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The flavoenzyme nikD, a 2-electron acceptor, catalyzes a remarkable aromatization of piperideine-2-carboxylate (P2C) to picolinate, an essential component of nikkomycin antibiotics. Steady-state kinetic data are indicative of a sequential mechanism where oxygen reacts with a reduced enzyme•dihydropicolinate (DHP) complex. The kinetics observed for complex formation with competitive inhibitors are consistent with a one-step binding mechanism. The anaerobic reaction with P2C involves three steps. The first step yields an enzyme•substrate charge transfer complex likely to contain the electron-rich P2C enamine. Calculated rates of formation and dissociation of the nikD•P2C complex are similar to those observed for the enzyme•1-cyclohexenoate complex. Formation of a reduced enzyme•DHP complex, (EH2•DHP)ini, occurs in a second step that exhibits a hyperbolic dependence on substrate concentration. The limiting rate of nikD reduction is at least 10-fold faster than the turnover rate observed with unlabeled or [4, 4, 5, 5, 6, 6-D6]-P2C and exhibits a kinetic isotope effect (KIE = 6.4). The observed KIE on Kd apparent (4.7) indicates that P2C is a sticky substrate. Formation of a final reduced species, (EH2•DHP)fin, occurs in a third step that is independent of P2C concentration and equal to the observed turnover rate. The observed KIE (3.3) indicates that the final step involves cleavage of at least one C-H bond. Tautomerization, followed by isomerization, of the initial DHP intermediate can produce an isomer that could be oxidized to picolinate in a reaction that satisfies known steric constraints of flavoenzyme reactions without the need to reposition a covalently tethered flavin or tightly bound intermediate.
NikD is a flavoprotein oxidase that plays an essential role in the biosynthesis of nikkomycins. Nikkomycins comprise a group of related peptidyl nucleoside antibiotics that resemble the natural substrate of chitin synthase. Chitin, the second most abundant polysaccharide in nature, is an integral component of the cell wall in fungi and the exoskeleton of invertebrates but is not found in mammals. Nikkomycins are potent antifungal agents that act by competitively inhibiting chitin synthase. Nikkomycins are therapeutically effective in treating human fungal infections that are especially prevalent in immunocompromised patients and are also useful in agriculture as easily degraded insecticides that are nontoxic for mammals (1).
The nikkomycin peptide is synthesized by a nonribosomal pathway and contains an N-terminal pyridyl moiety, derived from L-lysine, that is essential for antibiotic function. Synthesis of the pyridyl moiety is initiated by an α-aminotransferase that converts L-lysine to piperideine-2-carboxylate (P2C) (2), a compound that can exist in imine and enamine tautomeric forms (3–5). Picolinate is produced in a remarkable aromatization reaction catalyzed by nikD, involving an overall 4-electron oxidation of P2C (Scheme 1) (6). NikD contains covalently bound FAD (8α-S-cysteinyl-FAD) (7), acts as an obligate 2-electron acceptor (6), and is a member of a family of monomeric amino acid oxidases (Mr ~44 kDa) that all contain a single covalently bound flavin (8–13). A nikD-like reaction is implicated in the biosynthesis of the pyridyl moiety found in streptogramin antibiotics that are used to treat multidrug-resistant Gram-positive bacterial infections (14).
There are six possible paths for the initial 2-electron oxidation of P2C to dihydropicolinate (DHP), depending in part on whether nikD oxidizes the imine or enamine tautomer. Oxidation of the bond between N(1) and C(6) in the enamine tautomer, as indicated in Scheme 1, is the most likely path, as judged by results obtained in structural and biochemical studies (7, 15). The second 2-electron oxidation step is more problematic because it would seem to involve oxidation of a bond at a different location within the DHP intermediate, a scenario that requires an apparent change in regiospecificity. This feature would distinguish the nikD reaction from the overall 4-electron oxidation reactions catalyzed by the flavoenzymes choline oxidase, thiamine oxidase and glycolate oxidase because the two successive 2-electron oxidation steps, required to convert alcohol substrates to carboxylic acids, occur at the same carbon (16–18). Similarly, the three successive 2-electron oxidation steps in estrogen biosynthesis occur at the same exocyclic carbon in the impressive aromatization reaction catalyzed by aromatase cytochrome P450 (19).
NikD exhibits two absorption maxima in the visible region, as expected for a flavoprotein, plus an unusual long-wavelength absorption band due to charge transfer interaction of the flavin with Trp355 (20). Crystal structures determined for open and closed forms of the nikD•picolinate complex reveal two distinct ligand binding modes and two different orientations of the side chain of Trp355 (15). In the open form, the rings of picolinate and FAD are nearly perpendicular, a binding mode incompatible with redox catalysis. However, the rings of Trp355 and FAD are parallel in the open form, an orientation that promotes charge transfer interaction. In the closed form, the rings of FAD and picolinate are parallel, a binding mode compatible with redox catalysis. The rings of Trp355 and FAD are perpendicular in the closed form, an orientation that is incompatible with charge transfer interaction. The observed structures led us to propose a two-step mechanism for substrate binding: i) P2C binds to an open form with Trp355 stacked atop the flavin to produce an unstable redox-inactive open complex, otherwise similar to corresponding crystalline complex with picolinate; ii) The open nikD•P2C complex undergoes a conformational change to yield a more stable redox-active closed complex, similar to the crystalline closed nikD•picolinate complex (20).
In this paper, we investigate the mechanism of ligand binding and the initial 2-electron oxidation of P2C. Results obtained with analogs for the imine and enamine forms of P2C do not support a 2-step binding model but are consistent with a one-step approach to equilibrium in a reaction that is orders of magnitude slower than expected for a diffusion-controlled process. Steady-state and pre-steady-state kinetic studies with unlabeled and deuterium-labeled substrate have enabled spectral and kinetic characterization of two intermediates in the conversion of P2C to DHP. Unexpectedly, these studies also provide important insight into the conundrum of the second 2-electron step that produces picolinate.
[3, 3, 4, 4, 5, 5, 6, 6-D8]-L-lysine was purchased from CDN Isotopes Inc. Benzyl chloroformate and picolinate were purchased was Sigma. Methylselenoacetate was a generous gift from Dr. Louis Silks (National Stable Isotope Resource at Los Alamos). 1-Cyclohexenoate was obtained from Aldrich. -N-Carbobenzoxy-L-lysine was purchased from Fluka. L-Amino acid oxidase was isolated from Crotalus adamanteus venom (Biotoxins, Inc.), as previously described (21, 22). Unlabeled P2C was prepared as described by Bruckner et al. (6).
Recombinant wild-type nikD was isolated from cells [Escherichia coli Bl21(DE3)/pDV101] grown in Terrific Broth, as previously described (7). Steady-state kinetic studies with nikD at various concentrations of P2C and oxygen were conducted in 100 mM potassium phosphate buffer, pH 8.0, at 25 °C using screw-cap cuvettes (Spectrocell) that were equipped with a Teflon-silicon membrane and equilibrated at 25 °C with water-saturated gas mixtures containing 10, 21, 44 or 100 % oxygen (balance nitrogen), as previously described (23). Reactions were monitored by measuring picolinate formation at 264 nm ( = 3980 M−1 cm−1) (6). Steady-state kinetic parameters were estimated by fitting an equation for a sequential mechanism (equation 1, A = P2C, B = oxygen) to the data.
[3, 3, 4, 4, 5, 5, 6, 6-D8]-L-lysine was converted to a copper complex of ,-N-carbobenzoxy-L-lysine by following the method of Neuberger and Sanger (24). Free [3, 3, 4, 4, 5, 5, 6, 6-D8]--N-carbobenzoxy-L-lysine was prepared by treatment of the copper complex with sodium sulfide (25) and then converted to deuterium-labeled P2C by following a method previously used to prepare unlabeled P2C (6). Briefly, [3, 3, 4, 4, 5, 5, 6, 6-D8]- -N-carbobenzoxy-L-lysine was oxidized to [3, 3, 4, 4, 5, 5, 6, 6-D8]--N-carbobenzoxy-α-keto--amino-caproic acid using L-amino acid oxidase. The carbobenzoxy group was removed by treatment with acetic acid-HBr, forming [3, 3, 4, 4, 5, 5, 6, 6-D8]-α-keto--amino-caproic acid as a transient intermediate that spontaneously cyclizes to yield the hydrobromide salt of the imine tautomer of P2C (D8-P2C).
Absorption spectra were recorded using an Agilent Technologies 8453 diode array spectrometer. All spectra are corrected for dilution. Enzyme concentration was determined at pH 8.0 based on its absorbance at 456 nm ( = 11,200 M−1 cm−1) (7). The dissociation constant of the nikD•methylselenoacetate complex was determined by fitting a standard binding curve (ΔAobs = ΔAmax[ligand]/(Kd + [ligand]) to the data. The spectrum corresponding to 100% complex formation was calculated as previously described (26). Spectrophotometric titration data with picolinate or 1-cyclohexenoate were analyzed by using an equation for a tight binding inhibitor (equation 2, XT and ET are total ligand and enzyme concentrations, respectively). Fitting of binding equations was conducted by using Sigma Plot 10 (Systat Software).
Rapid reaction kinetic measurements were performed by using a Hi-Tech Scientific SF-61DX2 stopped-flow spectrometer. Data were collected in log mode to maximize the number of points acquired during the early phase of each reaction. All spectra or single-wavelength kinetic traces are the averages of at least three replicate shots. The kinetics of binding of picolinate, 1-cyclohexenoate or methylselenoacetate were monitored by using photomultiplier detection, unless otherwise indicated, in aerobic 100 mM potassium phosphate buffer, pH 8.0, at 25 °C. Reductive half-reactions with unlabeled or deuterium-labeled P2C were monitored by using diode array detection in anaerobic 100 mM potassium phosphate buffer, pH 8.0, containing 50 mM glucose and glucose oxidase (14.7 units/mL). All components, except glucose oxidase, were placed in the main compartment of a tonometer. Glucose oxidase was tipped from a sidearm into the main compartment after the solutions were made anaerobic by multiple cycles of evacuation, followed by flushing with oxygen-scrubbed argon. The entire flow circuit of the stopped-flow spectrometer was made anaerobic by an overnight incubation with anaerobic buffer containing 50 mM glucose and glucose oxidase (14.7 units/mL). All spectra are corrected for a small spectral contribution from P2C in the near-ultraviolet region. Fitting of single-wavelength kinetic traces was conducted by using Sigma Plot 10 (Systat Software), KinetAsyst 3 (TgK Scientifc) or Kinetic Studio (TgK Scientific). Global analyses of absorption spectra acquired during reductive half-reactions were performed using Specfit 3.0, a software package that generates calculated spectra of intermediates and rate constants for intermediate formation and decay.
Turnover rates at various concentrations of unlabeled P2C and oxygen were measured by monitoring picolinate formation at 264 nm (6). Double reciprocal plots of reaction rate versus P2C at different oxygen concentrations or versus oxygen at different P2C concentrations are linear and intersect to the left of the Y-axis, just below the X-axis (Figure 1). The results are consistent with a sequential mechanism where oxygen reacts with a reduced enzyme•DHP complex to yield an oxidized enzyme•DHP complex that can undergo a second redox cycle to produce picolinate. The minimal mechanism shown in Scheme 2 assumes that dissociation of the oxidized enzyme•DHP complex is negligible, consistent with other observations, as will be discussed. The steady-state kinetics parameters listed in Table 1 were obtained by fitting an equation for a sequential mechanism to the data.
Crystal structures observed for open and closed forms of the nikD•picolinate complex suggested that formation of the enzyme•P2C complex might occur via a two-step mechanism (15, 20). We reasoned that the kinetics of ligand binding to nikD might be more easily characterized in studies with competitive inhibitors, such as picolinate, the product of the catalytic reaction, or 1-cyclohexenoate (CHA), a 1-deaza analog of the enamine form of P2C. Fortunately, the binding of these inhibitors is fairly slow and can be readily monitored in a stopped-flow spectrometer at 25 °C.
The spectral course observed for the time-dependent formation of the nikD•picolinate complex in the presence of excess ligand (Figure 2A) is identical to that observed in steady-state titration studies (7). Picolinate binding results in a substantial increase in the intensity of the absorption peak at 456 nm, accompanied by the loss of a long-wavelength absorption band attributable to charge transfer interaction between FAD and Trp355 (20). Apparent first-order rate constants obtained for formation of the picolinate complex are independent of the observation wavelength (Figure 2B). The observed rate constants exhibit a linear dependence on inhibitor concentration (Figure 3A). In contrast, a hyperbolic dependence on ligand concentration with a finite Y-intercept is expected for a two-step binding mechanism (26). The results are, however, consistent with a simple one-step approach to equilibrium where kobs = kf[ligand] + kr. Rate constants for complex formation (kf) and dissociation (kr) were obtained from the slope and intercept, respectively, of this plot. The kinetically determined dissociation constant (Kd = kr/kf = 280 ± 30 µM) is in excellent agreement with a value determined by steady-state titration (Kd = 290 ± 40 µM) (Table 2). The rate constant determined for dissociation of the nikD•picolinate complex (kr = 8.0 ± 0.7 s−1) corresponds to the rate constant for the last step in the minimal steady-state kinetic mechanism (k6) (see Scheme 2). The results indicate that this step is only 3-fold faster than turnover (kcat = 2.3 ± 0.2 s−1), suggesting that product release may be partially rate-determining.
The spectral properties of the nikD•CHA complex are very similar to that observed for the picolinate complex (7) but the complex formed with CHA is ~20-fold more stable. Apparent first-order rate constants obtained for formation of the nikD•CHA complex exhibit a linear dependence on CHA concentration (Figure 3B), consistent with a one-step binding mechanism. Formation of the nikD•CHA complex is ~4-fold faster whereas dissociation of the complex is ~6-fold slower, as compared with the picolinate complex (Table 2).
The nikD complexes formed with picolinate, CHA or various other inhibitors (e.g., benzoate, pyrrole-2-carboxylate, 2-furoate) (7) do not exhibit significant long-wavelength absorption and hence provide no evidence for charge-transfer interaction between the ligands and FAD, as expected for these electron-poor substrate analogs. P2C can, however, exist as an electron-poor protonated imine or an electron-rich enamine (see Scheme 1). Methylselenoacetate (CH3SeCH2CO2 −) (MeSeA) can be viewed as an electron-rich analog of the reactive center in the enamine form of P2C. Importantly, MeSeA does form a spectrally detectable complex with nikD (Kd = 1.07 ± 0.04 mM), characterized by an increase in absorbance in both the 456 nm and long-wavelength regions (Figure 4A). The nikD•MeSeA complex exhibits a charge-transfer band at 581 nm, as estimated by the position of the peak observed in the difference spectrum (Figure 4A, inset). The results suggest that a nikD complex with the enamine tautomer of P2C would also exhibit evidence of charge transfer interaction. Apparent first-order rate constants obtained for formation of the nikD•MeSeA complex exhibit a linear dependence on ligand concentration (Figure 4B), as observed with picolinate and CHA. The charge-transfer complex formed with MeSeA is about 100-fold less stable as compared with the nikD•CHA complex, a difference that is mainly due to a decrease in the rate of complex formation (Table 2).
The kinetics of the initial 2-electron oxidation of P2C to DHP were investigated by monitoring the anaerobic reaction of nikD with various concentrations of P2C in a stopped-flow spectrometer. The reaction with 100 µM P2C exhibits an initial lag, followed by a decrease in absorption at 456 nm, as expected for flavin reduction. The lag becomes progressively smaller at higher substrate concentrations and is not detectable at 2500 µM P2C (Figure 5A). The observed decrease in absorbance at 456 nm exhibits biphasic kinetics (y = Ae−kfastt +Be−kslowt + C) at all tested P2C concentrations (100 to 2500 µM). The rate observed for the initial rapid phase (kfast) exhibits a hyperbolic dependence on the concentration of P2C (kobs = klim[P2C]/(Kd app +[P2C]) (Figure 5B). The limiting rate at saturating P2C (klim = 53 ± 1 s−1) is ~25-fold faster than the value obtained for kcat (2.3 ± 0.2 s−1) in steady-state kinetic studies (Table 1). The results indicate that oxidation of P2C to DHP is not rate-limiting during turnover. The apparent second-order rate constant for the reaction of nikD with P2C, as estimated from stopped-flow data (klim/Kd app = 2.0 ± 0.2 × 105 M−1 s−1), is in excellent agreement with a value calculated using steady-state kinetic parameters (kcat/Km P2C = 2.0 ± 0.3 × 105 M−1 s−1) (Table 1). About 7 % of the total absorbance decrease at 456 nm occurs in a second slow phase. The observed rate of the slow phase (kslow = 2.7 ± 0.1) is independent of the concentration of P2C (Figure 5B) and virtually identical to the value obtained for kcat (Table 1). The results show that the slow phase is kinetically competent and probably rate-determining during turnover. The slow phase is attributed to an isomerization of the initial reduced enzyme•DHP complex (Scheme 3), as will be discussed.
The spectral perturbations and kinetics observed for formation of nikD•inhibitor complexes suggested that binding of P2C would also occur in a relatively slow reaction, accompanied by a substantial increase in absorbance in the 456 nm region. In this case, ES complex formation might overlap with the onset of enzyme reduction, a scenario that could account for the initial lags observed when enzyme reduction is monitored at 456 nm. Preliminary evidence consistent with this hypothesis was obtained by comparing the absorption spectrum of free nikD with spectra observed 0.74 ms after mixing the enzyme with various concentrations of P2C. The resulting difference spectra do indeed exhibit an increase in absorption in the 456 nm region, with positive bands at ~440 and ~465 nm (Figure 6). Furthermore, an excellent fit to the total absorbance change observed at 456 nm during nikD reduction is obtained by fitting a triple-exponential equation to the data, as illustrated by results obtained for the reaction with 100 µM P2C which exhibits a prominent initial lag (see Figure 5A). The results are consistent with an initial, albeit undetected, time-dependent increase in absorbance at 456 nm due to ES complex formation. It is worth noting that difference spectra observed immediately after mixing nikD with P2C exhibit evidence of charge-transfer interaction, as judged by the substantial increase in long-wavelength absorption (λ > 575 nm) (see Figure 6). This feature strongly suggests that the ES complex contains the electron-rich enamine tautomer of P2C and not the electron deficient iminium zwitterion, a species unlikely to act as a charge-transfer donor.
A nearly isosbestic spectral course is obtained for the reaction of nikD with 100 µM P2C, exhibiting apparent concomitant loss of absorption in the 456 nm and long-wavelength regions (Figure 7A). However, closer inspection reveals a small transient increase in absorption at λ > ~535 nm during enzyme reduction (Figure 7A, inset). The magnitude and complexity of the small absorbance changes in the long-wavelength region become progressively more apparent at higher substrate concentrations, as illustrated by results obtained for the reaction with 2500 µM P2C (Figure 7B).
The kinetics of ES complex formation and enzyme reduction with unlabeled P2C appear to be poorly resolved, as judged by results described above. We reasoned that it might be possible to improve the kinetic resolution and directly observe a time-dependent formation of the ES complex by using deuterium-labeled P2C in order to slow the rate of substrate oxidation. As detailed in methods, we used [3, 3, 4, 4, 5, 5, 6, 6-D8]-L-lysine as the starting compound in a multi-step synthesis of deuterium-labeled P2C. The final step is conducted under acidic conditions and yields the hydrobromide salt of the imine tautomer, [3, 3, 4, 4, 5, 5, 6, 6-D8]-P2C (D8-P2C). Solvent exchange at position C(3) in D8-P2C will, however, occur at pH 8.0, owing to a facile interconversion between imine and enamine forms (3–5). Consequently, the deuterium-labeled substrate present in the pH 8.0 buffers used for kinetic studies is likely to contain 6 C–D bonds, [4, 4, 5, 5, 6, 6-D6]-P2C, and will be referred to as D6-P2C.
Double reciprocal plots that exhibit intersecting lines are obtained in steady-state kinetic studies with D6-P2C (data not shown), consistent with a sequential mechanism, as observed with unlabeled P2C. The steady-state kinetic parameters determined with the deuterium-labeled substrate are listed in Table 1. Deuterium substitution causes a nearly 3-fold decrease in turnover rate but does not significantly affect the apparent rate of reaction of the reduced enzyme with oxygen, as judged by values obtained for kcat and kcat/Km oxygen, respectively, with unlabeled and D6-P2C.
The effect of deuterium substitution on the kinetics of P2C oxidation to DHP was determined in reductive half-reaction studies. Importantly, an initial lag is not observed when the anaerobic reaction of nikD with D6-P2C is monitored at 456 nm. Instead, an initial time-dependent increase in absorbance is observed and attributed to ES complex formation (Figure 8A). This initial phase is followed by a biphasic decrease in absorbance at 456 nm, similar to that seen with unlabeled P2C. An excellent fit is obtained by fitting a triple-exponential equation (y = Ae−kinitialt + Be−kfastt +Ce−kslowt + D) to the total absorbance change observed at 456 nm, as illustrated by results obtained for the reaction with 100 µM D6-P2C (Figure 8A, inset)2. About 94% of the total observed absorbance decrease at 456 nm occurs in a fast phase. The observed rate of this phase (kfast) exhibits a hyperbolic dependence on the concentration of P2C (kobs = klim[P2C]/(Kd app +[P2C]) (Figure 8B). The observed rate of the slow phase (kslow) is independent of the concentration of P2C (Figure 8B)3. The latter two features are similar to that observed with unlabeled P2C (see Figure 5).
The limiting rate of nikD reduction at saturating D6-P2C (klim = 8.25 ± 0.01 s−1) is more than 6-fold slower than observed with unlabeled P2C (KIE = 6.4 ± 0.1)4 but nearly an order of magnitude faster than the turnover rate observed with D6-P2C (kcat = 0.89 ± 0.04 s−1) (Table 1). The results indicate that oxidation of deuterium-labeled P2C to DHP is not rate-limiting during turnover. The apparent second-order rate constant for the reaction of nikD with D6-P2C, as estimated from reductive half-reaction studies (klim/Kd app = 1.49 ± 0.02 × 105 M−1 s−1), is in moderately good agreement with a value calculated using steady-state kinetic parameters (kcat/Km P2C = 0.74 ± 0.07 × 105 M−1 s−1). The rate observed for the slow phase of the reductive half-reaction with D6-P2C (kslow = 0.822 ± 0.008 s−1) is more than 3-fold slower than observed with unlabeled P2C (KIE = 3.3 ± 0.1)4 but essentially identical to the value obtained for kcat with D6-P2C. The observed isotope effect indicates that the slow phase cannot simply reflect product release or a protein conformational change but must involve cleavage of one or more C–H bonds in the DHP intermediate, as will be discussed. The results also show that the slow phase observed with D6-P2C is kinetically competent and likely to be rate-determining during turnover with D6-P2C. This outcome agrees with results obtained with unlabeled P2C and indicates that deuterium substitution has not changed the apparent rate-determining step during turnover.
Maximal formation of the ES complex with 400 µM D6-P2C is observed 10 ms after mixing, as judged by the observed increase in absorbance at 456 nm (see Figure 8A). Spectra recorded during the initial 10 ms of this reaction show that ES complex formation results in isosbestic absorption increases in the 456 nm and long-wavelength regions (Figure 9A, inset). The rest of the reaction (t > 10 ms) proceeds with a concomitant loss of absorption in both regions, exhibiting a nearly isosbestic spectral course (Figure 9A, main panel). The data provide no evidence for a small transient increase in long-wavelength absorption during enzyme reduction, unlike results obtained with a low concentration of unlabeled P2C (see Figure 7A, inset).
The rate observed for the fast phase of enzyme reduction with 400 µM D6-P2C is close to the limiting rate of this step (see Figure 8B). Higher concentrations of D6-P2C mainly serve to increase the rate of ES complex formation. Consequently, the time required for maximal ES complex formation is shortened, the apparent maximal yield of the intermediate is increased and a greater proportion of the reaction occurs during mixing at higher substrate concentrations, as can be seen by inspection of the initial absorbance traces at 456 nm (see Figure 8A). These traces indicate that the maximal yield of ES complex observed in these studies is attained 2.24 ms after mixing with 2500 µM D6-P2C, the highest substrate concentration tested. The spectral perturbation due to ES complex formation with 2500 µM D6-P2C was estimated by subtracting the absorption spectrum of free nikD from that observed at 2.24 ms after mixing. The calculated difference spectrum is qualitatively similar, but much greater in magnitude, compared with that seen for the same reaction with unlabeled P2C (see Figure 6). This difference is attributed to the enhanced kinetic resolution of binding and oxidation steps that is achieved with deuterium-labeled substrate.
The spectral course observed for conversion of the ES complex, formed with 2500 µM D6-P2C, to the final reduced enzyme species (Figure 9B) is similar to that observed for the corresponding reaction with 400 µM D6-P2C (Figure 9A), except for a small transient increase in long-wavelength absorption that is observed during reduction at the higher substrate concentration (Figure 9B, inset). The small transient increase in long-wavelength absorption is, however, much less pronounced compared with that observed for the reaction with the same concentration of unlabeled P2C (see Figure 7B, inset). The reason for this difference is unclear.
Global analysis was performed by fitting a model, A → B → C → D, to spectra acquired in reductive half-reaction studies with D6-P2C. In this model, species A is free nikD, intermediate B is the putative oxidized enzyme•P2C charge transfer complex, intermediate C is the postulated reduced enzyme•DHP complex formed in the fast phase of enzyme reduction, and D is the reduced enzyme species observed at the end of the reductive half-reaction (see Scheme 3). A good fit of the proposed model was obtained. Kinetic parameters for the reductive half-reaction obtained by global analysis are in very good agreement with values obtained by analysis of absorbance changes at 456 nm (see Table 1). Similar absorption spectra are calculated for the intermediates by analysis of reactions at different substrate concentrations.
The calculated absorption spectrum of intermediate B exhibits increased absorption in the long-wavelength region, as expected for a charge-transfer complex, and is very similar to the spectrum observed 2.24 ms after mixing nikD with 2500 µM D6-P2C (Figure 10). The calculated absorption spectrum of intermediate C is consistent with that expected for a reduced flavoenzyme but exhibits modest differences, including higher absorbance in the 456 nm region, compared with the observed spectrum of the final reduced enzyme species (D). The calculated intermediate absorption spectra were used to estimate the fraction of the total absorbance decrease at 456 nm that would occur in the fast phase under conditions that promote quantitative ES complex formation [f = (B − C)/(B −D)] versus conditions where negligible accumulation occurs owing to poor kinetic resolution of binding and oxidation steps [f = (A − C)/(A −D)] . The calculated fractions for the two extremes are quite similar (0.957 and 0.949, respectively) and in good agreement with values obtained upon kinetic analysis of the biphasic absorbance decrease observed at 456 nm in reductive half-reactions with D6-P2C (0.944 ± 0.004) or unlabeled substrate (0.935 ± 0.004).
As described above, the rate observed for conversion of the ES complex to the initial reduced enzyme•DHP complex exhibits a hyperbolic dependence on the concentration of P2C (kobs = klim[P2C]/(Kd app +[P2C]). The value obtained for Kd app with D6-P2C is nearly 5-fold smaller than observed with unlabeled substrate (Table 1). For the mechanism shown in Scheme 3, Kd app is equal to the ratio, (kr + klim)/kf. Deuterium substitution is unlikely to affect the rate of ES complex formation (kf) or dissociation (kr). An isotope effect on Kd app is expected for a sticky substrate (kr << klim) but not for a nonsticky substrate (kr >> klim) where the value obtained for Kd app is equal to the true dissociation constant for the ES complex (Kd = kr/kf). The results strongly suggest that P2C is a sticky substrate for nikD. Values for kf, kr and Kd for the ES complex were calculated using values obtained for klim and Kd app with unlabeled and deuterium-labeled P2C. The calculated parameters for the nikD•P2C complex are very similar to kf, kr and Kd values obtained for the nikD complex formed with CHA, a competitive inhibitor and 1-deaza analog of the enamine form of P2C (Table 2).
The calculated value for the rate of dissociation of the nikD•P2C complex (kr = 3.8 ± 0.3 s−1) is more than an order of magnitude slower than the limiting rate obtained for oxidation of unlabeled P2C to DHP (klim = 53 ± 1 s−1), as expected for a sticky substrate. For the mechanism shown in Scheme 3, the ratio klim/Kd app (or kcat/Km P2C) is equal to a collection of rate constants, klimkf/(kr + klim). However, for a sticky substrate (kr << klim) the ratio will be equal to kf, the second-order rate constant for ES complex formation. Consistent with this prediction, the value observed for klim/Kd app (or kcat/Km P2C) with unlabeled P2C (2.0 ± 0.2 × 105 M−1 s−1) is, within experimental error, identical to the estimated value for kf (2.2 ± 0.2 × 105 M−1 s−1). The value observed for klim/Kd app with D6-P2C is about 30% smaller than the estimated value for kf. This outcome is consistent with the fact that the estimated rate of ES complex dissociation is only 2-fold slower than the limiting rate obtained for the 2-electron oxidation of D6-P2C (klim = 8.25 ± 0.01 s−1).
Substrate analogs bind to nikD in a relatively slow reaction and induce substantial changes in the visible absorption spectrum of the enzyme that can be readily monitored at 25 °C by using a stopped-flow spectrometer. Similar kinetics are observed for complex formation with several competitive inhibitors, including electron-poor (CHA) and electron-rich (MeSeA) analogs of the enamine form of P2C, and picolinate, the product of the 4-electron oxidation of P2C. In each case, apparent first-order rate constants observed for complex formation exhibit a linear dependence on ligand concentration with a finite Y-intercept, consistent with a simple one-step approach to equilibrium. The results provide no evidence for a two-step binding mechanism, previously proposed based on structures observed for open and closed forms of the nikD•picolinate complex (20). Trp355 is apparently stacked above the flavin ring in the major form present in solutions of ligand-free nikD at pH 8.0, as judged by the observed charge transfer band. In the two-step model, ligands were thought to form an initial complex where the indole ring of Trp355 remained parallel to the flavin ring, followed by displacement of Trp355 in a spectrally detectable step to produce a more stable complex with ligand stacked above the flavin ring. To account for the observed binding kinetics, we postulate that ligands bind to a minor form where Trp355 is perpendicular to the flavin to produce a complex with ligand stacked above the flavin in a single, spectrally detectable step. The second-order rate constant determined for complex formation (kf) with competitve inhibitors is positively correlated with complex stability: CHA > picolinate > MeSeA. However, even the most tightly bound inhibitor (CHA, Kd = 12.8 µM) binds at a rate (kf = 1.08 × 105 M−1 s−1) that is orders of magnitude slower than predicted for a diffusion-limited reaction (108 – 1010 M−1 s−1). As pointed out by Knowles (27), values below this range may be obtained if the ligand binds to a minor enzyme form, as proposed for nikD.
The initial phase observed during the anaerobic reaction of nikD with P2C is attributed to a relatively slow formation of an ES complex that exhibits higher absorbance at 456 nm and also in the long-wavelength region (λ > 575 nm), a diagnostic feature of charge transfer interaction. The complex is likely to contain the electron-rich enamine tautomer of P2C because the electron-deficient iminium zwitterion is unlikely to act as a charge transfer donor. Significantly, a charge transfer complex is also observed with MeSeA but not detected with electron-poor substrate analogs. The observed ES charge transfer complex is consistent with the proposal that the initial oxidation of P2C occurs at the bond between N(1) and C(6) (see Scheme 1), a reaction possible only with the enamine tautomer. Substrate binding and oxidation steps are poorly resolved when the reductive half-reaction is conducted with unlabeled P2C. Under these conditions, ES complex formation is detectable as an initial lag when the reaction is monitored at 456 nm and by the spectral perturbation observed immediately (t = 0.74 ms) after mixing. A time-dependent formation of the ES complex is, however, directly observed with D6-P2C owing to a slower rate of substrate oxidation.
The second phase of the reductive half-reaction is associated with conversion of the ES complex to a reduced enzyme•DHP complex, (EH2•DHP)ini (see Scheme 3). The observed rate of this step exhibits a hyperbolic dependence on substrate concentration (kobs = klim[P2C]/(Kd app +[P2C]). The limiting rate obtained for nikD reduction at saturating P2C is at least 10-fold faster than the turnover rate observed with labeled or unlabeled substrate and exhibits a substantial kinetic isotope effect (KIE = 6.4 ± 0.1)4. P2C is a sticky substrate, as judged by the observed kinetic isotope effect on Kd app (KIE = 4.7 ± 0.4)4. Values for the rate of formation (kf = 2.2 × 105 M−1 s−1), dissociation (kr = 3.8 s−1) and stability (Kd = 17 µM) of the nikD•P2C complex were estimated on the basis of the observed isotope effects on kcat and Kd app. The calculated parameters are very similar to those observed for the corresponding complex with CHA, a 1-deaza analog of the enamine tautomer (see Table 2). The estimated micromolar binding affinity of nikD for P2C, combined with the fact that P2C acts as a sticky substrate, may represent an evolutionary adaptation designed to prevent cellular accumulation of P2C, a compound that is fairly unstable under physiological conditions.
The calculated absorption spectrum of the initial reduced enzyme•DHP complex, (EH2•DHP)ini, is typical of that expected for a reduced flavoenzyme but exhibits somewhat higher absorbance in the 456 nm region and other differences compared with a final reduced species, (EH2•DHP)fin, that is formed in the third phase of the reductive half-reaction. The observed rate of the final step is independent of the substrate concentration, virtually identical to the turnover rate observed with labeled or unlabeled P2C, and exhibits a significant kinetic isotope effect (KIE = 3.3 ± 0.1)4. The results indicate that the last phase is kinetically competent and probably rate-determining during turnover. The observed isotope effect indicates that this step cannot involve a simple conformational change and/or product release. The results are consistent with a sequential mechanism where oxygen reacts with a reduced enzyme•DHP complex to yield an oxidized enzyme•DHP complex, as proposed on the basis of results obtained in steady-state kinetic studies. Additional evidence that argues against dissociation of any reduced enzyme•DHP complex during turnover is provided by oxidative half-reaction studies which show that free reduced nikD is not a kinetically competent intermediate5. Appreciable dissociation of the oxidized enzyme•DHP complex is also unlikely because DHP oxidation to picolinate is probably at least an order of magnitude faster than DHP release, as judged by values obtained for the corresponding reactions with P2C. Consistent with this hypothesis, the steady-state data provide no evidence for excess substrate inhibition, a feature that would be expected if DHP were released because substrate and the labile intermediate would then compete in binding to oxidized enzyme.
The kinetic isotope effect observed for the last step of the reductive half-reaction indicates that conversion of (EH2•DHP)ini to (EH2•DHP)fin must involve cleavage of one more C–H bonds in the DHP intermediate. A protonated DHP imine is produced during the proposed initial 2-electron oxidation of the P2C enamine (Scheme 4, step 1). Ionization of the protonated DHP imine will yield the corresponding neutral DHP enamine (Scheme 4, step 2), as judged by the analogous tautomerization reaction observed with free P2C (pKa = 8.2) (4, 5). Isomerization of the initially formed DHP enamine (A) will generate a more stable isomer with conjugated double bonds (B) (Scheme 4, step 3). We propose that nikD oxidizes the bond between N(1) and C(6) in isomer B to yield picolinate (Scheme 4, step 4). The proposed mechanism allows nikD to catalyze a remarkable aromatization reaction by effectively oxidizing the same bond twice, a scenario that circumvents the need to reposition FAD and/or the DHP intermediate. Such movements would be required to allow oxidation of different bonds while satisfying the geometric constraints observed for flavoenyzme-catalyzed reactions (15, 28). However, the flavin in nikD is covalently tethered to the protein and not likely to assume different positions during turnover, in contrast to the mobile flavin in p-hydroxybenzoate hydroxylase (29). Furthermore, motion of the DHP intermediate is probably highly constrained by an aromatic cage surrounding the DHP ring plus hydrogen bonding to the DHP carboxylate, as judged by the structure observed for the closed nikD•picolinate complex (15). These restrictions might be relaxed by conversion to an open complex (15) but this change in protein conformation is likely to promote a counterproductive release of the DHP intermediate. A catalytic strategy, similar to that proposed in Scheme 4 for nikD, may be utilized by other flavoenzymes that catalyze the aromatization of L-proline residues to yield pyrrole-2-carboxyl units that are essential components of various nonribosomally synthesized peptidyl antibiotics (e.g., chlorobiocin, pyoluteorin) (30, 31).
We are grateful to Dr. Louis Silks (National Stable Isotope Resource at Los Alamos) for his generous gift of methylselenoacetate. We thank Dr. Phaneeswara-Rao Kommoju for valuable technical assistance.
†This work was supported in part by Grant AI 55590 (M. S. J.) from the National Institutes of Health.
2The data do not permit accurate determination of values for kinitial owing to the small spectral change and the limited number of points contained within the initial phase in 456 nm absorbance traces extracted from diode-array data sets.
3Similar values for kfast and kslow were obtained by biphasic analysis of the absorbance decrease at 456 nm (see Table 1).
4The observed value is likely to include both primary and secondary KIE’s.
5P. R. Kommoju, R. C. Bruckner and M. S. Jorns, unpublished results.