Phosphate Utilization Efficiency Correlates with Expression of Low-Affinity Phosphate Transporters and Noncoding RNA, IPS1, in Barley1[W] Chun Y. Huang*, Neil Shirley, Yusuf Genc, Bujun Shi, and Peter Langridge Australian Centre for Plant Functional Genomics, University of Adelaide, Glen Osmond, South Australia 5064, Australia (C.Y.H., N.S., B.S., P.L.); and School of Agriculture, Wine, and Food, University of Adelaide, Adelaide, South Australia 5005, Australia (Y.G.)
Genetic variation in phosphorus (P) efficiency exists among wheat (Triticum aestivum) and barley (Hordeum vulgare) genotypes, but the underlying mechanisms for the variation remain elusive. High- and low-affinity phosphate (Pi) PHT1 transporters play an indispensable role in P acquisition and remobilization. However, little is known about genetic variation in PHT1 gene expression and association with P acquisition efficiency (PAE) and P utilization efficiency (PUE). Here, we present quantitative analyses of transcript levels of high- and low-affinity PHT1 Pi transporters in four barley genotypes differing in PAE. The results showed that there was no clear pattern in the expression of four paralogs of the high-affinity Pi transporter HvPHT1;1 among the four barley genotypes, but the expression of a low-affinity Pi transporter, HvPHT1;6, and its close homolog HvHPT1;3 was correlated with the genotypes differing in PUE. Interestingly, the expression of HvPHT1;6 and HvPHT1;3 was correlated with the expression of HvIPS1 (for P starvation inducible; noncoding RNA) but not with HvIPS2, suggesting that HvIPS1 plays a distinct role in the regulation of the low-affinity Pi transporters. In addition, high PUE was found to be associated with high root-shoot ratios in low-P conditions, indicating that high carbohydrate partitioning into roots occurs simultaneously with high PUE. However, high PUE accompanying high carbon partitioning into roots could result in low PAE. Therefore, the optimization of PUE through the modification of low-affinity Pi transporter expression may assist further improvement of PAE for low-input agriculture systems.
Phosphorus (P) is an essential macronutrient for plant growth and development. The availability of P is often low in soil, and a large amount of inorganic phosphate (Pi) fertilizers is applied to achieve high crop yields. Wheat (Triticum aestivum) and barley (Hordeum vulgare) production uses approximately 46% of the P fertilizers applied to cereals (FAO Fertilizer and Plant Nutrition Bulletin 17; http://www.fao.org). Extensive fertilization with P leads to rapid depletion of nonrenewable P resources and contributes to environmental pollution (Vance et al., 2003). P deficiency in cereal crops is widespread and causes significant yield reductions (Elliott et al., 1997; Gahoonia and Nielsen, 2004). The improvement of P efficiency in winter cereals through breeding is crucial for sustainable agriculture 1 This work was supported by the Grains Research and Development Corporation (to C.Y.H., N.S., B.S., P.L.), the Australian Research Council (to C.Y.H., N.S., B.S., P.L.), the South Australian Government (to C.Y.H., N.S., B.S., P.L.), and the University of Adelaide (to C.Y.H., N.S., Y.G., B.S., P.L.). * Corresponding author; e-mail
[email protected]. au. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Chun Y. Huang (
[email protected]). [W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.111.178459
and world food security (Godfray et al., 2010; Parry and Hawkesford, 2010). There are many definitions of P efficiency in the literature (Gourley et al., 1993; Hammond et al., 2009; Wang et al., 2010), but it can generally be divided into P acquisition efficiency (PAE) and P utilization efficiency (PUE). PAE refers to the ability of the plant to mobilize P from poorly soluble sources and/or to take up the soluble Pi available in the soil solution (Narang et al., 2000), whereas PUE is the amount of biomass produced per unit of acquired P (Ozturk et al., 2005; Wang et al., 2010). PAE is considered to be a major component of overall P efficiency (Ozturk et al., 2005; Ismail et al., 2007; Ramaekers et al., 2010), but PUE can influence PAE. P supply to plants is often fluctuating; therefore, P remobilization within the plant is critical for plant survival (Drew and Saker, 1984; Marschner and Cakmak, 1986; Jeschke et al., 1997; Vance et al., 2003; Huang et al., 2008). Remobilization and translocation of P from shoots to roots can lead to a larger root system, and hence greater exploitation of limited soil P resources for more P. However, the effect of PUE and simultaneous carbohydrate partitioning on PAE (Hermans et al., 2006; Liao et al., 2008) is not clear (Manske et al., 2001; Wang et al., 2010). PHT1 Pi transporters play a critical role in Pi acquisition from soil solution and Pi remobilization within the plant. Nine members of the PHT1 gene family have been found in the Arabidopsis (Arabidopsis thaliana)
Plant PhysiologyÒ, July 2011, Vol. 156, pp. 1217–1229, www.plantphysiol.org Ó 2011 American Society of Plant Biologists
1217
Huang et al.
genome (Mudge et al., 2002) and 13 in rice (Oryza sativa; Paszkowski et al., 2002). Functional characterization shows that some of the PHT1 members, such as AtPHT1;1, AtPHT1;4 (Shin et al., 2004), and OsPHT1;6 (Ai et al., 2009), are high-affinity transporters while others are low-affinity transporters, such as OsPHT1;2 (Ai et al., 2009). Eight PHT1 genes have been described in barley (Smith et al., 1999). The promoter::reporter fusions of HvPHT1;1 and HvPHT1;2 show that reporters are expressed in root hairs, epidermal and cortex cells, and vascular tissues of roots (Schu¨nmann et al., 2004; Glassop et al., 2005). Recently, it was confirmed that HvPHT1;1 is a high-affinity Pi transporter with a Km of 1.9 mM when expressed in Xenopus laevis oocytes (Preuss et al., 2011), suggesting that HvPHT1;1 is involved in Pi acquisition. HvPHT1;6 is expressed in both shoots and roots (Huang et al., 2008) and localized in the leaf phloem tissue (Rae et al., 2003). It displays unsaturated uptake in a millimolar range of Pi concentrations when expressed in oocytes (Preuss et al., 2010), suggesting that HvPHT1;6 is involved in Pi retranslocation. HvPHT1;7 is specifically expressed in green anthers (Druka et al., 2006), and HvPHT1;8 is a mycorrhiza-specific gene (Glassop et al., 2005). HvPHT1;3 and HvPHT1;4 are almost identical in nucleotide sequence. HvPHT1;3 is specifically expressed in roots (Smith et al., 1999). However, the functions of HvPHT1;3 and HvPHT1;4 in Pi homeostasis are not clear. Only one wheat PHT1 gene, TaPHT1;2D1, has been characterized so far (Miao et al., 2009), although several TaPHT1 genes are reported (Davies et al., 2002; Miao et al., 2009). Whole genomic sequences of barley and wheat are not yet available, which has hindered progress toward the functional characterization of PHT1 genes and their regulation in barley and wheat. Pi acquisition and translocation are highly regulated processes. Transcriptional activation of Pi transporter genes is believed to be the major control point in Pi transporter activity (Raghothama, 1999; FrancoZorrilla et al., 2004). IPS (for induced by phosphate starvation) genes have been shown to be involved in P homeostasis (Shin et al., 2006; Franco-Zorrilla et al., 2007; Doerner, 2008; Hammond and White, 2008; Liu et al., 2010). They are a class of non-protein-coding RNAs. Overall sequence identities among IPS genes are relatively low, but 22- to 24-nucleotide sequences are highly conserved among plant species (Hou et al., 2005; Shin et al., 2006). IPS genes are expressed primarily in vascular tissues of roots and shoots (Hou et al., 2005; Shin et al., 2006). Five IPS genes are found in the Arabidopsis genome (Franco-Zorrilla et al., 2007), and two IPS genes are present in monocotyledonous species such as rice, maize (Zea mays), and barley (Hou et al., 2005). The Arabidopsis loss-offunction mutant at4 shows high P accumulation in shoots (Shin et al., 2006). Inversely, overexpression of AtIPS1 and At4 decreases P accumulation in shoots (Franco-Zorrilla et al., 2007), indicating that IPS genes are involved in Pi remobilization and translocation. 1218
MicroRNAs represent a class of noncoding small RNAs that generally function as posttranscriptional negative regulators through base pairing to nearly complementary sequences in target mRNAs (JonesRhoades et al., 2006). miR399 is the first microRNA found to be involved in Pi homeostasis (Sunkar and Zhu, 2004; Fujii et al., 2005; Bari et al., 2006; Chiou et al., 2006). miR399-guided degradation of mRNA of the target gene PHO2, encoding for a ubiquitin E2 conjugase, regulates Pi homeostasis in the plant (Aung et al., 2006; Bari et al., 2006; Mallory and Bouche´, 2008). IPS genes are also involved in the regulation of the miR399-PHO2 pathway (Doerner, 2008). Incomplete sequence complementarity between AtIPS1 and AtmiR399 forms an RNA duplex with a mismatch loop, which inhibits the degradation of AtPHO2 mRNA (FrancoZorrilla et al., 2007). AtPHO2 regulates the expression of AtPHT1;8 and AtPHT1;9, and controls the remobilization and translocation of Pi via an unknown process (Aung et al., 2006; Bari et al., 2006; Chiou et al., 2006; Doerner, 2008). Considerable genetic variation in PAE and PUE has been reported in a wide range of barley and wheat genotypes (Gahoonia and Nielsen, 1996; Manske et al., 2001; Ozturk et al., 2005; Liao et al., 2008; George et al., 2011), but there have been few studies on the genetic variation in expression levels of Pi transporters and the relationships between the expression of Pi transporters and PAE or PUE (Ramaekers et al., 2010). To address this issue, we used barley, a diploid relative of hexaploid wheat, as a model system for winter cereals to study the functions of PHT1 genes involved in the acquisition and remobilization of Pi (Preuss et al., 2010, 2011). In this report, we show that differences in expression levels of four paralogs of the high-affinity Pi transporter gene HvPHT1;1, are not apparent among four barley genotypes differing in PAE, but there is a large difference in the expression of the lowaffinity Pi transporter HvPHT1;6 and a close homolog, HvPHT1;3. In addition, the expression of HvPHT1;6 and HvPHT1;3 correlates with the expression of HvIPS1 and PUE. The implications of these findings in the improvement of PAE for barley and wheat are discussed.
RESULTS Plant Growth, P Concentration, PAE, and PUE among Four Distinct Barley Genotypes
Growing plants in a soil is essential for the evaluation of PAE because it allows the efficient mechanisms operating at the root-soil interface to be functional. These mechanisms include greater exploitation of soil volume by roots, including root hairs, root exudates, and microbial activities. These morphological and biochemical changes can increase P availability by mobilizing sparingly soluble mineral P and organic P sources (Gahoonia and Nielsen, 1997; Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression
Bates and Lynch, 2001; Lambers et al., 2006). We grew barley plants in fine sand mixed with calcium carbonate to simulate calcareous sandy soil. Four distinct barley genotypes (Clipper, Sahara, Pallas, and a hairless mutant, brb, derived from Pallas) were selected. These four barley genotypes were grown in the simulated calcareous sandy soil with three rates of P. At high Pi supply (75 mg P kg21; referred to as P75), shoot dry weights of Pallas, brb, and Sahara were significantly higher than for Clipper, but there were no differences between Pallas, brb, and Sahara (Fig. 1A). At moderately low P supply (22.5 mg P kg21; referred to as P22.5), shoot dry weights of all four genotypes were significantly reduced compared with those at P75 (Fig. 1A). Shoot dry weight of Pallas was significantly higher than that of brb, Clipper, or Sahara. When P supply was further reduced (7.5 mg P kg21; referred to as P7.5), a further reduction in shoot dry weight was observed for all four genotypes, but the shoot dry weight of Pallas was significantly higher than that of brb and Sahara and the shoot dry weight of Clipper was significantly higher than that of Sahara (Fig. 1A). Root growth on a fresh weight basis was reduced by the low rates of P supply, but the reduction was much less than for shoot growth (Fig. 1, A and B). There was no significant interaction between P rates and genotypes. The root fresh weight of Pallas on average was significantly higher than that of the three other genotypes, but no differences were found among brb, Clipper, and Sahara. These results indicate that the low rates of P supply have a smaller impact on root growth than shoot growth. A genotypic difference in root growth was found between Pallas and brb but not between Clipper and Sahara. An increase in root-shoot ratio is often observed in plants grown at low P supply and is attributed to increased carbohydrate partitioning to the root (Zhu et al., 2002; Hermans et al., 2006; Liao et al., 2008). Root-shoot ratios on a fresh weight basis increased as P supply decreased in all genotypes (Fig. 1C). At low P supply (P7.5), root-shoot ratios of brb and Sahara were significantly higher than those of Pallas and Clipper, while the difference between Pallas and Clipper was not significant. At moderately low P supply (P22.5), a similar trend to that at low P supply (P7.5) was observed (Fig. 1C). At high P supply (P75), root-shoot ratios in brb and Clipper were significantly higher than that in Sahara and the root-shoot ratio of Pallas was between these two groups (Fig. 1C). P concentrations of shoots in all four genotypes at P75 were above the critical level of 4,000 mg P g21 dry weight (Fig. 2A), which is required for normal plant growth (Reuter and Robison, 1997; George et al., 2011). Genotypic differences in P concentrations of shoots were observed at P75. Clipper had a significantly higher P concentration than the other genotypes, and the P concentration in Pallas was higher than that in brb or Sahara. When P supply was reduced to P22.5, P concentrations of shoots were reduced to a level below the critical level (Fig. 2A). The shoot P concentration of Plant Physiol. Vol. 156, 2011
Figure 1. Shoot dry weight and root fresh weight of four barley genotypes. A, Shoot dry weight. B, Root fresh weight. C, Root-shoot ratio. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Means and SE values of four replicates are presented. There are significant differences in shoot dry weight and root-shoot ratio for interactions of P rate 3 genotype (P , 0.001). Error bars indicate LSD (LSD0.05) for shoot dry weight and root-shoot ratio. There are no significant differences in root fresh weight for interactions of P rate 3 genotype (P = 0.25).
1219
Huang et al.
action between P rates and genotypes. There was a significant difference among the four genotypes. The shoot P content was significantly higher in Pallas and Clipper than in brb or Sahara, but no difference between Pallas and Clipper was seen. These results indicate that Pallas and Clipper are efficient in P acquisition and that brb and Sahara are inefficient in P acquisition. When biomass production per unit of P in shoots of four genotypes was calculated for PUE, there were no significant differences at the high P supply (P75) among the four genotypes (Fig. 2C). The low rates of P supply at P22.5 and P7.5 increased PUE in all four genotypes, but differences in PUE were found among the four genotypes. The genotypes low in PAE (brb and Sahara) had significantly higher PUE than those with high PAE (Pallas and Clipper) at both P22.5 and P7.5 (Fig. 2C). These results show that all genotypes increase PUE when P supply is limited for optimal growth, and the genotypes with greater PUE at low P conditions have a higher root-shoot ratio (Fig. 1C). This suggests that high remobilization of P occurs simultaneously with enhanced carbohydrate partitioning into roots. Identification of Additional HvPHT1;1 Paralogs in the Barley Genome
Figure 2. P concentration, P content, and PUE of shoots in four barley genotypes. A, P concentration of shoots. B, P content of shoots. C, PUE of shoots. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Means and SE values of four replicates are presented. There are significant differences in P concentrations and PUE for interactions of P rate 3 genotype (P , 0.009). Error bars indicate LSD (LSD0.05) for P concentrations and PUE. There are no significant differences in shoot P content for interactions of P rate 3 genotype (P = 0.39).
Clipper remained higher than in the other genotypes, and Pallas had a significantly higher P concentration than either brb or Sahara. When P supply further decreased to P7.5, P concentrations of shoots were further reduced in all four genotypes (Fig. 2A). The shoot P concentration of Pallas remained higher than that of brb or Sahara. The shoot P concentration of Clipper was also higher than that of Sahara but similar to that of Pallas. Shoot P content, a measure of PAE (Ozturk et al., 2005), was significantly reduced at the two low rates of P supply (Fig. 2B), but there was no significant inter1220
Two paralogs of HvPHT1;1 genes have been described by Smith et al. (1999). We isolated two additional HvPHT1;1 paralogs from a Haruna Nijo bacterial artificial chromosome (BAC) library (Saisho et al., 2007) using a 388-bp probe derived from the coding sequence of HvPHT1;1. The first gene encoding a protein was identical to HvPHT1;1 (Fig. 3A) but differed in the 3# untranslated region (UTR). We designated this new HvPHT1 gene as HvPHT1;9 (GenBank accession no. AM904733). The transcript of HvPHT1;9 had a 56nucleotide deletion in the 3# UTR compared with that of HvPHT1;1 (Fig. 3B). The second new HvPHT1;1 paralog is similar to HvPHT1;2 and is designated as HvPHT1;10 (GenBank accession no. FN392213). The protein sequence of HvPHT1;10 differs only by four amino acid residues from HvPHT1;2 (Fig. 3A). Database searches identified an ortholog of HvPHT1;9 in wheat (GenBank accession no. BJ277773), which also has a 30-nucleotide deletion in the 3# UTR compared with that of TaPHT1;1 (GenBank accession no. CD871730). No Brachypodium sequences similar to either HvPHT1;1 or HvPHT1;9 were identified (Supplemental Fig. S1). Database searches using the coding sequences of HvPHT1;2 and HvPHT1;10 identified four orthologs in wheat and one ortholog in Brachypodium. All five orthologs of HvPHT1;2/HvPHT1;10 are 525 amino acid residues in length. The four wheat orthologs differ from each other only by two amino acid residues (data not shown). One of the four wheat orthologs (GenBank accession no. AJ344240) was used to represent TaPHT1;2 in the phylogenetic analysis of PHT1 proteins Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression Figure 3. Alignment of four HvPHT1 proteins and partial nucleotide sequences of HvPHT1;1 and HvPHT1;9. A, Alignment of four HvPHT1 proteins. Amino acid residues that differ among four HvPHT1 proteins are shown, and the residue numbers are indicated above. B, Alignment of nucleotide sequences of HvPHT1;1 and HvPHT1;9 in the 3# UTR. The nucleotide deletion in HvPHT1;9 is shown as X, and specific primers for quantitative RT-PCR of HvPHT1;1 and HvPHT1;9 are underlined. Accession numbers are as follows: AY188394 (HvPHT1;1), AY187020 (HvPHT1;2), AM904733 (HvPHT1;9), and FN392213 (HvPHT1;10).
(Supplemental Fig. S1). The phylogenetic tree showed that protein sequences of HvPHT1;1/HvPHT1;9, HvPHT1;2, HvPHT1;10, TaPHT1;1/TaPHT1;9, TaPHT1;2, and BdPHT1;2 form a closely related clade (Supplemental Fig. S1), suggesting that HvPHT1;1/HvPHT1;9 and TaPHT1;1/TaPHT1;9 originated from HvPHT1;10/ HvPHT1;2 and TaPHT1;2. Expression of Two HvIPS Genes among Four Barley Genotypes
IPS genes are highly responsive to P deficiency, and their transcript levels are a good biomarker for P deficiency responses (Liu et al., 1997; Martı´n et al., 2000; Wasaki et al., 2003). IPS transcripts also interact with miR399s, regulating P translocation (Franco-Zorrilla et al., 2007). Therefore, the transcript levels of two barley IPS genes (HvIPS1 and HvIPS2) in roots were determined for four barley genotypes. At the sufficient P supply (P75), a moderate level of HvIPS2 transcripts (4 to 7 3 106 normalized copies mg21 RNA) was detected in each of the four genotypes (Fig. 4). Lower P supply enhanced transcript levels of HvIPS2 in all four genotypes to 64 to 74 3 106 normalized copies mg21 RNA at P22.5 and 85 to 109 3 106 normalized copies mg21 RNA at P7.5 (Fig. 4). There was no significant interaction in the transcript levels of HvIPS2 between P rates and genotypes, but there was a significant difference at all P rates among the genotypes. The transcript level of HvIPS2 in brb was significantly higher at all P rates than that in Pallas, and the transcript level of HvIPS2 in Sahara was marginally higher at all P rates than that in Clipper. In contrast, the transcript levels of HvIPS1 at the high P supply (P75) were hardly detectable in any barley genotype (Fig. 4). When P supply was reduced to the moderately low level (P22.5), the transcript levels of HvIPS1 in brb and Sahara were sharply increased to approximately 30 3 106 normalized copies mg21 RNA and 80 3 106 normalized copies mg21 RNA, respectively. However, the transcript levels of HvIPS1 in Pallas Plant Physiol. Vol. 156, 2011
and Clipper at P22.5 increased only to less than 3 3 106 normalized copies mg21 RNA (Fig. 4). A further reduction in P supply to P7.5 increased the transcript level of HvIPS1 only slightly in brb, Pallas, and Clipper, but the transcript level of HvIPS1 in Sahara fell sharply relative to that at P22.5 (Fig. 4). These results show that HvIPS1 and HvIPS2 in all four genotypes are highly inducible under the low rates of P supply, indicating that the plants grown at P22.5 are moderately P deficient and those at P7.5 are severely P deficient. There is a basal level of HvIPS2 expression in the roots of the P-adequate plants but hardly any HvIPS1 transcripts in the roots of the P-adequate plants (Fig. 4). A large variation in the expression of HvIPS1 is found between the P-acquisition-efficient genotypes, Pallas and Clipper, and the P-acquisition-inefficient genotypes, brb and Sahara, especially at the moderately low P supply (P22.5), but this is not seen for HvIPS2 expression. Expression of Four HvPHT1;1 Paralogs in Four Barley Genotypes
At P75, a moderate level of HvPHT1;1 transcripts (3 to 12 3 106 normalized copies mg21 RNA) was detected in the roots of all four genotypes (Fig. 5). The transcript level of HvPHT1;1 in brb was the highest among the four genotypes, and the transcript level of HvPHT1;1 in Sahara was higher than that in Clipper or Pallas (Fig. 5). The decrease in P supply to P22.5 increased HvPHT1;1 transcripts in all four genotypes, especially for Sahara (Fig. 5). Sahara had a much higher level of HvPHT1;1 transcripts than the three other genotypes. The transcript level of HvPHT1;1 in brb was significantly higher than that in Pallas but was similar to that in Clipper. When P supply decreased to P7.5, the transcript levels of HvPHT1;1 further increased in all four genotypes (Fig. 5A). The transcript level of HvPHT1;1 in Sahara remained higher than in the three other genotypes, but there was no significant difference among Pallas, brb, and Clipper (Fig. 5). 1221
Huang et al.
similar size was present in Sahara. Instead, a larger PCR product was observed in Sahara (Supplemental Fig. S2). The sequence analysis of this larger PCR product of HvPHT1;9 from Sahara revealed that it was identical to that of HvPHT1;1. Therefore, the higher transcript levels of HvPHT1;1 detected in Sahara relative to the other genotypes (Fig. 5) could result from combined gene products of both HvPHT1;1 and HvPHT1;9. The transcript levels of HvPHT1;2 (2 to 9 3 106 normalized copies mg21 RNA) at P75 were detected in the roots of all four genotypes (Fig. 5). The moderately low P supply (P22.5) increased the transcript levels of HvPHT1;2 in all four genotypes to more than 4 times those at P75. The low P supply (P7.5) further increased the transcript levels of HvPHT1;2 in Pallas, brb, and Clipper, but the transcript levels of HvPHT1;2 were slightly reduced in Sahara (Fig. 5). There was a significant difference in the transcript level of HvPHT1;2 among the four genotypes. brb and Clipper had much higher transcript levels of HvPHT1;2 than Pallas and Sahara across all three rates of P supply (Fig. 5). The expression of HvPHT1;10 was similar to that of HvPHT1;2 in Pallas and brb but was different in Clipper and Sahara (Fig. 5). The transcript levels of
Figure 4. Transcript levels of two HvIPS genes in roots of four barley genotypes. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Quantitative real-time RT-PCR was used to determine transcript levels in roots. The means (n = 4) of normalized copies mg21 RNA and SE are presented. Log-transformed data were used in the statistical analysis for comparisons of means. There is a significant difference in the expression of HvIPS1 for interactions of P rate 3 genotype (P , 0.001) but not in the expression of HvIPS2 (P = 0.16).
Much lower levels of HvPHT1;9 transcripts (0.4 to 2.3 3 106 normalized copies mg21 RNA) were detected in the roots of Pallas, brb, and Clipper at the sufficient P supply (P75) but not in the roots of Sahara at any P rate (Fig. 5). The low rates of P supply at P22.5 and P7.5 increased the transcript levels of HvPHT1;9 in Pallas, brb, and Clipper, but the transcript levels were approximately 3 times lower than those of HvPHT1;1 (Fig. 5). A genotypic difference in the transcript level of HvPHT1;9 was found at P75 but not at the two low P rates. brb had a significantly higher level of HvPHT1;9 than Pallas or Clipper at P75. PCR amplification of genomic DNA using HvPHT1;9-specific primer pairs generated PCR products of HvPHT1;9 from Pallas, Clipper, and Haruna Nijo (note that the Haruna Nijo BAC library was used to obtain HvPHT1;9 sequence), but no PCR product of 1222
Figure 5. Transcript levels of four paralogs of HvPHT1;1 in roots of four barley genotypes. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Quantitative real-time RT-PCR was used to determine the levels of transcripts in roots. The means (n = 4) of normalized copies mg21 RNA and SE are presented. Log-transformed data were used in the statistical analysis for comparisons of means. There are significant differences in the expression of HvPHT1;1, HvPHT1;9, and HvPHT1;2 for interactions of P rate 3 genotype (P , 0.001 for all three genes) but not for HvPHT1;10 (P = 0.49). Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression
HvPHT1;10 in Clipper and Sahara were at least twice as much as those of HvPHT1;2 at all three rates of P. There was no difference in the transcript level of HvPHT1;10 between Clipper and Sahara across three rates of P supply (Fig. 5). The transcript levels of the four HvPHT1;1 paralogs reveal that all four genes are highly responsive to P deficiency. Although there were genotypic differences in the expression of the four HvPHT1;1 paralogs, the expression pattern was not apparent between two P-acquisition-inefficient genotypes and two P-acquisitionefficient genotypes. Expression of HvPHT1;6 and HvPHT1;3 in Four Barley Genotypes
At sufficient P supply (P75), the transcript levels of HvPHT1;6 in Pallas, brb, and Clipper were less than 0.5 3 106 normalized copies mg21 RNA, but a higher transcript level (9.3 3 106 normalized copies mg21 RNA) was detected in Sahara (Fig. 6). When moderately low P (P22.5) was supplied, the transcript levels of HvPHT1;6 increased to 9 3 106 normalized copies mg21 RNA in brb and 20 3 106 normalized copies mg21 RNA in Sahara (Fig. 6). However, the transcript levels of HvPHT1;6 increased only to 0.7 to 1.3 3 106 normalized copies mg21 RNA in Pallas and Clipper (Fig. 6). With low P supply (P7.5), the transcript level of HvPHT1;6 was further increased in brb to approximately 20 3 106 normalized copies mg21 RNA but was greatly reduced in Sahara relative to that at P22.5 (Fig. 6). The transcript levels of HvPHT1;6 in Pallas and Clipper were also increased at P7.5 but remained lower than 3.0 3 106 normalized copies mg21 RNA. HvPHT1;3 protein has the same number of amino acid residues as HvPHT1;6. HvPHT1;3 protein is more similar to HvPHT1;6 (77%) than the four HvPHT1;1 paralogs (70% identity). The transcript levels of HvPHT1;3 almost mimicked those of HvPHT1;6 (Fig. 6). A large difference in the expression of HvPHT1;6 and HvPHT1;3 was evident between the P-acquisition-efficient and P-acquisition-inefficient genotypes (Fig. 6). Interestingly, the difference in the expression of HvPHT1;6 and HvPHT1;3 is significantly correlated with the expression of HvIPS1 under the low P rates (P7.5 and P22.5) across the four genotypes (r = 0.91 and r = 0.95 at degrees of freedom = 30, respectively; P , 0.001) but not with that of HvIPS2. The P-acquisition-inefficient genotypes, brb and Sahara, had much higher expression levels of HvPHT1;3, HvPHT1;6, and HvIPS1 in roots under the low-P conditions than the P-acquisition-efficient genotypes, Pallas and Clipper (Figs. 4 and 6). Stability of RNA Duplexes Formed between Conserved 24-Nucleotide Motifs of HvIPS1 or HvIPS2 and HvmiR399s
The conserved 24-nucleotide motif of HvIPS1 differs from that of HvIPS2 by six nucleotides in the 11th to 16th nucleotide positions, which correspond to the Plant Physiol. Vol. 156, 2011
Figure 6. Transcript levels of HvPHT1;6 and HvPHT1;3 in roots of four barley genotypes. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Quantitative real-time RT-PCR was used to determine the levels of transcripts in roots. The means (n = 4) of normalized copies mg21 RNA and SE are presented. Log-transformed data were used in the statistical analysis for comparisons of means. There are significant differences in the expression of HvPHT1;6 and HvPHT1;3 for interactions of P rate 3 genotype (P , 0.001).
10th to 14th nucleotide positions of HvmiR399s from the 5# end (Fig. 7, A and B). These HvmiR399s were recently identified by deep sequencing of short reads of P-deficient barley plants of Pallas (Schreiber et al., 2011). By using RNAhybrid software (Rehmsmeier et al., 2004), minimum free energy (MFE) was calculated for the incomplete double-stranded RNA molecules formed between the conserved 24-nucleotide motif of HvIPS1 or HvIPS2 and HvmiR399s (Fig. 7, A and B). The HvIPS1 motif with HvmiR399s could form an RNA duplex with a mismatch loop of two nucleotides only at the 11th to 12th nucleotide positions of HvmiR399s, and the RNA duplexes possess MFE more negative than 230.9 kcal mol 21 (Fig. 7, A and C). In contrast, the 24-nucleotide motif of HvIPS2 with 1223
Huang et al.
between HvIPS1 and HvmiR399s are more stable than those between HvIPS2 and HvmiR399s. We also determined the transcript levels of a barley PHO2 ortholog, HvPHO2. Two primer pairs were used for quantitative reverse transcription (RT)-PCR amplification of both 3# and 5# ends of the HvPHO2 mRNA sequence, similar to the measurement of AtPHO2 described by Bari et al. (2006), given that HvPHO2 has a long mRNA (approximately 4,000 nucleotides) and multiple miR399 target sites in the 5# UTR (data not shown). When a pair of 5# end primers was used (HvPHO2-5), the transcript levels of HvPHO2 in roots were low in all four genotypes at the high rate of P supply (P75), but the transcript level of HvPHO2 in Clipper was significantly higher than in the three other genotypes (Fig. 8). When P supply was reduced to P22.5, the transcript levels of HvPHO2 were significantly increased in brb and Sahara but no change was found in Pallas and Clipper (Fig. 8). The further decrease in P supply to P7.5 enhanced the transcript levels of HvPHO2 in brb, Clipper, and Sahara to a similar level without much change in Pallas. In comparison, the transcript levels of HvPHO2 measured using a pair of primers at the 3# end (HvPHO2-3) were similar to those of HvPHO2-5 in response to P rates and genotypic differences, except for a higher transcript level (Fig. 8). These results indicate that the transcript levels of HvPHO2 in brb, Clipper, and Sahara are increased in the low P supply, and Pallas appears to be less responsive to P supply.
DISCUSSION Genetic Variation in P-Acquisition Efficiency in Four Distinct Barley Genotypes Figure 7. Nucleotide sequence alignment of conserved motifs of HvIPS1 and HvIPS2 with known HvmiR399 members and MFE of RNA duplexes. A, Alignment of the conserved motif of HvIPS1 with known HvmiR399 members. B, Alignment of the conserved motif of HvIPS2 with known HvmiR399s. C, An example of the structure of the RNA duplex formed between HvIPS1 (in red) and HvmiR399b (in green). D, An example of the structure of the RNA duplex formed between HvIPS2 (in red) and HvmiR399b (in green). MFE and the structures of RNA duplexes were obtained with RNAhybrid software (see “Materials and Methods”).
HvmiR399s could form an RNA duplex with a mismatch loop of five nucleotides at the 10th to 11th nucleotide positions of HvmiR399s and a second loop at the 12th to 15th nucleotide positions. The RNA duplexes possess MFE less negative than 227.1 kcal mol21 except for HvmiR399d (Fig. 7, B and D). A higher MFE value was predicted in the RNA duplex formed between HvIPS1 or HvIPS2 and HvmiR399d (Fig. 7, A and B), suggesting that HvIPS1 and HvIPS2 are more effective in sequestering HvmiR399d than the other HvmiR399s. Nevertheless, the RNA duplexes formed 1224
Three of the four genotypes used in this study have a diverse genetic background. Clipper is an Australian barley cultivar, Sahara is a landrace originating from Algeria (Karakousis et al., 2003), Pallas is a northern European barley cultivar (Gahoonia et al., 2001), and the fourth genotype, brb, is a root-hairless mutant derived from Pallas (Gahoonia et al., 2001). Shoot P content can be used as an indication of P acquisition efficiency (Manske et al., 2001; Zhu et al., 2002; Ozturk et al., 2005). The higher P content of shoots in Pallas and Clipper than in brb and Sahara indicates that Pallas and Clipper are P-acquisition-efficient genotypes whereas brb and Sahara are P-acquisition-inefficient genotypes. Our results are consistent with a previous study of Clipper and Sahara where they were grown in a calcareous soil (Zhu et al., 2002). A similar result was also observed for soil-grown Pallas and brb (Gahoonia and Nielsen, 2003). Reduced root surface area in brb is likely to be responsible for its low PAE (Gahoonia and Nielsen, 2003). However, it is not known what is responsible for the low PAE in Sahara. It is noteworthy that PUE in the P-acquisition-inefficient Sahara and brb is greater than that in the P-acquisition-efficient Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression
Figure 8. Transcript levels of HvPHO2 in roots of four barley genotypes. Barley seedlings were grown in a calcareous sandy soil with three rates of P (7.5, 22.5, and 75 mg P kg21 sand, referred to as P7.5, P22.5, and P75, respectively). Plants were harvested at day 16 after seed imbibition. Quantitative real-time RT-PCR was used to determine the levels of transcripts in roots. The means (n = 4) of normalized copies per mg21 RNA and SE are presented. There are significant differences in the expression of HvPHO2-5 and HvPHO2-3 for interactions of P rate 3 genotype (P = 0.006).
Clipper and Pallas under low-P conditions (Fig. 2, B and C). This shows that greater PUE could have an adverse effect on PAE due to a concomitant increase in carbohydrate partitioning into roots, indicated by higher rootshoot ratios (Fig. 1C). Expression Levels of the Genes Related to High-Affinity Pi Transporters in P-Acquisition-Inefficient Genotypes May Not Be the Limiting Factor
All four paralogs of the high-affinity Pi transporter, HvPHT1;1, were expressed in Pallas, brb, and Clipper, whereas expression of only three of the four HvPHT1;1 paralogs could be detected in Sahara (Fig. 5) due to the identical sequences in the 3# UTR between HvPHT1;9 and HvPHT1;1 (Figs. 3B and 5; Supplemental Fig. S2). Our recent work revealed that HvPHT1;1 is a highPlant Physiol. Vol. 156, 2011
affinity Pi transporter with a Km value of 1.9 mM when expressed in Xenopus oocytes (Preuss et al., 2011). Higher transcript abundance of HvPHT1;1 is found in the root hair zone than in the root tip (Preuss et al., 2011). The expression pattern of HvPHT1;1 in root tissues overlaps with HvPHT1;2 (Schu¨nmann et al., 2004; Glassop et al., 2005), suggesting that both HvPHT1;1 and HvPHT1;2 are involved in P acquisition. The two newly identified paralogs of HvPHT1;1 (HvPHT1;9 and HvPHT1;10) are either identical or very similar in protein sequences to HvPHT1;1 and HvPHT1;2 (Fig. 3A) and highly responsive to P deficiency (Fig. 5), suggesting that they have a role in P acquisition similar to HvPHT1;1 and HvPHT1;2. Orthologs (BdPHT1;2, TaPHT1;2, TaPHT1;1, and TaPHT1;9) group with the four barley HvPHT1;1 paralogs in the phylogenetic tree (Supplemental Fig. S1), suggesting that the PHT1 genes involved in P acquisition are conserved in Brachypodium and the Triticeae lineage. There is no apparent pattern in the expression levels of the four HvPHT1;1 paralogs between two P-acquisition-inefficient genotypes (Sahara and brb) and the two P-acquisition-efficient genotypes (Clipper and Pallas) under low-P conditions (Fig. 5). A similar result from two contrasting rice genotypes differing in PAE is also reported using a microarray analysis of seven OsPHT1 genes (Ismail et al., 2007). These data suggest that transcript levels of the high-affinity Pi transporters in P-acquisition-inefficient genotypes may not be a limiting factor if it is assumed that the high-affinity PHT1 is regulated predominantly at the transcript level. Interestingly, the absence of root hairs did not reduce the expression levels of any of the four HvPHT1;1 paralogs. Increased expression levels of HvPHT1;2 and HvPHT1;10 in brb relative to Pallas under the low P supply (Fig. 5) could be due to enhanced systemic Pi signaling initiated by plant low-P status (Figs. 2A and 4). However, the increased expression levels of the four HvPHT1;1 paralogs were also observed in brb relative to Pallas under the high P supply (Fig. 5), when plants have adequate P (Fig. 2A), and there was no increased expression of HvIPS1 (Fig. 4). This suggests that the interception of external Pi signal (Yang and Finnegan, 2010) may be impaired due to the absence of root hairs in brb. Expression Levels of the Genes Related to Low-Affinity Pi Transporters Are Correlated with PUE
HvPHT1;6 is a low-affinity Pi transporter (Preuss et al., 2010). It is expressed in both shoots and roots (Huang et al., 2008) and especially in phloem tissues of the leaf (Rae et al., 2003), suggesting that HvPHT1;6 plays a role in Pi remobilization in the whole plant. The protein sequence of HvPHT1;3 is more similar to HvPHT1;6 than to the HvPHT1;1 paralogs. HvPHT1;3 is grouped to the clade containing OsPHT1;8, BdPHT1;3, and TaPHT1;3 with a high bootstrap value (Supplemental Fig. S1). OsPHT1;8 and the low-affinity Pi transporter OsPHT1;2 have been shown to be involved 1225
Huang et al.
in Pi retranslocation and regulated by the miR399PHO2 signaling pathway (Wang et al., 2009; Liu et al., 2010). In addition, the expression of HvPHT1;3 in roots mimics that of HvPHT1;6 (Fig. 6). These data suggests that HvPHT1;3 may play a role in Pi remobilization in roots similar to HvPHT1;6. Higher expression of HvPHT1;6 and HvPHT1;3 is associated with higher PUE (Figs. 2C and 6). PUE has been proposed to be a bottleneck for further improvement of P efficiency in modern crop cultivars (Wang et al., 2010). Interestingly, the P-acquisition-inefficient genotype Sahara is a landrace, whereas P-acquisition-efficient Clipper is a modern cultivar. Higher remobilization of internal P to metabolically active tissues such as growing root tips and shoot meristems under low-P conditions could promote root growth for access to unexplored P resources in soil while maintaining shoot growth. This would lead to higher PAE. However, higher remobilization of internal P could be associated with higher carbohydrate partitioning into roots, as indicated by higher root-shoot ratios in the P-inefficient genotypes (Fig. 1C). A higher root-shoot ratio is also found in P-inefficient wheat genotypes (Liao et al., 2008). Reduced shoot growth would decrease the production of carbohydrates and consequently have an adverse effect on PAE. Therefore, optimization of Pi remobilization may be necessary for high PAE and high biomass production. Closely related orthologs (BdPHT1;3/BdPHT1;6 and TaPHT1;3/TaPHT1;6) of HvPHT1;3 and HvPHT1;6 were present in both Brachypodium and wheat (Supplemental Fig. S1), suggesting that the low-affinity Pi transporters are conserved within Brachypodium and the Triticeae lineage. Understanding the function and regulation of HvPHT1;3 and HvPHT1;6 could contribute to further improvement of PAE in both barley and other winter cereals. Correlation in the Expression of HvPHT1;3 and HvPHT1;6 with HvIPS1
The 24-nucleotide motifs are conserved in HvIPS1 and HvIPS2 (Fig. 7, A and B), and both genes were highly responsive to P deficiency (Fig. 4). However, genetic variation in gene expression was observed only for HvIPS1. The expression of HvIPS1 is highly correlated with the expression of HvPHT1;3 and HvPHT1;6 in the four barley genotypes tested under low-P conditions (Figs. 4 and 6), suggesting that HvIPS1 plays a unique role in Pi remobilization. The conserved motifs are more variable in two barley IPS genes than those of Arabidopsis (Franco-Zorrilla et al., 2007). The calculated MFE values of RNA duplexes formed between HvIPS1 or HvIPS2 and known HvmiR399s (Fig. 7, A and B) suggest that HvIPS1 is more effective in sequestering HvmiR399s than HvIPS2. Consequently, HvIPS1 could be more effective in the protection of HvPHO2 against HvmiR399-guided cleavage. In contrast to Arabidopsis, MFE values predicted for RNA duplexes formed between 24-nucleotide motifs 1226
of AtIPS genes and AtmiR399s were similar among five AtIPS genes (data not shown). This suggests that mRNA of all five AtIPS genes may have comparable effectiveness in sequestering AtmiR399s as that shown for AtIPS1 (Franco-Zorrilla et al., 2007). The transcript levels of the miR399 target gene HvPHO2 were generally higher in the low-P conditions (Fig. 8) and were parallel to the transcript levels of HvIPS1 to some extent (Fig. 4), suggesting that the miR399-PHO2 signaling pathway is conserved in barley. AtPHO2 has been shown to be involved in the regulation of PHT1 genes such as AtPHT1;8 and AtPHT1;9 in Arabidopsis (Aung et al., 2006; Bari et al., 2006) and OsPHT1;2 and OsPHT1;8 in rice (Wang et al., 2009). Therefore, it is likely that HvPHT1;6 and HvPHT1;3 are also subject to the regulation of the miR399-PHO2 signaling pathway. Notably, there are some differences in the transcript levels of PHO2 among Arabidopsis, rice, and barley. The transcript levels of HvPHO2 in roots are generally higher when the transcript levels of HvIPS1 reach the highest values in severely P-deficient plants except for Sahara (Fig. 8), whereas the expression of AtPHO2 in roots is decreased with an increase of AtIPS genes (Aung et al., 2006; Bari et al., 2006; Franco-Zorrilla et al., 2007). There is no change in the transcript levels of OsPHO2 in low-P conditions relative to high-P conditions when OsIPS1 and OsIPS2 are increased (Zhou et al., 2008). miR399-guided cleavage can decrease transcript levels of PHO2, while IPS transcripts can protect HvPHO2 mRNA against miR399-guided degradation (Franco-Zorrilla et al., 2007). A number of factors could have effects on the transcript levels of PHO2, including (1) PHO2 transcription rates under different P conditions, (2) PHO2 mRNA turnover rate, (3) the abundance of different members of the miR399 family (Lin et al., 2008), (4) the abundance of transcripts from different IPS genes, which vary in the conserved 24-nucleotide motif (Hou et al., 2005; Franco-Zorrilla et al., 2007) and the formation of incomplete doublestranded RNA complexes with different stability (Fig. 7, A and B), and (5) four to five complementarity sites for miR399 present in the 5# UTR of PHO mRNA (FrancoZorrilla et al., 2007), since different members of the miR399 family may have preference for these sites (Allen et al., 2005). The observed differences in the transcript levels of PHO2 between the three species could be due to any combination of these factors or other factors. Further studies are obviously required to elucidate the difference. Despite the differences in the transcript levels of PHO2 among these three species, the correlation in the expression levels between IPS and PHT1 genes related to Pi retranslocation is consistent across the three species. In addition, although the expression levels of HvPHT1;3 and HvPHT1;6 are highly correlated with those of HvIPS1 (Figs. 4 and 6), the correlation in the expression between HvPHT1;3/ HvPHT1;6 and HvPHO2 is not as tight as that with HvIPS1, especially in Sahara at P7.5 (Figs. 6 and 8), suggesting that additional mechanisms may be involved in the regulation of PHO2 and downstream PHT1 genes Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression
in Pi remobilization. There are other regulators involved in P signaling (Lin et al., 2009, 2010). It has been shown that OsSPX1 and OsPHR2 are involved in the regulation of low-affinity Pi transporters such as OsPHT1;2 (Wang et al., 2009; Liu et al., 2010). Sugars as a systemic signal can also regulate the expression of Pi transporters (Hammond and White, 2008). Therefore, the transcript levels of HvPHT1;3 and HvPHT1;6 could be modulated by additional regulators. A further analysis of barley orthologs of OsSPX1 and OsPHR2 could shed light on the regulation of HvPHT1;3 and HvPHT1;6 via the miR399-PHO2 pathway. In summary, Pi retranslocation within the plant is an important mechanism in plant adaptation to variable P supply. Low-affinity Pi transporters are responsible for Pi retranslocation within the plant. The higher expression of the low-affinity Pi transporter genes, HvPHT1;6/HvPHT1;3, could lead to greater PUE, but the concomitant increase in carbohydrate partitioning into roots could have an adverse effect on carbohydrate production and PAE. Therefore, optimization of PUE in plants may be required for high PAE and high yield in low-input agriculture systems. MATERIALS AND METHODS Plant Growth and P Analysis Four barley (Hordeum vulgare) genotypes, Pallas, brb (a mutant of Pallas), Clipper, and Sahara, previously shown to differ in P efficiency (Zhu et al., 2002; Gahoonia and Nielsen, 2003) were used in our experiments. All seeds were multiplied in a potting mix for similar nutrient contents. The weight of seeds used in the experiment was similar, in the range of 52 to 54 mg seed21 for three genotypes (Pallas, brb, and Clipper), but smaller for Sahara (45 mg seed21). The P content of Pallas, brb, Clipper, and Sahara was 259, 212, 285, and 193 mg P seed21, respectively. Siliceous sand was used in the experiments. Basal nutrients and plant growth conditions were as described by Genc et al. (2007). Calcium carbonate powder (0.5%, w/w) was added in the sand to simulate calcareous sandy soil and to reduce P availability (Ryan et al., 1985; Westermann, 1992). Three P rates (7.5, 22.5, and 75 mg P kg21 sand as KH2PO4) were used in the experiments for severely P-deficient, moderately P-deficient, and P-adequate plants, respectively. Plants were harvested at 16 d after seed imbibition. At harvest, shoots of two plants in each pot were cut above the soil surface, and the fresh weight of each plant was recorded. The shoots of one plant were oven dried for shoot dry weight and nutrient analysis, and the shoots of the other plant were frozen in liquid nitrogen for transcript analysis. Shoot P was determined with inductively coupled plasma emission spectrometry (Zarcinas et al., 1987). The roots of two plants in each pot were washed free of sand particles and separated. The excess water in roots was removed with tissue paper, and the fresh weight of roots was recorded. The roots of one plant were frozen immediately in liquid nitrogen for transcript analysis, and the roots of the other plant were used for measurements of root morphology.
roots was prepared using Trizol reagent according to the manufacturer’s instructions (Invitrogen) and treated with DNase I (Ambion). Then, RNA integrity was checked on an agarose gel. Two micrograms of total RNA from roots was used to synthesize cDNA with SuperScript III reverse transcriptase (Invitrogen). The transcript levels of four control genes (barley a-tubulin, heat shock protein 70, glyceraldehyde-3-phosphate dehydrogenase, and cyclophilin) were determined for all cDNA samples, and the most similar three of these four genes were used as normalization controls. Normalization was carried out as described by Vandesompele et al. (2002) and Burton et al. (2008) for the transcript levels of all root cDNA samples. Four biological replicates were used for transcript analysis. Three technical replicates were conducted for each cDNA sample. The normalized copies mg21 RNA were used to represent transcript levels. The primer sequences for all genes determined are listed in Supplemental Table S1.
MFE of RNA Duplexes The RNAhybrid software (http://bibiserv.techfak.uni-bielefeld.de/rnahybrid/ submission.html) was used with default settings to calculate MFE of RNA duplexes formed between HvIPS genes and HvmiR399s (Rehmsmeier et al., 2004).
Statistical Analysis The experimental setup was a completely randomized block design with four replications. Data of plant growth, P nutrition, and transcripts were analyzed using the Genstat Statistical Program (version 11.1; VSN International). The log-transformed data were used in analysis when treatment effects were larger than 10 times. The LSD at P = 0.05 was used for comparisons of means. Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers AM904733 for HvPHT1;9 and FN392213 for HvPHT1;10.
Supplemental Data The following materials are available in the online version of this article. Supplemental Figure S1. Phylogenetic tree of PHT1 Pi transporters in Arabidopsis, rice, Brachypodium, barley, and wheat. Supplemental Figure S2. PCR amplification of genomic DNA of four barley genotypes using HvPHT1;9-specific primers. Supplemental Table S1. Primer sequences used for quantitative, real-time RT-PCR analysis of transcript levels.
ACKNOWLEDGMENTS We are grateful to Dr. Julie Hayes for valuable comments on the manuscript, Dr. Iver Jakobsen for providing the seeds of Pallas and brb, and John Toubia, Margaret Pallotta, and Hui Zhou for technical assistance. Received April 18, 2011; accepted May 20, 2011; published May 23, 2011.
Identification of Two New Paralogs of HvPHT1;1
LITERATURE CITED
A 388-bp probe derived from the coding sequence of HvPHT1;1 (GenBank accession no. AY188394; 945–1,332 bp) was used for screening a Haruna Nijo BAC library (Saisho et al., 2007). Two unique BAC clones were isolated, one containing HvPHT1;10 and the other containing HvPHT1;1, HvPHT1;9, and HvPHT1;2. The two BAC clones were sequenced, and contigs were assembled for HvPHT1;9 and HvPHT1;10.
Ai P, Sun S, Zhao J, Fan X, Xin W, Guo Q, Yu L, Shen Q, Wu P, Miller AJ, et al (2009) Two rice phosphate transporters, OsPht1;2 and OsPht1;6, have different functions and kinetic properties in uptake and translocation. Plant J 57: 798–809 Allen E, Xie ZX, Gustafson AM, Carrington JC (2005) MicroRNA-directed phasing during trans-acting siRNA biogenesis in plants. Cell 121: 207–221 Aung K, Lin SI, Wu CC, Huang YT, Su CL, Chiou TJ (2006) pho2, a phosphate overaccumulator, is caused by a nonsense mutation in a microRNA399 target gene. Plant Physiol 141: 1000–1011 Bari R, Datt Pant B, Stitt M, Scheible WR (2006) PHO2, microRNA399, and
Real-Time Quantitative RT-PCR RNA isolation and real-time quantitative RT-PCR analysis of transcripts were conducted as described by Preuss et al. (2011). Briefly, total RNA from
Plant Physiol. Vol. 156, 2011
1227
Huang et al.
PHR1 define a phosphate-signaling pathway in plants. Plant Physiol 141: 988–999 Bates TR, Lynch JP (2001) Root hairs confer a competitive advantage under low phosphorus availability. Plant Soil 236: 243–250 Burton RA, Jobling SA, Harvey AJ, Shirley NJ, Mather DE, Bacic A, Fincher GB (2008) The genetics and transcriptional profiles of the cellulose synthase-like HvCslF gene family in barley. Plant Physiol 146: 1821–1833 Chiou TJ, Aung K, Lin SI, Wu CC, Chiang SF, Su CL (2006) Regulation of phosphate homeostasis by microRNA in Arabidopsis. Plant Cell 18: 412–421 Davies TGE, Ying J, Xu Q, Li ZS, Li J, Gordon-Weeks R (2002) Expression analysis of putative high-affinity phosphate transporters in Chinese winter wheats. Plant Cell Environ 25: 1325–1339 Doerner P (2008) Phosphate starvation signaling: a threesome controls systemic P(i) homeostasis. Curr Opin Plant Biol 11: 536–540 Drew MC, Saker LR (1984) Uptake and long-distance transport of phosphate, potassium and chloride in relation to internal ion concentrations in barley: evidence of non-allosteric regulation. Planta 160: 500–507 Druka A, Muehlbauer G, Druka I, Caldo R, Baumann U, Rostoks N, Schreiber A, Wise R, Close T, Kleinhofs A, et al (2006) An atlas of gene expression from seed to seed through barley development. Funct Integr Genomics 6: 202–211 Elliott DE, Reuter DJ, Reddy GD, Abbott R (1997) Phosphorus nutrition of spring wheat (Triticum aestivum L.). 1. Effects of phosphorus supply on plant symptoms, yield, components of yield, and plant phosphorus uptake. Aust J Agric Res 455: 855–867 Franco-Zorrilla JM, Gonza´lez E, Bustos R, Linhares F, Leyva A, Paz-Ares J (2004) The transcriptional control of plant responses to phosphate limitation. J Exp Bot 55: 285–293 Franco-Zorrilla JM, Valli A, Todesco M, Mateos I, Puga MI, RubioSomoza I, Leyva A, Weigel D, Garcı´a JA, Paz-Ares J (2007) Target mimicry provides a new mechanism for regulation of microRNA activity. Nat Genet 39: 1033–1037 Fujii H, Chiou TJ, Lin SI, Aung K, Zhu JK (2005) A miRNA involved in phosphate-starvation response in Arabidopsis. Curr Biol 15: 2038–2043 Gahoonia TS, Nielsen NE (1996) Variation in acquisition of soil phosphorus among wheat and barley genotypes. Plant Soil 178: 223–230 Gahoonia TS, Nielsen NE (1997) Variation in root hairs of barley cultivars doubled soil phosphorus uptake. Euphytica 98: 177–182 Gahoonia TS, Nielsen NE (2003) Phosphorus (P) uptake and growth of a root hairless barley mutant (bald root barley, brb) and wild type in lowand high-P soils. Plant Cell Environ 26: 1759–1766 Gahoonia TS, Nielsen NE (2004) Barley genotypes with long root hairs sustain high grain yields in low-P field. Plant Soil 262: 55–62 Gahoonia TS, Nielsen NE, Joshi PA, Jahoor A (2001) A root hairless barley mutant for elucidating genetic of root hairs and phosphorus uptake. Plant Soil 235: 211–219 Genc Y, Huang CY, Langridge P (2007) A study of the role of root morphological traits in growth of barley in zinc-deficient soil. J Exp Bot 58: 2775–2784 George TS, Brown LK, Newton AC, Hallett PD, Sun BH, Thomas WTB, White PJ (2011) Impact of soil tillage on the robustness of the genetic component of variation in phosphorus (P) use efficiency in barley (Hordeum vulgare L.). Plant Soil 339: 113–123 Glassop D, Smith SE, Smith FW (2005) Cereal phosphate transporters associated with the mycorrhizal pathway of phosphate uptake into roots. Planta 222: 688–698 Godfray HCJ, Beddington JR, Crute IR, Haddad L, Lawrence D, Muir JF, Pretty J, Robinson S, Thomas SM, Toulmin C (2010) Food security: the challenge of feeding 9 billion people. Science 327: 812–818 Gourley CJP, Allan DL, Russelle MP (1993) Defining phosphorus efficiency in plants. Plant Soil 155-156: 289–292 Hammond JP, Broadley MR, White PJ, King GJ, Bowen HC, Hayden R, Meacham MC, Mead A, Overs T, Spracklen WP, et al (2009) Shoot yield drives phosphorus use efficiency in Brassica oleracea and correlates with root architecture traits. J Exp Bot 60: 1953–1968 Hammond JP, White PJ (2008) Sucrose transport in the phloem: integrating root responses to phosphorus starvation. J Exp Bot 59: 93–109 Hermans C, Hammond JP, White PJ, Verbruggen N (2006) How do plants respond to nutrient shortage by biomass allocation? Trends Plant Sci 11: 610–617 Hou XL, Wu P, Jiao FC, Jia QJ, Chen HM, Yu J, Song XW, Yi KK (2005)
1228
Regulation of the expression of OsIPS1 and OsIPS2 in rice via systemic and local Pi signalling and hormones. Plant Cell Environ 28: 353–364 Huang CY, Roessner U, Eickmeier I, Genc Y, Callahan DL, Shirley N, Langridge P, Bacic A (2008) Metabolite profiling reveals distinct changes in carbon and nitrogen metabolism in phosphate-deficient barley plants (Hordeum vulgare L.). Plant Cell Physiol 49: 691–703 Ismail AM, Heuer S, Thomson MJ, Wissuwa M (2007) Genetic and genomic approaches to develop rice germplasm for problem soils. Plant Mol Biol 65: 547–570 Jeschke WD, Kirkby EA, Peuke AD, Pate JS, Hartung W (1997) Effects of P deficiency on assimilation and transport of nitrate and phosphate in intact plants of castor bean (Ricinus communis L). J Exp Bot 48: 75–91 Jones-Rhoades MW, Bartel DP, Bartel B (2006) MicroRNAS and their regulatory roles in plants. Annu Rev Plant Biol 57: 19–53 Karakousis A, Barr AR, Kretschmer JM, Manning S, Jefferies SP, Chalmers KJ, Islam AKM, Langridge P (2003) Mapping and QTL analysis of the barley population Clipper x Sahara. Aust J Agric Res 54: 1137–1140 Lambers H, Shane MW, Cramer MD, Pearse SJ, Veneklaas EJ (2006) Root structure and functioning for efficient acquisition of phosphorus: matching morphological and physiological traits. Ann Bot (Lond) 98: 693–713 Liao M, Hocking PJ, Dong B, Delhaize E, Richardson AE, Ryan PR (2008) Variation in early phosphorus-uptake efficiency among wheat genotypes grown on two contrasting Australian soils. Aust J Agric Res 59: 157–166 Lin SI, Chiang SF, Lin WY, Chen JW, Tseng CY, Wu PC, Chiou TJ (2008) Regulatory network of microRNA399 and PHO2 by systemic signaling. Plant Physiol 147: 732–746 Lin S-I, Santi C, Jobet E, Lacut E, El Kholti N, Karlowski WM, Verdeil J-L, Breitler JC, Pe´rin C, Ko S-S, et al (2010) Complex regulation of two target genes encoding SPX-MFS proteins by rice miR827 in response to phosphate starvation. Plant Cell Physiol 51: 2119–2131 Lin WY, Lin SI, Chiou TJ (2009) Molecular regulators of phosphate homeostasis in plants. J Exp Bot 60: 1427–1438 Liu CM, Muchhal US, Raghothama KG (1997) Differential expression of TPS11, a phosphate starvation-induced gene in tomato. Plant Mol Biol 33: 867–874 Liu F, Wang ZY, Ren HY, Shen CJ, Li Y, Ling HQ, Wu CY, Lian XM, Wu P (2010) OsSPX1 suppresses the function of OsPHR2 in the regulation of expression of OsPT2 and phosphate homeostasis in shoots of rice. Plant J 62: 508–517 Mallory AC, Bouche´ N (2008) MicroRNA-directed regulation: to cleave or not to cleave. Trends Plant Sci 13: 359–367 Manske GGB, Ortiz-Monasterio JI, van Ginkel M, Gonzalez RM, Fischer RA, Rajaram S, Vlek PLG (2001) Importance of P uptake efficiency versus P utilization for wheat yield in acid and calcareous soils in Mexico. Eur J Agron 14: 261–274 Marschner H, Cakmak I (1986) Mechanism of phosphorus-induced zinc deficiency in cotton. II. Evidence for impaired shoot control of phosphorus uptake and translocation under zinc-deficiency. Physiol Plant 68: 491–496 Martı´n AC, del Pozo JC, Iglesias J, Rubio V, Solano R, de La Pen˜a A, Leyva A, Paz-Ares J (2000) Influence of cytokinins on the expression of phosphate starvation responsive genes in Arabidopsis. Plant J 24: 559–567 Miao J, Sun JH, Liu DC, Li B, Zhang AM, Li ZS, Tong YP (2009) Characterization of the promoter of phosphate transporter TaPHT1.2 differentially expressed in wheat varieties. J Genet Genomics 36: 455–466 Mudge SR, Rae AL, Diatloff E, Smith FW (2002) Expression analysis suggests novel roles for members of the Pht1 family of phosphate transporters in Arabidopsis. Plant J 31: 341–353 Narang RA, Bruene A, Altmann T (2000) Analysis of phosphate acquisition efficiency in different Arabidopsis accessions. Plant Physiol 124: 1786–1799 Ozturk L, Eker S, Torun B, Cakmak I (2005) Variation in phosphorus efficiency among 73 bread and durum wheat genotypes grown in a phosphorus-deficient calcareous soil. Plant Soil 269: 69–80 Parry MAJ, Hawkesford MJ (2010) Food security: increasing yield and improving resource use efficiency. Proc Nutr Soc 69: 592–600 Paszkowski U, Kroken S, Roux C, Briggs SP (2002) Rice phosphate transporters include an evolutionarily divergent gene specifically acti-
Plant Physiol. Vol. 156, 2011
Genetic Variation in PHT1 and IPS Expression
vated in arbuscular mycorrhizal symbiosis. Proc Natl Acad Sci USA 99: 13324–13329 Preuss CP, Huang CY, Gilliham M, Tyerman SD (2010) Channel-like characteristics of the low-affinity barley phosphate transporter PHT1;6 when expressed in Xenopus oocytes. Plant Physiol 152: 1431–1441 Preuss CP, Huang CY, Tyerman SD (2011) Proton-coupled high-affinity phosphate transport revealed from heterologous characterization in Xenopus of barley-root plasma membrane transporter, HvPHT1;1. Plant Cell Environ 34: 681–689 Rae AL, Cybinski DH, Jarmey JM, Smith FW (2003) Characterization of two phosphate transporters from barley: evidence for diverse function and kinetic properties among members of the Pht1 family. Plant Mol Biol 53: 27–36 Raghothama KG (1999) Phosphate acquisition. Annu Rev Plant Physiol Plant Mol Biol 50: 665–693 Ramaekers L, Remans R, Rao IM, Blair MW, Vanderleyden J (2010) Strategies for improving phosphorus acquisition efficiency of crop plants. Field Crops Res 117: 169–176 Rehmsmeier M, Steffen P, Hochsmann M, Giegerich R (2004) Fast and effective prediction of microRNA/target duplexes. RNA 10: 1507–1517 Reuter DJ, Robison JB (1997) Plant Analysis: An Interpretation Manual. CSIRO Publishing, Melbourne, Australia, pp 86–92 Ryan J, Hassan H, Bassiri M, Tabbara HS (1985) Availability and transformation of applied phosphorus in calcareous soils. Soil Sci Soc Am J 51: 1215–1220 Saisho D, Myoraku E, Kawasaki S, Sato K, Takeda K (2007) Construction and characterization of a bacterial artificial chromosome (BAC) library from the Japanese malting barley variety ‘Haruna Nijo’. Breed Sci 57: 29–38 Schreiber AW, Shi BJ, Huang CY, Langridge P, Baumann U (2011) Discovery of barley miRNAs through deep sequencing of short reads. BMC Genomics 12: 129 Schu¨nmann PHD, Richardson AE, Smith FW, Delhaize E (2004) Characterization of promoter expression patterns derived from the Pht1 phosphate transporter genes of barley (Hordeum vulgare L.). J Exp Bot 55: 855–865 Shin H, Shin HS, Chen R, Harrison MJ (2006) Loss of At4 function impacts phosphate distribution between the roots and the shoots during phosphate starvation. Plant J 45: 712–726 Shin H, Shin HS, Dewbre GR, Harrison MJ (2004) Phosphate transport
Plant Physiol. Vol. 156, 2011
in Arabidopsis: Pht1;1 and Pht1;4 play a major role in phosphate acquisition from both low- and high-phosphate environments. Plant J 39: 629–642 Smith FW, Cybinski DH, Rae AL (1999) Regulation of expression of genes encoding phosphate transporters in barley roots. In Plant Nutrition: Molecular Biology and Genetics. Proceedings of the Sixth International Symposium on Genetics and Molecular Biology of Plant Nutrition, Elsinore, Denmark, August 17–21, 1998. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 145–150 Sunkar R, Zhu JK (2004) Novel and stress-regulated microRNAs and other small RNAs from Arabidopsis. Plant Cell 16: 2001–2019 Vance CP, Uhde-Stone C, Allan DL (2003) Phosphorus acquisition and use: critical adaptations by plants for securing a nonrenewable resource. New Phytol 157: 423–447 Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De Paepe A, Speleman F (2002) Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol 3: RESEARCH0034 Wang C, Ying S, Huang HJ, Li K, Wu P, Shou HX (2009) Involvement of OsSPX1 in phosphate homeostasis in rice. Plant J 57: 895–904 Wang XR, Shen JB, Liao H (2010) Acquisition or utilization, which is more critical for enhancing phosphorus efficiency in modern crops? Plant Sci 179: 302–306 Wasaki J, Yonetani R, Shinano T, Kai M, Osaki M (2003) Expression of the OsPI1 gene, cloned from rice roots using cDNA microarray, rapidly responds to phosphorus status. New Phytol 158: 239–248 Westermann DT (1992) Lime effects on phosphorus availability in a calcareous soil. Soil Sci Soc Am J 56: 489–494 Yang XJ, Finnegan PM (2010) Regulation of phosphate starvation responses in higher plants. Ann Bot (Lond) 105: 513–526 Zarcinas BA, Cartwright B, Spouncer LR (1987) Nitric-acid digestion and multielement analysis of plant-material by inductively coupled plasma spectrometry. Commun Soil Sci Plant Anal 18: 131–146 Zhou J, Jiao FC, Wu ZC, Li YY, Wang XM, He XW, Zhong WQ, Wu P (2008) OsPHR2 is involved in phosphate-starvation signaling and excessive phosphate accumulation in shoots of plants. Plant Physiol 146: 1673–1686 Zhu YG, Smith FA, Smith SE (2002) Phosphorus efficiencies and their effects on Zn, Cu, and Mn nutrition of different barley (Hordeum vulgare) cultivars grown in sand culture. Aust J Agric Res 53: 211–216
1229
