From the Department of Biomolecular Sciences,
Laboratory of Biochemistry, Wageningen Agricultural University,
Dreijenlaan 3, 6703 HA Wageningen, The Netherlands and
§ Hoechst Marion Roussel, Core Research Functions, building
G865A, D-65926 Frankfurt, Germany
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ABSTRACT |
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The conserved residues His-162 and Arg-269 of the flavoprotein p-hydroxybenzoate hydroxylase (EC 1.14.13.2) are located at the entrance of the interdomain cleft that leads toward the active site. To study their putative role in NADPH binding, His-162 and Arg-269 were selectively changed by site-specific mutagenesis. The catalytic properties of H162R, H162Y, and R269K were similar to the wild-type enzyme. However, less conservative His-162 and Arg-269 replacements strongly impaired NADPH binding without affecting the conformation of the flavin ring and the efficiency of substrate hydroxylation.
The crystal structures of H162R and R269T in complex with 4-hydroxybenzoate were solved at 3.0 and 2.0 Å resolution, respectively. Both structures are virtually indistinguishable from the wild-type enzyme-substrate complex except for the substituted side chains. In contrast to wild-type p-hydroxybenzoate hydroxylase, H162R is not inactivated by diethyl pyrocarbonate. NADPH protects wild-type p-hydroxybenzoate hydroxylase from diethylpyrocarbonate inactivation, suggesting that His-162 is involved in NADPH binding. Based on these results and GRID calculations we propose that the side chains of His-162 and Arg-269 interact with the pyrophosphate moiety of NADPH. An interdomain binding mode for NADPH is proposed which takes a novel sequence motif (Eppink, M. H. M., Schreuder, H. A., and van Berkel, W. J. H. (1997) Protein Sci. 6, 2454-2458) into account.
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INTRODUCTION |
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p-Hydroxybenzoate hydroxylase is the most thoroughly
characterized member of a group of inducible flavoprotein
monooxygenases which are involved in the biodegradation of aromatic
compounds by soil microorganisms (1). The enzyme catalyzes the
hydroxylation of 4-hydroxybenzoate to 3,4-dihydroxybenzoate,
i.e. the first step of the -ketoadipate pathway (2),
using NAD(P)H as electron donor shown in Scheme
1. The dihydroxylated aromatic product is readily subject to ring fission and further catabolism, allowing the
microbes to grow (2). p-Hydroxybenzoate hydroxylase has been
isolated from many microorganisms, and several gene sequences are
presently known (3). However, most information regarding its structure
and function comes from studies on the strictly NADPH-dependent enzymes from Pseudomonas strains
(4). The kinetic properties of p-hydroxybenzoate hydroxylase
have been elucidated (5), together with the catalytic mechanism (6, 7).
The catalytic cycle of p-hydroxybenzoate hydroxylase and
related flavoenzymes can be separated in two half-reactions, both
involving ternary complex formation. In the reductive part of the
reaction, the substrate acts as an effector, highly stimulating the
rate of flavin reduction by NADPH (8). After NADP+ release,
the substrate is hydroxylated in the oxidative part of the reaction
through electrophilic attack of a transiently stable oxygenated flavin
intermediate (6). Efficient hydroxylation requires substrate activation
upon binding (4). This prevents the uncoupling of the hydroxylation
reaction from oxygen reduction which would result in the production of
potential harmful hydrogen peroxide.
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The crystal structure of the enzyme-substrate complex of
p-hydroxybenzoate hydroxylase is known in atomic detail (9,
10). The enzyme is a homodimer of two independently acting 44-kDa
polypeptide chains containing a noncovalently bound FAD molecule (11).
Each subunit is built up from three domains, the FAD binding domain containing a characteristic -fold (12), the substrate binding domain, and the interface domain (10). The substrate is deeply buried
in the interior of the protein (Fig. 1)
and residues from all three domains are involved in catalysis.
Crystallographic studies have revealed that the flavin ring can attain
different orientations in and out of the active site (13-15). The
mobility of the flavin cofactor is thought to be essential for the
exchange of substrate and product during catalysis (13, 14) and for the
recognition of NADPH (15). In contrast to many other pyridine nucleotide-dependent enzymes, p-hydroxybenzoate
hydroxylase lacks a well defined domain for binding the coenzyme.
Sequence alignments suggest that this might be a common property of the
family of flavoprotein aromatic hydroxylases (16). So far, no crystal structures of p-hydroxybenzoate hydroxylase with NADP(H) or
pyridine nucleotide analogs have been obtained (17).
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Chemical modification studies of wtPHBH1 from Pseudomonas fluorescens have indicated the involvement of histidine (18), arginine (19), and tyrosine (20) residues in NADPH binding. Based on this information and the available crystallographic data, a potential mode of NADPH binding was proposed (20). In this model, the pyrophosphate moiety of NADPH binds in a cleft leading toward the active site and interacts with the side chains of His-162 of the FAD-binding domain and Arg-269 of the substrate binding domain (12). Despite their location near the protein surface (Fig. 1), both residues are highly conserved in p-hydroxybenzoate hydroxylase enzymes of known primary structure (3).
To investigate the validity of the proposed NADPH binding mode in more detail, we have undertaken the characterization of His-162 and Arg-269 variants of p-hydroxybenzoate hydroxylase from P. fluorescens. In this work we show that the His-162 and Arg-269 replacements weaken the NADPH binding without affecting the protein structure and the efficiency of substrate hydroxylation. Based on these and additional experiments, a refined model for the interdomain binding of NADPH is proposed. A preliminary account of this work has been presented elsewhere (1, 21).
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EXPERIMENTAL PROCEDURES |
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Materials-- Diethylpyrocarbonate was purchased from Janssen Chemicals. All other chemicals have been described elsewhere (22).
Site-specific Mutagenesis-- Site-specific mutagenesis of the gene encoding wtPHBH from P. fluorescens was performed in bacteriophage M13mp18 according to the method of Kunkel et al. (23). The oligonucleotide 5'-GCGATGGCTTCXXXGGCATCTCG-5' (where XXX denotes the replacement of CAC for GAC (H162D), AAC (H162N), TCC (H162S), ACC (H162T), TAC (H162Y), and AGG (H162R), respectively) was used as primer for the construction of His-162 mutants. The oligonucleotide 5'-GCGCCGCTGXXXAGCTTCGTGG-3' (where XXX denotes the replacement of CGC for GAC (R269D), AAA (R269K), AAC (R269N), TCC (R269S), ACC (R269T), and TAC (R269Y), respectively) were used as primers for the construction of Arg-269 mutants. The mutations were introduced into the Escherichia coli gene encoding the microheterogeneity-resistant mutant C116S (24). All mutations were confirmed by nucleotide sequencing according to Sanger et al. (25). For convenience and in view of identical catalytic properties (26), C116S is further referred to as wtPHBH.
Enzyme Purification--
Mutated p-hydroxybenzoate
hydroxylase genes were expressed in transformed E. coli TG2
grown in 5-liter batches of tryptone/yeast medium containing 100 µg/ml ampicillin and 20 µg/ml
isopropyl-1-thio--D-galactopyranoside at 37 °C under
vigorous aeration. The mutant enzymes were purified by a slightly
modified procedure of the purification protocol developed for wtPHBH
(26). The enzyme solution obtained after protamine sulfate treatment
was loaded onto Q-Sepharose FF, equilibrated in 20 mM
Tris/sulfate, pH 8.0. After washing, the enzyme was eluted with 0.2 M KCl and dialyzed against the starting buffer. The enzyme was then loaded onto Cibacron blue 3GA-agarose, equilibrated in 40 mM Tris/sulfate, pH 8.0. R269D, R269N, R269S, R269T, and
R269Y eluted during washing, whereas R269K and all His-162 mutants
eluted with 0.2 M KCl. After dialysis in 7 mM
potassium phosphate buffer, pH 7.0, the mutant enzymes were passed
through a hydroxyapatite column (27) and purified to apparent
homogeneity by fast protein liquid anion exchange chromatography (28).
Purified enzymes were stored as ammonium sulfate precipitates at
4 °C.
Analytical Methods-- Molar absorption coefficients of protein-bound flavin were determined in 50 mM sodium phosphate buffer, pH 7.0, by recording absorption spectra in the absence and presence of 0.1% SDS (29). Difference spectra with Cibacron blue 3GA were recorded with an automated Aminco DW-2000 spectrophotometer, essentially as described by Thompson and Stellwagen (30). Dissociation constants of enzyme-ligand complexes were determined fluorimetrically (31). Oxygen consumption experiments were performed as described previously (3). Aromatic product formation was analyzed by reverse phase HPLC (29). Kinetic experiments were performed at 25 °C in 100 mM Tris/sulfate pH 8.0, unless stated otherwise. The standard activity of p-hydroxybenzoate hydroxylase was measured as reported earlier (32). Steady-state kinetic parameters were determined as described previously (33). Rapid-reaction kinetics were performed with a High-Tech Scientific SF-51 stopped flow spectrophotometer (Salisbury, Wilts, United Kingdom), equipped with an anaerobic kit and interfaced to an Hyundai 486 microcomputer for data acquisition and analysis (22). pH-dependent rapid reaction experiments were performed in 40 mM Mes, pH 6-7, or 40 mM Hepes, pH 7-8. The ionic strength of buffers was adjusted to 50 mM with added sodium sulfate (34).
Chemical Modification with
Diethylpyrocarbonate--
Ethoxyformylation of histidine residues was
performed by adding 0.5 mM diethylpyrocarbonate to 20 µM enzyme in 80 mM Mes, pH 5.8 (I = 0.1 M) (18). Chemical modification was
interrupted by dilution or Bio-Gel P6DG filtration after addition of
excess imidazole. The amount of modified histidines was determined at 244 nm (244 = 3.6 mM
1
cm
1).
Crystallization-- Crystals of POHB complexed H162R and R269T were obtained using the hanging drop vapor diffusion method. The protein solutions contained 10-15 mg/ml enzyme in 100 mM potassium phosphate buffer, pH 7.0. The reservoir solution contained 40% saturated ammonium sulfate, 0.04 mM FAD, 0.15 mM EDTA, 2 mM POHB, and 30 mM sodium sulfite in 100 mM potassium phosphate, pH 7.0. Drops of 2 µl of protein solution and 2 µl of reservoir solution were allowed to equilibrate at 4 °C against 1 ml of reservoir solution. Crystals with dimensions of up to 0.3 × 0.2 × 0.1 mm3 grew within 5 days.
Data Collection--
X-ray diffraction data were collected using
a Siemens multiwire area detector and graphite monochromated CuK
radiation from an 18-kW Siemens rotating anode generator, operating at
45 kV and 100 mA. Data were processed using the XDS package (35). Data
collection statistics are given in Table
I.
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Refinement--
A starting electron density map was calculated
for H162R based on the structure of the wtPHBH·POHB complex (10),
after a correction had been made for the slightly different cell
dimensions (13). The 2Fo-Fc and
Fo-Fc maps clearly showed the
replacement of His-162 by Arg. His-162 was changed into Arg and fitted
in the electon density map with the graphics program O (36). The complete protein model was inspected and corrected where necessary. Refinement consisted of four macrocycles of map inspection and rebuilding using O with subsequent energy minimization and temperature factor refinement using the X-plor package (37). For the FAD we used the parameters as described by Schreuder et al.
(13). Water molecules were assigned by searching
Fo-Fc maps for peaks of at least
4, which were between 2.0 and 5.0 Å of other protein or water
atoms. Water molecules with temperature factors after refinement in
excess of 70 Å2 were rejected. The refined structure has
an R factor of 12.8% for 6695 reflections between 8.0 and
3.0 Å and contains 218 water molecules. The root mean square
deviations from ideal values are 0.010 Å for bond lengths and 1.52°
for bond angles.
GRID Calculations--
Energetically favorable binding sites for
the pyrophosphate moiety of NADPH in wtPHBH were computed with the GRID
program (38) using HPO42 and
PO43
as probe groups on a Silicon graphics
Indigo workstation. The parameters used to evaluate the nonbonded
interactions (including electrostatic, hydrogen bond, and Lennard-Jones
functions) of probe groups are based on the "extended" atom concept
used for the program CHARMM (39). Three-dimensional contour surfaces generated at selected energy levels were displayed with the program O
(36) together with wtPHBH (10).
NADPH Model--
The contour surfaces of the GRID calculations
were used as a start for building a three-dimensional model of the
enzyme-substrate complex in the presence of NADPH. The pyrophosphate
part of the NADPH molecule was fitted on the position of the GRID
contours with the graphics program O. The nicotinamide ring was placed at the re-side of the flavin ring by rotating around single
bonds. Similarly, the 2'-phosphate group was placed next to helix H2 as
indicated by mutagenesis studies (1). The model was then energy
minimized by using the conjugate gradient method using the XPLOR
package (37). The ternary complex was minimized allowing both NADPH and
enzyme-substrate complex to move, but the protein C-atoms were
fixed. The parameter set as determined by Engh and Huber (40) was used
for the protein part of the structure. The POHB, NADPH, and FAD
parameters were originally derived from CHARMM parameters (39). During
energy minimization the charges of the residues Arg, Lys, Glu, and Asp
were removed. The final model was obtained after 120 minimization
cycles with an initial drop in the energy of 40 kcal/mol.
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RESULTS |
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Enzyme Purification-- All His-162 mutants were expressed at high levels (5-10% of total protein) in E. coli TG2, and their yield after purification was comparable to that of wtPHBH (26). Comparable levels of protein expression were observed with the Arg-269 variants. However, with the exception of R269K, the Arg-269 variants were not retained on the Cibacron blue 3GA column used in the standard purification protocol. Such change in binding characteristics has also been observed with malate dehydrogenase variants (41). Since Cibacron blue 3GA is a strong inhibitor of wtPHBH, competitive with respect to NADPH (42), it was of interest to study the interaction of the His-162 and Arg-269 variants with the dye free in solution. Fig. 2 shows that the absorption perturbation difference spectra of wtPHBH and R269T, recorded in the presence of increasing concentrations of Cibacron blue 3GA, were nearly identical. It has been suggested that the shape of these difference spectra is indicative of electrostatic enzyme-dye interactions (43, 44). From analyzing the spectral data of Fig. 2 according to the procedure described by Thompson and Stellwagen (30), it was deduced that the dissociation constant of the wtPHBH-Cibacron blue 3GA complex (Kd = 0.34 ± 0.05 µM) is about one order of magnitude lower than the dissociation constant of the R269T-Cibacron blue 3GA complex (Kd = 4.6 ± 1.2 µM). Similar results as with wtPHBH were obtained for R269K and the His-162 variants. However, the affinity between Cibacron blue 3GA and the other Arg-269 mutants compared more favorable with that of R269T (not shown). Neither the dissociation constant of the enzyme-dye complexes nor the shape of the difference spectra changed significantly in the presence of 1 mM POHB or 1 mM NADPH. The dissociation constants of the enzyme-dye complexes were also determined from fluorimetric binding studies, using an enzyme concentation of 2 µM. With all His-162 and Arg-269 variants and similar to wtPHBH, quenching of flavin fluorescence was observed. From the titration curves and treating the data according to 1:1 binding (30), dissociation constants of 0.22 ± 0.05 µM and 1.2 ± 0.3 µM were estimated for the wtPHBH-Cibacron blue 3GA and R269T-Cibacron blue 3GA complex, respectively. These values are in reasonable agreement with the values deduced from the absorption difference spectral analysis. Altogether, the binding studies with Cibacron blue 3GA suggest that there is a corrrelation between the affinity of the mutant enzymes with the free and the immobilized dye. However, it is clear that the binding characteristics of the free dye do not simply predict the affinity of the mutant enzymes with the immobilized dye.
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Substrate Binding-- The His-162 and Arg-269 replacements did not change the optical properties of the flavin prosthetic group. The flavin absorption spectra of all mutants were identical to wtPHBH (26). Moreover, the flavin perturbation difference spectra in the presence of POHB indicated the flavin "in" conformation (14). Similar to the wild-type enzyme (26), the flavin fluorescence of the His-162 and Arg-269 mutants strongly decreased upon POHB binding. For all mutants, substrate binding could be described by simple binary complex formation with dissociation constants ranging from 20 to 40 µM.
Reaction Stoichiometry-- Oxygen consumption experiments revealed that in the presence of excess POHB and limiting NADPH, equal amounts of oxygen and NADPH were consumed. No hydrogen peroxide formation was detected when catalase was added at the end of the reactions. HPLC product analysis confirmed that all His-162 and Arg-269 mutants fully coupled enzyme reduction to substrate hydroxylation with stoichiometric formation of 3,4-dihydroxybenzoate.
Steady-state Kinetics-- The steady-state kinetic parameters of the His-162 and Arg-269 mutants were studied at pH 8.0, the optimum pH for turnover of wtPHBH (31). In agreement with the dissociation constants reported above, no significant changes in apparent Km values for POHB were observed (Table II). However, the apparent Km values for NADPH varied strongly (Table II), indicating that the type of amino acid residue engineered at position 162 or 269 drastically affects coenzyme binding. As can be seen from Table II, H162R, H162Y, and R269K are rather efficient enzymes with similar catalytic properties as wtPHBH. H162K and R269S are less efficient due to a clear increase in the apparent Km for NADPH. With H162D, H162N, H162S, H162T, R269D, R269N, R269T, and R269Y no reliable turnover rates could be estimated due to impaired NADPH binding.
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Pre-steady-state Kinetics--
The NADPH binding was further
investigated by stopped-flow kinetics (pH 8.0, 25 °C). In these
experiments, the anaerobic reduction of the enzyme-substrate complex
was followed by measuring the decrease in FAD absorption at 450 nm with
time as a function of the NADPH concentration (22, 45). The rate
constant of reduction of the His-162 mutants is strongly dependent on
the type of amino acid residue engineered (Fig.
3A). With H162R, the rate
constant of flavin reduction strongly increased with increasing
concentrations of NADPH and a maximal reduction rate constant of about
240 s1 and an apparent Kd for NADPH of
0.23 mM were estimated. These values are in the same range
as reported for the wild-type enzyme (26) (Table II), suggesting that
the H162R replacement does not significantly affect the effector role
of POHB and the mode of NADPH binding. With H162Y and to a lesser
extent with H162K, a high rate constant of reduction was found but with
a modestly elevated Kd for NADPH, revealing poor
NADPH binding (Table II). With the other His-162 mutants, the rate
constant of flavin reduction increased slightly with increasing NADPH
concentrations (Fig. 3A), and the almost linear dependence
of the reduction rate constant with NADPH concentration confirmed that
these mutants have lost the ability of proper coenzyme binding. In
agreement with the steady-state kinetic data, drastic changes in
catalysis were observed when the reductive half-reactions of the
Arg-269 mutants were studied. Except for R269K, all Arg-269 mutants
were rather slowly reduced due to impaired coenzyme binding (Fig.
3B, Table II).
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Chemical Modification Studies-- wtPHBH is readily inactivated by diethylpyrocarbonate at pH 6 (18). This inactivation was tentatively ascribed to the cooperative modification of His-162 and His-289, possibly both involved in NADPH binding (18). Similar to wtPHBH, incubation of H162R with diethylpyrocarbonate at pH 6 led to the modification of about four histidines (Fig. 5A). However, in the wild-type enzyme, the initial rate constant of ethoxyformylation is higher than with H162R and is accompanied with a much stronger decline in enzyme activity (Fig. 5B). During the initial stage of the chemical modification reactions, the ethoxyformylation of one histidine residue resulted in a loss of activity of more than 50% in wtPHBH and less than 10% in H162R (Fig. 5B). In contrast to wtPHBH (K'm NADPH modified enzyme >100 µM), ethoxyformylation of H162R did not change the apparent Km NADPH (cf. Table II), suggesting that His-162 is a main target of diethylpyrocarbonate modification which is involved in NADPH binding.
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Structural Properties--
All His-162 and Arg-269 variants were
tested by crystallization assays. Crystals with high and moderate
quality diffraction properties were obtained for R269T and H162R, both
in complex with POHB. The electron density maps of H162R and R269T
clearly identified the side chain substitutions (Figs.
6 and 7).
The refined three-dimensional structures of H162R and R269T are very
similar to the structure of wtPHBH (10) with root mean square
differences of respectively 0.25 and 0.20 Å for 391 equivalent C
atoms. The His-162 and Arg-269 side chains are situated near the
protein surface (Fig. 1) and do not form strong hydrogen bonds with
other residues. This feature is conserved in H162R and R269T, leaving the local structure near the sites of mutation unchanged. However, the
high B factors of the Arg-162 side chain suggest that this side chain
is much more flexible than the His-162 side chain in the wild-type
enzyme.
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GRID Calculations and Model Building--
To get more insight in
the binding mode of the pyrophosphate moiety of NADPH, we searched for
energetically favored binding sites of the "extended" phosphate
groups HPO42 and
PO43
in the wild-type enzyme (10). These
calculations were performed with the program GRID (38). Using a GRID
spacing of 1 Å we observed one large and a few small density peaks
(contoured at an energy level of
15 kcal/mol) in the active site
cleft. By using the graphics program O (36), the pyrophosphate moiety
of NADPH was modeled in the position of an extended GRID peak closely
located to His-162 and Arg-269 (Fig.
8A). Based on this position,
the other parts of the NADPH molecule were modeled in an extended conformation, similar to other NAD(P) complexes (46). In this model,
the cofactor reaches the active site through a cleft between the FAD
binding and substrate binding domains (Fig. 8B). This mode
of interdomain binding is new among known NAD(P)-dependent enzymes (46). The nicotinamide was placed at the re-side
(47) and parallel to the flavin ring (Fig. 8A). Attempts to
place the nicotinamide such that the pro-S hydrogen would be
transferred were unsuccessful due to steric hindrance of Pro-293. In
the pro-R configuration, the nicotinamide ring fitted well,
in full agreement with previous studies (48).
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DISCUSSION |
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In this report we have described the catalytic and structural properties of His-162 and Arg-269 variants of p-hydroxybenzoate hydroxylase from P. fluorescens. The results clearly establish that His-162 and Arg-269 play an important role in NADPH recognition. Flavin spectral analysis and substrate hydroxylation experiments revealed no significant changes in the active site. From this and the structural properties of H162R and R269T it is concluded that the poor catalytic efficiency of the majority of the mutant proteins can be ascribed to impaired NADPH binding.
His-162 is part of a novel conserved sequence motif in flavoprotein hydroxylases with a putative role in FAD and NAD(P)H binding (16). In p-hydroxybenzoate hydroxylase, this sequence motif (residues 153-166) extends from strand A4 to helix H7 and includes a large turn (residues 158-163), with His-162 and the structurally important Asp-159 and Gly-160. His-162 is conserved in p-hydroxybenzoate hydroxylases of known sequence and a positive charged residue is present at this position in other flavoprotein hydroxylases (16). Replacement of His-162 by Ser, Thr, Asn, and Asp results in inefficient flavin reduction due to impaired coenzyme binding. NADPH binding is only moderately affected in the H162R, H162Y, and H162K variants, suggesting that both the bulkiness and hydrogen bonding capacity of residue 162 are of importance in NADPH recognition. Our results agree with a chemical modification study of salicylate hydroxylase, which suggested that Lys-165, the equivalent of His-162 in p-hydroxybenzoate hydroxylase, is involved in binding the pyrophosphate moiety of NADH (51).
At pH 6, H162R interacts much weaker with NADPH than wtPHBH, while at pH 8, the interaction is about the same. Groups with a pKa value in this range are His-162 (free pKa ~ 6.2) and the 2'-phosphate moiety of NADPH (pKa ~ 6.5) (52). A possible explanation could be the following. At pH 6, His-162 is protonated which leads to a stronger interaction with the pyrophosphate moiety of NADPH. However, the 2'-phosphate of NADPH is also protonated which leads to a weaker interaction with Arg-33 and Arg-44 (Fig. 8B). At pH 8, the situation is reversed; His-162 is uncharged which leads to weaker interaction, while deprotonation of the 2'-phosphate of NADPH leads to a stronger interaction. These two effects are compensatory and may explain the nearly constant binding between pH 6 and 8 of NADPH and wtPHBH. For H162R, the situation is different. Arg-162 is protonated at both pH 6 and 8, favoring the interaction with the pyrophosphate of NADPH at both values of pH. At pH 8, also the deprotonated 2'-phosphate of NADPH will contribute to the binding strength. At this pH, the binding of NADPH to H162R is about as strong as with wtPHBH, suggesting that the extra ionic interaction of Arg-162 with the pyrophosphate compensates the intrinsic weaker NADPH binding of H162R. At pH 6, the 2'-phosphate of NADPH gets protonated, leading to weaker interaction not compensated by the creation of an additional ionic interaction elsewhere, which would nicely explain the observed weaker NADPH binding of H162R at pH 6.
Arg-269, located in the substrate binding domain, proved to be even more essential for NADPH recognition. Except for R269K, all Arg-269 variants have lost the ability of proper NADPH binding. This is in agreement with the poor catalytic properties of the R269A isoenzyme from P. fluorescens (53), and points to an electrostatic interaction between Arg-269 and NADPH. Furthermore, the high resolution x-ray structure of R269T shows that impaired NADPH binding is not caused by structural changes.
GRID calculations revealed an energetically favorable binding site for dianionic pyrophosphate near the N terminus of helix H7, supporting our earlier proposal (16) that the positive dipole of this helix is important for binding the pyridine nucleotide cofactor. The properties of the His-162 and Arg-269 variants and the GRID calculations suggest that the pyrophosphate moiety of NADPH interacts with the side chains of His-162 and Arg-269. This interdomain binding mode is in agreement with the involvement of Arg-42 (50) and Arg-44 (22) in NADPH binding, and the role of helix H2 in determining the coenzyme specificity (3).
Recent studies have shown that the flavin ring is mobile and that it can move in and out of the active site (13-15). In our model, we assumed that the flavin is in the out conformation in the NADPH complex. However, while our model fully explains all available mutagenesis and biochemical data, it is still very well possible that the flavin ring assumes an intermediate conformation in the NADPH complex. Further studies will be necessary to firmly establish the position of the flavin in the NADPH complex.
In conclusion, this report lends strong support for an interdomain binding of NADPH in p-hydroxybenzoate hydroxylase. There are only a few "non-Rossmann" fold enzymes of known three-dimensional structure with an interdomain binding mode of the pyridine nucleotide cofactor. These enzymes include isocitrate dehydrogenase from E. coli (54, 55), isopropylmalate dehydrogenase from Thermus thermophilus (56), the ribosome-inactivating protein trichosanthin isolated from root tuber (57), beef liver catalase (58), catalase from Proteus mirabilis (59), and glycogen phosphorylase b (60). In some of these proteins, a high flexibility of the nicotinamide nucleoside moiety is observed. In p-hydroxybenzoate hydroxylase, such flexibility could be essential for the recognition of the flavin ring, leading to the unique effector specificity (8).
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ACKNOWLEDGEMENT |
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We thank Denise Jacobs for assistance in the chemical modification experiments.
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FOOTNOTES |
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* 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.
¶ To whom correspondence should be addressed: Dept. of Biomolecular Sciences, Laboratory of Biochemistry, Wageningen Agricultural University, Dreijenlaan 3, 6703 HA Wageningen, The Netherlands. Tel.: 31-317-482868; Fax: 31-317-484801; E-mail: willem.vanberkel{at}fad.bc.wau.nl.
The abbreviations used are: wtPHBH, wild-type p-hydroxybenzoate hydroxylaseMes, 4-morpholineethanesulfonic acidPOHB, 4-hydroxybenzoateHPLC, high performance liquid chromatography.
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REFERENCES |
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