Expression analysis suggests novel roles for members of the Pht1 family of phosphate transporters in Arabidopsis


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The completion of the Arabidopsis thaliana genome has revealed that there are nine members of the Pht1 family of phosphate transporters in this species. As a step towards identifying the role of this gene family in phosphorus nutrition, we have isolated the promoter regions from each of these genes, and fused them to the reporter genes β-glucuronidase and/or green fluorescent protein. These chimeric genes have been introduced into A. thaliana, and reporter gene expression has been assayed in plants grown in soil containing high and low concentrations of inorganic phosphate (Pi). Four of these promoters were found to direct reporter gene expression in the root epidermis, and were induced under conditions of phosphate deprivation in a manner similar to previously characterised Pht1 genes. Other members of this family, however, showed expression in a range of shoot tissues and in pollen grains, which was confirmed by RT-PCR. We also provide evidence that the root epidermally expressed genes are expressed most strongly in trichoblasts, the primary sites for uptake of Pi. These results suggest that this gene family plays a wider role in phosphate uptake and remobilisation throughout the plant than was previously believed.


Phosphorus (P) is an essential mineral element for plants as it is involved in many key biochemical reactions and is a component of nucleic acids, phospholipids, and energy-providing ATP. P is acquired by plants in the form of inorganic phosphate (Pi), which is taken up from the soil solution by the roots. However, the concentration of Pi in the soil solution is typically less than 2 µm, while that in the cytoplasm of plant cells is generally greater than 10 mm (Mimura, 1999). This uptake therefore occurs against a large concentration gradient, and thus is an active process.

In the last few years, genes encoding proton-coupled Pi transporters (H+/H2PO4 symporters) have been isolated from a number of plant species, including Arabidopsis thaliana (Daram et al., 1999; Muchhal et al., 1996; Okumura et al., 1998; Smith et al., 1997), Solanum tuberosum (Leggewie et al., 1997; Rausch et al., 2001), Catharanthus roseus (Kai et al., 1997), Lycopersicon esculentum (Daram et al., 1998; Liu et al., 1998a), Medicago truncatula (Liu et al., 1998a), and Hordeum vulgare (Smith et al., 1999). These transporters have been classified into two families, namely Pht1 and Pht2 (Bucher et al., 2001). Based on the kinetic data that has been obtained for some of its members, the Pht1 family has been believed to contain the high affinity transporters. This family includes 17 out of the 18 plant phosphate transporter genes published to date. The remaining gene, ARAth;Pht2;1, shows low affinity kinetics, and is the first member of the Pht2 family to be isolated (Daram et al., 1999).

Amongst the 17 members of the Pht1 family published to date, Northern analysis has indicated that, with the exception of three genes from A. thaliana (for which no expression was detected), all members of this gene family are expressed in roots, and 11 are expressed solely in roots. In addition, the expression of at least 12 of these genes is induced in roots in response to Pi deprivation (Daram et al., 1998; Kai et al., 1997; Leggewie et al., 1997; Liu et al., 1998a, 1998b; Muchhal et al., 1996; Okumura et al., 1998; Smith et al., 1997, 1999). In contrast, ARAth;Pht2;1 is expressed mainly in the leaves, and appears to be little affected by Pi availability (Daram et al., 1999).

While the majority of Pht1 genes are known to be expressed in roots, the cell specificity of the expression has only been determined for 3 genes, LePT1, MtPT1 and StPT3. In situ hybridisation studies have shown that the tomato phosphate transporter gene LePT1 is expressed most strongly in the epidermis near the root tip and in the root cap, and also in the young stele of Pi starved roots (Daram et al., 1998; Liu et al., 1998a). In mature roots of mycorrhizal plants, this transporter (or one with high homology to it) was expressed in cortical cells containing arbuscules, suggesting a role for this transporter in Pi uptake from the fungus (Rosewarne et al., 1999). The StPT3 transporter from potato has recently been shown to be localised at the periarbuscular membrane of roots infected with arbuscular mycorrhizal fungi (Rausch et al., 2001), where it presumably plays a role in transport of Pi from the fungi to the host plant. The MtPT1 gene and protein have been localised in the epidermis and root hairs, using in situ hybridisation and immunolocalisation (Chiou et al., 2001).

These studies, along with the kinetic data that has been obtained for some of the members of the Pht1 gene family, have suggested that these transporters are involved in the uptake of phosphate from the soil solution. However, most of the Pht1 genes that have been isolated to date have been obtained from root cDNA libraries, which has introduced a bias towards root-expressed genes. The recent completion of the Arabidopsis thaliana genome has now allowed the analysis of the complete family of Pht1 transporters in this species. Towards this end, we have constructed promoter-reporter gene fusions for all nine Pht1 genes from A. thaliana, and have determined the expression patterns for each gene in transgenic plants. This has revealed that the Pht1 gene family plays a broader role in phosphorus nutrition than was previously believed.


Of the six A. thaliana Pht1 transporters described previously, only Pht1;1 and Pht1;4 have been demonstrated to function as phosphate transporters (Mitsukawa et al., 1997; Muchhal et al., 1996). BLAST searches using the coding sequence of these genes against the A. thaliana genome revealed the presence of nine members of the Pht1 gene family in this species. Our designated naming system for these genes (following the guidelines of the Commission for Plant Gene Nomenclature), along with their former name(s) and protein entry codes for MATDB (, are given in Table 1. An alignment of the amino acid sequences of the nine phosphate transporters encoded by these genes is shown in Figure 1.

Table 1.  Members of the Pht1 family of phosphate transporter genes from A. thaliana, and promoter lengths used in this study
Gene nameProtein entry
Former name(s)Size of promoter
relative to ATG
ARAth; Pht1;1At5g43350PHT1b, APT2c, AtPT1d− 2766 to − 1
ARAth; Pht1;2At5g43370PHT2b, APT1c− 2000 to − 3
ARAth; Pht1;3At5g43360PHT3b, AtPT4e− 1672 to − 23
ARAth; Pht1;4At2g38940PHT4b, AtPT2d− 2987 to + 38
ARAth; Pht1;5At2g32830PHT5b− 2713 to + 31
ARAth; Pht1;6At5g43340PHT6b− 2005 to + 24
ARAth; Pht1;7At3g54700− 2521 to − 2
ARAth; Pht1;8At1g20860− 2484 to + 39
ARAth; Pht1;9At1g76430− 1845 to − 36
Figure 1.

Alignment of the amino acid sequences of the 9 Pht1 phosphate transporters from Arabidopsis.

In order to determine the cell specificity and phosphate responsiveness of these nine genes, we amplified the promoter of each gene and fused it to the coding regions of the GFP and/or GUS reporter genes. These promoter-reporter gene fusions were introduced into A. thaliana plants, and progeny of the transgenic lines were assayed for reporter gene activity when grown in tissue culture and in soil containing high and low phosphate levels. At least 10 independent transgenic lines were analysed to determine cell specificity, and 2–3 representative lines were used to determine the response to Pi deprivation. We adopted this reporter gene approach, as the high homology amongst some members of the Pht1 gene family (Figure 1) may restrict the use of in situ hybridisation and immunolocalisation techniques. The increased root hair length observed on plants grown in the low Pi soil (Figure 2d,e) is consistent with the observations of Bates and Lynch (1996).

Figure 2.

Localisation of promoter-GFP expression in transgenic A. thaliana plants.

(a- e) Expression of Pht1;1-GFP in the hydathode of a cotyledon (a), axillary buds (b), peripheral endosperm of a germinating seed (c), and in roots from high Pi (d)and low Pi soil (e).

(f) Confocal microscope image showing expression of Pht1;1-GFP predominantly in the epidermis.

(g- h) Expression of Pht1;2-GFP in roots from high Pi (g) and low Pi soil (h).

(i- j) Confocal microscope images of Pht1;2-GFP expression in epidermis of young root (i) and cortex of old root (j).

(k- l) Expression of Pht1;2-GFP in the young lateral roots (yr), but not in the older roots (or) of a 5-week-old root system (l), and corresponding brightfield image (k).

(m- n) Expression of Pht1;3-GFP in primary and lateral roots (m), and expression in trichoblasts further along lateral root (n).

(o) Expression of Pht1; 3-GFP in hydathode of cotyledon.

(p)Confocal microscope image showing pericycle-specific expression of Pht1;3-GFP in the primary root and lateral root near junction with primary root. For the confocal microscopy images, the roots were stained with propidium iodide, allowing the cell walls of epidermal and cortical cells to be visualised by red fluorescence. Scale bars are 200 µm in (a) (k) (l) and (m); 100 µm in (b-e) (g-h) and (n); 20 µm in (f) (i-j) and (p); and 0.5 mm in (o). White dashes in (n) indicate edge of lateral root.

Transgenic plants containing the Pht1;1 promoter fused to either GFP or GUS showed reporter gene activity in roots, hydathodes of cotyledons and leaves, axillary buds and in the peripheral endosperm of germinating seeds (Figure 2a-e). No expression was observed in other tissues of the shoot. When grown in the low phosphate soil, there was an increase in expression of the reporter gene in the root hair zone of roots, and gene activity was induced in the root cap (Figure 2d,e). No increase in expression levels in hydathodes and axillary buds was observed in plants grown in low Pi soil. Confocal microscopy indicated that the expression in roots was strongest in the epidermis, with weak expression also occurring in cortical cells of the root hair zone (Figure 2f). Within the epidermis, strongest expression was observed in the trichoblast cells (Figure 3a). In Arabidopsis, trichoblasts occur in the epidermal files overlying the anticlinal cortical cell walls, and are separated by files of atrichoblasts (Dolan et al., 1993). Strong expression was also found throughout the columella and lateral root cap of roots from low P soil using confocal microscopy analysis (data not shown). In older, flowering plants, reporter gene expression was only observed in young lateral roots (data not shown).

Figure 3.

Localisation of promoter-reporter gene fusions in transgenic A. thaliana plants.

(a- e) Transverse sections derived from confocal microscope z-series through root of Pht1;1-GFP transgenic (a), root of Pht1;2-GFP transgenic (b), primary root of Pht1;3-GFP transgenic (c), lateral root of Pht1;3-GFP transgenic near junction with primary root (d), and further along this lateral root (e), all from plants grown in low Pi soil. Red fluorescence indicates cell walls stained with propidium iodide, while green indicates GFP fluorescence.

(f) Leaf from a Pht1;3-GUS transgenic plant, showing vascular-specific GUS activity.

(g- j) Localisation of GUS activity in Pht1;4-GUS transgenic plants, showing weak promoter activity in roots from high Pi soil (g), strong activity in roots from low Pi soil (h), activity in hydathodes located at leaf teeth (i), and strong activity in axillary buds (j).

(k) Transverse section of a wax-embedded Pht1;4-GUS root from low Pi soil, showing GUS activity in the epidermis (epi) and root hairs (rh), but not in cortex (cor), endodermis (end), phloem (ph) or xylem (x). Some wax crystals are also visible outside the epidermis.

(l) Pht1;4-GUS activity in stele, and in epidermis of lateral root.

(m) Pht1;4-GUS flowers, showing GUS activity in senescing anthers and abscission zones at the base of these anthers. Scale bars are 20 µm in (a-e) and (k); 100 µm in (g-h); 1 mm in (f, i and j); 50 µm in (l); and 0.5 mm in (m).

The Pht1;2 promoter directed expression of GFP and GUS only in root tissues, and showed weaker expression levels than the Pht1;1 promoter in plants grown in the presence of high phosphate levels. However, when grown in soil with low phosphate levels for 3 weeks, strong reporter gene expression was observed in epidermal cells and root hairs throughout the root hair zone (Figure 2g,h). In contrast to the Pht1;1 promoter, no expression was observed in the root tips. When observed using confocal microscopy, the expression was found to be mostly in the epidermis (Figure 2i), and was much stronger in the trichoblast cells than the atrichoblast cells (Figure 3b). In older regions of 3-week-old primary roots (near the hypocotyl junction), the strongest GFP expression was observed in cortical cells (Figure 2j). As the root systems aged further, reporter gene expression was only observed in young lateral roots (Figure 2k,l).

Reporter genes under the control of the Pht1;3 promoter were also expressed mainly in root tissues, and again were induced under Pi deprivation conditions. However, the cell specificity of this promoter differed from those described above. In 3-week-old seedlings, reporter gene expression was observed predominantly in the stele of the primary root (Figure 2m). Confocal microscopy of Pht1;3-GFP transgenic roots indicated that, in primary roots and in lateral roots near the junctions with primary roots, this expression is restricted to the pericycle cell layer (Figures 2p and 3c,d). In the lateral roots and near the tip of the primary root, however, the GFP expression was found to be strongest in the root hair-producing trichoblast cells, and the expression in the stele was much lower than in the primary root (Figures 2n and 3e). A small number of transgenic lines also showed weak reporter gene activity in vascular tissue of young leaves and in hydathodes (Figures 2o and 3f). As with the above promoters, similar expression patterns were observed with both the GFP and GUS reporter genes.

Transgenic plants containing the Pht1;4-GUS fusion showed GUS activity in roots, hydathodes and axillary buds (Figure 3g-j). In longer (overnight) assays, GUS activity extended throughout the cotyledons (data not shown). The activity of this promoter was also found to be induced in roots of plants grown in low Pi soil (Figure 3g,h), and reporter gene activity was observed in the epidermis (Figure 3k), root tips and in some cases in the stele, where it occurred in cell layers interior to the pericycle (Figure 3l). The Pht1;4-GUS transgenic plants also showed GUS activity in senescing anther filaments and in the abscission zone at the base of siliques (Figure 3m).

Quantitative GUS assays done on progeny of representative transgenic lines containing the Pht1;1, Pht1;2, Pht1;3 and Pht1;4 promoters linked to the GUS reporter gene confirmed the inducibility of these four promoters when grown in soil containing low levels of Pi (Figure 4). For each promoter, a significant increase in expression was observed in roots from low Pi soil, reflecting the results obtained with histochemical GUS assays, and with the GFP transgenics.

Figure 4.

GUS activity in root systems grown in high or low Pi soil.

Results show mean ± standard error for three replicates, each replicate consisting of root systems harvested from approximately 3–5 seedlings grown in one pot. Black bars show GUS activity in high Pi soil, while hatched bars show activity from low Pi soil.

In contrast to the above promoters, the Pht1;5 promoter directed GUS expression mainly in shoot tissues. In very young seedlings (3 days after germination), strong GUS activity was detected throughout the cotyledons and hypocotyl (Figure 5a). After a further 2 weeks growth, this activity was restricted to the vascular tissue of the cotyledons (Figure 5b,c). No GUS activity was observed in leaves until they started to senesce, at which point GUS activity was observed in the vascular tissue (Figure 5d). Transverse sections indicated that this GUS activity was strongest in the phloem (Figure 5e). GUS activity was also observed throughout young floral buds, but became restricted to the sepals later in floral development, where it was again strongest in the vascular bundles (Figure 5f). In plants grown under Pi deprivation, weak GUS activity was occasionally seen in the stele of some roots (data not shown).

Figure 5.

Localisation of promoter-reporter gene fusions in transgenic A. thaliana plants.

(a- f) Localisation of Pht1;5-GUS expression in 4-day-old seedling (a), 2- to 3-week-old seedlings (b- c), and in senescing leaves (d) and flower buds (f) of older plants. (e)Transverse section through senescing leaf from (d), showing GUS activity localised in the phloem.

(g-k) Expression of Pht1;6-GFP in tapetum of anthers (h) and corresponding brightfield image (g), in mature dry pollen grains (i), in pollen grains that have been allowed to germinate for 6 h (j), and in cotyledon hydathode (k). Scale bars are 1 mm in (a) and (c-d); 0.5 mm in (b) (f) and (k); 30 µm in (e); 100 µm in (g-h); and 50 µm in (i-j).

The Pht1;6 promoter directed GFP and GUS expression in anthers of transgenic plants (Figure 5g,h). Strong expression was observed in the tapetum of flowers from about Stage 9 to Stage 11 as defined by Smyth et al. (1990). Reporter gene expression was also observed in dry, mature pollen grains, and no change in expression level was detected during the first 6 h of pollen germination (Figure 5i,j). The pollen shown in these images is from a hemizygous plant, and is therefore segregating for the reporter gene construct. Approximately one third of the transgenic lines also showed weak expression in the hydathodes of cotyledons (Figure 5k), which in some cases extended throughout the vascular tissue of the cotyledons once the plants began flowering. The Pht1;7 promoter also directed strong GFP expression in mature pollen grains (data not shown).

We did not observe any reporter gene activity in any tissues with the Pht1;8 or Pht1;9 promoters under the conditions in which plants were grown in this study.

We have also analysed the expression patterns of each of the Pht1 genes using RT-PCR with gene-specific primers, except for Pht1;1 and Pht1;2, which were analysed together (with a single pair of primers), as gene-specific RT-PCR and Northern analysis has previously been published for these 2 genes (Smith et al., 1997). In general, the results obtained with RT-PCR confirm the organ specificity observed with the promoter fusions, with cDNA being amplified from tissues in which reporter gene expression was observed (Figure 6). The RT-PCR analysis did, however, provide evidence that the Pht1;7, Pht1;8 and Pht1;9 genes are expressed to some degree in Pi-deprived roots, despite the fact that no reporter gene expression was observed with the promoter fusions.

Figure 6.

RT–PCR analysis of the Arabidopsis Pht1 family. RT-PCR products were amplified from a range of Arabidopsis tissues using primers specific to each of the Pht1 genes except for Pht1;1 and Pht1;2, which were both amplified using a single set of primers.

The products of these reactions were visualized on an ethidium bromide-stained 1.2% agarose gel. The ‘no DNase treatment’ (top) revealed the presence of both cDNA (357 bp) and genomic DNA (508 bp) products prior to treatment of the RNA with DNase1, but this contaminating genomic DNA was effectively removed following DNase treatment (second from top), indicating that the amplified products represent cDNA. The diamond symbols indicate the 500 bp band of the 1 kb PLUS DNA ladder (Roche). Expected sizes of products from cDNA and genomic DNA are shown in Table 2.


The completion of the A. thaliana genome has revealed that the Pht1 family of phosphate transporter genes contains a total of nine closely related members. To study the expression of these genes, we have constructed promoter-reporter gene fusions, which have been assayed in transgenic A. thaliana plants. The results of this study provide some insight into the roles of the individual Pht1 phosphate transporters during plant development.

During seed germination, the mobilisation of lipid and protein reserves occurs first in the peripheral endosperm (Mansfield and Briarty, 1996). The phosphate required for this metabolic activity needs to be acquired through transporters in the plasma membranes of these cells, as there are no plasmodesmatal connections between the endosperm and maternal tissue. The observed activity of the Pht1;1 promoter in this tissue suggests that this transporter may perform this role.

The major pool of phosphate in dicot seeds is phytate (inositol hexaphosphate), which is stored in the embryo and endosperm during seed maturation (Lott et al., 1995). As the seed germinates, phytases break down this phytate, releasing Pi, which is then remobilised to growing parts of the plant for the synthesis of membrane lipids and nucleic acids (Lott et al., 1995). Our results suggest that the Pht1;5 transporter may be involved in this remobilisation, as the Pht1;5 promoter is expressed strongly throughout the cotyledons and hypocotyl in the first few days after germination when Pi is being released from phytate. The promoter activity subsequently drops in all tissues except those associated with the cotyledonary vascular bundles, through which the Pi that has been taken up by the surrounding tissues is moved to sink tissues elsewhere in the plant. In the A. thaliana seedlings, the vascular traces of at least the first 5 rosette leaves are connected to a cotyledonary vascular trace (Busse and Evert, 1999), and are therefore likely to act as sinks during development.

The promoter-reporter gene fusions in this study provide evidence that 4 members of the Pht1 family of phosphate transporters are involved in Pi uptake from the soil solution, as reporter genes driven by the promoters of Pht1;1, Pht1;2, Pht1;3 and Pht1;4 are all expressed in the root epidermis, and are induced when grown in soil containing low Pi concentrations. These genes are therefore playing a similar role to the previously characterised LePT1, LePT2 and MtPT1 genes (Chiou et al., 2001; Daram et al., 1998; Liu et al., 1998a). The Pht1;1 transporter of A. thaliana has been shown to function as a high affinity phosphate transporter when expressed in tobacco cells (Mitsukawa et al., 1997). The Pht1;2 protein is almost identical to Pht1;1 (Figure 1), and is therefore also likely to be a high affinity transporter. Such high affinity transporters are needed at the root-soil interface, due to the very low concentration of Pi in the soil solution. The affinity of the remaining Pht1 transporters from A. thaliana remains to be determined, but the presence of some of them in vascular tissues suggests that this family may also include lower affinity transporters. The presence of multiple transporters functioning at the root-soil interface reflects the importance of the root surface for Pi uptake by the plant.

Root hairs are the major site of Pi uptake from the soil solution (Gahoonia and Nielsen, 1998), and the confocal microscopy studies described here show that genes encoding the high affinity Pht1;1 and Pht1;2 transporters, as well as the Pht1;3 transporter (in lateral roots) are expressed preferentially in the root hair-producing trichoblast cells of the epidermis (Figure 3a,b,e). This pattern of gene expression has not been described for any other nutrient transporters, and reflects the importance of the root hairs for nutrient uptake, particularly phosphate. Furthermore, Arabidopsis is known to produce longer and more dense root hairs under low Pi conditions (Ma et al., 2001). This phenotype was also observed in the soil-grown plants in this study (e.g. Figure 2d,e). It is worth noting that the roots assayed in the present study were grown in soil, rather than in agar, hydroponic or aeroponic systems, which have been used for many of the other studies of members of the Pht1 gene family, and therefore the results obtained here may more accurately reflect the situation found in the field.

The observed weak activity of the Pht1;1 and Pht1;2 promoters in cortical cells of young roots would allow the uptake of Pi that has entered the root apoplastically. In older roots, we observed a change in the expression pattern of the Pht1;2 promoter, with the strongest expression being in cortical cells. This may reflect early stages of secondary thickening in such roots, during which the epidermis becomes non-functional. In even older root systems, we observed Pht1;1 and Pht1;2 promoter activity only in the young lateral roots, which had not undergone secondary thickening, and therefore still had functional epidermis layers (e.g. Figure 2k,l).

Once inside the root epidermal cells, radial movement of Pi occurs symplastically through the endodermis and into the stele, where it is loaded into the xylem in a process that appears to involve the product of the pho1 gene (Poirier et al., 1991). The activity of the Pht1;3 promoter in the pericycle of the primary root suggests that this transporter may also play a scavenging role, as suggested for the Sultr2;1 sulphate transporter of A. thaliana (Takahashi et al., 2000). The Pi flux in primary roots will increase as more lateral roots join the primary root, and therefore the leakage of Pi from the xylem vessels into the apoplastic spaces within the stele will be greater in primary roots. In such roots, the expression pattern of the Pht1;3 promoter switches from being epidermal, to being expressed mainly in the pericycle, where the Pht1;3 transporter may re-absorb the leaked Pi. In addition, the pericycle cells have the capacity to form new meristems (as occurs during lateral root initiation), and may therefore have greater requirements for Pi than the surrounding cells.

Several of these Pi transporter promoters were also found to direct reporter gene activity in the hydathodes of cotyledons and leaves. Similarly, an A. thaliana high affinity sulphate transporter (Takahashi et al., 2000) and potassium transporter (Lagarde et al., 1996) are expressed in hydathodes, and there is evidence for enhanced H+-ATPase activity in cells of the hydathode (Wilson et al., 1991). Hydathodes are the sites at which guttation water is lost from the leaves or cotyledons (Wilson et al., 1991). Nutrient transporters expressed in this tissue probably play a role in preventing nutrients from being lost from the leaf during guttation. We do not believe that the observed reporter gene activity in hydathodes has resulted from accumulation of the reporter gene product in this tissue, as the Pht1:5 promoter did not direct reporter gene activity in hydathodes.

A functional explanation for the observed activity of the Pht1;1 and Pht1;4 promoters in axillary buds is unclear. It may be that there is an unusually high demand for Pi, due to the high potential metabolic activity of this tissue. Interestingly, the A. thaliana Sultr1;1 sulphate transporter is also expressed in axillary buds (Takahashi et al., 2000).

The switch from vegetative to floral growth creates a new Pi sink, and at about this stage of development the Pht1;5 promoter becomes active in the phloem tissue of the oldest leaves, which are starting to senesce. This suggests that, in addition to its putative role in remobilisation of Pi from germinating seed, the Pht1;5 transporter may be involved in the redistribution of Pi out of old leaves (where it is no longer required), and up to the sink in the floral tissues. It has recently been shown that almost 80% of P is remobilised out of senescing Arabidopsis leaves (Himelblau and Amasino, 2001), and there is evidence for apoplastic phloem loading in source leaves of Arabidopsis (Gottwald et al. 2000), which explains the need for a transporter in this process. At least one senescence-associated RNase gene is known in Arabidopsis (Taylor et al., 1993), and this activity would generate additional Pi for remobilisation out of these old leaves. Similarly, the observed activity of the Pht1;5 promoter in sepals of opening flower buds may reflect a role for this transporter in remobilisation out of these organs, which are no longer metabolically active. Earlier in floral development, this promoter is expressed more uniformly throughout the floral buds, and so at this stage Pht1;5 may play a role in supplying Pi to these actively growing tissues.

Transport of Pi across the plasma membrane is also required within some tissues of the anther, which lack plasmodesmatal connections to adjacent cells. At the ‘post meiosis, precytokinesis’ stage of pollen development such connections between the tapetum and the middle layer are lost (Owen and Makaroff, 1995). Pi required for the active production of pollenkitt and tryphine by tapetal cells is therefore likely to be obtained via phosphate transporters expressed in the tapetum. Similarly, plasmodesmatal connections between the developing pollen and the tapetum are lost early in development (Owen and Makaroff, 1995), necessitating the action of transporters for Pi uptake into pollen grains. Previous studies have shown that phytate accumulates in pollen grains late in their development, and that germinating pollen grains are able to take up Pi from the surrounding medium (Helsper et al., 1984). The observed activity of the Pht1;6 and Pht1;7 promoters in tapetum and pollen grains suggests that these transporters are involved in Pi uptake in these tissues.

Finally, the Pht1;4 promoter was found to be active during floral organ abscission. This is the only example of abscission in A. thaliana (Patterson, 2001), and we speculate that this transporter may be involved in remobilisation of Pi out of the senescing floral organs (for subsequent transfer into newly forming flowers), and scavenging of any Pi that leaks out of the exposed vascular bundles in the abscission zone.

Sensitive RT-PCR analysis suggests that the Pht1;7, Pht1;8 and Pht1;9 genes are expressed at low levels in Pi-deprived roots. The fact that no reporter gene expression was observed with constructs containing the promoters of these genes may indicate that this expression was below the threshold of detection for reporter gene assays.

None of the Pht1 promoters tested here showed any activity in embryos, and we were unable to detect any RT-PCR products from any of these genes in silique cDNA. This is unexpected, as the embryo lacks plasmodesmatal connections to maternal tissue, and must therefore take up Pi from the surrounding apoplast. This may indicate that alternative transport mechanisms (such as delivery of phosphorus in an organic form) are functioning in embryos.

The promoter fusion technique used in the present study has revealed expression patterns for the Pht1;3, Pht1;5 and Pht1;6 promoters, despite the fact that Okumura et al. (1998) were unable to detect expression of these genes using Northern analysis. This is presumably due to the fact that these promoters are active only in specific cell types, and illustrates an advantage of this approach for analysing gene expression patterns.

The sequencing of the A. thaliana genome has revealed the presence of a single member of the Pht2;1 family of phosphate transporters in this species. This gene is expressed in all cells of the leaf, and encodes a low affinity Pi transporter (Daram et al., 1999). This transporter is therefore believed to play a role in Pi loading in leaves. Our results suggest that some members of the Pht1 family, including Pht1;5 and Pht1;6, also play a role in loading of Pi internally within tissues rather than uptake from the very low Pi concentrations found in the soil solution, suggesting these may also be low affinity transporters.

In conclusion, the results of this study suggest that members of the Pht1 family of phosphate transporters are expressed in a diverse range of tissues, and are therefore involved not only in uptake of Pi from the soil solution, but also in many other aspects of Pi translocation within the plant.

Experimental procedures

Construction of Pht1 promoter-reporter gene fusions

Pht1 promoters were cloned as transcriptional or translational fusions to the green fluorescent protein (GFP) or β-glucuronidase (GUS) reporter genes in the binary vectors pBI101.3 (Clontech Laboratories, Palo Alto, CA, USA) and pBI101.3-gfp, respectively. The latter vector was constructed by excising the GFP coding sequence and nopaline synthase terminator from pGEM.Ubi1-sgfpS65T (Elliott et al., 1999) using SacI and BamHI, blunting the SacI end using Klenow polymerase, and cloning this into pBI101.3 which had been digested with EcoRI, blunted with Klenow polymerase, and digested with BamHI.

DNA from A. thaliana ecotype Columbia was isolated using the Qiagen DNeasy kit (Qiagen, Pty. Ltd. Clifton Hill, Victoria, Australia), and used as a template to amplify the Pht1 promoter fragments. The promoters were amplified using Pfu high-fidelity polymerase (Stratagene, La Jolla, CA, USA) or Expand High Fidelity polymerase (Roche Diagnostics Australia, Castle Hill, NSW, Australia), according to the manufacturer's instructions. Primers were designed to amplify the promoter lengths shown in Table 1, and restriction sites were incorporated in the primers to facilitate cloning into the binary vectors. Further information on cloning strategies is available on request from the authors. Where present, introns in the untranslated leader sequence were included in the promoter-reporter constructs.

Plant transformation and growth conditions

Binary vectors were introduced into Agrobacterium tumefaciens strain AGL1 (Lazo et al., 1991) by electroporation. Transformation of A. thaliana was done using the floral dip procedure (Clough and Bent, 1998), and transgenic seedlings were selected on 0.5 × MS medium (Murashige and Skoog, 1962) containing 50 µg ml−1 kanamycin. At least 10 independent transgenic lines were generated for each construct.

Plants were grown at 24°C with a 16-h photoperiod and light intensity of 150 µmol m−2 sec−1. To test for promoter induction by phosphate deprivation, T1 or T2 progeny were germinated in a soil/sand mix containing high or low phosphate levels, and grown for 3 weeks. The low phosphate mix contained one part soil with low phosphorus (10 mg total P per kg) and nine parts fine washed sand. To create the high phosphate mix, finely powdered Ca(H2PO4)2.H2O was mixed through the above soil/sand mix at a rate of 60 mg phosphorus per kg of soil/sand mix. Plants growing in this soil/sand mix were watered with a nutrient solution containing 500 µm Ca(NO3)2, 500 µm KNO3, 250 µm MgSO4, 18 µm NaFeEDTA, 45 µm H3BO3, 4.5 µm MnCl2, 315 nm CuCl2, 750 nm ZnCl2 and 15 nm (NH4)6Mo7O24, but no Pi.

Pollen germination was carried out as described by Li et al. (1999).

Reporter gene assays

For reporter gene assays on soil-grown plants, the plants were carefully removed from the pots, soil was washed off the roots with water, and GFP fluorescence was observed in intact seedlings using a Leica MZ6 dissecting microscope with the GFP PLUS fluorescence module (Leica AG, Heerbrugg, Switzerland). Images were collected using a SPOT digital camera and associated software (Diagnostic Instruments, Sterling Heights, MI, USA). Confocal microscopy on intact roots was done using a Leica TCS SP2 confocal on an upright Leica DMRXE microscope.

Histochemical detection of GUS activity was done using the substrate 5-bromo-4-chloro-3-indolyl glucuronide (X-gluc; Jefferson, 1987), by vacuum-infiltrating seedlings in assay buffer (50 mm sodium phosphate pH 7.0, 0.1%Triton X-100, 0.5 mm potassium ferrocyanide, 0.5 mm potassium ferricyanide, and 10 mm EDTA) containing 0.05% X-gluc, and incubating at 37°C for 30 min to overnight. Green tissues were destained with ethanol prior to observation. For sectioning, tissue was briefly fixed in FAA (10% formalin/5% acetic acid/50% ethanol), dehydrated in ethanol, briefly equilibrated in Histoclear (National Diagnostics. Atlanta, GA, USA) and embedded in Paraplast embedding medium (Sigma-Aldrich Corp., St Louis, MO, USA). Embedded tissue was cut into 10 µm sections using a rotary microtome.

Quantitative fluorimetric GUS assays were done using the substrate 4-methyl umbelliferyl glucuronide (MUG) as previously described (Jefferson, 1987). Fluorescence was measured using a Fluoroskan Ascent microtitre plate reader. Protein concentrations were measured using the Bradford assay kit (Bio-Rad Laboratories, Hercules, CA, USA).

RT-PCR analysis

RNA was isolated from a range of A. thaliana tissues, including cotyledons and young leaves from 3-week-old soil-grown plants, old leaves from 5-week-old soil-grown plants, flowers, green siliques, roots from 3-week-old plants grown in high Pi soil, and roots from 3-week-old plants grown in low Pi soil, using an RNeasy kit (Qiagen). Approximately 1 µg of RNA was used as a template for first strand cDNA synthesis, using a Superscript First Strand cDNA Synthesis kit (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Contaminating genomic DNA was removed from the RNA samples using DNase 1 (Qiagen).

One µl of first strand cDNA was then used for PCR using gene-specific primers for each member of the Pht1 family except Pht1;1 and Pht1;2, which were both amplified with the same primer set. Primers were positioned on either side of introns when introns were present within the coding sequence. Details of primer sequences are available from the authors on request. PCR was carried out using EXPAND High Fidelity polymerase (Roche) according to the manufacturer's instructions, using 2.2 mm MgCl2 in 50 µl reactions. Thermal cycling consisted of an initial denaturation at 94°C for 2 min, followed by 10 cycles of denaturation at 94°C for 15 sec, annealing at 50°C for 30 sec, and extension at 72°C for 2 min, and then an additional 20 cycles during which the extension time was increased by 5 sec per cycle, followed by a final extension at 72°C for 7 min. The expected sizes of products amplified from cDNA and genomic DNA are shown in Table 2.

Table 2.  Expected sizes for RT–PCR products
GeneExpected product
from cDNA (bp)
Expected product from
genomic DNA (bp)
Pht1;1 + Pht1;2357508


We would like to thank Dr Rosemary White for assistance with the confocal microscopy, and Dr Agnieszka Mudge for assistance with the fluorimetric GUS assays. This work was supported by GrainGene.