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First published online December 7, 2007; 10.1104/pp.107.110809 Plant Physiology 146:418-430 (2008) © 2008 American Society of Plant Biologists
Identification, Biochemical Characterization, and Subcellular Localization of Allantoate Amidohydrolases from Arabidopsis and Soybean1,[W]Freie Universität Berlin, Institut für Biologie, Abteilung Biochemie der Pflanzen, 14195 Berlin, Germany (A.K.W., T.R., C.-P.W.); and Oxford Brookes University, School of Life Sciences, Oxford OX3 0BP, United Kingdom (I.A.S.)
Allantoate amidohydrolases (AAHs) hydrolize the ureide allantoate to ureidoglycolate, CO2, and two molecules of ammonium. Allantoate degradation is required to recycle purine-ring nitrogen in all plants. Tropical legumes additionally transport fixed nitrogen via allantoin and allantoate into the shoot, where it serves as a general nitrogen source. AAHs from Arabidopsis (Arabidopsis thaliana; AtAAH) and from soybean (Glycine max; GmAAH) were cloned, expressed in planta as StrepII-tagged variants, and highly purified from leaf extracts. Both proteins form homodimers and release 2 mol ammonium/mol allantoate. Therefore, they can truly be classified as AAHs. The kinetic constants determined and the half-maximal activation by 2 to 3 µM manganese are consistent with allantoate being the in vivo substrate of manganese-loaded AAHs. The enzymes were strongly inhibited by micromolar concentrations of fluoride as well as by borate, and by millimolar concentrations of L-asparagine and L-aspartate but not D-asparagine. L-Asparagine likely functions as competitive inhibitor. An Ataah T-DNA mutant, unable to grow on allantoin as sole nitrogen source, is rescued by the expression of StrepII-tagged variants of AtAAH and GmAAH, demonstrating that both proteins are functional in vivo. Similarly, an allantoinase (aln) mutant is rescued by a tagged AtAln variant. Fluorescent fusion proteins of allantoinase and both AAHs localize to the endoplasmic reticulum after transient expression and in transgenic plants. These findings demonstrate that after the generation of allantoin in the peroxisome, plant purine degradation continues in the endoplasmic reticulum.
Plant growth is often limited by nitrogen availability in the soil. While plants depend on efficient nitrogen uptake, they also require effective means to internally redistribute nitrogen during every stage of development (Rentsch et al., 2007  ). Most prominent are redistribution processes during germination or during senescence (Masclaux et al., 2000) when the great majority of nitrogen stored in the leaves is exported to sustain flowering and seed or storage organ development. However, "understanding the mechanisms of nitrogen remobilisation during leaf senescence is still at a preliminary stage" (p. 603) but will be required to increase nitrogen use efficiency and crop yields (Good et al., 2004).
Although proteins, mainly Rubisco, account for most of the reduced nitrogen in leaves, 8% to 19% of reduced nitrogen may be present as nucleic acids (data from wheat leaves; Peoples and Dalling, 1988
Purine catabolism can be divided into three phases. First, xanthine is generated from the purine nucleotides by deamination and oxidation of the nucleobases and by releasing the phosphate and Rib moieties (Zrenner et al., 2006
In many warm-climate legumes like French bean (Phaseolus vulgaris) or soybean, the reactions releasing ammonium from allantoate are not only required for nitrogen recycling from nucleic acids but also are of central importance to the plants' nitrogen supply under nitrogen-fixing conditions. Most of the symbiotically fixed nitrogen is used for de novo purine synthesis in the nodules. The purines are then degraded to the ureides allantoin and allantoate, which serve as nitrogen carriers and are exported to the xylem (Schubert, 1986
AAH activity in soybean extracts was activated by the addition of manganese, whereas the presence of manganese, calcium, iron, cobalt, and nickel did not activate the enzyme. The addition of metal chelators like EDTA and acetohydroxamate abolished the activity (Winkler et al., 1985
The subcellular localization of the enzymes participating in plant purine degradation has been almost exclusively studied in nodules (Schubert, 1986 In this article, we characterized AAHs from legume and non-legume plants biochemically using proteins purified to homogeneity from a plant source. The enzymatic activities releasing ammonium, the activation by several metal ions, and the direct inhibition by borate, fluoride, L-Asn, L-Asp, and a range of other substances were assessed. Our data shows that previous hypotheses regarding the mechanism of borate and L-Asn inhibition need to be rejected. The functionality of the AAHs and of Arabidopsis allantoinase (AtAln) was demonstrated in vivo by generating transgenic lines expressing tagged proteins in the corresponding mutants. Finally, we investigated the subcellular localization of the AAHs and AtAln. From this, we concluded that after the generation of allantoin in the peroxisome, plant purine degradation continues in the ER.
Cloning, Transient Expression, and Purification Database searches using the AAH sequences of Escherichia coli (allC, P77425) and Bacillus subtilis (allC, O32149) as query against the protein dataset of the Arabidopsis Genome Initiative identified two similar Arabidopsis AAH candidate proteins (Supplemental Fig. S1), tentatively called AtAAH (locus At4g20070) and AtAAH-2 (locus At5g43600). Amino acid identities to the enzyme from E. coli are approximately 26% to 27% (similarities 39%–43%) for AtAAH and AtAAH-2, respectively. A full-length reading frame for a soybean AAH candidate protein (GmAAH) was also identified by assembling five EST sequences. The predicted protein was 29% identical (43% similar) to the E. coli protein.
The crystal structure of the E. coli enzyme has recently been resolved (Agarwal et al., 2007
Coding sequences without a stop codon for the plant AAH candidate proteins were amplified by reverse transcription (RT)-PCR from ecotype Columbia for Arabidopsis and the cultivar ‘Williams 82’ for soybean. The products were cloned into the binary vectors pXCS-haemagglutinin-StrepII (HAStrep) and pXCS-yellow fluorescent protein (YFP; Witte et al., 2004
Agrobacteria carrying these binary vectors were injected into tobacco plants for transient expression. Nicotiana benthamiana was used for the transient expression of HAStrep-tagged proteins, while Nicotiana tabacum served for the transient expression of YFP-tagged proteins. Three to 4 d after infiltration, the Strep-tagged proteins were purified by StrepTactin affinity chromatography (Witte et al., 2004
Enzymatic Activity
In an enzymatic assay using allantoate as substrate, a linear increase of ammonium concentration with time was observed for reactions containing AtAAH and GmAAH but not for the reaction with AtAAH-2 (Fig. 2A
). We concluded that the protein encoded on locus At4g20070 is the Arabidopsis AAH and called it AtAAH in accordance with Todd and Polacco (2006)
The AtAAH reaction product, probably ureidoglycolate, is unstable under the assay conditions and decays to glyoxylate and urea, which was monitored by the inclusion of urease in the assay. Ureidoglycolate instability at different pH values and in the presence of divalent cations is well documented in the literature (Gravenmade et al., 1970 ). Nonenzymatic allantoate hydrolysis releasing either ammonium (Fig. 2A, AtAAH-2 curve) or urea (Fig. 2B, urease curve) is not observed under the employed assay conditions. To maintain activity, the presence of dithiothreitol (DTT) was required at all times. Under reducing conditions, the activity of purified AtAAH was stable for a minimum of 2 h, whereas GmAAH was more labile and slowly lost activity during storage on ice. Both purified proteins could be frozen in liquid nitrogen and stored in the freezer without activity loss. Activity assays for GmAAH were therefore always carried out with freshly thawed enzyme.
Using short 2-min assays, the initial activity at varying allantoate concentrations was measured. These assays required the optimization of the ammonium detection sensitivity (see "Materials and Methods"). The activity of both enzymes could be well described by Michaelis-Menten kinetics (Fig. 3, A and B , s/v against s graph). By fitting the data of the v against the s hyperbola, KM values of 30.1 ± 7.9 µM and 80.9 ± 12.3 µM (confidence interval [ci]; P = 95%) were determined for AtAAH and GmAAH, respectively. Specific activities of 30.7 units mg–1 and 58.1 units mg–1 corresponding to turnover numbers (kcat) of 27.2 s–1 and 50.2 s–1 were determined for AtAAH and GmAAH, respectively, using the data shown in Figure 3, A and B. In some extractions, up to 8-fold higher specific activities were found, but the basis for this variation could not be determined. The presence of Triton X-100 in the purified enzyme solution had a stimulating effect on enzyme activity (tested for AtAAH). A 10-fold reduction of the routinely used Triton X-100 concentration of 0.005% resulted reproducibly in >80% activity loss after a 30-min incubation on ice. It was not possible to recover the original activity by re-adding Triton X-100, even after incubation for several hours. Maximum activity was obtained by purifying the enzyme in the presence of 0.005 to 0.01% Triton X-100.
AAHs from bacterial and plant sources are activated by manganese (Vogels, 1966 ; Winkler et al., 1985). For AtAAH and GmAAH, manganese concentrations of 2.1 ± 0.6 µM and 2.9 ± 1.1 µM (ci, P = 95%) were required for one-half maximal activation, respectively, while approximately 50 µM was sufficient for full activation (Fig. 3C). The crystal structure of AAH from E. coli contained two metal ions per active center that were tentatively identified as zinc (Agarwal et al., 2007). We have used zinc, cobalt, and nickel in activation tests. In comparison to the activity with manganese, which was set to 100%, we found only marginal activity with zinc but up to 20% activity with cobalt or nickel in the concentration range of 5 to 50 µM. Higher concentrations inhibited the enzyme. In general, plants contain on average 100 µg manganese, 44 µg zinc, and 2 µg nickel per kilogram dry weight (dw; Epstein, 1999). Because manganese in plants is more abundant than nickel and cobalt, it appears likely that plant AAHs are activated by manganese in vivo.
The inhibition of allantoate-degrading activity by L-Asn and borate in soybean extracts has been reported by Lukaszewski et al. (1992) Purified GmAAH (Fig. 4A ) and AtAAH (data not shown) were inhibited by L-Asn and L-Asp but barely by D-Asn, demonstrating that a chelating effect cannot be responsible for L-Asn-mediated inhibition. L-Gln had no effect on activity. Structurally, L-Asn resembles allantoate or ureidoglycolate and may rather function as a competitive inhibitor. The inhibitory effect of L-Asn can be suppressed by increasing substrate concentrations (Fig. 4A), consistent with a competitive or mixed mechanism of inhibition. None of the tested amino acids were substrates for the enzymes.
Borate also inhibited AtAAH and GmAAH, but not by chelating manganese. At a limiting (50 µM) or abundant (750 µM) manganese concentration, GmAAH was inhibited to the same extent by borate concentrations from 0.5 to 8.0 mM (Fig. 4B). The same was observed for AtAAH (data not shown). Using a substrate concentration of 500 µM allantoate, 233 µM (ci, 194–279 µM; P = 95%) and 708 µM (ci, 611–820 µM; P = 95%) borate inhibited 50% of AtAAH and GmAAH activity, respectively (Fig. 4C).
Fluoride at a concentration of 1 mM was reported to inhibit the activity of AAH from B. fastidiosus by 20% (Xu et al., 1995
For a range of substances tested at 8 mM concentration in the presence of 500 µM allantoate, no inhibitory effect on AtAAH activity was observed. These included β-mercaptoethanol, allantoin, urea, N
We assessed the substrate specificity of the AAHs using several substances with similarity to allantoate at 8 mM concentration: N
Monomeric masses of 53.1 and 51.9 kD for AtAAH and GmAAH, respectively, were calculated based on the sequence of the tagged proteins with the predicted N-terminal signal anchor cleaved off (prediction by the SignalP 3.0 Web server; cleaved signal anchor underlined in Supplemental Fig. S1). To estimate the molecular mass of the native proteins, native gel electrophoresis with eight gels of different acrylamide concentrations was resolved. Ferguson plots were generated for eight molecular mass standards and AtAAH and GmAAH. In a secondary plot, the log (–slope) determined from the Ferguson analysis was plotted against the log (molecular mass) of the standards (Bryan, 1977
Using gel filtration analysis, native molecular masses of 196 kD for AtAAH and 143 kD for GmAAH were determined, respectively (Supplemental Fig. S2, B and C). The enzymes may either be able to form trimers/tetramers or have an atypical hydrodynamic radius for their molecular mass in this type of analysis. In light of the structural data from the E. coli enzyme, a trimeric structure appears very unlikely. A tailing effect of the elution peak observed during gel filtration of GmAAH (Supplemental Fig. S2C) may be caused by partial dissociation into monomers consistent with the lower stability of this enzyme.
A homozygous Ataah mutant (SALK 112631) was isolated from the mutant collection of the Salk Institute Genomic Analysis Laboratory (Alonso et al., 2003 The Ataah and Ataln mutants were unable to grow with 10 mM allantoin as the sole nitrogen source (Fig. 6A, 2 and 5 ), while the wild type grew and finally flowered (Fig. 6A, 1). The phenotype of the Ataah mutant was more severe than that of the Ataln mutant, which still showed some root elongation with a few plants reaching the four-leaf stage. Perhaps another enzyme in Arabidopsis can partially substitute for the lacking allantoinase under these growth conditions. With nitrate as nitrogen source, growth of the mutants and the wild type was identical (data not shown). The phenotype of the Ataah mutant could be complemented in transgenic plants expressing either AtAAH-HAStrep or GmAAH-HAStrep (Fig. 6A, 3 and 4). Equally, the defect in Ataln could be complemented in transgenic plants expressing AtAln-HAStrep (Fig. 6A, 6). Also, the expression of the corresponding YFP fusions rescued the mutants (data not shown). These results demonstrate that the T-DNA insertions in AtAAH or AtAln cause the growth defect on allantoin and that C terminally tagged versions of the corresponding proteins are biologically functional.
The allantoate content of Ataah mutant seedlings grown for 2 weeks on plates with nitrate as the nitrogen source was severalfold higher than in the wild type (Fig. 6B; Todd and Polacco, 2006 ). In two independent homozygous complementation lines expressing tagged AtAAH, allantoate amounts were similar to those found in the wild type, confirming the successful complementation. In two segregating complementation lines expressing tagged GmAAH, the allantoate concentrations were partially reduced compared to the concentrations found in the mutants. That only partial reduction was observed is probably due to the presence of mutant plants not carrying a transgene in the segregating population.
Plants were grown for 9 d on Murashige and Skoog medium with 9.35 mM KNO3 and were subsequently placed on medium containing: (1) KNO3; (2) an equimolar amount of KCl and no KNO3; or (3) KCl and no KNO3 but 10 mM of allantoin. After 2 and 5 d, plants were harvested for RNA extraction (five individuals per sample). Semiquantitative RT-PCR revealed that AtAln mRNA amounts increased due to nitrogen stress but were not further influenced by the presence of allantoin, whereas AtAAH mRNA levels did not significantly change in this experiment (Fig. 7 ).
Subcellular Localization of AtAAH, GmAAH, and AtAln
Currently, it is thought that the enzymatic reactions leading to allantoin take place in the peroxisome (summarized in Todd et al., 2006 Upon transient expression in tobacco, AtAln, AtAAH, and GmAAH localized to the ER in almost all cells (Fig. 8, A–I ), but were very occasionally also found in the Golgi or the peroxisomes (Supplemental Figs. S4 and S5). An N terminally truncated version of AtAAH-YFP lacking the first 71 amino acids exclusively localized to the cytosol (Fig. 8J), confirming that the N terminus is required for localization. A fraction of the full-length proteins was also present in the cytosol, maybe due to partial instability of the fusions. Western-blot analysis showed that the fluorescent fusion proteins were stable, although low amounts of free YFP were observed (data not shown).
Several seedlings from at least two independently transformed stable transgenic lines of AtAln and AtAAH fluorescent fusions were assessed. AtAln was found exclusively in the ER in shoot and root (Fig. 8L). AtAAH was also found in the ER (Fig. 8N) but was not uniformly distributed, localizing to possible substructures within the ER. Small punctate motile structures reminiscent of Golgi or peroxisomes were also occasionally observed.
The final steps of purine degradation have long been the focus of research, especially because in tropical legumes, these reactions are central to nitrogen supply under nitrogen-fixing conditions. However, it has been difficult to isolate the enzymes involved. Native AAHs have never been purified to homogeneity from a plant source, potentially due to the instability of enzymatic activities (Lukaszewski et al., 1992 ). Recently, Raso et al. (2007) reported that attempts to isolate allantoate-degrading activity from developing French bean fruits using several conventional purification techniques led to complete loss of activity. We show that C terminally StrepII-tagged enzymes can be successfully isolated from a plant source. The high purity obtained (Fig. 1) and the short processing time of <1 h illustrate the potential of StrepII tag-based purification for plant biochemistry. Earlier problems with insufficient binding to the StrepTactin matrix and problematic elution under certain conditions (Witte et al., 2004) were resolved here. Despite the rapid purification, the instability of the AAH enzymes was also noted in this work. A possible explanation for the instability may be the decay of the enzymes into monomers, as indicated by results from electrophoresis and gel filtration for the less-stable GmAAH (Supplemental Fig. S2).
Expression of AtAAH was shown to complement a yeast mutant defective in allantoate and urea degradation in a growth assay on allantoin as the nitrogen source (Todd and Polacco, 2006
The leaf allantoate content is several times lower in Arabidopsis than in soybean. In Arabidopsis, approximately 34 µmol kg–1 fresh weight (fw) was measured in this study (Fig. 6B), and approximately 400 µmol kg–1 fw was measured by Todd and Polacco (2006)
In soybean plants grown without the addition of manganese, a leaf manganese concentration of 11.9 mg kg–1 dw was measured (Vadez et al., 2000b
L-Asn inhibits AtAAH and GmAAH, likely by a competitive mechanism. The structural similarity of Asn to allantoate or ureidoglycolate also suggests that it may bind to the active site. The L-Asn concentration increases in leaf tissues of soybean during drought stress. King and Purcell (2005)
Borate as well as fluoride inhibit AtAAH and GmAAH (Fig. 4). No indication was found that borate functions by chelating manganese as previously postulated (Lukaszewski et al., 1992
Plant AAHs are multimers. A dimeric structure was found by an electrophoretic method in accordance with the dimeric structure of the E. coli enzyme (Agarwal et al., 2007
Arabidopsis mutants carrying T-DNA insertions in AtAAH or AtAln do not grow on allantoin as sole nitrogen source. This defect can be phenotypically (Fig. 6A) and molecularly (Fig. 6B) rescued by expressing C terminally tagged versions of the corresponding proteins. These findings show that AtAln functions as allantoinase, and AtAAH and GmAAH function as AAHs in vivo. The crystal structure of the E. coli enzyme (Agarwal et al., 2007
Nitrogen starvation induces genes involved in amino acid and nucleotide breakdown, while upon re-addition of nitrate these genes are repressed. However, the changes in gene expression are generally small (Scheible et al., 2004
Localization studies for enzymes involved in purine degradation were almost exclusively performed using nodules of legumes. It appears that the enzymes involved in the generation of allantoin locate to the peroxisomes (summarized in Todd et al., 2006
Plant Material and Growth Conditions
Mutants of Arabidopsis (Arabidopsis thaliana) from the SALK collection (Alonso et al., 2003 Agar plates were prepared with half-strength Murashige and Skoog nutrients without vitamins, sugars, or amino acids. NH4NO3 was omitted and CoCl2 was replaced by 0.5 µM NiCl2. Plates contained either 9.35 mM KNO3 or 10 mM allantoin as sole nitrogen source. To allantoin plates, 9.35 mM KCl was added to maintain the same molarity of potassium ions as on the nitrate plates. Plates were incubated for up to 6 weeks in a controlled growth room in long-day conditions (16 h light of 150 µmol m–2 s–1, 20°C day, 18°C night, 60% relative humidity).
RNA from plant leaves was prepared using TRI reagent (Sigma) and treated with DNaseI (Sigma) following the manufacturer's instructions. RT using 1.0 µg total RNA was performed with Moloney murine leukemia virus reverse transcriptase (Invitrogen) and a poly-T primer. AtAln cDNA was amplified by PCR using primers p1191: gaattcaaaATGGAGAGAACTTTGCTTC and p1192: cccgggAGTAGTTGCAAGTTGCAGAG introducing an EcoRI and XmaI site and a plant ribosome binding site consensus (all lowercase). The product was cloned via these restriction sites into modified versions of pXCS-HAStrep (AY457636; Witte et al., 2004 For semiquantitative RT-PCR, 400 ng of seedling RNA was reverse transcribed in a 20-µL reaction (as described above). Five microliters of 10-fold diluted cDNA was used as template in 20-µL PCR reactions. For the amplification of AtAln cDNA, the primers p1199: GACTTAGAGATTGAAGATGGTAGTGA and p1192 (see above) were used (53°C annealing temperature; 28 cycles). AtAAH cDNA was amplified with primers p1190: GCAGGACAGACAAGACTCAAG and p1176: CGTAAATCCACAGTGAAAGTTAC (53°C annealing temperature; 35 cycles), and Actin2 (At3g18780) cDNA was amplified with primers p1034: GAGAGGTTACATGTTCACCACAAC and p1033: GTGAACGATTCCTGGACCTGCCTC (53°C annealing temperature; 28 cycles). All primer sets would yield products of increased size with genomic DNA due to the presence of introns, but only amplification from cDNA was observed.
Mutants from the SALK and the SAIL collections were screened with a primer binding at the left border of the T-DNA insert (p488: TGGTTCACGTAGTGGGCCATC for SALK and p1197: TTTTCAGAAATGGATAAATAGCCTTGCTTC for SAIL) together with gene-specific primers. As gene-specific primers p1198: GAAGAGATCAAGAGAGAGATATGGAG and p1193: GATGTTTGTCATTGGAAAATC were used for AtAln, and p1190 and p1173 (see above) were used for AtAAH. The PCR products from the mutants were cloned and sequenced to map the exact position of the insertions. To determine the level of gene-specific mRNA in the mutants, cDNA from seedlings was prepared as described above. For amplification of AtAAH cDNA, the primers p1190 and p1173 were used (wild-type product size 678 bp; 53°C annealing temperature; 35 cycles). In the mutant, the T-DNA insertion is flanked by these primers. For amplification of AtAln cDNA, the primers p1198 and p1193 flanking the insertion were used (wild-type product size, 653 bp), and additionally the primers p1199 and p1192 not flanking the insertion were used (wild-type product size, 816 bp). The PCR was carried out at 53°C annealing temperature with 35 cycles. Control PCRs for Actin2 were performed as described above.
Transgenic plants were generated as described by Witte et al. (2004)
Transient expression in N. benthamiana was performed as described by Witte et al. (2004) For enzyme purification, 0.75 g of fresh leaf material was ground in 1.5 mL of extraction buffer consisting of 100 mM HEPES, pH 8.0, 100 mM NaCl, 5 mM EDTA, pH 8.0, 15 mM DTT, 100 µg mL–1 avidin, and 0.5% (v/v) Triton X-100. The extract was centrifuged at 21,000g, 15 min, 4°C, and 40 µL of StrepTactin Macroprep (50% suspension; IBA) was added to the supernatant. After incubation for 10 min (4°C) in a rotation wheel, the affinity matrix was pelleted by centrifugation for 30 s at 700g and the supernatant discarded. The matrix was washed five times with 500 µL of wash buffer (100 mM HEPES, pH 8.0, 100 mM NaCl, 0.5 mM EDTA, 2 mM DTT, 0.005% Triton X-100), discarding the supernatant after each wash. The proteins were eluted by adding 75 µL of elution buffer (wash buffer containing 10 mM biotin) and placing the vials for 5 min in an Eppendorf thermomixer (1,000 rpm) at room temperature. The elution was repeated with further 75 µL of elution buffer and the two elutions were pooled.
For the standard enzymatic assay, 50 µL of allantoate solution (routinely 6 mM in 10 mM HEPES, pH adjusted to 8.0) and 45 µL of wash buffer (see above) containing 2 mM MnCl2 were incubated separately for 5 min at 30°C. Two minutes before the start of the reaction, 5 µL of enzyme solution was added to the wash buffer. The enzyme solution was prepared by diluting purified enzyme in wash buffer such that a linear rate for the corresponding assay was obtained. Due to the variation of specific activities, dilutions needed to be adjusted for each purification batch. Dilutions of 2- to 20-fold were routinely employed corresponding to approximately 16 to 170 ng of protein per assay. The reaction was started by the addition of the preheated allantoate. In a time course, samples of 20 µL were taken and added to 200 µL of water, followed by 50 µL of phenol nitroprusside reagent and 100 µL of hypochloride reagent prepared as described by Witte and Medina-Escobar (2001) The determination of kinetic constants required short assay times and assays with low substrate concentrations. To increase the detection sensitivity, assays were carried out in 200-µL reactions to which 50 µL of phenol nitroprusside reagent and 100 µL of hypochloride reagent were added directly. Ten-fold diluted wash buffer was used in these reactions, but the Triton X-100 concentration in this diluted buffer was maintained at 0.005% and the DTT concentration was adjusted to 1 mM. The enzyme preparations were diluted such that at the lowest substrate concentration, a linear rate could still be measured within 2 min. Substrate concentrations of 37.5, 75, 150, 300, 600, and 1,200 µM were employed and measured in eight repeats each.
Inhibitors were generally included into the allantoate solution used to start the enzymatic reactions, with the exception of fluoride, which was present at the indicated concentrations in wash buffer and allantoate solution. As a result, the enzyme was already in contact with fluoride 2 min before the addition of allantoate (during the preincubation phase). This procedure was chosen because fluoride was reported to be a slow binding inhibitor for urease (Todd and Hausinger, 2000 Purified protein was quantified using the NanoOrange kit (N6666) from Invitrogen. Twenty microliters of eluted protein was mixed with 300 µL of working solution (in three replicates) and processed according to the manufacturer's instructions. Standards were constructed with bovine serum albumin solutions in elution buffer.
Seedlings were grown on half-strength Murashige and Skoog plates containing KNO3 as nitrogen source under long-day conditions. After 15 d, three samples of 0.1 g fw were taken from the seedlings of each plant type and extracted with 1 mL of 50 mM K/Na phosphate buffer, pH 7.0. Insoluble material was removed by centrifugation, and allantoin and allantoate concentrations were determined by differential glyoxylate analysis (Vogels and Van der Drift, 1970
SDS-gel electrophoresis, western blots for the detection of StrepII-tagged proteins, and Coomassie staining were performed according to Witte et al. (2004)
Statistical analyses were performed with the GraphPad Prism software package (www.graphpad.com). Michaelis-Menten parameters were determined by using the Michaelis-Menten equation to fit the enzymatic velocity against substrate concentration curve. Manganese activation data were fitted to a hyperbola: v = vmax ([Mn]/(Kd + [Mn])). Borate and fluoride inhibition data were fitted with a variable slope sigmoid equation (Hill equation) without constrains. The multiple alignment was generated with VectorNTI (Invitrogen) and shaded with the Boxshade program (www.ch.embnet.org/). Global pairwise alignments were performed with Needle from the EMBOSS open software suite. Subcellular targeting was predicted with the MultiLoc Web server (www-bs.informatik.uni-tuebingen.de/Services/MultiLoc/index_html). Signal anchor sequences were predicted with the SignalP 3.0 Web server (www.cbs.dtu.dk/services/SignalP). Gel filtration data was analyzed with the Unicorn software package (GE Healthcare).
Nicotiana tabacum was grown and epidermal cells infiltrated according to Sparkes et al. (2006) The GmAAH cDNA sequence was deposited in GenBank under the accession number AM773229.
The following materials are available in the online version of this article.
We thank Dr. Chris Weise from the Institute for Chemistry and Biochemistry of the FU Berlin for mass spectrometry analysis, and Renate Grübnau and Janet Evins for technical assistance. Received October 19, 2007; accepted November 26, 2007; published December 7, 2007.
1 This work was supported by the Alexander von Humboldt Foundation (to A.K.W., T.R., and C.P.W.), and by the Biotechnology and Biological Sciences Research Council (to I.A.S.).  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: Claus-Peter Witte (cpwitte{at}zedat.fu-berlin.de).
[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.107.110809 * Corresponding author; e-mail cpwitte{at}zedat.fu-berlin.de.
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-acetyl-L-Asn and N
