Control of C4a-Hydroperoxyflavin Protonation in the Oxygenase Component of <i>p</i>‑Hydroxyphenylacetate-3-hydroxylase

The protonation status of the peroxide moiety in C4a-(hydro)­peroxyflavin of <i>p</i>-hydroxyphenylacetate-3-hydroxylase can be directly monitored using transient kinetics. The p<i>K</i>_a for the wild-type (WT) enzyme is 9.8 ± 0.2, while the values for the H396N, H396V, and H396A variants are 9.3 ± 0.1, 7.3 ± 0.2, and 7.1 ± 0.2, respectively. The hydroxylation efficiency of these mutants is lower than that of the WT enzyme. Solvent kinetic isotope effect studies indicate that proton transfer is not the rate-limiting step in the formation of C4a-OOH. All data suggest that His396 may act as an instantaneous proton provider for the proton-coupled electron transfer that occurs before the transition state of C4a-OOH formation

The Transfer of Reduced Flavin Mononucleotide from LuxG Oxidoreductase to Luciferase Occurs via Free Diffusion

Bacterial luciferase (LuxAB) is a two-component flavin mononucleotide (FMN)-dependent monooxygenase that catalyzes the oxidation of reduced FMN (FMNH^–) and a long-chain aliphatic aldehyde by molecular oxygen to generate oxidized FMN, the corresponding aliphatic carboxylic acid, and concomitant emission of light. The LuxAB reaction requires a flavin reductase to generate FMNH^– to serve as a luciferin in its reaction. However, FMNH^– is unstable and can react with oxygen to generate H₂O₂, so that it is important to transfer it efficiently to LuxAB. Recently, LuxG has been identified as a NADH:FMN oxidoreductase that supplies FMNH^– to luciferase <i>in vivo</i>. In this report, the mode of transfer of FMNH^– between LuxG from <i>Photobacterium leiognathi</i> TH1 and LuxABs from both <i>P. leiognathi</i> TH1 and <i>Vibrio campbellii</i> (<i>Pl</i>LuxAB and <i>Vc</i>LuxAB, respectively) was investigated using single-mixing and double-mixing stopped-flow spectrophotometry. The oxygenase component of <i>p</i>-hydroxyphenylacetate hydroxylase (C2) from <i>Acinetobacter baumannii</i>, which has no structural similarity to LuxAB, was used to measure the kinetics of release of FMNH^– from LuxG. With all FMNH^– acceptors used (C₂, <i>Pl</i>LuxAB, and <i>Vc</i>LuxAB), the kinetics of FMN reduction on LuxG were the same, showing that LuxG releases FMNH^– with a rate constant of 4.5–6 s^–1. Our data showed that the kinetics of binding of FMNH^–to <i>Pl</i>LuxAB and <i>Vc</i>LuxAB and the subsequent reactions with oxygen were the same with either free FMNH^– or FMNH^– generated <i>in situ</i> by LuxG. These results strongly suggest that no complexes between LuxG and the various species are necessary to transfer FMNH^– to the acceptors. The kinetics of the overall reactions and the individual rate constants correlate well with a free diffusion model for the transfer of FMNH^– from LuxG to either LuxAB

Proton-Coupled Electron Transfer and Adduct Configuration Are Important for C4a-Hydroperoxyflavin Formation and Stabilization in a Flavoenzyme

Determination of the mechanism of dioxygen activation by flavoenzymes remains one of the most challenging problems in flavoenzymology for which the underlying theoretical basis is not well understood. Here, the reaction of reduced flavin and dioxygen catalyzed by pyranose 2-oxidase (P2O), a flavoenzyme oxidase that is unique in its formation of C4a-hydroperoxyflavin, was investigated by density functional calculations, transient kinetics, and site-directed mutagenesis. Based on work from the 1970s–1980s, the current understanding of the dioxygen activation process in flavoenzymes is believed to involve electron transfer from flavin to dioxygen and subsequent proton transfer to form C4a-hydroperoxyflavin. Our findings suggest that the first step of the P2O reaction is a single electron transfer coupled with a proton transfer from the conserved residue, His548. In fact, proton transfer enhances the electron acceptor ability of dioxygen. The resulting ·OOH of the open-shell diradical pair is placed in an optimal position for the formation of C4a-hydroperoxyflavin. Furthermore, the C4a-hydroperoxyflavin is stabilized by the side chains of Thr169, His548, and Asn593 in a “face-on” configuration where it can undergo a unimolecular reaction to generate H₂O₂ and oxidized flavin. The computational results are consistent with kinetic studies of variant forms of P2O altered at residues Thr169, His548, and Asn593, and kinetic isotope effects and pH-dependence studies of the wild-type enzyme. In addition, the calculated energy barrier is in agreement with the experimental enthalpy barrier obtained from Eyring plots. This work revealed new insights into the reaction of reduced flavin with dioxygen, demonstrating that the positively charged residue (His548) plays a significant role in catalysis by providing a proton for a proton-coupled electron transfer in dioxygen activation. The interaction around the N5-position of the C4a-hydroperoxyflavin is important for dictating the stability of the intermediate

Tuned Amperometric Detection of Reduced β‑Nicotinamide Adenine Dinucleotide by Allosteric Modulation of the Reductase Component of the <i>p</i>‑Hydroxyphenylacetate Hydroxylase Immobilized within a Redox Polymer

We report the fabrication of an amperometric NADH biosensor system that employs an allosterically modulated bacterial reductase in an adapted osmium­(III)-complex-modified redox polymer film for analyte quantification. Chains of complexed Os­(III) centers along matrix polymer strings make electrical connection between the immobilized redox protein and a graphite electrode disc, transducing enzymatic oxidation of NADH into a biosensor current. Sustainable anodic signaling required (1) a redox polymer with a formal potential that matched the redox switch of the embedded reductase and avoided interfering redox interactions and (2) formation of a cross-linked enzyme/polymer film for stable biocatalyst entrapment. The activity of the chosen reductase is enhanced upon binding of an effector, i.e. <i>p</i>-hydroxy-phenylacetic acid (<i>p</i>-HPA), allowing the acceleration of the substrate conversion rate on the sensor surface by in situ addition or preincubation with <i>p</i>-HPA. Acceleration of NADH oxidation amplified the response of the biosensor, with a 1.5-fold increase in the sensitivity of analyte detection, compared to operation without the allosteric modulator. Repetitive quantitative testing of solutions of known NADH concentration verified the performance in terms of reliability and analyte recovery. We herewith established the use of allosteric enzyme modulation and redox polymer-based enzyme electrode wiring for substrate biosensing, a concept that may be applicable to other allosteric enzymes

<i>p</i>‑Hydroxyphenylacetate 3‑Hydroxylase as a Biocatalyst for the Synthesis of Trihydroxyphenolic Acids

Trihydroxyphenolic acids such as 3,4,5-trihydroxycinnamic acid (3,4,5-THCA) <b>4c</b> and 2-(3,4,5-trihydroxyphenyl)­acetic acid (3,4,5-THPA) <b>2c</b> are strong antioxidants that are potentially useful as medicinal agents. Our results show that <i>p</i>-hydroxyphenylacetate (HPA) 3-hydroxylase (HPAH) from <i>Acinetobacter baumannii</i> can catalyze the syntheses of 3,4,5-THPA <b>2c</b> and 3,4,5-THCA <b>4c</b> from 4-HPA <b>2a</b> and <i>p</i>-coumaric acid <b>4a</b>, respectively. The wild-type HPAH can convert 4-HPA <b>2a</b> completely into 3,4,5-THPA <b>2c</b> within 100 min (total turnover number (TTN) of 100). However, the wild-type enzyme cannot efficiently synthesize 3,4,5-THCA <b>4c</b>. To improve the efficiency, the oxygenase component of HPAH (C₂) was rationally engineered in order to maximize the conversion of <i>p</i>-coumaric acid <b>4a</b> to 3,4,5-THCA <b>4c</b>. Results from site-directed mutagenesis studies showed that Y398S is significantly more effective than the wild-type enzyme for the synthesis of 3,4,5-THCA <b>4c</b>; it can catalyze the complete bioconversion of <i>p</i>-coumaric acid <b>4a</b> to 3,4,5-THCA <b>4c</b> within 180 min (TTN ∼ 23 at 180 min). The yield and stability of 3,4,5-THPA <b>2c</b> and 3,4,5-THCA <b>4c</b> were significantly improved in the presence of ascorbic acid. Thermostability studies showed that the wild-type C₂ was very stable and remained active after incubation at 30, 35, and 40 °C for 24 h. Y398S was moderately stable because its activity was retained for 24 h at 30 °C and for 15 h at 35 °C. Transient kinetic studies using stopped-flow spectrophotometry indicated that the key improvement in the reaction of Y398S with <i>p</i>-coumaric acid <b>4a</b> lies within the protein–ligand interaction. Y398S binds to <i>p</i>-coumaric acid <b>4a</b> with higher affinity than the wild-type enzyme, resulting in a shift in equilibrium toward favoring the productive coupling path instead of the path leading to wasteful flavin oxidation

Electron-donor substrates modeled in the active site of <i>Am</i>PDH.

<p>Carbons are colored yellow and numbered. Apostrophes indicate carbons on the non-reducing end sugar. Hydrogen atoms have been omitted for the purpose of clarity.</p

Comparison of related GMC dehydrogenases and oxidases.

a<p><i>Am</i>PDH, <i>A. meleagris</i> pyranose dehydrogenase; <i>Pc</i>CDH, <i>P. chrysosporium</i> cellobiose dehydrogenase; <i>Tm</i>P2O, <i>T. multicolor</i> pyranose 2-oxidase; <i>An</i>GOX, <i>A. niger</i> glucose 1-oxidase; <i>Ag</i>CHO; <i>A. globiformis</i> choline oxidase.</p>b<p>The <i>Ag</i>CHO mutant S101A shows increased efficiency in the oxidative half-reaction <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0053567#pone.0053567-Finnegan2" target="_blank">[68]</a>, stressing that the function of this side chain is different in <i>Ag</i>CHO compared with the sugar-oxidizing enzymes.</p

Data collection and crystallographic refinement statistics for <i>Am</i>PDH.

a<p>The outer shell statistics of the reflections are given in parentheses. Shells were selected as defined in <i>XDS</i><a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0053567#pone.0053567-Kabsch1" target="_blank">[54]</a> by the user.</p>b<p><i>R_sym</i> = [Σ<i>_hkl</i> Σ<i>_i</i> |<i>I–<i></i>|/Σ<i>_hkl</i> Σ<i>_i</i> |<i>I</i>| ] x 100%.</i></p><i>c<p>CC(1/2) = Percentage of correlation between intensities from random half-datasets. Values given represent correlations significant at the 0.1% level <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0053567#pone.0053567-Karplus1" target="_blank">[66]</a>.</p>d<p><i>R_factor</i> = Σ<i>_hkl</i> | |F_o|–|F_c| |/Σ<i>_hkl</i> |F_o|.</p>e<p>As determined by <i>MolProbity</i><a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0053567#pone.0053567-Chen1" target="_blank">[62]</a>.</p></i

Overall structure of <i>Am</i>PDH and similarity to AAO.

<p>Ribbon drawing of <i>Am</i>PDH (a) and aryl-alcohol oxidase (b), showing α-helices as spirals and β-strands as arrows. The covalently bound flavin cofactor is depicted as a stick model with carbon atoms in yellow. An overlay picture of the active site in <i>Am</i>PDH (beige carbon atoms) and AAO (green carbon atoms) is shown in (c).</p

Absorption spectra of reduced <i>Am</i>PDH reacting with molecular oxygen.

<p>Oxidation of PDH by oxygen at air-saturation as a function of time at 25°C. Line 1: reduced PDH at start of the reaction which lacks the flavin absorption shoulder around 490 nm; line 5, the spectrum of partially oxidized PDH (∼40% oxidation); line 10, the absorption spectrum of partially oxidized PDH after the reaction proceeded for 15 h.</p