Screening and Evaluation of New Hydroxymethylfurfural Oxidases for Furandicarboxylic Acid Production

HMFO is the only enzyme described to date that can catalyze by itself the three consecutive oxidation steps to produce FDCA from HMF. Unfortunately, only one HMFO enzyme is currently available for biotechnological application. This availability is enlarged here by the identification, heterologous production, purification, and characterization of two new HMFOs, one from Pseudomonas nitroreducens and one from an unidentified Pseudomonas species. Compared to the previously known Methylovorus HMFO, the new enzyme from P. nitroreducens exhibits better performance for FDCA production in wider pH and temperature ranges, with higher tolerance for the hydrogen peroxide formed, longer half-life during oxidation, and higher yield and total turnover numbers in long-term conversions under optimized conditions. All these features are relevant properties for the industrial production of FDCA. In summary, gene screening and heterologous expression can facilitate the selection and improvement of HMFO enzymes as biocatalysts for the enzymatic synthesis of renewable building blocks in the production of bioplastics.


RESULTS
Identification of new HMF-oxidizing enzymes. BLAST of the amino acid sequence of MetspHMFO (UniProt E4QP00) against the nonredundant protein sequences in GenBank revealed 41 entries with identities of Ͼ45%, all of them from the phylum Proteobacteria (Fig. 2). The largest numbers of HMFOs were found in the Pseudomonas (12 sequences), Bradyrhizobium (9 sequences), and Methylobacterium (8 sequences) genera, and those with the highest homology with MetspHMFO were HMFOs from Methylophylus, Pseudomonas, and Burkholderia species, with more than 60% identity with the query (Table S1 in the supplemental material).
All of the sequences described above conserve the ADP-binding domain and the consensus sequences PS00623 and PS00624, typical of the GMC oxidoreductase superfamily ( Fig. S1) (1). A more detailed analysis of conserved active-site amino acids (Fig.  3) reveals that all of them include the same histidine and asparagine residues, MetspHMFO His467 and Asn511, at the catalytic positions described for the latter enzyme (12). Asn102, Trp369, and Trp466 are also fully conserved, while substitutions are often observed at Met103, involved in the correct orientation of the substrate with respect to the active site, and at Val367, described as a site for improving the activity on carboxyl-containing substrates (12). From this repertoire of enzymes, four HMFO sequences from the two main HMFO clusters (Fig. 2)-one from P. nitroreducens (PseniHMFO), one from Pseudomonas sp. strain 11/12A (PsespHMFO), one from Xanthobacter sp. strain 126 (XanspHMFO), and one from Bradyrhizobium arachidis (BraarHMFO)-were selected and optimized for E. coli expression to produce them as recombinant proteins, together with MetspHMFO.
Expression and purification of two new HMFOs. The four optimized genes (Table  S2) were cloned into the pET23b and/or the His-tagged pET28a plasmid and transformed into E. coli C41(DE3). XanspHMFO and BraarHMFO could not be expressed using either the pET23b or the pET28a vector, while PseniHMFO and PsespHMFO were purified and characterized after pET28a and pET23b expression, respectively. To evaluate whether the His tag affects the properties of HMFOs, the optimized gene encoding MetspHMFO (Table S2) was also cloned into both plasmids, and the tagged/untagged forms were characterized in parallel. The four proteins were expressed as soluble, active enzymes by slow induction at 16°C during 72 to 96 h.
Untagged MetspHMFO and PsespHMFO were purified to electrophoretic homogeneity ( Fig. S2a and c, respectively) by two anion exchange chromatographic steps (see conditions in Table S3), yielding 34 and 10 mg of protein per liter of culture, respectively (Table 1). For the His-tagged proteins, anionic chromatography after the affinity chromatographic step provided pure enzyme ( Fig. S2b and d), with yields of 17 and 47 mg of protein per liter of culture of MetspHMFO His and PseniHMFO His , respectively ( Table 1). The specific activities of the purified enzymes were 3.3 to 3.6 U · mg Ϫ1 for MetspHMFO (similar for the untagged and the His-tagged forms), 2.0 U · mg Ϫ1 for PsespHMFO, and 0.2 U · mg Ϫ1 for PseniHMFO His . Vanillyl alcohol was used for the activity estimations described above, since it is oxidized with the highest catalytic efficiency, as described below.
The molecular weights (MW) of the different HMFOs, measured by matrix-assisted laser desorption and ionization-time of flight (MALDI-TOF) mass spectrometry, were ϳ57 kDa (Table 2), in good agreement with the theoretical masses calculated from their amino acid sequences (Table S1). This indicates that HMFOs are not purified as covalently bound dimeric enzymes. Also, the experimental isoelectric point (pI) values of 6.0 to 6.5 were similar to the theoretical ones (as shown in the tables mentioned above).
HMFO spectral properties. The secondary structures of the enzymes were evaluated by circular dichroism (CD) (Fig. S3a). The far-UV spectra showed minimums at ϳ222 nm, typical of ␣-helix structures, being more intense in the MetspHMFO and PsespHMFO spectra. The contents in ␣-helixes (27 to 47%), ␤-sheets (11 to 20%), and turns (20 to 24%) were obtained by deconvolution of the CD spectra (Table S4). As shown for MetspHMFO His , the molar ellipticity per residue (MER) intensity revealed an ␣-helix decrease (from 37% to 27%) and an increase of unordered structure (from 26% to 37%) in the His-tagged enzymes.
The enzymes showed UV-visible spectra characteristic of FAD-containing proteins ( Table 2 and Fig. S3b). Two peaks at 382 to 387 nm (band II) and 456 to 457 nm (band FIG 3 Conservation of residues around the HMFO active site after alignment of sequences included in the phylogenetic tree. Nonpolar residues are colored in brown, aromatic residues are colored in red, basic residues are colored in blue, and acidic residues are colored in green. The four new HMFO sequences optimized for gene expression in E. coli and that of MetspHMFO are labeled in orange and red, respectively. I) with a shoulder at 483 nm and A 279 /A 457 ratios of ϳ10 were observed, indicating the proper incorporation of the cofactor. The calculated extinction coefficients at the band I maximums ( band I ) of MetspHMFO, MetspHMFO His , PsespHMFO, and PseniHMFO His were 11,271, 11,667, 11,432, and 12,060 M Ϫ1 cm Ϫ1 , respectively. These values were used to estimate the corresponding enzyme concentrations.
Temperature and pH stability. As an estimation of thermostability, the melting temperature (T m ) for each enzyme was calculated from the changes of FAD fluorescence using the ThermoFAD assay (26). The denaturation curves indicated a T m of 48°C for the enzyme from Methylovorus (both MetspHMFO and MetspHMFO His ), while Ps-espHMFO and PseniHMFO appeared to be 8°C and 4°C less stable, respectively ( Fig. 4a and Table 2).
To determine the range of pHs in which the enzymes are active, we measured the residual activity on vanillyl alcohol after 72 h of incubation under different pH conditions. The enzyme from Methylovorus (both MetspHMFO and MetspHMFO His ) was quite stable at pH 6 to 9, conserving 70 to 85% activity ( Fig. 4b). At pH 10, the activities of MetspHMFO and MetspHMFO His decreased to only 5% and 24% activity, respectively, while PseniHMFO His kept ϳ65 to 70% of its initial activity between pH 6 and 10. The range in which PsespHMFO was stable was slightly narrower, pH 6 to 8, with only ϳ10% activity and total activity loss at pH 9 and pH 10, respectively. Outside these pH ranges, the activity of all the enzymes decreased abruptly in the first 24 h (data not shown).
Optimal temperature and pH. The pH and temperature optima were also calculated with vanillyl alcohol as the substrate. PsespHMFO and PseniHMFO His attained their maximal activities at 35 to 40°C and 40 to 45°C, respectively, and decreased to 50% below 15°C and 25°C, respectively, and above 50°C for both (Fig. 4c). Similar profiles  were observed for MetspHMFO, which attained its maximal activity at 45°C and kept more than 80% activity between 25 and 45°C (20 to 50°C for MetspHMFO His ). With regard to pH, none of the enzymes were active below pH 5 (Fig. 4d). The optimum for MetspHMFO (both for the untagged and His-tagged enzyme) was at pH 7.0, with the enzyme retaining ϳ70% of its activity at pH 6.5 to 8.0. Similarly, PsespHMFO showed its maximal activity at pH 7.5 to 8.0 and still showed 80% of its activity at pH 7 to 12. Although at basic pHs, the initial rates for MetspHMFO and PsespHMFO were high, the velocities decayed abruptly after a few seconds due to enzyme inactivation. However, PseniHMFO His exhibited a quite different pH behavior, since it was more active at basic pH, with the maximal value at pH 10.0. Below pH 9.0 and above pH 11.0, its activity decreased to less than 80%.
Effect of H 2 O 2 on HMFOs' stability. Since during HMFO activity, hydrogen peroxide is produced from O 2 reduction, we evaluated the effect that different amounts of H 2 O 2 (0.5 to 7.8 mM) had on the enzymes' stability. The results for MetspHMFO and MetspHMFO His suggest that the His tag has a positive effect on enzyme stability, even in the absence of H 2 O 2 ( Fig. 5a and b). Also, PseniHMFO His was more stable to peroxide exposure than MetspHMFO, keeping around 70% of its activity after 96 h in all the concentrations tested (Fig. 5d). In contrast, PsespHMFO was the least stable enzyme (Fig. 5c). In fact, low peroxide amounts (0.5 to 2.6 mM) had a slight stabilizing effect on PsespHMFO, while at the highest concentration, its activity decreased to only 10%.
Kinetic parameters for oxidation of HMF and its DFF and FFCA derivatives. The kinetic parameters for oxidation of vanillyl alcohol and furfuryl substrates HMF, DFF, and FFCA were determined for the four enzyme preparations ( Table 3). The untagged and His-tagged MetspHMFO showed similar k cat and K m values for vanillyl alcohol (measured by direct spectrophotometric estimation of the aldehyde product). PsespHMFO oxidized vanillyl alcohol with a catalytic efficiency (k cat /K m ) similar to that of the enzyme from Methylovorus sp., while PseniHMFO His was less efficient in oxidizing this substrate (with ϳ28-fold-lower values than the other enzymes).
Oxidation of HMF and DFF was followed by measuring H 2 O 2 release, using a coupled assay with horseradish peroxidase (HRP) at pH 7.0 (which is close to the HRP optimum). HMF was the best furfuryl substrate for all the enzymes. MetspHMFO and PsespHMFO showed similar k cat values, with a significant increase in the K m for PsespHMFO, while for PseniHMFO His , the K m was lower. The efficiencies for HMF oxidation by PsespHMFO and PseniHMFO were ϳ4-fold lower than for MetspHMFO. Concerning DFF oxidation, it was in all cases 10-to 14-fold slower than that of HMF, showing that the PsespHMFO enzyme had a positive cooperative effect (Hill coefficient, n H ϭ 2.4) and, again, a K m value significantly higher than for the rest of the enzymes.
Due to the very slow FFCA oxidation, the kinetic parameters for this compound could not be calculated in a continuous assay. Therefore, the production of FDCA was followed by high-performance liquid chromatography (HPLC) during 48 h of incubation with increasing concentrations of FFCA at 25°C and pH 7.0. The enzyme from Methylovorus showed strong substrate inhibition at FFCA concentrations higher than 3 mM. This made it impossible to assess the kinetic parameters, as no saturation by FFCA was attained and only k obs (observed rate constant)/concentration values were estimated for comparison of efficiencies. The FFCA inhibitory effect was much milder for the enzymes from Pseudomonas, and saturation was attained due to the lower K m values. These results showed that the FFCA oxidation efficiencies of PseniHMFO His and PsespHMFO are 1.4-and 2.2-fold higher, respectively, than those of MetspHMFO. However, the differences in catalytic efficiencies for FFCA oxidation with regard to those for HMF and DFF were significantly lower in PseniHMFO His and PsespHMFO than in MetspHMFO. This suggests a more efficient overall oxidation of HMF to produce FDCA by the Pseudomonas enzymes.
HMF conversion: effects of pH and temperature. Taking into account the similar kinetic parameters of MetspHMFO and MetspHMFO His and the higher stability under exposure to H 2 O 2 of the latter form, the following experiments with this enzyme were performed with the His-tagged form. As for MetspHMFO His , the enzymes from Pseudomonas were able to oxidize HMF to FDCA, catalyzing the three reaction steps. Taking into account their pH and temperature profiles ( Fig. 4c and d), the conditions for HMF conversion by each enzyme were investigated in 72-h reactions (Fig. 6). Maximal FDCA production was achieved at pH 7.0, pH 7.5, and pH 8.0 for MetspHMFO His , PsespHMFO, and PseniHMFO His , respectively (Fig. 6a). Under optimal pH conditions, MetspHMFO His and PsespHMFO showed half-lives (t 1/2 ) of 55 and 38 h, respectively, while PseniHMFO His was more stable, with a remarkable t 1/2 of ϳ100 h (Fig. 6b). Outside their optimal pH conditions, MetspHMFO His and PsespHMFO had reduced half-lives, while PseniHMFO His was quite stable in the range of pH 6 to 8 (Fig. S4, left).
Concerning temperature, PseniHMFO His and PsespHMFO kept maximal FDCA production (around 45% and 30%, respectively) in the range of 16 to 28°C, while MetspHMFO His showed maximal production in the 24 to 28°C range (Fig. 6c) (all reactions at optimal pH). At 28°C, the t 1/2 values for the three HMFOs were 61, 27, and 143 h for MetspHMFO His , PsespHMFO, and PseniHMFO His , respectively (Fig. 6d). Above 28°C, MetspHMFO His and PsespHMFO had decreased FDCA yields (Fig. 6c) due to enzyme inactivation, as shown by the t 1/2 decreases (Fig. S4, right). PseniHMFO His was the most thermostable enzyme for FDCA production (Fig. S4f), with t 1/2 values of more FDCA production: kinetics and long-term conversions. Taking into account the results described above, oxidation of HMF by the different HMFOs was followed first during 1 h at 25°C and the optimal pH of each enzyme (Fig. 7). For all of them, the oxidation of HMF was faster than that of DFF [k (HMF¡DFF) /k (DFF¡FFCA) was ϳ7-, 6-, and 13-fold faster for MetspHMFO His , PsespHMFO, and PseniHMFO His , respectively], with PseniHMFO His being the least efficient in these conversions, as expected from the HMF and DFF steady-state parameters (Table 3). For FFCA oxidation, reactions were followed during 4 days under the same conditions. As expected, this reaction was seen to be the limiting step for FDCA production, with k (FFCA¡FDCA) values of 0.0078, 0.0052, and 0.0092 h Ϫ1 for MetspHMFO His , PsespHMFO, and PseniHMFO His , respectively, indicating a more efficient FFCA oxidation by PseniHMFO His , which would therefore be the best enzyme for application.
Finally, long-term conversion of HMF was followed during 4 days (Fig. 8) at 28°C and the optimal pH for each enzyme. With a substrate/enzyme molar ratio of 300, the highest FDCA production was achieved by PseniHMFO His , with nearly complete conversion into FDCA after 96 h of reaction, while for MetspHMFO His and PsespHMFO, the yields were 89% and 65%, respectively (Table 4). Interestingly, under these conditions, PseniHMFO His showed a significantly longer half-life (150 h) than the other enzymes (ϳ40 to 60 h) and a higher total turnover number (TTN) for FDCA production.

DISCUSSION
HMFO screening, heterologous expression, and purification. The HMFO from Methylovorus sp. (MetspHMFO) is the only enzyme described to date that is directly related to HMF oxidation. This connection was established due to its homology with the related HmfH protein that forms part of a gene cluster involved in HMF metabolism in C. basilensis (3,6). In the present study, new HMFOs from P. nitroreducens and a Pseudomonas sp. strain were described, following their production as recombinant  Although an earlier attempt to express the gene responsible for HMF oxidation by C. basilensis and a homologous gene from Paraburkholderia phytofirmans was made, only MetspHMFO had been successfully produced as a functional protein (6) to date. Here, the two sequences from Pseudomonas strains, with 63 to 64% identity with the MetspHMFO sequence, were heterologously expressed in E. coli as active, soluble enzymes, together with MetspHMFO. The three recombinant HMFOs appeared as monomeric proteins, in agreement with a previous report on the latter enzyme (12), and were purified with good yields, increasing to three the number of members of the HMFO family currently available. New information on MetspHMFO. Comparison of the results obtained here for the Methylovorus enzyme purified without and with a His tag (MetspHMFO and MetspHMFO His ) indicated that the tag does not perturb the incorporation of the cofactor, as shown by the UV-visible spectra. Moreover, deconvolution of the CD spectra indicated an increase in the unordered structure, consistent with 90% of the His tag being disordered (27), and a slight decrease in the ␣-helix content. Neither activity nor the main physicochemical parameters were affected by the presence of the His tag, but a positive effect on stability under exposure to H 2 O 2 was found for the tagged MetspHMFO enzyme. This would represent an advantage for enzymes generating H 2 O 2 , which, as reported for other oxidases of the same superfamily, can inhibit the enzyme reactions or the enzymes themselves (28,29).
The kinetic parameters for vanillyl alcohol oxidation by MetspHMFO were similar to those previously reported for the same enzyme (14). However, the reported k cat values for HMF and DFF were significantly higher (594 min Ϫ1 and 96 min Ϫ1 , respectively) than those obtained in the present study. Considering the steady-state kinetic parameters obtained here, the estimated times for HMF and DFF consumption would be 7 and 98 min, respectively, which is consistent with the results obtained in 1-h reactions (1.5 mM HMF and 2.5 M HMFO). In contrast, Dijkman et al. (14) still detected DFF after 1 h of reaction between 2 mM HMF or DFF and 1 M HMFO, while according to their k cat value, all DFF should be consumed in ϳ20 min. In any case, the k cat and K m values for oxidation of these furfural derivatives by the MetspHMFO produced here are of the same order as those previously reported for other oxidases acting on these substrates, such as AAO (19) and GLOX (21). Regarding FFCA, the catalytic efficiency, estimated as k obs /[FFCA], was of the same order as that previously reported for MetspHMFO (12), although the strong inhibition by FFCA (above 3 mM) has not been described before.
Properties of two new HMFOs. The new enzymes from Pseudomonas strains were able to directly produce FDCA from HMF, with slight differences in their optimal reaction conditions, such as the higher activity and stability of PseniHMFO under slightly alkaline conditions. Although the new HMFOs do not show more activity on HMF and DFF than MetspHMFO, it is important to mention that they show higher efficiencies for FFCA oxidation, the rate-limiting step in FDCA production by HMFO (14) and other oxidases (16).
Strikingly, in terms of efficiency, PsespHMFO showed values for FFCA oxidation similar to those observed for DFF oxidation. The cooperative effect for DFF oxidation by this enzyme can be attributed to slow conformational changes that accompany substrate binding or product release. Similar cooperative effects have been described for other monomeric enzymes with single ligand-binding sites (30,31).
In the case of PseniHMFO, the efficiency for FFCA oxidation was only half of that of DFF oxidation. The results described above contrast with the much lower efficiency observed for MetspHMFO (FFCA oxidation 10-fold lower than DFF oxidation). The comparatively higher speed in rate-limiting FFCA oxidation by PseniHMFO results in higher final yields of FDCA in HMF bioconversions under optimized conditions. Moreover, its higher tolerance of the H 2 O 2 produced during the three oxidation steps results in a higher enzymatic stability during turnover, increasing its half-life. This stability provides an important advantage, since oxidases are often inhibited by large amounts of H 2 O 2 . Thus, our data reveal PseniHMFO as a robust and suitable biocatalyst for prolonged incubations, due to its longer half-life in wider ranges of pHs, temperatures, and H 2 O 2 concentrations.
Analysis of HMFO molecular models. Molecular modeling of PsespHMFO and PseniHMFO was undertaken at the Swiss-Model server (32), using the MetspHMFO crystal structure (PDB code 4UDP) (12) as the template. The predicted secondary structures were similar, as expected from their far-UV CD spectra.
However, inspection of their active sites revealed subtle differences in the environment (less than 6 Å) of the reactive N5 of FAD. Interestingly, in the MetspHMFO crystal structure, a water molecule is located at 3.7 Å from the above-named N atom and 4.6 Å from the catalytic His467 (Fig. 9a), at a position that is most probably occupied by the substrates during catalysis. Concerning neighbor residues, the pair Met103-Val104 in MetspHMFO (Fig. 9a) is replaced by Met103-Met104 in PsespHMFO (Fig. 9b) and by Val104-Leu105 in PseniHMFO (Fig. 9c). These two positions follow an active-site asparagine that is totally conserved in the GMC superfamily (1) as part of the conserved PS00623 sequence. Directed mutagenesis of MetspHMFO revealed how an M103A substitution (a changed of Met to Ala at position 103) significantly modified the enzyme turnover (for vanillyl alcohol) due to substrate pocket modification (12). A free space, potentially affecting the position of HMF and its partially oxidized derivatives (DFF and FFCA) with respect to the flavin cofactor, could also exist in PseniHMFO with a valine at this position.
The above-described differences do not include the presence of residues that could promote carbonyl activation in FFCA (33), as found in classical aldehyde-oxidizing oxidoreductases. However, changes in the flavin environment of PsespHMFO and PseniHMFO could modify the interaction of the flavin with the protein. Such interactions can modulate the redox potential of the oxidized isoalloxazine ring, promoting the enzyme's activity on less reactive substrates (34).
Pointing to the active-site cavity, Val367 is found in MetspHMFO. This residue is conserved in PsespHMFO, while Leu367 replaces the valine in PseniHMFO (Fig. 3). Interestingly, it has been reported that mutating this valine to arginine in MetspHMFO results in higher activity on FFCA (12), as found for PseniHMFO. However, the effect in the MetspHMFO variant is due to the introduction of a basic (arginine) residue, contributing to adequate positioning of the above-mentioned acidic substrate, while the natural change in PseniHMFO would only slightly reduce the pocket size.
On the other hand, the electrostatic surface potentials of the three HMFOs showed differences in charge distribution around the entrance of the flavin-access channel ( Fig.  9d to f), which could also modulate the access of substrates. Namely, the electronegative region partially covering the channel entrance in MetspHMFO could impede the access of acidic FFCA to the active site. However, the less electronegative entrances in PsespHMFO and especially in PseniHMFO would facilitate the entrance of this substrate, resulting in the observed higher activity of the new enzymes in catalyzing the ratelimiting step in HMF conversion to FDCA. The solvent access surfaces also showed that, although the O 2 substrate would more easily diffuse to attain the flavin ring of the buried cofactor in HMFOs, the entrance of the furfuryl-reducing substrates through the narrow access channel would most probably involve some side chain rearrangement at the residues forming the channel, as described in a related AAO (35,36).
Concluding remarks. Two enzymes from two Pseudomonas species have been added to the repertoire of HMFOs available, where only one member was purified and characterized to date. The enzyme from P. nitroreducens is a promising candidate for HMF oxidation due to its higher catalytic efficiency for FFCA oxidation, the bottleneck for FDCA production, compared to that of the previously described MetspHMFO. More importantly, in the context of industrial production of FDCA, additional advantages of the P. nitroreducens enzyme are its stability under exposure to H 2 O 2 and its robustness for long-term incubations, as shown by its high t 1/2 values under wider ranges of pH and temperature conditions. This feature would facilitate the enzyme's reuse and applicability in continuous operation for the production of FDCA as a building block for the production of bioplastics.

MATERIALS AND METHODS
Chemicals. HMF was kindly provided by AVA Biochem. DFF, FDCA, vanillyl alcohol, and vanillin were purchased from Sigma-Aldrich (St. Louis, MO, USA). FFCA was purchased from TCI America (Portland, OR, USA). AmplexRed and horseradish peroxidase were obtained from Invitrogen (Waltham, MA, USA).
Sequence search, phylogenetic analysis, and structure modeling. The screening of HMFO-like sequences was performed by BLAST of the amino acid sequence of MetspHMFO against the nonredundant protein sequences in GenBank. Multiple alignments of the enzymes found with Clustal Omega (37) were used to identify the conserved motifs (ADP binding domain and Prosite PS00623 and PS00624 sequences) in GMC proteins. A maximum-likelihood phylogenetic tree was constructed by MEGA X with 100-iteration bootstrapping (38) using the Whelan and Goldman (39) model of evolution using gammadistributed rate variation with empirical amino acid frequencies and invariant sites (WAGϩFϩIϩG). Finally, structural homology models of the protein sequences were generated using the Swiss-Model server (https://swissmodel.expasy.org) (32), and PyMOL version 2.033 (40) was used to examine the molecular structures obtained and generate solvent-accessible electrostatic surfaces.
Heterologous expression of HMFOs in E. coli. The codifying DNA sequences of the HMFOs from Methylovorus sp. M668, P. nitroreducens, Pseudomonas sp. 11/12A, Xanthobacter sp. 126, and Bradyrhizobium arachidis (NCBI accession numbers WP_013440946, WP_024766380, WP_047529632, WP_024277017, and WP_092217059, respectively) were manually optimized for E. coli expression and synthesized by ATG:biosynthetics. The sequences were subcloned from the pGH vector into the pET23b(ϩ) or pET28a(ϩ) expression vector. C41(DE3) E. coli cells were used for expression. E. coli cells containing the expression vectors were grown overnight in LB broth supplemented with the corresponding antibiotic at 37°C under continuous shaking. These precultures were used to inoculate 1-liter amounts of LB (supplemented with the antibiotic) and the cultures grown for 3 to 4 h at 37°C under continuous shaking until reaching an optical density at 500 nm of 0.9. Cultures were induced with 0.1 mM isopropyl-␤-D-thiogalactopyranoside, grown for 72 to 96 h at 16°C, and then harvested by centrifugation. Protein expression was monitored by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE).
Purification of recombinant HMFOs. The bacterial pellets were resuspended in the corresponding lysis buffer (Table S3 in the supplemental material), and the mixtures were incubated with 0.1 mg/ml of DNase I and 2 mg/ml lysozyme for 45 min. The solutions were sonicated and centrifuged to eliminate insoluble debris.
MetspHMFO and PsespHMFO (untagged proteins) were purified in a ResourceQ 6-ml column (GE Healthcare) with a linear NaCl gradient (60 to 160 mM in 4 column volumes [CV]) in 50 mM Tris-HCl, pH 8.0, followed by a MonoQ 5/50 GL column in which the protein fractions were eluted with a NaCl linear gradient (0 to 70 mM in 10 CV) in 50 mM Tris-HCl, pH 8.0 (Table S3).
His-tagged fusion proteins (MetspHMFO His and PseniHMFO His ) were purified by affinity chromatography in a HiTrap IMAC FF column (GE Healthcare) with a linear imidazole gradient (20 m to 300 mM in 6 CV) in 500 mM NaCl, 20 mM Tris-HCl, pH 7.5, followed by anionic exchange chromatography with a ResourceQ column (GE Healthcare) with an NaCl linear gradient (0 to 500 mM in 5 CV) in 50 mM Tris-HCl, pH 7.5 (Table S3).
The purified HMFOs were dialyzed against 50 mM Tris-HCl, pH 7.0, and stored at Ϫ80°C, being stable for several months.
Spectroscopic measurements. The UV-visible spectra of purified HMFOs were recorded between 250 and 700 nm in a Cary4000 spectrophotometer. Their extinction coefficients were calculated by heat denaturation (41) and estimation of the free FAD released ( band I ϭ 11,300 mM Ϫ1 cm Ϫ1 ) (42).
MW and pI determination. MW was analyzed by MALDI-TOF mass spectrometry at the Proteomics and Genomics Facility of CIB (CSIC), a member of the ProteoRed-ISCIII network. The experiments were performed in an Autoflex III MALDI-tandem-TOF instrument (Bruker Daltonics, Bremen, Germany) with a smart beam laser. Samples dissolved in 50 mM Tris-HCl, pH 7.0, were mixed with the matrix solution (3.5-dimethoxy-4-hydroxycinnamic acid dissolved in a mixture of acetonitrile-01% trifluoroacetic acid [1:2, vol/vol]). External calibration was performed using bovine albumin (Sigma), covering the range from 20,000 to 70,000 Da.
The pI was determined by 2-dimensional (2-D) electrophoresis. The first dimension was run on immobilized pH gradient strips (pH 3 to 10, linear, 7 cm) (Bio-Rad), and the second dimension was run on 12% SDS-PAGE gels. Protein bands were stained with a colloidal blue staining kit (Invitrogen).
pH and H 2 O 2 stability. The pH and H 2 O 2 stabilities were estimated by incubating the enzymes (5 to 40 M) in 100 mM Britton and Robinson (B&R) buffer at different pHs (pH 4.0 to 9.0) or in the presence of different H 2 O 2 concentrations (0.5 to 7.8 mM) in 50 mM Tris-HCl, pH 7.0. Residual activities were estimated by following the oxidation of saturating concentrations of vanillyl alcohol (3 mM) in 50 mM Tris-HCl, pH 7.0, at 25°C, immediately after mixing and after 24 h, 48 h, and 72 h of incubation at 25°C. For each enzyme, the highest activity after mixing was taken as 100% activity, and the percentages of residual activity at the different times and conditions were calculated according to this maximal value.
Thermal stability. The melting temperatures of the enzymes were analyzed by the ThermoFAD method (26). The increase in fluorescence due to FAD released as a consequence of enzyme unfolding was monitored with a real-time PCR thermocycler iQ5 (Bio-Rad). Denaturation curves were fitted to equation 1, describing a two-step thermal denaturation equilibrium: where ⌬G ϭ ⌬H͓1 Ϫ ͑T⁄T m ͔͒ Ϫ ⌬Cp͓͑T m Ϫ T͒ ϩ T ln͑T⁄T m ͔͒, F N and F D are the fluorescence signals of the native and denatured states, m N and m D are the slopes that describe their dependences with temperature, T and T m are the temperature and the melting temperature, respectively, R is the universal gas constant, and ΔG, ΔH, and ΔC p are the free energy, enthalpy, and specific heat of denaturation, respectively.
Optimal pH and temperature. The optimal temperatures and pHs for vanillyl alcohol oxidation by HMFOs were determined by measuring the oxidation of saturating concentrations of the substrate (3 mM) in 50 mM Tris-HCl buffer, pH 7.0, at different temperatures between 4°C and 70°C or in 100 mM B&R buffer in the range of pHs from 2.0 to 12.0. It was observed that the UV-visible spectra of vanillyl alcohol and vanillin changed with pH ( Fig. S5a and b) (46). Thus, in the case of the optimal pH, the activity was measured using the extinction coefficient for vanillyl alcohol oxidation to vanillin calculated for each pH (Fig. S5c).
Steady-state kinetics. Enzymatic activity was estimated by following the production of vanillin upon oxidation of vanillyl alcohol at 25°C in 50 mM Tris-HCl at the indicated pH in each case, using the corresponding extinction coefficient for vanillyl alcohol oxidation to vanillin (Fig. S5c).
Steady-state kinetic parameters for HMF and DFF oxidation by HMFOs were calculated by monitoring the production of H 2 O 2 during oxidation of the different substrates, using an HRP-coupled assay with AmplexRed as the final substrate in 50 mM Tris-HCl, pH 7.0, as previously described (16).