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Originally published In Press as doi:10.1074/jbc.M512385200 on April 20, 2006

J. Biol. Chem., Vol. 281, Issue 25, 17044-17053, June 23, 2006
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Kinetic Mechanisms of the Oxygenase from a Two-component Enzyme, p-Hydroxyphenylacetate 3-Hydroxylase from Acinetobacter baumannii*

Jeerus Sucharitakul{ddagger}1, Pimchai Chaiyen{ddagger}2, Barrie Entsch§, and David P. Ballou§

From the {ddagger}Department of Biochemistry and Center for Excellence in Protein Structure & Function, Faculty of Science, Mahidol University, Bangkok 10400, Thailand and the §Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109-06060

Received for publication, November 18, 2005 , and in revised form, April 19, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
p-Hydroxyphenylacetate hydroxylase (HPAH) from Acinetobacter baumannii catalyzes the hydroxylation of p-hydroxyphenylacetate (HPA) to form 3,4-dihydroxyphenylacetate (DHPA). The enzyme system is composed of two proteins: an FMN reductase (C1) and an oxygenase that uses FMNH– (C2). We report detailed transient kinetics studies at 4 °C of the reaction mechanism of C2.C2 binds rapidly and tightly to reduced FMN (Kd, 1.2 ± 0.2 µM), but less tightly to oxidized FMN (Kd, 250 ± 50 µM). The complex of C -FMNH–2 reacted with oxygen to form C(4a)-hydroperoxy-FMN at 1.1 ± 0.1 x 106 M–1 s–1, whereas the C -FMNH–2 -HPA complex reacted with oxygen to form C(4a)-hydroperoxy-FMN-HPA more slowly (k = 4.8 ± 0.2 x 104 M–1 s–1). The kinetic mechanism of C2 was shown to be a preferential random order type, in which HPA or oxygen can initially bind to the C -FMNH–2 complex, but the preferred path was oxygen reacting with C -FMNH–2 to form the C(4a)-hydroperoxy-FMN intermediate prior to HPA binding. Hydroxylation occurs from the ternary complex with a rate constant of 20 s–1 to form the C2-C(4a)-hydroxy-FMN-DHPA complex. At high HPA concentrations (>0.5 mM), HPA formed a dead end complex with the C2-C(4a)-hydroxy-FMN intermediate (similar to single component flavoprotein hydroxylases), thus inhibiting the bound flavin from returning to the oxidized form. When FADH– was used, C(4a)-hydroperoxy-FAD, C(4a)-hydroxy-FAD, and product were formed at rates similar to those with FMNH–. Thus, C2 has the unusual ability to use both common flavin cofactors in catalysis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Hydroxylation of aromatic compounds in bacteria by single component flavoprotein hydroxylases has been studied extensively for 40 years (1–3). Recently, several research groups, including ours, reported that hydroxylation of aromatic compounds can be catalyzed by two-protein or multiprotein monooxygenases. These include p-hydroxyphenylacetate 3-hydroxylase (HPAH)3 from Pseudomonas putida (4), Escherichia coli (5), and Acinetobacter baumannii (6), phenol hydroxylase (PheA) from both Bacillus stearothermophilus BR219 and Bacillus thermoglucosidasius A7 (7, 8), chlorophenol-4-monooxygenase from Burkholderia cepacia AC1100 (9), 2,4,6-trichlorophenol monooxygenase from Ralstonia eutropha JMP134 (10), pyrrole-2-carboxylate monooxygenase from Rhodococcus sp (11), styrene monooxygenase from Pseudomonas VLB120 (12, 13), and p-nitrophenol hydroxylase from Bacillus sphaericus (14). Most of these enzyme systems consist of reductase and monooxygenase components, where the reductase component provides reduced flavin for the monooxygenase component to use for hydroxylating the aromatic substrate. The number of enzymes known in this class continues to increase and many more hypothetical proteins derived from genome projects have also been identified (15).

Hydroxylation of p-hydroxyphenylacetate (HPA) to form 3,4-dihydroxyphenylacetate (DHPA) by HPAH is especially interesting because the same reaction is carried out by at least three types of two-component enzymes. The first HPAH purified was from P. putida, and it was shown to have FAD tightly bound to the smaller protein, and the larger protein (at that time) was thought to be a coupling protein enabling hydroxylation (4, 16). A different HPAH system was later isolated from E. coli W, and studies have shown that the smaller component (HpaC) is a flavin reductase that generates reduced FAD to be transferred to the larger component (HpaB) to hydroxylate HPA (5, 17). A detailed analysis of the mechanism of the E. coli-type HPAH is now in progress using the homologue from P. aeruginosa (18). The oxygenase in this system exhibits complex dynamics in catalysis (19).

Our group has isolated HPAH from A. baumannii and shown that the enzyme is quite different from the analogous HPAH enzymes from either P. putida or E. coli (6, 15, 20). The A. baumannii HPAH is a two protein enzyme system consisting of a smaller reductase component (C1) and a larger oxygenase component (C2) (6). Sequence and several catalytic properties indicate that both components are different from others in the two protein class of aromatic hydroxylases (15, 20). Our recent investigations of the reaction mechanisms of C1 have shown that HPA controls the reduction of C1-bound FMN by NADH by shifting the enzyme into a more active conformation (20). By contrast, HPA has no effect at all on the activity of the reductase from the E. coli-type HPAH from P. aeruginosa (18). The HPAH from P. putida (above) (16) requires fresh examination based upon our current knowledge. It is possible that this enzyme system operates in a manner similar to the system from A. baumannii, but the essential experiments to test this possibility have not been carried out.

C2 shows little sequence similarity to other oxygenases in the same class, and is unique for its ability to use reduced forms of riboflavin, FMN, or FAD to catalyze hydroxylations (6, 15). The overall reaction of C2 is described in Fig. 1. When C2 was mixed with reduced flavin and a limited amount of oxygen, an intermediate spectrum resembling that of a C(4a)-oxygen adduct of flavin was observed (6). Similar observations were made in the analogous reactions of the oxygenase component involved in biosynthesis of actinorhodin in Streptomyces coelicolor (ActVA) (21, 22) and with chlorophenol 4-monooxygenase (9). Despite preliminary observations that C(4a)-oxygenated intermediates are likely to be involved in oxygenation reactions of these oxygenase components, investigations by presteady state methods to elucidate the enzyme reaction mechanism in detail have never been carried out. In this article, we report investigations on the reaction of oxygen with C2 and reduced flavin using single mixing and double mixing stopped-flow spectrophotometry. The results comprehensively elucidate the reaction mechanism of C2, the order of substrate binding, and the binding constants of FMNH– and HPA to the enzyme.


Figure 1
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  		FIGURE 1.
The reaction catalyzed by the oxygenase component of p-hydroxyphenylacetate hydroxylase.

 

   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents—NAD+, NADH, FAD, glucose, and glucose oxidase were from Sigma. FMN was prepared by conversion of FAD to FMN with snake venom from Crotalus adamanteus (23). In brief, FAD (2.5 mg/ml) and venom (50 µg/ml) in 20 mM potassium phosphate buffer, pH 7.0 were incubated overnight in the dark. The reaction mixture was loaded onto a C18 Sep-Pak cartridge (Waters), previously equilibrated with 20 mM potassium phosphate buffer, pH 7.0, and the cartridge was washed with 10 mM potassium phosphate buffer, pH 7.0. FMN was eluted with water, and the solution was freeze-dried. Concentrations of the following compounds were determined using known extinction coefficients at pH 7.0: NADH, {epsilon}340 = 6.22 mM–1 cm–1; FAD, {epsilon}4500 = 11.3 mM–1 cm–1; FMN, {epsilon}446 = 12.2 mM–1 cm–1 and HPA, {epsilon}277 = 1.5 mM–1 cm–1 (6). C2 used in this study was cloned, expressed, and prepared as previously described (15). The concentration of C2 was estimated from the extinction coefficient (based on amino acid sequence) of {epsilon}280 = 56.7 mM–1 cm–1.

Enzyme Activity—Enzyme hydroxylation activity was detected in real time using a coupling reaction involving 3,4-dihydroxyphenylacetate dioxygenase (DHPAO) to convert the DHPA product of C2 to 5-carboxymethyl-2-hydroxy-muconate semi-aldehyde (CHS). This yellow compound has a maximum absorbance at 380 nm that is dependent upon pH (4, 6).

Spectroscopic Studies—UV-visible absorbance spectra were recorded with a Hewlett Packard diode array spectrophotometer (HP 8453A), or a Shimadzu 2501PC spectrophotometer. Fluorescence measurements were carried out with a Shimadzu RF5301PC spectrofluorometer. All these instruments were equipped with thermostatic cell compartments.

Determination of the Kd for Binding Oxidized FMN to C2—The measurements were performed by an ultrafiltration method using Centriprep® Y-M 10 from Amicon. Solutions were composed of 10 µM FMN in 50 mM sodium phosphate buffer, pH 7.0, and various concentrations of C2, (20, 40, 80, 160, and 200 µM) in a 10-ml total volume. Each solution was centrifuged at 3,200 rpm, 4 °C for 15 min to obtain a filtrate of ~1 ml (to minimize change in volume). The filtrate and retentate were analyzed for the amount of free and bound FMN, respectively. Ratios of the free and bound species were used to calculate the Kd value.

Rapid Reaction Experiments—Reactions were carried out in 50 mM sodium phosphate buffer, pH 7.0, 4 °C, unless otherwise specified. Rapid kinetics measurements were performed with a Hi-Tech Scientific Model SF-61DX in double mixing mode, or with either a model SF-61SX or a SF-61DX stopped-flow spectrophotometer in single mixing mode. The optical pathlengths of the observation cells were 1 cm. The stopped-flow apparatus was made anaerobic by flushing the flow system with an oxygen-scrubbing solution consisting of 400 µM glucose, 1 mg/ml glucose oxidase (15.5 unit/ml), and 4.8 µg/ml catalase in 50 mM sodium phosphate buffer, pH 7.0. The oxygen-scrubbing solution was allowed to stand in the flow system overnight and was then thoroughly rinsed with anaerobic buffer before experiments.

To study the oxidative half-reaction of C2, enzyme, or enzyme plus substrate and oxidized FMN in 50 mM sodium phosphate buffer, pH 7.0, were made anaerobic in glass tonometers by several cycles of evacuation followed by equilibration with argon that had been passed through an Oxyclear oxygen removal column (Labclear). Enzyme was anaerobically reduced with a solution of sodium dithionite (~5 mg/ml in 100 mM potassium phosphate buffer, pH 7.0) delivered from a syringe attached to the tonometer, and the reduction was monitored by UV-visible spectrophotometry. Solutions with various concentrations of oxygen were prepared by equilibrating buffer with air or with certified oxygen/nitrogen gas mixtures. Determinations of rate constants were obtained by fitting plots of apparent rate constants (kobs) versus concentration of substrate with a Marquardt-Levenberg non-linear fit algorithm that is included in the KaleidaGraph software (Synergy Software). The kobs from kinetic traces were calculated from exponential fits using KinetA-syst3 software (Hi-Tech Scientific, Salisbury, UK) or program A (written at the University of Michigan by Rong Chang, Jung-yen Chiu, Joel Dinverno, and D. P. Ballou).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Reaction of C2-FMNH– with O2 in the Absence of HPA—A solution of FMN (16 µM) plus C2 (25 µM) was placed in a glass tonometer equipped with a quartz cuvette, and made anaerobic as described under "Materials and Methods." An anaerobic solution of sodium dithionite was delivered into the tonometer to stoichiometrically reduce the FMN (see "Materials and Methods"). The resulting C2-FMNH– complex was loaded onto the stopped-flow spectrophotometer, where its reaction with oxygen was monitored at 380 and 446 nm (Fig. 2). A significant fraction of the first phase, which is shown by an increase in absorbance at 380 nm and no change at 446 nm, occurred during the dead time of the instrument (~0.002 s) and was complete by 0.006–0.02 s (high to low oxygen concentration, Fig. 2). The plot of kobs of this phase versus oxygen concentration was linear, yielding a second-order rate constant of 1.1 ± 0.1 x 106 M–1 s–1 (inset in Fig. 2). When absorbance values at the end of the first phase (reaction time of 0.01 s of highest oxygen reaction) at various wavelengths were plotted, the spectrum B in Fig. 3 was obtained. This spectrum has characteristics typical for a flavin-C(4a)-adduct with maximum absorbance at 380 nm. Based upon analogy to the reactions catalyzed by one component hydroxylases and the condition that HPA is absent, the spectrum B in Fig. 3 is likely to be C(4a)-hydroperoxy-FMN (1–3). Spectrum B is also similar to that of C(4a)-hydroperoxyflavins generally found in the class of single component aromatic hydroxylases (1, 3, 24–26), as well as for luciferase (the first two-component flavin-dependent oxygenase studied in detail) (27, 28), cyclohexanone monooxygenase (29) and model systems (30, 31). The hydroperoxide intermediate decayed slowly to yield oxidized FMN and probably H2O2 (at 0.037 ± 0.002 s–1, see traces at 446 nm from 1 to 100 s in Fig. 2). A small increase in absorbance at 380 nm was also observed between 0.01 and 1 s, and the kobs describing this phase was also dependent on oxygen concentration. Based on the Kd value of C2-FMNH– of 1.2 ± 0.2 µM (described in the next paragraph), ~1.6 µM free FMNH– is present under these reaction conditions. Therefore, this small absorbance change is likely to be because of free FMNH– reacting with oxygen.


Figure 2
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  		FIGURE 2.
Kinetics of reoxidation of C2-FMNH– complex. FMNH– (16 µM) plus C2 (25 µM) were mixed with buffer containing oxygen (0.13, 0.31, 0.61, and 1.03 mM from the lowest to the upper trace) in the stopped-flow spectrophotometer. All concentrations given are after mixing. The reactions were monitored at 380 and 446 nm (all traces at 446 nm superimposed). The inset shows the plot of kobs of the first phase against oxygen concentrations. The second order rate constant was calculated from the slope of the plot to be 1.1 ± 0.1 x 106 M–1 s–1.

 


Figure 3
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  		FIGURE 3.
Absorbance spectra observed during the reoxidation of C2-FMNH–. Results from the experiment described in the legend to Fig. 2 monitored at various wavelengths ranging from 310 to 550 nm, were used for plotting the intermediate spectrum. The dashed line (A) represents the beginning C2-FMNH– complex, and the solid line (C) represents the spectrum of the final species, oxidized FMN with C2. The intermediate spectrum (B) was obtained by plotting absorbance values occurring at the reaction time of 0.01 s. This is valid because the decomposition of the intermediate is very slow compared with its rate of formation.

 


Figure 4
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  		FIGURE 4.
Determination of the dissociation constant for C2 binding with reduced FMN. FMNH– (16 µM) was mixed with air-saturated buffer containing various concentrations of C2 (0, 2, 6, 10, 16, 20, 30, and 40 µM from lower to upper traces). The reactions were monitored at 380 nm, and all concentrations given are after mixing. The inset is a plot of the absorbance change that had occurred at 0.04 s versus the concentration of free C2. The free C2 concentration was calculated by subtraction of the concentration of C(4a)-hydroperoxy-FMN formed from the total amount of C2 present. The Kd for the C2-FMNH– complex was calculated to be 1.2 ± 0.2 µM.

 
When the same reaction of the C2-FMNH– complex was investigated with fluorescence detection using the excitation wavelengths of 360–446 nm and emission at wavelengths of greater than 500 nm, fluorescence increases were only observed with formation of the final species, oxidized FMN (data not shown). This result indicated that the C2-C(4a)-hydroperoxy-FMN was non-fluorescent.

Determination of Binding Constants of Reduced and Oxidized Flavin to C2—Free reduced FMN was mixed with air-saturated C2 solution in the stopped-flow apparatus, resulting in a reaction with kinetic traces nearly identical to those obtained when preformed C2-FMNH– was mixed with oxygen, as shown in Fig. 2. This result implies that binding of FMNH– to C2 is much faster than the oxidation of free FMNH– by oxygen (32), and also greater than the rate of formation of the C(4a)-flavin hydroperoxide at 0.13 mM oxygen, 185 ± 9s–1 (Fig. 2). Thus, the rate constant for C2 binding to FMNH– is likely to be ≥107 M–1 s–1 (k1 in Fig. 10).

Therefore, when FMNH– (16 µM) was mixed with various concentrations of C2 in air-saturated buffer in the stopped-flow spectrophotometer, the absorbance increase at 380 nm during the first phase (Fig. 4), because of the C(4a)-hydroperoxy FMN formed, was also directly dependent on the amount of C2-FMNH– complex initially present. In the absence of C2 (the lowest trace), the absorbance increased with a t1/2 ~0.7 s as free FMNH– oxidized to FMN in a complex autocatalytic reaction (32). As the concentration of C2 increased, less auto-oxidation of FMNH– is observed. Therefore, the increase in absorbance observed at 0.04 s represents the amount of C2-FMNH– present at the start of the reaction, and the plot of this change in absorbance versus the free C2 concentration represents the binding isotherm for FMNH–. The plot (inset in Fig. 4) shows that the absorbance increase is hyperbolically dependent on C2 concentration. A Kd (referred to as K Ad in the kinetic scheme in Fig. 10) value for the complex was calculated to be 1.2 ± 0.2 µM.


Figure 5
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  		FIGURE 5.
Reaction of oxygen with the C2-FMNH–-HPA complex. A solution containing FMNH– (16 µM), C2, (25 µM), and HPA (2 mM) (final concentrations) was mixed with various concentrations of oxygen containing HPA (2 mM) in the stopped-flow spectrophotometer. Reactions were monitored by absorbance at 380 nm. The arrow indicates enzyme absorbance (0.076 AU) before starting the reaction. Absorbance traces with final oxygen concentrations of 0.13, 0.31, 0.61, and 1.03 mM are shown from right to left. The dotted lines represent multiphasic exponential fits to the experimental data. The inset shows spectra of flavin intermediates formed in the oxygen reaction of the C2-FMNH–-HPA complex. Traces of reactions monitored at multiple wavelengths from 310–550 nm were used for calculating the spectra of flavin intermediates. The dotted line is a spectrum of the C2-FMNH–-HPA complex, whereas the upper solid line is the spectrum of oxidized FMN at completion of the reaction. Absorbance at 60 ms after the start of the reaction is plotted in the line with filled circles, and this represents predominately the spectrum of the C2-C(4a)-hydroperoxy-FMN-HPA complex. Absorbance at the reaction time of 150 ms was plotted in the empty circle line, and this represents predominately the spectrum of C2-C(4a)-hydroxy-FMN. Times were chosen on the basis of the rate constants in the reaction.

 
The Kd for the binding of oxidized FMN to C2 was determined by an ultrafiltration method described under "Materials and Methods." This experiment indicated that the Kd of C2-FMN was 250 ± 50 µM. The large uncertainty occurred because values could not be obtained at appropriately high C2 concentrations.

Reaction of the C2-FMNH–-HPA Complex with Oxygen—The reaction of C2 in the presence of substrate was investigated by mixing an anaerobic solution of FMNH– (16 µM), C2 (25 µM), and 2 mM HPA with buffer containing various oxygen concentrations in the stopped-flow spectrophotometer (Fig. 5). The rate of formation of C2 intermediate in presence of HPA was second order with respect to oxygen; however, the reaction is considerably slower than when no HPA is present (compare the increases at 380 nm in Fig. 2 to those in Fig. 5). An obvious interpretation of this result is that the reaction with oxygen is slower when HPA is bound to the enzyme. This interpretation was later verified (Figs. 7 and 8). In experiments where the concentration of HPA was varied in double mixing stopped-flow experiments, the rate of formation of C(4a)-hydroperoxy-FMN decreased with increasing concentrations of HPA (data not shown).

Fig. 5 shows that the reaction monitored at 380 nm consists of 4 phases. For example, with 130 µM oxygen (concentration after mixing), the first phase consisted of an increase of absorbance of ~0.026 AU occurring during the dead time, and continuing until ~0.012 s. With the highest oxygen concentration used (1.03 mM), this phase was complete during the dead time of the instrument. This first phase was dependent on oxygen concentration and is characterized by a rate constant of 1.1 ± 0.1 x 106 M–1 s–1 (data not shown). This value is the same as that observed in the reaction of C -FMNH–2 with oxygen in Fig. 2 (with no HPA present). The second phase (~0.012–0.6 s) of ~0.079 AU at 380 nm was also dependent on oxygen concentration and was characterized by a second order rate constant of 4.8 ± 0.2 x 104 M–1 s–1 (data not shown). Therefore, the first phase is likely to be the reaction of oxygen with C -FMNH–2 without HPA bound, whereas the second phase is due to the ternary complex C -FMNH–2 -HPA reacting with oxygen. The third phase is a lag in absorbance at 380 nm and corresponds to a process with a rate of ~17–22 s–1 (k6 in Fig. 10). This phase can only be resolved clearly in the reaction with 1.03 mM oxygen. The decrease in absorbance because of the fourth phase (0.7–20 s) was coupled with a large increase at 446 nm; this phase was fitted with a rate constant (0.35 ± 0.02 s–1) that was independent of oxygen concentration.

The absorption spectra of intermediates were calculated from the traces over a range of wavelengths of the reaction with 1.03 mM oxygen using the following rate constants: 54 s–1 for formation of the first intermediate, 20 s–1 for the second intermediate, and 0.35 s–1 for the final step in which oxidized FMN is formed. The analysis shows that spectra of the two intermediates are very similar (inset of Fig. 5), and have absorption characteristics similar to C(4a)-intermediates found for the single component flavoprotein hydroxylases (1, 24–26, 33). This also implies that the first and second intermediates are likely to be C2-C(4a)-hydroperoxy-FMN-HPA complex and C2-C(4a)-hydroxy-FMN-product complex, respectively (inset of Fig. 5). The slight increased absorbance in the region of 450 nm of the second intermediate in the inset to Fig. 5 is unlikely to belong to absorption of C2-C(4a)-hydroxy-FMN, but rather to a small amount of oxidized FMN resulting from an uncoupling pathway that does not result in hydroxylation (2, 24–26).

Therefore, we conclude that the first phase is the reaction of C -FMNH–2 (without HPA bound) whereas the second phase is the formation of C2-C(4a)-hydroperoxy-FMN-HPA complex. This also implies that the binding of HPA to the enzyme decreases the rate of formation of C2-C(4a)-hydroperoxy-FMN about 20-fold. We interpret the third phase to be the hydroxylation step where C2-C(4a)-hydroperoxy-FMN reacted with HPA to form the C2-C(4a)-hydroxy-FMN and DHPA. The C(4a)-hydroperoxy-FMN and C(4a)-hydroxy-FMN species have very similar spectra with this enzyme (see below), causing the absorbance change upon hydroxylation to be very small. Because of this small absorbance change, the rate constant for the hydroxylation step could not be determined accurately by this procedure. The fourth phase was caused by the dehydration of C2-C(4a)-hydroxy-FMN to yield the oxidized FMN species.

To verify further if the C -FMNH–2 -HPA complex has indeed led to hydroxylation as described above, DHPA product formed under the conditions used in stopped-flow experiments was determined. Reaction samples were collected from the stopped-flow instrument and quantified by HPLC methods. Solutions of FMNH– (50 µM), C2 (80 µM), and HPA (2 mM) were mixed with air-saturated buffer containing 2 mM HPA at 4 °C in the stopped-flow spectrophotometer. The reaction solutions from several shots were collected for analysis. The collected solutions were ultrafiltered using Centricons to remove the enzyme, and the samples were analyzed for DHPA by HPLC methods described previously (6). The analysis showed that 73 ± 4% of HPA was hydroxylated to form the DHPA product from the ternary complex under these conditions (Table 1).


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  		TABLE 1
Determination of hydroxylated product from single turnover reactions of C2

All reactions were carried out in 50 mM sodium phosphate buffer, pH 7.0, 4 °C, in a stopped-flow spectrophotometer. The percentage of product formed was calculated from the ratio of product determined to the amount of reduced flavin used. The reagents in syringe A were specified inside the brackets, whereas syringe B contained buffer with oxygen or reduced flavin. DHPA was determined by HPLC after enzyme was removed by ultrafiltration (6).

 
HPA Is Not the First Substrate Binding to C2—The reaction of C2 involves three substrates (Fig. 1), HPA, FMNH–, and oxygen. In this section we describe experiments to determine the sequence of binding of these compounds to C2. Fig. 6 shows experiments of the reaction to form the C2-C(4a)-hydroperoxy-FMN involving various premixing protocols. Trace A shows the reaction of the C -FMNH–2 -HPA complex with O2 (from Fig. 5), and trace B shows the reaction of the C -FMNH–2 complex with O2 and HPA. These demonstrate that the C -FMNH–2 -HPA complex reacts with O2 much more slowly than does the C -FMNH–2 complex. We used this information to examine whether C2 can effectively bind HPA in the absence of FMNH–. If such a complex does form, it would be expected that this complex in the presence of O2 would react with FMNH– to form C2-C(4a)-hydroperoxy-FMN at the slower rate, as seen in trace A. Upon mixing a solution containing C2, HPA, and oxygen with FMNH– in the stopped-flow spectrophotometer (trace D, Fig. 6), C2-C(4a)-hydroperoxy-FMN formed at the same rate as the reaction with free C -FMNH–2 (trace B) as well as that with mixing C2 in air-saturated solution with FMNH– (trace C). These results suggest that C2 alone does not bind effectively to HPA, and that FMNH– is the first species binding to the enzyme during catalysis.


Figure 6
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  		FIGURE 6.
Reduced FMN is the first ligand to bind to C2. Trace A is from a reaction in which C2-FMNH–-HPA complex (16 µM FMNH–, 25 µM C2, and 2 mM HPA final) was mixed with a solution of HPA (2 mM) containing 130 µM oxygen. Trace B is from the reaction in which the C2-FMNH– complex was mixed with a solution containing HPA and oxygen. Trace C was obtained from reacting a solution of C2 containing oxygen with a solution of FMNH–. Trace D is the result of premixing C2 with HPA in buffer containing oxygen, and then mixing with a solution of FMNH–.

 
Binding of HPA to C2FMNH–—After binding to C2 to form the C -FMNH–2 complex, the enzyme can in principle carry out the reaction through one of two paths (Fig. 10): A) C -FMNH–2 -HPA is formed prior to reacting with oxygen, and B) C2-C(4a)-hydroperoxy-FMN is formed prior to binding HPA. In this section, we report investigations of the kinetics and thermodynamics of binding HPA to C -FMNH–2 prior to reacting with oxygen. The binding kinetics of HPA to C -FMNH–2 were investigated by double mixing stopped-flow spectrophotometry, where the first mixing added HPA to C -FMNH–2 under anaerobic conditions to initiate the formation of the C -FMNH–2 -HPA complex, and after various times of aging, the second mix added oxygen to form the C2-C(4a)-hydroperoxy-FMN species, either in complex with HPA, or not. Final concentrations after double mixing were C2 (25 µM), FMNH– (16 µM), HPA (2 mM), and oxygen (1.03 mM). This resulted in ~16 µM of C2-FMNH– in the solution after double mixing. The reaction was monitored by absorbance at 380 nm to detect formation of C2-C(4a)-hydroperoxy-FMN. Any C2-FMNH– binary complex present reacted with this concentration of oxygen at >1000 s–1 and largely occurred in the dead time of the instrument, as shown by the dotted line in Fig. 7. Upon increasing the age time, formation of the C(4a)-hydroperoxy-FMN became slower as more C2-FMNH–-HPA complex formed. Any C2-FMNH–-HPA complex present reacted to form C2-C(4a)-hydroperoxy-FMN-HPA at ~54 s–1, and the reaction was completed by ~90 ms. This indicates that the more complete the formation of C2-FMNH–-HPA, the slower was the formation of C2-C(4a)-hydroperoxy-FMN. This result is also consistent with our previous interpretation in Fig. 5 that C2-FMNH–-HPA reacts with oxygen more slowly than does C2-FMNH–. Fig. 7 shows that the amount of C2-C(4a)-hydroperoxy-FMN formed between 7 and 80 ms (the slower reaction) was maximum when age times before mixing were ≥1s(inset in Fig. 7), indicating that binding of 2 mM HPAtoC2-FMNH– was complete by 1 s. The apparent rate constant (kobs) for binding of HPA to C2-FMNH–, calculated from the plot of the absorbance increase observed at 380 nm versus the age times after the first mixing, was 9.6 ± 2 s–1.


Figure 7
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  		FIGURE 7.
Kinetics of C2-FMNH– complex formation with HPA. Formation of the complex was investigated by double mixing stopped-flow spectrophotometry. In the first mix, HPA was added anaerobically to C2-FMNH–, and after various age times (0.05, 0.1, 1, 2, 4, 6, 8, and 10 s, upper to lower traces), oxygen was mixed with the aged solution. The final concentrations after double mixing were 16 µM C2-FMNH– (25 µM C2 plus 16 µM FMNH–), 2 mM HPA and 1.03 mM oxygen. Reactions were monitored by absorbance at 380 nm to assess the amount of C2-C(4a)-hydroperoxy-FMN formed. The dotted line represents the reaction of C2-FMNH– with oxygen under the same conditions. A plot of absorbance amplitudes that occurred at 380 nm at ~54 s–1 versus the age time (inset) represents the binding kinetics of HPA to C2-FMNH–.

 
The thermodynamics for binding of HPA to C2-FMNH– prior to reacting with oxygen were investigated by double mixing stopped-flow spectrophotometry, where various concentrations of HPA were mixed with C2-FMNH– anaerobically in the first mix, and oxygen was added to the preformed C2-FMNH–-HPA complex in the second mix (Fig. 8). The age time was 10 s to ensure complete formation of the substrate complex, and the reaction was monitored at 370 and 446 nm. The final concentrations were 16 µM C2-FMNH–, 1.03 mM oxygen, and a range of HPA concentrations. Fig. 8 shows that the kinetic traces are composed of four phases. The first phase observed is an increase in absorbance at 370 nm with an observed rate constant of 54 s–1, and the amplitude of this phase is dependent on concentration of HPA. It is known from the single mixing experiments (Fig. 2) that the reaction of C2-FMNH– and 1.03 mM oxygen is fast enough to be largely completed during the dead time of the stopped-flow instrument. With higher concentrations of HPA, larger fractions of the enzyme were in the form of the C2-FMNH–-HPA complex, resulting in smaller fractions of the enzyme reacting with oxygen during the dead time, and larger fractions reacting at 54 s–1 (Fig. 8). These amplitude changes were plotted against the concentrations of HPA, and a Kd value of 180 ± 3 µM (referred as K Bd in Fig. 10) was calculated for binding of HPA to C2-FMNH– (inset A in Fig. 8).


Figure 8
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  		FIGURE 8.
Dissociation constant for binding HPA to C2-FMNH–. This was measured by double mixing stopped-flow spectrophotometry. The first mix added various concentrations of HPA (80, 160, 400, 800, 2000, 4000, 8000 µM, from upper to lower traces) anaerobically to C2-FMNH– (16 µM), and the second mix added 1.03 mM oxygen to the resultant mixture. The age time between mixes was 10 s to ensure the complete formation of any C2-FMNH–-HPA complex, and the reactions were monitored at 370 nm. All concentrations given are after final mixing. The amplitude changes in the phase occurring at 54 s–1 (dead time to 80 ms) versus the HPA concentration are plotted in inset A and this was used to calculate the Kd (180 ± 30 µM) for binding of HPA to C2-FMNH–. Inset B shows the formation of oxidized FMN at 446 nm (see also Fig. 9).

 
The second phase in the reactions in Fig. 8 was a small decrease in absorbance with an observed rate constant of ~17–22 s–1 (k6 in Fig. 10), whereas the third phase was a small increase in absorbance with a rate constant of ~6–9 s–1 (k7 in Fig. 10). The absorbance at 370 nm decreased again in the fourth phase with the kobs values inversely dependent on the concentration of HPA used. The fourth phase was identified as the formation of the final oxidized FMN species because the traces coincided with a large increase in absorbance at 446 nm (shown in inset B). These results suggest that after formation of C2-C(4a)-hydroperoxy-FMN during the first phase, HPA was hydroxylated with the formation of C2-C(4a)-hydroxy-FMN during the second phase, similar to the results of Fig. 5. However, it is clear from this experiment that excess HPA can also bind to the enzyme to trap the C2-C(4a)-hydroxy-FMN-HPA species (Fig. 10) in the third phase, causing a slight increase in absorbance at 370 nm. With higher concentrations of HPA, more of this intermediate was trapped as the C2-C(4a)-hydroxy-FMN-HPA species, so that the dehydration to form the oxidized FMN was retarded (inset B in Fig. 8). Similar trapped C(4a)-hydroxyflavin-substrate species have also been observed in the oxidative half-reactions of several single component flavoprotein oxygenase enzymes (24–26, 34–35).

The Reaction of C2-C(4a)-hydroperoxy-FMN with HPA—Experiments from the previous section show that hydroxylation can occur via Path A in Fig. 10 where C2-FMNH– first binds to HPA and then reacts with oxygen to form C2-C(4a)-hydroperoxy-FMN-HPA or through Path B of Fig. 10, where the enzyme first forms C2-C(4a)-hydroperoxy-FMN and then binds to HPA in a following step. Therefore, the reaction of Path B was explored using a double mixing stopped-flow spectrophotometer, where the intermediate C2-C(4a)-hydroperoxy-FMN was generated by reacting C2-FMNH– with oxygen in the initial mixing; after aging for 0.1 s to fully form the C(4a)-hydroperoxy-FMN, the resultant intermediate was mixed with buffer containing various HPA concentrations. Reactions were monitored at 370 nm (Fig. 9), and the results indicate that binding to HPA (the small increase in absorbance from 2–20 ms) gave a phase with amplitudes and rates that were dependent on the concentration of HPA. Kinetic analysis showed that the observed rate constants (kobs) of this phase were hyperbolically dependent on HPA concentrations (inset A in Fig. 9). These results are consistent with binding being a two-step process, with a rapid equilibrium in the initial step and an isomerization in the following step (Path B in Fig. 10) (36). Data were analyzed according to Equation 1, yielding a Kd value for the initial binding of HPA of 0.35 ± 0.03 mM (K Cd, Fig. 10), and the rate constant for the subsequent isomerization of 208 ± 4s–1 (k4 in Fig. 10).

Formula 1(Eq. 1)
After HPA has bound, there are three more phases in the reaction similar to those seen in the double mixing experiments described in Fig. 8. In the second phase, C2-C(4a)-hydroperoxy-FMN-HPA converted to C2-C(4a)-hydroxy-FMN-DHPA with a rate of 17–22 s–1 (k6 in Fig. 10), and this was followed by a slight increase in absorbance during the third phase. The amplitude of the third phase is also dependent on HPA concentration. As before, the second phase is the hydroxylation step, and the third phase is the binding of HPA to C2-C(4a)-hydroxy-FMN, coincident with the release of DHPA. The fourth phase is the decrease in absorbance 370 nm and a large increase of absorbance 446 similar to those of Fig. 8B (data not shown). The fourth phase was interpreted as the dehydration of C2-C(4a)-hydroxy-FMN to form oxidized FMN, and this rate was inversely dependent on HPA concentration (as discussed in the next section and shown in inset B). When all relevant kinetic constants were considered, the results suggest that the oxygenation reaction of C2 occurs preferentially through Path B (Fig. 10) over Path A. Therefore, the kinetic mechanism for the oxygenation reaction of C2-FMNH– can be described as a preferential random order mechanism as shown in Fig. 10, and the preferred path utilized will be determined by the relative concentrations of substrates. (The formal nomenclature would be Uni Bi Uni Bi mechanism with a random segment for the second and third substrates, and probably the second and third products.)


Figure 9
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  		FIGURE 9.
Reaction of C2-C(4a)-hydroperoxy-FMN with HPA. The intermediate C2-C(4a)-hydroperoxy-FMN was generated by initially mixing C2-FMNH– (16 µM) with 0.31 mM oxygen in the double mixing stopped-flow spectrophotometer, and aging for 0.1 s. The resultant intermediate was then mixed with buffer containing various HPA concentrations of 80, 160, 400, 800, 2000, 4000, 8000 µM (lower to upper traces, respectively). Reactions were monitored by absorbance at 370 nm. All concentrations are given as final reaction conditions. The increase of absorbance during the first phase was dependent on HPA concentration and represents binding of HPA to C2-C(4a)-hydroperoxy-FMN. Inset A shows the plot of kobs for the first phase versus HPA concentrations. Inset B is a plot of kobs of the final step, dehydration of C2-C(4a)-hydroxy-FMN to the oxidized FMN versus HPA concentration.

 


Figure 10
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  		FIGURE 10.
C2 preferential random order reaction mechanism. Kinetic and thermodynamic constants relevant to each step are shown in the scheme.

 
To ensure that the hydroxylation reaction indeed occurred via Path B of Fig. 10, analyses were also carried out with reaction mixtures in which solutions of FMNH– (50 µM) and 2 HPA (2 mM) were mixed in the stopped-flow instrument with a solution of C2 (80 µM) and HPA (2 mM) in air-saturated buffer. Under these conditions, the reaction occurred via Path B in Fig. 10, where the intermediate C(4a)-hydroperoxyflavin was formed prior to binding of HPA. Samples collected from the stopped-flow instrument were analyzed by HPLC as described previously (Table 1). Results indicated that the DHPA product yield from Path B was 82 ± 3%, which is slightly greater than the value obtained from Path A as 73 ± 4% (Table 1 and Fig. 5).

Inhibition of the Dehydration of C2-C(4a)-hydroxy-FMN by HPA—Data from experiments shown in Figs. 8 and 9 indicate that the observed rate constants of the fourth phase were inversely related to the concentration of HPA, implying that HPA binds to the enzyme to form a trapped C2-C(4a)-hydroxy-FMN-HPA species and impedes it from dehydrating into oxidized FMN. Consistent with this interpretation, a plot of the observed rate constants of the last phase versus HPA concentration shows an inverse dependence on HPA. The rate extrapolated to zero with increasing concentrations of HPA (inset B in Fig. 9), implying that the trapped species is a dead-end complex and is not capable of dehydrating to form the oxidized enzyme. Because the observed rate constants for dehydration are specified by two reactions, the dehydration and the rebinding of HPA to the intermediate (Fig. 10), Equation 2 was used to analyze the observed rate constants for the dehydration. When the observed rate constants of the fourth phase were fitted to Equation 2, the K ind for dissociation of HPA from the C2-C(4a)-hydroxy-FMN was calculated to be 41 ± 1 µM (K ind in Fig. 10) and the dehydration rate constant obtained from extrapolation to zero HPA was 8.3 ± 2 s–1. This dehydration rate must represent the rate of dissociation of HPA from the enzyme.

Formula 2(Eq. 2)

Oxygen Reaction of C2-FADH– in the Presence and Absence of HPA—A unique property of C2 compared with other two-component flavin-dependent hydroxylases is its ability to use a variety of reduced flavin substrates. FADH– is as effective as FMNH– (6, 15). Therefore, we tested whether the mechanistic details for oxygenation by C2 with FADH– are similar to those in the reaction of C2 with FMNH–. Experiments similar to those described in Figs. 2 and 5 were carried out, but using C -FADH–2 instead of C2-FMNH–. The reaction of C2-FADH– with oxygen is very similar to the reaction of C -FMNH–2. The rate constant was 0.98 ± 0.05 x 106 M–1 s–1 for formation of C2-C(4a)-hydroperoxy-FAD versus 1.1 ± 0.1 x 106 M–1 s–1 for the reaction with FMNH– (Table 2, data not shown). In the presence of HPA, the C -FADH–2 -HPA complex reacted with oxygen more slowly than in the absence of HPA, similar to the reactions with FMNH–. The rate constant for formation of the C2-C(4a)-hydroperoxy-FAD-HPA complex is slightly smaller than that with FMN (3.7 ± 0.2 x 104 M–1 s–1 for FAD versus 4.8 ± 0.2 x 104 M–1 s–1 for FMN (see Table 2). The spectra of C2-C(4a)-hydroperoxy-FAD, both in the absence and presence of HPA, calculated using the method described in the FMN experiments, are very similar to those for the C2-FMNH reactions (data not shown).


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  		TABLE 2
Kinetic constants for reactions of C2–FMNH– and C2–FADH–

 
Double mixing experiments similar to those in Figs. 8 and 9, but using FADH– instead of FMNH–, yielded results similar to those with FMNH–. The Kd values for binding of HPA to C -FADH–2 or to C2-C(4a)-hydroperoxy-FAD are both similar to those with FMNH– (shown in Table 2), again emphasizing that FADH– can be used nearly as well as FMNH– by C2 with respect to both specificity and reactivity.

Single turnover reactions of C2 and FADH– were analyzed for the amount of hydroxylated product (Table 1) using the same protocols described in previous sessions of C2 and FMNH– reactions. Results in Table 1 indicate that FADH– can be used by C2 nearly as efficiently as FMNH–. The yields of DHPA obtained via Path A and B, 68 ± 3 and 74 ± 4%, were comparable to those for the FMNH– reaction (73 ± 4 and 82 ± 3%).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Our studies here have elucidated the detailed kinetic mechanism for the reactions of O2 with reduced flavin bound to the oxygenase component (C2) of HPAH from A. baumannii. The results and methods described can be used as prototypes for analyses of the two-component class of flavin-dependent oxygenases. These results clearly show that the oxygenation reaction of C2 occurs via C(4a)-oxygenated flavin intermediates, similar to the reaction of the single component aromatic flavoprotein hydroxylases, where existence of C(4a)-flavin intermediates is well documented (1, 2). It was previously found that C(4a)-hydroperoxyflavins reacted with aromatic substrates to form hydroxylated products in the reactions of p-hydroxybenzoate hydroxylase (3, 33), phenol hydroxylase (37–39), melilotate hydroxylase (40), anthranilate hydroxylase (41), 2-methyl-3-hydroxypyridine-5-carboxylic acid oxygenase (24–26), and 2-hydroxybiphenyl-3-monooxygenase (42). C(4a)-hydroperoxyflavins and C(4a)-hydroxyflavins were also detected in the oxygen reactions of HPAH from P. putida (16). The intermediates detected in the reaction of C2 are spectrally similar to those of the enzymes mentioned. However, the less common feature of C2 intermediates is that the C(4a)-hydroperoxyflavin and C(4a)-hydroxyflavin spectra are nearly identical; this characteristic has also been found in some mutant types of p-hydroxybenzoate hydroxylase (43). Partial resolution of spectra similar to those of C2-(C4a)-oxygenated intermediates was also obtained in studies of the reactions of 4-chlorophenol hydroxylase (9), the monooxygenase in the actinorhodin biosynthetic pathway (21, 22), and styrene monooxygenase (13) when the enzymes were mixed with reduced flavin and limited quantities of oxygen in the absence of substrate.

Although the reaction of C2 with O2 is similar to the reaction of single component aromatic hydroxylases with respect to using C(4a)-hydroperoxyflavin to hydroxylate the aromatic substrate, the overall kinetic mechanism of C2 is quite different (1–3). The first step of the reaction is binding of FMNH– to C2, followed by the reaction of the C -FMNH–2 complex with oxygen to form a quite stable C2-C(4a)-hydroperoxyflavin. Under conditions of catalytic turnover, an aromatic substrate binds to the preformed C2-C(4a)-hydroperoxyflavin intermediate (Path B in Fig. 10). This contrasts with the reactions of the single component aromatic hydroxylases where the aromatic compound must be bound to the enzyme prior to reduction and reaction with oxygen. The kinetic mechanism of C2 is remarkably similar to that for bacterial luciferases (Lux) in which the reaction of Lux-FMNH– with oxygen to form C(4a)-hydroperoxy-FMN occurs prior to binding of an aldehyde substrate (28). Although both C2 and Lux bind more tightly to the reduced than to the oxidized flavin, the Kd for the Lux-FMNH– complex is 80 nM (44), an order of magnitude smaller than that for C -FMNH–2 (1.2 µM). It is possible, however, that the C2-flavin complex becomes tighter after the reduced flavin is oxidized into C2-C(4a)-hydroperoxy FMN. The mechanism of C2 is also similar to that for the oxygenation half-reaction of cyclohexanone monooxygenase (CHMO), where cyclohexanone binds to the enzyme after formation of the FAD-C(4a)-peroxide (29).

The kinetic mechanism of C2 has similarities to the reaction of HPAH from P. putida. It was reported that in the reaction of O2 with the reduced flavoprotein plus the coupling protein of the P. putida HPAH, the rate of FAD-C(4a)-hydroperoxide formation is the same whether or not HPA was included in the oxygen-containing solution (1.1 x 106 M–1 s–1) (16). However, as shown above, the rate for formation of the C2-C(4a)-hydroperoxyflavin decreased from 1.1 x 106 M–1 s–1 to 4.8 x 104 M–1 s–1 when HPA was pre-bound to the C -FMNH–2 complex from A. baumannii (compare Figs. 3, 5, and 10). In the P. putida enzyme, it was also proposed that the reaction occurred via a pathway in which HPA bound to the oxygenase after the formation of C(4a)-hydroperoxy-FAD, similar to the reaction of C2 (Path B in Fig. 10). This was consistent with the rate of HPA binding to the reduced enzyme being rather slow (16). It is possible, however, that the P. putida enzyme is actually like the A. baumannii enzyme. The reported flavoprotein of P. putida might actually be a reductase regulated by HPA, similar to that from A. baumannii (20), whereas the coupling protein could be the oxygenase that receives reduced FAD from the reductase. Experiments to test this hypothesis have never been carried out. Thus, the lack of effect of HPA on the formation of the C(4a)-hydroperoxyflavin from P. putida HPAH could be caused by HPA not binding to the oxygenase until FADH– has bound.

Reduced flavin is reactive with oxygen. Therefore, to be effective, reduced flavin-utilizing enzymes such as C2 need to rapidly bind reduced flavin before auto-oxidation occurs. C2 was shown in this report to bind FMNH– very rapidly (Fig. 4) with an observed rate ≥200 s–1 (compare traces B and C in Fig. 6). Such a rate corresponds to a second order rate constant of at least 107 M–1 s–1, and this binding is quite tight (Kd of 1.2 µM under conditions studied). Therefore, the ability of C2 to catalyze reactions without being constantly bound to the cofactor like other flavoproteins can be explained by the preferential binding of the enzyme to the reduced rather than to the oxidized flavin. Similar binding properties were also observed for the oxygenase component (HpaB) of HPAH from E. coli; HpaB binds to FADH– with a Kd of 70 nM, whereas it binds to oxidized FAD with a Kd of 6 µM (45). Recently, a study of actinorhodin monooxygenase has shown that the oxygenase component, ActVA, binds to FMNH– with a Kd of 0.4 µM and to oxidized FMN with a Kd of 26 µM (21).

At high concentrations, HPA was found to form a dead-end complex with C2-C(4a)-hydroxyflavin and impede it from dehydrating to form the oxidized flavin (Figs. 8 and 9). Aromatic substrates were also found to bind to the C(4a)-hydroxy-FAD and inhibit the return to oxidized FAD in the reactions of several single component aromatic hydroxylases including phenol hydroxylase (34), 2-methyl-3-hydroxypyridine-5-carboxylic acid monooxygenase (24), and p-hydroxybenzoate hydroxylase (PHBH) (46). This type of substrate inhibition was also found in the reaction of P. putida HPAH (16). In the case of PHBH, it has been proposed that this inhibition is the natural consequence of the need for a conformational change from a solvent-free active site (for hydroxylation) to an "open" conformation for product and substrate exchange (3). Perhaps this inhibition is not important in cells, because cells are not likely to accumulate high concentrations of substrate that could cause such inhibition.

A unique property of C2 is the ability to use a variety of reduced flavin substrates; the enzyme works well with either FADH– or FMNH–, although less efficiently with reduced riboflavin (6, 15). Here we report that both C(4a)-hydroperoxy-FAD and C(4a)-hydroxy-FAD accumulated during the reaction of FADH– and C2 with oxygen (data not shown), implying that the reaction undergoes the same pathway as that of reduced FMN. Moreover, the kinetic constants for the reaction of FADH– and FMNH– are similar (Table 2), indicating that the reactivity of reduced FMN and FAD in each step of the C2 reaction is nearly the same. This also implies that C2 interacts with the reduced flavin primarily around the isoalloxazine where FAD and FMN share the same common structure. The flavin specificity of the HPAH from A. baumannii (C2) comes from the reductase, which binds more specifically and tightly to FMN (6, 20). This property contrasts to most other two-component monooxygenases, where the reductase is often less specific for the flavin whereas the oxygenase is specific for either FMNH– or for FADH–.

In conclusion, this study has elucidated the reaction mechanism of the oxygenase component (C2) of the enzyme HPAH from A. baumannii. The results clearly illustrate that C(4a)-oxygenated flavin intermediates are directly involved in the hydroxylation reaction. C2 binds to the reduced flavin (delivered from C1) in the initial step, reacts with oxygen to form the C2-C(4a)-hydroperoxyflavin, and finally binds HPA before hydroxylation occurs. This knowledge is needed to understand catalysis by the enzymes in this two-component class. This report will be followed by a subsequent article that explains in detail the transfer of the flavin between the two protein components of the enzyme.


   FOOTNOTES
 
* This work was supported by Grant GM64711 (to D. P. B.) from the National Institutes of Health, The Thailand Research Fund Grants RMU4880028 and RTA4780006, and Mahidol University (to P. C.). This study was also supported in part by a Research Team Strengthening Grant from BIOTECH (to Skorn Mongkolsuk). 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. Back

1 Recipient of a scholarship under the Commission on Higher Education Staff Development Project, Chulalongkorn University. Present address: Dept. of Biochemistry, Faculty of Dentistry, Chulalongkorn University. Back

2 To whom correspondence should be addressed: Dept. of Biochemistry and Center for Excellence in Protein Structure & Function, Faculty of Science, Mahidol University, Rama 6 Road, Bangkok 10400, Thailand. Tel.: 662-201-5596; Fax: 662-354-7174; E-mail: scpcy{at}mucc.mahidol.ac.th.

3 The abbreviations used are: HPAH, p-hydroxyphenylacetate hydroxylase; HPA, p-hydroxyphenylacetate; DHPA, 3,4-dihydroxyphenylacetate; C1, reductase component of HPAH from A. baumannii; C2, oxygenase component of HPAH from A. baumannii; HpaB, oxygenase component of HPA from E. coli; HPLC, high performance liquid chromatography. Back



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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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