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Originally published In Press as doi:10.1074/jbc.M312929200 on January 15, 2004

J. Biol. Chem., Vol. 279, Issue 14, 13547-13554, April 2, 2004
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Tandem Orientation of Duplicated Xanthine Dehydrogenase Genes from Arabidopsis thaliana

DIFFERENTIAL GENE EXPRESSION AND ENZYME ACTIVITIES*

Christine Hesberg, Robert Hänsch, Ralf R. Mendel{ddagger}, and Florian Bittner

From the Department of Plant Biology, Technical University of Braunschweig, 38023 Braunschweig, Germany

Received for publication, November 26, 2003 , and in revised form, January 13, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Xanthine dehydrogenase from the plant Arabidopsis thaliana was analyzed on molecular and biochemical levels. Whereas most other organisms appear to own only one gene for xanthine dehydrogenase A. thaliana possesses two genes in tandem orientation spaced by 704 base pairs. The cDNAs as well as the proteins AtXDH1 and AtXDH2 share an overall identity of 93% and show high homologies to xanthine dehydrogenases from other organisms. Whereas AtXDH2 mRNA is expressed constitutively, alterations of AtXDH1 transcript levels were observed at various stresses like drought, salinity, cold, and natural senescence, but also after abscisic acid treatment. Transcript alteration did not mandatorily result in changes of xanthine dehydrogenase activities. Whereas salt treatment had no effect on xanthine dehydrogenase activities, cold stress caused a decrease, but desiccation and senescence caused a strong increase of activities in leaves. Because AtXDH1 presumably is the more important isoenzyme in A. thaliana it was expressed in Pichia pastoris, purified, and used for biochemical studies. AtXDH1 protein is a homodimer of about 300 kDa consisting of identical subunits of 150 kDa. Like xanthine dehydrogenases from other organisms AtXDH1 uses hypoxanthine and xanthine as main substrates and is strongly inhibited by allopurinol. AtXDH1 could be activated by the purified molybdenum cofactor sulfurase ABA3 that converts inactive desulfo-into active sulfoenzymes. Finally it was found that AtXDH1 is a strict dehydrogenase and not an oxidase, but is able to produce superoxide radicals indicating that besides purine catabolism it might also be involved in response to various stresses that require reactive oxygen species.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Xanthine oxidoreductase (XOR)1 is a ubiquitous metalloflavo enzyme with a central role in purine catabolism where it catalyzes the oxidation of hypoxanthine to xanthine and xanthine to uric acid. The enzyme from higher eukaryotes is active as a homodimer composed of two identical subunits of 150 kDa, each being subdivided into three domains: a N-terminal domain of 20 kDa for binding of two iron-sulfur clusters of the [2Fe-2S] type, a 40-kDa domain harboring a FAD-binding site, and a C-terminal molybdenum cofactor (Moco)-binding domain of 85 kDa. XOR enzymes in mammals are present either as the predominantly existing xanthine dehydrogenase (XDH; EC 1.1.1.204 [EC] ) or as O2-dependent xanthine oxidase (XO; EC 1.1.3.22 [EC] ). Whereas XDH possesses high reactivity toward NAD+ and low reactivity toward O2 as electron acceptor, XO reacts in a strictly O2-dependent manner with negligible reactivity toward NAD+. Both forms can be interconverted reversibly by oxidation of cysteine residues (1), whereas the conversion of XDH into XO by limited proteolysis is irreversible (2). In contrast to mammalian XOR the avian enzyme is exclusively present in the dehydrogenase form (3). The ability of mammalian XO to produce superoxide and hydrogen peroxide by reducing molecular oxygen (4) led to the suggestion that it might play an important role in the pathogenesis of cellular injury (5, 6). Among all XOR enzymes studied so far the bovine enzyme from milk is most exhaustively studied and recently its crystal structure has been determined for both the XDH and the XO form (7).

Besides functional characterization of XDH/XO, the corresponding nucleotide and protein sequence information was published for organisms like humans (8), cow (9), rat (2), mouse (10), chicken (3), insects (11, 12), fungi (13), and bacteria (14, 15) also. With the exception of the silkworm all organisms analyzed so far possess one XOR gene.

In plants the XDH but not the oxidase form was purified from nodules of bean (16), from the green algae Chlamydomonas reinhardtii (17), from wheat leaves (18) and leaves of legumes (19), as well as from pea seedlings (20). All plant XDH proteins were found to be homodimers with a molecular mass of ~300 kDa and showed highest substrate specificity for hypoxanthine and xanthine but were also able to convert purines, pterines, and aldehydes at a much lower rate. Beside purine degradation, plant XDH is supposed to play a role in important cellular processes: (i) plant-pathogen interactions between phytopathogenic fungi, legumes, and cereals (21, 22); (ii) cell death associated with hypersensitive response (23, 24); and (iii) natural senescence (25). As all these processes require the formation of reactive oxygen species XDH was supposed to be able to produce superoxide anions and/or hydrogen peroxide (25). Supporting this hypothesis, XDH activity was found to be increased concomitant with superoxide dismutase and other oxygen-related enzymes in senescent pea leaves (25). Although much effort was spent at purification and biochemical characterization of plant XDH neither cloning of the corresponding cDNAs nor molecular data are published so far.

In this work we describe the cloning of two XDH cDNAs from Arabidopsis thaliana, their tandem arrangement in the genome, their mRNA expression levels as well as the enzymatic activities at various stresses and treatments, and the recombinant expression of AtXDH1 cDNA in the methylotrophic yeast Pichia pastoris with subsequent purification and characterization of the AtXDH1 protein.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Plant Material and Plant Growth—A. thaliana Col-O, Ler, and aba3.2 seeds were grown in pots containing low-nutrient soil in an AR-36L A. thaliana growth chamber (Percival Scientific, Perry, IA) at 13 h light/11 h darkness, at 21 °C, and 70% relative humidity for periods as given in the text.

Stress Treatment—For drought stress experiments, soil was completely removed from the roots prior to incubation under normal conditions in the chamber for 4 h or as given in the text (loss of fresh weight about 50%). Subsequently, roots and leaves were detached and used for RNA and activity analysis. ABA treatment in the case of plants without drought treatment was performed by spraying plants with 50 µM (±)-cis,trans-ABA in water uniformly onto the leaves for 4 h. In the case of combined ABA and drought treatment plants were sprayed with ABA prior to removal of plants from the soil and 2 h after removal. Treatment also lasted 4 h and control plants were sprayed with water instead of ABA solution. For NaCl treatment, 3-week-old plants were transferred to hydroponic culture 2 days before treatment and then incubated in nutrition solution containing 200 mM NaCl for 6 and 20 h. Cold stress treatment at 4 °C was performed in a chamber with ambient temperature for 6 and 20 h and freezing stress was applied by incubating plants in small chambers precooled to -4 °C. Because longer freezing stress will result in freezing of the soil freezing temperature treatment lasted only 6 h.

Preparation of RNA—Total RNA was prepared either as described by Ref. 26 or by using the NucleoSpin RNA Plant kit (Macherey & Nagel, Dueren, Germany) according to the manufacturers instructions.

Relative Quantitative Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR)—For each RT reaction 2 µgof A. thaliana total RNA was reverse-transcribed with avian myeloblastosis virus-reverse transcriptase (Promega, Madison, WI) and oligo-d(T)18-BamHI primer according to standard procedures (27). RT-PCR was performed on a PCR Express gradient cycler (Hybaid, Heidelberg, Germany) by using the SAWADY Taq DNA polymerase (Peqlab, Erlangen, Germany). AtXDH1-specific primers were AtXDH1+, 5'-CACATTTACTGAGCTAGTA-3', and AtXDH1-,5'-GTTTCCCCTCTGATGATGTTC-3'; AtXDH2-specific primers were AtXDH2+, 5'-TCTTCTCAAGGGTAATCCA-3', and AtXDH2-, 5'-TTCTCCCCTCTATTAAAGTTT-3'. The following PCR program was used: 3 min at 94 °C for initial denaturing of templates, 30 cycles including denaturing for 30 s at 94 °C, annealing for 1 min at 56 °C and elongation for 1 min at 72 °C, and a final elongation step for 6 min at 72 °C. RT-PCR generated fragments were directly ligated to pGEM-T Easy (Promega) and sequenced for ascertaining proper amplification.

Cloning of AtXDH2 cDNA—Two overlapping cDNA subfragments were generated by PCR from reverse transcribed total RNA of A. thaliana (Col-O) and subsequently fused by PCR according to standard procedures (27). The obtained full-length cDNA of AtXDH2 was directly ligated to pGEM-T Easy (Promega) and sequenced. Specific PCR primers for generation of the 5' subfragment (2080 base pairs) were AtXDH2-ATG, 5'-GTTCAGTGAAGATGGAGCAGAAC-3', and AtXDH2–2622rev, 5'-GCGACAAGCACACCAATA-3'. Primers for the 3' subfragment (2085 base pairs) were AtXDH2–2400for, 5'-TTATTTGCTACAGACGTG-3', and AtXDH2–3', 5'-TGATCCATCTTTCTCCCC-3'.

Generation of AtXDH1 Expressing P. pastoris—The cDNA clone AV548322 [GenBank] coding for full-length A. thaliana XDH1 was obtained from the Kazusa DNA Research Institute (Chiba, Japan). The yeast expression vector pPICZA with C-terminal His6 tag and P. pastors strain KM71 muts were purchased from Invitrogen (Carlsbad, CA). Standard molecular cloning techniques were used for DNA manipulation. The AtXDH1 cDNA was used as template for PCR to remove the 5'-untranslated region and stop codon and to generate a KpnI site at the 5' end and a ApaI site at the 3' end. Primers used for introducing restriction sites were AtXDH1 KpnI start, 5'-ATATATGGTACCATGGGTTCACTGAAAAAGGACGGC-3', and AtXDH1 ApaI stop, 5'-ATATATGGGCCCAACACTAAGATTAGGGTAGAAATCTGA-3'. The resulting PCR fragment containing the total coding region was cloned into pPICZA. P. pastoris was transformed with pPICZA/AtXDH1 and pPICZA (vector only) by electroporation according to the manual (EasySelect Pichia expression kit version A, Invitrogen). The presence of AtXDH1 cDNA in zeocin-resistant colonies was confirmed by PCR on the P. pastoris colonies.

Expression and Purification of AtXDH1—Several positive transformants were grown in 25 ml of BMGY (1% yeast extract, 2% peptone, 100 mM potassium phosphate, pH 6.0, 1.34% yeast nitrogen base without amino acids, 0.04% biotin, 1% glycerol, and 100 µg/ml zeocin) in a 250-ml baffled flask for 16–20 h (A600 2–3) at 30 °C and 150 rpm. Cells were collected by centrifugation and resuspended in 10 ml of BMMY (1% yeast extract, 2% peptone, 100 mM potassium phosphate, pH 6.0, 1.34% yeast nitrogen base without amino acids, 0.04% biotin, 0.3 mM sodium molybdate, and 0.5% methanol) in a 100-ml baffled flask and cultured again at 30 °C and 150 rpm. Cells were harvested after 0, 6, 10, 14, 18, 24, and 36 h of methanol induction by centrifugation and resuspended in breaking buffer (50 mM sodium phosphate, pH 7.4, 0.5 mM EDTA, 200 mM NaCl, 0.2 mM phenylmethylsulfonyl fluoride, and 5% glycerol). Cells were broken by vigorous vortexing with equal amounts of acid-washed glass beads (425–600 µm, Sigma) before cell debris and glass beads were removed by centrifugation. In the resulting supernatant XDH activity was examined by activity staining after native PAGE. Strongest intensity was detected 10 h after methanol induction with a gradual decrease until 36 h of incubation. The clone showing the highest XDH activity was selected for a large scale expression culture. Cells were grown in 250 ml of BMGY in a 1-liter baffled flask for 20 h, collected by centrifugation, and resuspended in 50 ml of BMMY in a 500-ml baffled flask. After cultivation for 10 h in BMMY the cells were harvested by centrifugation and resuspended in breaking buffer. Depending on the quantity of cells, they were broken either by vigorous vortexing with an equal volume of glass beads at 4 °C for a total of 30 min in bursts of 30 s alternating with cooling on ice or by three passages through a French press pressure cell with 14,000 p.s.i. operating pressure. After centrifugation the supernatant was used for purification of the His-tagged AtXDH1 protein by affinity chromatography with Ni-nitrilotriacetic acid (Ni-NTA)-superflow matrix (Qiagen, Hilden, Germany) under native conditions at 4 °C according to the manufacturers instructions. The sample was rebuffered to 50 mM Tris/HCl, pH 8.0, 5 mM EDTA, 2.5 mM dithiothreitol. For further purification AtXDH1 was subjected to anion exchange chromatography using a SourceTM 15Q column (Amersham Biosciences) equilibrated with 50 mM Tris/HCl, pH 8.0, 5 mM EDTA, 2.5 mM dithiothreitol (buffer A). Protein samples were applied to the column and eluted with buffer A followed by a linear gradient of 0 to 1 M NaCl in buffer A. Final purification and size determination was achieved by chromatography on a SuperdexTM 200 HR10/30 size exclusion column (Amersham Biosciences) equilibrated with 50 mM Tris/HCl, pH 8.0, 200 mM NaCl, 1 mM EDTA.

Determination of Protein Concentrations—Concentrations of total soluble protein were determined by use of Roti Quant solution (Roth, Karlsruhe, Germany) according to Ref. 28.

Wavescan of AtXDH1—Absorption spectroscopy was carried out using an Ultrospec 3000® spectrophotometer (Amersham Biosciences).

Sequence Analysis—Sequence analysis was performed with the ABI Prism Big Dye Terminator Cycle Sequencing Ready Reaction kit on an ABI Prism 310 cycle sequencer (PE Applied Biosystems, Warrington, UK) with a pop 6 polymer.

Expression of ABA3 and AO{alpha} from A. thaliana—Recombinant molybdenum cofactor sulfurase ABA3 and aldehyde oxidase AO{alpha} from A. thaliana were overexpressed and purified as described earlier in Refs. 29 and 30.

Enzyme Assays—For preparation of plant crude extracts plant material was squeezed at 4 °C in 2 volumes of extraction buffer (100 mM potassium phosphate, 2.5 mM EDTA, 5 mM dithiothreitol, pH 7.5), sonificated and centrifuged, and the supernatant was used for activity assays. XDH activity in plant crude extracts and recombinant AtXDH1 was visualized according to Ref. 31, except that native electrophoresis in the absence of SDS was run with 7.5% polyacrylamide gels and staining solution contained 1 mM hypoxanthine as substrate, 1 mM 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide and 0.1 mM phenazine methosulfate in 250 mM Tris/HCl, pH 8.5. For standard in gel XDH activity assays, each lane were loaded either with 80 µg of plant crude extract protein or with 1 µg of recombinant AtXDH1. The in vitro reconstitution of recombinant AtXDH1 by ABA3 was performed in a total volume of 0.4 ml of 50 mM Tris/HCl, pH 8.0. AtXDH1 (20 µg) was incubated with ABA3 (40 µg) in the presence of 1 mM L-cysteine for 1 h at 30 °C, followed by native PAGE with 1/5 volume of the reaction mixture and activity staining with hypoxanthine as substrate. Spectrophotometric determination of XDH activity was measured at 340 nm in a 1-ml reaction mixture containing 1 mM of the respective substrate, 1 mM NAD+, 50 mM Tris/HCl, pH 8.0, 1 mM EDTA and a suitable amount of recombinant AtXHD1. Reaction was started by addition of substrate. Inhibitors were preincubated with the enzyme for 5 min before starting the reaction. The xanthine-O2 reductase activity was measured under the same conditions but without NAD+, and O2-dependent production of uric acid was monitored at 295 nm. The production of superoxide radicals was monitored by following the reduction of cytochrome c at 550 nm. The specificity of -dependent reduction of cytochrome c was estimated by incorporating an excess of bovine superoxide dismutase in the assay mixture according to Refs. 32 and 33.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Cloning of AtXDH1 and AtXDH2 cDNAs and Their Tandem Arrangement in the Genome—Sequence similarity searches at "The Arabidopsis Information Resource" (TAIR)2 using human and bovine full-length XDH as query sequences detected two putative genes in the A. thaliana genome with higher similarity to XOR than to aldehyde oxidases (AO). Both genes were found in a tandem orientation on chromosome 4 with their reading frames pointing to the same direction (Fig. 1A). Based on the BAC clone AL079347 [GenBank] /ATF11I11 as reference both putative XDH genes, annotated as T11I11.130 and T11I11.140 and designated by us as AtXDH1 and AtXDH2, respectively, are located within the center region of this clone with an interspace of only 704 bp. The predicted genes range from position 50,304 (ATG) to 44,057 (TGA) for T11I11.130/AtXDH1 and from position 56,627 (ATG) to 51,009 (TGA) for T11I11.140/AtXDH2, thereby spanning regions of 6,248 and 5,619 base pairs, respectively. For both predicted genes expressed sequence tags were found indicating that both genes actually are transcribed. Although these expressed sequence tags clearly showed that annotation of the putative coding sequences is not fully correct they confirmed the predicted transcription start and stop sites. Neither extended data base searches nor genomic DNA hybridizations (data not shown) revealed more than these two XDH genes within the A. thaliana genome.



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  		FIG. 1.
Genomic and cDNA structure of AtXDH1 and AtXDH2. A, tandem orientation of AtXDH1 and AtXDH2 genes on chromosome 4, represented by the BAC clone ATF11I11. Large arrows indicate the orientation of genes. B, exon-intron structure of AtXDH1 and AtXDH2 genes. The overall sizes of the respective open reading frames (orf) are indicated and the relative sizes of exons (boxes) and introns (peaks) are shown; their lengths are indicated as numbers of base pairs.

 
Complete sequencing of the A. thaliana expressed sequence tag AV548322 [GenBank] has shown that the full-length open reading frame of AtXDH1 is interrupted by 13 introns and contains 4083 base pairs encoding a protein of 1361 amino acids (Fig. 1B) with 47% identity to human and bovine XDH. A full-length cDNA of AtXDH2 with an open reading frame of 4059 base pairs was obtained by reverse transcriptase-PCR using sequence information of the predicted AtXDH2 gene. The AtXDH2 gene also contains 13 introns highly conserved in exon/intron junctions compared with AtXDH1. The encoded AtXDH2 protein exhibits a length of 1353 amino acids with identities of 46% to human and bovine XDH. Both proteins, AtXDH1 and AtXDH2, reveal lower identities of 29–31% to AO proteins from A. thaliana but share overall identities of 93% to each other, indicating that they are similar to AO but functionally divergent. Full-length cDNA and protein sequences were deposited at GenBankTM (AY171562 [GenBank] for AtXDH1; AY518202 [GenBank] for AtXDH2).

Comparative primary structure analysis of AtXDH1 and AtXDH2 and XOR proteins from other organisms revealed a three-domain structure for both A. thaliana XDH monomers as is typical for XOR proteins. Like the chicken XDH (2) both A. thaliana XDH proteins contain a N-terminal domain including 8 strictly conserved cysteine residues for binding of two non-identical iron-sulfur clusters of the [2Fe-2S] type spanning amino acid positions 19 to 173 in AtXDH1 and 11 to 164 in AtXDH2, respectively. In each protein, the [Fe-S]-binding domain is followed by a FAD-binding domain (amino acids 260 to 440 in AtXDH1 and 252 to 432 in AtXDH2), whereas both domains are separated by hinge regions that are less conserved among all XOR proteins. FAD domains of both XDH proteins contain a FFLGYR motif (amino acids 417 to 422 in AtXDH1 and 409 to 414 in AtXDH2) that is supposed to be responsible for binding the second substrate NAD+ via the invariant tyrosine (2, 34). The third and C-terminal domain includes the Moco- and substrate-binding sites as well as the dimerization motif (35). In AtXDH1 it spans amino acid residues 612 to 1272 and in AtXDH2 604 to 1264, respectively, and is separated from the FAD/NAD domain by another hinge region. Within the Moco domain of XOR proteins both a strictly conserved glutamate and arginine residues are supposed to be essential for binding and proper positioning of purine substrates (36). AtXDH1 and AtXDH2 exhibit identical residues at the corresponding positions (Glu-831 and Arg-909 in AtXDH1, Glu-832 and Arg-901 in AtXDH2) indicating that the favored substrates of both proteins should be purines rather than aldehydes.

To find out at what evolutionary point XDH gene duplication might have occurred we analyzed the phylogenetic relationships of XDH proteins from various eukaryotic organisms (Fig. 2). Because AO is homologous to XDH but functionally divergent we have chosen three AO proteins from A. thaliana as an outgroup. The sequences used for this analysis show a splitting into three groups, among which plant sequences clearly form their own monophyletic subgroup besides animal and fungi XDH. Therein, A. thaliana XDH gene duplication appears to have happened long after the separation of dicots and monocots. Different from A. thaliana, none of the fully sequenced genomes of rice and C. reinhardtii were found to contain more than one XDH gene. Among the animals, vertebrates, insects, and nematodes group separately. Generally, the phylogenetic tree based on XDH protein similarities mirrors the species phylogeny and gives one more indication that AtXDH1 and AtXDH2 are in fact xanthine dehydrogenases rather than aldehyde oxidases.



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  		FIG. 2.
Phylogenetic neighbor joining tree of XDH proteins. Full-length sequences of XDH proteins and A. thaliana AOs were aligned using ClustalX, and phylogeny of XDH proteins was constructed by use of the neighbor joining method using PAUP 4.0. Numbers in the phylogenetic tree indicate 100 bootstrap replicates. A. thaliana AO1, AO2, and AO3 were used as an outgroup. The accession numbers of the XDH sequences are as follows: AtXDH1 (AY171562 [GenBank] ), AtXDH2 (AY518202 [GenBank] ), Oryza sativa (cDNA: AK065099 [GenBank] ), C. reinhardtii (not available), Emericella nidulans (CAA58034 [GenBank] , Neurospora crassa (EAA27223 [GenBank] , Homo sapiens (P47989 [GenBank] ), Rattus norvegicus (P22985 [GenBank] ), Mus musculus (CAA44705 [GenBank] , Bos taurus (CAA58497 [GenBank] , Gallus gallus (P47990 [GenBank] ), Drosophila melanogaster (S07245 [GenBank] ), Calliphora vicina (JQ0407), B. mori XDH1 (BAA21640 [GenBank] , B. mori XDH2 (BAB47183 [GenBank] , and Caenorhabditis elegans (NP_502747 [GenBank] ). Accession numbers for the AOs from A. thaliana are AtAO1 (BAA28624 [GenBank] , AtAO2 (BAA28625 [GenBank] , and AtAO3 (BAA82672 [GenBank] .

 
Differential Transcription and Enzyme Activities of AtXDH1 and AtXDH2—Because of the high similarities between the mRNAs of AtXDH1 and AtXDH2 and nearly identical transcript sizes, one cannot distinguish both transcripts by use of mRNA hybridization. Therefore we performed relative quantitative reverse transcriptase-PCR, using specific PCR primers designed for binding within the 3'-regions of each transcript and generating PCR fragments of 693 base pairs for AtXDH1 and 279 base pairs for AtXDH2, respectively. Proper amplification of the corresponding fragments was confirmed by sequencing.

As shown in Fig. 3A expression of AtXDH1 and AtXDH2 on the mRNA level can be detected in roots, leaves, stem, flowers, and siliques, indicating that both mRNAs are ubiquitously expressed in A. thaliana, although with varying amounts. Consistent with these findings also XDH activities were found in these organs. Unfortunately, discrimination of two XDH isoforms in non-denaturing polyacrylamide gels was impossible, either because of very similar physicochemical properties of both XDH proteins or because of the fact that only one isoform is actually translated. When analyzing the expression levels of AtXDH1 and AtXDH2 in plants of different age it turned out that mRNA levels of AtXDH1 increased in aging and senescent leaves, whereas AtXDH2 transcript levels remained unaltered (Fig. 3B) thereby simultaneously serving as an internal standard. In the same plants, a strong increase of XDH activity could be observed in senescent leaves but not at any other stage of development (Fig. 3B).



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  		FIG. 3.
Relative AtXDH1 and AtXDH2 mRNA expression and XDH activity at different conditions. A, relative mRNA expression (upper image) and enzymatic activity (lower image) of A. thaliana XDHs in different tissues; B, in leaves of different age (seedlings, 6 days; young, 2 weeks; adult, 3.5 weeks; aging, 6 weeks; senescent, 8 weeks); C, at salinity, cold and freezing stress; D, at drought stress treatment; and E, at ABA treatment in wild types and aba3.2 mutants. After RT-PCR, 2% agarose gels were loaded with 10 µl of the respective PCR reaction (M = 100 base pair ladder, upper band = 1000 base pairs; C = control; D = 4 h drought stressed). For XDH activity measurements, each lane on 7.5% native polyacrylamide gels was loaded with 80 µgof A. thaliana crude extract protein, and subsequently stained in the presence of hypoxanthine as substrate. Each figure represents one of at least 3 independent experiments that basically gave the same results.

 
When applying salt and cold stress to A. thaliana plants again the mRNA levels of AtXDH1, but not of AtXDH2, were altered. Whereas salinity (200 mM NaCl) caused a strong accumulation of AtXDH1 transcripts after 20 h, persistent cold at 4 °C as well as rapid freezing at -4 °C appeared to cause a decrease of AtXDH1 transcript levels (Fig. 3C). Although the change in mRNA amounts is more apparent under salt stress than at cold and freezing stress conditions no change in XDH activity could be observed at salinity, neither in leaves (Fig. 3C) nor in roots (data not shown). However, cold stress and rapid freezing clearly resulted in decreased activities of XDH.

Because dehydration is another common stress that plants have to cope with we exposed A. thaliana plants to drought stress that lasted for 4 h and resulted in a loss of fresh weight of about 50%. During this stress period mRNA amounts of AtXDH1 increased strongly in rosette leaves but at the same time dramatically decreased in roots (Fig. 3D), whereas again the levels of AtXDH2 mRNA basically remained unchanged. To find out whether this accumulation of AtXDH1 transcripts in leaves is because of the applied drought stress itself or because of associated stress-induced abscisic acid (ABA) synthesis we additionally analyzed mRNA amounts in wild type plants that were treated with exogenously applied ABA. This analysis was repeated with aba3 mutants that were exposed to the same drought stress, either with or without pretreatment of ABA. Because of a mutation in the Moco sulfurase gene aba3, mutants have lost the ability to activate XDH and AO by sulfuration, thereby rendered unable to respond to stresses that require AO-dependent ABA synthesis (29, 37, 38). Both the wild type and the aba3 mutants accumulated AtXDH1 transcripts upon ABA treatment but no increase was observed in aba3 mutants at drought stress without pretreatment of ABA (Fig. 3E). These data strongly suggest that transcript accumulation in A. thaliana leaves at drought stress is a consequence of stress-induced ABA synthesis and thereby solely indirectly related to drought. Concomitant to AtXDH1 transcript alteration XDH activity at drought stress was markedly increased in leaves and decreased in roots 4 and 20 h after starting stress treatment (Fig. 3D). ABA treatment without drought stress also resulted in slightly increased XDH activities in leaves of wild type plants (Fig. 3E), supporting that expression and activity of XDH are directly regulated by ABA rather than by drought stress itself.

Heterologous Expression and Purification of AtXHD1—The phylogenetic analyses (Fig. 2) indicate that AtXDH1 and AtXDH2 are in fact xanthine dehydrogenases rather than aldehyde oxidases, but unequivocal evidence comes only from the biochemical characterization of purified enzyme. First attempts to produce recombinant AtXDH1 in E. coli yielded only negligible amounts of recombinant protein, above all lacking cofactors and therefore being inactive. For this reason a eukaryotic system, the methylotrophic yeast P. pastoris, was chosen for heterologous expression of AtXHD1. The recombinant protein was purified by affinity chromatography followed by anion exchange chromatography. After this purification procedure the protein displayed one major band of 150 kDa in a Coomassie-stained SDS gel, corresponding well to the calculated molecular mass of 150.2 kDa for the deduced His6-AtXDH1 monomer. These data were confirmed by immunoblot analysis with anti-His antibody where the appropriate band was detected (Fig. 4A).



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  		FIG. 4.
Purification of AtXDH1 after expression in P. pastoris. A, SDS-PAGE analysis of AtXDH1 after different purification steps. Coomassie Brilliant Blue staining shows AtXDH1 as obtained from the P. pastoris crude protein extract (lane A,15 µg of protein), after Ni-NTA purification (lane B, 15 µg of protein), and after anion exchange chromatography (lane C, 5 µg of protein). Lane D reveals immunoblot analysis of Ni-NTA-purified AtXDH1 detected by use of anti-His antibody. B, activation of AtXDH1 by ABA3 and comparison of mobilities of AtXDH1 and XDH from leaf crude extracts on native PAGE. For XDH activation assay, 20 µg of AtXDH1 either were used as control (left lane) or were coincubated with 40 µg of ABA3 in the presence of 1 mM L-cysteine for 1 h at 30 °C before subjecting 1/5 volume of the reaction mixture to native PAGE (middle lane). Right lane was loaded with 80 µgof A. thaliana leaf crude extract and activity staining was performed with hypoxanthine as substrate. C, UV visible absorption spectrum of AtXDH1, recorded in 50 mM Tris/HCl, pH 8.0, containing 1 mM EDTA.

 
XOR and AO form a class of Mo-enzymes that are activated by sulfuration of their active center (39). This step is catalyzed by a highly specific enzyme named Moco sulfurase. Because we have recently cloned and purified the A. thaliana Moco sulfurase ABA3 (29) we wanted to know whether it could activate purified AtXDH1. After coincubation of AtXDH1 with the Moco sulfurase ABA3 an increase of AtXDH1 activity was observed (Fig. 4B), indicating that the recombinant protein as obtained from P. pastoris is not fully active. A second band appearing in this experiment above the typical AtXDH1 band might indicate the formation of an AtXDH1-ABA3 complex. When comparing gel mobility of purified recombinant AtXDH1 to native XDH from A. thaliana leaf crude extracts it was found that both bands appearing after native PAGE and subsequent activity staining showed identical mobilities (Fig. 4B). By use of gel filtration chromatography the molecular mass of recombinant AtXDH1 was found to range between 270 and 300 kDa, indicating that AtXDH1 in its native form is a homodimer (data not shown).

The absorption spectrum of AtXDH1 (Fig. 4C) is characterized by a maximum at 454 to 458 nm and a shoulder at about 540 to 590 nm, thereby corresponding to spectra of other molybdenum containing hydroxylases with typical peaks at 450 nm because of bound flavin chromophore, and a shoulder at 550 nm related to absorption of iron-sulfur centers. The absorption ratio of 2.8 at these two wavelengths is close to the ratio of 3 described for other XDH proteins revealing a FAD to FeS ratio of 1:4 (40, 41). Additionally, AtXDH1 shows strong absorption between 310 and 330 nm, which might be related to enedithiolene- and sulfo-molybdenum charge transfer bond.

Substrate Specificity of AtXDH1—XOR enzymes catalyze the oxidation of hypoxanthine to xanthine and of xanthine to urate with concomitant reduction of either NAD+ or molecular oxygen (42). When using hypoxanthine as substrate recombinant AtXDH1 reacted well with NAD+. However, molecular oxygen as the electron acceptor yielded a maximum substrate hydroxylation of about 2.5% compared with NAD+ (Table I) indicating that AtXDH1 occurs just in the dehydrogenase form. Based on the reaction with molecular oxygen, superoxide formation by AtXDH1 was measured by reduction of cytochrome c in the presence and absence of superoxide dismutase (32, 33). It was found that up to 22% of the electrons from xanthine were transferred to molecular oxygen to form superoxide radicals during catalysis.


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  		TABLE I
Use of molecular oxygen as electron acceptor and production of superoxide radicals by AtXDH1 Activity was assayed by monitoring the production of urate at 295 nm, superoxide-dependent reduction of cytochrome c was measured at 550 nm.

 
In addition to hypoxanthine and xanthine, AtXDH1 catalyzes the hydroxylation of purine and various aldehydes (Table II). Compared with hypoxanthine oxidation, hydroxylation rates of aldehydes ranged from 12.5% for heptaldehyde up to 31.3% for aromatic aldehydes like indole-3-carboxaldehyde under steady state conditions. AtXHD1 activity was found to be totally inhibited by allopurinol that is converted to alloxanthine and not released from the enzyme, as well as by p-hydroxymercuribenzoate, an inhibitor of SH groups, and potassium cyanide that removes the terminal sulfur ligand from the Moco of Mo-hydroxylases like XDH and AO. Comparing the substrate specificities of AtXDH1 to recombinant A. thaliana AO{alpha} demonstrated that hypoxanthine and xanthine are converted only by AtXDH1 (Fig. 5). Although AtXDH1 also converts several aldehydes conversion of these aldehydes by AO{alpha} is much more efficient. The pH optimum of AtXDH1 was found to range between 8.0 and 8.5 with NAD+ as oxidizing substrate.


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  		TABLE II
Substrate specificity of AtXDH1 Activity was measured as NAD+ reduction at 340 nm. AtXDHI and the respective inhibitor were coincubated 5 min prior to starting the reaction with hypoxanthine. Substrates were used in concentrations of 1 mM except for allopurinol and hydroxymercuribenzoate (each 0.1 mM) and potassium cyanide (50 mM).

 



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  		FIG. 5.
Substrate preferences of recombinant AtXDH1 and AO{alpha}. After native PAGE, activity bands were developed separately with strips from two lanes with equal concentrations (1 mM) of the following substrates: heptaldehyde (heptald.), 1-naphthaldhyde (naphthald.), indole-3-carboxaldehyde (indolecarb.), hypoxanthine (hypoxanth.), and xanthine. Each lane contained 10 µg of recombinant AtXDH1 (XDH) or AO{alpha} (AO).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The present study has demonstrated that A. thaliana possesses duplicated XOR genes that are arranged in tandem orientation in the genome. Both proteins represent xanthine oxidizing hydroxylases rather than AOs that do not require reduction of NAD+ for substrate conversion. Proving this assumption, recombinant AtXDH1 was shown to convert hypoxanthine and xanthine much more efficiently than aldehydes, although oxidation of aromatic indole-3-carboxaldehyde, abscisic aldehyde, and naphthaldehyde was still significant (Table II and Fig. 5). Recombinant AO{alpha} in turn was found to be unable to convert hypoxanthine and xanthine but oxidizes aldehydes much more efficiently than AtXDH1 (Fig. 5). Furthermore, AtXDH1 activity was completely inhibited by treatment with the typical XDH inhibitor allopurinol that is converted to alloxanthine, which remains tightly attached to the substrate binding pocket thereby preventing further substrate turnover (43).

XORs from mammals are present mainly in the dehydrogenase form but can readily be converted also to the oxidase form (44). Whereas XDH is the predominant enzymatic form in normal tissues and is involved in purine catabolism XO dominates in milk and in tissues subjected to injury where the production of reactive oxygen species is required. Both XDH and XO produce superoxide anions when O2 is used as electron acceptor. Although slower in comparison to XO, XDH produces more superoxide per mol of O2 but the formation of superoxide is nearly completely inhibited by the presence of NAD+. Therefore, the physiological significance of XDH-dependent superoxide generation in mammals is questionable because of excess amounts of NAD+ in cells under normal conditions (for review, see Ref. 6). However, depletion of the NAD+ available to XDH relative to O2 is supposed to be sufficient for XDH-catalyzed production of oxygen radicals (45). Interestingly, chicken XDH is not converted to XO and produces amounts of superoxide (40–44%; 46) very similar to bovine milk XDH (35–42%; 47) when using xanthine and O2 as substrates. Like chicken XDH, AtXDH1 appears to be present only in the dehydrogenase form indicated by the lack of two cysteine residues responsible for reversible conversion of rat XDH to XO (Cys-535 and Cys-992; 48) and the inefficient use of O2 as electron acceptor. Nevertheless, AtXDH1 transferred about 22% of the electrons from xanthine to O2 to produce superoxide radicals. Although the total amount of superoxide radicals produced by recombinant AtXDH1 appears to be too low for physiological significance the following points have to be highlighted: AtXDH1 transcript amounts increase dramatically at senescence accompanied by an even more obvious increase of XDH activity indicating that alteration of XDH activity in A. thaliana can be ascribed to alteration of AtXDH1 amounts and/or activity. Thus, under in vivo conditions higher amounts of native AtXDH1 protein that are activated far beyond normal levels are likely to produce also more superoxide. Whether or not superoxide production by AtXDH1 (and AtXDH2?) actually is of physiological importance remains to be investigated. But it is without doubt that plant XDH is involved in processes that require the formation of reactive oxygen species, such as pathogen defense (21, 22), cell death associated with hypersensitive reaction (23, 24), and natural senescence (25), thereby most likely fulfilling a function beyond purine degradation.

XDH gene duplication in A. thaliana, a situation that otherwise is known only from the silkworm Bombyx mori (12), forms the basis for differential gene regulation. Initially, expression studies on the mRNA level have shown that both AtXDH1 and AtXDH2 are ubiquitously expressed in all plant parts and distribution of XOR activity corresponds well with the mRNA distribution. However, in leaves of different age it became obvious that aging and natural senescence caused an increase of AtXDH1 transcripts only, but not of AtXDH2 transcripts. Concomitant to mRNA levels, only senescent leaves showed strongly increased XDH activities. These results are in accordance to Pastori and del Rio (25) who have observed a 7–8-fold increase of XDH activities in leaves of senescent pea plants concomitant to increasing activities of superoxide dismutase. Because oxidative processes during senescence require the generation of reactive oxygen species such as superoxide radicals it might well be that the function of plant XDH at senescence is the production of superoxide radicals rather than the degradation of purines, although the latter point should not be neglected because of the requirement of purine rescue for carbon and nitrogen remobilization during aging of plants. As found for AtXDH1, other degradative enzymes like RNases (49), proteinases (50–52), and lipases (53) are known to increase during the process of senescence. Interestingly, leaves of aging plants did not display altered XDH activities although senescent leaves, showing nearly the same increase of AtXDH1 transcripts, did so. This can be explained by the fact that XDH like AO requires a post-translational activation of the holoenzyme. According to changing environmental conditions the Moco sulfurase ABA3 is controlling the activities of XDH and AO (29, 37, 38) by changing the ratio of sulfurated/active to non-sulfurated/inactive enzymes, relatively independent from the total amount of holoprotein. It is most likely that the Moco sulfurase in aging leaves does not activate XDH beyond regular levels but is doing so at the stage of senescence.

Like in plants of different developmental stages also alteration of transcripts at various stresses is not mandatorily associated with alterations in XDH activities. But it appears that a decrease of XDH activity is regulated by transcript down-regulation like in cold-stressed plants and desiccated roots. On the other hand, increasing activities appear to result from two successive events, i.e. the accumulation of AtXDH1 transcripts and, presumably, subsequent sulfuration of XDH protein by its Moco sulfurase. As shown for AtXDH1, transcripts of ABA3 accumulate in drought-stressed leaves (29, 38) but not in roots,3 and ABA3 expression in leaves is stimulated by ABA treatment (38). In the case of AtXDH1 leaf-specific transcript accumulation at drought stress was found to be directly correlated to enhanced ABA levels rather than to the drought stress itself (Fig. 3E). This might indicate a common regulation for AtXDH1 and its activating enzyme ABA3 during processes that require ABA response. Similar to the situation at senescence the relevance of enhanced XDH activity at desiccation is not absolutely clear. It remains to be shown whether leaf-specific purine degradation is required for maintaining cell viability or whether also under drought stress conditions superoxide is produced in adaptation to this stress. Remarkably, at all conditions tested the amount of AtXDH2 transcripts remained more or less unaltered, whereas AtXDH1 displayed strong changes. Hence, one can conclude that altered XDH activities might be a consequence of increasing or decreasing amounts of AtXDH1 mRNA but not of AtXDH2 mRNA. Because final activation of XDH holoenzyme is carried out by ABA3 and therefore is not directly correlated to the amount of pre-existing protein it is so far unclear whether both XDH genes are actually translated and active or whether there is only one XDH protein in A. thaliana.

Up to now we have not been able to detect and distinguish two separate XDH activity bands in native PAGE supporting the possibility of only one active XDH isoenzyme. On the other hand, physicochemical properties of AtXDH1 and AtXDH2 might be nearly identical because of their high degree of homology and therefore might lead to identical migration in native PAGE making discrimination impossible. Nevertheless, we favor AtXDH1 either to be the only active XDH enzyme in A. thaliana or to be the one of greater physiological importance based on several observations: (i) at all stresses and treatments tested only AtXDH1 reacted significantly on the transcript level, whereas levels of AtXDH2 remained unaltered; (ii) concomitant to the alterations of AtXDH1 on transcript level also XDH activities changed the same way (except at salt stress); (iii) recombinant AtXDH1 and XDH from A. thaliana crude extracts displayed identical migration properties in native PAGE; and (iv) AtXDH1 was found to be able to produce superoxide, thereby likely being the protein reacting at senescence. In case of translated and active AtXDH2 we propose a more general and constitutive function during purine degradation but not at stress adaptation.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed. Tel.: 49-531-391-5870; Fax: 49-531-391-8128; E-mail: R.Mendel{at}tu-bs.de.

1 The abbreviations used are: XOR, xanthine oxidoreductase; ABA, abscisic acid; AO, aldehyde oxidase; Moco, molybdenum cofactor; XDH, xanthine dehydrogenase; XO, xanthine oxidase; RT, reverse transcriptase; Ni-NTA, nickel-nitrilotriacetic acid. Back

2 See www.arabidopsis.org. Back

3 F. Bittner, M. Bretthauer, and R. R. Mendez, unpublished data. Back


   ACKNOWLEDGMENTS
 
We thank the Kazusa DNA Research Institute (Chiba, Japan) for providing the expressed sequence tag AV548322 [GenBank] containing the AtXDH1 full-length cDNA, Tomokazu Koshiba (Tokyo, Japan) for kindly providing the AO{alpha}-expressing Pichia strain, and Jan Zeevaart (East Lansing, MI) for donating abscisic aldehyde. We are also grateful to Joern Petersen (Braunschweig, Germany) for help with phylogeny, Günter Schwarz for critically reading of the manuscript, and Saskia Helmsing for technical assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Stirpe, F., and Della Corte, E. (1969) J. Biol. Chem. 244, 3855-3863[Abstract/Free Full Text]
  2. Amaya, Y., Yamazaki, K., Sato, M., Noda, K., Nishino, T., and Nishino, T. (1990) J. Biol. Chem. 265, 14170-14175[Abstract/Free Full Text]
  3. Sato, A., Nishino, T., Noda, K., Amaya, Y., and Nishino, T. (1995) J. Biol. Chem. 270, 2818-2826[Abstract/Free Full Text]
  4. Hille, R., and Nishino, T. (1995) FASEB J. 9, 995-1003[Abstract]
  5. McCord, J. M. (1985) N. Engl. J. Med. 312, 159-163[Abstract]
  6. Nishino, T. (1994) J. Biochem. (Tokyo) 116, 1-6[Abstract/Free Full Text]
  7. Enroth, C., Eger, B. T., Okamoto, K., Nishino, T., Nishino, T., and Pai, E. F. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 10723-10728[Abstract/Free Full Text]
  8. Wright, R. M., Vaitaitis, G. M., Wilson, C. M., Repine, T. B., Terada, L. S., and Repine, J. E. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10690-10694[Abstract/Free Full Text]
  9. Berglund, L., Rasmussen, J. T., Andersen, M. D., Rasmussen, M. S., and Petersen, T. E. (1996) J. Dairy Sci. 79, 198-204[Abstract]
  10. Terao, M., Cazzaniga, G., Grezzi, P., Bianchi, M., Falciani, F., Perani, P., and Garattini, E. (1992) Biochem. J. 283, 863-870
  11. Keith, T. P., Riley, M. A., Kreitman, M., Lewontin, R. C., Curtis, D., and Chambers, G. (1987) Genetics 116, 67-73[Abstract/Free Full Text]
  12. Komoto, N., Yukuhiro, K., and Tamura, T. (1999) Insect Mol. Biol. 8, 73-83[CrossRef][Medline] [Order article via Infotrieve]
  13. Glatigny, A., and Scazzocchio, C. (1995) J. Biol. Chem. 270, 3534-3550[Abstract/Free Full Text]
  14. Leimkuehler, S., Kern, M., Solomon, P. S., McEwan, A. G., Schwarz, G., Mendel, R. R., and Klipp, W. (1998) Mol. Microbiol. 27, 853-869[CrossRef][Medline] [Order article via Infotrieve]
  15. Xi, H., Schneider, B. L., and Reitzer, L. (2000) J. Bacteriol. 182, 5332-5341[Abstract/Free Full Text]
  16. Boland, M. J. (1981) Biochem. Int. 2, 567-574
  17. Perez-Vicente, R., Alamillo, J. M., Cardenas, J., and Pineda, M. (1992) Biochim. Biophys. Acta 11117, 159-166
  18. Montalbini, P. (1998) Plant Sci. 134, 89-102[CrossRef]
  19. Montalbini, P. (2000) J. Plant Physiol. 156, 3-16
  20. Sauer, P., Frebortova, J., Sebela, M., Galuszka, P., Jacobsen, S., Pec, P., and Frebort, I. (2002) Plant Physiol. Biochem. 40, 393-400[CrossRef]
  21. Montalbini, P. (1992) J. Phytopath. 134, 218-228
  22. Montalbini, P. (1992) Plant Sci. 87, 225-231[CrossRef]
  23. Montalbini, P. (1995) Physiol. Mol. Plant Pathol. 46, 275-292[CrossRef]
  24. Montalbini, P., and Della Torre, G. (1996) Physiol. Mol. Plant Pathol. 48, 273-287[CrossRef]
  25. Pastori, G. M., and Del Rio, L. A. (1997) Plant Physiol. 113, 411-418[Abstract]
  26. Logemann, J., Schell, J., and Willmitzer, L. (1987) Anal. Biochem. 163, 16-20[CrossRef][Medline] [Order article via Infotrieve]
  27. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed., Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  28. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  29. Bittner, F., Oreb, M., and Mendel, R. R. (2001) J. Biol. Chem. 276, 40381-40384[Abstract/Free Full Text]
  30. Koiwai, H., Akaba, S., Seo, M., Komano, T., and Koshiba, T. (2000) J. Biochem. (Tokyo) 127, 659-664[Abstract/Free Full Text]
  31. Mendel, R. R., and Mueller, A. (1976) Biochem. Physiol. Pflanz. 170, 538-541
  32. McCord, J. M., and Fridovich, I. (1968) J. Biol. Chem. 243, 5753-5760[Abstract/Free Full Text]
  33. Ballou, D., Palmer, G., and Massey, V. (1969) Biochem. Biophys. Res. Commun. 36, 898-904[CrossRef][Medline] [Order article via Infotrieve]
  34. Nishino, T., and Nishino, T. (1989) J. Biol. Chem. 264, 5468-5473[Abstract/Free Full Text]
  35. Romao, M. J., Archer, M., Moura, I., Moura, J. J., LeGall, J., Engh, R., Schneider, M., Hof, P., and Huber, R. (1995) Science 270, 1170-1176[Abstract/Free Full Text]
  36. Glatigny, A., Hof, P., Romao, M. J., Huber, R., and Scazzocchio, C. (1998) J. Mol. Biol. 278, 431-438[CrossRef][Medline] [Order article via Infotrieve]
  37. Schwartz, S. H., Leon-Kloosterziel, K. M., Koornneef, M., and Zeevaart, J. A. (1997) Plant Physiol. 114, 161-166[Abstract]
  38. Xiong, L., Ishitani, M., Lee, H., and Zhu, J. K. (2001) Plant Cell. 13, 2063-2083[Abstract/Free Full Text]
  39. Wahl, R. C., Warner, C. K., Finnerty, V., and Rajagopalan, K. V. (1982) J. Biol. Chem. 257, 3958-3962[Abstract/Free Full Text]
  40. Coughlan, M. P. (1980) Moybdenum and Molybdenum-containing Enzymes, pp. 121-185, Pergamon Press, Oxford
  41. Mehra, R. K., and Coughlan, M. P. (1984) Arch. Biochem. Biophys. 229, 585-595[CrossRef][Medline] [Order article via Infotrieve]
  42. Hille, R. (1996) Chem. Rev. 96, 2757-2816[CrossRef][Medline] [Order article via Infotrieve]
  43. Robins, R. K., Revankar, G. R., O'Brien, D. E., Springer, R. H., Novinson, R., Albert, A., Senga, K., Miller, J. P., and Streeter, D. G. (1985) J. Heterocycl. Chem. 22, 601-634
  44. Della Corte, E., and Stirpe, F. (1968) FEBS Lett. 2, 83-84[CrossRef][Medline] [Order article via Infotrieve]
  45. Harris, C. M., and Massey, V. (1997) J. Biol. Chem. 272, 8370-8379[Abstract/Free Full Text]
  46. Nishino, T., Nishino, T., Schopfer, L. M., and Massey, V. (1989) J. Biol. Chem. 264, 2518-2527[Abstract/Free Full Text]
  47. Hunt, J., and Massey, V. (1992) J. Biol. Chem. 267, 21479-21485[Abstract/Free Full Text]
  48. Nishino, T., and Nishino, T. (1997) J. Biol. Chem. 272, 29859-29864[Abstract/Free Full Text]
  49. Taylor, C. B., Mariola, P. A., delCardayre, S. B., Raines, R. T., and Green, P. J. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 5118-5122[Abstract/Free Full Text]
  50. Hensel, L. L., Grbic, V., Baumgarten, D. A., and Bleecker, A. B. (1993) Plant Cell 5, 553-564[Abstract]
  51. Lohmann, K. N., Gan, S., John, M. C., and Amasino, R. M. (1994) Physiol. Plant 92, 322-328[CrossRef]
  52. Drake, R., John, I., Farrell, A., Cooper, W., Schuch, W., and Grierson, D. (1996) Plant Mol. Biol. 30, 755-767[CrossRef][Medline] [Order article via Infotrieve]
  53. Ryu, S. B., and Wang, X. (1995) Plant Physiol. 108, 713-719[Abstract]

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