Fungal AAO (aryl-alcohol oxidase) provides H2O2 for lignin biodegradation. AAO is active on benzyl alcohols that are oxidized to aldehydes. However, during oxidation of some alcohols, AAO forms more than a stoichiometric number of H2O2 molecules with respect to the amount of aldehyde detected due to a double reaction that involves aryl-aldehyde oxidase activity. The latter reaction was investigated using different benzylic aldehydes, whose oxidation to acids was demonstrated by GC-MS. The steady- and presteady state kinetic constants, together with the chromatographic results, revealed that the presence of substrate electron-withdrawing or electron-donating substituents had a strong influence on activity; the highest activity was with p-nitrobenzaldehyde and halogenated aldehydes and the lowest with methoxylated aldehydes. Moreover, activity was correlated to the aldehyde hydration rates estimated by 1H-NMR. These findings, together with the absence in the AAO active site of a residue able to drive oxidation via an aldehyde thiohemiacetal, suggested that oxidation mainly proceeds via the gem-diol species. The reaction mechanism (with a solvent isotope effect, 2H2Okred, of approx. 1.5) would be analogous to that described for alcohols, the reductive half-reaction involving concerted hydride transfer from the α-carbon and proton abstraction from one of the gem-diol hydroxy groups by a base. The existence of two steps of opposite polar requirements (hydration and hydride transfer) explains some aspects of aldehyde oxidation by AAO. Site-directed mutagenesis identified two histidine residues strongly involved in gem-diol oxidation and, unexpectedly, suggested that an active-site tyrosine residue could facilitate the oxidation of some aldehydes that show no detectable hydration. Double alcohol and aldehyde oxidase activities of AAO would contribute to H2O2 supply by the enzyme.
- aromatic aldehyde
- aryl-alcohol oxidase (AAO)
- enzyme kinetics
Lignin removal is a central process for recycling of the carbon fixed by photosynthesis in land ecosystems. Lignin protects plant polysaccharides against microbial attack and also represents a central issue in industrial utilization of lignocellulosic biomass for the production of renewable materials, chemicals and fuels in future lignocellulose biorefineries . High redox-potential peroxidases are characteristic of ligninolytic basidiomycetes (they are called white-rot fungi due to the colour of delignified wood) as recently confirmed by the comparison of fungal genomes [2,3]. Analysis of the white-rot fungal genomes available currently, including those of Phanerochaete chrysosporium and Pleurotus ostreatus, also confirmed the presence of genes encoding different types of oxidases that are responsible for providing the extracellular H2O2 required for lignin biodegradation as the peroxidase-oxidizing substrate. These include, among others, glyoxal oxidase (EC 184.108.40.206) first reported in P. chrysosporium, and AAO (aryl-alcohol oxidase; EC 220.127.116.11) mainly investigated in Pleurotus species [4,5].
Basidiomycetes also produce a variety of extracellular aromatic metabolites that have several roles in lignin degradation , and are also of interest for the flavour and fine-chemical industry, as in the case of aromatic aldehydes . Veratryl alcohol is synthesized by P. chrysosporium, where it acts as a lignin-peroxidase redox mediator, whereas the Pleurotus versatile peroxidase does not require such a mediator . Therefore, in Pleurotus species, the most abundant extracellular metabolite is p-anisaldehyde, together with its corresponding acid and alcohol, and some chlorinated metabolites (such as 3-chloro-p-anisaldehyde), with veratryl alcohol being found only in low levels . It has been shown that the above anisylic compounds are redox-cycled, with the participation of AAO and myceliar dehydrogenases, resulting in the continuous generation of extracellular H2O2 for lignin biodegradation at the expense of intracellular reducing power [9,10].
AAO is an extracellular FAD-containing enzyme in the GMC (glucose–methanol–choline) oxidoreductase family. It catalyses the oxidative dehydrogenation of a variety of aromatic (and some aliphatic polyunsaturated) alcohols with an α-carbon primary hydroxy group, with the concomitant reduction of O2 to H2O2 (Figure 1A), but it has also been suggested that it has the ability to oxidize aromatic aldehydes [11,12]. AAO was cloned for the first time in P. eryngii and, after that, has been expressed in Emericella nidulans (conidial state, Aspergillus nidulans) and Escherichia coli for further characterization of the enzyme and its alcohol oxidation mechanism [13–17]. However, to date the aldehyde reaction of AAO has remained basically unstudied.
In the present work we provide the first characterization of aldehyde oxidation by AAO using steady- and pre-steady state kinetics of wild-type AAO and some mutated variants. The kinetic studies have been accompanied by GC-MS and NMR analyses of substrates and products to obtain insights into the reaction mechanism of the enzyme.
Chemicals and commercial enzymes
p-Anisyl, veratryl, m- and p-chlorobenzyl, 3-chloro-p-anisyl, m- and p-fluorobenzyl alcohols, 2-4-hexadien-1-ol, benzaldehyde, p-anisaldehyde, veratraldehyde, m- and p-chlorobenz-aldehyde, 3-chloro-p-anisaldehyde, m- and p-fluorobenz-aldehyde, 3,4-difluorobenzaldehyde, p-nitrobenzaldehyde, 2,4-dinitrobenzaldehyde, 2,4-hexadienal, and benzoic, p-anisic, veratric, m-chlorobenzoic, 3-chloro-p-anisic, and m- and p-nitrobenzoic acids were obtained from Sigma–Aldrich. Glucose oxidase (from A. niger; type II) and aldehyde dehydrogenase (from Saccharomyces cerevisiae) were also obtained from Sigma–Aldrich. Amplex® Red was from Invitrogen. HRP (horseradish peroxidase) was from Roche. 5-Deazariboflavin was a gift from Dr G. Tollin (University of Arizona, Tucson, AZ, U.S.A.).
Recombinant AAO from P. eryngii was obtained by E. coli expression of the mature AAO cDNA (GenBank® accession number AF064069) followed by in vitro activation and purification as described previously . AAO concentrations were determined using the molar absorption coefficient (ε463= 11050 M−1·cm−1).
In addition to the above wild-type AAO, H502S, H546S and Y92F variants were prepared by PCR using the QuikChange site-directed mutagenesis kit (Stratagene). The cDNA of AAO cloned into the pFLAG1 vector (International Biotechnologies) was used as template and the following oligonucleotides, including mutations (underlined text) at the corresponding triplets (bold text), were used as primers (only direct constructions are shown): Y92F, 5′-GGTCTAGCTCTGTTCACTTCATGGTCATGATGCG-3′; H502S, 5′-GCCAACACGATTTTCAGCCCAGTTGGAACGGCC-3′; and H546S, 5′-CCCTTCGCGCCCAACGCAAGTACCCAAGGACCG-3′. The mutant plasmids were isolated and AAO cDNA was sequenced (GS-FLX sequencer from Roche) to confirm that only the designed mutations were introduced. The variants were produced as indicated above for wild-type AAO.
Steady-state kinetic constants for AAO aldehyde oxidation were calculated by monitoring H2O2 production using an HRP-coupled assay with Amplex® Red at 24 °C in air-saturated 0.1 M sodium phosphate buffer, pH 6. Turnover reactions were initiated by addition of AAO with an adder-mixer. In the presence of the H2O2 generated by AAO, HRP (6 units/ml) oxidizes Amplex® Red (60 μM) forming resorufin (ε563 = 52000 M−1·cm−1) . After non-linear fitting of data (three replicates) using SigmaPlot, mean and standard errors were obtained from the normalized Michaelis–Menten equation. p-Nitrobenzaldehyde (7 mM) oxidation by the H502S and H546S AAO variants and wild-type AAO were also estimated (as turnover numbers).
Oxidation rates of several alcohols by AAO were monitored both by aldehyde formation, using the molar absorbance coefficients , and by H2O2 generation, using an HRP-coupled assay with o-dianisidine, at 24 °C in air-saturated 0.1 M sodium phosphate buffer, pH 6. Assays were initiated by addition of AAO, and formation of the o-dianisidine oxidation product was monitored (ε460 6765 M−1·cm−1) . The results are presented as turnover rates (kcat in s−1) obtained as described above.
The enzymatic activity of commercial yeast aldehyde dehydrogenase on benzaldehyde, p-anisaldehyde and 4-nitrobenzaldehyde was measured at 25 °C in 0.1 M Tris/HCl, pH 8.0, containing 1 M KCl and 1 mM EDTA, by following the formation of NADH (ε340=6220 M−1·cm−1) from 5 mM NAD+.
For analysing the reaction products, 4 mM aldehyde solutions (1 mM in the cases of p-anisaldehyde and 3-chloro-p-anisaldehyde due to solubility constraints) were incubated with 0.3 μM AAO in 0.1 M sodium phosphate buffer, pH 6. After 3 h at 24 °C, under shaking, the reaction mixtures were acidified (pH 2–3) and liquid–liquid extracted with methyl t-butyl ether. Extracts were dried by evaporation and redissolved in chloroform. GC-MS analyses were performed directly, or after silylation with BSTFA [bis(trimethylsilyl)trifluoroacetamide] in the presence of pyridine. A Varian Star 3400 chromatograph equipped with a DB-5HT column (30 m×0.25 mm internal diameter; 0.1 μm film thickness) and coupled to an ion-trap detector (Varian Saturn 2000) was used. The oven was programmed to increase from 50 °C to 330 °C at 6 °C·min−1, and was held for 3 min. Both the temperature of the injector and the transfer line were 300 °C. Helium was used as the carrier gas at a flow rate of 2 ml/min. Compounds were identified by comparing their mass spectra with standards, which were also used to obtain response factors.
The hydration rates of aldehyde solutions (~10 μM) in 0.1 M sodium phosphate buffer, pH 6, prepared with 2H2O (isotopic purity 99.9%) was estimated by 1H-NMR using a Bruker Avance 500 MHz instrument. The signal of residual water proton (δH 4.701 p.p.m.) was used as internal reference for chemical shifts. For the hydration rate estimation, the signal of the H–C–(OH)2 proton in the gem-diol species was integrated and referred to that of the H–C=O proton of the aldehyde species. Spectra in DMSO (isotopic purity > 99.8%) were run as a reference, showing only the non-hydrated aldehyde.
Stopped-flow experiments were carried out on an Applied Photophysics SX17.MV spectrophotometer interfaced with an Acorn computer using the SX18.MV and Xscan software (from Applied Photophysics). Enzyme-monitored turnover experiments , as well as analysis of the reductive half-reactions under anaerobic conditions, were performed as described previously  in 0.1 M sodium phosphate buffer, pH 6, at 24 °C. Deconvolution of spectral data was performed by global analysis and numerical integration methods using the Pro-K software (Applied Photophysics).
For accurate estimation of observed rate constants (kobs) kinetic traces were recorded at 462 nm. These traces were fitted to the standard double-exponential or mono-exponential decays. kobs1 and kobs2 are the observed rate constants for the fast and slow phases of the reduction of the enzyme respectively. The kobs values at different substrate concentrations were fitted to either eqn (1) or eqn (2): (1) (2) where Kd is the dissociation constant for the enzyme–substrate complex, and kred and krev are the limiting rate and reverse rate of flavin reduction.
The solvent isotope effect in the reductive half-reaction of AAO with p-nitrobenzaldehyde was determined by measuring the rate constants using H2O and 2H2O. For assays in deuterated solvent, the reaction components were dissolved in deuterated 0.1 M sodium phosphate buffer, p2H 6. AAO was exhaustively dialysed against the deuterated buffer to remove all exchangeable protons.
Spectral characterization of AAO–acid complexes
Binding of aromatic acids was studied in 0.1 M sodium phosphate buffer, pH 6. The dissociation constants (Kd) for the complex of AAO with p-anisic acid and 3-chloro-p-anisic acid were determined from the absorption spectra during enzyme titration using eqn (3) for a 1:1 stoichiometry: (3) where ΔAbs is the absorbance difference values, E and L are the enzyme and ligand concentration respectively, and Kd is the dissociation constant . Finally, the complex of reduced AAO with p-anisic acid was prepared by photoreduction in the presence of EDTA and 5-deazariboflavin, as described previously .
Aldehyde oxidase activity during alcohol oxidation by AAO
The ability of AAO to continue oxidizing some aldehydes was deduced from the curves of H2O2 and aldehyde generated during reactions with eight aromatic alcohols (Figure 2). With p-anisyl alcohol, the two curves were similar and the final concentrations corresponded to the initial concentration of the alcohol, in agreement with the reaction stoichiometry (Figure 1A). However, with 3-chloro-p-anisyl alcohol the H2O2 concentration was nearly double that of 3-chloro-p-anisaldehyde, suggesting that a fraction of the aldehyde formed was oxidized further, releasing extra H2O2. Simultaneous alcohol oxidase and aldehyde oxidase activities were also observed in the reactions of AAO with other alcohols, e.g. with m- and p-fluorobenzyl alcohols (Table 1).
Steady-state kinetics with aldehydes
To characterize the aldehyde oxidase activity of AAO, a substrate specificity profile was constructed by determining apparent steady-state kinetic parameters with nine p- and m-substituted benzylic aldehydes, benzaldehyde and 2,4-hexadienal (Table 2) [the activity on 2,4-dinitrobenzaldehyde was too low (<1% that observed with p-nitrobenzaldehyde) to estimate kinetic constants (its water solubility was also low)]. Considering the AAO efficiency values, m-chlorobenzaldehyde was the best aldehyde substrate, whereas veratraldehyde was the worst substrate. AAO showed the highest kcat on p-nitrobenzaldehyde, whereas the three methoxylated benzaldehydes showed the lowest kcat values. Despite its low activity, AAO showed the highest affinity for p-anisaldehyde and 3-chloro-p-anisaldehyde.
Aldehyde oxidation was also investigated for three AAO variants obtained by site-directed mutagenesis of two histidine residues (His502 and His546) and one tyrosine (Tyr92) residue, which are conserved in the active sites of AAO and glucose oxidase . The two histidine residues seemed to be crucial for AAO oxidation of aldehydes, as the activities of the H502S and H546S variants on saturating concentrations of p-nitrobenzaldehyde were 0.22±0.02 min−1 and 0.11±0.01 min−1 respectively, compared with activity of 75.00±0.16 min−1 with the wild-type enzyme (their low activities prevented estimation of kinetic constants). On the other hand, removal of the phenolic hydroxy group of Tyr92 had distinctive effects on the oxidation of p-anisaldehyde and p-nitrobenzaldehyde (Table 3).
GC-MS identification of products from aldehyde oxidase activity
The reactions products of 12 aldehydes with AAO were analysed by GC-MS directly and after silylation, as shown in Figure 3 for the p-nitrobenzaldehyde reaction. The mass spectra of the product peaks revealed formation of p-nitrobenzoic acid by AAO. The corresponding acids were also found in the reactions of AAO with most of the aldehydes (see Supplementary Table S1 available at http://www.BiochemJ.org/bj/425/bj4250585add.htm for the characteristic MS fragments and the quantification of the acids formed). Under the reaction conditions used, the highest oxidation rates were obtained for p-chlorobenzaldehyde (48%), p-nitrobenzaldehyde (43%) and m-chlorobenzaldehyde (39%), followed by p-fluorobenzaldehyde (20%) and p-anisaldehyde (17%), no acids were detected from veratraldehyde and 2,4-dinitrobenzaldehyde and low oxidation rates (<7%) were obtained after incubation of AAO and the other five aldehydes.
NMR analysis of the gem-diol forms of the aldehyde substrates
To investigate the possible involvement of the gem-diol species in aldehyde oxidation, the hydration rates of the 12 aldehydes used as AAO substrates were estimated from their 1H-NMR spectra (Figure 4). The p-anisaldehyde spectrum showed only four signals (plus the residual water signal) corresponding to the methoxy, ortho (H2+6), meta (H3+5) and aldehyde (Hα) protons. The absence of the gem-diol benzylic proton signal (δH 6 p.p.m.) indicated that p-anisaldehyde was not hydrated. On the other hand, the p-nitrobenzaldehyde spectrum included the gem-diol signal (Hα′) together with that of the aldehyde form (Hα), as well as double signals for the ring protons for both the gem-diol (H2′+6′ and H3′+5′) and the aldehyde (H2+6 and H3+5) species, revealing partial hydration. Similar gem-diol signals were detected in all the other NMR spectra, with the exception of p-anisaldehyde and veratraldehyde, and the hydration rates were obtained by integrating the benzylic proton (Hα′ and Hα) signals (the chemical shifts and the hydration rates obtained are shown in Supplementary Table S2 available at http://www.BiochemJ.org/bj/425/bj4250585add.htm).
The hydration rates of all the aldehydes assayed were relatively modest, with the only exception being 2,4-dinitrobenzaldehyde. The highest values were obtained for the aromatic aldehydes bearing electron-withdrawing substituents, which facilitated the hydration reaction when located at the ortho or para positions, such as one or two nitro groups (20% and 83% hydration respectively), or at the meta position, such as chlorine (4%) or fluorine (3%) atoms. The presence of halogen atoms at the para position did not increase hydration with respect to benzaldehyde (1%), as expected from the above mentioned meta orientation of their electron-withdrawing effect, and the same was observed when comparing the hydration rates of 3,4-difluorobenzaldehyde and m-fluorobenzaldehyde (both being 3%). In contrast, the presence of methoxy substituents, acting as electron donors, especially at the para position, resulted in the complete absence of the gem-diol form, as found for p-anisaldehyde and veratraldehyde. Finally, the aliphatic polyunsaturated 2,4-hexadienal showed a very low hydration rate (below 1%).
Oxidation of benzylic aldehydes by yeast aldehyde dehydrogenase
The possible influence of different ring substituents on aromatic aldehyde oxidation by yeast aldehyde dehydrogenase, which does not oxidize the gem-diol forms, but acts on the non-hydrated aldehydes, was investigated. Interestingly, although the catalytic efficiency of aldehyde dehydrogenase oxidizing benzaldehyde was higher than on p-anisaldehyde, it was not increased further by the presence of an electron-withdrawing substituent in p-nitrobenzaldehyde (see Supplementary Figure S1 available at http://www.BiochemJ.org/bj/425/bj4250585add.htm) as found with AAO.
Stopped-flow spectrophotometric studies
By monitoring the redox state of the cofactor during AAO catalysis, information was obtained about the rate-limiting step. Using p-nitrobenzaldehyde as substrate, it was found that the enzyme is fully oxidized under air-saturated steady-state conditions (Figure 5). This corresponded to the first 1–2 min of the reaction (Figure 5, inset) when O2 availability is not a limiting factor, and this indicated that the reductive half-reaction by the aldehyde is much slower than the oxidative half-reaction by O2.
Pre-steady-state reduction of the oxidized enzyme by p-anisaldehyde, 3-chloro-p-anisaldehyde, veratraldehyde and p-nitrobenzaldehyde was investigated under anaerobic conditions (Figure 6, and Supplementary Figure S2 available at http://www.BiochemJ.org/bj/425/bj4250585add.htm). The first spectra, obtained after mixing AAO with the different aldehydes were in general different from those observed in the absence of substrate (Figure 6), showing displacement of the main flavin band (462 nm) and increased absorbance. These changes reflected formation of the oxidized enzyme complex with the aldehyde substrate (AAOox–S). The subsequent decrease at 462 nm corresponded to the reductive half-reaction. Formation of a charge-transfer complex between the reduced enzyme and the product (AAOred–P), characterized by a peak at 490 nm and a long-wavelength band, was also observed in the p-anisaldehyde reaction (Figure 6a), and the 3-chloro-p-anisaldehyde and veratraldehyde reactions (Supplementary Figure S2). The spectral course of the reductive half-reaction fitted to an irreversible two-step process without formation of any intermediate species for p-anisaldehyde, 3-chloro-p-anisaldehyde and veratraldehyde (with rate constants at saturating substrate concentrations of 1.32, 0.76 and 0.18 min−1 respectively) (Figure 6a and Supplementary Figure S2). However, AAO reduction by p-nitrobenzaldehyde best fitted a three-step process (Figure 6b), its initial phase being considerably faster (a rate constant over 90 min−1) than the reduction by the other aldehyde substrates (Figure 6c). The dependence of kobs on aldehyde concentration was determined with p-nitrobenzaldehyde (which showed 20% hydration and the highest turnover) and p-anisaldehyde (which showed low reactivity and no detectable gem-diol). These kobs dependencies exhibited hyperbolic saturation profiles with a good fit to eqn (1), suggesting an essentially irreversible flavin reduction and allowing the determination of transient constants for p-anisaldehyde (kred of 1.5±0.2 min−1 and Kd of 0.032±0.019 mM) and p-nitrobenzaldehyde (kred of 356±38 min−1 and Kd of 5.79±0.77 mM). In the case of p-nitrobenzaldehyde, AAO reduction is completed in a second slow phase (kred of approx. 0.74 min−1).
Using deuterated buffer, a solvent isotope effect with a 2H2Okred value of 1.5 (representing the ratio between kred in H2O and in 2H2O) was observed in the fast AAO reductive half-reaction with p-nitrobenzaldehyde.
Spectral characterization of AAO complexes with aromatic acids
It was found that aromatic acids bind at that point that the AAO absorption spectrum is modified (as illustrated in Supplementary Figure S3a, available at http://www.BiochemJ.org/bj/425/bj4250585add.htm, for the AAO titration with 3-chloro-p-anisic acid). Qualitatively similar difference spectra were also obtained with p-anisic, veratric, m-chlorobenzoic, m-fluorobenzoic and benzoic acids (results not shown). The tightest binding was observed with 3-chloro-p-anisic acid and p-anisic acid resulting in Kd of 31.5±0.1 μM and 94±3 μM respectively. The low AAO affinity for the other acids suggested Kd values in the millimolar range. The spectrum of the AAOred–(p-anisic acid) complex (Supplementary Figure S3b) was obtained by photoreduction of the AAOox–(p-anisic acid) complex. It was characterized by the displacement of the flavin maximum to approx. 490 nm and absorbance in the 520–600 nm region, being similar to that observed at the end of the stopped-flow experiments with p-anisaldehyde. This indicated that the fully reduced species observed after AAO reduction with this aldehyde, as well as with 3-chloro-p-anisaldehyde, is the AAOred–acid complex (Figures 6a and 6b). The other acids showed much looser binding and therefore no AAOred–acid complexes were detected at the end of AAO reduction (Supplementary Figure S2).
Aldehyde oxidase activity of AAO: aldehyde hydration and enzymatic oxidation
AAO typically oxidizes aromatic alcohols to the corresponding aldehydes, with the concomitant production of H2O2 [11,19] (Figure 1A). In the present study we demonstrated the ability of the enzyme to continue the oxidation of some alcohols to the corresponding acids, in agreement with preliminary 19F-NMR results . The aldehyde oxidase activity of AAO was confirmed by analysis of the mass spectra of the acids formed from a variety of aldehydes. The GC-MS results were in good agreement with the kcat values obtained, p-nitrobenzaldehyde and two chlorobenzaldehydes appeared to be the best AAO substrates in both cases.
In water solution, carbonyl compounds exist in equilibrium with their gem-diol forms. The hydration rates of the aromatic aldehydes are generally low compared with the aliphatic ones, and that of benzaldehyde is often considered as negligible. However, using 1H-NMR we demonstrated that all the aldehydes studied presented detectable hydration rates, with the exception of veratraldehyde and p-anisaldehyde. The rates could be correlated to the presence of electron-withdrawing or electron-donating substituents that stabilize or destabilize the gem-diol form.
Enzymatic oxidation of aldehydes often is catalysed using the unhydrated form, when an active-site residue forms a covalent adduct that favours oxidation. The most classical examples are aldehyde dehydrogenases, where a cysteine residue forms a thiohemiacetal linkage with the aldehyde substrate . Active-site serine, glutamate or lysine residues could also activate aldehydes by forming adducts under specific conditions . Alternatively, active-site metals can promote in situ hydration, as suggested for benzaldehyde oxidation by horse liver alcohol dehydrogenase  and mammalian aldehyde oxidases . The structure of the AAO active site (PDB entry 3FIM) shows that no residues susceptible to forming aldehyde adducts are present, and the enzyme does not contain metals . This, together with the observed correlation between the oxidation rates and the hydration of aldehydes, suggested that water is the activating molecule in aldehyde oxidation by AAO (Figure 1B). Oxidation of hydrated aldehydes has been reported for histidinol dehydrogenase , two mutated aldehyde dehydrogenases [28,29] and choline oxidase . Moreover, a positive correlation between aldehyde hydration and oxidation rates has also been found for Drosophila melanogaster alcohol oxidase . The similar activities obtained for benzaldehyde and p-nitrobenzaldehyde oxidation by yeast aldehyde dehydrogenase contrasted with those found for AAO, and supported different oxidation mechanisms by both enzymes.
The aldehyde oxidase activity of AAO is low compared with its alcohol oxidase activity . However, when the variable hydration rates were taken into account, the highest activity corresponded to the gem-diols of m-fluorobenzaldehyde, p-fluorobenzaldehyde and p-chlorobenzaldehyde and their corrected kcat values (100–130 s−1) were close to those obtained for the best alcohol substrates of AAO .
Mechanisms of aldehyde oxidation by AAO
If, as suggested above, the preferred aldehyde substrates of AAO are the gem-diol forms, which co-exist with the carbonyl ones, the mechanism of the rate-limiting reductive half-reaction would probably be similar to that recently reported for AAO oxidation of alcohols, characterized by the concerted transfer of the α-carbon hydride to flavin and the hydroxy proton to a catalytic base . The very slow AAO oxidation of 2,4-dinitrobenzaldehyde (despite its high hydration rate) can be explained because, once hydration has been promoted by the two nitro groups, their electron-withdrawing action will make the hydride transfer difficult. A similar situation was reported for oxidation of chlorinated acetaldehyde gem-diols by D. melanogaster alcohol dehydrogenase .
Cleavage of one of the gem-diol O–H bonds would involve an active-site histidine residue acting as a base, as suggested for conserved His502 or His546 (Supplementary Figure S4 available at http://www.BiochemJ.org/bj/425/bj4250585add.htm) in alcohol oxidation by AAO . This was confirmed by the analysis of the H502S and H546S AAO variants, which showed much lower activities on p-nitrobenzaldehyde than the wild-type enzyme. The 340-fold activity decrease of H502S was in the range of that reported for p-anisyl alcohol oxidation , but that of H546S (670-fold) was higher than that reported for alcohol oxidation by over 60-fold , indicating that this residue plays a more important role in the AAO-mediated oxidation of gem-diols compared with alcohols.
A concerted cleavage of gem-diol C–H and O–H bonds is suggested by the solvent kinetic isotope effect for the AAO reduction by p-nitrobenzaldehyde (2H2Okred 1.5), which was similar to that reported for its reduction by p-anisyl alcohol and 2,4-hexadien-1-ol . The above AAO oxidation mechanism contrasts with the sequential mechanism, involving previous alcohol activation to the alkoxide without a detectable solvent isotope effect, as reported for choline oxidase [34,35].
Substitution of Tyr92, which is located over the AAO flavin ring (see Supplementary Figure S4), for a phenylalanine residue did not strongly affect the oxidation of p-nitrobenzaldehyde, in agreement with previous alcohol oxidation results . However, the Y92F mutation caused a 5-fold reduction in the p-anisaldehyde kcat value, suggesting that the phenolic hydroxy group has some effect on the activation of aldehydes that are not spontaneously hydrated. Alternatively, Tyr92 could promote the oxidation of a minor gem-diol form below the detection level of 1H-NMR. Oxidation of small molar fractions of preformed gem-diol and enzyme-aided hydration have been proposed as two alternative hypotheses to explain aldehyde oxidation by some alcohol dehydrogenases [26,36] and histidinol dehydrogenase .
AAO is a versatile oxidase providing H2O2 for lignin degradation
A difference between AAO and other oxidases/dehydrogenases which catalyse oxidation of alcohols to acids concerns the eventual release of the aldehyde intermediates. As the specific reaction catalysed by choline oxidase is the conversion of choline into glycine betaine, the intermediate betaine aldehyde is not released from the enzyme under physiological conditions (although the enzyme is able to oxidize it when it is exogenously added) . The same occurs in the histidinol dehydrogenase reaction . In contrast, the physiological role of AAO is generation of extracellular H2O2, in a double-redox-cycle (alcohol↔aldehyde and aldehyde↔acid), where the oxidation products are not incorporated to metabolic pathways but reduced back by mycelium-associated dehydrogenases for the continuous supply of the peroxide required for lignin biodegradation [10,39].
Interestingly, those basidiomycetes producing AAO, typically Pleurotus and Bjerkandera species [11,40], also synthesize methoxylated and halogenated aromatic compounds, which can be detected in the extracellular medium (as alcohols, aldehydes and/or acids) [9,41]. p-Anisylic and 3-chloro-p-anisylic compounds are among the most abundant Pleurotus  and Bjerkandera [42,43] metabolites. On the other hand, although few nitroaromatics are synthesized by basidiomycetes compared with the extended production of haloaromatics , p-nitrobenzaldehyde biosynthesis has been reported  and p-nitrosalicylic acid was very recently found in Pleurotus cultures (Giovanni Sannia, personal communication). It seems that AAO has evolved to use differently substituted benzylic metabolites for H2O2 production. Depending on the compound available, AAO will behave basically as an alcohol oxidase (e.g. in the presence of veratrylic and anisylic metabolites), as an aldehyde oxidase (e.g. in the presence of p-nitrobenzylic metabolites) or as a double-oxidase being able to transform alcohols into the corresponding acids (e.g. in the presence of 3-chloro-p-anisylic metabolites) (reactions shown in Figures 1A and 1B respectively). This substrate versatility would help AAO to provide a maximal supply of H2O2 for lignin biodegradation under variable physiological and environmental conditions.
All of the authors performed the research and discussed the results obtained. Patricia Ferreira specifically contributed with the kinetic studies and design of the enzymatic part of the study; Aitor Hernandez with the preparation and characterization of the mutated variants; Beatriz Herguedas with some spectroscopic studies; Jorge Rencoret and Ana Gutiérrez with the GC-MS analyses; Jesús Jiménez-Barbero with the NMR analyses; María Jesús Martínez in enzyme characterization; and Milagros Medina in the interpretation of the pre-steady state kinetics and other results. Angel Martínez was responsible for the co-ordination of the study.
This work was supported by the Spanish Biotechnology Programme [grants numbers BIO2007-65890-C02-01, BIO2008-01533] and the Biorenew project of the European Union [grant number NMP2-CT-2006-026456]. P. F. was a recipient of a Programa Juan de la Cierva contract from the Spanish Ministry of Science and Innovation, and J. R. was a recipient of an I3P Fellowship of the Consejo Superior de Investigaciones Científicas.
The authors thank Antonio Romero (Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain) for providing access to the AAO crystal structure, and Vicenzo Lettera and Giovanni Sannia (University of Naples, Italy) for information on the Pleurotus nitroaromatic metabolites.
Abbreviations: AAO, aryl-alcohol oxidase; HRP, horseradish peroxidise; AAOox, oxidized AAO; AAOred, reduced AAO
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