Drought-Enhanced Xylem Sap Sulfate Closes Stomata by Affecting ALMT12 and Guard Cell ABA Synthesis

Reduced unloading and enhanced loading of sulfate increased xylem sap sulfate during early drought, which affects the opening of the ALMT12 channel and induces guard cell expression of the key step in ABA synthesis, NCED3.

Abstract

Water limitation of plants causes stomatal closure to prevent water loss by transpiration. For this purpose, progressing soil water deficit is communicated from roots to shoots. Abscisic acid (ABA) is the key signal in stress-induced stomatal closure, but ABA as an early xylem-delivered signal is still a matter of debate. In this study, poplar plants (Populus × canescens) were exposed to water stress to investigate xylem sap sulfate and ABA, stomatal conductance, and sulfate transporter (SULTR) expression. In addition, stomatal behavior and expression of ABA receptors, drought-responsive genes, transcription factors, and NCED3 were studied after feeding sulfate and ABA to detached poplar leaves and epidermal peels of Arabidopsis (Arabidopsis thaliana). The results show that increased xylem sap sulfate is achieved upon drought by reduced xylem unloading by PtaSULTR3;3a and PtaSULTR1;1, and by enhanced loading from parenchyma cells into the xylem via PtaALMT3b. Sulfate application caused stomatal closure in excised leaves and peeled epidermis. In the loss of sulfate-channel function mutant, Atalmt12, sulfate-triggered stomatal closure was impaired. The QUAC1/ALMT12 anion channel heterologous expressed in oocytes was gated open by extracellular sulfate. Sulfate up-regulated the expression of NCED3, a key step of ABA synthesis, in guard cells. In conclusion, xylem-derived sulfate seems to be a chemical signal of drought that induces stomatal closure via QUAC1/ALMT12 and/or guard cell ABA synthesis.

Leaves of vascular plants are equipped with stomata that actively control the exchange of CO₂, O₂, and water vapor between the leaf interior and the atmosphere. Drought induces stomatal closure via the release of K⁺ and anions from the guard cells of the stomata (Hedrich, 2012; Kollist et al., 2014). In this process the phytohormone abscisic acid (ABA), a drought-induced messenger, addresses the anion release via S-type/slow activated anion channel (SLAC1), SLAC1 Homolog 3 (SLAH3), and R-type/quick-activating anion channel1 (QUAC1/ALMT12) anion channels (Imes et al., 2013). Perception of ABA by the ABA receptors PYR/PYL/RCAR results in PP2C phosphatase (ABI1) inactivation. In turn, Open Stomata1 (OST1), a Ser/Thr-protein kinase, and calcium-dependent protein kinases released from ABI1 inactivation, phosphorylate and activate the anion channels SLAC1 and QUAC1/ALMT12. SLAC1 conducts the release of Cl⁻ and NO₃⁻, and QUAC1/ALMT12 the release of several anions such as Cl⁻, NO₃⁻, and SO₄²⁻ (Hedrich and Marten, 1993; Frachisse et al., 1999). The release of these anions depolarizes the plasma membrane of guard cells giving rise to voltage activation of the K⁺ release channel GORK (Ache et al., 2000) and deactivation of K_in channels (Hedrich, 2012). Inhibited influx of potassium along with enhanced efflux of potassium and associated osmotic water cause guard cell turgor loss and stomatal closure.

Roots perceive restricted soil water availability and communicate this environmental constraint as a stress signal toward the shoot, most likely via the xylem. Chemical (Schachtman and Goodger, 2008), hydraulic (Christmann et al., 2007, 2013), and electrical (Grams et al., 2007; Gil et al., 2008) signaling have been considered in drought stress signal transduction. Drought-related chemical signals, like ABA, are expected to be synthesized in the roots, then transported via the xylem to the leaves to induce stomatal closure (Hartung et al., 2002; Goodger and Schachtman, 2010; Wilkinson and Davies, 2010; Osakabe et al., 2014). However, grafting experiments with ABA-deficient tomato plants indicated the independency of stomatal closure on root-derived ABA (Holbrook et al., 2002). Besides ABA, also plant s-metabolism responds to drought (García-Mata and Lamattina, 2010; Chan et al., 2013; Misra et al., 2015). Recent experiments with maize (Zea mays; Ernst et al., 2010; Goodger and Schachtman, 2010) and flowering hop (Humulus lupulus; Korovetska et al., 2014) provided the first evidence, to our knowledge, that xylem-transported sulfate could be a root-to-shoot-transported chemical signal of water deficiency. Ernst et al. (2010) additionally reported that the significant decline in stomatal conductance was not accompanied by leaf hydraulic changes. Therefore, they concluded that at early stages of water deprivation, stomata respond to chemical rather than hydraulic signals from the roots. Enhanced sulfate contents in the xylem sap can originate from increased sulfate uptake by the roots (Ernst et al., 2010), remobilization of sulfate from storage pools inside the roots, or from diminished sulfate unloading from the xylem sap along the transport path. Thus, irrespective of the mechanism involved, sulfate transporters (SULTRs) provide for SO₄²⁻ homeostasis and stimulus-dependent sulfate concentration changes in the xylem sap (Takahashi et al., 2011; Rennenberg and Herschbach, 2014). Important SULTRs in poplar (Populus × canescens) are (1) PtaSULTR1,2, which is solely expressed in roots and, thus, thought to be responsible for sulfate uptake from the soil; (2) PtaSULTR1;1 and PtaSULTR3;3a, which are both abundant in parenchyma cells around xylem vessels of the wood and leaf veins and, thus, responsible for the xylem unloading of sulfate; and (3) SULTRs from group 4, PtaSULTR4;1 and PtaSULTR4;2, which are putative tonoplast localized and, thus, responsible for sulfate release from vacuoles (Kataoka et al., 2004; Dürr et al., 2010). In addition, sulfate permeation by anion channels has to be considered for xylem-loading of sulfate (Frachisse et al., 1999; Roberts, 2006).

Import of sulfate into the chloroplasts coregulates ABA biosynthesis. Sulfate reduction and Cys production in the chloroplasts feed-forward the sulfuration of the molybdenum cofactor (Moco) by Moco sulfurase (ABA3; Mendel, 2013; Cao et al., 2014). In Arabidopsis, oxidation of abscisic aldehyde to abscisic acid by the aldehyde oxidase (AAO3), the last step in ABA synthesis, requires the molybdenum cofactor activated by a Moco sulfurase (ABA3; Nambara and Marion-Poll, 2005). Furthermore, sulfate-promoted Cys synthesis can also act in stomatal movement via the gasotransmitter hydrogen sulfide (H₂S). H₂S is produced in the cytosol in enzymatic desulfuration of l-Cys, by l-Cys desulfurylase, and promotes stomatal closure in an ABA-dependent manner (García-Mata and Lamattina, 2010; Scuffi et al., 2014). Thus, it has been hypothesized that increased xylem sap delivered sulfate during drought can either trigger ABA synthesis in guard cells or/and induce stomatal closure in an ABA-dependent manner.

This study tested whether sulfate is an early xylem-delivered signal communicating drought stress to the shoot. Sulfate and ABA concentration changes in the xylem sap were monitored and related to stomatal conductance. Expression of SULTRs and putative SO₄²⁻-permeable ALMT12/QUAC1 anion channels were investigated to elucidate the origin of sulfate in the xylem sap during drought stress. Sulfate and ABA feeding experiments were performed to test their effects on stomatal conductance and to identify molecular targets of xylem-delivered sulfate. The loss of sulfate-channel function mutant, Atalmt12, was used to analyze sulfate impact on the ALMT12/QUAC1 anion channel. Effects on anion gating of the guard-cell-specific AtALMT12 channel were investigated by heterologous expression in Xenopus laevis oocytes. Furthermore, sulfate responses on guard-cell-specific gene expression of ABA receptors, drought-responsive genes, and transcription factors, as well as NCED3 (a key gene of ABA biosynthesis) were studied.

RESULTS

Sulfate, But Not ABA, Increases in the Xylem Sap during Early Drought

When poplar plants grown in sand culture were exposed to progressing water deprivation, sulfate starts to accumulate in the xylem sap after 48 h of drought. The increase in xylem sap sulfate preceded both ABA accumulation in the xylem sap that increased after 64 h of drought and the decline in stem water potential that increased after 72 h of drought (Fig. 1). Stomatal conductance, however, declined after 48 h of water deprivation, when xylem sap sulfate concentrations almost doubled (Fig. 1C), but ABA concentrations were not affected (Fig. 1D). Important, at this time, changes in water potential were not observed (Fig. 1B). Drought-induced xylem sap sulfate was independent of ABA biosynthesis, because poplar RNAi lines down-regulated in the expression of AAO and ABA genes also showed drought-enhanced xylem sap sulfate (Supplemental Fig. S1). The pH value of the xylem sap slightly decreased to pH 5.75 ± 0.33 and pH 5.65 ± 0.20 after 48 h and 56 h of water deprivation, respectively, compared to pH 5.99 ± 0.22 at the onset of the drought stimulus. In contrast to xylem sap sulfate that consistently increased over time, xylem sap phosphate increased after 72 h of drought when twig water potential increased (Fig. 2A) and stomatal conductance was ∼1/10th compared to the control. Malate that might also be a signal for stomatal closure (Hedrich and Marten, 1993; Hedrich et al., 1994) increased after 56 h of drought but declined thereafter (Fig. 2B). Potassium declined after 48, 56, and 64 h of water deprivation (Fig. 2C).

Figure 1.

Influence of water deprivation on stomatal conductance, stem water potential, and xylem sap composition of poplar. Stomatal conductance of the 10th leaf (A), stem water potential (B), sulfate concentration (C), and ABA concentration (D) of the xylem sap at 0 (control, n = 11), 24 (n = 4), 48 (n = 10–11), 56 (n = 5), 64 (n = 6), and 72 (n = 9) h of water deprivation. The presented values ± sd are relative values compared to the control at 0 h of water deprivation. Data of the controls without water stress (0 h) of both experiments are presented in the Supplemental Table S1. Asterisks indicate statistical significant differences of mean values ± sd (Student’s t test or Mann-Whitney U test) at P < 0.05 compared to the well-watered plants (control, 0 h).

Figure 2.

Influence of water deprivation on phosphate, malate, and potassium in the xylem sap of poplar. Phosphate (A), malate (B), and potassium (C) in the xylem sap of poplar at 0 (control, n = 10–11), 24 (n = 4), 48 (n = 11), 56 (n = 5), 64 (n = 6), and 72 (n = 8) h of water deprivation. Relative values of ion concentrations compared to their levels at 0 h of water deprivation are presented. Data of the controls without water stress (0 h) of both experiments are presented in the Supplemental Table S1. Asterisks indicating statistical significant differences (Student’s t test or Mann-Whitney U test) of mean values ± sd at P < 0.05 compared to the well-watered plants (control, 0 h).

Drought Affects Sulfate Transporter Expression

Changes in the xylem sap sulfate can be attributed to changes in the activity of distinct SULTRs responsible for sulfate uptake into the roots (PtaSULTR1;2; Dürr et al., 2010), unloading of xylem sap sulfate into xylem parenchyma cells (PtaSULTR1;1 and PtaSULTR3;3a; Dürr et al., 2010), and/or remobilization of sulfate stored in the vacuoles of xylem parenchyma cells (PtaSULTR4;1 and PtaSULTR4;2; Dürr et al., 2010). Therefore, SULTR expression was investigated in different root fractions (according to Herschbach et al., 2010) of drought-exposed poplar plants. Long white roots are roots emerging into the soil without developing side roots. Fine roots, which are roots with a diameter of approx. 0.5–1 mm that developed high density of side roots, were the most important root fraction within the root system of poplar (Herschbach et al., 2010). In addition, roots showing secondary growth were expected to be involved in nutrient storage. Expression of PtaSULTR1;2, the only root-specific SULTR expected to be responsible for sulfate uptake (Dürr et al., 2010), showed a temporarily increased expression after 24 h in fine and elongating roots that was not statistically significant, as well as after 48 h of drought in elongating roots (Fig. 3). Thus, the continuously increasing sulfate level in the xylem sap during water deprivation is unlikely to originate from enhanced sulfate uptake especially after prolonged water stress. mRNA levels of PtaSULTR1;1, responsible for xylem unloading (Dürr et al., 2010; Malcheska et al., 2013), significantly declined only after prolonged (72 h) water deprivation, but not in all three root fractions, i.e. white elongating roots, fine roots, and in roots showing secondary growth (Fig. 3). However, transcript levels of the second SULTR responsible for xylem unloading of sulfate, PtaSULTR3;3a (Dürr et al., 2010; Malcheska et al., 2013), declined after 24 h of water deprivation in elongating roots and after 48 h in elongating roots as well as in roots showing secondary growth (Fig. 3). After 56 h of water deprivation, PtaSULTR3;3a declined in all root fractions and thereby indicated reduced xylem unloading of sulfate.

Figure 3.

Transcript abundance of SULTRs in roots and leaves of poplar in response to drought. Relative expression levels of PtaSULTR1;2, PtaSULTR1;1, and PtaSULTR3;3a compared to their mRNA gene copies per μg RNA at 0 h of water deprivation in leaves, elongating roots, fine roots, and roots with secondary growth are presented. Data of controls without water stress (0 h) of both experiments are presented in the Supplemental Table S1. Asterisks indicate statistically significant differences of mean values ± sd (24 h, n = 3–4; 48 h, n = 9–11; 56 h, n = 5; 64 h, n = 6; and 72 h, n = 8–9; Student’s t test or Mann-Whitney U test) at P < 0.05 compared to well-watered poplar (control, 0 h of water deprivation; n = 10–11).

Transcript levels of PtaSULTR3;1b, a homologous SULTR gene of Arabidopsis group 3 SULTRs expressed in the chloroplast (Cao et al., 2013), start to increased significantly after 48 h in fine roots, and in addition, in both elongating roots and roots showing secondary growth after 56 h of water deprivation (Fig. 4). PtaSULTR3;1b transcript levels also increased in leaves after 64 h of water deprivation. Root expression of PtaSULTRs, which are likely localized in the tonoplast (PtaSULTR4;1 and PtaSULTR4;2), was observed for PtaSULTR4;1 that increased transiently but not continuously over time in elongating roots and roots showing secondary growth (Fig. 4). In leaves, increased expression of SULTRs was observed for both PtaSULTR3;1b and PtaSULTR4;2, after 64 and 72 h of drought (Fig. 4).

Figure 4.

Transcript abundance of SULTRs in roots and leaves of poplar in response to drought. Relative expression levels of PtaSULTR3;1b, PtaSULTR4;1, and PtaSULTR4;2 compared to their mRNA gene copies per μg RNA at 0 h of water deprivation in leaves, elongating roots, fine roots, and roots with secondary growth are presented. Data of controls without water stress (0 h) of both experiments are presented in the Supplemental Table S1. Asterisks indicate statistically significant differences of mean values ± sd (24 h, n = 3–4; 48 h, n = 9–11; 56 h, n = 5; 64 h, n = 6; and 72 h, n = 9; Student’s t test or Mann-Whitney U test) at P < 0.05 compared to well-watered poplar (control, 0 h of water deprivation; n = 10–11).

Expression of QUAC1/ALMT12-Type Sulfate Channel Is Drought Sensitive

Sulfate uptake by SULTRs depends on the proton-motive force established by the plasma membrane H⁺-ATPase (Lass and Ullrich-Eberius, 1984; Hawkesford et al., 1993). Thus, SULTRs can only transport sulfate from the apoplast into xylem parenchyma and/or guard cells. Anion channels (Frachisse et al., 1999; Roberts, 2006) must therefore perform efflux of sulfate from xylem parenchyma cells. In poplar, three putative ALMT channels that grouped with AtALMT12 in clade 3 of the ALMT1 anion channels/transporters family (and probably are able to gate sulfate) were identified (Barbier-Brygoo et al., 2011; Dreyer et al., 2012). In elongating roots, PtaALMT3a expression increased after 56 and 64 h of drought but recovered to the control mRNA level thereafter. PtaALMT3b, however, increased in elongating roots even after 24 h of water deprivation and remained high throughout the drought treatment (Fig. 5). Also, in fine roots, PtaALMT3b expression was higher than in the control, i.e. at 0 h of drought stress. As fine root biomass comprises more than 50% of the root’s biomass (Herschbach et al., 2010), PtaALMT3b seems to be involved in the sulfate release from xylem parenchyma cells upon drought and contributes to enhanced sulfate levels in xylem sap. In leaves, expression of PtaALMT3a continuously decreased after 48 h of drought, but PtaALMT3b was enhanced after 72 h of water deprivation (Fig. 5).

Figure 5.

Expression of ALMT channels. Transcript abundance of PtaALMT3a and PtaALMT3b that grouped with AtALMT12 in the clade 3 (Dreyer et al., 2012) and of the housekeeping gene PtaEF1B (Wildhagen et al., 2010) in leaves, elongating roots, fine roots, and roots with secondary growth. Data presented are relative mRNA abundance of gene copies per μg RNA of the respective gene compared to the control level at 0 h of water deprivation. Data of controls without water stress (0 h) of both experiments are presented in the Supplemental Table S1. Asterisks indicate statistically significant differences of mean values ± sd (24 h, n = 3–4; 48 h, n = 9–11; 56 h, n = 5; 64 h, n = 5–6; and 72 h, n = 9; Student’s t test or Mann-Whitney U test) at P < 0.05 compared to well-watered poplar (control, 0 h of water deprivation; n = 10–11).

Xylem-Delivered Apoplastic Sulfate Induces Stomatal Closure

To test whether sulfate directly affects stomatal movement, sulfate, ABA, and combinations of both were fed to detached poplar leaves and stomatal conductance was measured. Sulfate applied together with Mg at 2 mm (Fig. 6; Supplemental Fig. S2) significantly reduced stomatal conductance compared to the control [1 mm MgCl₂ plus 60 μm Mg(NO₃)₂]. When 3 µm ABA was fed, a similar decline in stomatal conductance was observed (Fig. 6). This effect was even stronger when 3 μm ABA was applied in combination with 2 mm sulfate (Fig. 6), suggesting that sulfate could serve as a positive regulator of ABA.

Figure 6.

Stomatal conductance of detached poplar leaves fed with sulfate and ABA via the petiole. The feeding solution containing 1 mm MgCl₂ plus 0.06 mm Mg(NO₃)₂ (pH 5.5) was taken as a control. Treatments were exposed to different concentrations of magnesium sulfate; 2 mm MgSO₄ plus 3 μm ABA; 3 μm ABA, 2 mm MgSO₄ plus 0.3 μm ABA; and 0.3 μm ABA via the petiole (n = 4–6). Mean values ± sd presented showed relative stomatal conductance after 60 min of incubation that were calculated in comparison to the stomatal conductance determined during preincubation when stable values are reached. Stomatal conductance was determined by gas exchange measurements and mean ± sd values of stomatal conductance (mmol m⁻² s⁻¹) are presented in the Supplemental Table S1. Asterisks indicate significant differences compared to the control feeding (*P < 0.05; **P < 0.01; ***P < 0.001; Student’s t test). The sulfate effect was independently observed if sulfate was either added to the control solution (Supplemental Fig. S2) or applied solely as MgSO₄. Capital letter (A) indicates statistical significant differences of mean values ± sd at P < 0.05 (Student’s t test) between two treatments as indicated. Stomatal conductance of leaves fed with the solution consisting of 1 mm MgCl₂ plus 0.06 mm Mg(NO₃)₂ ranged from 76 to 100 mmol m⁻² s⁻¹.

To confirm the impact of sulfate on stomatal conductance in poplar, similar feeding experiments were performed with Arabidopsis. When sulfate was fed to detached Arabidopsis leaves via the petiole, relative transpiration dropped within 40 min by ∼35% (Fig. 7). Thus, sulfate does not only trigger decreased stomatal conductance in the perennial woody poplar, but also in the annual herbaceous Arabidopsis.

Figure 7.

Transpirational water loss was determined by gas exchange measurements of detached leaves of Arabidopsis (Col-0). Partial stomatal closure was provoked by feeding MgSO₄ (final concentration 10 mm) via the petiole. The control feeding solution contained 0.06 mm MgNO₃ plus 1 mm MgCl₂. The gap in the x axes resulted from different time spans that individual plants needed to gain steady-state transpiration rates (n = 6; ± sd). Values were related to leaf area and normalized to the time point of MgSO₄ feeding (t = 0).

In search for a molecular explanation of the sulfate-induced decrease of stomatal conductance, the effect of apoplastic sulfate on the aperture of stomata in epidermis sections of Arabidopsis leaves was quantified. First, the interactive effect of sulfate on the action of ABA shown by Ernst el al. (2010) was confirmed using the identical experimental setup, but Arabidopsis epidermal peels were used instead of epidermal peels of maize (Fig. 8A). This experimental setup included a high potassium concentration (50 mm) in the incubation solution to ensure opening of stomata after peeling. Next, we showed that sulfate in the absence of ABA closed the stomata when this high potassium chloride concentration was decreased to zero (Fig. 8B) or to a potassium concentration found in the apoplastic space of leaf cells (10 mm potassium; Supplemental Fig. S3; Long and Widders, 1990). Without addition of artificially high potassium chloride in the experimental setup, sulfate induced stomatal closure in a dose-dependent manner (Fig. 8C). Remarkably, stomatal closure was significantly induced by apoplastic application of 2 mm sulfate, a concentration found in the xylem sap of drought-stressed maize plants (Ernst et al., 2010). These data demonstrate that sulfate fed via the petiole induce stomatal closure and suggest that drought-induced increase of sulfate in the xylem contributes to stomatal closure and may act as a long-distance transport signal from the root to the shoot (Fig. 1). These results do not exclude a role of other metabolites (e.g. malate, ABA) as additional root-to-shoot signals.

Figure 8.

Sulfate-induced stomatal closure in peeled epidermis of Arabidopsis. Peeled epidermal layers with intact stomata were prepared from 5-week-old Arabidopsis wild-type and Atalmt12 mutants as described in Ernst et al. (2010). A, Arabidopsis epidermal layers were floated on an artificial buffer (50 mm KCl and 10 mm MES, pH 5.5) with stomata facing the ambient air for 2 h. Subsequently, epidermal layers were exposed to increasing concentrations of ABA (0–0.3 μm) in the absence (white) or presence of 15 mm sulfate (black, MgSO₄) dissolved in potassium-containing solution (50 mm KCl and 10 mm MES, pH 5.3). B, Stomatal aperture of isolated epidermal cell layers in absence (white) or presence of 15 mm sulfate (black, MgSO₄) dissolved in water or water supplemented with 50 mm potassium and/or 10 mm MES at pH 5.3. C, Impact of different sulfate concentrations (0–15 mm MgSO₄) dissolved in potassium-buffered solution or water on stomatal aperture. D, Application of sulfate (15 mm MgSO₄) in water or sulfate and ABA (0.3 μm) in potassium-buffered solution to wild-type (gray) or Atalmt12 mutants (black-striped). Data represent mean values ± sd of five to seven epidermal layers (number of counted stomata for each epidermal layer = 10, number of analyzed stomata = 50–60). Statistical differences between treatments were analyzed by ANOVA on ranks followed by Student-Newman-Keuls and are marked with different letters (n = 5–7).

Sulfate-Induced Stomatal Closure Involves QUAC1/ALMT12

In the search for the molecular target of sulfate-induced stomatal closure, the responses of stomata from wild-type Arabidopsis and the loss-of-ALMT12 function mutant (Atalmt12) to sulfate were analyzed. Whereas stomatal aperture of wild-type Arabidopsis significantly decreased after sulfate application, stomata lacking ALMT12 failed to close in response to sulfate. Furthermore, Atalmt12 stomata were insensitive to ABA and a combination of ABA and sulfate (Fig. 8D). The insensitivity toward ABA is a characteristic feature of Atalmt12 (Meyer et al., 2010). To test if the sulfate insensitivity of the Atalmt12 mutant also affects its response to drought stress, the mutant line and wild-type Arabidopsis were subjected to water deprivation. When the gravimetric soil water content (water content per weight dried soil) dropped from ∼2.3 g g⁻¹ (well-watered) to ∼0.7 g g⁻¹ (drought-stressed; Supplemental Table S2), a significant reduction in stomatal conductance and transpiration was observed in wild-type Arabidopsis, but not in the mutant line (Fig. 9; Supplemental Fig. S4). Interestingly, sulfate fed to detached leaves of the mutant line Atalmt12 did not affect stomatal conductance (Fig. 9C).

Figure 9.

Drought and sulfate effects on stomatal conductance and photosynthesis of wild-type Arabidopsis and the Atalmt12 mutant line. The effect of drought on wild-type Arabidopsis and the mutant Atalmt12 on stomatal conductance (A) and on rate of photosynthesis (B) of leaves still attached. Whole plants of wild-type Arabidopsis (n = 10) and Atalmt12 mutants (n = 11) growing on commercial soil were continuously watered or exposed to drought (wild-type, n = 13; Atalmt12, n = 10) until the gravimetric soil water content reaches ∼0.7 g g⁻¹ for both lines (for biometrical data, see Supplemental Table S2). C, The effect of sulfate on stomatal conductance of detached leaves of wild-type Arabidopsis and the mutant Atalmt12. Detached leaves of wild-type Arabidopsis or the Atalmt12 mutant were exposed to deionized water adjusted to pH 5.5 (by NaOH and/or HCl) for 2 h preincubation and were subsequently exposed either to deionized water (n = 10) or 10 mm MgSO₄ (n = 10). Stomatal conductance was calculated from monitored weight loss after 60, 90, and 120 min, when water loss reaches stable values. Stomatal conductance was calculated mathematically (see Supplemental Materials and Methods). Statistics were prepared with one-way ANOVA Tukey after test for normal distribution with the Kolmogorov-Smirnov and Shapiro-Wilk tests. Different letters indicate significant differences of mean ± sd values between groups at P < 0.05. Data on stomatal conductance of detached leaves of wild-type Arabidopsis and the Atalmt12 mutant exposed to 1 mm MgCl₂ plus 0.06 mm Mg(NO₃)₂ (control feeding solution; pH 5.5 adjusted by NaOH and/or HCl) for 2 h preincubation and subsequently exposed either to the feeding solution (n = 10) or to feeding solution supplemented with 10 mm MgSO₄ (n = 10) are presented as Supplemental Fig. S4.

Sulfate Gates QUAC1/ALMT12 Open

The experiments presented above (Figs. 8D and 9) clearly show that the Atalmt12 mutant is less sensitive to sulfate. Studies showed that QUAC1/ALMT12 represents a sulfate-, nitrate-, and chloride-permeable anion channel with fast activation and deactivation kinetics (rapid-type anion channel, R-type; Hedrich and Marten, 1993; Meyer et al., 2010; Mumm et al., 2013) but only malate was tested in activating QUAC1/ALMT12 by shifting its voltage-dependent open probability to hyperpolarized membrane potentials. Therefore, the response of extracellular sulfate on QUAC1/ALMT12 steady-state conductance was investigated here. QUAC1/ALMT12 was coexpressed in X. laevis oocytes with its activating kinase OST1 (Imes et al., 2013). After 3 d of expression, two-electrode voltage-clamp experiments were conducted in the presence of either chloride- or sulfate-based media. In the presence of sulfate and a stimulating voltage pulse, QUAC1/ALMT12 activated with typical R-type channel fast-activation kinetics (Fig. 10A). Steady-state currents (I_SS) plotted as a function of the applied membrane potential underline the sulfate-dependent induction of anion efflux currents (Fig. 10B). Moreover, the peak currents in sulfate-based buffers exhibited similar amplitudes to those found in malate-based buffers. To elucidate whether the presence of sulfate shifts the voltage-dependent gating properties of QUAC1/ALMT12 to more negative membrane potentials, we inferred the relative open probability (rel. P_O) and described the curves with a Boltzmann equation. Like malate, sulfate shifted the rel. P_O of the anion channel to more negative membrane potentials following Boltzmann characteristics (Fig. 10C). The assumed long-distance signal sulfate thus shifted the half-maximal activation potential (V_1/2) by −50 mV compared to chloride controls.

Figure 10.

Sulfate activates QUAC1/ALMT12 anion channels. A, After 3 d of expression, representative whole-oocyte currents of QUAC1/ALMT12-expressing oocytes were recorded with the two-electrode voltage-clamp technique in the presence of either 10 mm sodium chloride or 10 mm sodium sulfate. In response to voltage jumps from the holding potentials of −200 to −80 mV, anion efflux currents were elicited. B, Steady-state currents (I_SS) recorded with QUAC1/ALMT12-expressing oocytes in either chloride-, sulfate-, or malate-based buffers (10 mm each) were plotted as a function of the applied membrane potential (n = 5, ±sd). C, The inferred rel. P_O was plotted as a function of the oocytes’ membrane potential (n = 4, ±sd). Fitting of the curves with a Boltzmann equation revealed a half-maximal activation potentials (V_1/2) of −68 ± 22 mV in sodium chloride-, −118 ± 34 mV in sodium sulfate-, and −159 ± 4 mV in sodium malate-based buffers.

Sulfate Triggers Gene Expression in Guard Cells in an ABA-Like Manner

Besides sulfate directly affecting stomatal closure, ABA-sulfate interaction could result from SO₄²⁻ feeding-forward ABA production or sensitivity. To test this hypothesis, Arabidopsis leaves were treated either with ABA, sulfate, or both stimuli together and changes in transcript abundance of typical ABA-regulated genes were analyzed in epidermis fraction enriched with guard cells (Geiger et al., 2011; Fig. 11). Two ABA-responsive genes, the PP2C protein phosphatase (HAI1) and ABAR (Bauer et al., 2013), the ABA down-regulated transcription factor MYB60 (Bauer et al., 2013), and the ABA receptors PYL2 and PYL4 (Gonzalez-Guzman et al., 2012), were quantified by qRT-PCR. After ABA or sulfate application, the ABA receptor genes (PYL2 and PYL4) and the ABA down-regulated transcription factor MYB60 were significantly down-regulated showing comparable expression levels with both stimuli as well as in combination (Fig. 11, C to E). The HAI1 and ABAR genes were significantly up-regulated only when feeding ABA and increased as well by sulfate feeding but not significantly. Combined treatment with ABA and sulfate did not further affect gene expression compared to solely ABA application. These responses indicate that both sulfate and ABA act via the same pathway. To test this assumption, we analyzed the expression of NCED3, a key enzyme of the ABA biosynthesis (Iuchi et al., 2001; Wan and Li, 2006; Melhorn et al., 2008) that catalyzes the cleavage of cis-xanthophylls (Nambara and Marion-Poll, 2005) during ABA synthesis and is feed-forward-regulated by ABA (Barrero et al., 2006). NCED3 was almost 3-fold up-regulated upon sulfate exposure, but not affected by ABA. Moreover, combined application of sulfate together with ABA prevents the sulfate effect. These data suggest that xylem-delivered sulfate enhances ABA biosynthesis in guard cells (Fig. 11F).

Figure 11.

Sulfate causes ABA-like gene expression regulation. Arabidopsis leaves were fed with 6 μm ± ABA (equivalent to 3 μm biologically active ABA), 10 mm MgSO₄, or both fed via the petiole for 4 h. Subsequently guard cell enrichment was performed as described by Bauer et al. (2013). A to E, Genes that are highly regulated by ABA showed the same features as sulfate. F, Strong induction of NCED3 indicates induced ABA synthesis. qPCR data were normalized to control plant values at 100% (mean ± sd, n ≥ 10) and analyzed by ANOVA on ranks followed by Tukey test. Only differences at P < 0.05 were taken into consideration and marked with different letters.

DISCUSSION

Is Sulfate a Xylem-Delivered Chemical Signal?

The presented experiments suggest that stomatal closure during drought is promoted by sulfate delivered via the xylem to the leaves. When stomatal conductance declined after 48 h of drought, only sulfate was increased in the xylem sap whereas ABA increased after 64 h and malate after 56 h of water stress. Furthermore, this drought-related sulfate signal preceded hydraulic signaling (Fig. 1). Velocity of xylem sap flow of 1.5-m-tall poplar plants (same clone as used in this study) was ∼5 to 6 m per hour (Windt et al., 2006), suggesting that sulfate can reach guard cells upon drought within 10 min with this approach, using poplar plants that are ∼80 cm in height. Although in the study of Christmann et al. (2007) water stress was evoked suddenly and hydraulic changes were observed, water stress in this study was imposed gradually and did not affect hydraulic conductivity. Holbrook et al. (2002) already showed that stomata close as the soil dries, independent of leaf-water deficit and root-produced ABA; these authors hypothesized that a chemical signal from the roots leading to a change in ABA levels in leaves may be responsible for stomatal closure. In this study, feeding of sulfate triggered stomatal closure, and thus, established causal relationship between sulfate and stomatal closure (Figs. 6–9; Supplemental Comment). Independent evidence for sulfate as a signal for stomatal control came from SO₂ fumigation experiments with Populus × canescens (Randewig et al., 2014). SO₂ reacts with water by dissociation to sulfite. Inside plant cells, sulfite is detoxified to sulfate by oxidation via sulfite oxidase (SO) in peroxisomes (Nowak et al., 2004; Brychkova et al., 2007; Lang et al., 2007) or directly in the apoplastic space by apoplastic peroxidases (Hamisch et al., 2012). Accumulation of SO₂-derived sulfate in the apoplastic solution around guard cells may trigger guard-cell turgor loss by opening of QUAC1/ALMT12. Recalculation of data from SO₂ exposure experiments (Randewig et al., 2014) revealed strong correlation between sulfate accumulation after SO₂ exposure and the relative decrease of stomatal conductance (Supplemental Fig. S5).

Irrespective of the species and methods, ABA-induced reduction in stomatal conductance was only slightly strengthened by sulfate (Figs. 6 and 8; Ernst et al., 2010; Korovetska et al., 2014) and vice versa, the sulfate-induced stomatal closure was only slightly promoted by ABA (Fig. 6). However, during drought when sulfate increased in the xylem sap of poplar, ABA remained low (Fig. 1). After continuing drought stress, sulfate in the xylem sap further increased, whereas ABA also starts to increase. Malate and potassium levels, both involved in stomatal movement, were affected in the xylem sap during early drought, i.e. malate increased whereas potassium decreased. Hence, this data cannot exclude that malate and/or potassium (Hedrich et al., 1994, Schachtman and Goodger 2008) could contribute to early effects on stomatal conductance under drought stress. However, this investigation indicates that sulfate might be one xylem-delivered chemical signal of stomatal closure in response to drought. It may be assumed that plants cultivated with GSH as their sole sulfur source would show impaired stomatal responsiveness to drought due to a lack of sulfate. However, such an effect was not observed when Arabidopsis plants were cultivated with GSH as sole sulfur source (data not shown). Both, plants cultivated on sulfate and on GSH contained comparable sulfate levels (data not shown) probably due to the oxidation of Cys to sulfate upon GSH degradation via sulfite oxidase (Rennenberg et al., 1982; Hänsch et al., 2007). Plant growth at low sulfate may be an alternative possibility to reduce sulfate levels in roots. However, culture conditions that support normal growth and prevent sulfate accumulation in the roots have so far not been achieved.

How Does Drought Trigger Enhanced Sulfate Levels in the Xylem Sap?

Malcheska et al. (2013) suggested that sulfate in the xylem sap can originate from sulfate uptake by roots or by sulfate release from xylem parenchyma and pith ray cells of the wood. Lateral roots of Quercus robur seedlings, however, showed sulfate uptake rates being independent of predrawn shoot water potential, but xylem loading strongly declined with decreasing leaf water potential (Seegmüller and Rennenberg, 2012). This effect should not increase (as found in this study), but instead decrease sulfate in the xylem sap upon drought. Expression analyses of different SULTRs in poplar roots further indicate that increasing sulfate uptake is not an important source of the sulfate accumulation in the xylem sap upon drought, but instead diminished unloading of sulfate from the xylem sap. mRNA abundance of PtaSULTR3;3a at early drought and after prolonged drought stress PtaSULTR1;1 decreased in all root fractions upon drought (Fig. 3). Both SULTRs are expressed in xylem parenchyma cells and thought to be involved in xylem unloading of sulfate (Dürr et al., 2010). However, at early drought PtaSULTR3;3a seems to possess a predominant role because it decreased first, whereas PtaSULTR1;1 may mediate reduced sulfate retrieval at later stages of drought. Expression of PtaSULTR3;3a decreased first in elongating roots exploiting new soil areas and then after 48 h in roots showing secondary growth. Because of the low half-life of sulfate transporter protein (Rennenberg et al., 1989), such a regulation would allow for a rapid response to low water availability.

Higher contents of sulfate in the xylem sap attributed to slower sap flow rates seems unlikely because during early water deprivation, consistent changes in the xylem sap concentrations of several other ions were not recorded (Fig. 2). PtaALMT3b, a homolog of AtALMT12 (Barbier-Brygoo et al., 2011; Dreyer et al., 2012), increased during early drought in elongating roots and fine roots (Fig. 5) and may mediate progressive release of sulfate from xylem parenchyma cells. The increased expression of PtaALMT3b in elongating roots exploiting new soil areas coincided with reduced expression of PtaSULTR3;3a that removes sulfate from the xylem. In Arabidopsis roots, expression of AtALMT12 also increased after 9 d of water stress (Rasheed et al., 2016). Because AtALMT12 is expressed in the root stele of Arabidopsis (Sasaki et al., 2010), sulfate gating from the cytosol of xylem parenchyma cells into the xylem sap by this channel may contribute to increased xylem sap sulfate concentrations. Thus, decreased sulfate retrieval from the xylem sap together with enhanced mobilization of sulfate from xylem parenchyma cells may jointly contribute to enhance sulfate levels in the xylem sap upon drought (Fig. 12). Increased sulfate in the xylem sap probably originated mostly from the cytosolic pool, because putative tonoplast-localized sulfate transporters (group 4) were not consistently affected by drought in any root fraction. mRNA of PtaSULTR4;2 increased in elongating roots only after 56 and 64 h of water withdrawal, whereas expression of PtaSULTR4;1 remained unaffected. Therefore, enhanced sulfate efflux from the vacuoles seems unlikely.

Figure 12.

Sulfate enrichment in the xylem sap during early drought in poplar results in enhanced apoplastic sulfate levels around guard cells. A, Under well-watered conditions, PtaSULTR3;3a (blue) is highly expressed in roots showing secondary growth, elongating, and fine roots. In combination with low expression of PtaALMT3b (gray), a putative sulfate-permeable channel (Barbier-Brygoo et al., 2011; Dreyer et al., 2012) in elongating and fine roots results in low sulfate concentrations in the xylem sap. The low expression of the putative chloroplast/plastid-localized PtaSULTR3;1b (gray, Cao et al., 2013, 2014) prevents enhanced sulfate transport into chloroplasts/plastids. B, Water stress results in enhanced sulfate concentration in the xylem sap and, as a consequence, in higher sulfate levels in the leaf apoplast. Increasing sulfate concentrations in the xylem sap are realized by enhanced sulfate efflux from root parenchyma cells, indicated by increased PtaALMT3b (orange) expression in elongating roots and fine roots, together with decreased sulfate uptake from the xylem sap, indicated by reduced expression of PtaSULTR3;3a (gray) in all three root fractions. In both leaves and roots, expression of the putative chloroplast/plastid-localized PtaSULTR3;1b (green; Cao et al., 2013, 2014) is enhanced during water stress as well. For a detailed view on the guard cells, see Figure 13. Enhanced expression is indicated in color, whereas low expression is indicated in gray.

Xylem-Delivered Sulfate Interacts with QUAC1/ALMT12 and ABA Synthesis in the Leaves

The Arabidopsis mutant Atalmt12 is impaired in stomatal closure to different signals, including ABA (Meyer et al., 2010; Sasaki et al., 2010). This study showed that this mutant is insensitive toward sulfate and as well toward joint sulfate-ABA-induced stomatal closure (Figs. 8 and 9). However, in contrast to a recently published study on Atalmt12 mutants (Medeiros et al., 2016), the rate of photosynthesis (Fig. 9; Supplemental Fig. S4) and growth (Supplemental Table S2) was not different between wild-type plants and the mutant in this study. Nevertheless, transpiration during drought remained higher in the Atalmt12 mutant in both studies (Fig. 9; Medeiros et al., 2016), supporting the view of impaired stomatal closure. Electrophysiological studies with QUAC1/ALMT12-expressing X. laevis oocytes revealed that extracellular and cytosolic malate and cytosolic sulfate activate the QUAC1/ALMT12 channel (Meyer et al., 2010; Sasaki et al., 2010). In addition, this study also showed that extracellular sulfate activates the QUAC1/ALMT12 channel (Fig. 10; Mumm et al., 2013). Enhanced xylem-delivered sulfate that enters the substomatal apoplastic space and gets into contact with guard cells either can activate QUAC1/ALMT12 (Figs. 12 and 13A) or can be taken up into guard cells. In the presence of extracellular sulfate, QUAC1/ALMT12 was activated with typical R-type channel fast-activation kinetics (Fig. 10A). Thus, anions released from guard cells by xylem-derived sulfate due to QUAC1/ALMT12 activation can depolarize the guard cell plasma membrane which, subsequently, leads to opening of potassium outward channels, such as GORK, releasing K⁺. This results in stomatal turgor loss and, consequently, stomatal closure (Ache et al., 2000). However, sulfate is not the sole signal affecting guard cell turgor. Other signals such as an endogenous control mechanism, the combination of several ion channels, and a complex signal transduction network also contribute to stomatal movement (Kim et al., 2010; Hedrich, 2012; Munemasa et al., 2015; Minguet-Parramona et al., 2016).

Figure 13.

Overview of four possible mechanisms by which xylem-delivered sulfate could trigger stomatal closure. Three sulfate-mediated mechanisms can result in the release of anions such as chloride, nitrate, and sulfate that depolarize the plasma membrane, which further leads to opening of potassium outward channels such as GORK (gray). Then GORK can release K⁺, thereby mediating stomatal turgor loss and, consequently, stomatal closure (Ache et al., 2000). A, First, xylem-delivered sulfate reaching the apoplast of guard cells can change voltage-dependent gating and thus triggers QUAC1/ALMT12 (pink) opening. B, Second, sulfate taken up into guard cells might trigger ABA synthesis via NCED3 (marked in blue), which is a key step in ABA synthesis in the chloroplasts (Bauer et al., 2013), thereby causing higher ABA concentration in the guard cell cytosol. C, Third, sulfate delivered through the xylem sap can be taken up into guard cells and further into the chloroplast by PtaSULTR3;1b (green). Here sulfate can be used for sulfate reduction and Cys synthesis. Increased expression of SULTR3;1, PtaSULTR3;1b in poplar, may enhance sulfate transport into chloroplasts. Cys is needed in the cytosol for the Moco sulfurase reaction (ABA3) that sulfurylates Moco, which is used by AAO3 in the last step of ABA synthesis (Seo and Koshiba, 2002; Nambara and Marion-Poll, 2005). This assumption is supported by a sultr3;1 loss-of-function Arabidopsis mutant that synthesizes less ABA (Cao et al., 2013) whereas high sulfate availability facilitates ABA synthesis. ABA can accumulate in the cytosol due to higher ABA synthesis in guard cells via (B) and/or (C) and can bind to the ABA receptors PYR/PYL/RCAR (gray). The activated ABA receptor inhibits PP2C (gray) phosphatases like ABI1 or ABI2. Ser/Thr-protein kinases, like OST1 (gray), are then released from inhibition and are able to activate target proteins like QUAC1/ALMT12 (pink) by phosphorylation (Imes et al., 2013; Osakabe et al., 2014). D, Sulfate uptake into guard cells and further into chloroplasts, indicated by enhanced expression of PtaSULTR3;1b (Cao et al., 2013), can result in Cys accumulation (Calderwood and Kopriva, 2014). In an additional mechanism, Cys can be used in the cytosol for H₂S synthesis by l-Cys desulfhydrase activity (Calderwood and Kopriva, 2014). H₂S is able to inhibit inward-rectifying K⁺ channels, thereby preventing stomatal opening (Papanatsiou et al., 2015). Labeling: All factors such as enzymes, proteins, channels, and metabolites and metabolic pathways not investigated in this study, are labeled in gray. Factors analyzed and/or expected to be affected, are marked either by color (investigated) or in black (assumed to be affected).

Another way of sulfate signaling becomes evident after sulfate enters the guard cells and may stimulate guard cell-autonomous ABA synthesis upon drought (Bauer et al., 2013; Fig. 13, B and C). A stimulating effect of sulfate on ABA synthesis has been shown with Arabidopsis seedlings by applying different sulfate concentrations to the growth medium (Cao et al., 2014). By working with a loss-of-function mutant deficient in the chloroplast-localized sulfate transporter SULTR3;1, Cao et al. (2014) demonstrated diminished AAO activity and ABA levels in Arabidopsis seedlings. This last step in ABA synthesis (Nambara and Marion-Poll, 2005) needs Cys in the cytosol that is synthesized via sulfate reduction in chloroplasts (Takahashi et al., 2011; Fig. 13D). Cys is also the precursor of H₂S, synthesized via l-Cys desulfhydrase, which affects stomatal closure as well (Calderwood and Kopriva, 2014; Scuffi et al., 2016). Although contradicting results, i.e. stomatal opening (Lisjak et al., 2010, 2011) and closure (García-Mata and Lamattina, 2010; Jin et al., 2013), have been reported, H₂S seems to affect stomatal closure in an ABA-independent way by inhibiting inward-rectifying K⁺ channels (Papanatsiou et al., 2015). Sulfate could be taken up into guard cells and further transported into the chloroplasts for Cys synthesis. Enrichment in Cys may also trigger H₂S formation (Fig. 13D; Calderwood and Kopriva, 2014) and, thus, stomatal closure from xylem-delivered sulfate.

In this study, a stimulating effect of sulfate on guard-cell autonomous ABA synthesis by increased NCED3 expression was strongly evident from feeding sulfate to Arabidopsis leaf petioles (Fig. 11). Even in Arabidopsis roots and shoots, expression of NCED3 increased after 3 and 5 d of water stress, respectively (Rasheed et al., 2016). ABA- or sulfate-treated Arabidopsis guard cells showed identical expression patterns of several ABA-controlled genes, but also revealed that the key transcript of ABA biosynthesis (NCED3; Wan and Li, 2006; Melhorn et al., 2008) is ∼3-fold induced by sulfate; this induction was counteracted by the addition of ABA (Fig. 11).

In summary, although other signals such as malate, potassium, and/or ABA might be involved in long-distance drought signaling, this data indicate that xylem-delivered sulfate contributes to stomatal closure by different mechanisms, or processes. For process 1, extracellular sulfate can promote stomatal closure by gating the guard cells’ anion channel QUAC1/ALMT12 open (Figs. 10 and 13A). For process 2, xylem-delivered sulfate taken up into guard cells can induce NCED3 expression and thereby up-regulates the key step in ABA synthesis in the chloroplasts (Figs. 11 and 13B). For process 3, xylem-delivered sulfate can enter the chloroplast of guard cells and can enhance Cys synthesis that in turn may promote ABA synthesis (Fig. 13C). For process 4, sulfate-induced Cys synthesis can result in H₂S production that effects inward-rectifying K⁺ channels (Fig. 13D). Via processes 1, 2, and 3, several ion channels, e.g. SLAC1, SLAH3, and QUAC1/ALMT12, on the plasma membrane of guard cells, are targeted that lead to ion efflux and plasma membrane depolarization (Imes et al., 2013). In turn, plasma membrane depolarization activates efflux of K⁺ from the guard cells by GORK (Ache et al., 2000). Efflux of ions and potassium from guard cells leads to turgor loss and subsequent stomatal closure upon drought. In contrast, process 4 will prevent stomatal opening. Which of these signaling pathways/processes is of superior importance during early drought needs further investigation.

Increasing ABA concentrations in the xylem sap after prolonged water stress (ABA in the xylem sap increased after 64-h drought in this study) could be the result of long-distance ABA cycling after ABA synthesis in leaves and phloem-to-xylem exchange along the transport to the roots (Goodger and Schachtman, 2010). Another explanation refers to the suggestion of Goodger and Schachtman (2010) that ABA accumulation at prolonged water stress may result from enhanced ABA synthesis in roots. In this study, the predicted chloroplast-localized sulfate transport PtSULTR3;1b showed low expression under well-watered conditions in all three root fractions whereas it continuously increased during ongoing water stress (Fig. 4). As the fine root fraction accounts for ∼50% of total root biomass (Herschbach et al., 2010), PtSULTR3;1b may contribute to sulfate transport into plastids of root parenchyma cells to promote the roots’ own ABA synthesis at ongoing drought as shown for salt stress in the roots of Arabidopsis (Cao et al., 2014; Ruiz-Sola et al., 2014). Further studies are required to unravel the contribution of such a molecular mechanism on sulfate-induced stomatal closure and to address the importance of xylem-delivered sulfate as an early drought stress signal from roots to the shoot as well as the contribution of sulfate to the roots’ own ABA synthesis.

MATERIALS AND METHODS

Plant Material

Poplar plants (Populus tremula × Populus alba; synonym P. × canescens) were micropropagated as described by Strohm et al. (1995) from sterile cultures. Cuttings were planted into quartz sand (1.0–2.0 mm size of the particles) and were cultured in a greenhouse under long-day conditions (Scheerer et al., 2010). The plants were fertilized once per week with 200 mL Hoagland solution (Honsel et al., 2012). If required, they were additionally watered with distilled water. After 3 months, plants at a height of ∼70 to 90 cm and an age of 15 to 18 weeks were placed into a controlled environmental growth chamber (HPS 1500; Heraeus Industrietechnik) for 2 weeks of acclimation (23/20°C d/n, a photoperiod of 16 h light) at a photosynthetic photon flux density of 150 ± 4 μmol m⁻² s⁻¹ at plant level and 60% relative air humidity. Arabidopsis (Arabidopsis thaliana) wild-type (Col-0) and Atalmt12 mutants (Meyer et al., 2010) were grown from seeds on commercial soils (Anzuchtsubstrat; Floragard Vertriebs) under short-day conditions (8-/16-h light/dark) in a controlled environmental growth chamber (23/20°C, 60% relative air humidity, 70 ± 5 μmol m⁻² s⁻¹ at plant level).

Leaf Gas Exchange Measurements

Transpiration, stomatal conductance, and photosynthesis rates of poplar and Arabidopsis (wild-type and Atalmt12 mutant) plants were measured at the 10th attached poplar leaf of four to six plants or at a mature Arabidopsis leaf from 11 to 14 plants using a portable gas exchange system (GFS 3000; Walz; Liu et al., 2015). Attached poplar leaves were placed into a leaf enclosure with 8-cm² leaf areas and attached mature Arabidopsis leaves into a leaf enclosure with 3-cm² leaf area (Walz). Enclosures were flushed with ambient air containing 400 μL L⁻¹ CO₂ and 10,000 μL L⁻¹ water vapor at a flow rate of 650 mL min⁻¹. Leaf temperature was maintained at 25°C and light intensity at 1,000 μmol m⁻² s⁻¹ photosynthetic photon flux density. After adaptation to enclosure conditions (∼5–10 min), gas exchange was determined for 10 min for both poplar and Arabidopsis.

The effect of sulfate on stomatal conductance of detached leaves of Arabidopsis wild-type (Col-1) and the Atalmt12 mutant was investigated by monitoring water loss over time by weighting the tubes containing the feeding solution plus two leaves. Stomatal conductance was then calculated as water loss per unit leaf area and time (mmol m⁻² s⁻¹). For detailed information on the calculation, see Supplemental Materials and Methods.

Furthermore, the effect of SO₄²⁻ application on transpiration of detached 5- to 6-week-old Arabidopsis wild-type (Col-0) leaves was measured with a different custom-made setup. Water vapor was recorded using two parallel water-cooled cuvettes with a gas stream of 1 L min⁻¹. The gas composition was controlled by mass flow meters, adjusted to 52.5% relative humidity at 20°C and 380 μL L⁻¹ CO₂, and detected by a HCM100 (Walz) as described by Bauer et al. (2013). This setup was designed to monitor stomatal movement only by relative measurement of transpiration related to the leaf surface. Leaves were illuminated by two LEDs providing light at 655 nm (3-W WEPDR3-S1 Power LED Star tiefrot; Winger) and 455 nm (Luxeon Royal Blue; Philips) at a photon flux density of 100 and 8 µmol m⁻² s⁻¹, respectively. The light beams were collected by two dichroic mirrors (Q525 LPXR and DCLP 425; Chroma Technology) and directed to the cuvettes via two fiber optics (Fiber Optic Illuminator FL-460). Detached Arabidopsis leaves from the wild type (Col-0) were cut under water and placed into tubes containing the feeding solution [1 mm MgCl₂ and 0.06 mm Mg(NO₃)₂] in the gas exchange cuvettes. After 30 to 60 min in darkness when the water vapor concentration reached equilibrium, leaves were illuminated and increased water loss was monitored. After stabilization, MgSO₄ (10 mm final concentration) or feeding solution [as control, 1 mm MgCl₂ and 0.06 mm Mg(NO₃)₂] was added via a tube into the feeding solution reservoir.

Drought Stress Experiments

Fifteen- to eighteen-week-old poplar plants were well watered up to the control day (day 0) and thereafter exposed to drought by terminating water supply. Plants were harvested at 0 h of water deprivation (day 0, control) and at 24, 48, 56, 64, and 72 h of water deprivation. The water status of plants was monitored (1) by measuring stomatal conductance of the 10th attached leaf and (2) by determining the stem water potential according to the method of Scholander et al. (1965) using a pressure vessel (Soilmoisture). Xylem sap samples were collected as described by Rennenberg et al. (1996). The 10th leaf and three different root fractions, i.e. elongating roots without side roots (up to 5 cm from the root tip), fine roots, and roots showing secondary growth, were harvested, frozen in liquid N₂, and stored at −80°C. The pH of the xylem sap was measured with a pH-Meter (pH 526 Multical using a SenTix Mic-D electrode; WTW) and was at 0 h 5.99 ± 0.22, at 24 h 5.95 ± 0.13, at 48 h 5.75 ± 0.33, at 56 h 5.65 ± 0.2, at 64 h 5.85 ± 0.18, and at 72 h 5.6 ± 0.3. Data presented are combined normalized data of two experiments with four to eleven replicates at each harvest. Original data of all parameters for the controls at 0 h of water deprivation of both experiments are presented in the Supplemental Table S1. Therefore, respective mean values received at the control, i.e. day 0, were set to 100% and used as reference to calculate relative values.

Drought stress of Arabidopsis plants was followed by determining pot weight. First visible symptoms of drought stress were detected when the gravimetric water content of the soil reached 0.67 ± 0.22 g g⁻¹ for the wild-type and 0.73 ± 0.14 g g⁻¹ for the mutant (Supplemental Table S2).

Feeding of Detached Leaves via the Petiole

Stomatal conductance of the 10th attached leaf of poplar was measured before the feeding experiment of detached poplar leaves. After detaching, the leaf petiole was directly placed into a solution consisting of 1 mm MgCl₂ and 0.06 mm Mg(NO₃)₂ adjusted to pH 5.5 with NaOH (pH 5.5 was chosen according to the pH of the xylem sap of poplar) for equilibrating stomatal conductance (preincubation). The solution was without potassium to avoid influence on stomatal behavior by this ion. After detached leaves reached a comparable stomatal conductance to that of being attached, after ∼60 min the solution was replaced by solutions containing different concentrations of MgSO₄ (0.2, 1, and 2 mm), ABA (0.3 and 3 μm), or a mixture of MgSO₄ plus ABA (2 mm MgSO₄ plus 3 μm ABA; 2 mm Mg₂SO₄ plus 0.3 μm ABA) adjusted to pH 5.5. Stomatal conductance was recorded for 1 h in four to six biological replicates. The stomatal conductance measured after 60 min of incubation was calculated as percentage compared to the stomatal conductance determined during the preincubation.

In accordance to the experiment with epidermis peels of Arabidopsis (see below), comparable treatments were performed with excised leaves. Therefore, 40 leaves of the wild-type Arabidopsis and of the Atalmt12 mutant were randomly collected and two of them were placed into a 0.5-mL tube (Eppendorf). Each tube contained 400 µL of deionized water at pH 5.5. Tubes were covered with Parafilm to avoid water loss by evaporation. After 2 h of preincubation, leaves were placed into new tubes with 400 μL of the solution containing either deionized water or to 10 mm MgSO₄ adjusted to pH 5.5 (n = 10 per line and treatment). Water loss from leaves and, thus, transpiration was determined by weighing the leaves together with the tubes every 30 min over 5 h. Control tubes without leaves did not show any evaporation if covered with Parafilm. Temperature and relative air humidity were recorded to calculate stomatal conductance (see below). Leaf area was determined from images taken after experiments by using the open source software ImageJ (National Institutes of Health). Transpiration rates were calculated as mmol of water loss per leaf area in s (mmol m⁻² s⁻¹). Stomatal conductance for water vapor (gH₂O) was calculated based on the transpiration rates, relative air humidity, and temperatures as described by De Kok et al. (1989) and Rennenberg et al. (1996) (equations are given in Supplemental Materials and Methods). The experiment was performed twice with comparable results that were combined.

Measurements of Stomatal Aperture in Experiments with Epidermis Peels

The impact of sulfate and ABA on stomatal aperture of 5-week-old wild-type Arabidopsis and Atalmt12 mutants grown on soil was determined according to Ernst et al. (2010) with minor modifications. Epidermal peels were floated for 2 h at room temperature and constant white light (200 µmol m⁻² s⁻¹) on water or on incubation buffer (50 mm KCl and 10 mm MES, pH 5.3) with the stomata side of the epidermal peel facing the ambient air. The effect of MgSO₄ (2–15 mm) and ABA (0.1–0.3 µm) on stomatal aperture was tested by transfer of epidermal peels to water or incubation buffer supplemented with respective compounds for 3 h. The stomatal aperture was determined with a conventional wide-angle microscope (DMIRB; Leica Microsystems).

Determination of the Anions

Sulfate and P contents in the xylem sap were determined by ion-exchange chromatography (DX-120 Ion Chromatograph; Dionex) as described by Herschbach et al. (2000). The xylem sap was diluted from 1:2 up to 1:20 with deionized water.

Potassium and malate was determined in 10-fold ultrapure-water-diluted xylem sap before isocratic separation of cations with 30 mm methanesulfonic acid at 43°C and a flow rate of 0.36 mL min⁻¹ for 27 min on an IonPac CS16 Column (2 mm; Thermo Fisher Scientific) connected to an ICS-1000 System (Dionex). Quantification was performed by conductivity detection after anion suppression (CERS-500 2 mm, suppressor current 43 mA) with the software Chromeleon version 6.6 (Dionex). Ions were quantified from the same sample according to Wirtz and Hell (2007).

Quantitative RT-PCR

Total RNA was isolated from 100 mg homogenized root and leaf material according to Kolosova et al. (2004) and cDNA synthesis was performed as described in Dürr et al. (2010), Honsel et al. (2012), and Malcheska et al. (2013). Gene expression analyses of PtaSULTRs, PtaALMT3a, and PtaALMT3b were performed by RT-qPCR, using a LightCycler 480 System (Roche Applied Science) as described by Dürr et al. (2010), Honsel et al. (2012), and Malcheska et al. (2013). Lengths of the PCR fragments and primer sequences of all analyzed genes, and for the selected housekeeping gene PtaEf1b, are given in Supplemental Table S3.

Transcript Analysis of Guard Cells Isolated from Arabidopsis Leaves Treated with ABA and/or Sulfate

Five to six leaves of 6- to 7-week-old Arabidopsis Col-0 were cut under water to avoid embolism, then transferred to incubation (50 mm KCl and 10 mm MES, pH 5.3) for 1 h. Then 6 μm ± ABA (equivalent to 3 μm biologically active ABA) or 10 mm MgSO₄, or both, were added (final concentrations). After 4 h incubation, guard cells were extracted using the blender method (described in detail in Hedrich et al. [1989] and Raschke and Hedrich [1989]) and qPCR was performed as described by Geiger et al. (2011). Transcripts were normalized to 10,000 molecules of actin 2:8 using standard curves calculated for the individual PCR products.

Determination of ABA

Two technical replicates of 50 µL of xylem sap were used for ABA extraction. Before the extraction procedure, 100 ng of [²H₆]-ABA (Plant Biotechnology Institute, National Research Council of Canada) were added to each sample, which was then incubated for 30 min under continuous shaking at 4°C. Subsequently, the aqueous sample was acidified to pH 3 with 1 m HCl and extracted two times with 600 µL ethyl acetate. For phase separation, the samples were centrifuged for 10 min at 13,000g. The ethyl acetate phases were carefully removed and combined. Ethyl acetate was then evaporated under a stream of N₂ and the samples were resuspended in 20 µL methanol. Methylation was performed by adding equal sample amounts of a 1:10 diluted solution (in diethylether) of trimethylsilyldiazomethane (Sigma-Aldrich) for 30 min at room temperature. The mixture was evaporated and resuspended in 50 µL ethyl acetate for GC-MS analysis.

GC-MS analysis was carried out on a Saturn 2100 Ion-trap Mass Spectrometer using electron impact ionization at 70 eV, connected to a gas chromatograph (model no. CP-3900) equipped with an autosampler (model no. CP-8400; all by Varian). For analysis, 1 µL of the methylated samples was injected in the splitless mode (splitter opening 1:100 after 1 min) onto a model no. ZB-5 column (30 m × 0.25 mm × 0.25 µm; Phenomenex) using He carrier gas at 1 mL min⁻¹. Injector temperature was 250°C and the temperature program was 60°C for 1 min, followed by an increase of 25°C min⁻¹ to 180°C, 5°C min⁻¹ to 250°C, and 25°C min⁻¹ to 280°C, then 5 min isothermically at 280°C. For higher sensitivity, the μSIS mode (the Varian Manual; Wells and Huston, 1995) was used. The endogenous hormone concentrations were calculated by the principle of isotope dilution (Cohen et al., 1986), using the ions at m/z 190:194 (endogenous and labeled standard; note that during fragmentation of ABA, two deuterium are lost) for methylated ABA (Walker-Simmons et al., 2000).

Two-Electrode Voltage-Clamp Experiments with Xenopus laevis Oocytes

For functional analyses, cRNA of QUAC1/ALMT12 and OST1 was prepared using the AmpliCap-Max T7 High Yield Message Maker Kits (EPICENTRE). Oocyte preparation and cRNA injection have been described by Becker et al. (1996). Oocytes were injected with 50 nL cRNA of QUAC1 (500 ng µL⁻¹) and OST1 (250 ng µL⁻¹) and incubated for 3 d at 16°C in ND96 solution. Before measurements, oocytes were injected with 50 nL of a 200 mm NaH-malate solution (pH 7.5) resulting in a final cytosolic malate concentration of 18 mm. Whole oocyte currents were recorded using the two-electrode voltage-clamp technique. The holding potential was clamped to −200 mV. Five-hundred-milliseconds-lasting test pulses ranged from 60 to −200 mV. The rel. Po was inferred from instantaneous current responses at a constant voltage pulse of −200 mV after to the test pulses. The half-maximal activation potential (V_1/2) was calculated by fitting the experimental data points with a single Boltzmann equation. Rel. P_O curves were normalized to the saturation values of the calculated Boltzmann distribution. Oocytes were perfused with solution containing 10 mm MES/Tris, pH 5.6, 1 mm Ca-Gluconate₂, 1 mm MgGluconate₂, 1 mm LaCl₃ and either 10 mm of NaH-malate, NaCl, or NaHSO₄. The osmolality was adjusted to 220 mOsmol kg⁻¹ with d-sorbitol.

Statistics

Statistics were performed according to the experimental design and data distribution. Statistical analyses and graphs were prepared with the software SigmaPlot version 11 (Systat Software) or with the software OriginPro version 9.1 (http://www.originlab.de). Comparisons between the control (day 0) and different periods of water deprivation in the drought-stress experiment with poplar were analyzed by Student’s t test for data with normal distribution (Figs. 1–5). The effects of different solutions on stomatal conductance of detached leaves were analyzed using Student’s t test for data with normal distribution (Fig. 6). Mann-Whitney U test was used when data were not normally distributed (Figs. 1–6). Normal distribution was tested with Shapiro-Wilk. One-way ANOVA using a Tukey test after testing for normal distribution with the Kolmogorov-Smirnov and Shapiro-Wilk tests was applied for statistical analyses between wild-type Arabidopsis and mutants exposed to drought (Fig. 9). ANOVA on ranks followed by the Tukey test was applied for the statistical analyses of relative expression levels of several ABA-induced genes in guard-cell-enriched extracts from peeled epidermis (Fig. 11). One-way ANOVA on ranks followed by Student-Newman-Keuls was used to identify significant differences between treatments of Arabidopsis epidermis peels. Significant differences at P < 0.05 were considered (Fig. 8). The strength of association between the relative difference in stomatal conductance for water vapor g(H₂O) and sulfate content of SO₂-exposed plants of poplar were tested with Pearson's product-moment correlation analyses. Data were log-transformed to achieve normal distribution and constant variance of the residuals. The correlation analyses and test of assumptions were performed with the software language R, version 3.0.1 (package “lme4”; R Core Team, 2013).

Accession Numbers

Sequences of PtaSULTR of P. × canescens are available in the NCBI database (https://www.ncbi.nlm.nih.gov) under the following accession numbers: DQ906929 (PtaSULTR1;1, Potri.005G169300), DQ174472 (PtaSULTR1;2, Potri.002G092500), DQ906928 (PtaSULTR3;1b, Potri.005G213500), DQ906924 (PtaSULTR3;3a, Potri.008G130400), DQ906930 (PtaSULTR4;1, Potri.008G049500), DQ906935 (PtaSULTR4;2, Potri.010G211400), and FJ372570 (PtaEf1b, Potri.015G094200). ALMT family members of clade 3 for analyses in P. × canescens were taken from P. trichocarpa (Barbier-Brygoo et al., 2011) from Phytozome version 9 (http://www.phytozome.net/). Arabidopsis ALMT12 homologs are: POPTR_0001s02700, Potri.001G144300 (PtaALMT3a), POPTR_0005s23000, and Potri.005G208500 (PtaALMT3b). Genes of ABA synthesis down-regulated by RNAi constructs in P. × canescens (Supplemental Fig. S1) are according to the P. trichocarpa analogs POPTR_007s08330, Potri.007G066400 (PtaABA3), POPTR_0004s20280, and Potri.004G191300 (PtaAO2). Arabidopsis transcripts that have been quantified by qPCR are PYL2 (At2g26040), PYL4 (At2g38310), MYB60 (At1g08810), HAI1 (At5g59220), ABAR (At3g02480), and NCED3 (At3g14440).

Supplemental Data

The following supplemental materials are available.

• Supplemental Figure S1. Stomatal conductance and xylem sap sulfate of poplar aba3 (Ptaba3-4, Ptaba3-7) and ao2 (Ptao2-22, Ptao2-53) RNAi lines (with halved ABA contents compared to wild-type poplar) during water deprivation.

• Supplemental Figure S2. Stomatal conductance of detached poplar leaves fed with sulfate.

• Supplemental Figure S3. Potassium dependency of sulfate-induced stomatal closure in epidermis layers of Arabidopsis.

• Supplemental Figure S4. Transpiration of wild-type Arabidopsis and of the Atalmt12 mutant in response to drought as well as stomatal conductance of detached leaves in response to sulfate.

• Supplemental Figure S5. Correlation analyses between stomatal conductance for water vapor g(H₂O) at daytime and sulfate concentrations of P. × canescens in response to SO₂ exposure.

• Supplemental Table S1. Experimental raw data (mean values ± sd) for the control plants of the drought (Figs. 1–5) and the feeding experiments with poplar (Fig. 6).

• Supplemental Table S2. Biometrics of wild-type Arabidopsis and of the Atalmt12 mutant as well as basic parameters of soil water content during drought of the gas exchange experiment presented in Figure 9.

• Supplemental Table S3. Primer sequences and length of amplified gene fragments in the real-time PCR approach.

• Supplemental Comment.

• Supplemental Materials and Methods.

• Supplemental References.