Introduction

The homeostasis of monovalent cations is a fundamental activity of living cells, with both permissive and regulatory roles in many cellular functions. The transport of K⁺, Na⁺ and H⁺ determines membrane potential, turgor, intracellular cation concentrations and pH, and these basic parameters are crucial for nutrient uptake, metabolism, cellular integrity, cell death, growth and differentiation (Hoffman, 1964; Harold, 1986; Hager et al, 1991; Yenush et al, 2002, 2005; Adams et al, 2006). Although the basic mechanisms of monovalent cation transport have been identified, our knowledge of their regulatory systems is rather fragmentary (Sanders and Bethke, 2000; Serrano and Rodriguez-Navarro, 2001; Pollard and Earnshaw, 2002).

Fungi and plants share general transport mechanisms at the plasma membrane based on an H⁺ chemiosmotic circuit. The primary pump is the electrogenic H⁺-pumping P-ATPase that generates an electrochemical H⁺ gradient (Serrano, 1989; Sussman and Harper, 1989; Morsomme and Boutry, 2000). This H⁺ gradient drives the secondary transport of K⁺ mediated by channels and carriers (Very and Sentenac, 2002; Rodriguez-Navarro and Rubio, 2006), as well as the uptake of nutrients by H⁺-symporters and the efflux of Na⁺ by H⁺-antiporters (Sanders and Bethke, 2000).

In plants, the regulation of H⁺-ATPases and K⁺ channels and carriers involves changes in both expression and activity of these systems. Auxin increases the levels of H⁺-ATPase (Hager et al, 1991; Frías et al, 1996; Rober-Kleber et al, 2003) and inward K⁺ channels (Philippar et al, 1999) in elongating tissues and blue light activates the H⁺-ATPase of guard cells by inducing its phosphorylation (Kinoshita and Shimazaki, 1999). The fungal toxin fusicoccin activates the H⁺-ATPase by binding to its phosphorylated C-terminus and recruiting 14-3-3 proteins (Würtele et al, 2003).

Abscisic acid inhibits the activity of the inward K⁺ channels of Arabidopsis guard cells (KAT1 and KAT2) through a signal transduction pathway involving reactive oxygen species and Ca²⁺ (Kwak et al, 2003). This hormone also represses the expression of the outward K⁺ channel (SKOR) at the root pericycle and xylem parenchyma of Arabidopsis (Gaymard et al, 1998). K⁺ starvation induces the expression of high-affinity K⁺ carriers such as Hak5 (Maathuis et al, 2003; Shin and Schachtman, 2004; Gierth et al, 2005) by a mechanism including hydrogen peroxide production (Shin and Schachtman, 2004). Again, the signal transduction pathways of these regulations remain largely unknown.

Na⁺ efflux in Arabidopsis is mediated by the SOS1 H⁺-antiporter, a transporter activated by the SOS2/CIP24-SOS3/CBL4 protein kinase–calcium sensor complex (Qiu et al, 2002; Quintero et al, 2002). Another protein kinase–calcium sensor complex, namely CIPK23-CBL1 (or CBL9), has recently been described to activate the AKT1 K⁺ uptake channel of Arabidopsis roots (Xu et al, 2006). These two protein kinases are the only regulatory components of plant cation homeostasis characterized at the molecular level.

One approach to identify novel regulators of cation transport consists of the identification of genes that upon gain of function improve tolerance to toxic cations (Serrano et al, 1999). Gain-of-function mutants obviate genetic redundancy and can identify bottlenecks in biological pathways. We have screened the collection of 'activation-tagging' Arabidopsis mutants (Weigel et al, 2000) for tolerance to the toxic cation norspermidine (Hamana et al, 1989), and report here that the QSO2 gene (At1g15020), encoding a quiescin-sulfhydryl oxidase (QSO), is a novel regulator of monovalent cation transport at the root symplast–xylem interface. QSOs are animal and plant enzymes proposed to participate in oxidative folding of disulfide-containing secreted proteins. Quiescins have been implicated in the regulation of growth and the elaboration of extracellular matrix of animal cells (Coppock et al, 1998; Thorpe et al, 2002). Our work suggests that these eukaryotic proteins may also regulate ion homeostasis.

Results

A screening based on tolerance to norspermidine resulted in the par1-1D mutant

We have screened the 'activation-tagging' mutant collection of the plant Arabidopsis thaliana (Weigel et al, 2000) for tolerance to toxic cations. Previous screenings for tolerance to Na⁺ in Arabidopsis resulted in mutants defective in either abscisic acid biosynthesis or perception. The high concentrations of NaCl required for toxicity during the germination assay (greater than 0.1 M) have significant osmotic effects, which trigger the biosynthesis of abscisic acid and this hormone inhibits germination and early growth (González-Guzmán et al, 2002). Alternatively, polyamines are toxic at millimolar concentrations, which pose no osmotic stress, and the presence of several positive charges per molecule makes their uptake very sensitive to membrane potential, negative inside. Norspermidine is a non-metabolizable polyamine (Hamana et al, 1989), and this type of toxic cation has been successfully used to demonstrate changes in membrane potential in yeast mutants affected in K⁺ transport (Forment et al, 2002).

We have screened 16 398 lines selecting for tolerance to 3.2 mM norspermidine, a concentration that completely inhibits growth of wild-type seeds. The most resistant mutant was named par1-1D, from 'polyamine resistant', gene 1, allele 1, Dominant (Meinke and Koornneef, 1997).

As indicated in Figure 1A and B, par1-1D seeds germinate and grow better than wild-type plants on plates containing norspermidine, spermidine, Li⁺ or Na⁺. Experiments in soil with adult plants also demonstrate tolerance to Li⁺ and Na⁺ (Figure 2). This tolerance is reflected both in the total size (Figure 2A) and in the discoloration of the leaves (Figure 2B and C) of stressed plants. Therefore par1-1D seems to have a pleiotropic phenotype of tolerance to toxic cations.

The mutation was monogenic and dominant, because F1 plants exhibited tolerance to toxic cations and there was a 3:1 segregation of the phenotype in the F2 generation. Actual data were 156:44 (c²=0.96).

The par1-1D mutation is caused by overexpression of At1g15020 (QSO2), a limiting factor for toxic cation tolerance

Homozygous par1-1D plants were crossed to Columbia wild-type plants. From the segregating F2 generation, norspermidine-tolerant plants were selected and scored for the presence of the T-DNA by Southern blot and PCR analyses. As a result, the par1-1D mutation was shown to be linked to the T-DNA because of 60 F2 norspermidine-tolerant plants analyzed, all of them contained T-DNA (data not shown). Plasmid rescue of the T-DNA and sequencing indicated that the insertion was in the genomic region depicted in Figure 3A, with the transcriptional enhancer close to the At1g15020 gene. Northern analysis confirmed that this gene was overexpressed about three-fold in the mutant (Figure 3B, left panel). As expected, the expression of the adjacent At1g15030 gene, the promoter of which was interrupted by the insertion, was greatly decreased (Figure 3B, right panel).

In order to test if overexpression of At1g15020 was responsible for the par1-1D phenotype, transgenic plants were generated, which overexpressed At1g15020 cDNA from the 35S promoter. Most (7 out of 11) of the transgenic lines overexpressed the Atg15020 gene (Figure 4A). Two overexpressing lines (numbers 2 and 10) were further analyzed and found to be more tolerant to toxic cations than controls (Figure 4B and C). This suggests that the par1-1D mutation was caused by overexpression of At1g15020.

At1g15020 encodes a sulfhydryl oxidase of the quiescin family and it has been previously named AtQSOX2, because there is a homologous gene, At2g01270, named AtQSOX1 (Thorpe et al, 2002). A recent review of the family (Houston et al, 2005) has reversed the numbers of the two genes. We propose the three-letter, italic name QSO2 to conform community standards for Arabidopsis genetics (Meinke and Koornneef, 1997), and maintain the numbers of the first publication (Thorpe et al, 2002). QSOs are flavoproteins containing a thioredoxin domain and which oxidize disulfhydryl groups in proteins to disulfides, with reduction of oxygen to hydrogen peroxide. They are secreted proteins located at the endoplasmic reticulum, Golgi and outside the cell. The domain structure of QSO2 is shown in Supplementary Figure 1S. QSOs are found only in multicellular organisms, plants and animals, while yeast cells contain enzymes lacking the thioredoxin domain but containing the sulfhydryl oxidase flavoprotein or ERV domain (Thorpe et al, 2002).

Three T-DNA insertion mutants of At1g15020 were identified in the TAIR collection (www.arabidopsis.org) and the homozygous lines named par1-2 (SALK_066130), par1-3 (SALK_025237) and par1-4 (SALK_072829). A scheme of the T-DNA insertions is presented in Figure 5A. Quantitative RT–PCR analysis indicated that QSO2 expression levels in the mutants were 13 (par1-2), 18 (par1-3) and less than 1% (par1-4) of wild type. The three mutant lines displayed more sensitivity to toxic cations than control plants, both as seedlings grown 'in vitro' (Figure 5B; details of seedlings shown for par1-2 in Figure 5C), and as plants grown in soil (details of plants shown for par1-2 in Figure 5D). Taken together with the results of the overexpression approach, these experiments demonstrate that the QSO2 sulfhydryl oxidase is a limiting factor for tolerance to toxic cations. On the other hand, a knockout mutant of the homologous gene At2g01270 has no detectable phenotype (data not shown).

QSO2 inversely regulates the accumulation of K⁺ and toxic cations

One mechanism of tolerance to toxic cations is based on decreasing cation accumulation. We have measured the initial rates of uptake (Figure 6A) and the final accumulation levels (Figure 6B) of norspermidine and Na⁺ in Arabidopsis plants (wild type, par1-1D and par1-2) incubated in liquid culture. While the initial rate of uptake (less than 20 min) was not significantly affected by either gain- or loss-of-function of QSO2, both the uptake at longer times (1 h) and the accumulation after 2 days of both toxic cations correlated inversely with QSO2 function. We have also measured the level of K⁺ (Figure 7A) and the initial rate of Rb⁺ uptake (as indication of K⁺ transport; Figure 7B). Again, the initial rate of uptake was not affected by QSO2 but, opposite to the results with toxic cations, both the uptake at long times and the accumulation of K⁺ were directly correlated with QSO2 function.

QSO2 modulates the loading of cations into the xylem

One determinant of cation accumulation, which could be affected by QSO2 is the plasma membrane electrical potential (Mulet et al, 1999). However, we have measured this parameter in root epidermal cells of wild type and plants with gain and loss of function of QSO2, and found no significant differences (Supplementary Table 1S). Another determinant of cation accumulation is the plasma membrane H⁺-ATPase, the primary pump that energizes all secondary transporters. However, neither the amount nor the activity of the enzyme was affected by QSO2 mutations (Supplementary Figure 2S). Also, the protein level of the root K⁺ uptake channel AKT1 (Hirsch et al, 1998) was not affected by QSO2 mutations (Supplementary Figure 3S A) and overexpression of QSO2 still confers tolerance to toxic cations in the akt1-1 mutant (Supplementary Figure 3S B).

As QSO2 mutations affect cation accumulation at long times but not the initial rate of uptake, we hypothesized that this QSO regulates cation loading at the root xylem and subsequent cation accumulation at the shoot. The results of Figure 8A and B indicate that, effectively, QSO2 positively regulates shoot accumulation of K⁺, while inhibiting shoot accumulation of toxic cations. Figure 8C and D show the same effect of QSO2 in the case of xylem sap concentrations. Therefore, a plausible mechanism of QSO2 action is the activation of K⁺ efflux at the root symplast–xylem interface while inhibiting the efflux of toxic cations at this location.

The skor-1 mutant is sensitive to toxic cations but overexpression of QSO2 still confers tolerance in this mutant

Although QSO2 could act on several transporters, the possibility exist that its primary target is K⁺ efflux at the symplast–xylem interface, and that activation of this system indirectly inhibits the efflux of toxic cations. The rationale is that activation of K⁺ efflux would hyperpolarize the potential difference across the symplast/xylem boundary (xylem positive), and this elevated potential would inhibit the efflux of toxic cations mediated by nonspecific channels.

One way to test our hypothesis was to compare the phenotypes of the par1-2- and skor-1-null mutants. SKOR is an outward K⁺ channel mediating K⁺ release into the xylem sap (Gaymard et al, 1998). As predicted by our model, the skor-1 mutant is sensitive to toxic cations in addition to K⁺ depletion (Figure 9A and B).

Although this result can be considered as a 'proof of concept' for the mechanism of QSO2 suggested above, SKOR is not the only system loading K⁺ into xylem. Actually, overexpression of QSO2 still confers tolerance to toxic cations and to K⁺ depletion in the skor-1 mutant (Figure 9A and B). Therefore, other unknown systems involved in xylem loading of K⁺ must be affected by QSO2.

Characterization of QSO2 expression and encoded protein

Quantitative RT–PCR analysis of Arabidopsis tissues indicates that QSO2 is mostly expressed in roots (Figure 10A). Transcription in roots is induced about two-fold by K⁺ depletion (Figure 10B), and this regulation is in agreement with the proposed activation by QSO2 of systems loading K⁺ into the xylem.

The localization of expression of more than 22 000 genes in the Arabidopsis root has been described by Birnbaum et al (2003). QSO2 (At1g15020) behaves as a housekeeping gene, because it is expressed at similar levels (within a factor of 2) in different root tissues (stele, endodermis, cortex and epidermis) and zones (meristematic, elongation and mature zone with root hairs).

A QSO2GFP fusion transiently expressed in Nicothiana benthamiana epidermal cells is present at the cell surface (Figure 11A), as expected from its N-terminal signal peptide. Plasmolysis of cells indicates that the fusion protein is associated with the cell wall and not the plasma membrane (Figure 11B). After expression in yeast about 30% of the recombinant QSO2 protein is solubilized by digestion of the cell wall in the process of making yeast protoplasts (Supplementary Figure 4S B). This fraction corresponds to the extracellularly secreted form of the enzyme.

We have purified QSO2 after expression in yeast (Supplementary Figure 4S B). Assay of sulfhydryl oxidase activity demonstrated specificity for dithiol compounds such as dithioerithrytol and dithiothreitol, and insignificant activity (less than 1%) with monothiols such as mercaptoethanol and glutathione. The turnover number measured with simple dithiols (14 min^-1) is within the range of values detected for sulfhydryl oxidases (Levitan et al, 2004). The optimum pH for activity is 7.5, but it has considerable activity (40–60% of optimum) at the pH values reported to prevail at the apoplast (from 5.5 to 6.5; Gao et al, 2004).

Discussion

QSOs are animal and plant enzymes proposed to participate in oxidative folding of disulfide-containing secreted proteins. The induction of human QSO1 when fibroblasts enter quiescence has suggested the name of the family and it has been proposed that the formation of the extracellular matrix may regulate growth (Thorpe et al, 2002). However, the lack of mutants in animal QSOs has prevented demonstration of the physiological roles of these enzymes. One report with neuroblastoma cells with gain and loss of function of a QSO has shown a role in the sensitization of these cells to apoptosis induced by interferon gamma (Wittke et al, 2003).

Our results with mutants of Arabidopsis QSO2 indicate that this QSO regulates cation homeostasis. Arabidopsis QSO2 is a positive regulator, directly or indirectly, of root stellar K⁺ efflux systems involved in xylem loading, and this physiological role is supported by the induction of QSO2 expression under conditions of K⁺ starvation.

Although the K⁺ efflux channel SKOR (Gaymard et al, 1998) and efflux systems for toxic cations (Li⁺, Na⁺, polyamines) could also be regulated, the simplest mechanism of QSO2 action is the activation of a K⁺ efflux system different from SKOR (because QSO2 overexpression still occurs in the skor-1 mutant). The activation of this system probably results from QSO2-mediated oxidation of sulfhydryl groups of the transporter exposed to the external side of the plasma membrane. The opposite effects of QSO2 mutations on the transport of K⁺ and toxic cations can be explained by changes in membrane potential. The activation of K⁺ efflux through this unknown system would hyperpolarize the symplast/xylem boundary, and this membrane potential, positive outside, would inhibit the efflux of toxic cations mediated by other channels. Unfortunately, the membrane potential of xylem parenchyma cells is not easily measured, because of the uncertainty of electrode location after penetrating the epidermal layer. The potential difference across the symplast/xylem boundary can be estimated from the trans-root electrical potential and the root epidermis potential (De Boer and Volkov, 2003), but such measurements have never been made in Arabidopsis. The measured membrane potentials at the symplast/xylem boundary are relatively low, about 80 mV, while the potential of epidermal and cortical cells are much higher, 150–170 mV (De Boer and Volkov, 2003). Therefore it is very likely that the membrane potential of xylem parenchyma cells is low because it is dominated by the 'leak' (channels or electrogenic cotransporters). Although SKOR does not seem to be target of QSO2, we have shown that the skor-1 mutant is sensitive to toxic cations, in addition to K⁺ depletion. This result was predicted by our model and can be considered as a 'proof of concept' for the mechanism of QSO2 suggested above.

We are investigating the nature of the Arabidopsis K⁺ transport system regulated by QSO2, by testing possible epistatic relationships between loss-of-function mutations of different monovalent cation transporters and the overexpression of QSO2. The antiporter SOS1 and many channels of the CNGC family have sulfhydryl groups susceptible of oxidation from the external side of the membrane and are therefore the first candidates.

Although we detect QSO2GFP at the cell wall during transient expression, the regulation of activity or stability of this unknown K⁺ transporter by formation of disulfide bridges (or by possible disulfide isomerase activity of QSO2; see Houston et al, 2005) could also occur during its movement to the plasma membrane. The identification of the K⁺ transporter regulated by QSO2 and electrophysiological studies of the symplast–xylem interface in QSO2 mutants may be crucial for the understanding of monovalent cation homeostasis in plants.

Finally, the observation that a plant QSO regulates ion homeostasis may be of general significance and deserves investigation in the case of animal QSOs (Thorpe et al, 2002).

Materials and methods

Plant material and growth conditions

A. thaliana plants (ecotype Columbia) were grown under greenhouse conditions (16 h light/8 h dark, at 232°C and 705% relative humidity) in pots containing a 1:2 vermiculite:soil mixture. For in vitro culture, seeds were surface sterilized by soaking in 70% ethanol containing 0.1% Triton X-100 for 15 min, followed by commercial bleach (2.5%) containing 0.05% Triton X-100 for 10 min and rinsing three times with sterile water. Stratification of the seeds was conducted during 3 days at 4°C.

The agar medium contained Murashige and Skoog (1962) salts (MS) with 3 or 1% (w/v) sucrose, 10 mM morpholino ethanesulfonic acid and 0.8% (w/v) agar, pH adjusted to 5.7 with Tris base. Different concentrations of norspermidine, spermine, NaCl and LiCl were prepared by adding appropriate amounts of reagents to the basal medium after autoclaving. For liquid culture, the MS medium was prepared without agar, and using 250-ml flasks or suspension-culture-6-well plates (Greiner). Plates were sealed and incubated in a controlled environment growth room at 23°C under long-day conditions (16 h light/8 h dark) at 80–100 mol/m²/s. When appropriate, seedlings (5–7 days old) were transferred to vermiculite:soil mixture and grown to maturity. Plants were irrigated twice a week with nutrient solution during 3 weeks (Naranjo et al, 2003). For stress experiments, NaCl and LiCl were added to the nutrient solution.

Isolation of mutants and genetic analysis

T-DNA 'activation tagging' lines were constructed in the laboratory of D Weigel (Salk Institute, La Jolla, CA) using the pSKI15 vector (Weigel et al, 2000). A total of 16 398 lines (stock numbers N21995 and N21991) was provided by the European Arabidopsis Stock Centre (NASC). The distributed T-DNA pools consisted of T4 seeds. These were screened at high density (170 Petri plates of 9 cm diameter containing 1000 seeds per plate) on MS medium (3% sucrose) with 3.2 mM norspermidine (Sigma). After 7 days, seedlings were considered norspermidine resistant only if they produce fully green expanded cotyledons. Two hundred and ten putative mutants were transferred to soil and grown to maturity. Aliquots of seeds (T5) from the putative mutants were screened again at low seed density (50–100 seeds per 9 cm diameter Petri plate) with 2.8 mM norspermidine. Of the 210 putative mutants, only eight exhibited normal phenotype on agar plates without norspermidine and fully green expanded cotyledons on plates with norspermidine. The best of them was called par1-1D for 'polyamine resistant' gene 1, allele 1, Dominant, following community standards for Arabidopsis genetics (Meinke and Koornneef, 1997).

The par1-1D mutant was crossed with the wild type (Columbia ecotype) by transferring pollen to the stigmas of emasculated flowers. F1 and F2 seeds were scored for germination in 3 mM norspermidine. From the segregating F2 generation, 60 resistant individuals were selected and DNA was extracted (see below) individually to check the correlation between T-DNA presence and resistance to norspermidine.

Plasmid rescue and sequencing

For plasmid rescue, 1 g of plant tissue was harvested from liquid culture and genomic DNA was extracted as described (Weigel and Glazebrook, 2002). DNA (2–3 g) was digested with EcoRI (rescue left T-DNA border) and BamHI (rescue right T-DNA border) in a 40 l reaction volume. After a phenol–chloroform extraction, samples were ligated overnight at 14°C in a total volume of 100 l. Ligated DNA was precipitated with ethanol–sodium acetate and a one-third portion was transformed by electroporation into XL1-MRF Escherichia coli cells.

Plasmids rescued across the left T-DNA border were sequenced with the following primers: 5'LB (5'-AGA TTT CCG AAT TAG AAT AA-3') and T7 (5'-TAA TAC GAC TCA CTA TAG GG -3'); for those with right T-DNA border: 5' ECORI (5'-GAA ATG GAT AAA TAG CCT TGC-3').

Extraction and analysis of nucleic acids

Genomic DNA was extracted from seedlings and young leaves, as described (Weigel and Glazebrook, 2002). For Southern blotting, 3–4 g DNA was digested with BamHI. CaMV 35S enhancer was used as a T-DNA-specific probe. RNA for Northern analysis was extracted from seedlings as described (Ecker and Davis, 1987). RNA (30 g) was resolved by electrophoresis on formaldehyde–agarose gel. The products were transferred by capillary action onto nylon membranes (Amersham) and hybridized with random-priming ³²P-labeled probes (Feinberg and Vogelstein, 1983). Signals were quantified with a Fujifilm BAS-1500 phosphorimager. The QSO2 and TUB (-tubulin) probes were prepared by PCR amplification from genomic DNA of wild-type Columbia plants. Primers were QSO2 (5'-TCC ATT TAA AAA GGC ACG TGA-3' and 5'-CAT CTC TCC TTT TCC CTT TCA-3') and TUB (5'-CCT GAT AAC TTC GTC TTT GG-3' and 5'-GTG AAC TCC ATC TCG TCC AT-3').

Overexpression of QSO2 in transgenic plants

QSO2 cDNA containing the complete open reading frame was obtained from Genomics Science Center, RIKEN, Japan. The cDNA was subcloned into the NotI site of pBS-SK+(Stratagene, USA), yielding to pBS-QSO2. A fragment including the cDNA from QSO2 was subcloned into pBIN121 (Jefferson et al, 1987) by replacing the GUS coding region between the XbaI and SacI sites. This results in a construct for overexpression of QSO2 under the control of the CaMV 35S promoter. The plasmid was introduced into Agrobacterium tumefaciens strain C58C1 by electroporation. A. thaliana (Columbia ecotype) wild-type plants were transformed by flower infiltration (Bechtold et al, 1993). Transgenic plants with 35S:QSO2 construction were screened on MS agar medium containing 50 mg/l kanamycin, and the expression of the transgene was further confirmed by Northern analysis, as described above.

Determination of K⁺, Na⁺ and norspermidine

Five-day-old seedlings were grown in liquid culture for 12 days and then transferred onto a second liquid culture supplemented with NaCl or norspermidine. For sampling at different times, seedlings were collected, rinsed twice briefly with cold 30 mM MgCl₂ and once with cold distilled water. For K⁺ and Na⁺ determination, samples were dried at 50°C for 3–4 days. After dry weight measurement, samples were extracted with 0.1 M nitric acid for 30 min at room temperature, cell debris were removed by filtration and monovalent cations were determined by atomic absorption spectrophotometry (Naranjo et al, 2003). In the case of norspermidine, samples were weighed and frozen in liquid nitrogen, and norspermidine was extracted and determined by HPLC, as described (Bellés et al, 1993).

Measurement of rubidium uptake

Potassium-free medium was prepared with the following ingredients: 2.5 mM Ca(NO₃)₂, 2 mM MgSO₄, 0.1 mM NaFeEDTA, 80 M Ca(H₂PO₄)₂, 25 M CaCl₂, 25 M H₃BO₃, 2 M ZnSO₄, 2 M MnSO₄, 0.5 M CuSO₄, 0.5 M Na₂MoO₄, 0.01 M CoCl₂, 1% sucrose and 2.5 mM MES. NH₄⁺ was added as NH₄H₂PO₄ at final concentration of 2 mM. The pH was adjusted to 5.7 with CaOH.

For measurement of potassium uptake, using Rb⁺ as a tracer, 5-day-old seedlings from MS agar plates were transferred to suspension-culture well containing 7 ml of liquid MS medium. After 10 days, the seedlings were collected, rinsed briefly with water and added to a 7 ml uptake solution containing potassium-free medium supplemented with 1 mM RbCl. The uptake was performed at 23°C under white fluorescent light. At the completion of uptake, the seedlings were rinsed twice with 7 ml of cold 30 mM MgCl₂ and once with water. Finally, the seedlings were treated similar to Na⁺ samples, and Rb⁺ content was determined by atomic absorption spectrophotometry.

Measurements of Li⁺, Na⁺ and K⁺ contents in shoots and xylem

Seeds were grown in MS medium for 5 days and transplanted to soil for 2–3 weeks. Plants were irrigated with NaCl or LiCl solutions as indicated, shoots collected, weighed and dried at 50°C for 3–4 days. Ions were extracted with 0.1 M nitric acid for 30 min at room temperature, cell debris were removed by filtration, ion content was determined by atomic absorption spectrophotometry.

Xylem sap was obtained from pressurized shoot of mature plants grown in MS medium for 5 days, and transplanted to soil for 3–4 weeks. Plants were then subjected to 50 mM NaCl or 10 mM LiCl by a single irrigation with nutritive solution for 3 or 2 days, respectively. The inflorescence stem was cut with a very sharp razor blade and placed in a pressure chamber, with the rest of the plant emerging out. Pneumatic pressure (10–30 bars) was applied. The first two drops emerging were discarded to prevent contamination of the xylem sap with contents from damaged cells or phloem sap. Ion content was determined by atomic absorption spectrophotometry.

Selection of At1g15020 and At3g02850 knock-out Arabidopsis lines

Three At1g15020 (QSO2) Arabidopsis independent knockout mutant lines (SALK_066130, SALK_025237 y SALK_072829) and one At3g02850 (SKOR) Arabidopsis knock-out mutant line (SALK_132944) were obtained from the Salk Institute Genomic Analysis Laboratory (<http://signal.salk.edu/cgi-bin/tdnaexpress>http://signal.salk.edu/cgi-bin/tdnaexpress). The seeds were sown and grown on MS medium with 50 g/ml of kanamycin for 6 days, resistant seedlings were transplanted to soil for 3 weeks, after which genomic DNA was extracted from the individual plants. Homozygous knockout mutant plants were selected by PCR using three primers. In all the cases, a T-DNA-specific primer pROKLbb1 5'-GCGTGGACCGCTTGCTGCAAC T-3' was used. For each line, two specific primers were used: SALK_066130 (par1-2LP 5'-TCAAGGCTCAGCAGACCCAAC-3' and par1-2RP 5'-TGCACATGATCGATACTTTTTGGTG-3') SALK_025237 (par1-3LP 5'-TTTGTTTCTCAACGGAAGGAG-3' and par1-3RP 5'-TTTAGGAGGAGCCCAAAAGAG-3'), SALK_072829 (par1-4LP 5'-CTTGTTGTTGGGTCTGCTGAG-3' and par1-4RP 5'-TGCCTATCTGCTTGTTGATCC-3') and SALK_132944 (skorLP 5'-TCAAGAATCTTTAGATCCGGACAGA-3' and skorRP 5'-ACAGGCGCCAATTTT AGGCAT-3').

Quantitative real-time RT–PCR analysis

Plant organs were harvested from four-week-old flowering plants grown in greenhouse. Organs were sampled as follows: opened flowers, all cauline leaves, a 1-cm section of the inflorescence stem and rosette leaves. Roots were sampled from 2-week-old seedlings grown vertically on MS. For the measurement of induced levels of QSO2, plants were grown on MS medium supplemented with 1% sucrose. After 5 days, 5–7 seedlings were transferred to liquid culture for 12 days, and then were either mock or treated with 2.8 mM spermidine, 25 mM LiCl, 120 mM NaCl, 10 mM H₂O₂, 10 M ABA and K⁺-free medium for 16 h. All plant tissues were frozen in liquid nitrogen. Total RNA was isolated as described (Verwoerd et al, 1989). For reverse transcription, 2 g of RNA previously treated with DNase I was incubated with Buffer RT 1 (Fermentas), 1 mM dNTPs, 0.2 g oligo(dT)₁₅ primer and 10 U M-MuLV reverse transcriptase (Fermentas), to finally obtain a 40 l cDNA solution. The sequence of the primers used for PCR amplifications were as follows: for QSO2 (At1g15020) 5'-AGG ACG CAG CAG AAG AAC CGG-3' and 5'-CAT CTC TCC TTT TCC CTT TCA-3', and for actin 8 (At1g49240) 5'-AGT GGT CGT ACA ACC GGT ATT GT-3' and 5'-GAG GAT AGC ATG TGG AAG TGA GAA-3'. The real-time PCR was performed in a total volume of 20 l, including 0.5 l of cDNA and gene-specific primers (500 nM final concentration). Amplification of PCR products was monitored via intercalation of Eva-green (Biotium), using an ABI PRISM 7000 sequence detection system (Perkin-Elmer Applied Biosystems). Amplification was performed for 40 cycles and the relative quantification of gene expression data in different treatments was carried using the comparative C_t method (Livak and Schmittgen, 2001). Expression levels were normalized using the C_t values obtained for the actin 8 gene. The relative transcript level for QSO2 was calculated by normalizing to actin 8 as follows: RTL=2^-C_t, where C_t=C_t(QSO2)–C_t(actin 8). The presence of a single PCR product was confirmed by dissociation analysis and agarose gel electrophoresis. Three independent replicate experiments were conducted.

Construction of QSO2:GFP fusion for expression in plants

The open reading frame of QSO2 gene used in this fusion was obtained by PCR from pBS-QSO2 construct (see above), using primers QSO2XbaI 5'-TGC TCT AGA GTA ATA AAC ACC AGA GAC CTT-3' and QSO2BamHI 5'-GCG GGA TCC TCT CTC CTT TTC CCT TTC ACT-3'. The amplified DNA was cloned into the XbaI and BamHI sites of a pBS-GFP vector (Chiu et al, 1996). The new construction pBS-QSO2GFP harbored a translational fusion between the QSO2 and GFP coding sequences. The QSO2GFP coding sequence and the NOS terminator was subcloned into XbaI–EcoRI doubly digested pBI121. In this new construct (pBI121-QSO2GFP), the expression of QSO2:GFP fusion was driven by 35S promoter. It was used to transform A. tumefaciens (strain C58C1).

Agrobacterium-mediated transient gene expression in N. benthamiana

Agrobacterium-based transient transformation was made as described in Wieland et al (2006) with some modifications. Briefly, A. tumefaciens culture was grown until saturation in LB medium (10 ml) at 28°C for 16 h. The culture was centrifuged and resuspended in the same volume of 10 mM MgCl₂, 10 mM MES pH 5.6 and 200 M acetosyringone. The culture was kept at room temperature for 3 h, without shaking. A syringe (without needle) was used to inject the Agrobacterium culture into the leaf. Plants were grown for 3–4 days in greenhouse before GFP localization was performed. To plasmolyze cells, the tissue samples were soaked in 1 M mannitol on glass slides for 30 min at room temperature. Fluorescence of GFP was observed by a Leica TCS-SL confocal microscope and laser scanning confocal imaging system. A 488 nm excitation wavelength and a 510 nm emission wavelength were used.

Construction of His-tagged QSO2 for expression in yeast

The coding region of the QSO2 cDNA was amplified by PCR from the pBS-QSO2 construct (see above), using primers QSO2-MssI5 5'-TTA TTA GTT TAA ACG TAA TAA ACA CCA GAG AC-3' and QSO2-MssI3 5'-TAT ATA GTT TAA ACT TTC TCT CCT TTT CCC-3'. The PCR product was subcloned into the PmeI site of pCM262 vector, yielding the pCM262-QSO2 construct. pCM262 vector is derived from pCM190 (Gari et al, 1997), and it contains tetracycline-regulable promoter, three copies of the HA epitope and 6 histidine tail fused to the C-terminus of the target gene. Once the pCM262-QSO2 construct was verified by sequencing, it was used to transform Saccharomyces cerevisiae strain W303 generating SA76 strain.

Purification of His-tagged QSO2 from yeast

Yeast SA76 strain was grown in minimal medium (2% sucrose, 0.7% yeast nitrogen base and 50 mM succinic acid pH 5.5) to an absorbance at 660 nm of 0.4–0.5, and cells were harvested by centrifugation, suspended in homogenization buffer (50 mM Tris–HCl, pH 7.6, 0.1 M KCl, 10% sucrose and protease inhibitor cocktail) and lysed by vortexing with glass beads (0.5 mm). The lysate was centrifuged first at 2000 r.p.m. for 5 min at 4°C and further centrifuged at 13 000 r.p.m. (16 000 g) for 30 min at 4°C. The final supernatant was used for purification.

Ni²⁺ affinity resin (His-Bind Resin, Novagen) was packed onto a column and washed with 3 ml H₂0 and 4 ml charge buffer (50 mM NiSO₄). After equilibration with 3 ml binding buffer (5 mM imidazole, 0.5 M NaCl, 20 mM Tris–HCl, pH 7.9) the centrifugated yeast extract was loaded onto the column and washed with 8 ml of binding buffer and 4 ml washing buffer (60 mM imidazole, 0.5 M NaCl, 20 mM Tris–HCl, pH 7.9). His-tagged QSO2 was eluted with 10 ml of elution buffer (0.3 M imidazole, 0.5 M NaCl, 20 mM Tris–HCl, pH 7.9). The peak fractions (0.5 ml) were dialyzed against 50 mM Tris–HCl, pH 8 and used immediately in a assay of sulfhydryl oxidase activity.

SDS–PAGE and protein immunoblotting

Protein content was measured with dye binding method (Bradford, 1976). BSA was employed as the protein standard. A 5 g weight of total protein was resuspended with Laemmli (1970) buffer and loaded onto 8% (w/v) linear acrylamide minigels. After electrophoresis, the gels were prepared for immunoblotting. SDS–PAGE-separated proteins were electrophoretically transferred onto nitrocellulose membranes (Millipore). Protein content on the membrane was detected by Ponceau S staining. Following transfer, membrane was blocked with Tris-buffered saline (100 mM Tris, 150 mM NaCl) containing 0.1% (v/v) Tween 20 and 2% non-fat milk powder, for 30 min on a rocker at room temperature. Blocked membrane was incubated overnight at 4°C on a rocker with anti-HA (1/10 000), the membrane was washed five times with Tris-buffered saline (100 mM Tris, 150 mM NaCl) containing 0.1% (v/v) Tween 20 for 10 min each on a rocker at room temperature, followed by the addition of a 1:5000 dilution of anti-mouse secondary antibody. Anti-HA blots were visualized using the ECL detection system (Amersham).

Enzymatic assay of QSO2 activity

Purified QSO2 protein (corresponding to 20 pmol/reaction) was incubated in 0.1 ml of phosphate-buffered saline buffer (75 mM potassium phosphate buffer pH 7.5 and 3 mM EDTA), together with reduced substrates corresponding to 50 nmol of reduced thiol groups. All the reactions were carried out for 5 min at 25°C. QSO2 activity was measured by determination of thiol content, as described (Levitan et al, 2004). Results are expressed as turnover numbers (per min).

Supplementary data

Supplementary data are available at The EMBO Journal Online (http://www.embojournal.org).

Acknowledgements

This work was supported by grants BFU2005-06388-C04-01 of the Spanish MEC (Madrid) and CPE03-006-C6-4 of the Spanish INIA (Madrid).

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