Enhanced salt tolerance mediated by AtHKT1 transporter-induced Na+ unloading from xylem vessels to xylem parenchyma cells


(fax +81 52 789 5206; e-mail uozumi@agr.nagoya-u.ac.jp; fax +1 858 534 7108; e-mail julian@biomail.ucsd.edu).


AtHKT1 is a sodium (Na+) transporter that functions in mediating tolerance to salt stress. To investigate the membrane targeting of AtHKT1 and its expression at the translational level, antibodies were generated against peptides corresponding to the first pore of AtHKT1. Immunoelectron microscopy studies using anti-AtHKT1 antibodies demonstrate that AtHKT1 is targeted to the plasma membrane in xylem parenchyma cells in leaves. AtHKT1 expression in xylem parenchyma cells was also confirmed by AtHKT1 promoter–GUS reporter gene analyses. Interestingly, AtHKT1 disruption alleles caused large increases in the Na+ content of the xylem sap and conversely reduced the Na+ content of the phloem sap. The athkt1 mutant alleles had a smaller and inverse influence on the potassium (K+) content compared with the Na+ content of the xylem, suggesting that K+ transport may be indirectly affected. The expression of AtHKT1 was modulated not only by the concentrations of Na+ and K+ but also by the osmolality of non-ionic compounds. These findings show that AtHKT1 selectively unloads sodium directly from xylem vessels to xylem parenchyma cells. AtHKT1 mediates osmolality balance between xylem vessels and xylem parenchyma cells under saline conditions. Thus AtHKT1 reduces the sodium content in xylem vessels and leaves, thereby playing a central role in protecting plant leaves from salinity stress.


Soil salinity is a major problem for agricultural productivity worldwide. High concentrations of sodium (Na+) in soils disturb ionic intracellular homeostasis, causing membrane dysfunction and inhibition of cellular metabolism, which leads to adverse effects on cell division, growth, photosynthesis and development (Flowers, 1999; Horie and Schroeder, 2004; Omielan et al., 1991). The absorption of inorganic ions takes place through root hairs, the epidermis and the cortex. Nutrients are further transported to root xylem parenchyma cells. Cations such as Na+ and K+ are loaded into the xylem vessels and are transported upward throughout the plant via the transpiration stream (De Boer and Volkov, 2003; Lacan and Durand, 1996; Raven et al., 1992; Wegner and De Boer, 1997; Wegner and Raschke, 1994). Ions that are unloaded by shoot/leaf xylem parenchyma cells can be transported into phloem sieves via symplastic diffusion (Alberts et al., 2002). Because the access to xylem and phloem tissues is difficult, the precise functions of many of the transporters that regulate long-distance transport of ion nutrients have remained largely unresolved.

The AtHKT1 channel/transporter is Na+ selective in Arabidopsis thaliana (Mäser et al., 2002a; Uozumi et al., 2000). Null mutations in AtHKT1 (Gong et al., 2004; Mäser et al., 2002b) and defective mutant alleles of AtHKT1 (Berthomieu et al., 2003) have been shown to cause excessive accumulation of Na+ in aerial parts of mature Arabidopsis plants, which results in chlorosis and inhibition of growth. Expression of AtHKT1 has been found in vascular tissues in leaves (Mäser et al., 2002b) and in particular in phloem tissues (Berthomieu et al., 2003). HKT transport proteins have been described in several plant types (Fairbairn et al., 2000; Garciadeblas et al., 2003; Golldack et al., 2002; Horie et al., 2001; Rubio et al., 1995; Schachtman and Schroeder, 1994; Su et al., 2003; Uozumi et al., 2000). AtHKT1 belongs to the Ktr/Trk/HKT superfamily of K+ transporters (Durell and Guy, 1999; Mäser et al., 2002a; Uozumi, 2001). Studies of the structure and function of AtHKT1 have determined its membrane topology (Kato et al., 2001) and ion selectivity filter, forming residues in the pore-forming regions (Mäser et al., 2002a). Two sets of Ktr systems, KtrAB and KtrCD, in Bacillus subtilis are important for the acquisition of Na+ tolerance (Holtmann et al., 2003). Recently, in the cyanobacterium Synechocystis sp. PCC 6803 the Ktr system was found to be essential for adaptation not only to salt stress but also to higher osmolality (Matsuda et al., 2004). The function of Ktr/Trk/HKT transporters indicates that these transporters may alleviate osmolality stress caused by higher salt concentrations.

The present study was undertaken to determine which cellular membrane AtHKT1 is targeted to and to explore the tissue and cell-type localization of AtHKT1. We further investigated the physiological functions of AtHKT1 in Na+/K+ transport in the xylem and phloem using athkt1 mutants. Here we demonstrate the plasma membrane localization of AtHKT1 in xylem parenchyma cells and show a major function of AtHKT1 in mediating unloading of Na+ from xylem vessels, thus protecting Arabidopsis leaves from sodium stress.


Immunological detection of AtHKT1 in the plasma membrane of xylem parenchyma cells

The membrane targeting of AtHKT1 or monocot HKT homologs has not yet been analyzed. To directly determine the location of AtHKT1 protein expression at the subcellular and plant cell-type levels, anti-AtHKT1 antibodies were generated. The hydrophilic 15-amino-acid sequence, located in the first pore-forming region, PA (Kato et al., 2001; Mäser et al., 2002a), was chosen as an epitope. The specificities of the anti-AtHKT1 antibodies were tested using AtHKT1 epitope–GST (glutathione-S-transferase) fusion constructs expressed in Escherichia coli or insect cells confirming antibody specificity (Supplementary Figure S1). Since the AtHKT1 antibodies probed against Arabidopsis plant extracts did not show any interaction, we predicted that AtHKT1 expression was restricted to a few cell types and/or the amount of natively expressed AtHKT1 protein was relatively small (data not shown). Therefore, we analyzed the tissue and subcellular localization of the AtHKT1 protein by immunoelectron microscopic studies. A cross-section of the vascular tubes of leaves of Arabidopsis plants grown in 1/2 MS medium showed immunological detection in the plasma membrane of xylem parenchyma cells when the antiserum was used at a 1:20000 dilution (Figure 1). Gold particles were located at the plasma membrane of xylem parenchyma cells, but not significantly found in other cellular membranes (Figure 1c). There were no significant immunological detection signals in other cells of the leaf. The plasma membrane localization of AtHKT1 was also supported by expression of green fluorescent protein (GFP)–AtHKT1 fusions in tobacco BY2 cells. These cells showed a GFP signal localized in the region of the plasma membrane compared with control tobacco cells expressing only the GFP protein, which showed expression throughout the cell and nucleus (Supplementary Figure S2).

Figure 1.

Immunological detection of AtHKT1.
Cross-section of Arabidopsis thaliana leaf immunostained using an anti-AtHKT1 antibody. (b) and (c) are enlarged sections of (a). The expression of AtHKT1 indicated by the appearance of gold particles [arrows in (b) and arrowheads in (c)] was detected in the xylem parenchyma cells (XP) but not in vessel tubes of xylem cells (Ve) or phloem cells (Ph). AtHKT1 was detected in the plasma membrane (PM), but AtHKT1 was not detected in the vacuole (V), the chloroplast (Chl) or the mitochondria (Mit).

Complementation of athkt1 alleles

As reported previously, athkt1 mutant plants showed hypersensitivity to Na+ under saline conditions (Berthomieu et al., 2003; Mäser et al., 2002b). We transformed the athkt1-3 mutant (Mäser et al., 2002b) with an AtHKT1 cDNA under the control of an 837 bp AtHKT1 promoter. Wild-type (Ws background), athkt1-3 and homozygous athkt1-3 mutant lines expressing the AtHKT1 cDNA were grown in media containing 75 mm NaCl (Na+ stress condition). The athkt1-3 mutant plants showed an increased Na+ sensitivity compared with wild-type plants (Figure 2a,b), due to over-accumulation of Na+ in shoots of athkt1 mutant plants (Mäser et al., 2002b). In contrast, introduction of AtHKT1 cDNA into athkt1-3 plants led to complementation of the Na+ sensitivity of athkt1-3 plants (Figure 2c). These data demonstrate complementation of athkt1 and suggest that the AtHKT1 promoter construct can be used for promoter–β-glucuronidase (GUS) gene analyses of tissue and cellular AtHKT1 localization.

Figure 2.

AtHKT1 cDNA expression in athkt1-3 complements the salt sensitive phenotype.
Wild type Ws (a), athkt1-3 plants (b) and the transgenic athkt1-3 lines expressing AtHKT1 (c) were analyzed for their salt sensitivity. After 5 days’ growth of 7-day-old seedlings on MS medium, plants were transferred onto minimal medium containing 75 mm NaCl (Mäser et al., 2002b). Pictures were taken after 16 days of incubation.

Expression of AtHKT1 in xylem parenchyma cells in leaves

To further test whether AtHKT1 is expressed in xylem parenchyma cells (Figure 1), we analyzed ProAtHKT1:GUS expression lines in transgenic Arabidopsis. Earlier experiments (Berthomieu et al., 2003; Mäser et al., 2002b) using transgenic ProAtHKT1:GUS plants had indicated that the AtHKT1 promoter is active in the vascular tissues of the root and leaf, but the specific expression in xylem parenchyma cells had not been investigated. Transgenic ProAtHKT1:GUS plants were grown for 15 days in 1/2 MS medium and leaves were analyzed. The cells adjacent to xylem vascular tubes, which were visible as spiral structures, were strongly stained, suggesting AtHKT1 expression in xylem parenchyma cells (Figure 3a,b). In addition, weaker GUS signals were observed in other cells in vascular tissues, consistent with AtHKT1 expression in additional cell types (Berthomieu et al., 2003). In leaf cross-sections strong GUS staining was observed in xylem parenchyma cells and weaker GUS staining was observed close to phloem vessels (Figure 3c). In roots GUS activity was generally observed in cells inside the endodermis (Figure 3d).

Figure 3.

Detection of GUS activity in xylem parenchyma cells.
(a) Arabidopsis thaliana plants expressing the GUS gene under the control of the AtHKT1 promoter were grown in 1/2 MS media containing 0.8% agar and 2% sucrose for 14 days. Strong blue GUS staining was detected in the cells surrounding xylem leaf tissues (a, b and c). GUS activity was detected in the vicinity of the xylem as well as the phloem in leaves (c), and inside the endodermis in roots (d). (b) is a magnification of (a).

Na+ accumulation in the xylem sap in AtHKT1-null plants

Since both antibody localization (Figure 1) and ProAtHKT1:GUS analyses (Figure 3) showed clear AtHKT1 localization in xylem parenchyma cells, we next examined the Na+ and K+ content in the vascular vessels in athkt1 loss-of-function mutants under saline and control conditions. The Na+ and K+ content in xylem and phloem sap of wild-type and athkt1 mutant plants grown with or without 75 mm Na+ were measured. Interestingly, disruption of AtHKT1 in both the athkt1-3 (Ws) and Columbia backgrounds, athkt1-1 and the fast neutron athkt1 disruption allele FN1148 (Gong et al., 2004), caused substantial increases in the Na+ content of the xylem sap (Figure 4a,b; P < 0.001). In the absence of Na+ stress lower levels of Na+ accumulated in the xylem sap of soil-grown plants. Nevertheless, under these control conditions significant increases in the Na+ content of athkt1 xylem sap samples were also found compared with the corresponding wild-type alleles. The average Na+ content of xylem sap samples derived from non-Na+-stressed Col wild type was 0.31 ± 0.05 μmol ml−1 sap (n = 3), whereas that of athkt1-1 was 3.38 ± 0.79 μmol ml−1 sap (n = 3) and that of FN1148 was 2.99 ± 0.32 μmol ml−1 sap (n = 4) (P < 0.03 for Col wild type and athkt1-1; and P < 0.001 for Col wild type and FN1148). Furthermore, the average Na+ content of xylem sap samples derived from Ws wild type was 1.69 ± 0.5 μmol ml−1 sap (n = 3), whereas that of athkt1-3 was 6.18 ± 1.43 μmol ml−1 sap (n = 4) (P < 0.005). These data suggest that AtHKT1 also functions in lowering Na+ in the xylem sap under non-stress conditions. A previous study had indicated that the sas2-1 (athkt1) non-loss-of-function allele did not affect the Na+ content in the xylem (Berthomieu et al., 2003). The Na+ content in the phloem sap was lower in all three athkt1 alleles than in wild-type controls (Figure 4c, P < 0.04 for athkt1-3; and Figure 4d, P < 0.001 for athkt1-1 and FN1148), which was also observed in the sas2-1 mutant allele (Berthomieu et al., 2003). These data demonstrate that AtHKT1 loss-of-function mutations cause not only a reduction in the Na+ content of the phloem sap but also a substantial increase in the Na+ content of xylem sap in athkt1 mutant alleles (Figure 4a,b). These findings suggest a new model by which AtHKT1 controls Na+ content in the xylem as well as in the phloem sap (see Discussion).

Figure 4.

AtHKT1 mutation causes an increase in the Na+ content of xylem sap and a decrease in the Na+ content of phloem sap under salt stress.
Soil-grown plants were subjected to 75 mm NaCl with one-twentieth MS salts for 2 days after bolting. The Na+ content in xylem sap extracted from wild-type Ws and athkt1-3 plants (n = 5) (Mäser et al., 2002b) (a) and from wild-type Col, athkt1-1 and FN1148 plants (n = 4) (Gong et al., 2004) (b). The Na+ content in phloem sap extracted from wild-type Ws and athkt1-3 plants (n = 3) (c) and from wild-type Col, athkt1-1 and FN1148 plants (n = 6) (d). Error bars represent ±SD.

We further analyzed the K+ content of xylem and phloem sap in wild-type and athkt1 mutant plants. In athkt1 mutants, the K+ content of xylem sap decreased slightly when soil-grown plants were exposed to 75 mm NaCl (Figure 5a, P < 0.01 for athkt1-3; Figure 5b, P < 0.01 for both athkt1-1 and FN1148). In addition, when plants were grown under hydroponic conditions and 10 mm NaCl was added to bolting mature plants for 3 days, a reduction in the K+ content of xylem sap was also observed in athkt1-1 and athkt1-3 mutant plants compared with wild type (data not shown; n = 6–10 experiments, P < 0.004). In control experiments in the absence of Na+ stress, no significant reduction was found in K+ content in xylem sap samples of athkt1 mutant plants in comparison with wild-type samples. Average xylem sap K+ content concentrations derived from WS wild type and athkt1-3 were 14.43 ± 1.59 (n = 3) and 12.32 ± 1.62 μmol ml−1 sap (n = 4) (P > 0.14), respectively, whereas those from Col wild type, athkt1-1 and FN1148 were 7.80 ± 1.11 (n = 3), 10.65 ± 3.71 (n = 3) and 8.81 ± 0.44 μmol ml−1 sap (n = 4), respectively (P > 0.30 for Col -wild type and athkt1-1 and P > 0.25 for Col wild type and FN1148). These data support a model in which the changes in K+ content of xylem sap result indirectly from AtHKT1-mediated Na+ transport (see Discussion).

Figure 5.

Effect of AtHKT1 mutation on K+ content in xylem sap and in phloem sap under salt stress.
Soil-grown plants were subjected to 75 mm NaCl with one-twentieth MS salts for 1 day [(a) and (c)] or for 2 days [(b) and (d)] after bolting. The K+ content in xylem sap extracted from wild-type Ws and athkt1-3 plants (n = 3) (a) and from wild-type Col, athkt1-1 and FN1148 plants (n = 4) (b). The K+ content in phloem sap extracted from wild-type Ws and athkt1-3 plants (n = 4) (c) and from wild-type Col, athkt1-1 and FN1148 plants (n = 4) (d). Error bars represent ±SD.

Unlike Na+ content, the K+ content of the phloem sap was not significantly affected by the athkt1 mutations compared with that of wild-type controls (Figure 5c,d). Consistent with the K+ levels of xylem sap (Figure 5a,b), the K+ content of shoots in athkt1 mutants grown under salt stress (75 mm NaCl) was revealed to be lower than the K+ content of wild-type plants (Figure 6a, P < 0.03 for athkt1-3; Figure 6b, P < 0.001 for athkt1-1 and FN1148). In addition the K+ content of roots was also affected in a manner such that athkt1 mutants maintain a higher K+ content than wild-type plants (Figure 6c, P < 0.03 for athkt1-3; Figure 6d, P < 0.001 for athkt1-1 and FN1148). Interestingly, the effects of athkt1 mutations on the K+ content of roots and shoots are opposite to the Na+ content phenotypes for roots and shoots (Mäser et al., 2002b), suggesting a coupling of Na+ unloading via AtHKT1 with K+ loading from xylem parenchyma cells under salt stress (see Discussion).

Figure 6.

K+ homeostasis is disturbed in athkt1 plants under salt stress.
The K+ content was reduced in shoots of athkt1 mutant alleles (a and b). The K+ content was increased in roots of athkt1 mutant alleles (c and d) [n = 6 for (a) and (c); n = 8 for (b) and (d)]. Six- to 7-day-old seedlings were transferred onto plugs and grown hydroponically in minimal medium (Mäser et al., 2002b) for 3 weeks with a solution change every 3 days. Then plants were subjected to 75 mm NaCl stress for 2 days. Error bars represent ±SD.

Osmolality in the medium modulates AtHKT1 expression

Because the present study suggests that AtHKT1 functions in the unloading of Na+ from xylem vessels into xylem parenchyma cells, we further investigated whether AtHKT1 may be regulated by ion concentrations and osmolality in the medium. Arabidopsis plants bearing ProAtHKT1:GUS constructs were germinated on 1/2 MS medium for 14 days. Then, seedlings were transferred into fresh synthetic medium containing basic components as described in Materials and Methods, with different concentrations of Na+, K+, sorbitol or mannitol. For Na+ treatments, K+ was supplied at 1 mm KCl, whereas Na+ was supplied at 0, 15, 30, 45, 60 or 75 mm NaCl (Figure 7a). For K+ treatments, Na+ was supplied at 1 mm NaCl, whereas K+ was supplied at 0, 15, 30, 45, 60 or 75 mm KCl (Figure 7b). For sorbitol or mannitol treatments, either compound was supplied at concentrations of 0, 50, 100, 150 or 200 mm in a medium containing 1 mm NaCl and 1 mm KCl (Figure 7c,d). Both Na+ (Figure 7a) and K+ (Figure 7b) induced the expression of AtHKT1 in leaves at concentrations up to 30 mm. The magnitude of the induction due to addition of both ions decreased at concentrations of more than 40 mm, and was similar to baseline levels at 75 mm in leaves (Figure 7a,b). These data provide evidence that AtHKT1 promoter activity is regulated by the Na+ content as well as by the K+ content in the growth medium. In addition, under stress, the intensity of the GUS signals also increased in the vein and at the edge of leaves compared with GUS signals under non-stress conditions (data not shown). The expression of AtHKT1 mRNA in roots and in shoots of plants was also examined by short-cycle (24 cycles) reverse transcriptase polymerase chain reaction (RT-PCR) (Figure 7e). Levels of AtHKT1 mRNA showed increases in the presence of 30 mm Na+ or K+ compared with levels found without supplementation of Na+ or K+ and in the presence of 75 mm Na+ or K+. Note that only 24 PCR cycles were used to analyze whether AtHKT1 mRNA levels were increased, and therefore baseline levels of AtHKT1 mRNA were below detection levels under low- and high-solute conditions (Figure 7e).

Figure 7.

Induction of AtHKT1 expression by elevated osmolality.
Two transgenic ProAtHKT1:GUS Arabidopsis lines (lines 2-1 and 4-7) were grown in 1/2 MS media for 14 days. The seedlings were transferred to fresh synthetic media containing 1 mm KCl and 1 mm NaCl. NaCl (a) or KCl (b) was supplied at concentrations of 0, 15, 30, 45, 60 and 75 mm. Sorbitol (c) or mannitol (d) was supplied at concentrations of 0, 50, 100, 150 and 200 mM. Plants were grown under these conditions for 14 days. GUS activity from roots (open symbols) or from shoots (filled symbols) was measured. Transgenic lines 2-1 (circles) and 4-7 (triangles) were used for this test. Error bars represent standard error.
(e) RT-PCR was performed using total RNA extract from roots (R) and shoots (S) of wild-type Arabidopsis, which were grown under the same conditions as in panels (a–d). The concentrations of NaCl, KCl, sorbitol and mannitol are indicated. b-TUB is the β-tubulin gene.

The cyanobacterium Ktr/Trk/HKT-type transporter is involved in osmoadaptation to higher osmolality stress (Matsuda et al., 2004). The effect of the non-ionic compounds sorbitol and mannitol on ProAtHKT1:GUS activity was tested (Figure 7c,d). Sorbitol induced an increase in GUS expression at 100 mm (Figure 7c). The GUS activity decreased at concentrations of more than 100 mm. On the other hand, data from short-cycle RT-PCR experiments show that the amount of AtHKT1 transcript was highest for 50 mm sorbitol but not 0 and 100 mm (Figure 7e). In the case of mannitol, the highest induction of GUS activity occurred at 50 mm (Figure 7d), and the expression exhibited a decrease at 100 mm mannitol. This tendency was also observed for the expression of AtHKT1 mRNA (Figure 7e). The upregulation of AtHKT1 ProAtHKT1:GUS activity in roots by the addition of sorbitol and mannitol was smaller than that in shoots. These data show that the AtHKT1 promoter is active at increased osmolality in the medium.


AtHKT1 is localized to the plasma membrane of xylem parenchyma cells

Prior to the present study, the membrane and cell targeting of plant HKT transporter proteins has remained largely unknown at the protein level due to the difficulty in obtaining antibodies to these highly hydrophobic proteins. In this study, AtHKT1-specific antibodies were generated against the peptides corresponding to the first pore domain of AtHKT1 (Kato et al., 2001; Mäser et al., 2002a). Immunoelectron microscopy using the antibodies showed the presence of AtHKT1 in the plasma membrane of xylem parenchyma cells in leaves (Figure 1). Plasma membrane localization of the mesenbryanthemum McHKT1 was recently detected; this also shows expression in xylem parenchyma cells (Su et al., 2003). Based on the immunological evidence, we examined the expression of AtHKT1 using ProAtHKT1:GUS plants and confirmed the GUS signals in the cells adjacent to xylem vascular cells (Figure 3). The expression of AtHKT1 is likely to be dependent on environmental conditions such as high salt or high osmolality (Figure 7).

AtHKT1 functions in removing Na+ from the xylem sap and enhancing Na+ content in phloem

Disruption mutations in AtHKT1 of both the Ws and Col Arabidopsis ecotypes resulted in dramatic increases in the Na+ content of xylem sap samples (Figure 4a,b), which can account for the increased accumulation of Na+ in aerial parts in athkt1 mutants (Alberts et al., 2002; Berthomieu et al., 2003; Mäser et al., 2002b). Na+ loaded into xylem parenchyma cells by AtHKT1 in wild-type plants may be transferred to the phloem (Alberts et al., 2002; Raven et al., 1992) and thus be recycled to the roots (Berthomieu et al., 2003) (Figure 8). The reduced uptake of Na+ into xylem parenchyma cells in athkt1 mutants may thus contribute to the reduced Na+ loading of the phloem as found here (Figure 4c,d), consistent with results showing low levels of Na+ in phloem sap in the sas2-1 allele (Berthomieu et al., 2003). Since lower GUS signals were observed in the vicinity of phloem tissues here (Figure 3c), our data do not exclude an additional function for AtHKT1 in phloem cells (Berthomieu et al., 2003). However, the presented data provide several lines of evidence that unloading of Na+ in the xylem sap is a major protective function of AtHKT1 during salinity stress in Arabidopsis.

Figure 8.

Model of AtHKT1 function in the removal of sodium from the xylem and recirculation of Na+ to roots via the phloem and protection of the leaf from sodium stress.
Sodium (Na+) is taken up by the root system. AtHKT1, which is operating in xylem parenchyma cells, unloads Na+ from xylem vessels into xylem parenchyma cells. Subsequently, Na+ moves through plasmodesmata from xylem parenchyma cells into the phloem via symplastic diffusion. The increase in Na+ accumulation in xylem sap of athkt1 mutants thus causes increased salinity stress in leaves, and decreased Na+ accumulation into xylem parenchyma cells can result in reduced Na+ loading of the phloem. In addition AtHKT1 has been proposed to mediate transport of Na+ into the phloem (Berthomieu et al., 2003). Na+/H+ antiporters in the plasma membrane (SOS1) and in the tonoplast (AtNHX1) are shown (Apse et al., 1999; Shi et al., 2002).

In the present study the collected xylem sap samples can be considered as the root exudate of loss-of-function athkt1 alleles (see Experimental procedures). On the other hand, xylem sap from the non-loss-of-function sas2-1 athkt1 allele was collected from shoots using a pressure chamber (Berthomieu et al., 2003). The methodological analyses of loss-of-function in the present study versus non-loss-of-function (Berthomieu et al., 2003) may lead to the observed differences in Na+ content in xylem sap between these two studies. Taken together, the immunolocalization of AtHKT1 in xylem parenchyma cells (Figures 1 and 3) and the increases in levels of Na+ in root exudate xylem sap in loss-of-function athkt1 alleles (Figure 4) reveal a physiological function for AtHKT1 in regulating the concentration of Na+ in the xylem sap.

Because osmolality is one of the major components in salinity stress, AtHKT1 may be involved in the adaptation of plants to hyperosmolality. Moderate osmolality stress (about 30 mm NaCl/KCl, about 50–100 mm sorbitol or about 50 mm mannitol) enhanced the intensity of GUS activity in leaves (Figure 7). These data and RT-PCR experiments suggest that at the level of transcription an enhancement of AtHKT1 promoter activity is regulated by hyperosmotic stress. AtHKT1-mediated transport of Na+ into xylem parenchyma cells may control the balance of ion concentrations and osmolality between xylem vessels and xylem parenchyma cells (Raven et al., 1992).

Experiments using the Xenopus laevis oocyte expression system showed a hypo-osmotic response of EcHKT1 and EcHKT2 from Eucalypus camaldulensis (Liu et al., 2001). The orthologs of HKT, Ktr transporters, are involved in osmoadaptation in Bacillus subtilis (KtrAB and KtrCD) and Synechocystis sp. PCC 6803 (KtrABE) (Holtmann et al., 2003; Matsuda et al., 2004). The Synechocystis KtrABE mutant could not grow under conditions of high Na+ and high osmolality (Matsuda et al., 2004). It was found that high osmolality induced loss of K+ from cells. KtrABE mediates Na+-stimulated K+ uptake to compensate for the K+ loss and generates an osmolality balance between the intracellular space and the medium. The yeast HKT ortholog, TRK1 of Saccharomyces cerevisiae, mediates K+ uptake (Ko and Gaber, 1991) and also contributes to modulating the ionic balance between the cytosol and the outside space and setting the membrane potential (Madrid et al., 1998). Na+ is well known as a cytotoxic cation in plants, but Na+ can be used as an osmolyte for vacuolar osmotic adjustment (Blumwald, 2000; Horie and Schroeder, 2004; Zhu, 2003). Tonoplast Na+ transporters compartmentalize Na+ from the cytosolic space (Apse and Blumwald, 2002; Apse et al., 1999) (Figure 8). An increase in the Na+ content in vacuoles may cause increases in the volume of the vacuoles, and accordingly a decrease in the volume of the cytosolic space, which leads to an increase in the cytosolic osmolality of cells. We predict a possible additional role for AtHKT1 in response to increases in osmolality in the xylem: AtHKT1 may counteract high osmolality stress by causing retrieval of Na+ from the xylem and by increasing the osmolality of the xylem parenchyma cells when high levels of Na+ are present in the xylem.

Model for AtHKT1 regulation of xylem and shoot K+ content

As a consequence of the absence of AtHKT1 in athkt1 plants, the amount of K+ in the xylem sap and in shoots showed a slight decrease under salt stress (Figures 5a,b and 6a,b), which is opposite to the effect on Na+ content found in athkt1 mutants (Figure 4a,b) (Berthomieu et al., 2003; Gong et al., 2004; Mäser et al., 2002b). Lacan and Durand (Lacan and Durand, 1996) have proposed that in the presence of Na+, the transport system, which resides in the xylem parenchyma, absorbs Na+ from the xylem sap and releases K+. Even relatively moderate salinity stress (up to 37.5 mm NaCl) allowed removal of Na+ from the xylem stream and release of K+ into the xylem (Lacan and Durand, 1996). Consistent with this hypothesis, we have found that levels of K+ in the xylem sap and in leaves of athkt1 mutants were decreased (Figures 5a,b and 6a,b).

Interestingly, the root/shoot K+ content of athkt1 showed an opposite characteristic to that of Na+ under salt stress (Mäser et al., 2002b; Rus et al., 2004), showing that athkt1 mutants maintain lower K+ levels in shoots (Figure 6a,b) but higher K+ in roots (Figure 6c,d). In roots, efflux of K+ from xylem parenchyma cells into the xylem sap has been proposed to be carried out by outwardly rectifying K+ channels called KORC (De Boer and Volkov, 2003; Gaymard et al., 1998; Roberts and Tester, 1995; Wegner and De Boer, 1997; Wegner and Raschke, 1994). Indeed uptake of Na+ via AtHKT1 would depolarize xylem parenchyma cells, which in turn could activate K+ efflux channels, resulting in release of K+ into the xylem sap. This model is consistent with our findings that athkt1 loss-of-function mutations slightly reduce the K+ content of xylem sap and of shoots (Figures 5a,b and 6a,b). Furthermore, the reduced K+ loading of xylem vessels in athkt1 mutants (Figure 5a,b) may also cause the higher retention of K+ in roots found in athkt1 mutants (Figure 6c,d).

SOS1 mRNA is expressed in epidermal cells in roots and in xylem parenchyma cells in roots and in shoots (Shi et al., 2002). The sos1 mutants show Na+-sensitive inhibition of root elongation and Na+ over-accumulation (Wu et al., 1996). SOS1 has been proposed to reduce the net Na+ flux into cells in the root tip (Shi et al., 2003). Under conditions of high Na+, athkt1 mutants showed chlorosis and inhibition of growth of aerial tissues (Berthomieu et al., 2003; Gong et al., 2004; Mäser et al., 2002b). The sos1 mutant exhibited a higher Na+ content in xylem sap compared with the wild type (Shi et al., 2002), which is similar to the phenotype of athkt1 mutants. Nevertheless, mutation of AtHKT1 rescued the salt-sensitive phenotype of sos1 plants (Rus et al., 2004). The present findings and derived model (Figure 8), and other recent findings (Berthomieu et al., 2003; Gong et al., 2004; Mäser et al., 2002b), indicate that AtHKT1 is involved in Na+ homeostasis in roots. Further analyses of the physiological mechanisms mediating salinity resistance in sos athkt1 double-mutant plants will be required and may lead to identification of further complex interactions.

In summary, the present study demonstrates plasma membrane targeting and expression in xylem parenchyma cells of the AtHKT1 protein. Together with the observed dramatic increase in Na+ content in the xylem sap of three athkt1 loss-of-function mutants, we propose a major role for AtHKT1 in removing excess sodium from the xylem during osmotic and salinity stress, thus protecting leaves from sodium toxicity.

Experimental procedures

Antibody production

A peptide corresponding to the sequence KITKPRTTSRPHDFD (amino acid positions 41–55 of AtHKT1) was synthesized and was used to generate antiserum containing the anti-AtHKT1 antibodies (Sawadei Corp., Tokyo, Japan). The DNA encoding the peptide sequence was inserted into the BamHI–EcoRI site in the vector pGEX-2T (Amersham Biosciences, Little Chalfont, UK). From E. coli containing the resultant plasmid, the GST fusion peptide was purified using a glutathione SepharoseTM 4B column (Amersham Biosciences). For production of the GST fusion peptide in Sf9 cells, the DNA fragments encoding the same fusion protein were inserted into the NcoI–XhoI site in the pFASTBac HTa plasmid (Invitrogen, Carlsbad, CA, USA). The gene was transfected into Sf9 cells using the Baculovirus system (Invitrogen). The recombinant proteins were electrophoresed on SDS/12.5% PAGE gel, stained with Coomassie Brilliant Blue or electroblotted onto polyvinylidene difluoride (PVDF) microporous membrane (Millipore, Billerica, MA, USA). The horseradish peroxidase (HRP)-conjugated goat antirabbit immunoglobulin G was used as a secondary antibody. The protein was detected with an ECLTM Western blotting detection reagents system (Amersham Biosciences).

Immunoelectron microscopy

The specimens for immunoelectron microscopy (IEM) were immersed in a mixture of 0.5% glutaraldehyde and 4% paraformaldehyde in 50 mm sodium phosphate buffer (pH 7.2) and were kept at 4°C for 1.5 h. After washing three times with PBS, the specimens were dehydrated through an ethanol series (30–99.5% v/v), and embedded in LR white resin (London Resin Company Ltd, Theale, UK) followed by polymerization for 24 h at 50°C. Immunostaining with rabbit anti-AtHKT1 antiserum (rabbit IgG; diluted 1:20 000) was performed on ultrathin sections picked up on nickel grids after blocking with normal goat IgG (chromatographically purified; Zymed Laboratories, Inc., San Francisco, CA, USA) diluted 1:30 in 50 mm Tris-buffered saline containing 0.1% BSA for 15 min as described in Kamasawa et al. (1992). An ultra-small colloidal gold-conjugated goat antirabbit IgG (diluted 1:100, Aurion, Wageningen, The Netherlands) was used as a secondary antibody. After immunostaining, the sections were fixed with 1% glutaraldehyde (GA) in 0.1 m phosphate buffer, and the colloidal gold particles were enhanced by silver deposition using high-efficiency silver enhancement reagents (R-GENT SE-EM, Aurion) as described (Humbel et al., 2001). The grids were stained in 6% uranyl acetate and viewed by transmission electron microscopy (H-800; Hitachi, Tokyo, Japan) at 125 kV.

Subcellular localization of GFP–AtHKT1

AtHKT1 was fused at its N-terminus with GFP and Tobacco Bright Yellow 2 (BY2) cells were transformed via Agrobacterium (strain C58). Fluorescence of GFP was observed by confocal microscopy (Eclipse TE2000-U; Nikon, Tokyo, Japan) using a 488 nm excitation wavelength and a 510 nm emission wavelength. The pJFH vector, which includes a 35S CaMV promoter and a green fluorescence protein (GFP) gene, was kindly provided by Dr Jeffrey F. Harper (University of Nevada).

Salt sensitivity analyses

Seeds of wild-type (Wassilewskija: Ws), athkt1-3 and three independent transgenic athkt1-3 plants expressing AtHKT1 cDNA under the control of a native 0.8 kb promoter (Mäser et al., 2002b) were germinated on MS medium. Two of the three transgenic lines were heterozygous and one was homozygous. For heterozygous lines MS plates containing 25 mg l−1 hygromycin B were used to select transgenic plants. Four healthy seedlings, which were 6–7 days old, were then transferred onto fresh MS medium using a nylon mesh with 10 μm pores. Five days later meshes were transferred onto minimal medium (Mäser et al., 2002b) containing 75 mm NaCl and incubated for another 16 days.

AtHKT1 promoter–GUS expression

Seeds of Arabidopsis thaliana expressing the AtHKT1 promoter–β-glucuronidase (GUS) gene were germinated in half-strength MS media containing 0.8% agar and 2% sucrose at 20°C on a day/night cycle of 16 h light/8 h dark (Mäser et al., 2002b). At day 15, the seedlings were transferred into fresh media containing 2 mm MgSO4, 1 mm CaCl2, 5 mm Ca(NO3)2, 1 mm H3PO4, 0.1 mm FeEDTA, 5 mmβ-morpholinoethanesulfonic acid (MES), 7 μm H3BO3, 1.4 μm MnSO4, 1 μm ZnSO4, 4.5 μm KI, 0.1 μm CuSO4, 0.2 μm Na2MoO4, 10 nm CoCl2, 0.8% agar and 2% sucrose. As indicated in treatments for Na+ or K+, the basic medium was supplemented with 0, 15, 30, 45, 60 or 75 mm NaCl or KCl. In addition, for sorbitol and mannitol treatments, the basic medium was supplemented with 0, 50, 100, 150 or 200 mm sorbitol or mannitol. The ProAtHKT1:GUS plants were analyzed for GUS expression according to standard procedures (Jefferson et al., 1987). The whole plant for each treatment was incubated at 37°C for 12 h in the staining solution containing 0.5 mm X-Gluc (5-bromo-4-chloro-3-indolyl-β-d-glucuronide).

Reverse transcriptase-PCR

Total RNA was isolated according to the same procedure described previously (Mäser et al., 2002a; Uozumi et al., 2000). Reverse transcription was conducted according to the Superscript III (Invitrogen). A primer set used for the β-tubulin 2 gene was 5′-CTCAAGAGGTTCTCAGCAGTA-3′; 5′-TCACCTTCTTCATCCGCAGTT-3′ and that for AtHKT1 was 5′-TGACGTTGAGACTGTTACTG-3′; 5′-CTTTCGGTGATTGAAATGAG-3′. The PCR was performed using ExTaq polymerase (Takara, Ootsu, Japan) under the following conditions: 94°C for 60 sec; 24 cycles of 94°C for 30 sec, 55°C for 45 sec, 72°C for 90 sec.

Determination of GUS activities

One hundred milligrams of roots or shoots from plants grown in the indicated treated media for 14 days was ground in liquid nitrogen and extracted with 1 ml lysis buffer containing 50 mm phosphate buffer (pH 7), 10 mm EDTA, 0.1% Triton X-100, 0.1% sodium lauryl sarcosine and 10 mmβ-mercaptoethanol. The extracts were centrifuged at 11 000 g at 4°C for 10 min. Then, the GUS activities in the supernatant of root or shoot were determined according to Jefferson et al. (1987) using 4-methylumbelliferyl-β-d-glucuronide (MUG) as a substrate. Histochemical assays of GUS activity were conducted using the samples that were already stained according to Jefferson et al. (1987).

Collection of xylem and phloem sap

Plants were grown on soil (McConkey Co., Sumner, WA, USA) for 3–4 weeks until they were at the bolting stage. Plants were then subjected to 75 mm NaCl by a single irrigation with one-twentieth MS salts for 1 or 2 days. For collection of xylem sap all rosette leaves were removed with scissors, and the inflorescence stem was cut with a very sharp razor blade (Gaymard et al., 1998; Shi et al., 2002). The tray and plants were then covered with a transparent plastic dome to maintain high humidity. Drops of xylem sap that accumulated at the cutting surface of the inflorescence stem were then collected using a micropipette. Collected sap samples were later diluted directly in 5% HNO3 solution. Sodium and potassium ion content was measured by inductively coupled plasma-optic emission spectroscopy (ICP-OES, Perkin Elmer Optima 3000XL, Applied Biosystems, Foster City, CA, USA) (Mäser et al., 2002b). For collection of phloem sap (Berthomieu et al., 2003; Corbesier et al., 2003), four to five mature rosette leaves were detached at their petiole bases. The petioles were recut under 20 mm EDTA-(NH4)2 (pH 7.5). Four leaves collected from one plant were placed in a 1.5 ml microcentrifuge tube with their petioles immersed in 1.25 ml of 15 mm EDTA-(NH4)2 (pH 7.5). Then the tubes were placed for 4 h in an illuminated growth room in airtight transparent plastic containers, in which the atmosphere was water saturated (to prevent uptake of EDTA solution by the leaves), to dissolve the phloem sap in EDTA solution (Berthomieu et al., 2003; Corbesier et al., 2003). Then the EDTA solution was diluted with equal volumes of 10% HNO3 solution and used for ICP measurements.


We thank Naomi Kamasawa, Mamiko Sato and Emiko Kobayashi (Japan Women's University) for expert assistance, and Alice Chen (UC San Diego) and Hiroshi Miyake (Nagoya University) for helpful discussions. This work was supported by a grant-in-aid for COE Research (to N.U.), Department of Energy grant DOE-DE-FG02-03ER15449 (to J.I.S), the 21st Century COE Program (to N.U.), and in part by NIEHS grant 1P42 ES010337 (to J.I.S.), grants-in-aid for scientific research (1604660, 17078005, and 17380064 to N.U.) from the MEXT and the JSPS (to N.U.). Immunoelectron microscopical experiments were supported by the Open Research Center of JWU established in private universities in Japan with support of the Ministry of Education, Culture, Sports, Science and Technology.