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J. Biol. Chem., Vol. 281, Issue 26, 17977-17988, June 30, 2006

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Lys-D48 Is Required for Charge Stabilization, Rapid Flavin Reduction, and Internal Electron Transfer in the Catalytic Cycle of Dihydroorotate Dehydrogenase B of Lactococcus lactis^*

Jonathan P. Combe, Jaswir Basran, Parvinder Hothi, David Leys, Stephen E. J. Rigby^¶, Andrew W. Munro^||, and Nigel S. Scrutton¹

From the Manchester Interdisciplinary Biocentre, Faculty of Life Sciences, University of Manchester, Manchester M60 1QD, United Kingdom, the Department of Biochemistry, University of Leicester, Henry Wellcome Building, Lancaster Road, Leicester, LE1 7RH, United Kingdom, the ^¶Department of Biological Sciences, Queen Mary College, University of London, London E1 4NS, United Kingdom, and the ^||Manchester Interdisciplinary Biocentre, School of Chemical Engineering and Analytical Science, University of Manchester, Manchester M60 1QD, United Kingdom

Received for publication, February 14, 2006 , and in revised form, April 3, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Dihydroorotate dehydrogenase B (DHODB) catalyzes the oxidation of dihydroorotate (DHO) to orotate and is found in the pyrimidine biosynthetic pathway. The Lactococcus lactis enzyme is a dimer of heterodimers containing FMN, FAD, and a 2Fe-2S center. Lys-D48 is found in the catalytic subunit and its side-chain adopts different positions, influenced by ligand binding. Based on crystal structures of DHODB in the presence and absence of orotate, we hypothesized that Lys-D48 has a role in facilitating electron transfer in DHODB, specifically in stabilizing negative charge in the reduced FMN isoalloxazine ring. We show that mutagenesis of Lys-D48 to an alanine, arginine, glutamine, or glutamate residue (mutants K38A, K48R, K48Q, and K48E) impairs catalytic turnover substantially (50–500-fold reduction in turnover number). Stopped-flow studies demonstrate that loss of catalytic activity is attributed to poor rates of FMN reduction by substrate. Mutation also impairs electron transfer from the 2Fe-2S center to FMN. Addition of methylamine leads to partial rescue of flavin reduction activity. Nicotinamide coenzyme oxidation and reduction at the distal FAD site is unaffected by the mutations. Formation of the spin-interacting state between the FMN semiquinone-reduced 2Fe-2S centers observed in wild-type enzyme is retained in the mutant proteins, consistent with there being little perturbation of the superexchange paths that contribute to the efficiency of electron transfer between these cofactors. Our data suggest a key charge-stabilizing role for Lys-D48 during reduction of FMN by dihydroorotate, or by electron transfer from the 2Fe-2S center, and establish a common mechanism of FMN reduction in the single FMN-containing A-type and the complex multicenter B-type DHOD enzymes.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The dihydroorotate dehydrogenases (DHOD)² are flavoproteins that participate in the de novo biosynthesis of pyrimidines. They are a heterogeneous family of enzymes that catalyze the conversion of dihydroorotate to orotate (Scheme 1) and the transfer of electrons to various redox acceptors (1). Class 1A DHODs (2, 3) are soluble dimeric enzymes, contain a FMN prosthetic group and are able to use fumarate as an electron acceptor. They are found in anaerobic yeasts, milk-fermenting bacteria, and some protozoa. The soluble class 1B enzymes, which are found in Gram-positive bacteria, contain FMN, FAD, and a 2Fe-2S center and use NAD⁺ as electron acceptor (4, 5). Lactococcus lactis and related milk-fermenting bacteria like Enterococcus faecalis are unusual in possessing two DHOD enzymes (i.e. DHODA (class 1A) and DHODB (class 1B)). The class 2 enzymes, which are found in eukaryotes and Gram-negative bacteria, are membrane-bound, contain FMN and are oxidized by ubiquinone (6–8).

The multicenter DHODB (EC 1.3.3.1 [EC] ) enzymes have been purified from a number of organisms including L. lactis, Bacillus subtilis, E. faecalis, and Clostridium oroticum (4, 9–11). A crystallographic structure for L. lactis DHODB is available (5). The enzyme is a dimer of heterodimers, with a catalytic subunit (termed subunit D) responsible for the oxidation of dihydroorotate to orotate and reduction of the FMN cofactor. Electron transfer subunits (termed K subunits) are involved in electron transfer and the binding/reduction of the electron acceptor NAD⁺. The K subunits bind a 2Fe-2S center and FAD, and the electron transfer pathway is from FMN (subunit D) to 2Fe-2S center (subunit K) to FAD (subunit K) and NAD⁺. Substrate binding in the active site leads to conformational changes that occur in specific regions of the catalytic D-subunit (5). Notable among these are the relatively large movements for the side chains of Lys-D48 and Tyr-K232.

Kinetic isotope effect studies and analysis of the pH dependence of the reaction catalyzed by E. faecalis DHODB indicate a reversible reaction. Rate-limiting macroscopic ionizations have been identified in the reductive and oxidative half-reactions (11). Studies of double isotope effects on ^DV and ^D(V/K_DHO) obtained for deuteration at C₅-proS and C₆ of dihydroorotate have been used to propose a mechanism in which C₅-proS proton transfer and C₆-hydride transfer are concerted. This mode of oxidation of dihydroorotate prevents formation of an unstable carbanion intermediate (pK_a 20–21, Ref. 10). Active site cysteine and lysine residues have been proposed to act as a general base and electrostatic catalyst in C. oroticum DHODB (10).

We have reported reductive titrations of DHODB from L. lactis and demonstrated a five electron capacity (12). The mid-point reduction potential of the 2Fe-2S center (–212 mV ± 3 mV), the oxidized/semiquinone (E₁) and semiquinone/hydroquinone (E₂) couples for the FMN (E₁ =–301 ± 6mV; E₂ =–252 ± 8 mV) and FAD (E₁ =–312 ± 6mV; E₂ =–297 ± 5 mV) are known from UV-visible spectroelectrochemical titration and EPR studies. A spin-interacting state between the FMN semiquinone species and the reduced 2Fe-2S center forms during reductive titration with dithionite. This spin-interacting state is characterized by an unusual EPR signal with very small rhombic anisotropy and g values 2.02, 1.99, and 1.96 (12). To begin to address the mechanism of flavin reduction and internal electron transfer in DHODB we have performed a series of stopped-flow studies of the wild-type L. lactis enzyme and mutants altered in the region of Lys-D48, which is located in the FMN-binding region. Herein, we report the first detailed kinetic characterization of DHODB using stopped-flow methods and demonstrate a key role for Lys-D48 in FMN reduction and internal electron transfer from the 2Fe-2S center to FMN. Our studies are consistent with a charge-stabilization role for Lys-D48 during electron transfer rather than the result of major perturbation of the reduction potential of the redox cofactors, or disruption of superexchange pathways that contribute to the efficiency of electron transfer between redox cofactors.

View larger version (5K): [in this window] [in a new window]   SCHEME 1

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Steady-state Kinetic Assays—DHOD activity was measured by monitoring the formation of orotate using a lactate dehydrogenase-coupled assay (10). The method used, which is a modified version of the published procedure, involved incubating freshly purified enzyme with 50 mM cysteine at –20 °C for at least 10 days prior to carrying out steady-state assays. This procedure abolished the cysteine-dependent lag phase observed in the steady-state kinetic traces of wild-type DHOD and thus negated the need to incubate the reaction mixture with additional cysteine for 20 min prior to measurement of enzyme activity (see supplemental data, Figs. S1 and S2). The steady-state activities of wild-type and mutant DHOD enzymes were determined using a Hewlett-Packard 8452A single beam diode array spectrometer with a 1-cm light path. Reactions were performed at 25 °C in 50 mM potassium phosphate/50 mM bis-Tris propane buffer pH 8.0, containing 1 mM EDTA, 2 mM pyruvate, 0.02 units/ml rabbit muscle L-lactate dehydrogenase (LDH), 900 µM NAD⁺ and a fixed concentration of DHOD. The reaction was initiated by the addition of L-dihydroorotate and initial rates were calculated from the absorbance increase at 290 nm (₂₉₀ = 5960 M^–1 cm^–1). One unit of activity is defined as the amount of enzyme that catalyzes the production of 1 µmol of orotate per second. Apparent K_m and k_cat values for L-dihydroorotate were determined at a fixed and saturating concentration of NAD⁺ (900 µM). The data were fitted to the Michaelis-Menten equation using the Grafit software package (14).

DHODB activity using either NADH/orotate or NADH/molecular oxygen as substrates was measured by monitoring the change in absorption at 340 nm that accompanies oxidation of NADH. Reactions were performed at 25 °C in 50 mM potassium phosphate/50 mM bis-Tris propane buffer pH 8.0 in a total volume of 1 ml. Steady-state kinetic measurements were performed using a Jasco V530 UV/VIS spectrophotometer with a 1-cm light path. Assays in which orotate and NADH were used as substrates were carried out under strict anaerobic conditions (<5 ppm O₂) to prevent the oxidase activity of DHODB. This was achieved by housing the spectrophotometer within a glove box (Belle Technology, Dorset, UK). The apparent k_cat and K_m values for orotate were determined at a constant and saturating NADH concentration (140 µM) and the apparent k_cat and K_m values for NADH were determined at a constant (and saturating) orotate concentration of 200 µM. DHODB activity using NADH and molecular oxygen was measured aerobically; the apparent k_cat and K_m values for NADH were measured at atmospheric concentrations of oxygen (258 µM). Initial rates of reaction were calculated from changes in absorption at 340 nm using an extinction coefficient of 6220 M^–1 cm^–1 (oxidation of NADH) and fitted to the Michaelis-Menten equation using the Grafit software package (14).

Preparation of Anaerobic Samples—Buffer (50 mM potassium phosphate/50 mM bis-Tris propane, pH 8.0) was made anaerobic by bubbling argon gas through it for >2 h. Solutions were then placed in an anaerobic glove box (Belle Technology) overnight to remove any residual traces of oxygen. Protein samples were made anaerobic by passing through a small gel filtration (Bio-Rad 10 DG) column (housed in the glove box), which had been pre-equilibrated with anaerobic buffer. Substrate and coenzyme solutions were made by adding the appropriate solid to anaerobic buffer.

Stopped-flow Kinetic Measurements—Unless stated otherwise, all kinetic studies were carried out under strict anaerobic conditions (<5 ppm O₂) within a glove box environment (Belle Technology). Experiments were performed in 50 mM potassium phosphate/50 mM bis-Tris propane buffer, pH 8.0, at the stated temperatures. Rapid reaction kinetic experiments were performed using an Applied Photophysics SX.18MV-R stopped-flow spectrophotometer contained within the glove box. Stopped-flow, multiple-wavelength absorption studies were carried out using a photodiode array detector and X-SCAN software (Applied Photophysics Ltd). Spectral deconvolution was performed by global analysis and numerical integration methods using PROKIN software (Applied Photophysics). For single-wavelength studies, absorption and fluorescence data collected were analyzed using nonlinear least squares regression analysis on an Acorn Risc PC microcomputer using Spectrakinetics software (Applied Photophysics). In the reductive halfreaction, experiments were performed by mixing enzyme in the appropriate buffer with an equal volume of reducing substrate/cofactor in the same buffer at the desired concentration. The concentration of substrate was always at least 10-fold greater (5-fold for fluorescence measurements) than the enzyme concentration, thereby ensuring pseudo first-order conditions. For each substrate concentration, at least four replica measurements were collected and averaged. NADH fluorescence experiments were carried out using an excitation wavelength of 340 nm and a 450 nm emission band-pass filter (g35–3367; from Coherent Optics, Greycain Road, Watford, Herts., UK).

In single wavelength studies of wild-type DHODB, reduction by DHO was observed at 452 nm. Under pseudo first-order conditions, transients were triphasic in nature and were fitted using the standard triple exponential expression (Equation 1),

(Eq. 1)

where k_obs1, k_obs2, and k_obs3 are the observed rate constants for the fast, intermediate, and slow phases respectively, C₁, C₂, and C₃ are the relative amplitude values for the three phases, and b is an offset value to account for a non-zero baseline. Observed rate constants for absorption changes occurring in the reduction of DHODB by NADH were obtained from fits of the data to a standard double-exponential expression. NADH fluorescence transients were monophasic and analyzed using a standard single exponential expression. Analysis of the concentration dependence of flavin reduction rate constants was accomplished by fitting plots of observed rate constant versus substrate concentration to a hyperbolic dependence. Derived rate constants are attributable to resolved kinetic phases, and do not necessarily report on a single microscopic step in the reaction mechanism.

Time-dependent spectral changes that occur during the course of steady-state turnover of wild-type and mutant DHODBs were analyzed using the enzyme monitored turnover method (15) and multiple wavelength stopped-flow spectroscopy. Reactions were performed using a photodiode array detector and X-SCAN software interfaced with an Applied Photophysics SX18MV-R reaction analyzer. Solution conditions are described under "Results."

EPR Spectroscopy—During redox titrations with dithionite (see above), samples (200 µl) of DHODB (32.5 µM heterodimer) were withdrawn for EPR spectroscopic analyses. The samples were placed in standard 3-mm quartz EPR tubes, and sealed with parafilm inside the glove box. Immediately after removing tubes from the glove box they were placed into a dry ice/ethanol bath to freeze the samples, and then transferred into liquid nitrogen. Samples were stored in liquid nitrogen until analyzed. The EPR spectra were recorded on a Bruker ELEXSYS E500 spectrometer fitted with an Oxford Instruments ESR900 liquid helium cryostat. All samples were contained in 100 mM potassium phosphate buffer, pH 7.0. Quantitation of the paramagnetic species formed was achieved using double integration of the appropriate spectra or difference spectra as previously described in Ref. 12. Quantitative analysis of reduction potentials was achieved by UV-visible potentiometric analysis using the method of Dutton (16), as described previously for studies with wild-type DHODB (12).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Lys-D48, Conformational Mobility, and Interactions with the FMN Isoalloxazine and Tyr-K232/2Fe-2S Center in Native and Orotate-bound Enzyme—Crystal structures of native and orotate-bound L. lactis DHODB have indicated differences between the two structures that potentially have major implications for the mechanism of flavin reduction and internal electron transfer (5). These structural differences involve (i) a flexible loop that contains the catalytic cysteine residue that is proposed to abstract a proton from the C5 atom of dihydroorotate, (ii) a re-orientation of Lys-D170 so that it hydrogen bonds to the FMN in the orotate-bound structure, and (iii) a re-orientation of Met-D247 so that the thioether group interacts with the FMN ribityl O3 and Lys-D170 (Fig. 1). Residue Lys-D48, however, undergoes the largest structural change. In the structure of the native enzyme, the Lys-D48 side chain is positioned away from the FMN. In this conformation, the N atom of Lys-D48 is located close to the C1 and C1 atoms of Tyr-K232. Additionally, the N atoms of Lys-K247 and Lys-D170 are also positioned close to Tyr-K232 (Fig. 1B). The binding of orotate to the active site of DHODB brings about a re-positioning of Lys-D48 such that the side chain now makes hydrogen bonding interactions with the N5 and O4 atoms of the FMN isoalloxazine ring. Moreover, the carboxyl group of orotate forms a hydrogen bond with the side chain of Lys-D48, suggesting this interaction is important in binding of orotate to the enzyme. In this conformation, Lys-D48 occupies a position similar to that of the equivalent lysine residue found in L. lactis DHODA (3, 17). We hypothesized that in the orotate-bound form of DHODB, the hydrogen bonds formed between the N group of Lys-D48 and the FMN N5 and O4 atoms might modulate the reductive half-reaction chemistry through a variety of effects. These might include (i) alteration of the electronic properties of the FMN, (ii) an influence on the geometry of substrate binding, (iii) charge stabilization of the FMN (at the O4 position) during FMN reduction, and (iv) possible disruption of the degree of electronic coupling between the FMN and 2Fe-2S center which might affect the rate of electron transfer from FMN to the 2Fe-2S center. The implications of this conformational mobility for mechanism are unclear, and for this reason we have investigated the importance of this Lys-D48 in the catalytic cycle by directed mutagenesis and analysis of the electron transfer properties of four mutant forms of the enzyme (K48A, K48Q, K48R, and K48E) designed to alter the charge and size of the side chain at position D48. Each mutant enzyme was purified as described previously for the wild-type enzyme and was assembled stoichiometrically with FAD, FMN, and the 2Fe-2S center. The spectral properties of the purified mutant proteins were essentially identical to those of the wild-type enzyme (12). The electronic absorption spectra of the oxidized forms of wild-type and all K48 mutant DHODB enzymes had absorption maxima at 453 nm and 376 nm. The absorption shoulder at 540 nm from the iron-sulfur cluster is also observed in all oxidized forms (12). Initially, we studied in detail, by stopped-flow and steady-state methods, the kinetics of flavin reduction and electron transfer in wild-type DHODB. These studies are presented below and form a framework for comparison with the mutant enzymes (also reported below).

View larger version (27K): [in this window] [in a new window]   FIGURE 1. The environment of the FMN in the D subunit of DHODB. A, the FMN environment in native enzyme (PDB accession code 1EP1). B, the FMN environment in orotate-bound enzyme (PDB accession code 1EP2). C, a close up view of the environment of the FMN and 2Fe-2S center in DHODB illustrating the interactions between Lys-D48 and the FMN isoalloxazine ring (orotate-bound enzyme) and Lys-D48 and Tyr-K232 (native enzyme). Oxygen atoms of active site water molecules are displayed as gray spheres.

To obtain values of rate constants for each kinetic phase as a function of NADH concentration, we performed stopped-flow studies with detection at a single wavelength or by fluorescence detection. Absorption changes monitored at 452 nm following rapid mixing of DHODB with NADH were biphasic, consistent with the photodiode array data (Fig. 3A). The concentration dependence of both kinetic phases was investigated under pseudo first-order conditions at this wavelength (Fig. 3B). The first kinetic phase showed a linear dependence on NADH concentration with rate constant 6.9 ± 0.1 x 10⁶ M^–1 s^–1, which represents reduction of the FAD cofactor of DHODB by NADH. The second kinetic phase was essentially independent of NADH concentration over the range 10–35 µM, and probably reflects internal electron transfer to the 2Fe-2S center and FMN and/or loss of NAD⁺. Above 40 µM the apparent independence was lost (Fig. 3B), but the mechanistic origin for this is uncertain. Also, the relatively small amplitude changes associated with the slow phase (Fig. 2C) complicates any attempt to rigorously analyze the concentration dependence of this phase. Rapid mixing studies of DHODB with NADH using fluorescence detection (excitation 340 nm, emission 450 nm) to monitor coenzyme oxidation produced monophasic reaction transients (Fig. 3C). Observed rate constants calculated from these fluorescence transients were essentially the same as those measured in the fast phase of the absorption transients (Fig. 3A). These data confirm that the first kinetic phase observed in the absorption transients is attributable to coenzyme oxidation.

View larger version (19K): [in this window] [in a new window]   FIGURE 2. Reduction of DHODB by NADH. A, time-dependent spectral changes that occur on mixing 4.5 µM wild-type DHODB with 4.5 µM NADH. Reaction conditions: 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0; 4 °C. The time course shown is 1 s after the initial mixing event. For clarity, only selected spectra are shown. B, deconvoluted spectra for the reaction shown in A. The data were fitted to a two-step model A B C. The rate constants (s–1; obtained from global fitting) are A B, 52 ± 0.7; B C, 13 ± 0.1. C, deconvoluted spectra for the reaction of DHODB with a 10-fold excess (45 µM) of NADH. The data were fitted to a two-step model A B C. The rate constants (s–1; obtained from global fitting) are A B, 273 ± 2; B C, 2.3 ± 0.2.

View larger version (25K): [in this window] [in a new window]   FIGURE 3. Dependence of the observed rates of flavin reduction/oxidation on NADH/NAD+ concentration for wild-type DHODB. Reaction conditions: 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0; 4 °C. A, absorption transient at 452 nm observed on mixing wild-type DHODB (2.3 µM) with NADH (30 µM). Transient is fitted to a standard biphasic expression (rate constants, 196 s–1 and 8.7 s–1). Inset, plot of residuals from the fit to a biphasic rate expression. B, plot illustrating the NADH concentration dependence of the rate for the fast (filled circles) and slow (open circles) phases of the absorption change observed at 452 nm and for the fluorescence change (monitored at 450 nm; open squares) with wild-type DHODB. C, fluorescence transient (excitation 340 nm, emission 450 nm) observed on mixing wild-type DHODB (5 µM) with NADH (30 µM). Transient is analyzed using a standard single exponential expression (rate constant, 172 s–1). Inset, plot of residuals from the fit to a monophasic rate expression. D, absorption transient at 452 nm observed on mixing wild-type 5 electron-reduced DHODB (2.3 µM) with NAD+ (400 µM). Transient is fitted to a standard biphasic expression (rate constants, 30.6 s–1 and 4.8 s–1). Inset, plot of residuals from the fit to a biphasic rate expression. E, plot illustrating the NAD+ concentration dependence of the rate for the fast (filled circles) and slow (open circles) phases of the absorption change observed at 452 nm on mixing 5 electron-reduced DHODB with NAD+.

View larger version (30K): [in this window] [in a new window]   FIGURE 4. Reduction of DHODB by DHO. A, time-dependent spectral changes that occur on mixing 4.5 µM wild-type DHODB with 4.5 µM DHO. Reaction conditions: 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0; 4 °C. Data were acquired over a 1-s period after the initial mixing event. For clarity, only selected spectra are shown. B, deconvoluted spectra for the reaction shown in A. The data were fitted to a two-step model A B C. The rate constants (s–1; obtained from global fitting) are A B, 12 ± 0.1; B C, 4.7 ± 0.1. C, time-dependent spectral changes that occur on mixing 4.5 µM wild-type DHODB with 1 mM DHO. Reaction conditions: as for A. Data were acquired over a 0.5-s period after the initial mixing event. For clarity, only selected spectra are shown. D, deconvoluted spectra for the reaction shown in C. The data were fitted to a three-step model A B C 3 D. The rate constants (s–1; obtained from global fitting) are A B, 251 ± 2; B C, 28 ± 0.1; C D; 6 ± 0.1.

The concentration dependence of the three kinetic phases was investigated in single wavelength stopped-flow studies at 452 nm. The first kinetic phase shows a hyperbolic dependence on dihydroorotate concentration (Fig. 5A). Fitting to plots of observed rate constant versus DHO concentration using a hyperbolic expression produced a dissociation constant for the complex of oxidized enzyme and dihydroorotate of 244 ± 9 µM and a limiting rate constant for flavin reduction of 522 ± 8s^–1. The second (B C, 22 s^–1) and third (C D, 1.6 s^–1) kinetic phases are essentially independent of substrate concentration in the pseudo first-order regime and are attributed to internal electron transfer and further reduction of the enzyme by DHO (Fig. 5B). The lack of a dependence of the rate constants for these two phases on DHO concentration is consistent with the intrinsic rate of FMN reduction by substrate being faster than the rate of internal electron transfer within DHODB.³

Steady-state Reactions of Wild-type and Mutant Enzymes—Replacement of Lys-D48 in DHODB with Glu, Arg, Gln, or Ala has very little effect on the apparent Michaelis constant for the substrate (Table 1). At most, a 4-fold increase in K_m was observed for the K48E mutant when compared with the wild-type enzyme. Mutation of Lys-D48 in DHODB has a substantial effect, however, on the catalytic activity of the enzyme with an almost 500-fold decrease in the turnover number for K48A DHOD compared with wild-type DHODB. This indicates a key role for Lys-D48 in the catalytic cycle of DHODB.

View larger version (15K): [in this window] [in a new window]   FIGURE 5. Concentration dependence of observed rate constants for each kinetic phase in the reaction of DHODB with DHO. Main panel, plot illustrating the DHO concentration dependence of the rate (kfast) of the fast phase observed at 452 nm for wild-type DHODB. The limiting rate for flavin reduction for the fast phase (estimated using the Strickland equation) was calculated to be 522 ± 8 s–1 and the dissociation constant for the oxidized enzyme-dihydroorotate complex is 244 ± 9 µM. Inset, concentration dependence of the rate for the intermediate (open circles) and slow (closed circles) phases of the triphasic transients observed at 452 nm. Reaction conditions: 50 mM bis-Tris propane/50 mM phosphate buffer pH 8.0; 4 °C. Enzyme concentration, 2.3 µM.

View larger version (23K): [in this window] [in a new window]   FIGURE 6. Time-dependent spectral changes for the reaction of K48R DHODB with DHO. A, time-dependent spectral changes that occur on mixing 4.5 µM K48R DHODB with 1 mM DHO. Reaction conditions: 50 mM bis-Trispropane/50mM phosphatebuffer, pH 8.0; 25 °C. The experiment was performed over 50 s after the initial mixing event. For clarity, only selected spectra are shown. B, deconvoluted spectra for the reaction shown in A. The data were fitted to a three-step model A B C D. The rate constants (s–1; obtained from global fitting) are A B, 1.24 ± 0.01; B C, 0.18 ± 0.01; C D, 0.014 ± 0.001.

View larger version (23K): [in this window] [in a new window]   FIGURE 7. Enzyme-monitored turnover of wild-type DHODB and the K48Q mutant with NADH as reducing substrate. A, 4 µM wild-type DHODB was mixed with 500 µM NADH and 65 µM O2 in the stopped-flow apparatus and spectra recorded with a photodiode detector at the start of the reaction (spectrum 1), during the steady-state phase (spectrum 2), and at the end of the reaction (spectrum 3). B, reaction conditions as for A but with 4 µM K48Q DHODB. C, 4 µM wild-type DHODB was rapidly mixed with 500 µM NADH and 50 µM orotate and spectra recorded at the start of the reaction (spectrum 1), during the steady-state phase (spectrum 2) and at the end of the reaction (spectrum 3). D, reaction conditions as for C but with 4 µM K48Q DHODB. All experiments were carried out under anaerobic conditions in 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0, at 25 °C.

Flavin Reduction Levels during Steady-state Turnover of Wild-type and Mutant Enzymes Revealed by Enzyme-monitored Turnover—We also probed steady-state turnover in the wild-type and K48Q enzymes using the enzyme monitored turnover method to assess the effects of mutating Lys-D48 on the reduction level of the enzyme. Profiles from enzyme monitored turnover studies using (i) NADH and oxygen and (ii) NADH and orotate as substrates with wild-type and K48Q enzyme are shown in Fig. 7. With wild-type enzyme, the steady-state spectrum (spectrum 2) indicates that the enzyme is essentially fully reduced during turnover with oxygen (Fig. 7A) or orotate (Fig. 7C) as electron acceptors, indicating that enzyme reduction and internal electron transfer within wild-type enzyme are fast relative to reoxidation of the cofactors by orotate or oxygen. Comparable studies with the K48Q enzyme indicate that the flavins are approximately only half reduced during steady-state turnover (Fig. 7, B and D), suggesting more of a balance between the rates of reductive and oxidative processes. The FAD reduction kinetics of the K48Q mutant are essentially unaffected by mutation as demonstrated by stopped-flow mixing studies of FAD reduction by NADH (Fig. 8). That the flavin reduction level during steady-state turnover is less in the mutant enzyme compared with wild-type suggests a bottleneck on the rate of electron transfer from the 2Fe-2S center to FMN. The steady-state spectrum, therefore, predominantly represents reduced FAD and oxidized FMN.

View larger version (11K): [in this window] [in a new window]   FIGURE 8. Plot illustrating the NADH concentration dependence of the rate for the fast phase of the absorption change observed at 452 nm for wild-type DHODB (filled circles) and the K48Q mutant (open squares). Reaction conditions: 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0; 4 °C.

View larger version (30K): [in this window] [in a new window]   FIGURE 9. Enzyme-monitored turnover of wild-type DHODB and the K48Q mutant with DHO as reducing substrate. A, 4 µM wild-type DHODB was mixed with 1 mM DHO and 65 µM O2 in the stopped-flow apparatus and spectra recorded with a photodiode detector at the start of the reaction (spectrum 1), during the steady-state phase (spectrum 2), and at the end of the reaction (spectrum 3). B, reaction conditions as for A but with 4 µM K48Q DHODB. Inset, time course of the absorption change monitored at 452 nm for the reaction shown in B. C, 4 µM wild-type DHODB was rapidly mixed with 1 mM DHO and 50 µM NAD+ and spectra recorded at the start of the reaction (spectrum 1), during the steady-state phase (spectrum 2) and at the end of the reaction (spectrum 3). D, reaction conditions as for C but with 4 µM K48Q DHODB. Inset, time course of the absorption change monitored at 452 nm for the reaction shown in D. All experiments were carried out in 50 mM bis-Tris propane/50 mM phosphate buffer, pH 8.0, at 25 °C.

View larger version (34K): [in this window] [in a new window]   FIGURE 10. A, EPR spectra of mutant (K48Q) DHODB reduced with the number of electron equivalents of sodium dithionite as indicated. The features marked * arise from the 2Fe-2S cluster and that marked arises from the spin-coupled signal (see text, and Ref. 12). Parameters: microwave power 0.2 milliwatt, modulation amplitude 5G, modulation frequency 100 KHz, temperature 15 K. B, plot of relative integral of the spin coupled signal against electron equivalents added. C, plot of relative integral of the flavosemiquinone signal recorded at high temperature (spectra not shown; parameters: microwave power 10 microwatts, modulation amplitude 1G, modulation frequency 100 KHz, temperature 120 K) against electron equivalents added. In B and C, , K48R; •, K48Q; , K48A; , K48E.

View larger version (15K): [in this window] [in a new window]   FIGURE 11. Molecular graphics representation of the change in side-chain location for Lys-D48 and interactions formed on binding dihydroorotate. The arrows indicate the flow of electrons following proton abstraction from the C5 atom of dihydroorotate by Cys-D135. In this scheme, proton transfer is shown from the -amino group of Lys-D48 to the O4 atom of the FMN isoalloxazine ring (i.e. the proposed charge neutralization scheme). A and B represent crystallographically defined water molecules. The figure was produced using PyMol software (23).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Our kinetic data demonstrate that FMN reduction by (i) dihydroorotate, or (ii) electron transfer from the 2Fe-2S, is impeded in the different Lys-D48 mutant enzymes. The change in the position of the Lys-D48 side-chain on binding orotate in the active site positions the -amino group of the side chain within hydrogen bonding distance of the FMN O4 atom, the substrate carboxyl group and a water molecule (Fig. 11). Additionally, the -amino group of Lys-D170 is in close proximity to the FMN N10 atom. The immediate environment of Lys-D170 suggests hydrogen bonds form to a ribityl alcohol group, the FMN O₂ atom and a water molecule, which might have a role in stabilization of the reduced FMN, although the geometry is less favorable for this role compared with that observed for Lys-D48. The location of Lys-D48 in orotate-bound DHODB suggests that neutralization of the semi- or dihydroquinone FMN is possible through formal proton transfer to the FMN O4 atom by the -amino group of Lys-D48 (Fig. 11). The compromised kinetics of FMN reduction by dihydroorotate (Table 3), and reverse electron transfer from the 2Fe-2S center (Fig. 7, B and D), are consistent with such a role for Lys-D48. Moreover, the partial rescue of FMN reduction activity observed on the addition of methylamine to the mutant DHODB enzymes (Fig. S3, supplemental data) is also consistent with this proposed role for Lys-D48.

We were surprised to find that our EPR (Fig. 10) and potentiometry studies (supplemental data Fig. S4) indicate mutation of Lys-D48 does not perturb substantially the midpoint reduction potentials of the flavin and 2Fe-2S cofactors. Given the proposed role of Lys-D48 in charge neutralization (through formal proton transfer to the FMN O4 atom) one might expect the midpoint potentials of the FMN to be considerably perturbed as a result of mutation. We emphasize, however, that both the potentiometry and EPR measurements were made with enzyme reduced by dithionite and not the substrate dihydroorotate. At this stage, it is not known if reduction of the cofactors by dithionite is sufficient to recruit the side chain of Lys-D48 to the position observed in the crystal structure of the DHODB-orotate complex, or whether additional interactions (e.g. the interaction of Lys-D48 with the carboxylate group of dihydroorotate) are also required to promote this conformational change. If reduction by dithionite is unable to recruit Lys-D48 to the position seen in the enzyme-orotate complex, then the environment of the FMN in both wild-type and mutant forms of DHODB will be similar in our EPR and potentiometric titrations. In this case, one might expect only minor perturbation of the midpoint potentials of the FMN and 2Fe-2S centers, consistent with our experimental observations. We suggest that if charge neutralization of the O4 atom requires proton transfer, and in a situation where reduction by dithionite does not recruit Lys-D48 to fulfill this role, it is likely this proton could be recruited from solvent under equilibrium titration conditions. In a stopped-flow turnover situation using dihydroorotate as substrate, we expect this proton transfer to be from Lys-D48 in wild-type enzyme. In the mutant proteins, however, this proton transfer would need to be from solvent and this could account for the slow FMN reduction rates observed with the mutant enzymes.

The retention of the spin-interacting state in the mutant DHODB enzymes indicates clearly that superexchange pathways between the FMN_sq and reduced 2Fe-2S center are essentially unperturbed. We infer, therefore, that the slow rates of electron transfer from the 2Fe-2S center to FMN, as observed in our enzyme monitored turnover studies (Fig. 7, B and D), are not attributed to weaker electronic coupling between these two centers. This is consistent with the robust nature of electronic coupling in proteins over relatively short distances, which buffers against adverse affects on the coupling matrix element following mutation of an electron tunneling pathway (21).

Studies with the class 2, membrane-associated human DHOD have also implied a major role for an active site lysine residue in charge stabilization during reduction of the enzyme-bound FMN by dihydroorotate (22). In this case, the redox chemistry involves only a single FMN cofactor, as electron transfer is from substrate to FMN and then to ubiquinone. The crystal structure of the class 1A DHOD enzyme from L. lactis also reveals an active site lysine residue that is appropriately positioned to assist in charge stabilization/protonation in the reductive half-reaction. Our studies with the more complex class 1B enzyme have established similarities in the requirement for an active site lysine residue to stabilize or neutralize negative charge in the FMN isoalloxazine ring, and thus emphasize a common mechanism for facilitating rapid electron transfer from substrate to the FMN cofactor in both soluble and membrane-bound DHOD enzymes.

Concluding Remarks—Lys-D48 is required to maintain rapid rates of FMN reduction and internal electron transfer in DHODB through a mechanism that involves electrostatic stabilization of, or charge neutralization by proton transfer to, the O4 atom of the FMN isoalloxazine ring. Mutation of Lys-D48 does not affect electronic coupling between the 2Fe-2S and FMN centers as superexchange pathways are maintained in mutant enzymes, giving rise to the characteristic spin-interacting state seen in wild-type enzyme. Our study establishes a common mechanism for enhancing electron transfer in both soluble (class 1B) and membrane-bound DHOD (class 2) enzymes and emphasizes the need to stabilize the reduced form of the flavin isoalloxazine ring in flavoprotein catalysis.

	FOOTNOTES

 
^* The work was funded by the UK Biotechnology and Biological Sciences Research Council and the Royal Society. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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

¹ To whom correspondence should be addressed: Manchester Interdisciplinary Biocentre and Faculty of Life Sciences, University of Manchester, Jackson's Mill, PO Box 88, Sackville St., Manchester M60 1QD, United Kingdom. Tel.: 0161-306-5152; E-mail: nigel.scrutton{at}manchester.ac.uk.

² The abbreviations used are: DHOD, dihydroorotate dehydrogenase; DHODB, dihydroorotate dehydrogenase B; sq, semiquinone; hq, hydroquinone; ox, oxidized; SHE, standard hydrogen electrode.

³ Alternatively, some step in the course of flavin reduction subsequent to formation of the Michaelis complex (e.g. product release) could also rate-limit electron transfer and account for the lack of dependence of these rate constants on DHO concentration.

⁴ Given the slow rate constant for the fast phase in the mutant enzymes, it was not possible to measure enzyme-substrate dissociation constants using stopped-flow methods because the first and second kinetic phases could not be resolved at lower substrate concentrations.

⁵ We emphasize that EPR studies are performed at low temperature unlike kinetic and potentiometric measurements (using spectroelectrochemical methods). Also, in kinetic studies we cannot rule out the possibility that charge-transfer states might contribute to long wavelength absorption. For these reasons, a comparison and rationalization of the degree of semiquinone stabilization using these different methods is unreliable.

	ACKNOWLEDGMENTS

 
We thank Dr K. F. Jensen for the gift of plasmid pFN3.

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TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

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