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First published online July 24, 2009; 10.1104/pp.109.143180 Plant Physiology 151:199-209 (2009) © 2009 American Society of Plant Biologists OPEN ACCESS ARTICLE
Molecular and Biochemical Characterization of AtPAP15, a Purple Acid Phosphatase with Phytase Activity, in Arabidopsis1,[W],[OA]School of Biological Sciences, University of Hong Kong, Pokfulam, Hong Kong, China (R.K., K.-H.C., B.L.L.); and Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada T2N 1N4 (E.Y.)
Purple acid phosphatase (PAP) catalyzes the hydrolysis of phosphate monoesters and anhydrides to release phosphate within an acidic pH range. Among the 29 PAP-like proteins in Arabidopsis (Arabidopsis thaliana), AtPAP15 (At3g07130) displays a greater degree of amino acid identity with soybean (Glycine max; GmPHY) and tobacco (Nicotiana tabacum) PAP (NtPAP) with phytase activity than the other AtPAPs. In this study, transgenic Arabidopsis that expressed an AtPAP15 promoter::β-glucuronidase (GUS) fusion protein showed that AtPAP15 expression was developmentally and temporally regulated, with strong GUS staining at the early stages of seedling growth and pollen germination. The expression was also organ/tissue specific, with strongest GUS staining in the vasculature, pollen grains, and roots. The recombinant AtPAP purified from transgenic tobacco exhibited broad substrate specificity with moderate phytase activity. AtPAP15 T-DNA insertion lines exhibited a lower phytase and phosphatase activity in seedling and germinating pollen and lower pollen germination rate compared with the wild type and their complementation lines. Therefore, AtPAP15 likely mobilizes phosphorus reserves in plants, particularly during seed and pollen germination. Since AtPAP15 is not expressed in the root hair or in the epidermal cells, it is unlikely to play any role in external phosphorus assimilation.
At pH in the range of 4 to 7, purple acid phosphatases (PAPs) catalyze the hydrolysis of a wide range of activated phosphoric acid monoesters and diesters and anhydrides (Klabunde et al., 1996  ). They are distinguished from the other phosphatases by their insensitivity to L-(+) tartrate inhibition and therefore are also known as tartrate-resistant acid phosphatases. Their characteristic pink or purple color derives from a charge transfer transition between a Tyr residue and the "chromophoric" ferric ion in the binuclear Fe(III)-Me(II) center, where the metal (Me) is iron, zinc, or manganese (Schenk et al., 1999). PAP proteins are also characterized by seven conserved amino acid residues (shown in boldface) in the five conserved motifs DXG, GDXXY, GNH(D/E), VXXH, and GHXH, which are involved in the coordination of the dimetal nuclear center (Li et al., 2002).
PAPs are widespread in mammals, fungi, bacteria, and plants. Interestingly, while only a few copies of PAP-like genes are present in mammalian and fungal genomes (Mullaney and Ullah, 2003
Some PAP members can hydrolyze phytic acid (myoinositol hexakisphosphate [InsP6]) to inorganic phosphate and free or lower phosphoric esters of myoinositol. Since the major storage form of phosphorus in plant seeds and pollen grains is phytate, PAPs with phytase activity may play a role in seed and pollen germination. However, not all PAPs exhibit phytase activity. The first plant phytase PAP, GmPHY, was isolated from the cotyledon of germinating soybean seedlings (Hegeman and Grabau, 2001
Relatively little is known about the biochemical properties and physiological roles of the 29 PAP-like Arabidopsis genes (del Pozo et al., 1999 In this study, AtPAP15 expressed in a plant (tobacco) system was biochemically characterized, and its temporal and spatial expression patterns in Arabidopsis were examined. The physiological roles of AtPAP15 in phosphorus mobilization were also delineated.
Overexpression and Purification of AtPAP15 in Transgenic Tobacco Plants The soluble GST-AtPAP15 protein did not show any enzymatic activity. Therefore, a His-tagged AtPAP15 protein was stably overexpressed in tobacco plants using an explant method. Gene expression was confirmed by PCR (data not shown) and western-blot analysis using specific anti-AtPAP15 antiserum (Fig. 1A ). Phytase activity was approximately 3-fold greater in transgenic tobacco leaves compared with wild-type leaves (Fig. 1B). Leaves from permanent lines were used for further protein purification.
The purification of the AtPAP15 protein was achieved by ion-exchange, affinity, and gel filtration chromatography. The purification table of a representative run is shown in Table I . A single polypeptide band of approximately 60 kD was detected by silver staining (Fig. 2A , lane 5), which confirmed the homogeneity of the protein. The apparent molecular mass of the native enzyme was estimated to be approximately 58 kD, using preparative-grade gel filtration chromatography (data not shown), confirming that the native enzyme is a monomeric protein. Western-blot analysis (Fig. 2B) and mass spectrometry were employed to confirm the identity of the protein. The sequence coverage of the peptide mass fingerprint reached 25% (134 of 532 residues) and the MOWSE score was 66 (Supplemental File S1), confirming its identity as AtPAP15. The enzyme had been purified approximately 344-fold with an overall recovery of 2.4%; it exhibited a phytase activity of 10 units mg–1 protein (Table I).
Biochemical Properties of Purified AtPAP15
pH and Temperature Effects on AtPAP15 Phytase Activity
Effects of Ions and Inhibitors on Enzymatic Activity of AtPAP15 We investigated the influence of various ions on the enzymatic activity of AtPAP15. Ca2+ and Zn2+ stimulated the phytase activity of AtPAP15 (127.5% and 129.4%, respectively), whereas Cu2+ and SO32– inhibited this activity (reduced to 17.8% and 43.1%, respectively). Other ions (Mn2+, Mg2+, Ni2+, SO42–, and NO3–) exhibited no effects on phytase activity. The most prominent PAP inhibitor, molybdate, was also the most notable inhibitor of AtPAP15; 0.25 mM MoO4– was sufficient to completely abolish enzymatic activity. The presence of 5 mM F– or PO43– ions reduced the residual activities of the enzyme to 20% or 0%, respectively. The purified enzyme was significantly resistant to treatment with tartrate. Other compound supplements such as EDTA and citrate exerted no influence on enzymatic activity (Fig. 4 ).
Substrate Specificity of Purified AtPAP15 Purified AtPAP15 exhibited broad substrate specificity (Table II ), with p-nitrophenyl-phosphate (pNPP), phosphoenolpyruvate (PEP), and Na-pyrophosphate being the most effective substrates. Compared with activity toward pNPP, the activities toward various deoxyribonucleotide triphosphates were approximately 31% to 52% and the activity toward phytate was 7%. Very low activity was seen when monophosphates (AMP or GMP) were used as substrates. Negligible activity was detected on Glc-1-P.
Kinetic Parameters of Purified AtPAP15 The kinetic parameters of AtPAP15 were measured when pNPP, PEP, or Na-phytate was used as substrate (Table III ). AtPAP15 had relatively higher affinity (Km = 278 µM) and lower Vmax (13.44 units mg–1) toward Na-phytate than toward pNPP or PEP. The catalytic efficiencies (kcat/Km) of the enzyme toward pNPP and PEP were approximately 10-fold higher than that toward Na-phytate.
The biochemical properties of the recombinant AtPAP15 from plant were very different from that presented by Zhang et al. (2008) , who reported the specific activities of a GST:AtPAP15 toward a few substrates. In our study, the GST:AtPAP15 produced from E. coli was inactive. Zhang et al. (2008) reported the specific activity of GST:AtPAP15 toward pNPP to be only 0.546 units mg–1, which was far lower than the plant-derived AtPAP15 level (165 units mg–1; Table III) and that reported for the other plant PAPs (222–368 units mg–1 for tomato [Solanum lycopersicum] PAPs [Bozzo et al., 2002] and 266 units mg–1 for KbPAP [Vogel et al., 2002]). Regarding specific activities toward phytate, the two recombinant AtPAPs gave 10.0 (plant) and 0.7 (bacterial) units mg–1, respectively.
To determine the expression patterns of AtPAP15 with respect to specific tissue and developmental stages, transgenic Arabidopsis plants expressing an AtPAP15 promoter::GUS fusion protein were produced. All three promoter regions directed the same pattern of GUS expression under normal growth conditions (data not shown). Intense GUS staining was observed at the early stages of seedling establishment and in the cotyledon, radicle, and hypocotyl (Fig. 5, A–D ). The signal was stronger and more diffuse in the cotyledon leaves but weaker and more restricted in younger immature leaves (Fig. 5E). In mature, fully expanded rosette leaves of soil-grown Arabidopsis transgenic lines, GUS staining was predominantly detected in the vasculature (Fig. 5F). In roots, intense staining could be observed at all stages but was mainly restricted to the vascular cylinder (Fig. 5G). No expression could be found in the root hairs (Fig. 5G) or in the root cap cells (Fig. 5H). GUS staining was consistently detected in the vasculature of both the stems and the roots. Cross-sectional analysis revealed that the staining was mainly localized to the xylem and phloem tissues in leaf, hypocotyl, and root sections (Fig. 5, I–L).
In reproductive organs, GUS staining was obvious in the flowers (Fig. 5M); however, little or no staining was evident in the developing seeds or siliques (data not shown). Blue color was observed in the anther and also the stigma papillae (Fig. 5N). Pollen squash analysis indicated that the majority of the GUS staining was from pollen grains (Fig. 5O). To further analyze GUS expression in flowers, AtPAP15 promoter::GUS transgenic Arabidopsis anthers were sectioned to reveal the stage at which GUS expression was evident. Cross-sections of Arabidopsis anthers revealed little or no GUS gene expression in the early stage of pollen development (Fig. 6, A–D ). This expression increased at later stages of pollen development and was obvious in mature pollen grains (Fig. 6, E and F). Staining could also be detected in in vitro-germinated pollen (Fig. 5P).
The promoter::GUS data indicated that AtPAP15 expression is developmentally and temporally regulated, with strong GUS staining at early stages of seedling growth and at late stages of pollen development. The expression was also organ/tissue specific, with the vasculature, pollen grains, and seedlings showing the strongest staining.
To analyze AtPAP15::GUS expression in response to a variety of stimuli, two independent homozygous transgenic lines were used for in situ analysis of GUS activity. In general, the overall GUS staining patterns in shoots and roots were not affected by these treatments. The only exception was at the root tip, where an increase of GUS activity was observed under salicylic acid, abscisic acid, NaCl, sorbitol, or mannitol treatment (Fig. 7 ). In contrast, no alterations in GUS staining were observed under nutrient stresses, such as phosphorus, nitrogen, or potassium starvation (data not shown).
Molecular Characterization of AtPAP15 T-DNA Insertion Mutants and Their Complementation Six T-DNA insertion lines at the AtPAP15 locus are available from the Arabidopsis Biological Resource Center, of which we chose SAIL_529_D01 (T9) and SALK_061597 (T2), which insert into exon 1 and exon 2, respectively, to be used in this study. T2 and T9 seeds from the distribution stock were germinated, and segregation of the T-DNA-encoded antibiotic resistance marker was tracked. Genomic DNA was then extracted from the antibiotic-resistant T2 progeny, and this DNA was screened for T-DNA insertion by PCR using gene-specific primers and primers anchored in the T-DNA borders. Six (T2-1, -3, -5, -6, -8, and -9) and three (T9-2, -7, and -9) individual plants were selected as homozygotes by the absence of the AtPAP15 PCR product (data not shown). Gene silencing of the homozygous plants was confirmed by reverse transcription (RT)-PCR and western blotting. In RT-PCR assays, a single amplified DNA fragment with the predicted size (approximately 1.6 kb) was absent in T-DNA mutants when compared with wild-type plants (Fig. 8A ). Protein was extracted from seedlings for western-blot analysis via AtPAP15 antiserum. No AtPAP15 expression was observed in these insertion mutants compared with wild-type plants (Fig. 8B). A slightly smaller protein band was recognized in all lines by the anti-PAP15 antiserum (Fig. 8, B and C). This band is not AtPAP15 because it was present in all T-DNA lines, which were shown not to express AtPAP15 mRNA by RT-PCR (Fig. 8A). This band, which was not present in tobacco (Fig. 2B), could be an AtPAP protein with high amino acid sequence homology to AtPAP15. Western blotting of wild-type plants using the preimmune serum did not give any band (Supplemental Fig. S1).
To verify that the phenotypic difference was due to the disruption of AtPAP15, complementation lines were produced by introducing the construct AtPAP15 in the NOS promoter-containing pCAMBIA1300 into the mutants. Four homozygous T3 generation lines bearing one copy of the complemented gene were selected based on segregation of hygromycin resistance; these lines were verified by PCR screening and western blotting (Fig. 8C). Two lines (PC1 and PC2) were chosen for subsequent studies (Fig. 8, A and B).
Based on results of the above GUS assay, AtPAP15 is highly expressed in early stages of germination, implying that it may play a role(s) in seed germination or seedling growth. However, the insertion mutants and their complementation lines did not differ in growth performance, phenotypes, or seed germination rate when planted in soil (data not shown). Therefore, the phosphatase activities of 2-d-old seedlings were measured. It was observed that both phytase activity and acid phosphatase (APase) activity were significantly lower in the T-DNA mutants than in the wild type, whereas the complementation line showed comparable value to the wild type (Table IV ). Since GUS studies indicated that AtPAP15 expression was substantial during pollen germination, the enzyme activities in germinating pollen were also assayed, which showed similar results (Table IV). In general, the specific activities of phytase and phosphatase in pollen were higher than that in the seedlings among all lines, possibly due to a lower complexity of total protein in pollen.
An in vitro pollen germination experiment was then conducted. As shown in Figure 9 , the pollen germination rate was significantly reduced in the mutants (30%–35%) compared with wild-type plants (78%). Complementation of AtPAP15 mutants with pCAMBIA1300-PAP15 resulted in recovery of the mutagens relative to the wild-type response (65%). These results indicated that the pollen germination phenotype of the AtPAP15 mutant correlates to AtPAP15 deficiency.
Only two PAP-like genes (Flanagan et al., 2006 ) are present in animal genomes, compared with the greatly larger numbers in plants (Li et al., 2002). It is intriguing that so many different PAPs are required for plant metabolism and how different the physiological functions of these PAPs are. In this study, we focused on AtPAP15, a PAP with phytase activity. Not all PAPs exhibit phytase activity; for instance, no phytase activity has been reported for mammalian PAPs. In addition, some plant PAPs, including KbPAP (Cashikar et al., 1997), two secreted tomato PAPs (Bozzo et al., 2002), and an intracellular tomato PAP (Bozzo et al., 2004), lack phytase activity.
Our results confirmed that AtPAP15 is an acidic phosphatase with phytase activity. Phytate (InsP6), which is primarily complexed with metal ions, is the principal storage form of phosphorus in seeds (Otegui et al., 2002
The seedlings of T-DNA lines only exhibited 35% to 55% of the phytase activity of wild-type seedlings, but this reduction did not affect the germination rate of the seedlings. The expression of redundant phytase genes and the abundance of phosphorus in the seeds may explain the normal seed germination rate of the T-DNA lines. During pollen germination, pollen of T-DNA lines only exhibited 25% to 57% phytase activity and 59% to 71% APase activity compared with the wild type (Table IV), and this reduction is correlated with a lower in vitro germination rate of the pollen (Fig. 9). These results indicated that AtPAP15 is a key phytase/phosphatase during pollen germination and implicated a physiological role in the mobilization of phosphorus reserves in pollen. It is unlikely that AtPAP15 is secreted from pollen for external phosphorus assimilation. During in vitro germination of Lilium longiflorum pollen, in-gel acid phosphatase activity staining of germination medium only generated a single 32-kD tartrate-resistant acid phosphatase; therefore, it is unlikely to be an AtPAP15-like protein (approximately 60 kD; Ibrahim et al., 2002
PAPs with phytase activity have been identified in soybean (Hegeman and Grabau, 2001
AtPAP23 also exhibits phytase activity (Zhu et al., 2005 AtPAP15 expression was not restricted to the reserve storage organs; its expression was also strong in the plant vascular tissues (Fig. 5, E–L), where phytate amounts are scarce. Furthermore, it is noteworthy that phytate was not the only substrate of AtPAP15 (Table II). Rather, it was active toward many organic phosphorus compounds. Vascular tissues allow the transport of energy compounds from the shoots to the roots and the transport of nutrients from the roots to the shoots. AtPAP15 may mediate the remobilization of inorganic phosphates from organic phosphorus compounds and may help maximize the phosphorus efficiency of the plant by redistributing surplus phosphorus from the mature to the growing tissues. This possibility is consistent with the lack of AtPAP15 expression in root tips and hairs (Fig. 5, G and H), where very large amounts of inorganic phosphate are consumed for building biomolecules such as DNA and RNA.
Arabidopsis does not secrete phytases from its roots (Richardson et al., 2001
A recent report proposed that AtPAP15 helps to supply the plant with myoinositol for ascorbate synthesis (Zhang et al., 2008
AtPAP15 Cloning, Vector Construction, and Tobacco Transformation The Columbia ecotype of Arabidopsis (Arabidopsis thaliana) was used throughout this study. Total RNA was extracted using the TRIzol method (Invitrogen) from 3-week-old Arabidopsis plants. Total cDNA was transcribed using Moloney murine leukemia virus reverse transcriptase (Promega) according to the manufacturer's instructions. The open reading frame (ORF) of AtPAP15 was amplified by forward (5'-TATGTCGACATGACGTTTCTACTACTTCTAC-3') and reverse (5'-GACTAGTTCAGTGGTGGTGGTGGTGGTGGCAATGGTTAACAAGGCGGT-3') primers by Pfx polymerase (Roche). A 6x His tag was appended to the reverse primer to create a C-terminal His tag.
The AtPAP15 ORF was then subcloned into a pBa002a-derived plant expression vector carrying a Basta-resistant gene and a cauliflower mosaic virus 35S promoter. The expression construct of pBa002a-PAP15 was mobilized into Agrobacterium tumefaciens strain GV3101 by freeze-thaw transformation (Hofgen and Willmitzer, 1988
The signal peptide of AtPAP15 was predicted by the SignalP 3.0 server (Bendtsen et al., 2004
Fifty grams of transgenic tobacco leaf tissue was ground in liquid nitrogen and extracted with 50 mM sodium acetate buffer (pH 5.0) freshly supplemented with 1 mM phenylmethylsulfonyl fluoride and 5 mM dithiothreitol. The leaf tissue extracts were then centrifuged at 12,000g twice, and the supernatant was used for protein concentration and enzyme activity measurements. The enzyme was further purified to homogeneity by three chromatographic procedures (Table I). All chromatography was carried out on an ÄKTA purifier FPLC system (GE Healthcare), sequentially with a cation-exchange column (HiPrep 16/10 CM FF, 16 x 100 mm), a Ni2+ affinity column (HiTrap HP, 1 mL), and a gel filtration column (Superdex 75, 10 x 300 mm) following the manufacturers' instructions. The purified protein was concentrated and stored at –80°C for further assays. The purified protein was fractionated by 10% (w/v) SDS-PAGE (MiniII; Bio-Rad) and visualized by silver staining; the protein was further confirmed by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry analysis (Lung et al., 2008
The native molecular mass of the purified enzyme was estimated by gel filtration (Superdex 75, 10 x 300 mm) using the ÄKTA purifier FPLC system (GE Healthcare) as described above. It was calculated from a plot of Kav (partition coefficient) against log molecular mass, which was calibrated using five protein standards: IgG (molecular mass, 150 kD), albumin (molecular mass, 55 kD), ovalbumin (molecular mass, 45 kD), chymotrypsinogen A (molecular mass, 20 kD), and ribonuclease A (molecular mass, 15 kD).
Phytase activity was estimated colorimetrically by monitoring the release of inorganic phosphate from phytic acid (Na-InsP6; Sigma-Aldrich). One unit of phytase activity was defined as the release of 1 µmol of phosphate per minute under the described conditions. Fifty-microliter samples were reacted at 37°C for 1 h in a 100 mM NaOAc (pH 4.5) assay buffer containing 1 mM Na-InsP6; the reaction was terminated by addition of an equal volume of 4% (w/v) TCA. The liberated inorganic orthophosphate was quantified spectrophotometrically with molybdenum blue (Murphy and Riley, 1962
pH Profile
Temperature and Thermal Stability Profiles
Substrate Specificity
Effect of Ions and Inhibitors on Enzymatic Activity
Enzyme Kinetics
To generate AtPAP15 promoter::GUS fusion constructs, AtPAP15 promoter sequences of various lengths (0.47, 1.1, and 1.6 kb) were first amplified by PCR using forward primers (F1, 5'-ATATTCTAGACATGCTCTGGTATATATTAAACTCC-3'; F2, 5'-ATATTCTAGACGATGCTACGTAGATGAAACG-3'; F3, 5'-ATATTCTAGAAGTG CCTAAACGAGTCCATTTA-3') and a reverse primer (PR, 5'-ATATCTCGAGCGTTCCAGAGGGTGGT-3'). The amplified fragments were cloned into the pGEM-T vector and sequenced. The AtPAP15 promoter fragments were released by XhoI and XbaI digestion and were subcloned into pBa002a-GUS to make transcriptional fusion sequences with the reporter gene GUS. Binary vectors containing the transgene inserts were mobilized into A. tumefaciens GV3101 by freeze-thaw transformation. Transformation of Arabidopsis was performed by the floral dip method (Clough and Bent, 1998
Eight to 15 T0 transgenic lines from each construct line were examined for GUS activity. All transgenic lines displayed identical patterns of GUS staining but with different intensities. Further analyses were performed on two homozygous T3 transgenic lines with strong GUS expression.
For the histological examination of tissues and the histochemical localization of GUS staining in seedlings and anthers, fixed plant samples of different developmental stages of pBa002a-AtPAP15 F2-GUS lines were dehydrated through a graded ethanol series and embedded in Historesin (Yeung, 1999
Homozygous pBa002a-AtPAP15 GUS lines (T3) were used as plant materials. For phytohormone treatment and for salt and osmotic stresses, seeds were germinated on MS agar plates for 7 d prior to treatment and incubated in liquid medium for another 12 to 48 h. The treatments were as follows: abscisic acid (100 µM), gibberellic acid (100 µM), methyl jasmonate (50 µM), salicylic acid (100 µM), hydrogen peroxide (4 mM), sodium chloride (250 mM), sorbitol (300 mM), and mannitol (300 mM). For nutrient starvation analysis, plants were first germinated on MS agar plates for 5 d, then transferred to MS medium lacking a specific nutrient (phosphorus, nitrogen, or potassium), and grown for another 2, 4, or 6 d. For all treatments, MS medium was used in parallel as a control. After harvest, plants seedlings were GUS stained.
T-DNA insertion lines were obtained from the Arabidopsis Biological Resource Center (Ohio State University). Seeds of two AtPAP15 insertion mutants, SALK_061597 (T2) and SAIL_529_D01 (T9), were germinated in the presence of kanamycin (50 mg L–1) or Basta (5 mg L–1) to follow the segregation of the antibiotic resistance marker. Antibiotic-resistant T2 plants were screened for T-DNA insertion by PCR using gene-specific primers and primers anchored in the T-DNA borders. Interruption of the AtPAP15 gene was further confirmed by RT-PCR and western-blot analysis. To generate the construct for complementation, an AtPAP15 construct, including the NOS promoter and the AtPAP15 cDNA, was subcloned into a pCAMBIA1300 plant expression vector. Empty pCAMBIA1300 vectors were employed as controls. The construct was transferred into A. tumefaciens strain GV3101, and T-DNA mutants were transformed using the floral dip method as described previously. T1 plants were selected using MS plates with hygromycin (20 mg L–1) and verified by PCR screening. Homozygous T3 lines were later selected by hygromycin resistance and verified by PCR and western-blot analysis.
T-DNA homozygous (T2 and T9), wild-type, and complementation lines were planted in soil for pollen collection. In vitro Arabidopsis pollen germination experiments were conducted as described previously (Fan et al., 2001
To measure phytase and APase activity during seed germination, around 100 seeds of different lines were surface sterilized with 20% Clorox and then sown in sterilized MQ water. Seeds were collected after 48 h, and protein was extracted using 100 mM sodium acetate buffer (pH 4.5) and 1 mM phenylmethylsulfonyl fluoride. Internal substrates, phosphate, and salts were removed from the protein fraction by Microcon (molecular weight cutoff 10,000; Millipore). Phytase and APase activity were detected using 1 mM Na-phytate and pNPP as substrates, respectively. For pollen, different lines were planted in soil for pollen collection. The in vitro Arabidopsis pollen germination experiment was carried out as described above. After 8 h of germination, the pollen was collected by centrifugation and the protein was extracted for enzyme assays as described above. Experiments were repeated twice, and at least four replicates were carried out.
All data were analyzed by one-way ANOVA using the LSD at the level of 5% (P < 0.05) to identify the significant differences between the observations, with the aid of the statistical program SPSS 10.0. For in silico analysis, homology searches in GenBank were done using the BLAST server (http://www.ncbi.nlm.nih.gov/BLAST/). Multiple alignments of protein sequences were performed using the ClustalX and N-J plot programs. For prediction of protein expression, Genevestigator was employed (http://www.genevestigator. ethz.ch). Sequence data from this article can be found in the GenBank/EMBL data libraries under accession number At3g07130.
The following materials are available in the online version of this article.
We thank Dr. W.K. Yip's laboratory at the University of Hong Kong for kindly providing plant vectors and technical support for plant cultures. We also thank Dr. Clive Lo at the University of Hong Kong for his support with photomicroscopy. Received June 18, 2009; accepted July 20, 2009; published July 24, 2009.
1 This work was supported by the University Research Committee (grant no. 10206029) and by a Discovery Grant from the Natural Sciences and Engineering Research Council of Canada to E.Y. 
2 These authors contributed equally to the article. 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: Boon Leong Lim (bllim{at}hku.hk).
[W] The online version of this article contains Web-only data.
[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.109.143180 * Corresponding author; e-mail bllim{at}hku.hk.
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