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First published online May 15, 2003; 10.1104/pp.103.020941 Plant Physiology 132:578-596 (2003) © 2003 American Society of Plant Biologists Changes in Gene Expression in Arabidopsis Shoots during Phosphate Starvation and the Potential for Developing Smart Plants1Horticulture Research International, Wellesbourne, Warwick CV35 9EF, United Kingdom (J.P.H., H.C.B., M.R.B., D.C.E., C.R., K.E.W., P.J.W.); and Plant Science Division, Nottingham University, Sutton Bonington Campus, Loughborough LE12 5RD, United Kingdom (J.P.H., M.J.B., S.T.M., R.S.)
Our aim was to generate and prove the concept of "smart" plants to monitor plant phosphorus (P) status in Arabidopsis. Smart plants can be genetically engineered by transformation with a construct containing the promoter of a gene up-regulated specifically by P starvation in an accessible tissue upstream of a marker gene such as  -glucuronidase (GUS). First, using microarrays, we identified genes whose expression changed more than 2.5-fold in shoots of plants growing hydroponically when P, but not N or K, was withheld from the nutrient solution. The transient changes in gene expression occurring immediately (4 h) after P withdrawal were highly variable, and many nonspecific, shock-induced genes were up-regulated during this period. However, two common putative cis-regulatory elements (a PHO-like element and a TATA box-like element) were present significantly more often in the promoters of genes whose expression increased 4 h after the withdrawal of P compared with their general occurrence in the promoters of all genes represented on the microarray. Surprisingly, the expression of only four genes differed between shoots of P-starved and -replete plants 28 h after P was withdrawn. This lull in differential gene expression preceded the differential expression of a new group of 61 genes 100 h after withdrawing P. A literature survey indicated that the expression of many of these "late" genes responded specifically to P starvation. Shoots had reduced P after 100 h, but growth was unaffected. The expression of SQD1, a gene involved in the synthesis of sulfolipids, responded specifically to P starvation and was increased 100 h after withdrawing P. Leaves of Arabidopsis bearing a SQD1::GUS construct showed increased GUS activity after P withdrawal, which was detectable before P starvation limited growth. Hence, smart plants can monitor plant P status. Transferring this technology to crops would allow precision management of P fertilization, thereby maintaining yields while reducing costs, conserving natural resources, and preventing pollution.
Phosphorus (P), an essential mineral nutrient for plants, is required in large amounts to maintain growth (Raghothama, 1999  ; Abel et al., 2002). Plants acquire P in the form of phosphate from the soil solution via their roots. Phosphate exists in the soil solution as one of three orthophosphate species (H3PO4, H2PO4–, and HPO42–) depending on the pH (Bohn et al., 1979). The predominant form taken up by plants is thought to be H2PO4– (Ullrich-Eberius et al., 1984; Schachtman et al., 1998). Plants have evolved many morphological and enzymatic adaptations to tolerate low phosphate availability. Increased root to shoot ratios (Lynch, 1995; Williamson et al., 2001), production of roots in P-rich patches (Drew, 1975; Robinson, 1994), and the proliferation of long root hairs (Bates and Lynch, 2001; Jungk, 2001) are often observed when plants are starved of P. These adaptations enhance the exploitation of the soil volume. In addition, the associations of some plant species with symbiotic mycorrhizal fungi are stimulated by P starvation (Harrison, 1999). The transcription and activity of RNases (Bariola et al., 1994) and acid phosphatases (Goldstein, 1992; Duff et al., 1994; del Pozo et al., 1999; Haran et al., 2000; Baldwin et al., 2001; Miller et al., 2001; Li et al., 2002) are increased by P starvation. These enzymes release phosphate from both cellular (Bariola et al., 1994) and extracellular (Duff et al., 1994) organic compounds. The roots of P-deficient plants also exude organic acids into the soil solution to increase the availability of phosphate bound to soil particles and from phosphate salts (Lipton et al., 1987; Holford, 1997; Raghothama, 1999; López-Bucio et al., 2000). The transcripts and activity of phosphate transporters are increased to optimize uptake and remobilization of phosphate in P-deficient plants (Muchhal et al., 1996; Daram et al., 1999; Kai et al., 2002; Karthikeyan et al., 2002; Mudge et al., 2002; Versaw and Harrison, 2002). The internal P economy of P-starved plants is also improved by increasing the activity of enzymes that replace P in metabolites (Malboobi and Lefebvre, 1997; Plaxton and Carswell, 1999; Ciereszko et al., 2001) and structural compounds, for example the proteins involved in the replacement of chloroplast phospholipids with sulfolipids encoded by the SQD genes (Essigmann et al., 1998; Yu et al., 2002). It is thought that these morphological and enzymatic responses to P starvation are coordinated by both general stress-related and P-specific signaling cascades.
It has been suggested that manipulating the expression of genes to optimize root morphology for exploiting the soil volume, to maximize the uptake of phosphate, or to improve P use efficiency could reduce the phosphate fertilizer requirement of crops (López-Bucio et al., 2000
In addition to their use in laboratory studies to elucidate the signaling cascades initiated by P starvation, smart plants also have a practical use. To realize the growth potential of crop plants, the phosphate concentration in the rhizosphere must be maximized, and to maintain crop yields and quality, the horticultural and agricultural industries in the United States and western Europe routinely apply excessive amounts of phosphate fertilizer (Goldstein, 1992 In this paper, we have identified Arabidopsis genes whose expression increases specifically in response to P starvation, when the P content of plant tissues begins to decline but before the lack of P affects growth. The promoters for these genes are suitable for the generation of smart plants. We have generated transgenic Arabidopsis bearing a construct containing a marker gene (GUS) under the control of the promoter sequence for one of these P-sensitive genes (SQD1), and we demonstrate that these smart plants can be used to monitor P deficiency in plants. Transferring this technology to crop plants will help to manage the application of phosphate fertilizers for sustainable agriculture.
We are interested in developing smart plant technology as both a practical tool to monitor crop P status and as a laboratory tool to discover the regulatory cascades that alter gene expression in response to P deficiency. Smart plants can be genetically engineered by transformation with a construct containing the promoter of a gene up-regulated by P starvation upstream of a gene encoding a visible marker. Detection of the visible marker in the leaves of smart plants indicates P stress and the need for fertilizer application. Because any period of P deficiency that reduces crop growth incurs a yield penalty (Broadley et al., 2002 ), it is important to detect P deficiencies in crops before they become acute and impact on growth. Thus, the P-sensitive promoter driving marker gene expression in the smart plant must become active early enough in response to phosphate depletion to enable remedial fertilizer application. Transcript profiling of Arabidopsis leaves using high-density oligonucleotide arrays was used to identify promoters active in the early stages of incipient P deficiency.
The effects of removing P supply on shoot P concentration and growth was determined on 28-d-old Arabidopsis growing hydroponically (Fig. 1). The shoot fresh weight of plants growing in a solution lacking P (–P solution) was comparable with that of control plants grown in a complete nutrient solution for at least 72 h after P was withdrawn. However, the shoot fresh weight of plants grown without P was significantly (P < 0.001) lower than plants grown in a complete nutrient solution 216 h after P was withdrawn. A decline in P concentration was observed in shoots of Arabidopsis growing in –P solutions. A significant decrease in shoot P concentration in plants lacking a P supply was observed before there was any significant difference in the shoot fresh weight of plants growing in complete and –P solutions. There was no significant difference in the shoot P concentrations of plants growing in complete and –P solutions 24 h after P was withdrawn. However, the shoot P concentrations of plants grown in a –P solution for 72 h were significantly less than those of plants grown in a complete nutrient solution. Thus, between 24 and 72 h after P withdrawal, Arabidopsis grown in –P solutions had a significantly lower shoot P concentration but retained comparable growth to plants grown in a complete nutrient solution. Hence, promoters from genes with up-regulated expression between 24 and about 72 h after P withdrawal could be used to drive the expression of a marker gene in smart plants because they would allow the plant to respond to P deficiency before growth was affected.
To study changes in the Arabidopsis shoot transcriptome in response to P withdrawal, total RNA samples from shoot material were used to challenge Affymetrix GeneChips representing 8,100 Arabidopsis genes. Total RNA was isolated from rosette leaves of Arabidopsis accession Columbia-5 (Col-5) at growth stage 3.90 (Boyes et al., 2001 Between 3,483 and 5,043 transcripts were declared present in the RNA samples used to challenge the Affymetrix GeneChips. Changes in transcript abundance after P starvation were calculated as their abundance in mRNA from shoots of plants grown in –P solutions divided by their abundance in control plants grown in complete nutrient solutions and harvested at the same growth stage. Differentially regulated transcripts were defined as those with a -fold change > 2.5 in two biological replicates. To reduce the number of transcripts falsely classified as differentially regulated, GeneChip signal values less than 20 were reset to 20 (see "Materials and Methods").
For the majority of genes represented on the GeneChip, expression levels appeared unchanged after phosphate withdrawal (Fig. 2). The expression of no gene was differentially regulated at all time points after P withdrawal, and only 10 genes overlapped between two time points (Fig. 2; Tables I and II). The expression of 64 genes was altered >2.5-fold in two biological replicates 4 h after withdrawing P (Table I). This group of genes was termed "early" P response genes. Sixty genes increased their expression (between 2.5- and 59.6-fold), and four genes decreased their expression. There was great variability in the relative expression of some genes at this time point. This can be attributed to biological variability and may be a consequence of the heterogeneous or asynchronous responses of individual plants to P withdrawal. The differences in gene expression between biological replicates are unlikely to arise from experimental error, because the microarray data were normalized and neither biological replicate showed consistently greater changes in expression. Also, variability in expression was observed for genes that were highly expressed. Twenty-eight h after P was withdrawn, the expression of only four genes differed between shoots of plants grown in –P solutions and shoots of plants grown in complete nutrient solutions (Table II). This lull in differential gene expression has also been observed several hours after other environmental challenges (Reymond et al., 2000
Smart plant technology to monitor crop P status requires promoters that increase the expression of a marker gene specifically in response to P starvation. It was therefore necessary to confirm that the genes up-regulated by P starvation in our experiments were unaffected by the withdrawal of other mineral elements (K and N). The shoot K concentration of plants grown in solutions lacking K for 28 h (73.2 µmol g–1 fresh weight) was lower than the shoot K concentration of plants grown in full nutrient solution (83.6 µmol g–1 fresh weight). However, the shoot N concentration of plants grown in solutions lacking N for 28 h was not significantly lower than the shoot N concentration of plants grown in full nutrient solution (3.93% dry weight), although it declined to 3.71% dry weight 100 h after N was withdrawn. The P concentrations in shoots of plants starved of K or N for 28 h did not decline (data not shown). Thus, the changes in gene expression 28 h after the withdrawal of K or N will be responses to a decline in the shoot concentration of K or N specifically. Only one of the early genes (At1g75040; Table I) and three of the late genes (At4g21620, At1g21250, and At2g17840; Table II) responding to P withdrawal responded also to K or N withdrawal (Fig. 2). Furthermore, only two genes responding to P withdrawal (At5g64100 and At1g21750) were found to respond to S starvation (Nikiforova et al., 2003
A literature survey revealed that the expression of several genes that responded to P withdrawal was also affected by other environmental challenges (Tables I and II). Many genes responding to P withdrawal also increased after wounding or pathogen attack (Uknes et al., 1992 Promoters from genes whose expression is increased by P withdrawal but is unaffected by other environmental or developmental challenges can be used in smart plant technologies. The expression of 13 early genes increased specifically in response to P withdrawal (Table I). These encoded proteins with diverse cellular functions, including transcriptional regulation (At2g28710). However, the increase in their expression was transient. Consequently, smart plants based on their promoters would not be suitable for the periodical monitoring of plant P status. The expression of 18 genes was increased 100 h after P withdrawal and appeared unaffected by other stimuli (Table II). One of these genes, a subtilisin-like Ser protease (At2g04160; AIR3), was up-regulated both 28 and 100 h after P withdrawal. Twelve late genes encoded proteins involved in aspects of cellular metabolism (At1g09500, At1g16410, At1g17020, At1g17740, At2g04400, At2g15090, At2g22780, At2g43820, At4g16190, At4g35630, At4g36670, and At5g50850). A further late gene, whose expression responded solely to P withdrawal, encoded a scarecrow-like transcription factor (At4g17230; SCL13). Because increased expression of late genes was sustained and their up-regulation occurred as tissue P declined but before growth was compromised, their promoters would be suitable for smart plant technologies.
The functional categories of the gene products whose expression was altered by P starvation were obtained from the Munich Information on Protein Sequences (MIPS) database using their AGI identifier (Tables I and II). When no category was assigned by functional characterization, the most probable category based on sequence homology was assigned. Approximately 20% to 30% of both the early and late P response gene products were assigned no known function (Fig. 3), which is comparable with the 38% of the 8,100 gene products represented on the Affymetrix GeneChip whose function is unknown.
The early gene products (Table I) were dominated by those involved in metabolism (31%) and in cell rescue and defense (20%). Of the 20 metabolism genes, 13 were involved in primary metabolic functions, such as amino acid metabolism and carbohydrate metabolism, and seven were involved in the production of secondary metabolites involved in plant defense, such as alkaloids and phenylpropanoids. The gene products involved in cell rescue and defense included five chitinases, four peroxidases, a PR-1-like protein, and a tobacco (Nicotiana tabacum) hin-1 homolog. Other significant functions of the early gene products included transcription (6%), protein fate (3%), cellular communication (2%), and transport facilitation (2%). There were nine gene products common to both early and late responses. These included a cytochrome P450 (At4g37370), two glutathione-S-transferases (At1g02920, At2g29460), a FAD-linked oxidoreductase (At1g30700), two peroxidases (At5g64120 and At4g37520), the tobacco hin-1 homolog (At2g35980), and a lectin (At3g15356). Curiously, the expression of no chitinase or endochitinase genes remained elevated 100 h after P withdrawal. In a way similar to the early gene products, roles in metabolism (33%) and cell rescue and defense (16%) dominated the late gene products (Table II). At this time, however, the number of gene products involved in secondary metabolism (11) was greater than those involved in primary metabolism (10). The gene products involved in cell rescue and defense included three peroxidases, a non-race-specific disease resistance protein (NDR1), a putative nematode-resistance protein, thionin, and a respiratory burst oxidase protein. A greater proportion of the late gene products was involved in transcription (8%), protein fate (5%), cellular communication (6%), and transport facilitation (5%). Interestingly, the identity of these genes differed between 4 and 100 h after P withdrawal (Tables I and II). For example, the expression of five transcription factors (three zinc finger proteins, a heat shock transcription factor, and a blue copper-binding protein) was up-regulated 4 h after P withdrawal, but the expression of two other transcription factors (for SCL13 and EREB2) was up-regulated, and three other transcription factors (IAA5, a SUPERMAN-like protein, and a putative transcription factor protein) were down-regulated 100 h after P withdrawal. Because auxin orchestrates plant morphology, it is noteworthy that four of the 12 genes whose expression was down-regulated after 100 h of P starvation were potentially auxin regulated.
To confirm the gene expression patterns observed using microarray technology, quantitative PCR was performed. The expression of several genes with potentially contrasting magnitude and relative changes in expression during P starvation was monitored (Table III). Genes with contrasting levels of expression that responded to the withdrawal of P but not K or N were chosen, which were identified as up-regulated (At4g12470) or down-regulated (At1g10060) solely at the 4-h time point, up-regulated (At2g22780; At5g26340; At1g17740; At4g35630; At1g61800) or down-regulated (At2g18890) solely at the 100-h time point, or up-regulated at both the 4- and 28-h time points (At2g02990) in the microarray analyses. The expression of two genes previously identified as being up-regulated immediately (IPS1; Martín et al., 2000
In general, the results from the quantitative PCR experiments confirmed the changes in gene expression after the withdrawal of P observed using microarray technology (Table III). The temporal changes in gene expression after P withdrawal followed similar patterns of change in experiments using microarrays and quantitative PCR, but the magnitude and kinetics of these changes apparently differed. This difference may be a consequence of using different biological samples, contrasting primers, or normalizing data to different genes. Most of the discrepancies between experiments performed using microarrays and quantitative PCR occurred at the 4-h time point. This probably reflects biological variability and may be a consequence of the general nature of any initial shock response to an environmental perturbation (Desikan et al., 2001
Promoters for genes that are expressed early enough in response to P starvation to enable remedial fertilizer application (< 100 h after withdrawing P), but whose expression is not affected by transient vagaries in environmental conditions (< 4 h after withdrawing P) or by other stimuli are suitable for the generation of smart plants. One such gene is SQD1 (Table III), a gene involved in sulfolipid biosynthesis, originally isolated from a cDNA library with probes homologous to the bacterial sqdB genes by Essigmann et al. (1998 Transgenic Arabidopsis harboring constructs containing the promoter for SQD1 driving the expression of a GUS marker gene showed a gradual increase in leaf GUS activity after P withdrawal (Fig. 4). No GUS activity was observed 20 h before the withdrawal of P. Four hours after P was withdrawn, GUS activity had increased noticeably and appeared to remain constant as plants approached incipient P deficiency (28–100 h). A further marked increase in GUS activity was observed 220 h after the withdrawal of P. These observations demonstrate the potential for smart plant technology based on the SQD1::GUS construct to monitor plant P status.
Changes in Gene Expression after P Withdrawal
Changes in gene expression in Arabidopsis shoots during P starvation assayed using microarray technology (Tables I and II) were confirmed using quantitative PCR techniques (Table III). When P was with-held from Arabidopsis plants growing hydroponically, changes in gene expression occurred that were either specific to P starvation or common to a variety of environmental perturbations (Tables I and II). The expression of 60 genes was transiently up-regulated 4 h after withdrawing P. Many of these genes are ubiquitous "shock" response genes up-regulated by various pathogens and environmental perturbations (Table I; Desikan et al., 2001
Surprisingly, the expression of only four genes differed between nutrient-replete plants and plants starved of P for 28 h (Table II). This lull in differential gene expression has also been observed in the time course of changes in gene expression after other environmental perturbations (Reymond et al., 2000
Smart plants can be used to assay crop P status. They can be genetically engineered in transgenic plants by harnessing a phosphate-sensitive promoter to the expression of a gene encoding a visible marker. Typical marker genes that could be used are GUS or green fluorescent protein. Detection of the visible marker in the leaves of a smart plant would indicate a physiological P stress and the need for fertilizer application. It is important that the expression of the marker gene responds solely to P and increases as tissue P concentration declines but before P starvation affects plants growth. In the experiments described here, the shoot P concentration of Arabidopsis plants had decreased without an effect on growth between 24 and 72 h after P withdrawal (Fig. 1). Promoters for genes whose expression is increased during this time period are suitable for smart plant technology. One such gene is SQD1 (Table III; Essigmann et al., 1998 The smart plant concept was tested using the promoter from the gene SQD1 to control the expression of the GUS marker gene. It was observed that GUS expression in leaves of transgenic Arabidopsis bearing the SQD1::GUS construct increased after the withdrawal of P (Fig. 4) and that an increase could be detected before P starvation affected plant growth. However, because the GUS activity continued to increase with acute P starvation, smart plants based on the SQD1::GUS construct would require careful calibration. In practice, a grower may also need to know whether P stress had been relieved after remedial P fertilization. Therefore, it would be necessary for the expression of any marker gene in a smart plant to return rapidly to basal levels after the application of P. This was not investigated in the present study, but the responsiveness of SQD1 promoter activity to remedial P fertilization is under investigation. In addition, the marker gene product itself would need to have a rapid turnover time that would allow the grower to determine quickly whether remedial P fertilization had been successful. A maximal half-life on the order of a few days is probably necessary.
It is possible that promoters for other P-responsive genes might be used for smart plant technologies. Because P stress can be induced by a variety of factors and because many different signaling and metabolic pathways are implicated, it might be wise to identify a suite of promoters responding differentially to P starvation to allow plant P stress to be monitored in all environments. These promoters could be stacked in a single construct in which each controlled the expression of a different marker gene, for example other colored or fluorescent proteins, such as yellow fluorescent protein, cyan fluorescent protein, and red fluorescent proteins (Lansford et al., 2001
Smart plants can be used as a basic research tool as well as for more applied ends. In the laboratory, smart plants could be used (a) after mutation of the transgenic lines to identify components of signaling cascades impacting on nutrient-specific gene regulation (Martín et al., 2000
Plant Material and Growth Conditions Seeds of Arabidopsis accession Col-5 were obtained from the Nottingham Arabidopsis Stock Centre (N1644; Nottingham, UK). Seeds were washed in 70% (v/v) ethanol-water, rinsed in distilled water, and surface sterilized using 50% (v/v) domestic bleach-water. Seeds were then rinsed and imbibed for 3 to 5 d at 4°C in sterile distilled water to break dormancy.
After imbibition, seeds were sown in un-vented, polycarbonate culture boxes (Broadley et al., 2001 After 21 d, plants were transferred, still on polycarbonate discs, to a hydroponics system situated in a Saxcil growth cabinet (S.K. Saxton Ltd., ARC Works, Cheshire, UK). Plants were grown with a 16-h photoperiod at a temperature of 24°C during the day and 16°C at night. Illumination was provided by a bank of OSRAM L 58 W/23 fluorescent tubes (OSRAM, Langley, UK) giving a photon flux density between 400 and 700 nm of 75 µmol photons m–2 s–1 at plant height. Relative humidity was approximately 80%. Each polycarbonate disc containing plants was placed on a light-proof 500-mL beaker over 450 mL of aerated complete nutrient solution (pH 5.6) containing: 8.0 mM NO3–, 4.025 mM Ca2+, 0.764 mM SO42–, 0.75 mM K+, 0.75 mM Mg2+, 0.25 mM H2PO42–, 0.1mM FeNaEDTA, 0.05 mM Cl–, 0.03 mM H2BO3–, 0.01 mM Mn2+, 0.001 mM Na+, 0.001 mM Zn2+, 0.003 mM Cu2+, and 0.0005 mM MoO42–. Nutrient solution was recirculated using a peristaltic pump at a flow rate of 30 mL min–1 (M045 OEM peristaltic pump, Autoclude, Essex, UK) through four beakers and a central reservoir of 6 L. Four hydroponic units could be operated simultaneously. Nutrient solutions were replaced twice a week. Plants were grown hydroponically for 7 d in complete nutrient solution before experimentation.
To determine the effects of removing the P, K, or N supply on shoot growth and mineral content, the complete nutrient solution was replaced with nutrient solutions lacking these elements. These are referred to as –P, –K, and –N solutions. In solutions lacking P, KH2PO4 was replaced with KH2SO4 to maintain potassium levels. In solutions lacking K, KH2PO4 and KOH were replaced with Ca(H2PO4)2 to maintain phosphate levels. In solutions lacking N, Ca(NO3)2 was replaced with CaSO4 to maintain calcium levels. In the first experiment, which was repeated three times, plants were harvested 1, 3, 8, and 11 d after P was removed from the nutrient solution. At each harvest, the fresh weights of the shoots of 25 plants were determined. The shoot material was then dried at 80°C for 48 h, and the dry weights of individual shoots were determined. Shoot P, K, and N contents were determined from bulked samples of 25 plants. To determine P and K, samples were ashed overnight at 490°C, and the ash was dissolved in 1 mL of nitric acid and 5 mL of water. Solutions were filtered through No. 5 paper (Whatman, Kent, UK) and analyzed for P and K by inductively coupled plasma optical emission spectrophotometry (J Y Horiba Ultima 2 ICP-OES, Jobin Yvon, Middlesex, UK). To determine N, an aliquot of the bulked sample was loaded directly into a combustion analyzer (CN 2000, LECO UK, Stockport, Cheshire, UK) and analyzed for percentage of N via an internal thermal conductivity detector. In the second experiment, which was also repeated three times, plant material was harvested 1, 3, 8, and 11 d after removing N or K from the nutrient solution and also in nutrient-replete plants. Shoot fresh and dry weights and shoot P, K, and N contents were determined as described above.
To determine changes in gene expression in response to phosphate starvation, the complete nutrient solution was replaced by –P nutrient solution 4 h before the midpoint of the light-period 28 d after sowing. Shoot material was harvested 4, 28, and 100 h after the phosphate supply was removed, mid-way through the light period. Shoot material from control plants supplied with complete nutrient solution was also harvested at these time points and also 20 h before P starvation. To identify genes up-regulated specifically in response to P starvation, rather than as a nonspecific response to mineral deficiency, the complete nutrient solution was replaced by –K or –N nutrient solutions. Again, the nutrient solutions were changed 4 h before the midpoint of the light period, and shoot material was harvested 28 h after K or N was removed, mid-way through the light period. At each harvest, shoot material from eight to 12 plants from each treatment was bulked into 1.5-mL colorless, sterile, screw-cap polypropylene tubes and was snap-frozen in liquid nitrogen. Tissue samples were stored at –70°C before the extraction of total RNA. The entire experiment was performed twice.
Tissue samples, previously stored at –70°C, were placed in liquid nitrogen before grinding. To each sample, 1 mL of TRIzol reagent was added, and total RNA was subsequently extracted according to the manufacturer's instructions (Invitrogen Life Technologies, Paisley, UK). Due to the high content of proteoglycans and polysaccharides in plant material, the following modifications were made: (a) After homogenization with the TRIzol reagent, the samples were centrifuged to remove any remaining plant material. The supernatant was then transferred to a clean Eppendorf tube. (b) To aid precipitation of RNA from the aqueous phase, 0.25 mL of isopropanol and 0.25 mL of 1.2 M NaCl solution containing 0.8 M sodium citrate were added. This procedure precipitates the RNA while maintaining the proteoglycans and polysaccharides in a soluble form. Samples of total RNA were then sent to AROS Applied Biotechnology (Aarhus, Denmark) for labeling and Arabidopsis GeneChip analysis (Affymetrix Inc., Santa Clara, CA).
All data manipulations were performed using Microsoft Excel (Microsoft, Redmond, WA). Signal values (indicating the relative abundance of a particular transcript) and detection call values (indicating the probability that a particular transcript is present) were generated by Microarray Analysis Suite 5.0 software (Affymetrix Inc., Santa Clara, CA). Transcripts called "Absent" in both experiments were removed from subsequent analyses. Differences in transcript abundance were expressed as -fold changes, which were calculated as the ratio of signal values in different experiments or treatments. A difference in transcript abundance of 2.5-fold was taken as indicating a difference in gene expression. It has been estimated by challenging Arabidopsis GeneChips with identical RNA samples that the frequency of false calls (differences in transcript abundance 2.5-fold) due to technical reasons approximates 0.04% to 0.2% (Fowler and Thomashow, 2002
The AGI numbers for transcripts present on the Affymetrix Arabidopsis GeneChip were cross-referenced against probe sets on the GeneChip to remove duplicates. The annotation of transcripts was confirmed using the AGI and GenBank numbers given by Affymetrix. The target nucleotide sequences used by Affymetrix to design probes for the GeneChip (Available from http://www.affymetrix.com) were used to BLAST the GenBank sequence database to confirm the identity of the assayed transcript. A similar confirmation of probe fidelity has been undertaken independently by Ghassemian et al. (2001
The identification of cis-regulatory elements (five to 10 bases, with one or no single nucleotide discrepancies) that occurred with a greater frequency within the 10 to 1,000 bases upstream of genes whose expression increased early or late after the withdrawal of P in our study, compared with the regulatory sequences contained upstream of all genes on the Affymetrix GeneChip (P value of 0.05), was performed by the GeneSpring subprogram "Find potential regulatory sequences" (Silicon Genetics, Redwood City, CA). The occurrence of cis-regulatory elements previously identified as common to genes whose expression responds to P starvation (Mukatira et al., 2001
Primers were designed to SQD1 based on its published cDNA sequence (GenBank accession no. AF022082). These consisted of an initial pair of primers (CCCTTCAGTTATCTCACTTAGCAG and AAGCTGAAAGTAGATGCGCCAT) to perform inverse PCR (IPCR) on circularized genomic DNA (Collins and Weissman, 1984
A 1.6-kb region upstream of the SQD1 gene was inserted into the pMOG24 plasmid (Mogen International, Leiden, The Netherlands) upstream of the GUS marker gene. The pMOG24 plasmid was then transformed into Agrobacterium tumefaciens C58 pGV3850 via electroporation (Wei-jun and Forde, 1989
To determine changes in GUS activity in untransformed and transgenic plants bearing the SQD1::GUS construct in response to phosphate starvation, fully expanded leaves of hydroponically growing plants were removed 20 h before and 4, 28, 100, and 220 h after complete nutrient solution was replaced by –P nutrient solution 28 d after sowing. Excised leaves were vacuum infiltrated with staining solution containing 520 mg L–1 5-bromo-4-chloro-3-indolyl
Reverse transcription was performed on 1 µg of total RNA, isolated from plants grown under the same conditions as those grown for microarray analysis, using the ThermoScript RT-PCR system (Invitrogen Life Technologies). The cDNA synthesis reaction was carried out using random hexamers (50 ng µL–1) according to the manufacturer's instructions. Primers for quantitative PCR were designed to the cDNA sequences of genes identified by the microarray screening process and 18S rRNA gene control using the Primer Express Software (v2.0, Applied Biosystems, Cheshire, UK; Table IV). Analysis of transcript levels of each gene was achieved by quantitative PCR using an ABI Prism 7900 HT sequence detector and SYBR Green fluorescent dye. Triplicate reactions (15 µL volume) were performed for each gene and sample (control and –P plants, 4, 28, and 100 h after the withdrawal of P) using 384-well plates consisting of 20-ng cDNA sample, 1 µM forward and reverse primer, and 7.5 µL of 2x SYBR Green PCR master mix (Applied Biosystems). The quantitative PCR reaction conditions were 50°C (2 min) followed by 95°C (10 min) for 1 cycle, then 95°C (15 s) followed by 60°C (1 min) for 40 cycles. This was followed by a dissociation step of 95°C for 15 s, 60°C for 15 s and 95°C for 15 s. The dissociation step was included to generate data for melting curve analysis so that primer dimers or nonspecific products could be detected in the reaction. A control reaction was carried out on each sample using 2 ng of total RNA, to confirm that there was no contaminating chromosomal DNA in the original sample. The expression pattern of individual genes, as suggested by microarray data, dictated which cDNA samples were used to generate standard curves, i.e. the standard curves for genes whose expression increased 100 h after P withdrawal were generated from cDNA samples isolated from plants 100 h after the withdrawal of P. Samples were diluted to give a cDNA concentration range from 10 ng µL–1 to 0.01 ng µL–1. The cycle threshold and normalized fluorescence (
All novel materials described in this publication will be made available upon request in a timely manner for noncommercial research purposes, subject to the requisite permission from any third-party owners of all or parts of the material. Obtaining any permissions will be the responsibility of the requestor.
We thank Andy Bradshaw, Martin Holdsworth, and Steve Coggins (Horticulture Research International [HRI]) for constructing the Arabidopsis hydroponics system; Simon Elliott and Mark Powell (HRI) for mineral analyses; Thomas Thykjaer (AROS Applied Biotechnology, Aarhus, Denmark) for microarray analyses; Martin Sergeant (HRI) for assistance with quantitative PCR; and Andrew Thompson and Ken Manning (HRI) for their comments on the original manuscript. Received January 24, 2003; returned for revision February 28, 2003; accepted March 9, 2003.
Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.103.020941.
1 This work was supported by the Biotechnology and Biological Sciences Research Council (UK), by the Department for Environment, Food and Rural Affairs (UK; project nos. HH0915SFV, HH3502SFV, and HH3501SFV), and by a Horticulture Research International Gordon Browning Studentship to J.P.H. * Corresponding author; e-mail philip-j.white{at}hri.ac.uk; fax 01789–470552.
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ABSTRACT
RESULTS
) or a solution lacking P (
). Plants were 28 d old at the beginning of the experiment. Data are expressed as mean ± SE (n = 3 experiments).
Rn) values were determined for each sample during the quantitative PCR cycling reaction using the ABI Prism sequence detector software (v2.0). The cycle threshold value was calculated using a threshold value set at 0.2. Transcript levels of each gene were normalized to the 18S rRNA control gene. 