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
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ABSTRACT |
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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 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 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.
Organisms and Culture Conditions--
E. coli DH5 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 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 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.
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
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 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.
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 ( 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").
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.
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).
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
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).
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).
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
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-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).
EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
(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.
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.
-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).
RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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.
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.
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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.
Representative purifications of PtxD from E. coli BL21(DE3)/pWM302 and
from P. stutzeri WM536
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.
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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.
View larger version (10K):
[in a new window]
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.
Substrate specificity and inhibition of PtxD
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).
View larger version (15K):
[in a new window]
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 (
), 100 (
), 167 (
), and 500 (
)
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
View larger version (14K):
[in a new window]
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 ( ), 25 (
), 50 (
), 75 (
), 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.
View larger version (17K):
[in a new window]
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 ( ), 5 (
), 10 (
),
15 (
), 20 (
), 25 (
), and 30 (
) 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
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[in a new window]
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
(
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.
|
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.
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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.
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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.
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
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ABBREVIATIONS |
---|
The abbreviations used are:
MOPS, 3-N-morpholinopropanesulfonic acid;
PAGE, polyacrylamide gel electrophoresis;
MES, 4-morpholine-ethanesulfonic
acid;
IPTG, isopropyl-1-thio--D-galactopyranoside.
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