalling purposes via AA-mediated HO· generation in the apoplast can also be proposed. In this context, a new signalling role for apoplastic AA is emerging. This possibility has not been addressed in higher plants. It can be hypothesized that apoplastic AA functions as a signalling molecule in plants. This can be due to redox properties of this molecule or potentially via some receptor-like mechanisms. In this context, the testing of potential elevations in cytosolic free Ca2+ ([Ca2+]cyt.) using Ca2+ aequorin luminometry is crucial; it can directly demonstrate AA involvement in central reactions of plant signalling (via generation of Ca2+ signals). If AA is a signalling molecule, it can have a system for release from plant cells and breakdown in the extracellular space. The breakdown can be catalysed by ascorbate peroxidase (De Gara, 2004) and ascorbate oxidase (De Tullio et al., 2013), while its release can hypothetically be mediated by anion channels. The negative electric potential difference across the plasma membrane of typical plant cell is –120 mV to –60 mV (Demidchik et al., 2002). This provides a driving force for passive ascorbate efflux from cells, if the plasma membrane is permeable to this anion. In order to answer this question, the plasma membrane conductance mediated by the AA efflux should be measured. The permeability of plant plasma membranes to Asc– has not previously been examined by electrophysiological techniques, although this question is definitely of fundamental importance for plant biology. Endogenous AA represents as much as 10% of soluble carbohydrates in some plant tissues (Noctor and Foyer, 1998); therefore, a tight control over transport of AA is crucial for the organism. In animal cells, anion channels dominate ascorbate efflux, although other mechanisms are also involved (Corti et al., 2010). Thus, it can be hypothesized that anion channels may also be involved in plant AA efflux. The aim of this study was to examine AA effects on Ca2+ signalling and explore the potential role of transition metals, Cu+/2+ and Fe2+/3+, in the development of these effects. The AA efflux currents were measured by conventional patch-clamp techniques using protoplasts isolated from roots. The ascorbate release was assessed by examining apoplastic ascorbyl radicals using EPR spectroscopy. The model plant, Arabidopsis thaliana, was used as a convenient system for electrophysiological, cell death, luminometry, and EPR spectroscopy assays. Materials and methods Plant material Wild-type (WT) A. thaliana, ecotype Columbia-0 (Col-0), was used to study programmed cell death (PCD) symptoms (microscopy), ionic currents (patch-clamp), and formation of Asc·– (EPR spectroscopy). Transgenic Col-0 were used to measure changes in [Ca2+]cyt. by aequorin luminometry. Col-0 seeds were obtained from the European Arabidopsis Stock Centre (arabidopsis.info). For all experiments, the plants were grown vertically on a surface of sterilized gel (in Petri dishes), which contained full-strength Murashige and Skoog (MS) medium (Murashige and Skoog, 1962), 1% sucrose (w/v), and 0.35% Phytagel (w/v, Sigma-Aldrich), pH 6.0/KOH. Constant temperature and irradiance photosynthetically active radiation (PAR) conditions were maintained in growth cabinets (22 ± 1 °C, 16 h daylight, LED lamps, with PAR wavelength, 200 μmol m−2 s−1 irradiance). For patch-clamp and luminometry measurements, root protoplasts were isolated using techniques developed by Demidchik and Tester (2002) and Demidchik et al. (2002, 2010). Briefly, roots were excised from 50–100 A. thaliana seedlings (7–12 d old) and cut into 0.5–2 mm long pieces using a razor blade in a plastic dish, which contained 4 ml of enzymatic solution comprising 1.5% (w/v) Cellulase Onozuka RS (Yakult Honsha, Tokyo, Japan), 1% (w/v) cellulysin (CalBiochem, Nottingham, UK), 0.1% (w/v) pectolyase Y-23 (Yakult Honsha), 0.1% (w/v) BSA (Sigma), 10 mM KCl, 10 mM CaCl2, 2 mM MgCl2, 2 mM MES, pH 6.0 with Tris; 290–300 mOsM adjusted with 330 mM sorbitol. After gentle shaking (45 rpm) in the enzyme solution for 30–50 min at 28 °C, protoplasts were filtered (30 µm pore mesh) and rinsed with holding solution (HS; 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM sucrose, 10 mM glucose, 2 mM MES, pH 6.0 with Tris; 290–300 mOsM with sorbitol). Protoplasts were collected by 5 min centrifugation at 200 g and ~0.5 ml was diluted with 8–10 ml of HS. Luminometric measurements of [Ca2+]cyt. Seven- to 12-day-old seedlings of A. thaliana plants constitutively expressing apoaequorin, under the control of the 35S Cauliflower mosaic virus (CaMV) promoter, were used (Knight et al., 1996, 1997; Polisensky and Braam, 1996). Seeds were kindly provided by Professor Marc Knight (Durham University). Luminometric measurements were carried out directly on these seedlings or on protoplasts. Ca2+ activity was recorded continuously up to 15 min with a single-tube Turner BioSystems Luminometer Model TD-20/20 controlled by Spreadsheet Interface Software (Promega). Standard aequorin chemiluminometry techniques were used to examine changes in [Ca2+]cyt. (Knight et al., 1996, 1997; Demidchik et al., 2011; Sosan et al., 2016). Coelenterazine, native (4 µM; Prolume Ltd, USA) was used in all measurements. The remaining aequorin was discharged for 5 min using a mixture containing 2 M CaCl2, 20% ethanol (v/v), and 0.01% Triton X-100 (v/v). The amount of free Ca2+ entering the cells was calculated using the calibration equation derived empirically: pCa=0.332588(−log k)+5.5593, where k=(luminescence counts s−1)/(total luminescence counts remaining) (Knight et al., 1996, 1997). Curves showing dose dependence of [Ca2+]cyt. in response to AA were fitted and analysed using SigmaPlot 10.0 (Systat Software Inc.). Patch-clamp measurements Conventional patch-clamp and protoplast isolation techniques were used (Demidchik et al., 2010). The standard bathing solution contained (in mM): 20 CaCl2, 2 Tris, 0.1 NaCl, adjusted to pH 6.0 with MES, and 290–300 mOsM, with sorbitol. A freshly prepared mixture of this solution with AA was applied in whole-cell outside-out patches. All pipette solutions (PSs) contained at least 10 mM Cl– and 100 nM Ca2+ adjusted with 2 mM Tris/MES, pH 7.2. Other saline solutions are indicated in the figure legends. Liquid junction potentials were calculated by JPCalc which is included in the Axon Clampex 10.6 software (Molecular Devices, USA) and corrected. The voltage was held at –90 mV, then square 7.6 seconds-long depolarizing or hyperpolarizing voltage pulses were applied. Currents were measured using PC-ONE Patch/Whole Cell Clamp (CORNERSTONE Series) amplifier (Dagan Corporation, USA) controlled by Digidata 1320/Clampex 10.6 (Molecular Devices, USA). Imaging symptoms of programmed cell death Standard bright-field and epi-fluorescent microscopy were used for the analysis of morphological (irreversible plasmolysis, cytoplasm shrinkage, plasma membrane rupture, appearance of dark bodies, etc.) and biochemical (caspase-like proteolytic activity; CaspACE FITC-VAD-fmk in situ marker kit; Promega, UK) cell death symptoms in root cells as described by Demidchik et al. (2010) and Hogg et al., (2011). An inverted Nikon TS100F microscope (Nikon, Japan) powered by NIS-Elements AR software and a standard fluorescein isothiocyanate (FITC) filter were used. The exposure time was 0.5 s. The bathing solution contained (in mM): 0.1 KCl, 0.1 CaCl2, pH 6.0 (4 MES/2 Tris). Five minutes staining with 0.5% Evans blue (Roth; w/v) was applied to AA-treated roots; then the dye was washed five times. To analyse protease activity, 10 μM FITC-VAD-fmk was applied together with the AA. After treatment with AA, the roots were washed three times for 5 min and observed under the microscope. Detection of Asc·– by EPR spectroscopy Standard EPR spectroscopy in liquid phase (20 °C) was used for the examination of Asc·– in intact roots, in control conditions, and after treatment by 100 mM NaCl (Liszkay et al., 2004; Demidchik et al., 2010). The cell-free tests were also carried out as required. Preliminary cell-free tests and analysis of the literature (Halliwell and Gutteridge, 2015) showed that the ascorbate radical signal is stable at pH 8.0; therefore, this pH level was used throughout the EPR tests. The basic solution for EPR spectroscopy tests included 2 mM Tris/MES (pH 8.0), 0.1 mM CaCl2, 0.1 mM KCl. For all tests, reagents were mixed 60 s before recording the EPR spectrum in oxygen-depleted conditions. ESR Grade Water (Noxygen) was used, and all solutions were additionally purified using ultrafine activated charcoal powder (Sigma-Aldrich) to remove any residues of transition metals. The EPR spectra were recorded on a Bruker EMX (X-band) spectrometer equipped with an ER 4103TM cylindrical mode resonator and analysed using WinEPR (Bruker). For in vitro tests, the EPR Spectrometer SPINSCAN (ADANI) was used with a standard cuvette set. Asc·– spectra were detected without the spin-trap as described by Buettner and Jurkiewicz (1993). In cell-free conditions, AA was directly mixed with buffer solution containing the substances being tested. To measure Asc·– generation in intact roots, 150 intact 1-week-old seedlings were removed from the medium, carefully washed several times, and adapted for 1 h in a 1 ml vial filled with buffer solution containing 0.1 mm KCl and 0.1 mM CaCl2 (pH 8.0; Tris/MES), then 2 cm lengths of the root tips were immersed for 5 min in the fresh buffer solution. After treatment, the eluate from the vial was directly collected in a Bruker AquaX system (the multiple bore design of closely spaced capillaries allows for much higher sensitivity compared with the capillary or flat-cell designs) for measurements of liquid samples. Results Exogenous l-ascorbate induces free radical- and transition metal-dependent elevation of cytosolic free Ca2+ To assess whether AA can trigger [Ca2+]cyt. elevations, 0.01–30 mM AA was added to excised roots of intact A. thaliana plants constitutively expressing aequorin (Fig. 1). Starting from 100 µM, AA induced a significant transient increase in [Ca2+]cyt., with a peak during 4–7 min (ANOVA test, P<0.001; Fig. 1A). The addition of buffer solution without AA or application of low AA concentrations (0.1–10 µM) did not cause significant changes in the cytosolic Ca2+ (Fig. 1B). Fig. 1. View largeDownload slide The effect of exogenously applied l-ascorbic acid (AA) on the activity of cytosolic Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 10 µM to 30 mM AA (I.–VII.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA are indicated in the panels. (IX.) [Ca2+]cyt. elevation in response to combined application of 1 mM Cu2+ and 1 µM AA. ‘DC’: time, when the discharge solution was added to roots (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different AA concentrations (mean ±SE; n=10–11). (C) Dependence of peak [Ca2+]cyt. on the concentration of Cu2+ applied with (open circles) or without (filled circles) 1 mM AA (mean ±SE; n=5–10). (D) Effect of iron ions applied as Fe(II)EDTA on peak AA-induced [Ca2+]cyt. transients (mean ±SE; n=4). The bathing solution in all tests contained 10 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). Fig. 1. View largeDownload slide The effect of exogenously applied l-ascorbic acid (AA) on the activity of cytosolic Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 10 µM to 30 mM AA (I.–VII.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA are indicated in the panels. (IX.) [Ca2+]cyt. elevation in response to combined application of 1 mM Cu2+ and 1 µM AA. ‘DC’: time, when the discharge solution was added to roots (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different AA concentrations (mean ±SE; n=10–11). (C) Dependence of peak [Ca2+]cyt. on the concentration of Cu2+ applied with (open circles) or without (filled circles) 1 mM AA (mean ±SE; n=5–10). (D) Effect of iron ions applied as Fe(II)EDTA on peak AA-induced [Ca2+]cyt. transients (mean ±SE; n=4). The bathing solution in all tests contained 10 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). The basal [Ca2+]cyt. level was 70.8 ± 3.8 nM (±SE; n=15). Peak values were calculated by subtracting this level (tested in each individual trial) from the obtained maximal (peak) increase of [Ca2+]cyt.. The relationship between the mean peak [Ca2+]cyt. increase and the concentration of applied AA is demonstrated in Fig. 1B. This ‘dose–response curve’ did not show saturation even at very high levels of AA in the bathing solution. The maximal [Ca2+]cyt. elevation was induced by 30 mM AA (321 ± 40.1 nM; ±SE; n=10). At the same time, physiological levels of AA, which in roots reach 5–10 mM (Smirnoff and Wheeler, 2000; Kawa et al., 2016; Alscher and Hess, 2017), triggered effects that were 2–5 times smaller (Fig. 1). Hypothetically, exogenous AA can undergo transition metal-catalysed oxidation leading to formation of HO·, activation of Ca2+-permeable cation channels, and elevation of [Ca2+]cyt. (Demidchik et al., 2003). Copper ions are the most powerful catalysts of Fenton-like reactions (generating HO·) compared with iron and other transition metals (Fry, 1998; Fry et al., 2002; Halliwell and Gutteridge, 2015). Addition of Cu2+ together with AA resulted in [Ca2+]cyt. elevation, which demonstrated parameters similar to AA-induced [Ca2+]cyt. but had much higher peak values (Fig. 1). Very low Cu2+ concentrations (10–7 M and 10–6 M) caused significant stimulation of AA-induced [Ca2+]cyt. transients. This is aligned with plant cell catalytic activities of transition metals (Cu2+ and Fe3+), which can reach 0.3–15 µM (Becana and Klucas, 1992). The minimal Cu2+ concentration which caused statistically significant stimulation of the AA effect was 0.1 µM (n=7; P<0.01; ANOVA; Fig. 1C). Iron [Fe(II)-EDTA] also stimulated AA-induced [Ca2+]cyt. elevation, but it was less effective than copper, causing significant stimulation at millimolar levels (Fig. 1D). To examine the involvement of free radicals, such as HO·, in the AA-induced [Ca2+]cyt. increase, AA was added together with a free radical scavenger, thiourea (Fig. 2). Thiourea effects on HO· generation and HO·-dependent processes have previously been observed in different preparations, including EPR studies on A. thaliana roots (Demidchik et al., 2003, 2010; Liszkay et al., 2004; Halliwell and Gutteridge, 2015). Addition of 3 mM thiourea to aequorin-expressing roots without AA caused a small increase in [Ca2+]cyt. (peak: 15 ± 6.2 nM; mean ±SE; n=6). When 3 mM thiourea was added together with 1 mM or 3 mM AA, the elevation of [Ca2+]cyt caused by AA decreased dramatically (up to eight times; Fig. 2). This test with thiourea, which is considered by some authors as a hydroxyl radical-specific scavenger (Liszkay et al., 2004), strongly suggests that AA acts on cytosolic free Ca2+ via a free radical-dependent pathway, such as generation of HO·, causing activation of Ca2+-permeable cation channels (Demidchik, 2015). Similar results (inhibition of AA-induced [Ca2+]cyt. transients; n=5; data not shown) were obtained here for DMSO (0.03%, v/v), which is a non-specific scavenger of free radicals (Demidchik et al., 2010; Halliwell and Gutteridge, 2015). Fig. 2. View largeDownload slide Inhibitory action of thiourea (free radical scavenger) on the l-ascorbate-induced elevation of the cytosolic free Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 1 mM and 3 mM AA (I. and III.) and combined application of these concentrations with 3 mM thiourea (II. and IV.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA and thiourea are indicated in the panels. DС: time of applying the discharge solution (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different treatments by AA and combined application of AA and thiourea (mean ±SE; n=6–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a) and (b)). The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Fig. 2. View largeDownload slide Inhibitory action of thiourea (free radical scavenger) on the l-ascorbate-induced elevation of the cytosolic free Ca2+ ([Ca2+]cyt.) in Arabidopsis thaliana roots expressing aequorin. (A) Typical [Ca2+]cyt. transients induced by 1 mM and 3 mM AA (I. and III.) and combined application of these concentrations with 3 mM thiourea (II. and IV.). The time of AA addition to the bathing solution is indicated by a dashed arrow. Concentrations of AA and thiourea are indicated in the panels. DС: time of applying the discharge solution (indicated by an arrow; see the Materials and methods for details). (B) Dependence of peak [Ca2+]cyt. elevation on different treatments by AA and combined application of AA and thiourea (mean ±SE; n=6–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a) and (b)). The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Two classical blockers of plasma membrane Ca2+ influx channels, Gd3+ and La3+ (Demidchik et al., 2002), were used to define the involvement of cation channels in AA-induced [Ca2+]cyt. elevation (Fig. 3). The results showed that Gd3+ and La3+ were very effective inhibitors of cytosolic calcium burst induced by both 1 mM and 3 mM AA. This confirms that the effect on cytosolic Ca2+ was related to the activation of Ca2+-permeable cation channels. Moreover, the decrease of extracellular Ca2+ from 10 mM to 50 µM caused a decrease of peak [Ca2+]cyt. elevations induced by 1 mM AA from 44.4 ± 7.2 nM (mean ±SE; n=9) to 9.5 ± 3.6 nM; mean ±SE; n=6). Fig. 3. View largeDownload slide Peak transient [Ca2+]cyt. increase induced by 3 mM AA in the presence of lanthanide cations (La3+ and Gd3+), transition metal chelators (bathocuproine and deferoxamine), or after removal of the cell wall by isolation of protoplasts [mean ±SE; n=7–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a)]. Protoplast isolation techniques were the same as in patch-clamp analyses (see the Materials and methods). Concentrations of salines are indicated in the figure. The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Fig. 3. View largeDownload slide Peak transient [Ca2+]cyt. increase induced by 3 mM AA in the presence of lanthanide cations (La3+ and Gd3+), transition metal chelators (bathocuproine and deferoxamine), or after removal of the cell wall by isolation of protoplasts [mean ±SE; n=7–10; ***P<0.0001, ANOVA; data which were compared are indicated as (a)]. Protoplast isolation techniques were the same as in patch-clamp analyses (see the Materials and methods). Concentrations of salines are indicated in the figure. The bathing solution contained 10 mM CaCl2, pH 6.0 adjusted by 2 mM Tris/4 mM MES. Based on data presented in Fig. 2, the AA-induced [Ca2+]cyt. elevation is related to HO· generation in the cell wall, which can be catalysed by transition metals (copper and iron) residing in the cell wall. To examine this hypothesis, plants were pre-treated by copper (0.3 mM bathocuproine) and iron (0.3 mM defereoxamine) chelators (Fig. 3). This reduced the AA-induced [Ca2+]cyt. elevations by 40–60%. Combined application of AA with both 0.3 mM bathocuproine and 0.3 mM defereoxamine resulted in complete inhibition of AA-induced [Ca2+]cyt transients (Fig. 3), demonstrating that copper and iron (and HO· generation mediated by these metals) are central to the AA-induced Ca2+ signalling. Apart from the application of chelators, cell wall metals were depleted by the removal of the cell wall using enzymatic treatment (protoplast were isolated by the same protocols as were used for patch-clamping; Fig. 3). The protoplasts lacked AA-induced [Ca2+]cyt elevation. This additionally confirmed that the cell wall was crucial for AA effects. Cell death symptoms in roots treated by l-ascorbic acid PCD induced by a mixture of AA and Cu2+ and stressors associated with oxidative burst (NaCl, pathogens, heavy metals, etc.) has recently been studied in detail (Demidchik et al., 2010, 2017; Petrov et al., 2015). If AA added to the cell wall is capable of producing HO· and stimulating Ca2+ influx, then the induction of PCD in the AA-treated roots is a possible scenario. Stresses can induce a number of detectable morphological symptoms of PCD, such as irreversible plasmolysis, cytoplasm shrinkage, plasma membrane rupture, and appearance of dark spots at the site of the nucleus (Demidchik et al., 2010, 2017; Hogg et al., 2011). Plasma membrane damage can be monitored by staining with membrane-impermeant dyes, such as Evans blue. Hydrolytic ‘self-digestion’ reactions accompanying PCD can be assessed by staining the cell death proteases using the CaspACE FITC-VAD-fmk in situ marker kit (Promega, UK; Demidchik et al., 2010). Fluorescently labelled protease inhibitor zVAD-fmk (FITC-VAD-fmk) is a widely used tool for the observation of PCD protease activation in intact tissues, because, at micromolar concentrations, this substance does not significantly inhibit proteases, although it binds to and fluorescently labels them (Elbaz et al., 2002; Bonneau et al., 2008). In this case, an increased intracellular fluorescence indicates a higher protease activity. For AA-treated roots, morphology, Evans blue, and FITC-VAD-fmk tests demonstrated that long-term treatment with AA (15 h and 40 h) triggered a statistically significant increase in the number of cells with PCD symptoms (Fig. 4). Trichoblasts (root hairs) were more sensitive to both 15 h and 40 h treatment by AA than mature epidermal cells (atrichoblasts). The addition of 0.3 mM bathocuproine and 0.3 mM defereoxamine together with AA prevented the development of morphological PCD symptoms in all epidermal cell types. Treatment by 1 mM AA caused an increase of FITC-VAD-fmk fluorescence by 30–40%, which was abolished by the combined addition of 0.3 mM bathocuproine and 0.3 mM defereoxamine (Fig. 4C). Overall, these tests showed that physiological levels of exogenous AA induced a very moderate increase in the level of PCD symptoms in roots. AA was mainly targeting root hairs, where the cell wall is thinner, and metabolic activities, including those that are catalysed by copper and iron, are higher. Fig. 4. View largeDownload slide Symptoms of programmed cell death (PCD) in Arabidopsis thaliana L. roots treated with l-ascorbic acid. (A) Changes in Arabidopsis root cell morphology (‘Morphological symptoms’) and activity of cell death proteases (‘FITC-VAD-fmk’) induced by 1 mM l-ascorbic acid. Arrows show areas with condensed bodies within irreversibly plasmolysed cells treated over 15 h by 1 mM AA or 1 mM AA with a mixture of copper and iron chelators (0.3 mM bathocuproine, 0.3 mM deferoxamine). (B) Mean ± SE numbers of cells with morphological PCD symptoms in root hairs (trichoblasts) and atrichloblasts (data of 10 independent trials). (C) Mean ±SE FITC-VAD-fmk fluorescence intensities (15 independent trials) in the control and after 15 h treatment by 1 mM l-ascorbic acid or 1 mM l-ascorbic acid with a mixture of copper and iron chelators. In all tests, the bathing solution contained 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). **P<0.001 and ***P<0.0001, ANOVA, respectively [data which were compared are indicated as (a)]. Treatment by 1 mM l-ascorbate with a mixture of transition metal chelators did not cause a statistically significant increase in the number of cells with morphological PCD symptoms and FITC-VAD-fmk fluorescence intensities. Fig. 4. View largeDownload slide Symptoms of programmed cell death (PCD) in Arabidopsis thaliana L. roots treated with l-ascorbic acid. (A) Changes in Arabidopsis root cell morphology (‘Morphological symptoms’) and activity of cell death proteases (‘FITC-VAD-fmk’) induced by 1 mM l-ascorbic acid. Arrows show areas with condensed bodies within irreversibly plasmolysed cells treated over 15 h by 1 mM AA or 1 mM AA with a mixture of copper and iron chelators (0.3 mM bathocuproine, 0.3 mM deferoxamine). (B) Mean ± SE numbers of cells with morphological PCD symptoms in root hairs (trichoblasts) and atrichloblasts (data of 10 independent trials). (C) Mean ±SE FITC-VAD-fmk fluorescence intensities (15 independent trials) in the control and after 15 h treatment by 1 mM l-ascorbic acid or 1 mM l-ascorbic acid with a mixture of copper and iron chelators. In all tests, the bathing solution contained 0.1 mM KCl, 0.1 mM CaCl2, pH 6.0 (adjusted by 2 mM Tris/4 mM MES). **P<0.001 and ***P<0.0001, ANOVA, respectively [data which were compared are indicated as (a)]. Treatment by 1 mM l-ascorbate with a mixture of transition metal chelators did not cause a statistically significant increase in the number of cells with morphological PCD symptoms and FITC-VAD-fmk fluorescence intensities. Measurement of Asc–-mediated currents by the patch-clamp technique Demidchik et al. (2010) have shown that exogenously applied AA (1 mM) does not activate Ca2+ influx currents in Arabidopsis root epidermal protoplasts in conditions of high [K+] in the pipette solution (PS: 80 mM K+). Here, using the same experimental system, the effect of 1 mM AA was examined in conditions of high Na+ concentration in PS (50 mM Na+; Fig. 5). AA treatment during 15–30 min did not cause statistically significant modifications of Ca2+ influx currents (P>0.01; ANOVA; n=7). This confirmed that an intact cell wall is required for AA-induced activation of Ca2+ influx. Fig. 5. View largeDownload slide Changes in plasma membrane currents induced by intracellular and extracellular l-ascorbate in Arabidopsis thaliana root cell protoplasts. (A) Typical whole-cell currents obtained in different conditions. ‘40 gluconate in PS’: pipette solution (PS) contained 40 mM Na-gluconate, 10 mM NaCl, 0.75 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; bathing solution (BS) contained 20 mM CaCl2, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1 mM Tris. ‘40 ascorbate in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 gluconate in PS’. ‘40 gluconate in PS+1 mM exogenous AA’: PS salines were the same as in ‘40 gluconate in PS’; BS contained 20 mM CaCl2, 1 mM l-ascorbic acid, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1.5 mM Tris. ‘40 ascorbate, 1 mM A9C in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 1 mM anthracene-9-carboxylic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 ascorbate in PS’. Holding voltages were corrected by the JPCalc command in Clampex 10.6. (B) Mean ±SE current–voltage relationships (I–V curves) measured in protoplasts patch-clamped using different PSs (n=6–10). PS and BS compositions are shown in (A). Fig. 5. View largeDownload slide Changes in plasma membrane currents induced by intracellular and extracellular l-ascorbate in Arabidopsis thaliana root cell protoplasts. (A) Typical whole-cell currents obtained in different conditions. ‘40 gluconate in PS’: pipette solution (PS) contained 40 mM Na-gluconate, 10 mM NaCl, 0.75 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; bathing solution (BS) contained 20 mM CaCl2, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1 mM Tris. ‘40 ascorbate in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 gluconate in PS’. ‘40 gluconate in PS+1 mM exogenous AA’: PS salines were the same as in ‘40 gluconate in PS’; BS contained 20 mM CaCl2, 1 mM l-ascorbic acid, 0.1 mM NaCl, pH 6.0 adjusted by 2 mM MES/1.5 mM Tris. ‘40 ascorbate, 1 mM A9C in PS’: PS contained 40 mM NaOH, 40 mM l-ascorbic acid, 1 mM anthracene-9-carboxylic acid, 0.75 mM BAPTA, 0.3 mM CaCl2 (100 nM Ca2+), pH adjusted by 10 mM Tris; BS contained the same salines as in ‘40 ascorbate in PS’. Holding voltages were corrected by the JPCalc command in Clampex 10.6. (B) Mean ±SE current–voltage relationships (I–V curves) measured in protoplasts patch-clamped using different PSs (n=6–10). PS and BS compositions are shown in (A). Organic anions, including malate–, oxalate–, and citrate–, can move through the plant plasma membranes, including those of Arabidopsis roots, but the Asc– permeability has not been tested so far (Diatloff et al., 2004; Gruber et al., 2010; Hedrich, 2012). To assess Asc– efflux currents, gluconate– was substituted with Asc– in PS, and the test with an anion channel blocker, anthracene-9-carboxylic acid (A9C), was carried out (Fig. 5). Negative inwardly directed currents, which are responsible for anion efflux, were recorded and analysed. When a pipette was filled with 50 mM gluconate– (‘Gluconate pipette’ in Fig. 5A), a small inwardly directed current was observed, which corresponded to Ca2+ influx through constitutive Ca2+-permeable non-selective cation channels (Demidchik et al., 2002). Patch-clamp studies have demonstrated that gluconate appears to be ‘virtually impermeant’ to anion channels (Alvarez-Leefmans and Russell, 1990). In this respect, addition of Asc– instead of gluconate– to PS should reveal the Asc– conductance. Indeed, the addition of 40 mM Asc– in PS resulted in the appearance of a large inwardly directed current, which was three times larger than the current measured using the ‘Gluconate pipette’ (Fig. 5). The Asc– efflux current was completely blocked by the addition of 1 mM A9C to the PS (Fig. 5). A9C also caused a shift of the current–voltage curve in the direction of the reversal potential for Ca2+ (calculated reversal potentials were as follows: ECl= –33.5; ECa=44.2; ENa= –156.5). Thus, these data provided evidence for a relatively good Asc– permeability through the plasma membrane of A. thaliana root cells. It should be noted that the focus of this study was not on the analysis of ascorbate-permeable ion channels. Therefore, further research is required to investigate the biophysical properties and molecular nature of root Asc– efflux conductances. Testing ascorbate release: EPR spectroscopy of root Asc·– in normal and NaCl-stressed plants EPR spectroscopy techniques were applied to assess Asc– efflux qualitatively (Fig. 6). This approach does not show AA release per se but it provides information about the occurrence of its oxidized form, Asc·–, in the apoplast of intact roots. In this case, NaCl (250 mM) was used as an inducer of AA release, while A9C (1 mM) was applied as an inhibitor of AA efflux. Fig. 6. View largeDownload slide The effect of salt stress on the generation of ascorbyl radicals (Asc·–) in intact Arabidopsis thaliana roots studied with EPR spectroscopy. (A) Typical Asc·– spectra in the solution containing 1 mM l-ascorbic acid (‘1 mM AA’), the same solution with 10 mM H2O2 (‘1 mM AA, 10 mM H2O2’), and in root exudates from untreated roots [‘Roots’, corresponding to ‘Control’ in (B)], roots treated by 250 mM NaCl (‘Roots, 250 mM NaCl’), and roots treated with the same NaCl concentration in the presence of 1 mM anthracene-9-carboxylic acid (‘Roots, 250 mM NaCl, 1 mM A9C’) or reduced glutathione (‘Roots, 250 mM NaCl, 1 mM GSH’). (B, C) Mean ±SD EPR signal intensities (Asc·– spectra; n=6–9) obtained in the experimental conditions indicated in (A). Note the different scale on the x-axis in (C) and (B). All experiments were carried out using buffer solution (BS) containing 10 mM Tris (titrated by MES to pH 8.0; Sigma Ultra). Statistical analyses: ANOVA, comparison of control and NaCl-treated roots (B; ***P<0.0001), NaCl-treated roots and NaCl-treated roots with addition of A9C (B; ***P<0.001), cell-free 1 mM AA and 1 mM AA with 10 mM H2O2 (C; ***P<0.0001). Data which were compared are indicated as (a). Fig. 6. View largeDownload slide The effect of salt stress on the generation of ascorbyl radicals (Asc·–) in intact Arabidopsis thaliana roots studied with EPR spectroscopy. (A) Typical Asc·– spectra in the solution containing 1 mM l-ascorbic acid (‘1 mM AA’), the same solution with 10 mM H2O2 (‘1 mM AA, 10 mM H2O2’), and in root exudates from untreated roots [‘Roots’, corresponding to ‘Control’ in (B)], roots treated by 250 mM NaCl (‘Roots, 250 mM NaCl’), and roots treated with the same NaCl concentration in the presence of 1 mM anthracene-9-carboxylic acid (‘Roots, 250 mM NaCl, 1 mM A9C’) or reduced glutathione (‘Roots, 250 mM NaCl, 1 mM GSH’). (B, C) Mean ±SD EPR signal intensities (Asc·– spectra; n=6–9) obtained in the experimental conditions indicated in (A). Note the different scale on the x-axis in (C) and (B). All experiments were carried out using buffer solution (BS) containing 10 mM Tris (titrated by MES to pH 8.0; Sigma Ultra). Statistical analyses: ANOVA, comparison of control and NaCl-treated roots (B; ***P<0.0001), NaCl-treated roots and NaCl-treated roots with addition of A9C (B; ***P<0.001), cell-free 1 mM AA and 1 mM AA with 10 mM H2O2 (C; ***P<0.0001). Data which were compared are indicated as (a). EPR spectroscopy-based measurements of Asc·– in intact A. thaliana roots have recently been developed in our laboratories (Sosan et al., 2016). We have also analysed the potential of other sensitive techniques, such as measuring the radioactively labelled Asc– efflux (Parsons and Fry, 2010) and electrospray ionization MS (Grillet et al., 2014); however, these techniques had significant technical limitations in studying intact roots. In normal conditions, a constitutive Asc·– level was detected in A. thaliana roots (Fig. 6), suggesting that some Asc· is present in the apoplast. The characteristic ascorbyl radical EPR doublet (Laroff et al., 1972; Halliwell and Gutteridge, 2015) was comparable with those that were generated in buffer solution containing 1 mM AA (pH 8.0). Previously, Parsons and Fry (2010) have demonstrated that AA leaks from plant cells stressed by H2O2 for a few seconds. Here, treatment of roots during 3 min by 250 mM NaCl increased Asc·– peak intensity up to 3–4 times. Longer treatment times did not increase Asc·– signal intensity, although, in some cases, it started to decline after treatment for 3 min. The Asc·– signal was sensitive to reduced glutathione in both control and NaCl-treated roots. Reduced glutathione was also effective in vitro, quenching the Asc·– signal measured in solution containing 1 mM AA and 10 mM H2O2 (Fig. 6). Addition of the anion channel blocker, A9C (1 mM), which inhibited Asc– efflux currents in root protoplasts (Fig. 5), together with NaCl, caused a decrease of the NaCl-induced Asc·– signal. Overall, these data demonstrated that the amount of Asc·– increases in the apoplastic space after treatment by NaCl in an A9C-dependent manner. This is indicative of Asc·– release in response to the depolarizing action caused by NaCl (as the counterion for K+). Discussion The data presented here demonstrate that AA, starting from 30 µM, induces elevation of [Ca2+]cyt. in intact Arabidopsis roots. The shape of AA-induced [Ca2+]cyt. transients was similar to that of a mixture AA and Cu2+, which is capable of generating HO·, although the magnitudes of AA-induced [Ca2+]cyt. peaks were smaller (Fig. 1). The overall reaction of [Ca2+]cyt. to AA closely resembles the reaction to ADP (Demidchik et al., 2011) and some phytohormones (Straltsova et al., 2015), which potentially act via stimulation of ROS/HO· production by NADPH oxidase (Demidchik and Shabala, 2018). The peak [Ca2+]cyt. induced by physiological apoplastic AA concentrations (10 µM to 3 mM; Smirnoff and Wheeler, 2000) was reproducible and statistically significant, although it did not exceed 100–150 nM. Such a moderate [Ca2+]cyt. transient is supposedly a signal for finely tuned adjustment of cell metabolism and gene expression. Interestingly, long-term exposure to 1 mM AA (15 h and 40 h; 1 mM AA) moderately increased PCD symptoms in root cells but did not cause complete death of the root system even after 40 h exposure (Fig. 4). Potentially, this can also have a regulatory function. Here the application of AA alongside the mixture of copper and iron chelators prevented AA-induced PCD symptoms, clearly demonstrating the role of these metals in AA-induced PCD. Intriguingly, PCD symptoms evoked by AA were more often observed in trichoblasts than in atrichoblasts, coinciding with the reported higher density of ROS-induced Ca2+ influx current in root hair protoplasts (Demidchik et al., 2003). Hypothetically, AA levels >3 mM, which caused high [Ca2+]cyt. peaks (Fig. 1), can occur in the apoplast, when cells collapse and an intracellular medium and organelles containing high AA levels are excreted. Plant leaf intracellular AA levels vary from 10 mM to 75 mM, while roots accumulate up to 5–10 mM AA; therefore, a lot of ascorbate can potentially be released locally (Smirnoff and Wheeler, 2000; Kawa et al., 2016; Alscher and Hess, 2017). AA levels >10 mM triggered very strong Ca2+ influx according to the dose–response curve presented in Fig. 1. Release of AA can be caused by mechanical injury (wounding stress) and/or PCD. Inducing PCD itself (Fig. 4), AA may be a trigger of self-release, with a further induction of Ca2+ signals. This mechanism can hypothetically play a role in systemic signal transduction stimulating propagation of Ca2+ signals in the case of pathogen attack or wounding stress. Removal of the cell wall abolished the AA-induced activation of Ca2+ signals (Fig. 3), demonstrating that factors needed for this activation reside in the cell wall. Addition of copper or iron chelators halved peak [Ca2+]cyt. transients, while their combined application fully suppressed them (Fig. 3). Taken together, these data indicate that AA effects on [Ca2+]cyt. critically depend on the cell wall Cu+/2+ and Fe2+/3+. These metals are abundant in the apoplast, where they form one of the biggest pools in plants (Sattelmacher, 2001; Printz et al., 2016). In the cell wall, Cu+/2+ and Fe2+/3+ can bind to negatively charged residues of polysaccharides and other organic substances, such as pectin, hemicellulose, and lignin (Fry et al., 2002; Fry, 2004; Wertz et al., 2017). Normally, the apoplastic activities of ionic Cu+/2+ and Fe2+/3+ are very low, while their catalytic activities are maintained at a relatively high level and increase under stress conditions, such as drought (Becana and Klucas, 1992; Moran et al., 1994, 1997). Grillet et al. (2014) have recently shown that AA efflux from root cells is critically important for reduction and mobilization of apoplastic iron and that it is a necessary step for iron acquisition in Pisum sativum. Moreover, Guo et al. (2017) have demonstrated that AA efflux alleviated iron deficiency in an abscisic acid (ABA)-dependent manner. These authors have speculated that AA can stimulate the ABA biosynthesis regulating plant adaptation to iron deficiency. The data presented here demonstrate that AA in the presence of cell wall-bound transition metals can induce Ca2+ signals, which are also known as major modulators of ABA-mediated processes. Here, a very wide range of copper and iron concentrations was tested individually and in combination with AA. Application of CuCl2 at a level higher than 1 µM caused significant stimulation of the AA-induced [Ca2+]cyt. elevation, while addition of Fe(II)-EDTA showed similar effects only at levels >1 mM. This difference can be explained by much higher copper reactivity in catalysis of Fenton-like reactions compared with iron (Halliwell and Gutteridge, 2015) as well as better solubility of copper salts as compared with Fe(II/III)-EDTA complexes. Nevertheless, iron chelators were effective in aequorin luminometry, suggesting that catalytically active ‘bio-available’ iron, similar to copper, is important for induction of Ca2+ signals by exogenous AA. In animal cells, AA efflux is dominated by volume-sensitive and Ca2+-dependent anion channels, but some other mechanisms can also be involved, including gap junction hemichannels, exocytosis of secretory vesicles, and potentially homo- and hetero-exchange systems (Corti et al., 2010). In this study, addition of AA to a patch-clamp pipette instead of gluconate evoked large inward currents, which were sensitive to 1 mM A9C (a blocker of anion channels; Fig. 5). Ascorbate efflux currents showed rapid activation kinetics in response to voltage steps, similar to R-type anionic conductances reported in a number of plant preparations (Diatloff et al., 2004; Roberts, 2006; Kollist et al., 2011; Hedrich, 2012). Ascorbate currents were not previously measured in plant protoplasts, but they can be compared with currents mediated by citrate, malate, and other organic anions (Gruber et al., 2010). Among the major classes of plant anion channels (encoded by ALMT, CLC, and SLAC), members of the ‘aluminium-activated malate transporter’ (ALMT) gene family are the best candidates for mediating ascorbate efflux. These channels are responsible for transport of large organic anions, such as malate, oxalate, and citrate. They have been discovered as A9C-sensitive systems releasing malate and other organic acids to chelate soil Al3+ (Ryan et al., 1995, 1997). Here, we provided the evidence that these channels can also be involved in another important process, which is release of AA to the extracellular space. AA does not form complexes with Al3+; therefore, this reaction should have a different evolutionary route from Al tolerance. Current–voltage curves of ascorbate efflux conductance (Fig. 5) were very similar to those that were reported in Al-free conditions (control) for malate efflux channels from different preparations, including Arabidopsis thaliana ALMT1 expressed in Xenopus oocytes (Hoekenga et al., 2006; Piñeros et al., 2008; Gruber et al., 2010) or Triticum aestivum ALMT1 expressed in tobacco culture cells (Zhang et al., 2008). Using EPR spectroscopy allowed us to resolve monodehydroascorbate reductase (MDHAR) spectra in control plants and the 9AC-sensitive burst of the MDHAR spectrum intensity induced by the addition of NaCl (3 min, 250 mM NaCl; Fig. 6). After 10 min and 15 min exposure to NaCl, a 2- to 3-fold decrease in MDHAR spectrum intensity was noted. We may speculate that by that time released AA was oxidized by mobilized iron and copper. AA release from plant cells has been reported several times in a variety of circumstances in the plant literature (Luwe et al., 1993; Luwe and Heber, 1995; Parsons and Fry, 2010; Grillet et al., 2014). Luwe and Heber (1995) measured AA content in control and O3-treated leaves of spinach. They found that leaf AA content increased after O3 treatment from 1–2 mM to 2–3 mM. Grillet et al. (2014) found that embryos of pea and A. thaliana excrete AA for iron reduction and uptake. Parsons and Fry (2010) tested the effect of 1–10 mM H2O2 on AA efflux in rose and A. thaliana suspension cultures in relation to mechanisms of antioxidant defence in cell cultures. They showed that H2O2 induces loss of 20% of cell AA in a few minutes. Peak AA efflux occurred in the first 100 s of H2O2 application. These authors also measured H2O2-induсed electrolyte leakage, which accompanied AA release, but was not related to the loss of cell integrity and viability. According to the literature and data shown here, plant plasma membranes have good permeability to AA. Therefore, ascorbate can function as a counterion during ROS-induced K+ efflux under oxidative stress conditions (Demidchik et al., 2014; Demidchik, 2015). This partially explains the very fast kinetics of AA release found by Parsons and Fry (2010). Potassium is the most abundant ion in the cytosol and a major species that leaves a plant cell in response to key stresses, such as NaCl, drought, pathogens, hypoxia, O3, UV, xenobiotics, heavy metals, etc. (Shabala et al., 2006; Demidchik et al., 2014). The potassium efflux channel GORK is activated by ROS (H2O2 and HO·), which are generated in response to stresses (Demidchik et al., 2010, 2018). This activation normally results in stress-induced leakage of K+ accompanied by leakage of anions. Salt stress (50 mM NaCl) causes the decrease of cytosolic K+ from 70–80 mM to 20–30 mM (Shabala et al., 2006). An equal amount of the anions should leave the cell to equilibrate changes in electric charge and ionic balance caused by K+ loss. A number of anions can potentially function as counterions, and ascorbate could be one of them. If GORK-mediated K+ efflux and ascorbate release are related processes, this can have important consequences for plants, including (i) additional stimulation of the ROS-induced K+ efflux via HO· production by released AA; (ii) involvement of GORK in systemic ROS-dependent responses and signal transduction; (iii) loss of cytosol antioxidant capacity in response to stresses that are accompanied by K+ leakage; and (iv) ascorbate can participate in the adjustment of plant metabolism under stress. These opportunities and other implications of AA release are summarized in Fig. 7. We hypothesize that ascorbate efflux can be part of the plant cell response to stresses initiating redox-dependent Ca2+ signalling and K+ efflux at the plant plasma membrane (Fig. 7). Ascorbate release can also be a reason for a moderate increase in PCD, which can have an adaptive role or originate the long-distance signalling. Additionally, AA release can be an iron-mobilizing reaction, which is necessary for iron uptake by root cells, and AA, when released, may stimulate cell growth via the ROS–Ca2+ hub (Fig. 7; Demidchik and Shabala, 2018). Fig. 7. View largeDownload slide Hypothetical scheme of plant plasma membrane signalling and transport reactions involving l-ascorbate. Stresses or developmental signals can trigger NADPH oxidase activity, leading to production of ROS, such as O2·–, H2O2, and HO·, which activate Ca2+ influx and K+ efflux channels (see Demidchik and Shabala, 2018; Demidchik et al., 2003, 2010, 2018, for details). Stress-induced depolarization results in K+ efflux, which can potentially be accompanied by Asc– (ascorbate-) efflux. In this case, ascorbate– is transported as the counterion to maintain cell ionic and charge balance. Ascorbate accumulation in the apoplast results in reduction of transition metals, such as copper and iron, leading to generation of HO·, which can cause activation of Ca2+ influx channels in the same cell and neighbouring cells. Overall, ascorbate efflux stimulates the self-amplifying ROS–Ca2+ hub, which is based on activation of NADPH oxidase by cytosolic Ca2+ and stimulation of Ca2+ influx by transition metal/ascorbate-catalysed conversion of O2·–/H2O2 to HO·, which, in turn, activates Ca2+ influx channels. Moreover, ascorbate efflux may cause iron and copper reduction for nutritional needs. Please note that some other transporters, such as HO·-activated non-specific ion channels, which are permeable to both cation and anions (Zepeda-Jazo et al., 2011) and hypothetically mediate Ca2+ influx and AA efflux, can also be involved in AA signalling reactions. Fig. 7. View largeDownload slide Hypothetical scheme of plant plasma membrane signalling and transport reactions involving l-ascorbate. Stresses or developmental signals can trigger NADPH oxidase activity, leading to production of ROS, such as O2·–, H2O2, and HO·, which activate Ca2+ influx and K+ efflux channels (see Demidchik and Shabala, 2018; Demidchik et al., 2003, 2010, 2018, for details). Stress-induced depolarization results in K+ efflux, which can potentially be accompanied by Asc– (ascorbate-) efflux. In this case, ascorbate– is transported as the counterion to maintain cell ionic and charge balance. Ascorbate accumulation in the apoplast results in reduction of transition metals, such as copper and iron, leading to generation of HO·, which can cause activation of Ca2+ influx channels in the same cell and neighbouring cells. Overall, ascorbate efflux stimulates the self-amplifying ROS–Ca2+ hub, which is based on activation of NADPH oxidase by cytosolic Ca2+ and stimulation of Ca2+ influx by transition metal/ascorbate-catalysed conversion of O2·–/H2O2 to HO·, which, in turn, activates Ca2+ influx channels. Moreover, ascorbate efflux may cause iron and copper reduction for nutritional needs. Please note that some other transporters, such as HO·-activated non-specific ion channels, which are permeable to both cation and anions (Zepeda-Jazo et al., 2011) and hypothetically mediate Ca2+ influx and AA efflux, can also be involved in AA signalling reactions. Acknowledgements Financial support from the Russian Science Foundation (grant #15-14-30008 to VD) is gratefully acknowledged. 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