ArticlePDF Available

Creating peroxidase-oxidase fusion enzymes as toolbox for cascade reactions



In this work we prepared a set of bifunctional oxidase-peroxidases by fusing four distinct oxidases to a peroxidase. While such fusion enzymes have not been observed in nature, they could be expressed and purified with good yields. Characterization revealed that the artificial enzymes retained the capability to bind the two required cofactors and were catalytically active as oxidase and peroxidase. The peroxidase fusions of alditol oxidase and chitooligosaccharide oxidase could be used for selective detection of xylitol and cellobiose with a detection limit in the low µM range. The peroxidase fusions of eugenol oxidase and 5-hydroxymethylfurfural oxidase could be used for dioxygen-driven one-pot two-step cascade reactions to convert vanillyl alcohol into divanillin and eugenol into lignin oligomers, respectively. The designed oxidase-peroxidase fusions represent attractive biocatalysts that allow efficient biocatalytic cascade oxidations that only require molecular oxygen as oxidant.
Creating Oxidase–Peroxidase Fusion Enzymes as aToolbox
for Cascade Reactions
Dana I. Colpa,[a] NikolaLonc
ˇar,[b] Mareike Schmidt,[a] and Marco W. Fraaije*[a]
Aset of bifunctional oxidase–peroxidases has been prepared
by fusing four distinct oxidases to aperoxidase. Although such
fusion enzymeshave not been observed in nature,they could
be expressed andpurified in good yields. Characterization re-
vealed that the artificial enzymes retained the capability to
bind the two required cofactors and were catalytically activea
oxidaseand peroxidase. Peroxidase fusions of alditol oxidase
and chitooligosaccharide oxidasecould be used for the selec-
tive detection of xylitol and cellobiose with adetection limit in
the low-micromolar range. The peroxidase fusions of eugenol
oxidaseand 5-hydroxymethylfurfural oxidase could be used for
dioxygen-driven, one-pot,two-step cascade reactions to con-
vert vanillylalcohol into divanillin and eugenol into lignin olig-
omers. The designed oxidase–peroxidase fusions representat-
tractive biocatalysts that allow efficient biocatalytic cascade ox-
idations that only require molecular oxygen as an oxidant.
In nature,most enzymes take part in metabolic pathwaysin
which each formed product is asubstrate for the next enzy-
matic reaction. To optimize the efficiency of such intricate bio-
catalytic cascades,the enzymes are often brought together to
form enzyme complexes, for example, the pyruvate dehydro-
genasecomplex,the microbial type Ifatty acid synthasecom-
plex, andthe cellulosome.[1–3] The cellulosome is found in
anaerobic microorganisms and consists of ascaffolding pro-
tein, which brings together the required hydrolytic enzymes to
degradecellulosic biomass.[3] In some cases, this has even led
to the fusion of two or more enzymes to create abi-/multi-
functional protein,[4] for example, pyrroline-5-carboxylate syn-
thase, which features both glutamate kinase and g-glutamyl
phosphate reductase activities, or the pentafunctional AROM
complexfrom Aspergillus nidulans,which is involved in aromat-
ic amino acid biosynthesis.[5, 6]
Inspired by the latter observation, variousartificial enzyme
fusions have been createdinrecent years to engineerefficient
multifunctional biocatalysts. The first artificial bifunctional
fusion enzyme, ahistidinol dehydrogenase/aminotransferase,
was published in 1970.[7] Several fusion enzymes have been
made since.[4,8, 9] For example, afusion between afatty acid de-
carboxylase cytochrome P450 (OleTJE) and alditol oxidase
(AldO)was made to fuel the reactions of OleTJE with hydrogen
peroxide produced by the oxidase.[10] To enable efficient cofac-
tor regeneration, we have shown that various nicotinamide ad-
enine dinucleotide (phosphate) (NAD(P)H)-dependent monoox-
ygenases can be produced fused to phosphite dehydrogenas-
es, which efficiently regenerate NAD(P)H.[11,12]
Fusionenzymes have several advantagesover separate en-
zymes.They are cheaper and less labor intensive concerning
their production because only one enzyme needs to be ex-
pressedand purified. Another advantage is the close proximity
of the catalytic sites, which enables substrate channel-
ing.[4,9, 13, 14] Substrate channeling circumvents diffusion of the
intermediate product in the solution,and hence, increases the
combined reactionrate.
We were particularly inspired by the interplay between oxi-
dases and peroxidases also found in nature. Oxidases and per-
oxidasesare often coexpressed. Oxidases produce hydrogen
peroxide, whichagain is asubstrate for peroxidases. Well-
knownexamples of such interplay between oxidases and per-
oxidasesare found in fungi.[15,16] Many fungi secrete specialized
peroxidases (e.g.,lignin peroxidase and manganese perox-
idase) that aid in biomass degradation.[15] Except for secreting
these heme-containing enzymes,these fungi also secretevari-
ous oxidases (e.g.,pyranose oxidase and aryl alcohol oxidase)
to serve as hydrogen peroxideproducing enzymes to fuel the
peroxidases.[15, 16] The consecutivereactions of oxidasesand
peroxidases are also applied in enzyme activity screening ap-
proachesand biosensors. Numerous assays and biosensors are
based on the combination of an oxidaseand aperoxidase, for
instance, for the detection of glucoseoruric acid levels in
blood serum.[17–19] The activities of various oxidases were stud-
ied in coupled assays,inwhich,typically,horseradish perox-
idase (HRP) is employed.[20–23] HRP,however,isstill extracted
from horseradish because of difficulties in the heterologous ex-
pression of this plant peroxidase.[24] Therefore, for this study,
we selectedarecently discovered bacterial peroxidase, SviDyP,
which waseasily produced by Escherichia coli.[25, 26]
Fusion enzymes betweenoxidases and peroxidases have not
been made previously,althoughthey form catalytically logical
combinationsbecause the oxidase-formed hydrogen peroxide
will drive the fused peroxidase (Figure 1).
[a] D. I. Colpa,M.Schmidt, Prof. Dr.M.W.Fraaije
Molecular EnzymologyGroup,University of Groningen
Nijenborgh 4, 9747AG Groningen (The Netherlands)
[b] Dr.N.Lonc
Groningen Enzyme and CofactorCollection (GECCO)
University of Groningen, Nijenborgh 4
9747AG Groningen (The Netherlands)
Supporting information and the ORCID identification numbers for the
authors of this article can be found under https ://
T2017 The Authors. Published by Wiley-VCH Verlag GmbH&Co. KGaA.
This is an open access article under the terms of the Creative Commons At-
tributionNon-Commercial NoDerivs License, which permitsuse and distri-
bution in any medium, provided the original work is properly cited, the use
is non-commercial and no modifications or adaptations are made.
ChemBioChem 2017,18,2226 –2230 T2017 The Authors. PublishedbyWiley-VCH Verlag GmbH &Co. KGaA, Weinheim2226
Herein, we fused abacterial peroxidase (SviDyP from Saccha-
romonospora viridis DSM 43017, EC to four different
bacterialoxidases (EC 1.1.3.x).[25, 26] SviDyP belongs to the family
of DyP-type peroxidases, which are known fortheir activity on
dyes and phenolic compounds.[27–30] The peroxidase is easily
expressed in abacterial hostand is also avery robust enzyme.
SviDyP was fused to two flavin adenine dinucleotide (FAD)-
containing oxidases that were active towards sugars: alditol
oxidase(HotAldO) from Acidothermus cellulolyticus 11Band chi-
tooligosaccharide oxidase (ChitO) from Fusarium graminea-
rum.[20, 23] In this study,aChitO triple mutant, Q268R/G270E/
S410R (ChitO*), was used because of its increased catalytic effi-
ciency towards glucose, lactose, cellobiose, and maltose. In ad-
dition to these oxidases, SviDyP was fused to two other flavo-
protein oxidases that featured apartially overlapping sub-
strate/product scope to SviDyP:eugenol oxidase (EugO) from
Rhodococcus sp. strain RHA1 and 5-hydroxymethylfurfuralox-
idase (HMFO) from Methylovorus sp. strain MP688.[21, 22] This
overlap in substrate/product scope, with both fusion partners
active on phenolic compounds, would allow one-pot cascade
reactions. Thus, we were able to produce four novel bifunc-
tional fusion biocatalysts that could either serve arole in bio-
sensing or act as acatalyst for one-pot,two-step cascade reac-
The four DyP-type peroxidase/oxidase fusion enzymes
(which we termed P-oxidases) were made by cloning the
genes of the individual oxidases ChitO*,EugO, HMFO, and
HotAldO Cterminally to the gene encoding for His-tagged
SviDyP.The resulting fusion enzymes were overexpressed and
subsequently purified by affinity chromatography to yield 26–
60 mg of enzyme per liter of culture broth medium. The fusion
enzymes displayed an intense red–brown color that was indi-
cative of binding of the heme and flavin cofactors. Analysis by
UV/Vis absorbance spectroscopy revealed absorbance maxima
at l=280 (protein) and 406 nm (heme) for all enzymes. The
Reinheitszahl (Rzvalue) of the fusion enzymes varied between
0.61 and 0.97, andsuggested effective incorporation of the
heme cofactor.The typicalabsorbance maxima of FAD, l
&350–385 and 440–460nm,[20–22, 31] couldnot be observed due
to the high absorbance of the heme cofactor.Toconfirmbind-
ing of the FADcofactor,the purified ChitO*,EugO, and Hot-
AldO fusion enzymes were analyzedfor in-gel fluorescence
after SDS-PAGE. This revealed that all three fusion enzyme con-
tained acovalently bound flavin cofactor. Such analysis was
not feasible forthe HMFO fusion enzyme because this flavo-
protein oxidase contained adissociable FAD. Nevertheless, ac-
tivity measurements (see below)confirmed that this fusion
enzymewas also functional as an oxidase, thus confirming the
presence of the flavin cofactor.
To verify that the prepared fusion enzymes were fully func-
tional,the activities of both fusion partnerswere measured
(Table 1). The observed peroxidase activities for all fusion en-
zymeswere in good agreement with the kcat. values deter-
mined for the isolated peroxidase. Accordingly,itcan be con-
cluded that the activity of the peroxidase was unaffected by
fusing it to the oxidases. Also, the oxidasesdisplayed activity
when fused to SviDyP,although the activities were somewhat
lower than the activitiesofthe non-fused enzymes.Oxidase ac-
tivities of 15–43 %were observed for the fused oxidases
ChitO*,EugO, HMFO, and HotAldO.This could be partly ex-
plained because the activity was measured at afixed substrate
concentration (kobs), whichwould yield lower rates when com-
pared with kcat. values taken from the literature.Another ex-
planation of the lower observed rates may lie in incomplete
flavin cofactor incorporation.Yet, prolonged incubation of the
fusion enzymes did not result in higheractivities. The some-
what lowered oxidase activities may also be caused by struc-
tural effects of bringing the enzymes together.Nonetheless, it
can be concluded that both fusion partnersofthe created
fusion enzymesshow significant activities. Therefore, we start-
ed to explore their use as bifunctional biocatalysts.
There are numerousapplications in whichthe combined use
of aperoxidaseand oxidase is exploited for detection purpos-
es. One known application that uses such aP-oxidase couple is
the combined use of glucose oxidaseand HRP in biosensors to
determine the glucoselevel in blood.[17] Glucoseoxidase oxi-
dizes glucose to gluconic acidinthe presence of molecular
oxygen, and the formed hydrogen peroxide is subsequently
used to translate the oxidase activity into areadout. Weex-
plored SviDyP–oxidase fusion enzymesfor their use in detect-
Figure 1. Fused oxidase–peroxidases (P-oxidases)enable O2-drivenoxidative
cascade reactions. The cascade reaction from vanillyl alcohol to divanillin is
shownasanexample. Table 1. Peroxidaseand oxidase activities of the fusion enzymes.[a]
Fusion kobs [s@1]Fusion kobs [s@1]
enzymePeroxidaseOxidase enzyme Peroxidase Oxidase
P-ChitO* 7.7 (6.6) 1.0 (6.5[23])P-HMFO 7.1 (6.6) 9.0 (21[21])
P-EugO 5.0 (6.6) 2.5 (12[22])P-HotAldO8.6 (6.6) 0.43 (1.9[20])
[a] The peroxidase activity was measuredbyusing Reactive Blue 19 as a
substrate at pH 4.0. The oxidase activities of P-EugO and P-HMFO towards
vanillylalcohol weremeasured at pH 7.5 and 8.0, respectively.The activity
of P-HotAldOtowards xylitol was measured at pH 7.5,and the activity of
P-ChitO* towardscellobiosewas measured at pH 7.6. The values in paren-
theses indicate the kcat. valuesofthe separate enzymes, as determined for
SviDyP (seethe Supporting Information) or as reported in the literature.
ChemBioChem 2017,18,2226 –2230 T2017 The Authors. PublishedbyWiley-VCH Verlag GmbH &Co. KGaA, Weinheim2227
ing sugars. SviDyP is arepresentative of anewly discovered
class of peroxidases, the DyP-type peroxidases, which have the
advantage over HRPthat they are typicallyeasily overex-
pressed andpurified from aheterologous host, such as
E. coli.[24, 27] First, we produced and probednative SviDyP for its
performance, and found it to be mainly activeatpH3–7 at
ambient temperature, with an optimum for activity towards
Reactive Blue 19 at pH 4.0. SviDyP is active towards 4-aminoan-
tipyrine(AAP) and 3,5-dichloro-2-hydroxybenzenesulfonic acid
(DCHBS), which are commonly used as chromogenicsubstrates
in peroxidase assays (AAP/DCHBS assay). The P-ChitO* and P-
HotAldO fusion enzymes were tested with the AAP/DCHBS
assay for their use in detecting sugars. ChitO is active towards
mono-, di-, and oligosaccharides and is the only oxidase
known to be able to oxidizeN-acetylated carbohydrates.[23, 31]
Various ChitO mutants have been engineered that display dis-
tinct preferences for different carbohydrates. This would allow
the generation of dedicated P-oxidase fusions for the detection
of specific mono-and oligosaccharides. HotAldO is mainly
active on alditols, such as xylitoland sorbitol, which would
allow its use for xylitol or sorbitol sensing.[20] To test the fusion
enzymes, pH 6was used because this was the value at which
optimaofthe oxidases and peroxidase overlapped.With satu-
rating concentrationsoftest sugars (24 mmcellobiose for P-
ChitO* and1.4 mmxylitol for P-HotAldO), aclear and rapid
color developed at arate of 0.3 s@1for both sugars. The rate of
color formation was close to the observed rate when native
SviDyP was tested in the AAP/DCHBS assay (0.4 s@1). This indi-
cates that under the employed conditions the peroxidase is
rate determining in the assay.Amore detailed analysis of the
sensitivity of P-ChitO*and P-HotAldO revealed that the fusion
enzymes were able to detect low levels of cellobiose (25 mm)
and xylitol (10 mm;Figure S3 in the Supporting Information).
When using Amplex Red as afluorogenic peroxidase substrate,
we could even lower the detection limit by one order of mag-
nitude (Figure S4). This shows that such peroxidase–oxidase
fusion enzymes are perfectly suited for sensing purposes by
harboring the full catalytic arsenal for an oxygen-driven bio-
For the generated P-HMFOand P-EugO fusion enzymes,we
exploredtheir use in fully linked cascade reactions. We imag-
ined that, except for the use of the oxidase-generated hydro-
gen peroxide, the aromatic product formed by the oxidases
could also be used as asubstrate for the fused peroxidase.
DyP-type peroxidases have been shown to act on various aro-
matic compounds,whereas HMFO and EugO are, among other
substrates, active on monophenolic compounds.[21, 22, 27–30] This
overlap in substrate/productscope is perfect for one-pot cas-
cade reactions. In earlier work, we showed that another DyP-
type peroxidase, TfuDyP,dimerized vanillyl alcohol, vanillin,
and vanillyl acetone.[28] Dimerization of phenolic compounds is
aknown reaction for peroxidases and laccases, and involves
oxidative phenolic coupling and keto–enol tautomeriza-
tion.[32,33] Divanillin is adesired taste/flavor enhancer and is re-
ported to give an impression of creaminess to food and to
mask the sense of bitterness.[33] In this work, we examined
whether P-EugO and P-HMFO could produce divanillin from
vanillyl alcohol, through acascade reaction in which vanillyl al-
cohol was oxidized to vanillin by an oxidaseand subsequently
dimerized to divanillin by SviDyP (Figure 1). P-EugO and P-
HMFO were incubated with vanillylalcohol at pH 5.5, and the
reaction mixtures were subsequently analyzed by LC-MS (Fig-
ures S5–S14). Both P-EugO andP-HMFO were found to convert
vanillylalcohol. After 21 h, P-HMFO had oxidized 90 %ofvanill-
yl alcohol into vanillin (69 %) and divanillinand related oligo-
mers (21%). Under the same conditions, P-EugO converted
92%ofvanillyl alcohol into vanillin(53 %) and ahigher
amount of oligomers (39 %), of which the most dominant
product was divanillin (see the Supporting Information). These
resultsdemonstrate that the fusion enzymes are suitable for
the productionofthe taste enhancer divanillin.[33] Apart from
being recognized as flavors, vanillin and divanillin are also con-
sidered as renewable buildingblocks for the production of
bio-based plastics.[34–36] Furthermore, divanillin and related phe-
nolic dimers have an antimetastatic potential.[37] Recently,we
developed aone-pot, two-step cascade reaction in which
EugO and HRP or SviDyP were combined to produce low-mo-
lecular-weight lignin-like oligomers from eugenol.[38] The creat-
ed fusion enzymeP-EugO simplifies this newly developed ap-
proach to synthesize lignin oligomer from eugenol. HPLC anal-
ysis revealed that incubation of eugenolwithP-EugO gave the
same lignin products:phenyl coumaran, pinoresinol, coniferyl
alcohol, dieugenol, and alignin tetramer (Figure S15).
In conclusion, we made four active fusion enzymes of DyP-
type peroxidase, SviDyP,and four different oxidases that we
termedP-oxidases. All designed fusion enzymes could be over-
expressed by E. coli as asoluble protein. SviDyP proved to be a
good substitute for HRP in the HRP-coupledassay and could
be appliedatanacidic pH. This SviDyP assay could be applied
to explore the substrate scope of oxidases or as abiosensor
for the detection of, for instance, sugars. SviDyP has an over-
lapping substrate/product scope with multiple oxidases, which
is perfect for cascade reactions. Fusion enzymes P-HMFOand
P-EugO were used in one-pot, two-step cascade reactions. P-
HMFO could be used to prepare divanillin as the main product,
whereas P-EugO could be used for the synthesis of lignin olig-
omers. For future work, it would be interesting to shift thepH
optimaofthe oxidaseand peroxidase closer together.The pH
optimaofseveralenzymes were previously shifted through
site-directed mutagenesis.[39–41] By optimizing these artificial fu-
sions of redox enzymes,novel, effective, bifunctional biocata-
lysts can be developed.
Experimental Section
Chemicals, reagents, and enzymes:Chemicals, media compo-
nents, and reagents were obtained from Sigma(–Aldrich), Merck,
BD, Acros Organics, TCI, Alfa Aesar,Thermo Fisher,and Fisher Sci-
entific. Amplex Red (Amplisyn Red) was obtained from SynChem.
Oligonucleotides and HRP were obtained from Sigma. Restriction
enzyme HindIII was obtained from New England Biolabs. The Pfu-
Ultra Hotstart PCR master mix was from Agilent Te chnologies, and
the In-Fusion HD EcoDry cloning kit was obtained from Clontech.
ChemBioChem 2017,18,2226 –2230 T2017 The Authors. PublishedbyWiley-VCH Verlag GmbH &Co. KGaA, Weinheim2228
Cloning:The genes of oxidases ChitO* (ChitO triple mutant,
Q268R/G270E/S410R), EugO, HMFO, and HotAldO were amplified
and cloned Cterminally to the SviDyP gene in vector pBAD His-
SviDyP;[26] for original and new plasmids, see Ta ble S1 in the Sup-
porting Information. pBAD His-SviDyP contains Cterminally to the
SviDyP gene, astop codon, aHindIII restriction site, and another
stop codon. The vector was linearized by using restriction enzyme
HindIII. The gene of HotAldO (including aC-terminal His6tag, with-
out the first codon for methionine) was cloned into pBAD His-
SviDyP by using restriction free cloning.[42] The obtained plasmid
contained astop codon between the genes of SviDyP and HotAldO.
The stop codon was mutated to serine by QuikChange PCR to
yield vector pBAD His-SviDyP-HotAldO-His. The above-mentioned
stop codon was mutated to serine before cloning of the other ox-
idase genes. The oxidase genes were subsequently amplified and
cloned into the obtained plasmid by In-Fusion cloning (In-Fusion
HD EcoDry cloning kit, Clontech). The HindIII restriction site was re-
tained on both sides of the oxidase genes. E. coli strain TOP10 (In-
vitrogen) was transformed by the obtained plasmids.
Culture growth and enzyme purification:Precultures were grown
on lysogeny broth medium (LB 5mL) at 37 8C, 135 rpm, overnight.
To inoculate 400 mL Te rrific Broth (TB) medium, 1:100 preculture
was added. These cultures were grown at 37 8C, with shaking at
135 rpm, until the OD600 reached about 0.4–0.6, after which they
were induced by 0.02% l-arabinose and grown at 17 8Cand
135 rpm for 70 h. All cultures were supplemented with ampicillin
(50 mgmL@1). Cells were harvested by centrifugation at 6700 gand
48Cfor 20 min (Beckman Coulter,Avanti JE centrifuge, JLA 10.500
rotor). Pellets were washed with buffer A(50 mmpotassium phos-
phate, 0.5mNaCl, pH 8.0), harvested by centrifugation (3000 g,
48C, 40 min, Eppendorf centrifuge 5810R), and stored at @208C
before use. Prior to enzyme purification, the pellets were thawed
and resuspended in buffer Asupplemented with phenylmethane-
sulfonyl fluoride (PMSF;0.1 mm). Cells were disrupted by sonica-
tion (70%amplitude, 5min total on time with cycles of 5son and
10 soff) and the cell-free extract was obtained by centrifugation at
16000 gand 4 8Cfor 15 min (VWR, Micro Star 17R centrifuge). The
enzymes were purified from the cell-free extract by using a5mL
His-Trap HP column (GE Health care). The columns were washed
with buffer Aand buffer Asupplemented with 6, 12, and 24 mm
imidazole. The enzymes were eluted with 300 mmimidazole in
buffer A. Subsequently,the buffer was exchanged to buffer B
(20 mmpotassium phosphate, 150 mmNaCl, pH 7.5) by using a
10 mL Econo-Pac 10 DG desalting column (BioRad). The purified
enzymes were flash frozen with liquid nitrogen and stored at
@208C. UV/Vis absorbance spectra of the enzymes were recorded
between l=250 and 800 nm at ambient temperature (V-660 spec-
trophotometer,Jasco). The protein concentrations were deter-
mined by using the Lambert–Beer law and the predicted molecular
extinction coefficients (ExPASy ProtParam tool[43])were as follows:
e280 nm =48 470 m@1cm@1for SviDyP, e280 nm =124915m@1cm@1for P-
ChitO* (SviDyP-ChitO*, in case it contained one disulfide bond),
e280 nm =127 770 m@1cm@1for P-EugO (SviDyP-EugO), e280 nm =
126850 m@1cm@1for P-HMFO (SviDyP-HMFO), and e280 nm =
116880m@1cm@1for P-HotAldO (SviDyP-HotAldO).
Steady-state kinetic analysis of SviDyP:The steady-state kinetic
parameters of SviDyP were determined for Reactive Blue 19
(e595 nm =10 mm@1cm@1[26])insodium citrate buffer (50 mm,pH4.0)
with H2O2(100 mm)and enzyme (20 nm). SviDyP was added to start
the reaction. Oxidation of Reactive Blue 19 was followed spectro-
photometrically (JASCO V-660) at ambient temperature.
Oxidase and peroxidase activity of the fusion enzymes:The ac-
tivities of both fusion partners were determined separately.For all
reactions, asaturating substrate concentration of 20 times the KM
value was used. For SviDyP,the same reaction conditions were
used as described above, with asubstrate concentration of 100 mm
Reactive Blue 19 (KM=4.6 mm). For the oxidases, the same reaction
mixtures and pH values were used as described before.[20–23] Prior
to the reactions, the oxidases were incubated with 100 mmFADfor
1h at ambient temperature. Vanillyl alcohol was used as asub-
strate for EugO[22] and HMFO,[21] d-(+
+)-cellobiose for ChitO*,[23] and
xylitol for HotAldO.[20] The oxidation of vanillyl alcohol was fol-
lowed spectrophotometrically at l=340 nm (vanillin, e340 nm =
14 mm@1cm@1at pH 7.5[22] and 8.0[21]). The oxidation of d-(++)-cello-
biose and xylitol were followed by means of aHRP-coupled assay.
In this assay,hydrogen peroxide was produced by the oxidases
and used by HRP to couple DCHBS and AAP to apink product
(e515 nm =26 mm@1cm@1).[20, 23]
SviDyP coupled assay for the detection of oxidase substrates:
This assay was avariant of the HRP coupled assay mentioned
above and made use of the peroxidase activity of dye-decolorizing
peroxidase SviDyP instead of HRP.The coupled activity of fusion
enzymes P-ChitO* and P-HotAldO were determined at pH 5
(50 mmsodium citrate buffer) and 6(50 mmpotassium phosphate
buffer). The reaction mixtures contained 0.1 mmAAP,1.0 mm
DCHBS, and 23.8 mm(20 VKM)d-(++)-cellobiose for ChitO* or
1.4 mmxylitol (20VKM)for HotAldO. The formation of the pink
product was followed spectrophotometrically at ambient tempera-
ture (e515 nm =26 mm@1cm@1). To determine whether the oxidase or
peroxidase was the limiting factor in these reactions, the reactions
were repeated in the presence of H2O2(100 mm)todetermine the
optimal reaction rate of SviDyP.
Analysis of the sensitivity of the SviDyP assay:The sensitivity of
the coupled assay was studied by determining the lower concen-
tration limit for substrate detection, as described before.[44] Reac-
tion mixtures (200 mL) contained 0.1 mmAAP,1.0 mmDCHBS,
150 nmfusion enzyme, and varying substrate concentrations
(0.5 mm–1 mm)in50mmpotassium phosphate buffer pH 6.0.
d-(++)-Cellobiose and xylitol were used as substrates for P-ChitO*
and P-HotAldO, respectively.The enzymes were added to start the
reaction. Reactions were performed in triplicate and the absorb-
ance at l=515 nm (pink product, e515 nm =26 mm@1cm@1)was fol-
lowed at ambient temperature for 15 min by using aSynergyMX
(BioTek) plate reader.The obtained values after 15 min were cor-
rected for both the path length and the blank. For comparison, the
sensitivity of the coupled assay was also determined by using
60 mmAmplex Red (10-acetyl-3,7-dihydroxyphenoxazine, Amplisyn
Red) instead of AAP/DCHBS. Astock of 6.0 mmAmplex Red was
prepared in DMSO. The oxidation of Amplex Red was followed by
measuring the fluorescence of the product, resorufin (lex =530 nm,
lem =590 nm), for 15 min at ambient temperature.
One-pot cascade reaction for the synthesis of divanillin and re-
lated dimers and oligomers:Vanillyl alcohol was dissolved in
water at aconcentration of 50 mm.Reaction mixtures (2.0 mL) con-
tained 2mmvanillyl alcohol and 1.0 mmSviDyP,P-HMFO, or P-
EugO in sodium citrate buffer (50 mm,pH5.5). In the case of
SviDyP,500 mmH2O2was added. Reaction mixtures were incubated
in 15 mL closed tubes at 30 8Cand 100 rpm, for 21 h. Control reac-
tions were prepared without enzyme. After 2, 3, and 21 h, samples
were taken. Enzymes were heat-inactivated at 95 8Cfor 10 min,
after which time the samples were centrifuged for 5min at
16,100Vg. Reaction products were analyzed by reversed-phase
HPLC by using aJasco HPLC system. Samples (10 mL) were injected
ChemBioChem 2017,18,2226 –2230 T2017 The Authors. PublishedbyWiley-VCH Verlag GmbH &Co. KGaA, Weinheim2229
onto aGrace Altima HP C18 column (5 mm, 2.1V150 mm, with
1.0 cm precolumn of the same material). Solvents used were as fol-
lows:A:water with 0.1 %formic acid;B:acetonitrile. HPLC
method:2min 10%B,2–20 min gradient to 70 %B,20–23 min
70%B,23min 10 %B,followed by 7min re-equilibration. Detec-
tion by aUVdetector at l=280 nm and flow rate of 0.5 mL min@1.
LC-MS analysis was performed on aSurveyor HPLC-DAD instru-
ment coupled to an LCQ Fleet detector by using scanning for both
positive and negative modes. Samples were injected onto aGrace
Altima HP C18 column (3 mm, 2.1 V100 mm, with 1.0 cm precolumn
of the same material), flow rate 0.3 mL min@1.Solvents used were
as follows:A:water with 0.1%formic acid;B:acetonitrile with
0.08%formic acid. LC-MS method:2min 100%A,2–32 min gradi-
ent to 80%B,32–37 min 80 %B,37–38 min 100 %A,38–48 min
One-pot cascade reaction for the synthesis of lignin-like oligo-
mers from eugenol:The activity of P-EugO towards eugenol was
assayed as described before.[38] Reaction mixtures (2.0 mL) con-
tained 1.0 mmP-EugO, 10 mmeugenol, and 5% DMSO (v/v) in po-
tassium phosphate buffer (20 mm,pH6.0). Astock solution of
300 mmeugenol was prepared in DMSO. For comparison, areac-
tion mixture containing SviDyP (1.0 mm)and EugO (1.0 mm)was as-
sayed. All reactions were performed in duplicate and compared
with areaction without enzyme. Reaction mixtures were incubated
at 308Cand 50 rpm in 20 mL Pyrex tubes with aheadspace to
volume ratio of 10:1. Samples (200 mL) were taken after 24 and
96 h. These samples were heat-treated and analyzed by reversed-
phase HPLC, as described above, for the production of divanillin
and related oligomers.
This work was supported by the NWOgraduate program: syn-
thetic biology for advanced metabolic engineering, project
number022.004.006, The Netherlands.
Conflict of Interest
The authors declare no conflict of interest.
Keywords: biocatalysis ·domino reactions ·enzymes ·protein
engineering ·sensors
[1] E. Schweizer,J.Hofmann, Microbiol. Mol. Biol. Rev. 2004,68,501 –517.
[2] M. S. Patel,N.S.Nemeria, W. Furey,F.Jordan, J. Biol. Chem. 2014,289,
[3] R. H. Doi, A. Kosugi, Nat. Rev.Microbiol. 2004,2,541–551.
[4] S. Elleuche, Appl. Microbiol. Biotechnol. 2015,99,1545–1556.
[5] I. P8rez-Arellano,F.Carmona-]lvarez, A. I. Mart&nez, J. Rodr&guez-D&az, J.
Cervera, ProteinSci. 2010,19,372–382.
[6] I. G. Charles, J. W. Keyte, W. J. Brammar,M.Smith, A. R. Hawkins, Nucleic
Acids Res. 1986,14,2201 –2213.
[7] J. Yourno, T. Kohno, J. R. Roth, Nature 1970,228,820–824.
[8] K. Yu,C.Liu, B.-G. Kim, D.-Y.Lee, Biotechnol. Adv. 2015,33,155–164.
[9] R. J. Conrado, J. D. Varner,M.P.Delisa, Curr.Opin. Biotechnol. 2008,19,
[10] S. Matthews, K. L. Tee, N. J. Rattray,K.J.Mclean, D. Leys,D.A.Parker,
R. T. Blankley, A. W. Munro, FEBS Lett. 2017,591,737 –750.
[11] D. E. To rres PazmiÇo, R. Snajdrova, B.-J. Baas, M. Ghobrial, M. D. Mihovi-
lovic, M. W. Fraaije, Angew.Chem. Int. Ed. 2008,47,2275–2278; Angew.
Chem. 2008,120,2307 –2310.
[12] N. Beyer,J.K.Kulig, A. Bartsch, M. A. Hayes, D. B. Janssen,M.W.Fraaije,
Appl. Microbiol. Biotechnol. 2017,101,2319 –2331.
[13] I. Meynial Salles, N. Forchhammer,C.Croux,L.Girbal, P. Soucaille,
Metab.Eng. 2007,9,152–159.
[14] H. S. Seo, Y. J. Koo, J. Y. Lim, J. T. Song, C. H. Kim, J. K. Kim, J. S. Lee, Y. D.
Choi, Appl. Environ. Microbiol. 2000,66,2484–2490.
[15] A. M. Abdel-Hamid, J. O. Solbiati, I. K. O. Cann, Adv.Appl. Microbiol.
[16] P. Ander,L.Marzullo, J. Biotechnol. 1997,53,115 –131.
[17] D. Barham, P. Trinder, Analyst 1972,97,142–145.
[18] H. J. Chun, Y. M. Park, Y. D. Han, Y. H. Jang, H. C. Yo on, BiochipJ.2014,8,
[19] R. Mundaca-Uribe, F. Bustos-Ram&rez, C. Zaror-Zaror,M.Aranda-Bustos,
J. Neira-Hinojosa, C. PeÇa-Farfal, Sens.Actuators B 2014,195,58–62.
[20] R. T. Winter,D.P.H.M.Heuts, E. M. A. Rijpkema, E. van Bloois,H.J.
Wijma, M. W. Fraaije, Appl. Microbiol. Biotechnol. 2012,95,389 –403.
[21] W. P. Dijkman, M. W. Fraaije, Appl. Environ. Microbiol. 2014,80,1082–
[22] J. Jin, H. Mazon, R. H. H. van denHeuvel, D. B. Janssen,M.W.Fraaije,
FEBS J. 2007,274,2311–2321.
[23] A. R. Ferrari, M. Lee, M. W. Fraaije, Biotechnol. Bioeng. 2015,11 2,1074
[24] F. W. Krainer, A. Glieder, Appl. Microbiol. Biotechnol. 2015,99,1611–
[25] W. Yu,W.Liu, H. Huang,F.Zheng, X. Wang, Y. Wu, K. Li, X. Xie, Y. Jin,
PLoS One 2014,9,e110319.
[26] D. I. Colpa, M. W. Fraaije, J. Mol. Catal. B 2016,134,372–377.
[27] D. I. Colpa, M. W. Fraaije, E. van Bloois, J. Ind. Microbiol. Biotechnol. 2014,
[28] N. Lonc
ˇar,D.I.Colpa, M. W. Fraaije, Te trahedron 2016,72,7276 –7281.
[29] H. J. O. Ogola, T. Kamiike,N.Hashimoto, H. Ashida,T.Ishikawa,H.Shiba-
ta, Y. Sawa, Appl. Environ. Microbiol. 2009,75,7509 –7518.
[30] S. J. Kim, M. Shoda, Appl. Environ. Microbiol. 1999,65,1029–1035.
[31] D. P. H. M. Heuts, D. B. Janssen,M.W.Fraaije, FEBS Lett. 2007,581,
[32] R. T. Nishimura, C. H. Giammanco,D.A.Vosburg, J. Chem. Educ. 2010,
87,526 –527.
[33] U. Krings, V. Esparan, R. G. Berger, Flavour Fragrance J. 2015,30,362
[34] M. Fache, B. Boutevin, S. Caillol, Eur.Polym. J. 2015,68,488–502.
[35] A. Llevot, E. Grau, S. Carlotti, S. Grelier,H.Cramail, Polym. Chem. 2015,
6,7693 –7700.
[36] A. S. Amarasekara, A. Razzaq, ISRN Polym. Sci. 2012,2012,1–5.
[37] P. Jantaree, K. Lirdprapamongkol, W. Kaewsri, C. Thongsornkleeb,K.
Choowongkomon, K. Atjanasuppat, S. Ruchirawat, J. Svasti, J. Agric.
Food Chem. 2017,65,2299–2306.
[38] M. H. M. Habib, P. J. Deuss, N. Lonc
ˇar,M.Trajkovic, M. W. Fraaije, Adv.
Synth. Catal. 2017,
[39] S. Pokhrel, J. C. Joo, Y. J. Yo o, Biotechnol. BioprocessEng. 2013,18,35–
[40] S. Mendes, V. Brissos, A. Gabriel, T. Catarino, D. L. Turner,S.Todorovic,
L. O. Martins, Arch. Biochem.Biophys. 2015,574,99–107.
[41] A. To mschy,R.Brugger,M.Lehmann, A. Svendsen,K.Vogel, D. Kostre-
wa, S. F. Lassen,D.Burger,A.Kronenberger,A.P.G.M.van Loon,etal.,
Appl. Environ. Microbiol. 2002,68,1907 –1913.
[42] F. van den Ent, J. Lçwe, J. Biochem. Biophys. Methods 2006,67,67–74.
[43] E. Gasteiger,C.Hoogland, A. Gattiker,S.Duvaud, M. R. Wilkins, R. D.
Appel, A. Bairoch in Proteomics Protocols Handbook (Ed. :J.M.Walker),
Humana, Totowa, 2005,pp. 571 –607.
[44] A. R. Ferrari, Y. Gaber,M.W.Fraaije, Biotechnol. Biofuels 2014,7,37.
Manuscript received:September 6, 2017
Accepted manuscript online:September 8, 2017
Version of record online :October 11,2017
ChemBioChem 2017,18,2226 –2230 T2017 The Authors. PublishedbyWiley-VCH Verlag GmbH &Co. KGaA, Weinheim2230

Supplementary resource (1)

... As a result, several variants of ChitO were created with distinct substrate acceptance profiles, including variants that behave as a D-glucosamine oxidase or a cellobiose oxidase . A variant of the ChitO enzyme having high specificity for cellobiose was fused to a peroxidase (Colpa et al., 2017). This allows detection of low amounts of cellobiose. ...
... Also, enzyme engineering can be of use. Recently, the fusion of carbohydrate oxidases to peroxidases was shown to be possible, allowing a sensitive biosensing system for specific carbohydrates which relies only on one biocatalyst (Colpa et al., 2017). ...
Full-text available
Carbohydrates are widely abundant molecules present in a variety of forms. For their biosynthesis and modification, nature has evolved a plethora of carbohydrate-acting enzymes. Many of these enzymes are of particular interest for biotechnological applications, where they can be used as biocatalysts or biosensors. Among the enzymes catalysing conversions of carbohydrates are the carbohydrate oxidases. These oxidative enzymes belong to different structural families and use different cofactors to perform the oxidation reaction of CH-OH bonds in carbohydrates. The variety of carbohydrate oxidases available in nature reflects their specificity towards different sugars and selectivity of the oxidation site. Thanks to their properties, carbohydrate oxidases have received a lot of attention in basic and applied research, such that nowadays their role in biotechnological processes is of paramount importance. In this review we provide an overview of the available knowledge concerning the known carbohydrate oxidases. The oxidases are first classified according to their structural features. After a description on their mechanism of action, substrate acceptance and characterisation, we report on the engineering of the different carbohydrate oxidases to enhance their employment in biocatalysis and biotechnology. In the last part of the review we highlight some practical applications for which such enzymes have been exploited.
... Although oxidase-peroxidase fusion enzymes have not been observed in nature, they are often co-expressed because they are interconnected since oxidases produce hydrogen peroxide, which in turn is a substrate for peroxidases (Colpa et al., 2017). ...
Full-text available
Fusion proteins, understood as those created by joining two or more genes that originally encoded independent proteins, have numerous applications in biotechnology, from analytical methods to metabolic engineering. The use of fusion enzymes in biocatalysis may be even more interesting due to the physical connection of enzymes catalyzing successive reactions into covalently linked complexes. The proximity of the active sites of two enzymes in multi-enzyme complexes can make a significant contribution to the catalytic efficiency of the reaction. However, the physical proximity of the active sites does not guarantee this result. Other aspects, such as the nature and length of the linker used for the fusion or the order in which the enzymes are fused, must be considered and optimized to achieve the expected increase in catalytic efficiency. In this review, we will relate the new advances in the design, creation, and use of fused enzymes with those achieved in biocatalysis over the past 20 years. Thus, we will discuss some examples of genetically fused enzymes and their application in carbon‑carbon bond formation and oxidative reactions, generation of chiral amines, synthesis of carbohydrates, biodegradation of plant biomass and plastics, and in the preparation of other high-value products.
... It has been pointed out that the spatial confinement of different biocatalysts operating in parallel or in sequence can drastically reduce diffusion distances of the reagents and thereby accelerate the rate of biocatalytic cascade reactions [2]. Next to co-immobilization of different enzymes [3] or confinement in vesicles [4] also genetically fused enzymes [2,5,6] have been investigated. From these studies it can be concluded that generally, spatial proximity represents an advantage for cascade catalysis as diffusion distances of the individual reagents are reduced. ...
Hydrocarbon synthesis from (waste)oils enabled by a cascade of lipase-catalysed hydrolysis and decarboxylase-catalysed decarboxylation has become an active area of research en route to alternative, biobased fuels. However, Poor substrate transport efficiency is a major issue causing low reaction rates. This study focused on a protein self-assembly strategy based on SpyTag/SpyCatcher to overcome diffusion limitations. For this, two fusion proteins, TLL-Linker-SpyCatcher based on the lipase from Thermomyces lanuginosus and CvFAP-Linker-SpyTag based on the fatty acid photodecarboxylase from Chlorella variabilis were designed. A covalent multi-enzyme complex (TLL-CvFAP) was formed spontaneously by self-assembly of each enzyme. The effects of temperature, pH and molar ratio of self-assembled components on assembly efficiency were investigated. The results showed that the multi-enzyme complex TLL-CvFAP reached about 60% after 12 h of assembly, and the enzyme activity of the multienzyme complex was increased by about 50% compared to that of the corresponding non-assembled enzymes. Under optimized conditions 10 mM soybean oil were converted into 25 mM of the corresponding hydrocarbons, suggesting a good potential of biofuel synthesis.
... OD is an enzyme of the oxidoreductases family which catalyzes an oxidation reduction reaction, exclusively involving molecular dioxygen (O 2 ) as the electron acceptor. 48 The reaction mechanism keeps on with the donation of a hydrogen atom, and the molecular oxygen gets reduced to hydrogen peroxide (H 2 O 2 ) or water (H 2 O). 49 OD-like activities of CeZrO 4 were estimated by the colorimetric assay, performing the reaction between a chromogenic substrate TMB, dopamine, and catalyst without a supplementary oxidizing agent (H 2 O 2 ). ...
Full-text available
The myth of inactivity of inorganic materials in a biological system breaks down by the discovery of nanozymes. From this time, the nanozyme has attracted huge attention for its high durability, cost-effective production, and easy storage over the natural enzyme. Moreover, the multienzyme-mimicking activity of nanozymes can regulate the level of reactive oxygen species (ROS) in an intercellular system. ROS can be generated by peroxidase (POD), oxidase (OD), and Fenton-like catalytic reaction by a nanozyme which kills the cancer cells by oxidative stress; therefore, it is important in CDT (chemo dynamic therapy). Our current study designed to investigate the enzyme mimicking behavior and anticancer ability of cerium-based nanomaterials because the cerium-based materials offer a high redox ability while maintaining nontoxicity and high stability. Our group synthesized CeZrO4 nanoparticles by a green method using β-cyclodextrin as a stabilizer and neem leaf extract as a reducing agent, exhibiting POD- and OD-like dual enzyme activities. The best enzyme catalytic activity is shown in pH = 4, indicating the high ROS generation in an acidic medium (tumor microenvironment) which is also supported by the Fenton-like behavior of CeZrO4 nanoparticles. Inspired by the high ROS generation in vitro method, we investigated the disruption of human kidney cells by this nanoparticle, successfully verified by the MTT assay. The harmful effect of ROS in a normal cell is also investigated by the in vitro MTT assay. The results suggested that the appreciable anticancer activity with minimal side effects by this synthesized nanomaterial.
... Since the peroxide sensitivity is an issue presented by many different heme-containing proteins [17,18], strategies like the combination with oxidases for in situ production of H2O2 have been attempted [19]. Another option has been the modification of the enzyme to increase its stability towards its substrate since it is generally considered that the presence of sensitive amino acids close to the heme group is responsible for the peroxide inactivation. ...
Full-text available
The dye-decolorizing peroxidases (DyP) are a family of heme-dependent enzymes present on a broad spectrum of microorganisms. While the natural function of these enzymes is not fully understood, their capacity to degrade highly contaminant pigments such as azo dyes or anthraquinones make them excellent candidates for applications in bioremediation and organic synthesis. In this work, two novel DyP peroxidases from the organism Rhodococcus opacus 1CP (DypA and DypB) were cloned and expressed in Escherichia coli. The enzymes were purified and biochemically characterized. The activities of the two DyPs via 2,2′-azino-bis [3-ethylbenzthiazoline-6-sulphonic acid] (ABTS) assay and against Reactive Blue 5 were assessed and optimized. Results showed varying trends for DypA and DypB. Remarkably, these enzymes presented a particularly high tolerance towards H2O2, retaining its activities at about 10 mM H2O2 for DypA and about 4.9 mM H2O2 for DypB.
Human health is significantly impacted by fluorides in water.Too much fluoride consumption through drinking water causes chronic dental and bones diseases. A trace number of fluorides in drinking water can be determined, which is immensely vital for human health. In the current study, we have designed a visual colorimetric chemosensor assay using the brilliant oxidase mimicking activity of biosynthesized FeMnO4@GQD nanocomposite. FeMnO4@GQD nanocomposites showed excellent mimicking activity in mild acid medium pH‐5, and obtained a Vmax value of 1.71603×10−6 and an important Km value of 0.0684 mM based on Hanes‐Woolf kinetics. The robust nanocomposites were shown to be significant stability for detecting oxidases under conditions. As a consequence, oxidase‐like activity was inhibited in the presence of fluoride ions. A linear correlation was established by the development of blue colour with fluoride ions concentration and appeared at a low limit of detection of 1.2 μg/ml without interfering with other common ions. Oxidase‐like activity was inhibited in the presence of fluoride ions. A linear correlation was established by the development of blue colour with fluoride ions concentration and appeared at a low limit of detection of 1.2 μg/ml without interfering with other common ions.
Biomass being a renewable source of energy, has emerged as an attractive target for manufacturing valuable products. These possibilities can be explored to meet the current need for degradable plastic, 2,5-furandicarboxylic acid (FDCA). Integration of chemical and biological approaches for direct biomass conversion into FDCA was evaluated in this study. 5-hydroxymethylfurfural (5-HMF) was chemo-catalytically obtained from fructose using recyclable amberlyte IR-120 as a catalyst with >98% purity. Klebsiella oxytoca NCIM 2694 bacterial strain showed the potential of converting 98% of 5-HMF into FDCA with 58% selectivity at 96 h. With optimized conditions of pH 7, 37 °C, 2 g whole cells, we reported 99% 5-HMF conversion to FDCA with 95% selectivity at 72 h and 2667 mg L⁻¹ yield, with 39 mg L⁻¹ h⁻¹ productivity. This is the highest yield obtained with the substrate concentration as high as 3000 mg L⁻¹ reported till date. While the bacterial tolerance to 5-HMF observed was for the highest 5-HMF concentration of 4000 mg L⁻¹, with 99% conversion however, compromising the FDCA yield to 2447 mg L⁻¹ and 32 mg L⁻¹ h⁻¹ productivity. Atom economy of 85% and E factor of 17.71 g g⁻¹ was obtained as a measure of its efficiency and sustainability of the process. The developed process will decrease the cost by excluding any extra nutrient supplement, complete substrate utilization, highest FDCA selectivity/productivity and higher tolerance by K. oxytoca, sequentially catalyzing the oxidations by a single route for FDCA synthesis from renewables.
Oxidative biotransformations find a prominent role in the fine chemical industry and the valorization of renewable feedstocks. Implementation of oxygen-dependent reactions faces some challenges across scales and at different levels of development. First, the fruitful development of enzyme candidates and identification of reaction possibilities is not in consonance with the implementation in process engineering. Second, reaction engineering faces a complex interplay of reaction kinetic, oxygen transfer and process stability. Third, given the advances in synergic fields such as molecular biology, chemistry, material sciences and (micro)process engineering, an interdisciplinary assembly from a consistent discipline around heterogeneous biocatalyst engineering would be of strategic value. We show advances in design of active and robust immobilized enzyme catalysts to be applied in (continuous) intensified processes. A framework based on joint design of catalyst and reactor will be discussed for the design and optimization of the catalysts and biotransformations involved.
Within the cellular microenvironment of organisms, the surface‐ and volume‐ enclosed multiple enzymes perform highly complex series of chemical reactions for very selective and efficient synthesis of biomolecules. Several efforts have been carried out to mimic these timely and spatially defined natural enzyme cascade reactions onto the artificial multienzyme nano‐assemblies via immobilization of two or more enzymes on nanosupport, while maintaining the catalytic activity. These biocatalytic systems undergo cascade reactions for the efficient production of chemicals and materials of interest. Their increasing interest is based on their numerous potential applications in the area of environmental, biomedical and applied chemistry. A variety of nanosupports and multienzyme immobilization techniques have been optimized and reported. Nanoreactors usually show molecular crowding and anomalous diffusion of substances that modify the mass action kinetic laws. Thus, the kinetics is different than those shown by free diluted enzymes. Consequently, the catalytic activity considerations in cascade reactions on nanoassemblies are of great importance. Thus, in this work, important parameters of enzyme kinetics in cascade reaction and the molecular tools for the design and optimization of bi‐ and multi‐enzymatic nanobioreactors reactions are discussed.
Full-text available
Jeotgalicoccus sp. 8456 OleTJE (CYP152L1) is a fatty acid decarboxylase cytochrome P450 that uses hydrogen peroxide (H2 O2 ) to catalyse production of terminal alkenes, which are industrially important chemicals with biofuel applications. We report enzyme fusion systems in which Streptomyces coelicolor alditol oxidase (AldO) is linked to OleTJE . AldO oxidizes polyols (including glycerol), generating H2 O2 as a co-product and facilitating its use for efficient OleTJE -dependent fatty acid decarboxylation. AldO activity is regulatable by polyol substrate titration, enabling control over H2 O2 supply to minimise oxidative inactivation of OleTJE and prolong activity for increased alkene production. We also use these fusion systems to generate novel products from secondary turnover of 2-OH and 3-OH myristic acid primary products, expanding the catalytic repertoire of OleTJE . This article is protected by copyright. All rights reserved.
Full-text available
To facilitate the wider application of the NADPH-dependent P450BM3, we fused the monooxygenase with a phosphite dehydrogenase (PTDH). The resulting monooxygenase-dehydrogenase fusion enzyme acts as a self-sufficient bifunctional catalyst, accepting phosphite as a cheap electron donor for the regeneration of NADPH. The well-expressed fusion enzyme was purified and analyzed in comparison to the parent enzymes. Using lauric acid as substrate for P450BM3, it was found that the fusion enzyme had similar substrate affinity and hydroxylation selectivity while it displayed a significantly higher activity than the non-fused monooxygenase. Phosphite-driven conversions of lauric acid at restricted NADPH concentrations confirmed multiple turnovers of the cofactor. Interestingly, both the fusion enzyme and the native P450BM3 displayed enzyme concentration dependent activity and the fused enzyme reached optimal activity at a lower enzyme concentration. This suggests that the fusion enzyme has an improved tendency to form functional oligomers. To explore the constructed phosphite-driven P450BM3 as a biocatalyst, conversions of the drug compounds omeprazole and rosiglitazone were performed. PTDH-P450BM3 driven by phosphite was found to be more efficient in terms of total turnover when compared with P450BM3 driven by NADPH. The results suggest that PTDH-P450BM3 is an attractive system for use in biocatalytic and drug metabolism studies. Electronic supplementary material The online version of this article (doi:10.1007/s00253-016-7993-7) contains supplementary material, which is available to authorized users.
Full-text available
A simple paper-based optical biosensor for glucose monitoring was developed. As a glucose biosensing principle, a colorimetric glucose assay, using glucose oxidase (GOx) and horseradish peroxidase (HRP), was chosen. The enzymatic glucose assay was implanted on the analytical paper-based device, which is fabricated by the wax printing method. The fabricated device consists of two paper layers. The top layer has a sample loading zone and a detection zone, which are modified with enzymes and chromogens. The bottom layer contains a fluidic channel to convey the solution from the loading zone to the detection zone. Double-sided adhesive tape is used to attach these two layers. In this system, when a glucose solution is dropped onto the loading zone, the solution is transferred to the detection zone, which is modified with GOx, HRP, and chromogenic compounds through the connected fluidic channel. In the presence of GOx-generated H2O2, HRP converts chromogenic compounds into the final product exhibiting a blue color, inducing color change in the detection zone. To confirm the changes in signal intensity in the detection zone, the resulting image was registered by a digital camera from a smartphone. To minimize signal interference from external light, the experiment was performed in a specifically designed light-tight box, which was suited to the smartphone. By using the developed biosensing system, various concentrations of glucose samples (0–20 mM) and human serum (5–17 mM) were precisely analyzed within a few minutes. With the developed system, we could expand the applicability of a smartphone to bioanalytical health care. © 2014, The Korean BioChip Society and Springer-Verlag Berlin Heidelberg.
Synthetic lignin was prepared biocatalytically in a one-pot two-step reaction using an oxidase/peroxidase cascade enzyme system. Using eugenol in combination with eugenol oxidase and a peroxidase, lignin-like material was produced. The cascade reaction takes advantage of the ability of the oxidase to produce coniferyl alcohol and hydrogen peroxide from eugenol and molecular oxygen. The hydrogen peroxide is used by the peroxidase for the for-mation of crosslinks that typify lignin. As eugenol oxidase has a broad substrate acceptance profile, also 4-allylphenol (chavicol) and 4-allyl-2,6-dimethoxyphenol could be used as precursors of the synthetic lignin. As a result, all three naturally occurring monolignols could be prepared and incorporated in the synthetic lignin. The reaction was optimized in order to achieve the highest possible yield of insoluble lignin oligomers and scaled up to 1 gram. Analysis of the water-insoluble product by gel permeation chromatography revealed the formation of relatively small lignin oligomers (~1000 dalton). By using two-dimensional heteronuclear single quantum coherence nuclear magnetic resonance spectroscopy and gas chromatography-mass spectrometry analyses it could be demonstrated that the material contained α-O-4/β-O-4, β-O-4, β-β, β-5 linkages and dibenzodioxocin units. All these features indicate that the biocatalytically produced material closely resembles natural lignin. While 54% of eugenol was converted into water-insoluble oligomers, the remaining substrate was converted into water soluble dimers and tetramers which are important lignin model compounds. Therefore, the presented method represents a valuable and facile biocatalytic approach for the preparation of lignin-like material and potentially valuable chemicals.
The spread of cancer cells to distant organs, in a process called metastasis, is the main factor that contributes to most death in cancer patients. Vanillin, the vanilla flavoring agent, has been shown to suppress metastasis in a mouse model. Here, we evaluated the anti-metastatic potential of the food additive divanillin, the homo-dimer of vanillin, and their structurally related compounds, apocynin and diapocynin, in hepatocellular carcinoma cells. The Transwell invasion assay showed that the dimeric forms exhibited higher potency than vanillin and apocynin in inhibiting invasion, with IC50 values of 23.3±7.4 to 41.3±4.2 μM for the dimers, which are 26-34 fold lower than IC50 values of vanillin and apocynin (p<0.05). Both monomeric and dimeric forms target regulation of the invasion process, by inhibiting phosphorylation of FAK and Akt. Molecular docking studies suggested that the dimers should bind more tightly than vanillin and apocynin to the Y397 pocket of FAK FERM domain. Thus, the food additive divanillin has greater anti-metastatic potential than the flavoring agent vanillin.
The heterologous overexpression level of the bacterial dye decolorizing peroxidase TfuDyP in Escherichia coli was increased sixty fold to approximately 200 mg of purified enzyme per liter culture broth by fusing the enzyme to the small ubiquitin-related modifier protein (SUMO). The highly overexpressed SUMO-TfuDyP was, however, almost inactive. Analysis of the enzyme by UV–vis absorption spectroscopy and high-resolution mass spectrometry showed that a large fraction of the highly overexpressed enzyme contained the iron deficient heme precursor protoporphyrin IX (PPIX) instead of heme. Here we show that the activity of the enzyme was dependent on the expression level of the protein.
Dye-decolorizing peroxidases (DyPs) represent a new class of oxidative enzymes for which the natural substrates are largely unknown. To explore the biocatalytic potential of a DyP, we have studied the substrate acceptance profile of a robust DyP peroxidase, a DyP from Thermobifida fusca (TfuDyP). While previous work established that this bacterial peroxidase is able to act on a few reactive dyes and aromatic sulfides, this work significantly expands its substrate scope towards lignin related compounds, flavors, and various dyes.
Four biphenyl monomers derived from vanillin or eugenol were synthesized and polymerized by the ADMET methodology. The biphenyl compounds were produced by enzymatic dimerization of 2-methoxy-4-methylphenol, methyl vanillate, vanillin and eugenol. Further chemical modifications of the obtained dimers such as transesterification, Wittig reaction or allylation led to α,ω-dienes. The reactivity of these bio-based monomers towards ADMET polymerization employing several catalysts was investigated. Only oligomers were obtained when the diallylated compound was employed. The polymers based on dieugenol and the bis-unsaturated diester exhibit good thermal stability and Tg at 17 and 54 °C, respectively. Polymerization of the divinyl compound obtained by the Wittig reaction of divanillin showed a reasonable molar mass of 30000 g mol-1, a high Tg at around 160 °C and thermostability with a 5% weight loss occurring at 380 °C.
The use of vanillin for the preparation of renewable polymers is reviewed in this work. The synthesis of polymers from renewable resources is a burning issue that is actively investigated. Vanillin is currently one of the only biobased and aromatic compounds that are industrially available. For this reason, vanillin recently gained much attention from the polymer community. The first part of this work aims at giving an overview of the different existing sources of vanillin, and of their relevance in the context of a potential use in polymer science. The second part of this work sums up the efforts of the scientific community to prepare a wide range of vanillin-based polymers, e.g. phenolic, epoxy and benzoxazine resins, polyesters, acrylate and methacrylate polymers. The interest in the use of vanillin to prepare renewable polymers is recent but the number of contributions on this subject is growing fast.
Dehydrodivanillin, the symmetrical dimer of vanillin, is a taste enhancer which imparts pleasant impressions of creaminess to food. Found in vanilla pods in traces only, a co-substrate independent dimerization of vanillin, conducted in a co-solvent system to improve the solubility of vanillin, was developed using iso-active fungal laccases from Meripilus giganteus, Agaricus bisporus and Funalia trogii. The yields were compared with a peroxidase from Marasmius scorodonius (MsP2) and the reference enzyme horseradish peroxidase (HRP), both supplied with hydrogen peroxide. Using laccase catalysis, the kinetically preferred reaction product, 5,5'-dehydrodivanillin, rapidly reached saturation and precipitated in situ, thus shifting the reaction equilibrium to the product. Yields of > 95 % were obtained with the high-redox-potential laccase of Funalia trogii, while HRP gave 18 %. Copyright © 2015 John Wiley & Sons, Ltd.