Purification and Characterization of a Novel Phosphorus-oxidizing Enzyme from Pseudomonas stutzeri WM88*,

Amaya M. Garcia CostasDagger, Andrea K. White§, and William W. Metcalf

From the Department of Microbiology, University of Illinois, Chemical and Life Sciences Laboratory, Urbana, Illinois 61801

Received for publication, December 28, 2000, and in revised form, February 20, 2001


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The ptxD gene from Pseudomonas stutzeri WM88 encoding the novel phosphorus oxidizing enzyme NAD:phosphite oxidoreductase (trivial name phosphite dehydrogenase, PtxD) was cloned into an expression vector and overproduced in Escherichia coli. The heterologously produced enzyme is indistinguishable from the native enzyme based on mass spectrometry, amino-terminal sequencing, and specific activity analyses. Recombinant PtxD was purified to homogeneity via a two-step affinity protocol and characterized. The enzyme stoichiometrically produces NADH and phosphate from NAD and phosphite. The reverse reaction was not observed. Gel filtration analysis of the purified protein is consistent with PtxD acting as a homodimer. PtxD has a high affinity for its substrates with Km values of 53.1 ± 6.7 µM and 54.6 ± 6.7 µM, for phosphite and NAD, respectively. Vmax and kcat were determined to be 12.2 ± 0.3 µmol min-1 mg-1 and 440 min-1. NADP can substitute poorly for NAD; however, none of the numerous compounds examined were able to substitute for phosphite. Initial rate studies in the absence or presence of products and in the presence of the dead end inhibitor sulfite are most consistent with a sequential ordered mechanism for the PtxD reaction, with NAD binding first and NADH being released last. Amino acid sequence comparisons place PtxD as a new member of the D-2-hydroxyacid NAD-dependent dehydrogenases, the only one to have an inorganic substrate. To our knowledge, this is the first detailed biochemical study on an enzyme capable of direct oxidation of a reduced phosphorus compound.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Phosphorus is widely reported to be a redox conservative element in biological systems, with the sum total of phosphorus biochemistry consisting of the formation and hydrolysis of phosphate-ester bonds. These reports imply that reduced phosphorus compounds are not important in living systems and that enzymatically catalyzed redox reactions of phosphorus compounds do not occur; however, an increasing body of evidence indicates that this is not the case. Although it is true that inorganic phosphate (P valence +5) is the principal form of phosphorus in living systems and that phosphate-esters play a critical role in phosphate biochemistry, it is now clear that reduced phosphorus compounds of both natural and xenobiotic origin play important roles in numerous biological systems. Accordingly, many organisms have been shown to possess metabolic pathways for reduction of phosphate to a variety of reduced phosphorus compounds (1-3); others have been shown to possess metabolic pathways for oxidation of reduced phosphorus compounds (4-9). Among the most striking of these is a recently isolated sulfate-reducing bacterium that obtains all of the energy it requires for growth from the oxidation of phosphite (+3 valence) to phosphate (10).

Unfortunately, detailed studies examining the mechanisms of biological phosphorus oxidation and reduction are scarce. This is particularly true with regard to the biochemical characterization of enzymes involved in reduced phosphorus metabolism. A few of the enzymes involved in the biosynthesis of the reduced phosphorus antibiotic bialaphos (3, 11, 12) as well as the enzyme phosphoenolpyruvate phosphonomutase from Tetrahymena (13) have been purified and characterized. However, these carbon-phosphorus bond-synthesizing enzymes catalyze phosphorus reduction indirectly via intramolecular rearrangements; they do not catalyze direct redox reactions of phosphorus moieties. A similar situation exists for most enzymes involved in carbon-phosphorus bond cleavage. Thus, the electron-withdrawing nature of the beta -carbonyl groups in phosphonoacetate and phosphonoacetaldehyde renders the carbon-phosphorus bond in each of these compounds susceptible to hydrolytic cleavage by the enzymes phosphonoacetate hydrolase and phosphonoacetaldehyde hydrolase (14-17). The mechanism of the broad substrate specificity enzyme carbon-phosphorus lyase probably does not involve a simple hydrolytic mechanism, based on the examination of various substrates and their products (18). However, the mechanism of this enzyme remains obscure because in vitro activity of the enzyme has never been achieved, despite numerous attempts and the identification and characterization of the genes that encode it (18, 19).

Two biochemical studies of enzymes that presumably do catalyze direct phosphorus redox reactions have been reported. Malacinski and Konetzka (20, 21) did cell suspension studies and partially purified an NAD-dependent phosphite oxidoreductase from Pseudomonas fluorescens 195, and Heinen and Lauwers (8) did cell suspension studies with a hypophosphite oxidase from Bacillus caldolyticus. While these studies clearly demonstrate the enzymatic nature of the process, they do not greatly add to our understanding of the biochemistry of phosphorus redox reactions. In no case of which we are aware has an enzyme that catalyzes a direct phosphorus redox reaction been biochemically characterized in pure form.

Recently, we isolated an organism, Pseudomonas stutzeri WM88, that is capable of oxidizing phosphite and hypophosphite to phosphate (22). Molecular and genetic analyses suggested that oxidation of hypophosphite to phosphate in this organism occurs through a phosphite intermediate. These analyses also showed that there are two distinct chromosomal loci responsible for these oxidations: ptxABCD, required for phosphite oxidation, and htxABCDE, required for hypophosphite oxidation. Sequence analysis of these loci suggests that phosphite oxidation is catalyzed by the product of ptxD, a putative protein with significant homology to members of the D-isomer-specific 2-hydroxyacid dehydrogenase family. Hypophosphite oxidation is probably catalyzed by the product of htxA, which is predicted to encode an enzyme with significant homology to known 2-oxoglutarate-dependent dioxygenases. Our genetic findings encouraged us to study this system biochemically to learn more about the role of these enzymes in the oxidation of phosphite and hypophosphite. In this paper, we report the overexpression of the NAD-dependent phosphite dehydrogenase, PtxD, in E. coli, its purification to homogeneity, and its biochemical characterization.

    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Organisms and Culture Conditions-- E. coli DH5alpha (23) was used as the host for DNA cloning experiments, and E. coli BL21(DE3) (24) was used as the host for overexpression from plasmid pET11a (Novagen, Inc., Madison, WI) and its derivatives. These strains were grown in standard LB medium supplemented with ampicillin (50 µg/ml) or carbenicillin (100 µg/ml) as needed. All P. stutzeri strains are derivatives of the phosphite- and hypophosphite-oxidizing bacterium P. stutzeri WM88 (22). P. stutzeri WM536 is a mutant that does not produce extracellular capsule. P. stutzeri WM567 is a streptomycin-resistant derivative of P. stutzeri WM536. P. stutzeri WM581 (rpsL, del3(BsiWI)::aph) is a derivative of P. stutzeri WM567 that carries a deletion of the ptxABCDE operon and is unable to utilize either phosphite or hypophosphite as sole phosphorus sources. P. stutzeri strains were grown at 37 °C in 0.4% glucose-MOPS1 medium containing the indicated phosphorus source at 0.5 mM unless otherwise noted (25). Phosphite and hypophosphite were always prepared fresh and filter-sterilized prior to use. Cells were grown in 0.4% glucose-MOPS medium with 0.1 mM phosphate for studies involving phosphate-limited growth. Cells were grown in 0.12% glucose-MOPS medium with 2.0 mM phosphate for studies involving phosphate-excess growth. For large scale protein purifications, P. stutzeri WM536 was grown in a 30-liter stainless steel bioreactor (model P30A, B. Braun Biotech, Allentown, PA) at 30 °C in glucose-MOPS medium containing 2 mM hypophosphite. Antifoam 289 (Sigma) was added as needed. To ensure that no residual phosphate was present in the media, all glassware was soaked and rinsed with ultrapure deionized water prior to use. The bioreactor was rinsed with 0.1 M nitric acid prior to use for the same purpose.

Cloning and Overexpression of ptxD-- Standard methods for isolation and manipulation of plasmid DNA were used throughout (26). The ptxD gene was amplified by polymerase chain reaction from plasmid pWM294 (22) using Vent DNA polymerase (Life Technologies, Inc.) and the primers 5'-CACACACATATGCTGCCGAAACTCG-3' and 5'-AGCGGATAACAATTTACAGG-3'. The forward primer was designed to introduce an NdeI site (underlined) at the ptxD initiation codon. The resulting polymerase chain reaction product was digested with NdeI and BamHI and cloned into the same sites in the expression vector pET11a (Novagen, Inc., Madison, WI) to form pWM302. The ptxD gene in pWM302 was sequenced with standard T7 promoter and terminator primers at the W. M. Keck Center for Comparative and Functional Genomics (University of Illinois) and is identical to the previously determined sequence (22).

To induce overexpression of plasmid-borne genes, E. coli BL21 (DE3) transformants carrying either pWM302 or pET11a were grown in LB medium containing carbenicillin at 37 °C. Upon reaching midlog phase (A600 ~0.6), IPTG (1 mM final concentration) was added, and the cultures were incubated for an additional 1.5 h, at which time they were harvested by centrifugation. For large scale overexpression experiments, cultures were grown in the 30-liter stainless steel bioreactor at 30 °C.

Purification Steps-- All purification steps took place at 4 °C. Approximately 20 g (wet weight) of IPTG-induced BL21 (DE3)/pWM302 cells were resuspended in 35 ml of freshly made buffer A (20 mM MOPS buffer, pH 7.25, 10% glycerol, 1 mM dithiothreitol). DNase I (~10 mg) was added, and the suspension was passed twice through a chilled French pressure cell at 13,000 p.s.i. The broken cell slurry was then centrifuged at 20,000 × g for 30 min to pellet debris and unbroken cells, and the supernatant fraction was collected as the crude cell extract. The crude extract was separated into soluble and membrane fractions by centrifugation at 270,000 × g for 45 min. The pellet was discarded, and the supernatant fraction (high speed extract) was used in subsequent steps.

High speed extracts containing ~180-350 mg of protein were loaded onto an NAD-affinity column (~10 ml of swollen resin) with attachment of the ligand at C-8 (catalog no. N1008; Sigma) at a flow rate of 0.5 ml/min. Fractions from the flow-through containing PtxD activity were pooled, adjusted to 1 M NaCl, and loaded at the same flow rate onto an NAD affinity column (~15 ml of swollen resin) with attachment of the ligand at N-6 (catalog no. N9505; Sigma). Unbound proteins were eluted from the second column with 10 column volumes of buffer B (20 mM MOPS, pH 7.25, 10% glycerol, 1 mM dithiothreitol, 1 M NaCl) followed by 10 column volumes of buffer A. PtxD was then eluted with an NAD gradient (0-3 mM) in buffer A over 5 column volumes. Active fractions that were homogenous as determined by visual inspection of SDS-PAGE gels were pooled and then desalted and concentrated by ultrafiltration (Centriplus membrane; molecular mass cut-off 30,000 Da; Amicon, Beverly, MA).

PtxD from P. stutzeri WM536 was purified following the same tandem affinity protocol. Eluted fractions with specific activity higher than about 3.0 units/mg were pooled and purified through the tandem affinity protocol a second time. Active fractions from the second purification that were ~90% pure as determined by visual inspection of SDS-PAGE gels were pooled and concentrated as described above.

Protein and Enzyme Assays-- PtxD activity was assayed spectrophotometrically by continuously monitoring the absorbance of NADH at 340 nm. The extinction coefficient of 6220 M-1 cm-1 was used to calculate the concentration of NADH. Standard enzyme units (µmol of NADH produced min-1) are used throughout. Unless otherwise noted, the assay mixture contained 20 mM MOPS, pH 7.25, 0.5 mM NAD, 1 mM phosphite, and 10-100 µl of enzyme extract in a 1-ml volume. Most assays were carried out at room temperature. Characterization assays were carried out at 30 °C. For the temperature studies, acetylated bovine serum albumin (10 µg/ml final concentration) was added to the assay buffer. For the pH studies, the MOPS buffer was replaced by a Tris/acetate/MES buffer (100 mM Tris, 50 mM glacial acetic acid, and 50 mM MES), and the pH was adjusted with HCl or NaOH. The ionic strength of this buffer was calculated to be 0.1 at all pH values (27). Phosphate production was assayed colorimetrically by end point assays as described by Lanzetta et al. (28). Protein concentrations were assayed with Coomassie Plus reagent from Pierce according to manufacturer protocols with bovine serum albumin as the standard.

Gel Electrophoresis-- SDS-PAGE was carried out as described by Laemmli (29) in 12% polyacrylamide slab gels. Proteins were visualized by staining with Coomassie Blue. Native PAGE was carried out at 4 °C in 6% polyacrylamide continuous gels using a 35 mM HEPES, 43 mM imidazole buffer (final pH 7.1). Two activity stains were used. To detect phosphite-dependent NADH production, native PAGE gel slabs were incubated for 30 min at 30 °C in 100 ml of 100 mM Tris, pH 8.5, containing 10 mM phosphite, 25 mg of NAD, 30 mg of nitro blue tetrazolium, and 2 mg of phenazine methanosulfate as described by Heeb and Gabriel (30). Chemical reduction of the nitro blue tetrazolium dye by enzymatically produced NADH results in precipitation of a dark blue product, which is easily seen in the stained gels. To detect phosphate production from phosphite and NAD, native PAGE gel slabs were incubated in 100 ml of 100 mM Tris, pH 8.5, containing 10 mM phosphite, 25 mg of NAD, and 50 mM calcium chloride. The gels were then rinsed and stained with ammonium molybdate and methyl green as described by Cutting (31). Phosphate produced by the enzymatic oxidation of phosphite is precipitated as CaHPO4, which is visualized as a dark green band by the staining procedure.

Gel Filtration and Mass Spectrometry-- Gel filtration was carried out in a XK 16/70 column (Amersham Pharmacia Biotech) with Sephacryl S-300 as the matrix. The mobile phase was buffer A with 0.5 M NaCl, and the flow rate was 0.5 ml/min. A mixture of purified PtxD and the following standards was applied to the column for estimation of the native molecular mass of PtxD: bovine thyroglobulin (670,000 Da), bovine gamma -globulin (158,000 Da), chicken ovalbumin (44,000 Da), horse myoglobin (17,000 Da), and vitamin B12 (1350 Da). Mass spectrometry was carried out at the University of Illinois Mass Spectrometry facility using matrix-assisted laser desorption ionization in a Voyager-DE STR mass spectrometer (PerSeptive Biosystems, Framinghan, MA).

Amino Terminus Sequencing-- Purified PtxD was separated by electrophoresis under denaturing conditions in 12.5% polyacrylamide gels. The protein was then transferred onto a polyvinylidene difluoride membrane (Bio-Rad) using a Hoeffer Scientific semidry blotter according to manufacturer protocols and using Tris-glycine/methanol/SDS as the blotting buffer. Protein was visualized with Coomassie Blue and sequenced by Edman degradation at the University of Illinois Protein Sciences Facility.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The ptxD Gene Encodes an NAD:Phosphite Oxidoreductase-- Sequence analysis of the P. stutzeri ptxD gene suggests that it encodes an NAD:phosphite oxidoreductase (22). To test this hypothesis, we cloned the ptxD gene into a T7 expression plasmid and attempted to overexpress the PtxD protein in E. coli. Crude cell extracts were prepared from IPTG-induced strains carrying the ptxD overexpression plasmid, pWM302, and from control cells carrying the overexpression vector, pET11a, without an insert. Initial attempts to demonstrate phosphite-dependent NAD reduction in these crude extracts failed. However, phosphite-dependent NAD reduction (specific activity ~0.2 units mg-1) was observed in extracts prepared from the PtxD overexpression strain after high speed centrifugation to remove the membrane-associated NADH oxidase activity (high speed extracts). No activity was observed in high speed extracts of the vector only control, indicating that this activity was dependent on the ptxD gene.

Similarly, phosphite-dependent NAD reduction was detected in high speed cell extracts of P. stutzeri WM567 grown in media with either phosphite or hypophosphite as sole phosphorus sources (specific activity ~0.02 units mg-1 for both). However, the observed enzyme activity was significantly lower than that observed in extracts of the overproducing E. coli strain. Phosphite-dependent NAD reduction (specific activity ~0.02 units mg-1) was also observed in high speed extracts prepared from P. stutzeri WM567 grown in medium with a growth-limiting concentration of phosphate as the sole phosphorus source, while PtxD activity was not detected in extracts of cells grown in medium with excess phosphate. No activity was detected in extracts of the ptxD mutant P. stutzeri WM581 grown in phosphate-limiting medium, which again demonstrates that this activity requires the ptxD gene. Taken together, these data clearly indicate that the ptxD gene encodes an NAD:phosphite oxidoreductase. Further, the data obtained from P. stutzeri extracts indicate that ptxD expression is induced by phosphate starvation.

Purification of Native and Recombinant PtxD-- A two-step NAD-affinity protocol was developed that allows purification of recombinant PtxD after overexpression in E. coli. PtxD does not bind an NAD affinity column with C-8 attachment of the ligand, but this step is critical in eliminating other putative NAD-binding enzymes present in the high speed cell extract (data not shown). PtxD does bind a second NAD affinity column with attachment of the ligand at N-6. This binding occurs even in the presence of 1 M NaCl, which is required to prevent binding of unwanted proteins. An elution gradient of 0-3 mM NAD is used to recover the adsorbed protein from this second column. Other putative NAD-binding enzymes co-elute with PtxD for about half of the elution gradient. These fractions, estimated to be 95% pure, were used for preliminary analyses. During the second half of the elution gradient, the fractions contained homogenous PtxD as shown by SDS-PAGE (Fig. 1). The routine purification yield is ~50% (Table I). Native PtxD could be purified by the same protocol from extracts of hypophosphite-grown P. stutzeri WM536. As with recombinant PtxD, native PtxD did not bind the first affinity column, but it did bind the second one in the presence of 1 M NaCl. A preparation that had been purified through the tandem affinity protocol twice gave a preparation that was ~90% pure as estimated by SDS-PAGE (data not shown), with a yield of 9.3% (Table I). This preparation was sufficiently pure to allow mass spectrometric and amino terminus analyses. Importantly, the specific activity of the recombinant protein is essentially identical to that of the native protein.


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Fig. 1.   Overexpression and purification of recombinant PtxD. Protein samples from various stages of the purification were separated by SDS-PAGE and stained with Coomassie Blue as described. A two-step affinity protocol yields homogeneous recombinant enzyme. Lanes 1 and 9, marker proteins (size in kDa is shown); lane 2, lysed cells before IPTG induction; lane 3, lysed cells after IPTG induction; lane 4, crude cell extract; lane 5, cell-free crude extract; lane 6, high speed supernatant; lane 7, flow-through from first NAD affinity column; lane 8, purified enzyme (4.5 µg) from second NAD affinity column.

                              
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Table I
Representative purifications of PtxD from E. coli BL21(DE3)/pWM302 and from P. stutzeri WM536
Cell growth and enzyme purifications were as described under "Experimental Procedures."

Mass Spectrometry and Amino Terminus Sequencing-- To verify that PtxD produced in E. coli is identical to that produced by the native host, we sequenced the first 15 residues of the PtxD amino terminus from each preparation. Both preparations yielded the sequence MLPKLVITHRVHDEI, which is in complete agreement with the amino acid sequence predicted from the DNA sequence. We also carried out mass spectrometry analyses to examine whether PtxD is modified in either of the two organisms. The native and recombinant proteins gave peaks of 36413 ± 18 and 36,430 ± 18 daltons, respectively, in agreement with the predicted molecular mass of PtxD of 36,415 daltons. These results indicate that both organisms produce the same unmodified enzyme. In addition, both samples had an additional peak of approximately similar height corresponding to a mass ~190 daltons smaller than the predicted molecular mass (36,239 ± 18 daltons for the native preparation and 36,226 ± 18 daltons for the recombinant preparation). Because we obtained a unique amino-terminal sequence from both preparations, we believe this smaller peak represents a modified form of PtxD rather than a contaminating protein of nearly identical molecular weight. Further, the unique amino-terminal sequence suggests that the lower molecular weight peak is not the result of amino-terminal processing of PtxD. The loss of the two C-terminal residues (-AC, 174 daltons) is a possible explanation for this result. Importantly, a mixture of 50% native and 50% recombinant PtxD gave only the same two peaks, indicating that whatever the nature of the smaller peak it is not an artifact of overexpression in E. coli. Because the recombinant enzyme is apparently identical to the native enzyme and because the recombinant enzyme is produced in much higher amounts, this form was used for all of the remaining studies.

Characterization of PtxD-- Homogeneous preparations of PtxD catalyze the oxidation of phosphite to phosphate coupled to the reduction of NAD to NADH. NADH and phosphate production is strictly dependent on the presence of PtxD, NAD, and phosphite. Heat-denatured PtxD is incapable of catalyzing phosphite oxidation and NAD reduction (data not shown). In addition, production of phosphate and NADH was shown to be catalyzed by a single protein using enzymatic activity stains (Fig. 2). When assayed under standard conditions, the specific activity of PtxD, measured independently by the production of either phosphate or NAD, was 10.6 and 10.3 units/mg, respectively, indicating that phosphate and NADH production is stoichiometric. The reverse reaction, as measured by phosphate-dependent NADH oxidation, was not observed (with 4 mM phosphate and 1 mM NADH); however, this reaction is not expected to be significant based on thermodynamic considerations (see "Discussion").


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Fig. 2.   Native gel stained for PtxD activity. PtxD was separated by nondenaturing gel electrophoresis in a 6% continuous gel in HEPES/imidazole buffer. The gel was cut into three identical slices and stained either for total protein or for enzymatic activity. Total protein was detected by staining with Coomassie Blue (lane 1). To detect phosphite-dependent NAD reduction, gel slabs were incubated in Tris buffer with phosphite, NAD, and nitro blue tetrazolium as described. Production of NADH is detected by precipitation of the reduced tetrazolium dye as a purple band (lane 2). To detect phosphate production, gel slabs were incubated in Tris buffer with phosphite, NAD, and CaCl2 and stained with methyl green as described. Production of phosphate is indicated by a green stained band of precipitated CaHPO4 (lane 3). A single band is seen in each lane, indicating that a homogeneous preparation of PtxD catalyzes production of phosphate and NADH from phosphite and NAD.

Gel filtration analyses (data not shown) of purified PtxD suggest a native molecular mass of ~69 kDa, consistent with enzyme being a homodimer (the predicted molecular mass of the homodimer is 72.8 kDa). PtxD has a temperature optimum of 35 °C with a sharp decrease in activity at higher temperatures (Fig. 3A). It is active through a wide pH range (pH 5-9) with maximum activity from 7.25 to 7.75 (Fig. 3B). The addition of NaCl to the assay buffer has a pronounced negative effect on enzyme activity, with only 37% of the activity left at 200 mM NaCl (Fig. 3C). The addition of either EDTA or EGTA (10 mM final concentrations) to the assay buffer has no effect on enzyme activity, indicating that loosely bound metals are not involved in catalysis.


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Fig. 3.   Characterization of PtxD with respect to temperature, pH, and salt concentration. A, PtxD activity was assayed in the presence of 20 mM MOPS, pH 7.25, 1 mM phosphite, 0.5 mM NAD, and 10 µg/ml bovine serum albumin at increasing temperatures; B, PtxD activity was assayed in the presence of a 100 mM Tris, 50 mM acetate, 50 mM MES buffer at different pH (adjusted with HCl or NaOH), 1 mM phosphite, and 0.5 mM NAD; C, PtxD activity was assayed in the presence of 20 mM MOPS, pH 7.25, 1 mM phosphite, 0.5 mM NAD, and increasing concentrations of NaCl. The results shown are the average of three experiments.

Several alternative substrates were tested for their ability to substitute for either NAD or phosphite (Table II). NADP is able to substitute for NAD but only at substantially higher concentrations and substantially reduced rates. None of the compounds we tested is able to substitute for phosphite. These include several compounds that are substrates for homologous enzymes (glycerate, phosphoglycerate, lactate, 2-hydroxyisocaproate, and formate) and others (hypophosphite, methylphosphonate, arsenite, sulfite, and nitrite) that are structurally or chemically similar to phosphite. We also examined the ability of PtxD to utilize alternate substrates in the reverse direction. As described above, PtxD is unable to catalyze the reverse reaction (phosphate reduction) using NADH as an electron donor. PtxD is also unable to catalyze the reduction of nitrate, arsenate, sulfate, acetate, bicarbonate, methylphosphonate, aminoethylphosphonate, glycerate, or pyruvate (potential substrates were tested at 4 mM with 1 mM NADH; the limit of detection is ~0.025 units/mg under these conditions). PtxD did catalyze the reduction of hydroxypyruvate (4 mM hydroxypyruvate, 1 mM NADH), albeit at a very low level (0.14 units/mg).

                              
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Table II
Substrate specificity and inhibition of PtxD
All assays were conducted at 30 °C in 20 mM MOPS, pH 7.25. The concentration of additional components was varied as indicated in the footnotes. The results shown are the average of two experiments.

Finally, we examined whether a variety of compounds could act as enzyme inhibitors (Table II). One of the products of the reaction, phosphate, did not inhibit the activity of this enzyme; however, the other product of the reaction, NADH, is a strong inhibitor of PtxD activity, with complete inhibition observed at 4 mM. Several substrate analogs were examined for inhibitory activity. Sulfite was a strong inhibitor of PtxD activity, while nitrite, formate, D-glycerate, D-2-hydroxy-4-methylvalerate, hydroxyisocaproate, and arsenite moderately inhibited the activity. Several of the cofactor analogs tested were weak enzyme inhibitors, including ATP, ADP, ADP-ribose, and NADP. AMP does not inhibit PtxD. Detailed kinetic studies of enzyme inhibition are described below.

Initial Rate Studies-- PtxD activity was determined with varying levels of substrates in the absence of products (Fig. 4), and the data were fit to various kinetic models using a modified version (32) of the program of Cleland (33). These initial rate data show that the enzyme follows Henri-Michaelis-Menten kinetics and suggest that the reaction proceeds via a sequential mechanism. The Km values were determined to be 53.1 ± 6.7 and 54.6 ± 6.7 µM for phosphite and NAD, respectively. The Vmax is 12.2 ± 0.3 µmol min-1 mg-1, and kcat is 440 min-1 (per monomer). Data from fits to the sequential mechanism and to alternative mechanisms are presented in the supplementary material (Tables S1 and S2).


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Fig. 4.   Initial velocity patterns with NAD and phosphite. The reaction was initiated by adding 3.5 µg of PtxD to the reaction mixture. Left, the concentration of phosphite was varied at the fixed concentrations of NAD. Right, the concentration of NAD was varied at the fixed phosphite concentrations. Concentrations used for both substrates were 45 (), 56 (), 71 (open circle ), 100 (black-triangle), 167 (black-square), and 500 (black-diamond ) µM. Duplicate assays were performed at each concentration. The curve fits shown represent linear regression analysis of the data from each fixed concentration. Model fitting using the entire data set is described under "Results" and shown in the supplemental material, Tables S1 and S2

To distinguish between the random and ordered sequential mechanisms, initial rate studies were also carried out in the presence of products and in the presence of the dead end inhibitor sulfite. The type of inhibition and kinetic constants were determined by fitting the data to various kinetic models (33, 34). As described above, phosphate does not inhibit the PtxD reaction at a concentration of 4 mM; therefore, we attempted to show inhibition at higher levels of phosphate. No inhibition of PtxD activity by phosphate was observed with both phosphite and NAD held at concentrations approximating their respective Km values (50 µM each), even at phosphate concentrations of 100 mM. Thus, phosphate does not inhibit the PtxD reaction. In contrast, NADH does inhibit the PtxD reaction (Fig. 5). Initial rate studies in the presence of NADH suggest that it is a competitive inhibitor with respect to both phosphite (Kis = 115 ± 6 µM) and NAD (Kis = 233 ± 15 µM). Initial velocity studies in the presence of the dead end inhibitor sulfite suggest that it is a competitive inhibitor with respect to phosphite (Kis = 16.1 ± 0.1 µM) and an uncompetitive inhibitor with respect to NAD (Kii = 10.8 ± 0.1 µM) (Fig. 6). Data from fits to the indicated mechanisms and to alternative inhibition mechanisms are presented in the supplementary material for all experiments (Tables S3-S6).


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Fig. 5.   Initial velocity patterns in the presence of the product NADH. The reaction was initiated by adding 3.5 µg of PtxD to the reaction mixture. NADH was included in the assay mixtures at concentrations of 0 (black-diamond ), 25 (black-square), 50 (black-triangle), 75 (open circle ), and 100 () µM. Left, NAD was held constant at 50 µM with phosphite varied. Right, phosphite was held constant at 50 µM with NAD varied. Duplicate assays were performed at each concentration. The curve fits shown represent linear regression analysis of the data from each fixed NADH concentration. Model fitting using the entire data set is described under "Results" and shown in the supplemental material, Tables S3 and S4.


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Fig. 6.   Initial velocity patterns in the presence of the dead end inhibitor sulfite. The reaction was initiated by adding 3.5 µg of PtxD to the reaction mixture. Sulfite was included in the assay mixtures at concentrations of 0 (black-diamond ), 5 (black-square), 10 (black-triangle), 15 (open circle ), 20 (), 25 (), and 30 (triangle ) µM. Left, NAD was held constant at 50 µM with phosphite varied. Right, phosphite was held constant at 50 µM with NAD varied. Duplicate assays were performed at each concentration. The curve fits shown represent linear regression analysis of the data from each fixed sulfite concentration. Model fitting using the entire data set is described under "Results" and shown in the supplemental material, Tables S5 and S6.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Biological redox reactions involving phosphorus compounds are poorly understood, at best, due to a dearth of biochemically characterized enzymes. In this study, we purified to homogeneity PtxD, an enzyme that catalyzes oxidation of the reduced inorganic phosphorus compound phosphite. PtxD is an NAD-dependent dehydrogenase that oxidizes phosphite to phosphate with stoichiometric reduction of NAD to NADH. To our knowledge, this study is the first to characterize an enzyme in pure form that catalyzes direct oxidation of a reduced phosphorus compound; however, PtxD does resemble a phosphite oxidoreductase that was partially purified from P. fluorescens strain 195 (21). This enzyme also used NAD as a cofactor, was specific for phosphite and was inhibited by sulfite. It is possible that this enzyme is closely related to PtxD; however, these studies were incomplete, and because P. fluorescens strain 195 has been lost we cannot draw any definitive conclusions.

It could be argued that phosphite is not the true substrate of PtxD and that the observed oxidation of this compound represents a side activity of an enzyme that catalyzes a different reaction in vivo; however, several lines of evidence argue against this possibility. First, PtxD has a very high affinity toward phosphite, with a Km of ~50 µM. This low Km is even more significant given the observation that the ptxD gene forms an operon with a putative binding protein-dependent phosphite transport system (22). Because such transport systems are known to concentrate their substrates up to 100,000-fold (35), it seems almost certain that the observed Km is within the range of physiological significance. Second, we were unable to demonstrate activity of PtxD with any substrate other than phosphite, although numerous analogous compounds and compounds that are substrates of homologous enzymes were tested. Third, and most importantly, we showed that PtxD activity is induced by phosphate starvation. This strongly implies that the true substrate of PtxD is a phosphorus compound and that the function of PtxD is to provide the cell with an alternate source of phosphorus.

The enzymatic oxidation of phosphite is unprecedented. Two general chemical mechanisms can be envisioned for this reaction. The first involves nucleophilic attack at the phosphorus center and subsequent displacement of the hydride to the NAD acceptor. In this mechanism, the nucleophile could arise either from water (Fig. 7, Scheme 1) or from an amino acid side chain on the enzyme (Fig. 7, Scheme 2). In the latter case, a phosphoanhydride-linked enzyme intermediate requiring subsequent hydrolysis would be formed during the reaction. The second mechanism involves initial transfer of the hydride to the NAD acceptor and concomitant formation of the unstable compound metaphosphate (Fig. 7, Scheme 3). The studies presented here do not discriminate between these possibilities. More detailed studies designed to address this issue are beyond the scope of this initial enzyme description and are reported elsewhere (53).


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Fig. 7.   Possible chemical mechanisms for the PtxD reaction. Three possible chemical mechanisms for the concomitant oxidation of phosphite and reduction of NAD are shown. Schemes 1 and 2 involve initial nucleophilic attack at the phosphorus center and subsequent loss of the hydride. Scheme 3 involves initial loss of the hydride to produce the unstable intermediate metaphosphate. See "Results" for discussion.

Unlike most NAD-dependent dehydrogenases, PtxD does not appear to catalyze the reverse reaction (i.e. reduction of phosphate with NADH) to a measurable extent. This is not, however, a surprising result based on the thermodynamics of the reaction. At pH 7, the reduction potential (E) of the phosphate-phosphite couple is -650 mV (36), while that of the NADH-NAD couple is -320 mV (37). Thus, the reduction of NAD by phosphite is a significantly exergonic reaction (Delta G0' = -63.32 kJ/mol). Using this value, the equilibrium constant for the forward reaction is calculated to be 1.34 × 1011, and hence, the reduction of NAD by phosphite is essentially irreversible under physiological conditions. This suggests that PtxD may be particularly useful as a coenzyme regenerating enzyme for applications that require continuous regeneration of NADH (38).

Amino acid sequence comparisons indicate that PtxD is a member of the D-isomer-specific, 2-hydroxyacid NAD-dependent dehydrogenase protein family (39), the first with an inorganic substrate. An alignment of PtxD with several members of this family shows that it shares many of their characteristics, including the conserved NAD binding site and one of the Prosite signature sequences for this enzyme family (40) (Fig. 8). Chemical modification, site-directed mutagenesis, and crystallographic studies of several D-isomer-specific dehydrogenases have pointed to three residues, His292, Glu266, and Arg237 (PtxD numbering) essential for catalysis in this family of enzymes (41-45). Each of these residues is conserved in the enzyme family, and each is also present in PtxD. Formate dehydrogenase is the only exception in that it has a glutamine residue instead of the glutamate (46). These three residues correspond to the catalytic residues His195, Asp168, and Arg171 from L-lactate dehydrogenases (47). Similar to the proposed roles of these residues in lactate dehydrogenase, His292 is believed to act as a proton donor, Glu266 is believed to stabilize the positive charge from the protonated histidine, and Arg237 is believed to bind the carboxylate moiety of the hydroxyacid. In the case of PtxD, it seems plausible that Arg237 could bind the ionized hydroxyl groups of phosphite.


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Fig. 8.   Alignment of PtxD with D-hydroxyacid NAD-dependent dehydrogenases. FastA searches with PtxD against the nonredundant Swiss Protein Database show that PtxD is highly homologous to members of the D-hydroxyacid NAD-dependent dehydrogenases family (26-34.5% identical to the top 50 matches, most of which are known or putative members of the family; data not shown). Well studied representatives (crystal structures are available for five of the six sequences used) from this family were aligned with PtxD using ClustalW (52), clearly showing conservation of important features. Solid arrow, the NAD-binding motif; asterisks, the putative catalytic residues; dashed arrow, a conserved signature sequence for the D-isomer-specific 2-hydroxyacid family (see "Discussion" for details). Proteins used were as follows. PtxD, phosphite dehydrogenase from P. stutzeri WM88; FDH, formate dehydrogenase from Pseudomonas sp. 101; LDH, D-lactate dehydrogenase from Lactobacillus helveticus; GDH, D-glycerate dehydrogenase from Hyphomicrobium methylovorum GM2; SerA, D-3-phosphoglucerate dehydrogenase from E. coli; PdxB, erythronate-4-phosphate dehydrogenase from E. coli; HICDH, D-2-hydroxyisocaproate dehydrogenase from Lactobacillus casei. Swiss Protein accession numbers for the sequences used are O69054, P33160, P30901, P36324, P08328, P05459, and P17584, respectively.

In addition to these structural features, PtxD shares certain mechanistic features with other NAD-dependent dehydrogenases. Many NAD-dependent dehydrogenases follow a steady state ordered mechanism in which NAD is the first substrate to bind (47, 48). Among the D-isomer 2-hydroxyacid-specific family, the kinetic mechanism has been determined for only a single member. This enzyme, 2-hydroxyisocaproate dehydrogenase, follows the rapid equilibrium ordered mechanism with NAD binding first in the forward direction but follows the steady state ordered mechanism with NADH binding first in the reverse direction (49). Our initial rate studies are most consistent with PtxD following the steady state ordered mechanism, with NAD binding first and NADH being released last. The initial rate data from experiments conducted in the absence of products give reasonably good fits to the sequential mechanism. Further, these data clearly rule out the rapid equilibrium ordered and ping-pong mechanisms, each of which result in unique initial rate patterns that are distinct from the sequential mechanism (see supplemental material, Tables S1 and S2, for fitting data). That the mechanism is ordered rather than random is shown by the results of the inhibition experiments in the presence of the dead end inhibitor sulfite. The only mechanism consistent with the finding that sulfite is a competitive inhibitor of phosphite and an uncompetitive inhibitor of NAD is an ordered mechanism in which NAD binds first.

Although the initial rate studies appear to rule out a rapid equilibrium mechanism, the data from the product inhibition studies do not completely fit the pattern expected for the steady state sequential ordered mechanism; nor do they fit the patterns expected for any of the standard mechanistic patterns (50). One of the products, phosphate, does not inhibit the enzyme at any concentration tested. This observation would be easily explained by the irreversibility of the PtxD reaction if phosphate were the first product to be released from the enzyme in an ordered mechanism. Thus, phosphate would not be connected to the substrate side of the equation by a reversible step and, therefore, should not be an inhibitor. A similar observation was made regarding the NAD-dependent formate dehydrogenase from Phaseolus aureus. This enzyme also has essentially irreversible thermodynamics, follows a sequential ordered mechanism, and is not inhibited by its apparent product CO2 (Ref. 51; see also discussion in Ref. 34). Alternatively, lack of product inhibition by phosphate is the expected result for a rapid equilibrium ordered mechanism in which phosphate is the first product released; however, as described above, this result is inconsistent with the initial rate studies conducted in the absence of products. The other product, NADH, is a competitive inhibitor with respect to both phosphite and NAD. This observation is more difficult to explain. The only mechanism consistent with this result is the rapid equilibrium ordered mechanism without abortive ternary complexes. Again, this result is inconsistent with the initial rate data. The reason for this unusual pattern of NADH inhibition is unclear at this time.

Finally, the discovery of enzymes that are specific for reduced phosphorus compounds provides additional evidence for the existence of phosphorus redox cycling in nature. Although neither phosphite nor hypophosphite has been detected in the environment, the fact that there are organisms such as P. stutzeri that oxidize these phosphorus compounds to phosphate in a very specific fashion strongly suggests that these compounds must be present in the environment. Further biochemical and genetic studies are essential to increase our understanding of this interesting and important process.

    ACKNOWLEDGEMENTS

We are indebted to the Miller laboratory for generous sharing of equipment and to Rachel Larsen, Charles Miller, Biswarup Mukhopadhyay, Ralph Wolfe, and John Cronan for guidance. We thank Wilfred van der Donk for many helpful discussions and for critically reading the manuscript and Bryce Plapp for useful discussions on potential enzyme mechanisms.

    FOOTNOTES

* This work was supported in part by National Institutes of Health Grant GM59334. The Voyager mass spectrometer used by the University of Illinois Mass Spectrometry facility was purchased in part with Division of Research Resources, National Institutes of Health, Grant RR 11966.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains six tables.

Dagger Supported by a DeBeor Fellowship from the University of Illinois Department of Microbiology.

§ Supported by National Institutes of Health Grant GM07283-26.

To whom correspondence should be addressed: Dept. of Microbiology, University of Illinois, B103 Chemical and Life Sciences Laboratory, 601 S. Goodwin, Urbana, IL 61801. Tel.: 217-244-1943; Fax: 217-244-6697; E-mail: metcalf@uiuc.edu.

Published, JBC Papers in Press, February 22, 2001, DOI 10.1074/jbc.M011764200

    ABBREVIATIONS

The abbreviations used are: MOPS, 3-N-morpholinopropanesulfonic acid; PAGE, polyacrylamide gel electrophoresis; MES, 4-morpholine-ethanesulfonic acid; IPTG, isopropyl-1-thio-beta -D-galactopyranoside.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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