| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
|
First published online April 30, 2004; 10.1104/pp.103.036459 Plant Physiology 135:334-345 (2004) © 2004 American Society of Plant Biologists Phosphatase Under-Producer Mutants Have Altered Phosphorus Relations1Intercollege Program in Plant Physiology (J.L.T., J.D., J.P.L., M.J.G.), Department of Biology (M.C.T., J.D., M.J.G.), and Department of Horticulture (J.P.L., M.J.G.), Penn State University, University Park, Pennsylvania 16802
Phosphorus (P) acquisition and partitioning are essential for plant homeostasis. P is available for plant uptake when in its inorganic form (H2PO4–, or Pi), but Pi is often limiting in soils. Plants secrete acid phosphatases (APases) into the apoplastic space, which may be important for obtaining Pi from organic P sources; however, the relative importance of these enzymes for plant P nutrition has yet to be determined. We demonstrate that the root-associated APase pool is increased in Arabidopsis when Pi is limiting and document five APase isoforms secreted from Arabidopsis roots. Previously, we presented the identification of the phosphatase under-producer (pup) mutants, which have decreased in vivo root APase staining when grown under low P conditions. Here, we present the characterization of one of these, pup3, and further studies with pup1. pup3 has 49%, 38%, and 37% less specific APase activity in exudates, roots, and shoots, respectively. Root-associated APase activity is decreased by 16% in pup1 and 25% in pup3, regardless of P treatment. Two APase activity isoforms are reduced in pup3 exudates, and root and shoot isoforms are also affected. One of the two exudate isoforms is recognized by a polyclonal antibody raised to an Arabidopsis purple APase recombinant protein (AtPAP12); however, AtPAP12 transcript levels are unaffected in the mutant. The pup3 mutation was mapped to 68.4 ± 6.0 centimorgans on chromosome 5. Although P concentrations were not altered in pup1 and pup3 tissues when grown in nutrient solution in which Pi was the sole source of P, the mutants had 10% (pup1) and 17% (pup3) lower shoot P concentrations when grown in a peat-vermiculite mix in which the majority of the total P was present as organic P. Therefore, the pup defects, which include secreted APases, are functionally important for plant P nutrition.
Phosphorus (P) deficiency is a major limitation to plant growth (Marschner, 1995  ; Vance et al., 2003). P is taken up by plants in its inorganic form, Pi or orthophosphate (H2PO4–). Although a macronutrient for plants, the availability of soil Pi is often below that of micronutrients because Pi is immobilized within soil organic complexes, clay complexes, and precipitated salts (for review, see Marschner, 1995). Sparingly available soil Pi can be ameliorated with Pi fertilizers, and applied fertilizers represent more than 80% of world Pi use (Steen, 1998). Modern agriculture relies on crops that provide maximum yields with these fertilizers; however, world resources of extractable Pi are limited, nonrenewable, and increasingly ecologically hazardous to obtain (Steen, 1998). As extractable world Pi stores become depleted, agriculture will be forced to adjust to a lack of Pi fertilizer while continuing to feed an expanding population. Alternatively, the organic P (Po) component of agricultural soil is abundant, representing up to 50% of total soil P (for review, see Vance et al., 2003). Plants may mobilize Po by secreting enzymes into the rhizosphere, including acid phosphatases (APases), ribonucleases, and deoxyribonucleases (for review, see Abel et al., 2002; Vance et al., 2003). Developing plants with enhanced abilities to access the Po component of soils is important to decrease agriculture's reliance on Pi fertilizers. Toward that goal, our group focused on the potential of plant-derived secreted APases to increase Po availability for plant nutrition.
APases may be active against a wide array of organic molecules present in soil Po. These enzymes are nonspecific orthophosphoricmonoester phosphohydrolases (EC 3.1.3.2), cleaving Pi from ester linkage sites. Secreted plant phosphatases preserve >50% activity over a broad pH range (4.0–7.6), maintain >80% activity over a broad temperature range (22°C–48°C), and are stable at temperatures as high as 60°C (LeBansky et al., 1991
The significance of plant-derived secreted APases to plant P nutrition needs to be determined because microbes and fungi within the rhizosphere also contribute to the overall pool of soil phosphatases (for review, see Richardson, 2001
APases are abundant in Arabidopsis and are represented by at least four gene families. A recent survey of the annotated Arabidopsis genome identified sequences for one His APase, four phosphatidic APases, 10 vegetative storage protein APases, and 29 purple APases (PAPs; Li et al., 2002
Two Arabidopsis mutants have been described that are defective in P starvation responses, pho3 (Zakhleniuk et al., 2001
Phosphatase In Vivo Activity Staining Is Reduced in pup3 Roots
The pup mutants were identified by screening for reduced secreted APase activity when seedlings were grown in –P medium containing the APase stain 5-bromo-4-chloro-3-indolyl phosphate (BCIP; Trull and Deikman, 1998
Specific APase Activity Is Reduced in pup3 But Not pup1
As a first step toward characterization of the pup3 defect, specific APase activity measurements were carried out on concentrated root exudates and protein extracts from root and shoot tissues (Fig. 2A). Relative to Col control plants, the pup3 mutant had 49%, 38%, and 37% less specific APase activity in exudates, roots, and shoots, respectively. pup1 exudates showed no change in specific APase activity (data not shown), and APase activity in tissue extracts was shown previously to be normal (Trull and Deikman, 1998
The Rhizospheric APase Pool Is Reduced in pup Mutants
An in vivo root-associated APase activity assay was developed to quantify the reduction in APase activity staining and determine whether rhizospheric APase is affected in the pup mutants (Fig. 2B; Table I). The root-associated APases measured in this assay (and discussed throughout the text) consist of those bound to cell walls at the root surface, those located within the apoplastic space of the root, and those secreted from the root during the course of the assay. Similar assays have been performed on whole root systems of tomato (Boutin et al., 1981 Using this method, we observed that Arabidopsis, like other plants studied with this technique, increases its root-associated APase activity in response to low P conditions (Fig. 2B). A 2 d –P treatment imposed on adult tissues previously grown under sufficient P conditions increased root-associated APase activity by 22% in Col and 25% in Wassilewskija (WS; Fig. 2B; Table I, P < 0.0001). The pup mutants responded to the –P treatment with the same relative magnitude as their controls (no genotype x P treatment interaction; Table I). Taking both P treatments together, root-associated APase activity was reduced by 16% in pup3 and by 25% in pup1 (Table I, P < 0.0001). These decreases in root-associated APase activity indicate that the pup mutants are defective in the rhizospheric APase pool. P concentrations were measured from tissues used in the root-associated APase activity assays (Table I). Although the –P treatment dramatically lowered P tissue concentrations by approximately 25% in roots and more than 30% in shoots (data not shown), mutant plants did not differ from their controls in P accumulation under these conditions (no genotype x P treatment interaction; Table I). This result rules out the possibility that the pup mutants' decreased root-associated APase activity is an indirect result of increased Pi accumulation and subsequent repression of APase activity.
APase isoform analysis was carried out to identify the nature of the decreased activity in the pup mutants (Fig. 3). Concentrated root exudates (Fig. 3A), shoot protein extracts (Fig. 3B), and root protein extracts (Fig. 3C) were run on native PAGE and stained for APase activity. Heavy and heterogeneous glycosylation (Stahl et al., 1994
When extracts were run on SDS-PAGE under nonreducing and otherwise native conditions, pup3 shoot proteins were missing a 90-kD APase isoform (Fig. 3D). This is different from the 160-kD isoform shown previously to be missing in pup1 under these conditions (Trull and Deikman, 1998 ). APase isoforms from pup1 exudates were not different from controls (data not shown).
Protein and transcript accumulation of a low P-regulated PAP (AtPAP12) was carried out to analyze the effects of the pup mutations on this secreted APase. In exudates, one APase isoform, E3, was recognized by the AtPAP12 antibody (Fig. 4A). The decreased APase activity of E3 in pup3 extracts (Fig. 3A) correlated with reduced reactivity of E3 with the AtPAP12 antibody. The other isoform with reduced pup3 APase activity, E2, was not recognized by the AtPAP12 antibody.
The AtPAP12 antibody reacted with three isoforms in Col shoots (Fig. 4B) and two isoforms in Col roots (Fig. 4C). The multiple isoforms recognized in extracts from these tissues may have been due to AtPAP12 dimerization (Schenk et al., 2000 ), cross-reactivity with other PAPs (this antibody cross-reacts with a PAP from Spirodela oligorrhiza; Nakazato et al., 1998), or cross-reaction with carbohydrates from other glycoproteins (Miller et al., 2001). As in exudates, these patterns were altered in the pup3 mutant. The AtPAP12 antibody had reduced reactivity with the upper two isoforms in pup3 shoot extracts (Fig. 4B). Of these two bands, the lower one corresponds to the APase isoform with decreased activity in pup3 shoot extracts (Fig. 3B). The AtPAP12 antibody did not recognize the lower isoform in pup3 root extracts (Fig. 4C), and this isoform migrated at the same location as the isoform that had reduced APase activity in pup3 (Fig. 3C).
Accumulation of AtPAP12 transcripts was analyzed to determine if the pup3 defects occur at this level of regulation (Fig. 4D). The expressed sequence tag (EST) used as a probe is the last 550 bp of the 3' end of the AtPAP12 cDNA, and only one band was recognized on the blot. As shown previously by two other groups (Haran et al., 2000 Proteins from the pup1 mutant were also tested for differences with regards to immunoreactivity with the AtPAP12 antibody and AtPAP12 transcript accumulation, and they were not different from controls (data not shown).
Although the pup mutants accumulated normal P concentrations when grown in a nutrient solution in which P was supplied entirely as Pi (Table I), the pup mutants exhibited altered P relations when P was supplied as Po within a soil substrate (Fig. 5; Table II). The peat-vermiculite soil mix used in this experiment had 2.67 ± 0.14 µg P/g soil; however, only 0.88 ± 0.02 µg P/g soil was available as Pi. Therefore, the majority of P in this substrate was Po. Plants were also watered with a weak fertilizer solution (Somerville and Ogren, 1982
Shoot P concentrations were lower for both the pup3 and pup1 mutants when compared to their controls (Fig. 5A; Table II). In pup3, this decrease was 17% lower with the +Pi fertilizer treatment and 13% lower for the –Pi fertilizer treatment. For pup1, this decrease was not as pronounced but still significant at 13% for the +Pi fertilizer treatment and 7% for the –Pi fertilizer treatment. Total P accumulation was also altered in pup shoots (Fig. 5B; Table II). For pup3 the decrease was 28%/+Pi fertilizer and 27%/–Pi fertilizer, while pup1 accumulated 31% less P in the +Pi fertilizer treatment but the same amount of P in the –Pi fertilizer treatment.
The fertilizer treatment in this study highlights differences between the two ecotypes in their response to applied Pi. While the +Pi fertilizer treatment increased shoot P concentrations in the Col ecotype (Fig. 5A; Table II), the total amount of P accumulated in shoots remained the same (Fig. 5B; Table II). The WS ecotype exhibited the opposite strategy: Shoot P concentrations remained constant with applied Pi fertilizer (Fig. 5A; Table II), while the total amount of P accumulated increased by 30% (Fig. 5B). These results agree with other findings that different Arabidopsis ecotypes demonstrate growth plasticity with regards to P uptake and accumulation (Krannitz et al., 1991
A mapping population to localize the pup3 mutation to a chromosomal region within the Arabidopsis genome was developed using an outcross to the WS ecotype. Since APase activity staining was not a reliable method to determine the pup3 phenotype, the genotype of F2 progeny was determined based on reduced APase activity from crude shoot extracts (Fig. 6). Of the 110 plants assayed, roughly one-quarter had 70% of the crude shoot APase activity of their control parents, which is comparable to the assays with purified pup3 shoot proteins (Fig. 2A). The dispersion pattern of this crude assay implies that pup3 is either a recessive or codominant mutation (Fig. 6). Plants within the lower quartile were confirmed for pup3 isoform phenotypes by native PAGE before codominant amplified polymorphic sequences (CAPS) mapping (Table III). Three markers on chromosome 5 (NIT4, RBCS-B, and ASB2) are linked to the pup3 mutation, with RBCS-B demonstrating the tightest linkage to pup3. Based on the directionality from the other two markers, the pup3 mutation is located at 68.4 ± 6.0 cM on chromosome 5. This position places the pup3 mutation in a different location from both the pup1 mutation and AtPAP12. pup1 was mapped previously between 34.0 cM and 54.8 cM on chromosome 2 (Trull and Deikman, 1998
Plants alter the phosphatase activity of soils. Within the rhizosphere, phosphatase activity is increased and correlates with a zone of Po depletion around plant roots. Roots, microbes, and mycorrhizal fungi all contribute to rhizospheric phosphatase activity. Free-living soil microbes concentrate around root systems because lysed plant cells and root secretions are sources of carbon and nutrients (Marschner, 1995 ). While microbes secrete phosphatases that liberate Pi for plants, they also compete with plants for that resource. Mycorrhizal fungi are also a significant source of soil phosphatases, and they can be major providers of Pi to plants when available P is limiting (Miyasaka and Habte, 2001). However, not all plants form mycorrhizal associations, and in nonmycorrhizal roots (such as Arabidopsis), APase activity is associated with the root surface and rhizosphere (McLachlan, 1980a; Dodd et al., 1987). Plants grown in solution culture mobilize a variety of Po substrates to fulfill their complete P nutritional requirements when deprived of Pi, so plant-derived phosphohydrolases have the capacity to provide for P growth demands (Richardson et al., 2001; Abel et al., 2002). The work presented in this study addresses the importance of plant-derived APases in plant P nutrition using two Arabidopsis mutants that are defective in rhizospheric APase activity.
Previously, the importance of secreted plant APases to plant P nutrition has only been implied. Increased expression of APases under low P conditions is part of an overall P stress response that functions to increase P availability to the plant and is analogous to the microbial PHO regulon (Abel et al., 2002
We establish that Arabidopsis, like other plants previously studied (McLachlan, 1980a
We also present evidence that plant-derived APase activity can be altered by a single mutation, pup3. The pup3 mutant has reduced APase activity as demonstrated by in vivo staining (Fig. 1), root-associated assay (Fig. 2B), and specific activity (Fig. 2A). The amount of reduction in pup3 APase activity, ranging from 25% to 49% (depending on the localization of the assay within the plant), was unexpected because there are more than 20 actively transcribed APase genes within the Arabidopsis genome (Li et al., 2002
The pup3 mutation maps to chromosome 5 within the Arabidopsis genome between At5g29584 and At5g36210, an area encompassing 2.7 Mb of sequence. This region includes AtPAP26, which is closely related to AtPAP12 (Li et al., 2002
The pup1 mutant was described previously (Trull and Deikman, 1998
Although the rhizospheric APase pool is affected by the pup mutations, APase activity in other parts of the plant may be important as well. The effects of PUP3 are not limited to APases secreted into the rhizosphere because the APase activity pool and APase activity isoforms are affected in both root and shoot tissues. Similarly, the PUP1 isoform is present in shoots as well as roots (Trull and Deikman, 1998
The pup mutants demonstrate that secreted and/or cell wall-bound APases are important for P nutrition. When grown in a soil mix containing Po as a major source of P, pup1 and pup3 have lower shoot P concentrations (Fig. 5). However, P concentrations are not decreased when Pi is the sole source of P (Table I). Therefore, the pup1 and pup3 phenotypes are functionally important when Po can be utilized as a P source. This implies that proteins affected by the pup mutants are important for obtaining and/or maintaining shoot P concentrations when grown in soil. Even when Pi is added to the soil mix, the pup mutants cannot concentrate as much Pi in their shoots as their wild-type counterparts. The hydrolysis of Po into Pi adds to the total amount of Pi available for uptake, and in soils Pi uptake is a diffusion-limited process resulting in a zone of Pi depletion around plant roots (Marschner, 1995
Plant Materials, In Vivo Activity Staining, and Growth Conditions
Identification of the pup mutants was based on decreased staining with the APase substrate BCIP or XP (Sigma, St. Louis; Trull and Deikman, 1998
For the in vivo APase staining, seedlings were grown to 5 d postgermination (dpg) in solid medium (0.5x modified Hoagland salts [Johnson et al., 1957
Seedlings were grown to 7 dpg on +P solid medium (0.5x modified Hoagland salts [Johnson et al., 1957 For the assay, plants were removed from the shaking flasks, briefly rinsed in –P medium, and transferred in a time-dependent manner to rocking Magenta boxes containing 10 mL of reaction buffer (–P medium with 19.1 mM MES, 1.3% Suc, and 5 mM p-nitrophenol phosphate, or pNPP). Reactions proceeded for 30 to 50 min at approximately 22°C, then 185 µL of the reaction buffer was removed to 832 µL of 1 N NaOH. p-Nitrophenol (pNP) accumulation was read as A410 and converted to nanomoles by plotting values against a pNP standard curve generated with assay reagents. Roots and shoots were dissected, transferred to preweighed aluminum foil envelopes, and dried for 2 d at 65°C. Root-associated phosphatase activity was calculated as nanomoles pNP liberated by the root system per min per root dry weight (grams). Because the two mutants were generated from different ecotype backgrounds, separate experiments were performed with pup1 and pup3 with their respective controls. Completely randomized designs were used, and each of these experiments was replicated at least twice.
Plants used as material for protein extractions were grown as described (Muchhal et al., 1996 For the protein extraction, roots and shoots were ground in liquid N, then ice-cold extraction buffer (0.1 M K-acetate, pH 5.5, 20 mM CaCl2, 2 mM EDTA, and 0.1 mM phenylmethylsulfonyl fluoride) added to 4 mL/g tissue. Polyvinylpolypyrrolidone was added at 60 mg/g tissue, and the samples were gently agitated at 4°C for 1 h. Samples were centrifuged at 27,000g and 4°C for 30 min, the supernatant removed to a fresh tube, and glycerol added to 20% (v/v). Proteins were quantified with the Bradford method using Coomassie Plus protein assay reagent (Pierce, Rockford, IL) and stored at –80°C. Proteins were isolated from three separate plantings.
Growth medium from the 2-d induction period of the root-associated phosphatase study was passed through a 0.2-µm filter to remove debris, then concentrated >200x with Centriplus-10 centrifugal concentrators (Millipore, Bedford, MA) at 4°C according to manufacturer's instructions. Growth media from four separate flasks were bulked together per exudate sample (therefore, each sample, n, consisted of exudates from 12 plants). Buffer exchange was carried out with protein extraction buffer during the concentration, and glycerol added to 20% (v/v) after concentration was complete. Exuded proteins were stored at –80°C. Exudates were collected from three separate experiments.
Extracted proteins were dialyzed overnight at 4°C in dialysis tubing (6,000–8,000 Mr cutoff; Spectra/Por; Spectrum Laboratories, Los Angeles) against protein extraction buffer with 20% glycerol to remove Pi and other possible APase inhibitors. To measure specific APase activity, 0.5 µg of protein (<40 µL) was added to 300 µL of prewarmed 10 mM pNPP in 50 mM sodium acetate, pH 5.5. Reactions proceeded for 10 min at 25°C, were stopped with 600 µL of 1 N NaOH, and were quantified as for the root-associated APase assay. Samples from each of three protein harvests were assayed simultaneously.
Protein electrophoresis was carried out on 5% stacking/10% resolving (w/v) native acrylamide gels at 30 to 60 V and 4°C. Equal amounts of concentrated exuded proteins (7.5 µg), roots (10 µg), or shoots (28 µg) were loaded for comparison between samples. APase staining was carried out with Fast Black K salt and
Plants for RNA isolation were grown vertically until 16 dpg on either +P (1.2 mM) or –P (9 µM) 0.5x modified Hoagland medium with P-purified phytagel (Trull et al., 1997
Seeds were planted in 6-cm diameter pots in a standard 55% to 65% sphagnum peat moss/perlite mix (Sunshine Aggregate Plus Mix 4; SunGro, Bellevue, WA). The soil mix contained no additional fertilizer. Prepared pots were stored at 4°C for 2 d and placed in the growth chamber under standard conditions (above). Plants were thinned to a density of four plants per pot at 7 dpg and thereafter fertilized twice weekly by subirrigation with either +P or –P fertilizer (Somerville and Ogren, 1982
Total P determinations were carried out on dried tissues using the molybdate method (Murphy and Riley, 1962
Statistical analysis was performed with StatView version 5.0.1 (SAS Institute, Cary, NC). The cutoff was P > 0.1 for values termed not significant.
A pup3 mapping population was generated by outcross to the WS ecotype. F2 mutant segregants were identified first by their decreased APase activity in crude shoot protein extracts. Crude proteins were isolated by grinding individual small leaves from 14 dpg seedlings (grown in peat-vermiculite soil mix) in 500 µL of 50 mM Tris, pH. 7.5, pelleting debris by centrifugation (14,000g, 4°C, 5 min), and collecting the supernatant. APase activity measurements were carried out on these extracts by adding 200 µL of extract to 2 mL of 2.5 mM pNPP in 50 mM sodium acetate, pH 4.8, and the reaction carried out at 37°C for 30 min. Reactions were stopped with the addition of 4 mL of 1 N NaOH. A420 was read with the Brinkmann PC-800 colorimeter (Westbury, NY). APase activity was calculated as A420 per shoot fresh weight per minute and expressed as a percentage of the average between the two control genotypes. Candidate pup3 protein samples with <75% APase activity compared to Col and WS control samples were quantified using the Bradford method (Bradford, 1976
The pup3 mutation was mapped using CAPS as markers (Konieczny and Ausubel, 1993
We thank Dr. Thomas McKnight for use of the AtPAP12 antibody. Discussions with Drs. Jack Schultz, Eva Pell, Seogchang Kang, and Kathleen Brown were also very helpful. Received November 20, 2003; returned for revision February 25, 2004; accepted March 7, 2004.
1 This work was supported by the National Science Foundation (Plant Responses to the Environment Research Training Grant no. 9413204 to John C. Schultz and Eva J. Pell at Penn State University) and the U.S. Department of Agriculture (grant no. 95–37100–1567 to M.C.T., J.P.L., M.J.G., and J.D.). 
2 Present address: Biology Department, CB 3280, University of North Carolina, Chapel Hill, NC 27599.
3 Present address: Monsanto Company, 1920 Fifth Street, Davis, CA 95616. Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.103.036459. * Corresponding author; e-mail tomscha{at}email.unc.edu; fax 919–962–1625.
Abel S, Ticconi CA, Delatorre CA (2002) Phosphate sensing in higher plants. Physiol Plant 115: 1–8[CrossRef][Medline] Berger S, Bell E, Sadka A, Mullet JE (1995) Arabidopsis thaliana Atvsp is homologous to soybean VspA and VspB, genes encoding vegetative storage protein acid phosphatases, and is regulated similarly by methyl jasmonate, wounding, sugars, light and phosphate. Plant Mol Biol 27: 933–942[CrossRef][Web of Science][Medline] Boutin J, Provot M, Roux L (1981) Effect of cycloheximide and renewal of phosphorus supply on surface acid phosphatase activity of phosphorus deficient tomato roots. Physiol Plant 51: 353–360 Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248–254[CrossRef][Web of Science][Medline] Burns R (1986) Interactions of enzymes with soil mineral and organic colloids. In PM Huang, M Schnitzer, eds, Interaction of Soil Minerals with Natural Organics and Microbes, Vol 17. Soil Science Society of America, Madison, WI, pp 429–451 Cashikar AG, Rao NM (1996) Role of the intersubunit disulfide bond in the unfolding pathway of dimeric red kidney bean purple acid phosphatase. Biochim Biophys Acta 1296: 76–84[Medline] Coello P (2002) Purification and characterization of secreted acid phosphatase in phosphorus-deficient Arabidopsis thaliana. Physiol Plant 116: 293–298[CrossRef] del Pozo JC, Allona I, Rubio V, Leyva A, de la Pena A, Aragoncillo C, Paz-Ares J (1999) A type 5 acid phosphatase gene from Arabidopsis thaliana is induced by phosphate starvation and by some other types of phosphate mobilising/oxidative stress conditions. Plant J 19: 579–589[CrossRef][Web of Science][Medline] Dodd J, Burton C, Burns R, Jeffries P (1987) Phosphatase activity associated with the roots and the rhizosphere of plants infected with vesicular-arbuscular mycorrhizal fungi. New Phytol 107: 163–172 Duff SM, Lefebvre DD, Plaxton WC (1991) Purification, characterization, and subcellular localization of an acid phosphatase from black mustard cell-suspension cultures: comparison with phosphoenolpyruvate phosphatase. Arch Biochem Biophys 286: 226–232[CrossRef][Web of Science][Medline] Guame A, Machler F, De Leon C, Narro L, Frossard E (2001) Low-P tolerance by maize (Zea mays L.) genotypes: significance of root growth, and organic acids and acid phosphatase root exudation. Plant Soil 228: 253–264[CrossRef] Guiltinan MJ, Velten J, Bustos MM, Cyr RJ, Schell J, Fosket DE (1987) The expression of a chimeric soybean beta-tubulin gene in tobacco. Mol Gen Genet 207: 324–328
Haran S, Logendra S, Seskar M, Bratanova M, Raskin I (2000) Characterization of Arabidopsis acid phosphatase promoter and regulation of acid phosphatase expression. Plant Physiol 124: 615–626 Hayman AR, Jones SJ, Boyde A, Foster D, Colledge WH, Carlton MB, Evans MJ, Cox TM (1996) Mice lacking tartrate-resistant acid phosphatase (Acp 5) have disrupted endochondral ossification and mild osteoporosis. Development 122: 3151–3162[Abstract] Hunter DA, McManus MT (1999) Comparison of acid phosphatases in two genotypes of white clover with different responses to applied phosphate. J Plant Nutr 22: 679–692 Johnson C, Stout P, Broyer T, Carlton A (1957) Comparative chlorine requirements of different plant species. Plant Soil 8: 337–353[CrossRef] Konieczny A, Ausubel FM (1993) A procedure for mapping Arabidopsis mutations using co-dominant ecotype-specific PCR-based markers. Plant J 4: 403–410[CrossRef][Web of Science][Medline] Koornneef M, Stam P (1992) Genetic analysis. In C Koncz, NH Chua, J Schell, ed, Methods in Arabidopsis Research. World Scientific, River Edge, NJ, pp 81–99 Krannitz PG, Aarssen LW, Lefebvre DD (1991) Relationships between physiological and morphological attributes related to phosphate uptake in 25 genotypes of Arabidopsis thaliana. Plant Soil 133: 169–175[CrossRef] LeBansky B, McKnight T, Griffing L (1991) Purification and characterization of a secreted purple phosphatase from soybean suspension cells. Plant Physiol 99: 391–395
Li D, Zhu H, Liu K, Liu X, Leggewie G, Udvardi M, Wang D (2002) Purple acid phosphatases of Arabidopsis thaliana: comparative analysis and differential regulation by phosphate deprivation. J Biol Chem 277: 27772–27781 Li M, Tadano T (1996) Comparison of characteristics of acid phosphatases secreted from roots of lupin and tomato. Soil Sci Plant Nutr 42: 753–763 Marschner H (1995) Mineral Nutrition of Plants. Academic Press, New York McLachlan KD (1980a) Acid phosphatase activity of intact roots and phosphorus nutrition in plants. I. Assay conditions and phosphatase activity. Aust J Agric Res 31: 429–440[CrossRef] McLachlan KD (1980b) Acid phosphatase activity of intact roots and phosphorus nutrition in plants. II. Variations among wheat roots. Aust J Agric Res 31: 441–448
Miller SS, Liu J, Allan DL, Menzhuber CJ, Fedorova M, Vance CP (2001) Molecular control of acid phosphatase secretion into the rhizosphere of proteoid roots from phosphorus-stressed white lupin. Plant Physiol 127: 594–606 Miyasaka SC, Habte M (2001) Plant mechanisms and mycorrhizal symbioses to increase phosphorus uptake efficiency. Commun Soil Sci Plant Anal 32: 1101–1147[CrossRef]
Muchhal U, Pardo J, Raghothama K (1996) Phosphate transporters from the higher plant Arabidopsis thaliana. Proc Natl Acad Sci USA 93: 10519–10523 Murphy J, Riley JP (1962) A modified single method for the determination of phosphate in neutral waters. Anal Chim Acta 27: 31–36[CrossRef]
Nakazato H, Okamoto T, Nishikoori M, Washio K, Morita N, Haraguchi K, Thompson GA, Jr., Okuyama H (1998) The glycosylphosphatidylinositol-anchored phosphatase from Spirodela oligorrhiza is a purple acid phosphatase. Plant Physiol 118: 1015–1020 Nishikoori M, Washio K, Hase A, Morita N, Okuyama H (2001) Cloning and characterization of cDNA of the GPI-anchored purple acid phosphatase and its root tissue distribution in Spirodela oligorrhiza. Physiol Plant 113: 241–248[Medline] Richardson AE (2001) Prospects for using soil microorganisms to improve the acquisition of phosphorus by plants. Aust J Plant Physiol 28: 897–906 Richardson AE, Hadobas PA, Hayes JE (2001) Extracellular secretion of Aspergillus phytase from Arabidopsis roots enables plants to obtain phosphorus from phytate. Plant J 25: 641–649[CrossRef][Web of Science][Medline]
Rubio V, Linhares F, Solano R, Martin AC, Iglesias J, Leyva A, Paz-Ares J (2001) A conserved MYB transcription factor involved in phosphate starvation signaling both in vascular plants and in unicellular algae. Genes Dev 15: 2122–2133 Schenk G, Guddat LW, Ge Y, Carrington LE, Hume DA, Hamilton S, de Jersey J (2000) Identification of mammalian-like purple acid phosphatases in a wide range of plants. Gene 250: 117–125[CrossRef][Web of Science][Medline] Somerville CR, Ogren WL (1982) Mutants of the cruciferous plant Arabidopsis thaliana lacking glycine decarboxylase activity. Biochem J 202: 373–380[Web of Science][Medline] Stahl B, Klabunde T, Witzel H, Krebs B, Steup M, Karas M, Hillenkamp F (1994) The oligosaccharides of the Fe(III)-Zn(II) purple acid phosphatase of the red kidney bean. Eur J Biochem 220: 321–330[Web of Science][Medline] Staswick PE, Papa C, Huang JF, Rhee Y (1994) Purification of the major soybean leaf acid phosphatase that is increased by seed-pod removal. Plant Physiol 104: 49–57[Abstract] Steen I (1998) Phosphorus availability in the 21st century. Phosphorus and Potassium 217: 25–31
Thomas C, Sun Y, Naus K, Lloyd A, Roux S (1999) Apyrase functions in plant phosphate nutrition and mobilizes phosphate from extracellular ATP. Plant Physiol 119: 543–551 Trull MC, Deikman J (1998) An Arabidopsis mutant missing one acid phosphatase isoform. Planta 206: 544–550[CrossRef][Web of Science][Medline] Trull MC, Guiltinan MJ, Lynch JP, Deikman J (1997) The responses of wild-type and ABA mutant Arabidopsis thaliana plants to phosphorus starvation. Plant Cell Environ 20: 85–92 Vance CP, Uhde-Stone C, Allan DL (2003) Phosphorus acquisition and use: critical adaptations by plants for securing a nonrenewable resource. New Phytol 157: 423–427[CrossRef]
Yan X, Liao H, Trull MC, Beebe SE, Lynch JP (2001) Induction of a major leaf acid phosphatase does not confer adaptation to low phosphorus availability in common bean. Plant Physiol 125: 1901–1911 Zakhleniuk OV, Raines CA, Lloyd JC (2001) pho3: a phosphorus-deficient mutant of Arabidopsis thaliana (L.) Heynh. Planta 212: 529–534[CrossRef][Web of Science][Medline] This article has been cited by other articles:
|
|||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| ASPB Publications | PLANT PHYSIOLOGY® | THE PLANT CELL | |
|---|---|---|---|
TOP
ABSTRACT
RESULTS
8 for each treatment. Each sample (n) is the activity of three plants grown together in liquid culture. Error bars are SE. Statistical analyses were performed with ANOVA (
-naphthyl acid phosphate. Markers are native protein electrophoresis markers (left), and APase isoforms are noted on the right. Isoforms designated with an asterisk have decreased activity in pup3 exudates. Samples were 7.5 µg of concentrated root exudate proteins (A), 28 µg of shoot proteins (B), and 10 µg of root proteins (C). D, Shoot extracts run under SDS-PAGE but otherwise native conditions and subsequently stained for APase activity also show a reduced pup3 isoform. Markers for D are Rainbow high Mr markers (Amersham Pharmacia Biotech).
AtPAP12 antibody. Markers are native protein electrophoresis markers (left). Arrows to the right indicate proteins with reduced reactivity in pup3. D, Total RNA (9.25 µg/lane) isolated from 16-dpg plants grown on a continuous treatment of either high or low P was probed sequentially with an EST corresponding to AtPAP12 or 
