Rational engineering of a flavoprotein oxidase for improved direct oxidation of alcohols to carboxylic acids

Article abstract of DOI:10.3390/molecules22122205

The oxidation of alcohols to the corresponding carbonyl or carboxyl compounds represents a convenient strategy for the selective introduction of electrophilic carbon centres into carbohydrate-based starting materials. The O₂-dependent oxidation of prim-alcohols by flavin-containing alcohol oxidases often yields mixtures of aldehyde and carboxylic acid, which is due to "over-oxidation" of the aldehyde hydrate intermediate. In order to directly convert alcohols into carboxylic acids, rational engineering of 5-(hydroxymethyl)furfural oxidase was performed. In an attempt to improve the binding of the aldehyde hydrate in the active site to boost aldehyde-oxidase activity, two active-site residues were exchanged for hydrogen-bond-donating and -accepting amino acids. Enhanced over-oxidation was demonstrated and Michaelis-Menten kinetics were performed to corroborate these findings.

Full text of DOI:10.3390/molecules22122205

molecules
Article
Rational Engineering of a Flavoprotein Oxidase for
Improved Direct Oxidation of Alcohols to
Carboxylic Acids
Mathias Pickl 1, Christoph K. Winkler 2, Silvia M. Glueck 2, Marco W. Fraaije 3
ID
and Kurt Faber 1,
*
1
Department of Chemistry, University of Graz, Heinrichstrasse 28, A-8010 Graz, Austria;
Mathias.Pickl@edu.uni-graz.at
Austrian Centre of Industrial Biotechnology, ACIB GmbH c/o Department of Chemistry, University of Graz,
Heinrichstrasse 28, A-8010 Graz, Austria; winkler_christoph@hotmail.com (C.K.W.);
Si.Glueck@uni-graz.at (S.M.G.)
Molecular Enzymology Group, Groningen Biomolecular Sciences and Biotechnology Institute,
University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands; m.w.fraaije@rug.nl
Correspondence: kurt.faber@uni-graz.at; Tel.: +43-316-380-5332
2
3
*
Received: 21 November 2017; Accepted: 8 December 2017; Published: 12 December 2017
Abstract: The oxidation of alcohols to the corresponding carbonyl or carboxyl compounds
represents a convenient strategy for the selective introduction of electrophilic carbon centres
into carbohydrate-based starting materials. The O2-dependent oxidation of prim-alcohols by
flavin-containing alcohol oxidases often yields mixtures of aldehyde and carboxylic acid, which is due
to “over-oxidation” of the aldehyde hydrate intermediate. In order to directly convert alcohols into
carboxylic acids, rational engineering of 5-(hydroxymethyl)furfural oxidase was performed. In an
attempt to improve the binding of the aldehyde hydrate in the active site to boost aldehyde-oxidase
activity, two active-site residues were exchanged for hydrogen-bond-donating and -accepting amino
acids. Enhanced over-oxidation was demonstrated and Michaelis–Menten kinetics were performed
to corroborate these findings.
Keywords: biocatalysis; alcohol oxidation; aldehyde oxidation; flavoprotein oxidase; protein design
1. Introduction
Oxidases are prominent biocatalysts due to their ability to utilize molecular oxygen as oxidant [
In contrast to alcohol dehydrogenases, they do not suffer from an equilibrium problem, as the
utilization of O2 as electron acceptor leads to a practically irreversible reaction [ ]. Recently,
1,2].
1,3
5-(hydroxymethyl)furfural oxidase from Methylovorus sp. MP688 (HMFO, EC: 1.1.3.37), a well
expressing, stable flavoprotein, that is active on a variety of benzylic or allylic prim-alcohols was
described [
be obtained from hexoses on a large scale, and its oxidation to 2,5-furandicarboxylic acid furnishes a
bio-based monomer for PEF as a substitute for petroleum-derived PET [ ]. Furthermore, the oxidation
activity of HMFO on prim-thiols has been described [ ]. Crystal structures of HMFO containing
oxidized and reduced flavin are available (Figure 1, PDB: 4UDP and 4UDQ), which makes it a
perfect template for rational protein redesign [ ]. Analogous to other structurally related flavoprotein
4], which was termed for its ability to oxidize 5-(hydroxymethyl)furfural [5]. The latter can
6
7
8
oxidases, the reaction proceeds through proton abstraction from OH through a histidine residue
(His467), with concomitant hydride transfer to the N5 of the FAD cofactor. Re-oxidation of FADH by
O2 leads to the formation of hydrogen peroxide and regenerates FAD. In order to avoid detrimental
effects of the formed hydrogen peroxide and to regenerate molecular oxygen, catalase was added to
the standard reaction setup in this study.
Molecules 2017, 22, 2205; doi:10.3390/molecules22122205
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Figure 1. Wild-type HMFO active site (PDB 4UDP): FAD (yellow), residues surrounding flavin-N5
are shown in green (Met103, Asn511, His467), residues selected for site-directed mutagenesis are
highlighted in pink (Val465, Trp466), and benzaldehyde hydrate (gem-diol of 1b) is shown in blue.
When prim-alcohols are used as substrates, the oxidation reaction does not necessarily stop
at the aldehyde stage, but may proceed to the corresponding carboxylic acid. This takes place
via spontaneous (non-enzymatic) formation of the aldehyde (gem-diol) hydrate. In contrast to the
aldehyde, this hydrated species is accepted as a substrate by HMFO and furnishes the carboxylic acid
via “over-oxidation” (Scheme 1). Similar activities are known for several flavin-dependent alcohol
oxidases [
9–11]. In contrast, Cu-containing galactose oxidase is unable to oxidize aldehydes and/or
the corresponding hydrates [12
–
14]. For such enzymes, an additional aldehyde oxidase is required to
produce carboxylic acids from alcohols [15].
Scheme 1. Two-step oxidation of benzylic alcohols to carboxylic acids catalyzed by HMFO variants.
Although several flavin-dependent alcohol oxidases are capable of oxidizing an alcohol to the
corresponding carboxylic acid, they show astonishing differences: some alcohol oxidases readily
oxidize the aldehyde to the carboxylic acid [16,17], while others (e.g., aryl alcohol oxidase) render the
carboxylic acid only in minor amounts [10]. Furthermore, the equilibrium and rate of (non-enzymatic)
aldehyde hydration strongly depend on the electronic properties of the substrate and the solvent
system [1]. Overall, HMFO oxidizes aldehydes rather slowly. With the aim to develop an oxidase for the
direct oxidation of alcohols to the corresponding carboxylic acids via the aldehyde stage, we engineered
HMFO oxidase to enhance its catalytic performance for the second aldehyde oxidation step.

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2. Results and Discussion
Our strategy to improve the oxidation of the aldehyde hydrate via protein engineering focuses on
the introduction of hydrogen-bond-donating or -accepting amino acids that support stabilization of the
gem-diol in the active site, bearing in mind that an additional H-bond adds ~2 kcal
mol−1 of binding
·
energy [18]. Similar patterns are found in aldehyde dehydrogenases, where a polar amino acid active
site residue promotes aldehyde hydration and hence favors carboxylic acid formation [19]. Potential
active-site residues for mutation were identified by considering the orientation and distance of their
side chain to the substrate's hydroxy-groups. Five amino acids in the vicinity of FAD are highlighted
in Figure 1.
From those, two residues interacting with the (hydrated) aldehyde moiety of the substrate via
hydrogen bonding by positioning of the benzylic hydrogen atom in close proximity of N5 were selected
(for docking, see SI): Val465, positioned directly on top of N5 of FAD and Trp466, performing
stacking with the benzylic moiety of the isoalloxazine ring of FAD (Figure 1) [8,20].
π-π
To eliminate potential sources of errors arising from different protein expression levels
and background reactions only purified enzymes were tested [21 23]. In an initial experiment,
the performance of wild-type HMFO in the oxidation of benzylic alcohols was evaluated with
substrates bearing electron-withdrawing and -donating groups in the p-position (1a 5a) after 24 h
(Figure 2). All substrates were readily oxidized to the corresponding aldehydes 1b 5b. To our
delight, the highly activated p-nitrobenzyl alcohol 5a was over-oxidized to carboxylic acid 5c with 29%
conversion, while substrates 1a 4a showed only traces of carboxylic acid formation (<1%). This can
–
–
–
–
be explained by the strong electron-withdrawing p-nitro-group of 5a, which leads to ~15% hydrated
gem-diol in equilibrium [24]. According to the literature, the ratio between the aldehyde and its
(gem-diol) hydrate remains constant during a similar enzymatic redox reaction [24,25], which is
experimentally supported by the fact that identical endpoint results were obtained for reactions using
pre-incubated aldehyde (see SI). This means that the rate of aldehyde hydration is not rate limiting,
but the position of the equilibrium.
Figure 2. Oxidation of benzylic alcohols 1a–5a (25 mM) by HMFO (wild type, 0.7 µM) in sodium
phosphate buffer (100 mM, pH 7.0) at 30 ◦C with atmospheric oxygen in the presence of catalase
after 24 h.
In order to enhance binding of the aldehyde hydrate, Trp466 and Val465 in wild-type HFMO
(Figure 1, purple) were systematically exchanged to amino acids that may serve as H-bond donors or
-acceptors, i.e., tyrosine, glutamine, asparagine, histidine, threonine, serine, lysine, arginine, aspartate,
and glutamate. In order to differentiate H-bonding from steric effects, also apolar amino acids alanine
and phenylalanine were included. Since wild-type HMFO already showed measurable formation

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of the carboxylic acid after 2 h (5a, 4%), this time point was chosen as a reference point for the
comparison of variants, because product inhibition and problems of enzyme instability are negligable.
With non-activated substrates 1a–4a, all variants gave only traces of over-oxidized product. However,
using 5a as substrate, encouraging results were obtained (Figure 3). Although activities dropped
in general, several mutants bearing polar residues in place of Val465 and Trp466 showed enhanced
chemoselectivity for carboxylic acid formation relative to wild type HMFO (Table 1).
Figure 3. Oxidation of 4-nitrobenzyl alcohol (5a) (25 mM) to 4-nitrobenzaldehyde (5b) and
4-nitrobenzoic acid (5c) by HMFO variants (0.7
30 C with atmospheric oxygen in presence of catalase after 2 h (HPLC).
µ
M) in sodium phosphate buffer (100 mM, pH 7.0) at
◦
Table 1. Enhanced carboxylic acid formation (5c) from 5a with HMFO variants.
Variant
Ratio (%) a
Wild type
Trp466Gln
Trp466His
Trp466Ser
Trp466Arg
Val465Ser
Val465Thr
Val465Asp
6
5
18
18
17
14
7
16
Val465Thr/Trp466His 37
a
Ratio of carboxylic acid (5c) to aldehyde formation (5b).
Most variants showed a similar chemo-selectivity for carboxylic acid/aldehyde formation of 1:12
to 1:15 as the wild type. Although Trp466Ser, Trp466Arg, and Val465Asp variants showed enhanced
over-oxidation, their overall activity was disappointing (<5%). In contrast, Trp466His and Val465Ser
mutants showed 2–3-fold enhanced acid formation by maintaining reasonable overall activities (~25%).
In an attempt to combine good activity with enhanced chemoselectivity, a double variant was
constructed by combining Val465Thr (most active) with Trp466His (highest chemoselectivity) (Figure 3).
By this way, the ratio of aldehyde-formation/overoxidation could be further pushed to a 5-fold
improvement of the chemoselectivity at the expense of some loss of activity.
The potency of the exchange of Trp466 to His may be explained by two effects: The introduction
of a hydrogen bond interaction with the substrate while maintaining π-π interaction with FAD for
re-oxidation (similar to Trp at this position in the wild type) [26]. The positive effect of Ser in position
465 may be attributed to maintaining Trp466 in place for efficient re-oxidation [20] combined with
favorable H-bonding of the aldehyde hydrate.

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For further characterization of the activities of enzyme variants in the individual oxidation steps,
Michaelis–Menten kinetics were determined. Hence, alcohol (5a) and aldehyde (5b) were tested with
the wild-type enzyme and the most active variants Val465Thr and Trp466His (Table 2). The catalytic
−1
efficiency of the wild type enzyme was found to be high for 5a (kcat/KM 28 s
·
mM−1) and modest for
5b (~1 s−1
·
mM−1), accounting for the high chemoselectivity (28-fold) between both oxidation steps.
Since the KM for alcohol and aldehyde (hydrate) remained about the same (0.26 versus 0.31 mM),
chemoselectivity was (almost entirely) caused by differences in kcat (7.7 vs. 0.3 s−1). The doubled
catalytic efficiency of Val465Thr for 5a was caused by a 10-fold lower KM, which over-compensates
a ~4-fold reduction in kcat compared to the wild type. The Trp466His variant showed a three orders
of magnitude lower catalytic efficiency, which is due to the loss in kcat, because the KM remained
almost unaffected. Surprisingly, whereas Trp466His exhibited a measurable catalytic efficiency for 5b
,
no activity could be found with Val465Thr, even at higher enzyme concentrations. Since for Val465Thr
an overall performance similar to the wild-type HMFO was expected, it is remarkable that no activity
was detected on substrate 5b. A possible explanation is that there is an extremely low KM and kcat
,
which would be in line with the observed lowered KM for the alcohol substrate with Val465Thr. It is
striking that the efficiency of Trp466His for 5a and 5b is within a close range, between 50 s−1 M−1
·
and 20 s−1
of magnitude.
·
M
−1, respectively, but for the wild-type enzyme these values differed by three orders
Table 2. Steady state kinetics of wild-type HMFO and selected variants in the oxidation of p-nitrobenzyl
alcohol (5a) and p-nitrobenzaldehyde (5b).
kcat (app)/KM (app)
−1
)
KM (app) (mM)
Substrate
HMFO Variant
kcat (app) (s
−1
−1
(s ·M
)
5a
Wild type
Trp466His
Val465Thr
7.4
0.02
1.7
0.26
0.40
0.03
28,000
50
57,000
4-nitrobenzyl alcohol
5b
Wild type
Trp466His
Val465Thr
0.3
0.003
<0.0001
0.31
0.16
n.d.
970
20
-
4-nitrobenzaldehyde
Kinetic parameters of wild-type HMFO and variants were determined in sodium phosphate
buffer (100 mM, pH 7.0) at 30 ◦C by monitoring H2O2 formation in a coupled assay [27].
3. Materials and Methods
3.1. Bacterial Cultivation and Enzyme Production
Protein expression and purification were performed after an adapted protocol from Dijkman
and Fraaije [
the SUMO-HMFO encoding plasmid (Champion
CA, USA) was diluted 1:100 in 200 mL of Terrific Broth containing 50
was grown at 37 C until an optical density at 600 nm (OD600) of 0.8–1.0 was reached. Cells were
induced with isopropyl-
Cells were harvested by centrifugation at 3730
Hettich, Tuttlingen, Germany) and resuspended in Tris-HCl (35 mL, 100 mM, pH 8.0) supplemented
with glycerol (10% v/v), 150 mM NaCl (150 mM) and FAD (10 M). Cells were disrupted with a
4
]. For HMFO expression, an overnight culture of E. coli BL21(DE3) cells containing
pET SUMO, ThermoFisher Scientific, Carlsbad,
™
µg
mL−1 kanamycin and
·
◦
β
-D-thiogalactopyranoside (IPTG, 1.0 mM) and grown overnight at 20 ◦C.
g for 15 min (Hettich® Rotina 420R centrifuge, 4 ◦C,
×
µ
Branson Digital Sonifier 250 (30% amplitude, 1 s pulse, 4 s pause for 2 min). The lysate was cleared by
centrifugation (20,000× g for 15 min).
His-Tagged HMFO was purified by Ni-affinity chromatography (5 mL HiTrap FF column, GE
Healthcare, Little Chalfont, UK) applying a 5–500 mM gradient of imidazole (binding buffer: Tris-HCl
50 mM, pH 8.0 with 150 mM NaCl containing 5 mM imidazole; elution buffer: Tris-HCl 50 mM,

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pH 8.0 with 150 mM NaCl containing 500 mM imidazole). Fractions containing HMFO (analysed by
SDS-PAGE) were pooled, concentrated by ultrafiltration (20 mL, 50 kDa cut-off, Vivaspin, Göttingen,
Germany), and desalted (Sephadex
™ G-25 M, GE Healthcare, Little Chalfont, UK). The sample was
◦
lyophilized and stored at
−
20 C. For activity tests, the lyophilized enzyme preparation was dissolved
in sodium phosphate buffer (pH 7.0, 100 mM, 20 µL/mg) without cleaving off the SUMO-tag.
3.2. Site-Directed Mutagenesis
Site-directed mutagenesis of the wild-type HMFO gene was performed using two-step
whole-plasmid PCR. For the creation of Val465Thr-Trp466His, the HMFO-Trp466His plasmid was used
as template. The primers were ordered at IDT (Leuven, Belgium) and are listed in Table S1.
After three cycles of linear PCR, the mixture containing the forward primer and the mixture with
the reverse primer were combined for additional 15 cycles. Template DNA was cleaved with DpnI
(New England Bio-Labs, Ipswich, MA, USA). The plasmid was purified with a PCR purification kit
(Qiagen, Hilden, Germany) and transformed into E. coli TOP10 cells. The introduction of the mutations
was confirmed by sequencing.
3.3. Biotransformation
Lyophilized purified HMFO variants were dissolved in phosphate buffer (100 mM, pH 7.0),
after determining the protein concentration by Bradford analysis. HMFO (0.7
µM) variants and
catalases from M. lysodeikticus (10 L, 1700 U) were added to phosphate buffer (1 mL, pH 7.0, 100 mM)
µ
supplemented with FAD (20
in a horizontal position. The reaction mixture was shaken at 30 C and 120 rpm for 24 h at atmospheric
µ
M). The substrate (25 mM) was added, and vials were placed into a shaker
◦
oxygen concentration. The mixture was then vortexed, MeCN (500 µL) was added, the mixture was
vortexed again and centrifuged, and the supernatant was directly measured by HPLC-UV.
3.4. Aldehyde Hydration
A solution of 1b
24 h at room temperature. After incubation, wild-type HMFO and catalase (10
added. The vials were placed in a shaker in a horizontal position and shaken at 30 ◦C and 120 rpm
for 24 h at atmospheric oxygen concentration. The mixture was then vortexed, MeCN (500 L) was
–
5b (25 mM) in phosphate buffer (1 mL, 100 mM, pH 7.0) was incubated for
µL, 1700 U) were
µ
added, the mixture was vortexed again and centrifuged, and the supernatant was directly measured
by HPLC-UV. As a control, the reaction was performed without preincubating the aldehyde (see SI).
3.5. Kinetics
The kcat and KM values of the HMFO variants were determined by measuring H2O2 production
during the reaction. In a coupled H2O2 detection assay, horseradish peroxidase (HRP 40 U/mL)
oxidized 4-aminoantipyrine (0.1 mM) and 3,5-dichloro-2-hydroxybenzenesulfonic acid (1 mM) to form
a pink product that was monitored at 515 nm (ε515 = 26 mM−1
·
cm−1) or with Amplex® Red (50
M) was assayed at
µM,
monitored at 563 nm, ε563 = 52 mM−1 cm−1). The activity of HMFO (0.1 to 10
·
µ
different substrate concentrations (0.05 to 10 mM), depending on the substrate used. All reactions were
performed at 30 ◦C in phosphate buffer (100 mM, pH 7.0).
4. Conclusions
HMFO is a potent biocatalyst for the O2-dependent oxidation of substituted benzylic alcohols
to the corresponding aldehydes and/or carboxylic acids. The relative ratio of both oxidation steps
strongly depends on the electronic properties of the p-substituent on the aromatic ring, which is due to
its effect on the hydration of the aldehyde forming the gem-diol substrate for the second oxidation step.
As a result, 4-nitrobenzyl alcohol was efficiently converted to the corresponding carboxylic acid. With
the introduction of hydrogen bonding amino acid residues within the active site, we expected to boost

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the second oxidation step by improving the binding of the gem-diol substrate. Indeed, the relative
activity of the two oxidation steps was improved up to ~5-fold in favor of over-oxidation to the
carboxylic acid for substrate 5a, albeit at the cost of overall activity. However, for substrates 1a–4a,
lacking an electron-withdrawing (activating) group, no such improvement was achieved due to the
unfavorable position of the aldehyde-hydration equilibrium. If carboxylic acids are desired from
alcohols, engineered HMFO variants showing enhanced over-oxidation activities are advantageous,
because they do not form toxic levels of aldehyde intermediates through direct formation of carboxylic
acids, which are better tolerated by whole-cell biocatalysts [28].
Supplementary Materials: Supplementary materials are available online.
Acknowledgments: Funding from the Austrian Science Fund (FWF) within the DK Molecular Enzymology
(project W901) and the Austrian BMWFW, BMVIT, SFG, Standortagentur Tirol, Government of Lower Austria,
and ZIT through the Austrian FFG-COMET-Funding Program is gratefully acknowledged.
Author Contributions: M. Pickl and C. K. Winkler conceived and designed the experiments; M. Pickl performed
the experiments, and M. Pickl, S. M. Glueck, M. W. Fraaije and K. Faber analyzed the data and wrote the paper.
Conflicts of Interest: The authors declare no conflict of interest.
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,
Sample Availability: Samples of the compounds 1a–5a and 1c–5c are available from the authors.
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