Citation: Bauer, J.A.; Zámocká, M.;
Majtán, J.; Bauerová-Hlinková, V.
Glucose Oxidase, an Enzyme
“Ferrari”: Its Structure, Function,
Production and Properties in the
Light of Various Industrial and
Biotechnological Applications.
Biomolecules 2022, 12, 472. https://
doi.org/10.3390/biom12030472
Academic Editor: Stanislav Stuchlík
Received: 28 February 2022
Accepted: 17 March 2022
Published: 19 March 2022
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biomolecules
Review
Glucose Oxidase, an Enzyme “Ferrari”: Its Structure, Function,
Production and Properties in the Light of Various Industrial and
Biotechnological Applications
Jacob A. Bauer
1
, Monika Zámocká
1
, Juraj Majtán
1,2
and Vladena Bauerová-Hlinková
1,
*
1
Institute of Molecular Biology, Slovak Academy of Sciences, 845 51 Bratislava, Slovakia;
2
Department of Microbiology, Faculty of Medicine, Slovak Medical University, Limbová 12,
833 03 Bratislava, Slovakia
* Correspondence: [email protected]; Tel.: +421-2-5930-7434
Abstract:
Glucose oxidase (GOx) is an important oxidoreductase enzyme with many important roles
in biological processes. It is considered an “ideal enzyme” and is often called an oxidase “Ferrari”
because of its fast mechanism of action, high stability and specificity. Glucose oxidase catalyzes the
oxidation of
β
-D-glucose to D-glucono-
δ
-lactone and hydrogen peroxide in the presence of molecular
oxygen. D-glucono-
δ
-lactone is sequentially hydrolyzed by lactonase to D-gluconic acid, and the
resulting hydrogen peroxide is hydrolyzed by catalase to oxygen and water. GOx is presently known
to be produced only by fungi and insects. The current main industrial producers of glucose oxidase
are Aspergillus and Penicillium. An important property of GOx is its antimicrobial effect against
various pathogens and its use in many industrial and medical areas. The aim of this review is to
summarize the structure, function, production strains and biophysical and biochemical properties of
GOx in light of its various industrial, biotechnological and medical applications.
Keywords:
glucose oxidase; catalytic mechanism; GOx producing organisms; FAD binding domain;
biosensors; nanosensors; antimicrobial effect
1. Introduction
Glucose oxidase is an enzyme that has widespread applications in industry and
biotechnology. Due to this, a deep understanding of its structure and function are war-
ranted. Glucose degradation is the most universal metabolic process. In addition to its
breakdown in glycolysis, glucose can also be directly oxidized to glucono-
δ
-lactone by a
number of enzymes.
These fall into two classes: (1) the dehydrogenases glucose dehydrogenase (
β
-D-
glucose: NAD(P)
+
1-oxidoreductase, E.C. 1.1.1.47) [
1
] and quinoprotein glucose dehydro-
genase (D-glucose:ubiquinone oxidoreductase, E.C. 1.1.5.2) [
2
] and (2) the oxidases glucose
oxidase (GOx;
β
-D-glucose:oxygen 1-oxidoreductase, E.C. 1.1.3.4) [
3
] and pyranose oxidase
(pyranose:oxygen 2-oxidoreductase, E.C. 1.1.3.10) [4].
The dehydrogenases oxidize glucose in one step using a co-factor, either nicotinamide
adenine dinucleotide (phosphate) (NAD(P)
+
) or pyrroloquinoline quinone (PQQ), as the
electron sink while the oxidases use a two-step mechanism in which a bound flavin adenine
dinucleotide (FAD) co-factor is used to oxidize glucose to form glucono-
δ
-lactone and an
enzyme-FADH
2
intermediate followed by electron transfer to O
2
to form H
2
O
2
(Figure 1).
The principal difference between GOx and pyranose oxidase is that the former is specific to
β-D-glucose while the latter is also able to act on D-xylose, L-sorbose and D-galactose.
Biomolecules 2022, 12, 472. https://doi.org/10.3390/biom12030472 https://www.mdpi.com/journal/biomolecules
Biomolecules 2022, 12, 472 2 of 25
Figure 1. The general reaction of GOx.
The high specificity, high turnover and high stability of GOx make it an ideal enzyme
for biosensor applications [
5
], some of which will be described below. Although its rate
constant is still several orders of magnitude below the diffusion limit, GOx has a much
higher
k
cat
/K
M
(on the order of 10
6
M
−1
· s
−1
) compared with most other oxidoreductases,
prompting at least one researcher to call it “the Ferrari of the oxidases” [6].
GOx is a member of the glucose-methanol-choline oxidoreductase (GMC oxidoreduc-
tase) superfamily. The members of this family are all FAD-dependent oxidoreductases
that share a common fold [
7
,
8
]. They consist of two functional domains, an N-terminal
FAD-binding domain, which contains a strictly conserved
βαβ
mononucleotide-binding
motif and a more variable substrate binding-domain. As the name suggests, the mem-
bers of this family oxidize a variety of substrates containing hydroxyl functional groups,
including mono and di-saccharides, alcohols, cholesterol and choline. GOx is perhaps
the most thoroughly characterized of these, and its mechanism will be described more
thoroughly below.
2. Glucose Oxidase Structure
Glucose oxidase has widespread applications in industry and biotechnology. Due to
this, a deep understanding of its structure and function are warranted. Perhaps surprisingly
for such a well-studied enzyme, there are only seven crystal structures of glucose oxidase
available, six from A. niger and one from P. amagasakiense [
9
–
12
]. The A. niger GOx structures
range in resolution from 2.4–1.2 Å (Table 1); five of them are in hexagonal space groups
(P3
1
21 or P3
2
21), and one is in an orthorhombic one (C222
1
). Although the active form in
solution is a dimer, the asymmetric unit of each structure contains only monomer with the
second monomer occupying a symmetry-related site.
These structures are all very similar, with a mean RMSD of 0.31
±
0.06 Å over an
RMSD range of 0.15–0.40 Å. The P. amagasakiense structure is unique in having a complete
dimer in the asymmetric unit of space group P2
1
2
1
2
1
. This is very similar to the A. niger
GOx structures, with an average RMSD of 0.68
±
0.04 Å over a range of 0.64–0.77 Å. As
it has been the most thoroughly studied, A. niger GOx will be described below; however,
because of the high structural similarity, much of the description of the overall structure
will also apply for P. amagasakiense GOx.
2.1. Overall Structure
The A. niger GOx monomer forms a rough, rounded rectangular prism shape, mea-
suring roughly 60
×
52
×
37 Å. The monomer is comprised of a single folding domain
with a rather complicated topology. When viewed in a manner that emphasizes the sec-
ondary structure elements while minimizing the contribution of the unstructured loops
(as in
Figure 2a,b
), two subdomains can be seen, each one centered around a five-stranded
β
-sheet. These two subdomains can be associated with FAD-binding on the one hand and
substrate binding on the other.
Biomolecules 2022, 12, 472 3 of 25
Table 1. Tertiary structures of glucose oxidase determined by X-ray crystallography.
PDB ID Resolution (Å) Source Reference Remarks
1GAL 2.3 A. niger [9]
The earliest and lowest resolution
structure.
1CF3 1.9 A. niger [10]
1GPE 1.8 P. amagasakiense [10]
The only structure with a complete
dimer in the asymmetric unit
3QVP 1.2 A. niger [11]
The highest resolution structure
available.
3QVR 1.3 A. niger [11]
5NIT 1.9 A. niger [12]
A2 Mutant engineered for higher
stability and turnover: T30V, I94V,
A162T, R537K, M566V
5NIW 1.8 A. niger [12]
A9 mutant: T30V, R37K, I94V, V106I,
A162T, M566V.
Figure 2.
The overall structure of A. niger glucose oxidase. (
a
) The overall structure of the GOx
monomer. The two central
β
-sheets are colored orange (parallel
β
-sheet A) and light brown (anti-
parallel
β
-sheet C) and labeled. The two loops that participate in the dimer interface are colored
magenta. (
b
) The GOx dimer. The two monomers are colored green and cyan, the FAD co-factor
is shown as van der Waals spheres, the carbohydrate chains attached to Asn-89 that participate in
the dimer interface are shown as sticks, and the loops that participate in the dimer interface are
colored magenta. The more important of the two loops is labeled. (
c
) A closer view of parallel
β
-sheet
A, which lies at the topological center of the enzyme. The
β
-sheet, together with the
α
-helix that
forms part of the
βαβ
motif, is colored by residue from the N-terminus to the C-terminus based
on sequence position to illustrate that all regions of the sequence take part in its formation. The
α
-helices and anti-parallel
β
-sheet that flank
β
-sheet A are shown in light blue. (
d
) A closer view of
the anti-parallel
β
-sheet C, which forms a Greek-key motif. The residues surrounding
β
-sheet C form
the second sub-domain and are colored by position from the N-terminal end to the C-terminal. The
cytosolic-facing
α
-helices following
β
-strands C2 and C4 lie behind the central
β
-sheet in this view.
All panels show the PDB structure 1CF3 [10].
Biomolecules 2022, 12, 472 4 of 25
The first, FAD-binding domain is centered on a five-stranded parallel
β
-sheet (
β
-sheet
A in Figure 2c), which is flanked by a smaller, three-stranded anti-parallel
β
-sheet on one
side and three
α
-helices on the other. It contains the FAD-binding site and is similar to the
FAD-binding motifs found in other FAD-binding, GMC oxidoreductase proteins, several of
which are listed in Table 2. The most highly conserved part of this motif is the
βαβ
motif
comprising strands A3, A4 and the
α
-helix between them; a
ψ
-BLAST search returns only
local alignments to this region on the second iteration.
Table 2.
Proteins from the PDB similar in structure to A. niger glucose oxidase. Structures were
identified by a BLAST search against the PDB using the A. niger sequence. Listed structures covered
at least 75% of the query sequence and represent unique structures. All these proteins are members
of the GMC oxidoreductase family.
PDB ID RMSD Description Identity (%) Reference
6XUT 1.2704
Trametes cinnabarina Oligosaccharide
dehydrogenase
35.89 [13]
4YNT 1.2841 Aspergillus flavus FAD glucose dehydrogenase 35.69 [14]
6ZH7 1.4654
Chlorella variabilis Fatty acid
Photodecarboxylase
27.99 [15]
5OC1 1.5672 Pleurotus Eryngii aryl-alcohol oxidase 29.67 [16]
6ZE2 1.5827
Chaetomium thermophilum FAD-dependent
oxidoreductase
31.24 [17]
4H7U 1.6328 Agaricus meleagris pyranose dehydrogenase 28.60 [18]
5HSA 1.6562 Pichia pastoris Alcohol Oxidase 23.56 [19]
6H3G 1.6725 Phanerodontia chrysosporium Alcohol oxidase 24.81 [20]
3NNE 1.7055 Arthrobacter globiformis choline oxidase 26.31 [21]
4HA6 1.7182 Mesorhizobium loti pyridoxine 4-oxidase 26.85 [22]
6F97 1.7508
Methylovorus sp. 5-(hydroxymethyl)furfural
oxidase
28.52 [23]
3Q9T 1.7866 Aspergillus oryzae formate oxidase 25.55 [24]
6O9N 2.1470
Myceliophthora thermophila aryl-alcohol oxidase
30.75 [25]
The parallel
β
-sheet at the center of this subdomain lies at the topological center of
the molecule. Starting with strand A3, the chain traces out the
βαβ
motif (that is, A3–
α
-
helix–A4) followed by a long loop forming part of the dimer interface and an extension
forming about one-quarter of the second subdomain. The trace then returns to form strand
A5. After A5, the chain forms a three-stranded anti-parallel
β
-sheet that forms part of the
FAD-binding domain, and this is followed by strand A2. After A2 comes the remainder of
the second subdomain followed by the last strand of the parallel
β
-sheet (A1). This is then
followed by a long C-terminal α-helix.
In this context, it can be seen that the residues responsible for forming the FAD-binding
pocket come from all parts of the polypeptide chain. The majority of the binding site is
formed by residues from the N-terminal part of the sequence. Residues from the central
region provide additional support, while most of the residues from the C-terminal region
form the edges of the flavin binding pocket and also contribute two of the three active
site residues.
The second subdomain is centered around a five-stranded anti-parallel
β
-sheet, which
is supported by six
α
-helices (Figure 2d). The first of the five
β
-strands, C1, is much shorter
than the others and arises from the extension following strand A4 of the FAD-binding
domain. The remaining four strands are arranged in a Greek-key motif: strands C5 and C2
are connected through a loop that passes over strands C3 and C4, which are joined together
by a hairpin loop.
Strands C2 and C3 are connected through a large excursion that forms three of the
supporting
α
-helices. Following C4 is a long loop that forms the second part of the dimer
interface followed by the last to helices of the subdomain. The six
α
-helices lie on the
Biomolecules 2022, 12, 472 5 of 25
cytosolic side of the
β
-sheet, while the
β
-sheet itself forms one side of a deep pocket that
has the active site and the flavin at the bottom.
Overall, this subdomain is similar to the binding sheet subdomain of cholesterol
oxidase [
26
]. Several of the residues thought to be important for binding
β
-D-glucose are
found here, including Trp-426, Phe-414 and Glu-412. The active site will be described more
closely in Section 3 where the catalytic mechanism is described.
2.2. Dimer Interface
The A. niger GOx dimer measures approximately 60
×
52
×
77 Å. The FAD groups
are located near the dimer interface but are more than 22 Å apart, making it unlikely that
they communicate with one another through allostery (Figure 2b). The dimer interface is
predominantly formed by residues from the long loop following strand A4 of the parallel
β
-sheet of the FAD-binding subdomain (residues 75–98) and from the loop following strand
C4 of the anti-parallel
β
-sheet of the substrate-binding subdomain (residues 432–455). The
loop containing residues 75–98 covers part of the FAD-binding pocket.
It was previously thought that the apo form of GOx was a monomer [
27
] and that FAD
binding was coupled to dimer formation [
28
,
29
], and the position of this loop appeared to
support those conclusions. More recent work has shown, however, that after the dissocia-
tion of FAD, the enzyme is not in a monomeric state but appears to form aggregates [
30
]
and that the dimer does not dissociate during thermal denaturation [
31
]. GOx does ap-
pear to dissociate into monomers at pH 5 and below in the presence of sodium
n
-dodecyl
sulfate [32].
In addition to forming part of the FAD-binding site, residues 75–98 also from part of
the wall of the active site cavity of the second monomer in the dimer: the complete dimer
is needed to properly assemble a complete active site cavity, which may account at least in
part for the dimeric form of the holoenzyme [27,33].
2.3. Active Site
In the complete dimer, the active site lies at the bottom of a deep, cone-shaped cavity
whose apex is centered on the N5 atom of the middle flavin ring, the one that accepts the
hydride during reduction. Only three amino-acid side-chains are near this center, His-516,
His-559, and Glu-412 (Figure 3). His-559 is fixed by a strong hydrogen-bond to Glu-412 and
Glu-412 is largely fixed by the side-chains of the surrounding residues (Ala-349, Phe-351,
Phe-414, Trp-426 and Leu-428). The side-chain of His-516 is less constrained and is more
conformationally flexible [12].
As all these structures were produced from crystals grown at pH 5.1–6.9, it seems
likely that both histidine residues are protonated on both N atoms. All but one of the
GOx crystal structures also have a water molecule at the center of the active site, forming
hydrogen-bonds to both His-516 and His-559 and lying 3.0 Å away from the N1 ring of
FAD (Figure 3a). The location of this water molecule has been taken to represent the
likely position of the O1 hydroxyl group of
β
-D-glucose [
10
]. Kinetic, structural and
thermodynamic data have allowed likely roles to be assigned to these residues.
Despite numerous attempts, no structure of GOx with a bound substrate has been
determined [
10
,
12
]. Wohlfahrt et al. [
10
] used manual docking and molecular dynamics
simulation to characterize a likely enzyme–substrate complex using oxidized A. niger
GOx and
β
-D-glucose. Their modeling suggested that the residues most important for the
enzyme–substrate complex included the catalytic His-516 and His-559 along with Tyr-68,
Thr-110, Phe-414, Trp-426, Arg-512 and Asn-514. His-516 and His-559 formed hydrogen-
bonds directly with the O1 hydroxyl group, which is expected to lose a proton in the
catalytic step, while Asn-514, Arg-512, Tyr-68 and O4 of the FAD all form hydrogen-bonds
to the remaining glucose hydroxyl groups, thereby, anchoring the substrate in position.
Biomolecules 2022, 12, 472 6 of 25
Figure 3.
The active site of A. niger GOx. (
a
) The active site of the oxidized form of the enzyme in the
absence of any substrate. The residues potentially important for catalytic activity are colored magenta;
those which are thought to be involved in binding
β
-D-glucose are colored cyan. The water molecule
near the center is thought to indicate the approximate location of the
β
-D-glucose O1 hydroxyl in the
substrate-bound conformation. PDB structure 1CF3 [
10
] is shown. (
b
) The active site of an engineered
GOx mutant showing an O
2
molecule bound to the active site (PDB structure 5NIT from Petrovi ´c
et al. [12]).
3. Catalytic Mechanism
The catalytic mechanism of GOx has been studied for many years, primarily using
kinetic methods [
34
]. These, together with the enzyme structure, provide the identities
of the active-site residues and allow a description of the mechanism of the reaction to be
proposed. Many many different substrates and electron-acceptors have been employed
(see Table 3); however, we will concentrate on the “natural” reaction using β-D-glucose as
the substrate and O
2
as the electron acceptor.
Table 3. Some glucose oxidase substrates.
Substrate
GOx Activity (%)
1
Reference
β-D-glucose 100 [35–38]
2-deoxy-D-glucose 25–30 [36–38]
4-O-methyl-D-glucose 15 [37]
6-deoxy-D-glucose 10 [37]
4-deoxy-D-glucose 2 [37]
2-deoxy-6-fluoro-D-glucose 1.85 [38]
3,6-methyl-D-glucose 1.85 [38]
4,6-dimethyl-D-glucose 1.22 [38]
3-deoxy-D-glucose 1 [37]
6-O-methyl-D-glucose 1 [37]
α-D-glucose 0.64 [37,38]
mannose 0.2; 1 [36–38]
altrose 0.16 [37,38]
galactose 0.08 [36–38]
xylose 0.03 [36–38]
idose 0.02 [37,38]
1
Percent relative to β-D-glucose.
GOx operates using a Ping-Pong Bi Bi mechanism in which
β
-D-glucose oxidation and
O
2
reduction occur in two different steps [
35
,
39
–
41
] (Figure 4). This allows the oxidation
and reduction half reactions to be analyzed independently using steady state kinetics.
pH profiling showed that the p
K
a
of the amino-acid side-chains in the active site of the
enzyme–substrate complex is between 6.9–7.8 [35,42].
Biomolecules 2022, 12, 472 7 of 25
Several lines of evidence ascribed this p
K
a
value to His-516 [
40
,
41
]. The most likely
mechanism involves a base-catalyzed hydride transfer from the glucose C1 to the flavin N5.
Glucose binding is thought to displace the water molecule observed in the active site of the
GOx crystal structures, which abstracts a proton from His-516 as it leaves (this abstraction
appears to be necessary, according to modeling studies, because His-516 cannot adopt a
proper catalytic position when doubly-protonated) [10].
Hydride transfer occurs in a concerted step in which a proton is removed from the
glucose O1 hydroxyl group by a basic group on the enzyme while a hydride is transferred
from the glucose C1 to the flavin N5 [
43
]. The kinetic rate constants associated with this step
show a large H/D kinetic isotope effect, suggesting that this transfer is the rate-limiting step
of the reductive half-reaction [
44
]. The basic group on the enzyme is likely to be His-516: a
His-516-Ala mutant had practically no catalytic activity [41].
Considerations of resonance suggest that hydride transfer should create a negative
charge around the FAD N1 atom, which have been confirmed by NMR studies of GOx at
pH 5.6 in the absence of oxygen [
45
]. Following the reduction half-reaction, the enzyme is
left in a reduced form with a bound glucono-
δ
-lactone. The product is then displaced by
water or possibly O
2
and the oxidation half-reaction follows.
Figure 4.
A scheme illustrating the overall glucose oxidase reaction. A indicates glucose, P indicates
glucono-
δ
-lactone, B represents O
2
, Q represents H
2
O
2
, E represents the oxidized form of GOx bound
to FAD, and F stands for the reduced form of GOx bound to FADH
2
. The individual components of
the reaction are illustrated below the scheme; from left to right they are reduced GOx (E), the most
likely mechanism of the GOx reduction half-reaction (EA
↔
FP), the GOx–O
2
complex (FB) and the
GOx–H
2
O
2
complex (EQ).
The kinetic and thermodynamic parameters for the oxygen-binding step suggest that
O
2
easily and rapidly diffuses into the enzyme [
35
,
44
,
46
]. Indeed, structural studies on
mutant GOx enzymes engineered to be more stable and have greater catalytic efficiency
found O
2
bound in the active site [
12
] (Figure 3). The oxygen bound in a small pocket
formed by shifting His-516 up towards the active-site opening and displacing the active-site
water molecule so that it lay outside hydrogen-bonding range of FAD (its closest approach
was 3.7 Å from the N5 atom of FAD).
In its ground state, molecular oxygen is a paramagnetic triplet, making its insertion
into diamagnetic organic molecules a spin-forbidden process [
47
]. In the active site, one
of the O
2
atoms is 2.7 Å away from the N1 flavin ring while the second is 3.0 Å away
from the His-516 ring. This position led Petrovi´c et al. [
12
] to suggest that the oxidative
half-reaction might rely on orbital coupling between the oxygen and the
π
electrons in the
His-516 side-chain to overcome this limitation.
Generally, O
2
oxidizes organic substrates by transferring electrons one at a time,
forming free-radical intermediates, and most analyses of of the oxidative half-reaction
assumed a step-wise electron transfer [
34
,
48
]. The rate-limiting step appears to be the
transfer of the first electron from flavin to O
2
to produce the flavin semiquinone radical
Biomolecules 2022, 12, 472 8 of 25
and the superoxide anion [
41
]. Kinetic isotope effect experiments show that no proton
transfer takes place during this step [
35
,
44
]. The second electron transfer step, from the
flavin semiquinone radical to the superoxide anion, happens quickly, with a second-order
rate constant of 10
9
M
−1
· s
−1
[49].
Glycosylation
Consistent with its role as an extracellular protein, GOx is glycosylated, with carbohy-
drates, mostly mannose-like sugars, comprising between 10 and 16% of its final molecular
weight [
27
,
36
,
50
]. Both
N
-linked and
O
-linked sugar chains are present. The removal of 95%
of the GOx carbohydrate content had a noticeable effect on the kinetics of glucose oxidation,
its stability at low pH and the number of available isoelectric forms but did not affect its
thermal stability or the optimal pH and temperature of the catalytic reaction [50,51].
More particularly, a study using H/D isotope substitution found that reduced glyco-
sylation appeared to lower the enthalpy of activation and, therefore, increase the activity of
the enzyme [
52
]. Deglycosylation typically removes all of the
O
-linked sugars at their link-
age to the amino-acid side chain but leaves the first
N
-acetylglucosamine (Nag), potentially
allowing those sites to be identified in the crystal structure [53].
Potential glycosylation sites can be identified by the consensus sequence N-X-T/S,
where X is any amino-acid residue except proline [
54
]. Eight such sites are present in
A. niger GOx (residues Asn-43, 89, 161, 168, 258, 355, 388 and 473); seven of these sites
have been confirmed in at least one of the known A. niger GOx structures. Asn-43 does not
appear to be glycosylated in any of these structures, and peptide sequencing work also
suggested that no carbohydrate is attached here [54].
4. Natural Sources of Glucose Oxidase
GOx is primarily produced by fungi and insects [
55
]. There were early reports of
GOx activity in extracts from red algae [
56
], citrus fruits [
57
], mammalian tissues [
58
] and
bacteria [
59
–
62
]. A careful examination of these studies suggests that the activities from
plants and mammals are not due to GOx and that several of the activities reported for
bacteria appear to be due to the NAD-dependent glucose dehydrogenase. Two early studies
on extracts from Burkholderia pseudomallei [
59
] and Acetobacter suboxydans [
60
] may be due
to GOx; however, there were no later studies confirming this.
All these studies made use of what would now be considered rather crude cell-free
preparations, which would make it difficult to distinguish between the glucose oxidase
activity of GOx and that of pyranose oxidase. To date, there appears to have been no
confirmed glucose 1-oxidase identified in any bacterial strain, and a recent phylogenomic
study of the GMC oxidases found that pyranose oxidase is only one of the GMC proteins
present in both fungi and bacteria [8].
To unambiguously identify those organisms that are presently known or suspected of
harboring GOx, the NCBI Protein database, UniProtKB and the Brenda Enzyme Database
were searched for entries classified as having E.C. number 1.1.3.4, and some of the recent
literature was surveyed. After discarding duplicate entries and removing those that were
mis-annotated (for example, a Streptomyces coelicolor alditol oxidase, E.C. 1.1.3.41, was
incorrectly classified as GOx in one of these databases), a total of 50 unique species were
identified; 45 species were from fungi, and five were from insects.
The insect species include Apis melifera (the honey bee) [
63
], Helicoverpa armigera
(Cotton bollworm) [
64
], Heliothis viriplaca (marbled clover, a moth), Mythimna separata
(Oriental armyworm) and Spodoptera exigua (Beet armyworm) [
65
]. No mammalian, plant,
or bacterial species were found. The results are listed in Table 4.
Only 27 of the 50 species had sequences available in one of these three databases.
Table 4 shows the similarities of these sequences to A. niger GOx according to BLAST. All
sequences are roughly the same length (generally around 600 residues) and the alignment
covers at least 89% of the A. niger sequence. They can be divided into three main groups:
The two Aspergillus species share 85% identity, next are the Penicillium and Talaromyces
Biomolecules 2022, 12, 472 9 of 25
species with 63–67% identity to A. niger (with one exception), and last are all other species,
which have only 27–35% identity.
Table 4. GOx-producing organisms in at least one major database or at least one reference.
Organism Name Accession or Reference
1
Identity
2
Fungi
Alternaria alternata [66] –
Aspergillus carbonarius U V9SH09 85.79
Aspergillus niger U P13006 [3,55] 100.00
Aureobasidium pullulans U A0A221SAG9 34.09
Aureobasidium sp. U A0A1V0E5A9 31.69
Ceratocystis fimbriata U A0A0F8CXS8 28.28
Cladosporium neopsychrotolerans U A0A5Q2UVJ5, U A0A5Q2USS5 31.80, 32.56
Cystobasidium laryngis [67] –
Dioszegia sp. [67] –
Flavodon flavus [68] –
Fusarium oxysporum [69] –
Goffeauzyma gastrica [67] –
Goffeauzyma gilvescens [67] –
Leucosporidium fragarium [67] –
Leucosporidium creatinivorum [67] –
Malassezia restricta U A0A3G2S2X3, U A0A3G2SBT7 29.44, 32.64
Mucor circinelloides [70] –
Penicillium adametzii U A2I7K9 [71] 64.13
Penicillium amagasakiense U P81156 [72] 65.74
Penicillium canescens [73] –
Penicillium chrysogenum U K9L4P7 [74] 62.91
Penicillium expansum [75] –
Penicillium janthinellum [76,77] –
Penicillium viticola U A0A0Y0IDS5 63.00
Penicillium sp. U A0A7L7T1A0 65.51
Penicillium sp. RFL-2021a N KAF7733001 34.33
Phanerochaete chrysosporium [78] –
Pleurotus ostreatus [79] –
Pycnoporus cinnabarinus [80] –
Rasamsonia emersonii U A0A0F4YPS7 34.78
Rhizopus stolonifer [81] –
Schizophyllum commune U D8QJE7 [82] 34.31
Serendiptia indica N CAG7851011 33.56
Sporidiobolus salmonicolor [67] –
Talaromyces flavus U Q92452 [83] 63.97
Talaromyces funiculosus [84] –
Talaromyces pinophilus [85] –
Talaromyces purpureogenus [86] –
Talaromyces stipitatus U B8MDS4 [87] 63.85
Talaromyces variabilis U Q70FC9 [88] 66.61
Thanatephorus cucumeris U M5BNG8 [89] 30.98
Wickerhamomyces anomalus [67] –
Xylona heveae [90] –
Yarrowia sp. B02 N KAG5360348 26.87
Insects
Apis melifera U Q9U8X6 [63] 26.66
Helicoverpa armigera U B2MW81 [64] 29.40
Heliothis viriplaca U A0A142I707 29.27
Mythimna separata U A0A218N0E8 28.07
Spodoptera exigua U D9ZFI1 [65] 28.74
1
Key: N—NCBI Protein, U—UniprotKB followed by accession number.
2
Relative to the A. niger GOx sequence
(U P13006). – indicates no sequence available.
Biomolecules 2022, 12, 472 10 of 25
As shown in Table 2 above, a sequence similarity of this is generally the same iden-
tity shared between A. niger GOx and other, functionally non-equivalent GMC oxidases.
This means that it will generally not be possible to identify as GOx uncharacterized se-
quences from more distantly related organisms even within the same kingdom using
BLAST searches alone.
A phylogenetic tree based on an alignment of all these sequences generally maintains
the grouping seen in Table 4, putting the insect GOxes together in one branch and separating
the fungal GOxes into two separate branches (Figure 5). One branch clusters the Aspergillus,
Talaromyces, and most of the Penicillium species, while the other branch contains all the
remaining species. Curiously, the Yarrowia sp., Thanatephorus cucumeris, and Ceratocystis
fimbriata species are closer to the insect species than the fungal species.
Figure 5.
A phylogenetic tree based on an alignment of all sequences given in Table 4. The separation
between the insect and fungal GOxes can clearly be seen. Within the fungal group, the Aspergillus,
Talaromyces, and most of the Penicillium species cluster in their own clade, while the other species
cluster in a different one. This tree was generated by PhyML [
91
] based on an alignment prepared by
Clustal Ω [92].
A multiple-sequence alignment of three fungi (A. niger, P. amagasakiense and M. restricta)
and two insect (A. melifera and S. exigua) GOx sequences highlights the conserved areas
(Figure 6). The overall sequence identity relative to the consensus sequence reached
approximately 37%; however, there is a clear division between the fungal (43%) and insect
(28%) sequence groups (see also Figure 5).
The largest differences occur at the N-terminus where signaling pre-sequences may be
located. Overall, the alignment shows several large blocks of relatively conserved residues
separated by regions of low identity. As described above (Section 2.1), the residues forming
the FAD binding site are distributed throughout the N-terminal, central and C-terminal
part of the molecule, and the residues involved in FAD binding all occur in one of these
relatively conserved blocks.
Biomolecules 2022, 12, 472 11 of 25
Figure 6.
Amino-acid sequence alignment of GOx from A. niger (UniProt ID P13006), P. amagasakiense
(P81156), M. restricta (A0A3G2SBT7), A. melifera (Q9U8X6) and S. exigua (D9ZFI1) performed by
T-COFFEE [
93
]. The overall amino-acid sequence identity is 36%. The alignment is over the full-
length sequences found in the UniProt database, which includes likely N-terminal presequences. The
secondary structure elements shown are based on those derived from the 1.9 Å wild-type A. niger
GOx structure (PDB ID 1CF3). Red and pink cylinders represent
α
-helices and 3
10
helices, respectively;
blue and purple arrows indicate parallel and anti-parallel
β
-sheets, respectively. Individual
β
-strands
within each
β
-sheet (A–E) are numbered according to their position in the structure rather than the
sequence (see Figure 2). The A. niger GOx presequence cleavage site (R22) is marked with a vertical
line. The positions of the crucial catalytic residues His-516, His-559 and Glu-412 (corresponding
respectively to residues 538, 581 and 434 in the full-length sequence) are shown in larger and bold-face
type. Residues participating in FAD binding are in boxed. Cysteines forming a disulfide bond in the
A. niger and P. amagasakiense structures are shown in larger, bold-face, pink type. The glycosylation
sites observed in the A. niger and P. amagasakiense structures (Asn-89, 161, 355 and 388, corresponding
respectively to residues 111, 183, 377 and 410 in the full-length sequence) are shown in larger, deep
blue type. The positions of potentially glycosylated asparagines in the M. restricta, A. melifera and S.
exigua GOx sequences are shown in larger, light blue type.
Outside the N-terminus, the regions of lowest identity correspond to residues 83–93,
145–189, 340–408 and 490–509 in the A. niger GOx structure. Residues 145–189 form part of
the GOx outer surface distant from the active site; however, the remaining three appear to
be structurally important. Residues 83–93 form an important part of the dimer interface and
the insect sequences have a five-residue insertion in this area. Residues 490–509 occur near
the dimer interface and also feature a five-residue insertion in the insect and M. restricta
sequences. This region is also in contact with the loop containing residues 83–93, so these
insertions may be correlated. Finally, residues 340–408 include the loop between
β
-strands
C5 and C2 and the extension following C2.
The M. restricta and insect sequences have a poorly-conserved insertion between
β
-
strands C5 and C2 and are lacking the sequences that comprise the either just the first two
Biomolecules 2022, 12, 472 12 of 25
(insect sequences) or all three (M. restricta) of the
α
-helices between C2 and C3. Generally,
the catalytic residues His-516, His-559 and Glu-412 are conserved, although only His-516 is
absolutely conserved in all five sequences.
In the insect sequences, His-559 is replaced with an asparagine (a substitution that is
not uncommon in the GMC family in this position generally), while Glu-412 is replaced
with an aspartate in A. melifera. These substitutions suggest that there are subtle differences
in the conformation of the active sites in these proteins; they also provide additional support
to the designation of His-516 as the primary catalytic residue as described in Section 3.
The sequence alignment also shows that the positions of Cys-164 and Cys-206, which
form disulfide bonds in the A. niger and P. amagasakiense structures, are conserved only
in these two sequences. The cysteines in M. restricta, A. melifera and S. exigua appear in
different locations in the sequence, suggesting either differences in disulfide bond formation
or no disulfide bonds occur in these proteins.
Finally, it may be worth noting that the A. niger and P. amagasakiense glycosylation sites
identified structurally (Asn-89, 161, 355 and 388 in the A. niger 1CF3 structure corresponding
to Asn-111, 183, 377 and 410 in Figure 6) do not appear to be generally conserved. Only an
equivalent to Asn-89 appeared in the same general location in M. restricta and A. melifera
GOx. The positions of the remaining potential glycosylation sites are either shifted or
missing in the other three GOx sequences, suggesting alternative patterns of glycosylation.
In particular, the alignment in Figure 6 shows that M. restricta and A. melifera do have
potential N-X-(T/S) glycosylation motifs in the neighborhood of residues Asn-355 and
Asn-388, suggesting that at least one glycosylation site might be expected in this location.
5. Industrial and Medical Applications of GOx
GOx is used in many branches of industry because of its ability to oxidize glucose and
produce hydrogen peroxide. Its rapid turnover and high stability finds it many applications
in the food, pharmaceutical, medical, textile and power industries. For many of these
applications GOx is used in a biosensor or nanosensor [
94
–
99
], in nanoparticles [
100
–
102
]
or in nanosheets [103].
In many modern applications, GOx is often used in combination with other enzymes,
for example, tyrosinase in the analysis and discrimination of musts and wines [
104
],
α
-
amylases and xylanases for improving the quality of dough and bread [
105
], peroxidase
for accurately measuring the level of glucose in blood and saliva [
106
] and tears [
94
],
the autophagy inhibitor chloroquine in cancer intervention therapy [
101
] and insulin for
regulating blood glucose levels in diabetes [
107
,
108
]. Finally, it has been combined with
the anti-cancer drug tirapazamine and human serum albumin to create a nanoreactor
capable of increasing the levels of hypoxia and reactive oxygen species and inhibiting
tumor growth [109].
The applicability of GOx primarily depends on its quantity, thermal stability and
activity. Many studies focused on identifying which fungal strains are better for biosensor
development, which are better for clinical studies and which are better for biochemical
diagnostic tests [
55
]. Optimal GOx utilization also requires consideration of the type of
matrix on which GOx is bound and type of media and conditions under which it is used.
Consequently, in addition to identifying the best GOx producers and the ideal conditions
for its stability and activity, the development of different binding materials, environmental
conditions and detection systems are also very important for expanding the range of GOx’s
industrial applications.
5.1. Industrial Production of GOx
Industrially, the most important producers of GOx are species belonging to the genera
Aspergillus and Penicillium [
55
,
110
,
111
], especially A. niger and P. amagasakiense. These
genera are important because of their metabolic versatility and because they are “generally
regarded as safe” by regulatory agencies [112].
Biomolecules 2022, 12, 472 13 of 25
Overall, P. amagasakiense GOx is catalytically more effective than A. niger GOx, with
a 6
×
lower Michaelis constant (
K
m
) for
β
-D-glucose and 10
×
higher catalytic efficiency
(
k
cat
/K
m
) [
113
]; however, it is also less stable and has a lower antimicrobial activity [
114
].
This prompted at least one group of researchers to create a hybrid GOx with improved
stability and catalytic efficiency by genetically combining elements from both A. niger and
P. amagasakiense GOxes [
115
]. Other researchers have attempted to improve the antioxidant
capabilities [116] and thermal stability [117] of A. niger GOx.
Fungal GOx is produced by solid state fermentation (SSF) and submerged fermentation
(SmF) [
118
,
119
]. Submerged fermentation is the more useful method because environmental
factors can be controlled more easily. A study on the variability of GOx from Aspergillus
tubingensis CTM507 produced by SSF and SmF found that SmF produced GOx more
efficiently but that the GOx produced by SSF had a higher activity (170 U mL
−1
for SSF
versus 43.73 U mL
−1
for SmF) [
120
]. Both of these methods have limited capabilities, so
more efficient ways of producing GOx are being researched using genetic recombination
techniques, mutagenesis and immobilization methods [3,55].
Glucose and saccharose are the carbon sources most often used for the industrial
production of GOx [
3
,
121
]. GOx production can be induced from A. niger by glucose [
121
],
CaCO
3
[122], Mn
2+
, Co
2+
, thioglycolic acid, gluconic acid [123], EDTA (ethylene diamine
tetra-acetic acid) Zn
2+
and Fe
2+
[
116
]. Inhibitors of A. niger GOx include the Ag
+
, Hg
2+
,
Cu
2+
and Mg
2+
ions, CaCl
2
[
124
,
125
], hydrogen peroxide accumulation [
116
], arsenates,
p
-chloro-mercapto-benzoate, phenyl mercuric acetate [
125
], hydroxylamine, hydrazine,
phenyl hydrazine, dimedone, NaHSO
4
[
126
], guanidinium chloride, urea and SDS (sodium
dodecyl sulfate) [116].
The low fermentation capacity, complicated purification process and effectiveness
of GOx limit the applicability of GOx from natural sources. Attempts to improve GOx
production through the mutagenesis and screening of production strains has enhanced
GOx production in A. niger by about 77% [
127
]. Genetic recombination has also been
used to express GOx heterologously. The most effective cloning techniques and highest
overexpression levels were reported for Saccharomyces cerevisiae and Escherichia coli and
other Aspergillus and Penicillium species have also been used as hosts [128–130].
Other candidates for the industrial production of GOx are H. polymorpha and P. pas-
toris [
131
,
132
]. Along with S. cerevisiae, these yeasts have the advantages of rapid growth
and extracellular protein production; the highest level of A. niger GOx production from
S. cerevisiae is currently 9 g L
−1
. Unfortunately, GOx produced by S. cerevisiae and H.
polymorpha tend to produce over-glycosylated forms of GOx with reduced activities [
133
].
However, P. pastoris has been successfully used for the production of a GOx from A. niger
and P. variable P16, which was not over-glycosylated [134,135].
5.2. Use of Glucose Oxidase in the Food Industry
GOx has several important applications in the food industry [
55
], including the baking
industry, the production of drinks, the production of gluconic acid and food preservation.
In the baking industry, GOx is used as an oxidant to improve the quality of bakery prod-
ucts [
136
–
138
]. The hydrogen peroxide produced by GOx makes the dough more elastic
and viscous [
139
] and lipase and GOx can increase the quality and durability of bread [
140
].
Generally, bread quality depends on wheat quality and dough rheology depends on
the quantity of enzymes added [
141
,
142
]. GOx causes the formation of protein fibers in
dough. Basal additives, such as GOx, ascorbic acid and
α
-amylases decrease bread fragility
and enhance chewability, adhesion, elasticity and cohesion [
143
]. The addition of fungal
xylanase has a positive effect on flours and crumb firmness [105,144].
GOx also finds use in the reduction of the alcohol content of wine. Warmer tem-
peratures during the growth season are likely to increase the level of glucose in wine
grapes. By decreasing the level of glucose, which would otherwise be transformed into
alcohol through anaerobic fermentation, GOx could decrease the quantity of alcohol in the
resulting wine [
145
]. The hydrogen peroxide produced by GOx has a bactericidal effect on
Biomolecules 2022, 12, 472 14 of 25
the corrosive acidic and dairy microbes produced during the fermentation process. The
hydrogen peroxide produced can be removed by the enzyme catalase, which transforms
H
2
O
2
into oxygen and water. GOx in combination with catalase decreases the quantity of
alcohol better than GOx alone by converting glucose into gluconic acid [146].
For measuring the quantity of glucose in liquids, GOx has also been incorporated into
analytical devices known as biosensors. These devices combine a biological component,
frequently some kind of enzyme or antibody, with a physical or electrical transducer and
an electrical component to provide a measure of the analyte of interest [
147
]. For example,
Garcia-Hernandez et al. [
104
] constructed a bioelectronic “tongue” consisting of enzymes
(tyrosinase and glucose oxidase) and polypyrrole or polypyrrole/Au nanoparticles to
measure the alcoholic degree in musts and wine.
The information from this sensor can help to predict the characteristics of the finished
wine at the beginning of the vinification process. Lopes et al. [
148
] created a biosensor that
consisted of GOx and immobilized horseradish peroxidase to determine the quantity of
glucose in beverages, such as orange juice and energy drinks. Finally, Mason, Longo and
Scampicchio [
149
] created an electrochemical biosensor that consisted of GOx immobilized
on a nylon nano-fiber membrane to determine the level of glucose in brewed beer.
Such devices also allow the level of glucose to be measured in “sugar-free” foods
intended for diabetics. For example, Kalaivani et al. [
98
] constructed a reliable sensor for
detecting nanomolar amounts of glucose in food products.
Its production of gluconic acid also finds some uses for GOx in the food industry.
Gluconic acid is used as an acid regulator, color stabilizer, antioxidant and chelating agent
in food and drinks [
150
]. Gluconic acid is also applied in the dairy industry in cheese curd
production, improving thermal stability of milk and cleaning of aluminum tins. The most
popular application of gluconic acid in the food industry is its use as an acid regulator and
antioxidant [151].
By removing oxygen and glucose from food, GOx can be used to lengthen shelf-life
through two reasons. First, during food preservation, an unwanted reaction may occur
between the nuceophilic functional groups on amino acids with the active carbonyls on
sugars to cause non-enzymatic browning through the Maillard reaction. By removing
unwanted sugars from the food to be preserved, this browning is prevented.
For example, using GOx to remove glucose residues from dried eggs improves their
durability [
152
]. The hydrogen peroxide produced during this reaction also helps to destroy
all pathogen microbes potentially present in raw eggs. Extra hydrogen peroxide can then
be removed by the enzyme catalase [
153
]. The combination of GOx and catalase can also be
used to control the non-enzymatic browning of fruit and tomato paste.
The second reason is that the extra oxygen can support bacterial growth, and removing
the excess oxygen will inhibit the growth of aerobic bacteria. The removal of excessive
oxygen is essential for canned foods [
154
] and Karimi et al. [
155
] studied the removal of
dissolved oxygen in water through its reduction by glucose, catalyzed by glucose oxidase
and catalase. GOx can also be used to remove of oxygen from canned drinks (such as
beer and wine) to help them keep their color and taste [
156
,
157
]. The decomposition of
mayonnaise is connected with lipid peroxidase [
158
] and GOx and catalase can slow down
lipid peroxidation: oxygen removed during glucose reduction will not be available for
lipid metabolism.
GOx is also used directly as an antimicrobial agent in the food industry. It was
demonstrated to decrease the growth of many pathogenic bacteria, including Clostridium
perfringens, Campylobacter jejuni, Salmonella infantis, Staphylococcus aureus and Listeria mono-
cytogenes [
159
]. Vartiainen, Ratto and Paulussen [
160
] found that immobilized GOx inhibits
the growth of Escherichia coli and Bacillus subtilis and Malherbe et al. [
130
] demonstrated
that S. cerevisiae producing GOx from A. niger had antimicrobial activity against bacteria
that produce lactic acid and acetic acid.
Polyamide and ionomer films with immobilized GOx inhibited the growth of E. coli
CNCTC 6859, Pseudomonas fluorescens CNCTC 5793, Lactobacillus helveticus CH-1, Listeria
Biomolecules 2022, 12, 472 15 of 25
ivanovii CCM 5884 and Listeria innocua CCM 4030 on agar medium [
161
]. Yuan et al. [
162
] de-
veloped a photodynamic antimicrobial system consisting of glucose, GOx and horseradish
peroxidase to inactivate bacterial and fungal pathogens. Finally, Xu et al. [
163
] developed a
biosensor comprising of antibodies, GOx, gold nanoparticles, shell magnetic beads, poly-
dopamine and polymeric nanocomposites for the detection of pathogens in foods. This
biosensor was able to detect E. coli O157:H7 with a detection limit of 10
2
cfu/mL.
5.3. Glucose Oxidase Biosensors in Medicine Applications
5.3.1. Cancer
The ability of GOx to consume intracellular glucose and oxygen to produce hydrogen
peroxide and gluconic acid might allow it to be used in certain cancer therapy combinations.
By consuming glucose, GOx could reduce the available metabolic energy sources of cancer
cells, thereby, inhibiting their proliferation, and by consuming oxygen and producing
gluconic acid, it could increase the hypoxia and acidity of the tumor microenvironment.
In several of these proposed combination treatments, GOx is embedded in nanocom-
posites. These come in a variety of forms, including hollow mesoporous silica nanoparticles,
metal–organic frameworks, organic polymers, and magnetic nanoparticles are used for
the construction of GOx-based nanocomposites for multi-modal synergistic cancer ther-
apy [164].
In one proposed treatment [
101
], GOx chloroquine are attached to the surface and
loaded into the cavity of rattle-structured polydopamine core-hollow mesoporous silica
shell nanoparticles (PDA@hm). These nanoparticles are then used for a therapy combining
energy metabolism regulation (by GOx), autophagy inhibition (by chloroquine) and low-
temperature photothermal therapy (induced by the PDA nanocore). Photothermal therapy
uses light-absorbing materials to convert photoenergy into local hyperthermia to destroy
cancerous tissues [165,166].
In this treatment, GOx served to starve the tumor and directly suppressed the expres-
sion of the heat-shock proteins HSP70 and HSP90. In another proposed treatment [
99
],
GOx–MnO
2
nanosheets were developed to destroy cancer cells using a combination of
starvation and self-oxygenation by GOx and photothermal therapy by the MnO
2
nanosheet.
In these nanostructures, the GOx catalytic activity could be enhanced by the hyperthermia
triggered by near-infrared laser radiation.
These GOx–MnO
2
structures exhibited pH and glucose responsive performance, ac-
tivated by magnetic resonance and photoacoustic dual-modal imaging. Finally, GOx
formed one component of a recently-designed “nanoreactor” consisting of a tirapazamine
(TPZ)–human serum albumin–GOx mixture combined with a metal–polyphenol network
consisting of Fe
3+
ions and tannic acid [109].
This complex kills cancer cells by producing HO
•
, a process termed chemodynamic
therapy (CDT) and TPZ
•
radicals. The HO
•
radicals are produced from H
2
O
2
by Fe
2+
/Fe
3+
ions through the Fenton reaction [
167
], while TPZ develops into a toxic radical under
conditions of tumor hypoxia. The role of GOx in this assembly is to consume oxygen to
increase the tumor’s hypoxia level, to produce H
2
O
2
for conversion to HO
•
and to consume
glucose for starvation therapy.
5.3.2. Diabetes Treatment
The earliest suggestion for a glucose sensor involving GOx was in 1962 [
168
]. Today,
GOx is extensively used in the most common methods for measuring blood glucose lev-
els [
169
,
170
]. Here, we limit ourselves to some relatively recent developments. GOx activity
is highly oxygen dependent, which can lead to inaccuracies in amperometric
β
-D-glucose
determinations.
Fokkert et al. [
171
] investigated the performance of fluorescence sensor-based and
GOx-based glucose measurement during intensive exercise, when oxygen consumption is
expected to be higher, and normal daily activities. They found that both methods were less
Biomolecules 2022, 12, 472 16 of 25
accurate during exercise than during daily activities, and this finding persisted over the
whole range of glucose concentrations examined.
Gutierrez et al. [
172
] attempted to overcome this problem by engineering GOx variants
with lower oxygen dependency through random mutagenesis using error-prone PCR and
sequence saturation. They discovered which positions seemed to be vital for oxygen
sensitivity and for oxygen activity. One of their variants had a 37-fold reduced oxygen
dependency but maintained the same β-D-glucose specificity and thermal resistance.
Other studies have attempted to use GOx to measure effective blood glucose levels
non-invasively by monitoring other body fluids. Thus, Mohammadnejad et al. [
106
]
developed a more precise, accurate and rapid method to measure blood glucose using
GOx and peroxidase to oxidize a 4-[(hydroxy-3-methoxyphenyl)-azo]-benzenesulfonic acid
(GASA) substrate. The stability of GASA and its oxidized products along with its direct
and fast consumption by peroxidase, not only made it possible to determine blood glucose
concentration with high reproducibility but also allowed it to be used for salivary samples.
A low-cost non-invasive paper-based biosensor for glucose measurements from tears
has also recently been developed [
94
]. Finally, to eliminate some limitations and errors
in some of the more commonly used monitoring systems, J˛edrzak et al. [
97
] developed a
biosensor consisting of magnetite, lignin and polydopamine bound to GOx together with
ferrocene and a dedicated carbon paste electrode. The results showed that this biosensor has
a potential for application in the determination of glucose in various commercial products.
One of the more interesting developments in the use of GOx in glucose monitoring
is its potential to be coupled to an insulin delivery system, which could improve the
health and quality of life for many diabetics. Yang et al. [
173
] developed a glucose and
magnetic-responsive microvesicle delivery system, which can both regulate glucose levels
and generate nitric oxide.
The injectable microvesicles are loaded with GOx, which can reduce hyperglycemia
by consuming excess glucose. An applied magnetic field can then alter the permeability
of the microvesicle shell, allowing the H
2
O
2
produced by GOx to react with L-Arginine
within the vesicle to produce nitric oxide, which has been shown to be important in the
early stages of glucose-stimulated insulin secretion [174].
A glucose-responsive “closed-loop” insulin delivery system mimicking the function
of pancreatic cells has potential to improve quality of life and health in diabetics. Yu
et al. [
107
] created a glucose-responsive insulin delivery “closed-loop” system using a
painless microneedle-array patch containing glucose responsive vesicles that are loaded
with insulin and GOx. Under conditions of hyperglycemia, the GOx consumes glucose and
oxygen, creating hypoxic conditions, which then trigger the release of the insulin.
This device effectively regulated the blood glucose in a mouse model of chemically
induced type 1 diabetes. In a later refinement, the same group made an enhanced system
where insulin release contingent on both hypoxia and H
2
O
2
[
108
]. This system could
effectively regulate blood glucose in mice with chemically-induced type 1 diabetes for 10 h.
Several glucose-responsive nanoparticles are based on phenylboronic acid, which has
better stability compared with protein-comprised systems [
175
]. Chai et al. [
100
] created a
glucose-responsive insulin delivery system comprised of poly(acrylamido phenylboronic
acid)/sodium alginate nanoparticles loaded with GOx. The GOx-laded nanoparticles
showed greater glucose sensitivity and faster glucose-responsive insulin release than
nanoparticles loaded with insulin alone. These nanoparticles can also be easily prepared
and have good biocompatibility.
5.4. GOx in Wound Healing: From Bench to Bedside
As mentioned above, GOx has several important applications in the food and medical
industries. GOx’s ability to continuously produce hydrogen peroxide at low concentrations
has attracted a great attention in the area of wound management. One wound treatment
that makes use of the GOx-mediated release of hydrogen peroxide involves honey. Honey
Biomolecules 2022, 12, 472 17 of 25
has been used for the treatment of a wide range of injuries, including acute and chronic
wounds and burns [176].
A plethora studies have provided compelling evidence that hydrogen peroxide is the
major antibacterial compound found in diluted honey. GOx, likely together with some
polyphenols found in honey, is primarily responsible for the hydrogen peroxide generated
during nectar processing and honey ripening. GOx, together with other enzymes catalyzing
the metabolism of sugar, is produced by the hypopharyngeal glands of worker bees and
secreted into honey [177].
Therefore, GOx is a regular but quantitatively variable compound in natural honey [
178
].
The final amount of hydrogen peroxide in dilute honey is a result of the enzymatic activity
of both GOx and pollen-derived catalase enzymes. Interestingly, honeydew honeys, which
are rich in polyphenols, are able to generate a higher level of hydrogen peroxide than many
other honey types. It is believed that polyphenols, including flavonoids, might significantly
contribute to higher levels of hydrogen peroxide [179].
The efficacy of the topical application of honey in the treatment of various wounds has
been well-documented enough to allow it to be registered as a medical device in wound
care management. GOx itself has become an interesting therapeutic platform in wound
care [180–182].
There are several clinically tested GOx-based products available in the market for
wound application containing a GOx enzyme immobilized in a suitable carrier (e.g., a
hydrogel), which gradually releases a controlled amount of hydrogen peroxide
[183,184]
.
However, a GOx-embedded hydrogel does not perfectly replicate antimicrobial honey-
mimetic mechanisms because glucose is not added into the hydrogel.
Furthermore, the high concentration of H
2
O
2
in the hydrogel could damage the
normal tissues around the wound site. Most importantly, reactive oxidative species (ROS),
such as HO
•
are more effective in inhibiting and killing bacteria than H
2
O
2
. A variety of
nanomaterials exhibiting the enzyme-mimicking activity (called nanozymes) are able to
convert H
2
O
2
into ROS species.
The development of nanozymes, such as metal-based nanoparticles and carbon-metal
hybrid nanomaterials, which can simultaneously exhibit dual or multienzyme mimetic
activity, are a promising antibacterial therapy in wound care.
Nanozymes with embedded GOx have very recently become the object of several
studies investigating their wound-healing properties [
185
–
187
]. In a study by Du et al. [
187
],
a nanozyme comprised of clinically approved iron oxide nanoparticles coated with GOx
exhibited GOx, catalase and peroxidase-like activities.
Interestingly, in neutral and acidic microenvironments, it exhibited pH-switchable
GOx/peroxidase and GOx/catalase cascade reactions, respectively. Thus, a fabricated
Fe
3
O
4
–GOx nanozyme can, on the one hand, eradicate bacterial biofilm and shorten the
inflammatory phase of wound healing and, on the other hand, accelerate the epithelializa-
tion and remodeling phase of wound healing. In addition, in vivo testing reveals that the
Fe
3
O
4
–GOx nanozyme had superior healing activities against diabetic wounds caused by
methicillin-resistant Staphylococcus aureus compared to unjoined Fe
3
O
4
nanoparticles and
GOx.
Although these nanozymes show effective antibacterial and wound healing properties,
their direct use in clinical settings is considered questionable. Their unique characteristics,
including their small size, chemical composition and solubility, may create a high risk and
a hazard for human health. To address the limits of nanozymes in wound care, nanozymes
can be incorporated into suitable wound dressings [188].
Zhang and co-workers [
186
] developed a novel wound dressing prepared from a
nanozyme constructed by assembling GOx onto a hollow mesoporous carbon nanosphere
doped with single-atom Fe and bacterial cellulose enveloped polypropylene composites.
This nanozyme-based wound dressing exhibited outstanding breathability, biocompatibility
and water uptake and antibacterial and antibiofilm ability.
Biomolecules 2022, 12, 472 18 of 25
6. Conclusions
Glucose oxidase catalyzes the oxidation of
β
-D-glucose to D-glucono-
δ
-lactone, which
spontaneously hydrolyzes to D-gluconic acid and hydrogen peroxide in the presence of
molecular oxygen. Due to its fast turnover and high stability and specificity, glucose oxidase
has found many uses in industry and medicine.
Uses have been found for all aspects of its reaction: its consumption of glucose,
production of H
2
O
2
, consumption of oxygen and production of gluconic acid have all
found practical applications. Glucose oxidase is among the earliest of the enzymes to be
used in this way, and new applications continue to be found.
Although the biochemical and biophysical characteristics of GOx have been thoroughly
studied, there are still areas where additional study would be beneficial. For example, all
but one of the structures of GOx presently known are variations of A. niger GOx. Although
a number of industrial strains with enhanced stability or activity have been prepared, there
is no structural analysis of what makes these forms more stable. Despite numerous efforts,
there is still no structure of a GOx–substrate complex.
Moreover, some uncertainty still remains as to whether GOx might also be found in
some bacteria or whether animals other than insects might produce it. Industrial research
is also expected to develop new materials for biosensors and nanosensors and new ways of
attaching GOx to them. In addition, GOx represents a promising molecule, which, in the
form of nanozymes incorporated into wound dressings, may be of great use in wound care.
Author Contributions:
Writing—original draft preparation, J.A.B., M.Z., J.M. and V.B.-H.; writing—
review and editing, J.A.B. and V.B.-H.; visualization, J.A.B. and V.B.-H.; project administration, V.B.-H.
and J.M.; funding acquisition, V.B.-H. and J.M. All authors have read and agreed to the published
version of the manuscript.
Funding:
This research was funded by Vedecká Grantová Agentúra MŠVVaŠ SR and SAV grant
numbers 2/0131/20 and 2/0022/22.
Acknowledgments:
The authors wish to thank Eva Kutejová for general support during the writing
of this article.
Conflicts of Interest: The authors declare no conflict of interest.
Abbreviations
The following abbreviations are used in this manuscript:
GOx Glucose Oxidase
FAD Flavin Adenine Dinucleotide
NAD(P) Nicotinamide Adenine Dinucleotide (Phosphate)
PQQ PyrroloQuinoline Quinone
GMC Glucose-Methanol-Choline
RMSD Root-Mean-Squared Deviation
NMR Nuclear Magnetic Resonance
NCBI National Center for Biotechnology Information
SSF Solid State Fermentation
PDA PolyDopAmine
HSP70/90 Heat-Shock Protein 70/90
TPZ Tirapazamine
CDT ChemoDynamic Therapy
GASA 4-[(hydroxy-3-methoxyphenyl)-azo]-benzenesulfonic acid
ROS Reactive Oxygen Species
References
1.
Stolarczyk, K.; Rogalski, J.; Bilewicz, R. NAD(P)-dependent glucose dehydrogenase: Applications for biosensors, bioelectrodes,
and biofuel cells. Bioelectrochemistry 2020, 135, 107574. [CrossRef]
2.
Anthony, C. The quinoprotein dehydrogenases for methanol and glucose. Arch. Biochem. Biophys.
2004
, 428, 2–9. [CrossRef]
[PubMed]
Biomolecules 2022, 12, 472 19 of 25
3.
Bankar, S.B.; Bule, M.V.; Singhal, R.S.; Ananthanarayan, L. Glucose oxidase—An overview. Biotechnol. Adv.
2009
, 27, 489–501.
[CrossRef] [PubMed]
4.
Abrera, A.T.; Sützl, L.; Haltrich, D. Pyranose oxidase: A versatile sugar oxidoreductase for bioelectrochemical applications.
Bioelectrochemistry 2020, 132, 107409. [CrossRef] [PubMed]
5. Wilson, R.; Turner, A.P.F. Glucose oxidase: An ideal enzyme. Biosens. Bioelectron. 1992, 7, 165–185. [CrossRef]
6.
Mattevi, A. To be or not to be an oxidase: Challenging the oxygen reactivity of flavoenzymes. Trends Biochem. Sci.
2006
,
31, 276–283. [CrossRef]
7.
Cavener, D.R. GMC oxidoreductases: A newly defined family of homologous proteins with diverse catalytic activities. J. Mol.
Biol. 1992, 223, 811–814. [CrossRef]
8.
Sützl, L.; Foley, G.; Gillam, E.M.J.; Bodén, M.; Haltrich, D. The GMC superfamily of oxidoreductases revisited: Analysis and
evolution of fungal GMC oxidoreductases. Biotechnol. Biofuels 2019, 12, 118. [CrossRef]
9.
Hecht, H.J.; Kalisz, H.M.; Hendle, J.; Schmod, R.D.; Schomburg, D. Crystal Structure of Glucose Oxidase from Aspergillus niger
Refined at 2.3 Å Resolution. J. Mol. Biol. 1993, 229, 153–172. [CrossRef]
10.
Wohlfahrt, G.; Witt, S.; Hendle, J.; Schomburg, D.; Kalisz, H.M.; Hecht, H.J. 1.8 and 1.9 Å resolution structures of the Penicillium
amagasakiense and Aspergillus niger glucose oxidases as a basis for modelling substrate complexes. Acta Crystallogr. D Struct. Biol.
1999, 55, 969–977. [CrossRef]
11.
Kommoju, P.R.; Chen, Z.; Bruckner, R.C.; Mathews, F.S.; Jorns, M.S. Probing Oxygen Activation Sites in Two Flavoprotein
Oxidases Using Chloride as an Oxygen Surrogate. Biochemistry 2011, 50, 5521–5534. [CrossRef] [PubMed]
12.
Petrovi´c, D.; Frank, D.; Kamerlin, S.C.L.; Hoffmann, K.; Strodel, B. Shuffling Active Site Substate Populations Affects Catalytic
Activity: The Case of Glucose Oxidase. ACS Catal. 2017, 7, 6188–6197. [CrossRef] [PubMed]
13.
Cerutti, G.; Gugole, E.; Montemiglio, L.C.; Turbé-Doan, A.; Chena, D.; Navarro, D.; Lomascolo, A.; Piumi, F.; Exertier, C.; Freda,
I.; et al. Crystal structure and functional characterization of an oligosaccharide dehydrogenase from Pycnoporus cinnabarinus
provides insights into fungal breakdown of lignocellulose. Biotechnol. Biofuels. 2021, 14, 161. [CrossRef] [PubMed]
14.
Yoshida, H.; Sakai, G.; Mori, K.; Kojima, K.; Kamitori, S.; Sode, K. Structural analysis of fungus-derived FAD glucose dehydroge-
nase. Sci. Rep. 2015, 5, 13498. [CrossRef]
15.
Sorigué, D.; Hadjidemetriou, K.; Blangy, S.; Gotthard, G.; Bonvalet, A.; Coquelle, N.; Samire, P.; Aleksandrov, A.; Antonucci, L.;
Benachir, A.; et al. Mechanism and dynamics of fatty acid photodecarboxylase. Science
2021
, 372, eabd5687. [CrossRef] [PubMed]
16.
Carro, J.; Martínez-Júlvez, M.; Medina, M.; Martínez, A.T.; Ferreira, P. Protein dynamics promote hydride tunnelling in substrate
oxidation by aryl-alcohol oxidase. Phys. Chem. Chem. Phys. 2017, 19, 28666–28675. [CrossRef] [PubMed]
17.
Švecová, L.; Østergaard, L.H.; Skálová, T.; Schnorr, K.M.; Koval’, T.; Kolenko, P.; Stránský, J.; Sedlák, D.; Dušková, J.; Trundová, M.;
et al. Crystallographic fragment screening-based study of a novel FAD-dependent oxidoreductase from Chaetomium thermophilum.
Acta Crystallogr. D Struct. Biol. 2021, 77, 755–775. [CrossRef]
18.
Tan, T.C.; Spadiut, O.; Wongnate, T.; Sucharitakul, J.; Krondorfer, I.; Sygmund, C.; Haltrich, D.; Chaiyen, P.; Peterbauer, C.K.;
Divne, C. The 1.6 Å Crystal Structure of Pyranose Dehydrogenase from Agaricus meleagris Rationalizes Substrate Specificity and
Reveals a Flavin Intermediate. PLoS ONE 2013, 8, e53567. [CrossRef]
19.
Koch, C.; Neumann, P.; Valerius, O.; Feussner, I.; Ficner, R. Crystal Structure of Alcohol Oxidase from Pichia pastoris. PLoS ONE
2016, 11, e0149846. [CrossRef]
20.
Nguyen, Q.T.; Romero, E.; Dijkman, W.P.; de Vasconcellos, S.P.; Binda, C.; Mattevi, A.; Fraaije, M.W. Structure-Based Engineering
of Phanerochaete chrysosporium Alcohol Oxidase for Enhanced Oxidative Power toward Glycerol. Biochemistry
2018
, 57, 6209–6218.
[CrossRef]
21.
Finnegan, S.; Yuan, H.; Wang, Y.F.; Orville, A.M.; Weber, I.T.; Gadda, G. Structural and kinetic studies on the Ser101Ala variant of
choline oxidase: Catalysis by compromise. Arch. Biochem. Biophys. 2010, 501, 207–213. [CrossRef] [PubMed]
22.
Mugo, A.N.; Kobayashi, J.; Yamasaki, T.; Mikami, B.; Ohnishi, K.; Yoshikane, Y.; Yagi, T. Crystal structure of pyridoxine 4-oxidase
from Mesorhizobium loti. Biochim. Biophys. Acta 2013, 1834, 953–963. [CrossRef] [PubMed]
23.
Pickl, M.; Swoboda, A.; Romero, E.; Winkler, C.K.; Binda, C.; Mattevi, A.; Faber, K.; Fraaije, M.W. Kinetic Resolution of sec-Thiols
by Enantioselective Oxidation with Rationally Engineered 5-(Hydroxymethyl)furfural Oxidase. Angew. Chem. Int. Ed. Engl.
2018
,
57, 2864–2868. [CrossRef] [PubMed]
24.
Doubayashi, D.; Ootake, T.; Maeda, Y.; Oki, M.; Tokunaga, Y.; Sakurai, A.; Nagaosa, Y.; Mikami, B.; Uchida, H. Formate Oxidase,
an Enzyme of the Glucose-Methanol-Choline Oxidoreductase Family, Has a His-Arg Pair and 8-Formyl-FAD at the Catalytic Site.
Biosci. Biotechnol. Biochem. 2011, 75, 1662–1667. [CrossRef]
25.
Kadowaki, M.A.S.; Higasi, P.M.R.; de Godoy, M.O.; de Araújo, E.A.; Godoy, A.S.; Prade, R.A.; Polikarpov, I. Enzymatic versatility
and thermostability of a new aryl-alcohol oxidase from Thermothelomyces thermophilus M77. Biochim. Biophys. Acta Gen. Subj.
2020
,
1864, 129681. [CrossRef]
26.
Coulombe, R.; Yue, K.Q.; Ghisla, S.; Vrielink, A. Oxygen Access to the Active Site of Cholesterol Oxidase through a Narrow
Channel Is Gated by an Arg-Glu Pair. J. Biol. Chem. 2001, 276, 30435–30441. [CrossRef]
27.
Tsuge, H.; Natsuaki, O.; Ohashi, K. Purification, properties, and molecular features of glucose oxidase from Aspergillus niger. J.
Biochem. 1975, 78, 835–843. [CrossRef]
28.
Swoboda, B.E.P. The relationship between molecular conformation and the binding of flavin-adenine dinucleotide in glucose
oxidase. Biochim. Biophys. Acta 1969, 175, 365–379. [CrossRef]
Biomolecules 2022, 12, 472 20 of 25
29.
Cioci, F.; Lavecchia, R. Effect of polyols and sugars on heat-induced flavin dissociation in glucose oxidase. Biochem. Mol. Biol. Int.
1994, 34, 705–712.
30.
Gouda, M.D.; Singh, S.A.; Rao, A.G.A.; Thakur, M.S.; Karanth, N.G. Thermal Inactivation of Glucose Oxidase. Mechanism and
stabilization using additives. J. Biol. Chem. 2003, 278, 24324–24333. [CrossRef]
31.
Zoldák, G.; Zubrik, A.; Musatov, A.; Stupák, M.; Sedlák, E. Irreversible Thermal Denaturation of Glucose Oxidase from Aspergillus
niger Is the Transition to the Denatured State with Residual Structure. J. Biol. Chem.
2004
, 279, 47601–47609. [CrossRef] [PubMed]
32.
Jones, M.N.; Manley, P.; Wilkinson, A. The dissociation of glucose oxidase by sodium n-dodecyl sulphate. Biochem. J.
1982
,
203, 285–291. [CrossRef]
33.
Ye, W.N.; Combes, D. The relationship between the glucose oxidase subunit structure and its thermostability. Biochim. Biophys.
Acta 1989, 999, 86–93. [CrossRef]
34.
Leskovac, V.; Trivi´c, S.; Wohlfahrt, G.; Kandraˇc, J.; Periˇcin, D. Glucose oxidase from Aspergillus niger: The mechanism of action
with molecular oxygen, quinones, and one-electron acceptors. Int. J. Biochem. Cell Biol. 2005, 37, 731–750. [CrossRef]
35.
Gibson, Q.H.; Swoboda, B.E.P.; Massey, V. Kinetics and Mechanism of Action of Glucose Oxidase. J. Biol. Chem.
1964
,
239, 3927–3934. [CrossRef]
36.
Swoboda, B.E.P.; Massey, V. Purification and Properties of the Glucose Oxidase from Aspergillus Niger. J. Biol. Chem.
1965
,
240, 2209–2215. [CrossRef]
37.
Pazur, J.H.; Kleppe, K. The Oxidation of Glucose and Related Compounds by Glucose Oxidase from Aspergillus niger. Biochemistry
1964, 3, 578–583. [CrossRef]
38.
Kunst, A.; Draeger, B.; Ziegenhorn, J. Colorimetric methods with glucose oxidase and peroxidase. In Methods of Enzymatic
Analysis, 3rd ed.; Bergmeyered, H.U., Ed.; Chemie: Weinheim, Germany, 1984.
39.
Voet, J.G.; Coe, J.; Epstein, J.; Matossian, V.; Shipley, T. Electrostatic Control of Enzyme Reactions: Effect of Ionic Strength on the
pK
a
of an Essential Acidic Group on Glucose Oxidase. Biochemistry 1981, 20, 7182–7185. [CrossRef]
40.
Su, Q.; Klinman, J.P. Nature of Oxygen Activation in Glucose Oxidase from Aspergillus niger: The Importance of Electrostatic
Stabilization in Superoxide Formation. Biochemistry 1999, 38, 8572–8581. [CrossRef]
41.
Roth, J.P.; Klinman, J.P. Catalysis of electron transfer during activation of O
2
by the flavoprotein glucose oxidase. Proc. Natl. Acad.
Sci. USA 2003, 100, 62–67. [CrossRef]
42.
Wohlfahrt, G.; Trivi´c, S.; Zeremski, J.; Periˇcin, D.; Leskovac, V. The chemical mechanism of action of glucose oxidase from
Aspergillus niger. Mol. Cell. Biochem. 2004, 260, 69–83. [CrossRef]
43.
Bright, H.J.; Appleby, M. The pH Dependence of the Individual Steps in the Glucose Oxidase Reaction. J. Biol. Chem.
1969
,
244, 3625–3634. [CrossRef]
44.
Bright, H.J.; Gibson, Q.H. The Oxidation of 1-Deuterated Glucose by Glucose Oxidase. J. Biol. Chem.
1967
, 242, 994–1003.
[CrossRef]
45.
Sanner, C.; Macheroux, P.; Rüterjans, H.; Müller, F.; Bacher, A.
15
N- and
13
C-NMR investigations of glucose oxidase from
Aspergillus niger. Eur. J. Biochem. 1991, 196, 663–672. [CrossRef] [PubMed]
46.
Weibel, M.K.; Bright, H.J. The Glucose Oxidase Mechanism. Interpretation of the pH dependence. J. Biol. Chem.
1971
,
246, 2734–2744. [CrossRef]
47. Sawyer, D.T. Oxygen Chemistry; Oxford University Press: Oxford, UK, 1991.
48.
Prabhakar, R.; Siegbahn, P.E.M.; Minaev, B.F.; Ågren, H. Activation of Triplet Dioxygen by Glucose Oxidase: Spin-Orbit Coupling
in the Superoxide Ion. J. Phys. Chem. B 2002, 106, 3742–3750. [CrossRef]
49.
Palfey, B.A.; Ballou, D.P.; Massey, V. Oxygen Activation by Flavins and Pterins. In Active Oxygen in Biochemistry; Valentine, J.S.,
Foote, C.S., Greenberg, A., Liebman, J.F., Eds.; Springer: Dordrecht, The Netherlands, 1995; pp. 37–83. [CrossRef]
50.
Kalisz, H.M.; Hecht, H.J.; Schomburg, D.; Schmid, R.D. Effects of carbohydrate depletion on the structure, stability and activity of
glucose oxidase from Aspergillus niger. Biochim. Biophys. Acta 1991, 1080, 138–142. [CrossRef]
51.
Kalisz, H.M.; Hecht, H.J.; Schomburg, D.; Schmid, R.D. Crystallization and preliminary X-ray diffraction studies of a deglycosy-
lated glucose oxidase from Aspergillus niger. J. Mol. Biol. 1990, 213, 207–209. [CrossRef]
52.
Kohen, A.; Jonsson, T.; Klinman, J.P. Effects of Protein Glycosylation on Catalysis: Changes in Hydrogen Tunneling and Enthalpy
of Activation in the Glucose Oxidase Reaction. Biochemistry 1997, 36, 2603–2611. [CrossRef]
53.
Edge, A.S.B. Deglycosylation of glycoproteins with trifluoromethanesulphonic acid: elucidation of molecular structure and
function. Biochem. J. 2003, 376, 339–350. [CrossRef]
54.
Frederick, K.R.; Tung, J.; Emerick, R.S.; Masiarz, F.R.; Chamberlain, S.H.; Vasavada, A.; Rosenberg, S. Glucose Oxidase from
Aspergillus niger. Cloning, gene sequence, secretion from Saccharomyces cerevisiae and kinetic analysis of a yeast-derived enzyme.
J. Biol. Chem. 1990, 265, 3793–3802. [CrossRef]
55.
Dubey, M.K.; Zehra, A.; Aamir, M.; Meena, M.; Ahirwal, L.; Singh, S.; Shukla, S.; Upadhyay, R.S.; Bueno-Mari, R.; Bajpai, V.K.
Improvement Strategies, Cost Effective Production, and Potential Applications of Fungal Glucose Oxidase (GOD): Current
Updates. Front. Microbiol. 2017, 8, 1032. [CrossRef] [PubMed]
56.
Bean, R.C.; Hassid, W.Z. Carbohydrate oxidase from a red alga, Iridophycus flaccidum. J. Biol. Chem.
1956
, 218, 425–436. [CrossRef]
57.
Bean, R.C.; Porter, G.G.; Steinberg, B.M. Carbohydrate Metabolism of Citrus Fruits. II. Oxidation of sugars by an aerodehydroge-
nase from young orange fruits. J. Biol. Chem. 1961, 236, 1235–1240. [CrossRef]
58. Heyningen, R.V. Metabolism of Xylose by the Lens. Calf lens in vitro. Biochem. J. 1956, 69, 481–491. [CrossRef]
Biomolecules 2022, 12, 472 21 of 25
59.
Dowling, J.H.; Levine, H.B. Hexose oxidation by an enzyme system of Malleomyces pseudomallei. J. Bacteriol.
1956
, 72, 555–560.
[CrossRef]
60.
King, T.E.; Cheldelin, V.H. Glucose oxidation and cytochromes in solubilized particulate fractions of Acetobacter suboxydans. J.
Biol. Chem. 1957, 224, 579–590. [CrossRef]
61.
Bentley, R.; Slechta, L. Oxidation of mono- and disaccharides to aldonic acids by Pseudomonas species. J. Bacteriol.
1960
, 79, 346–355.
[CrossRef]
62. Hauge, J.G. Glucose dehydrogenation in bacteria: A comparative study. J. Bacteriol. 1961, 82, 609–614. [CrossRef]
63.
Ohashi, K.; Natori, S.; Kubo, T. Expression of amylase and glucose oxidase in the hypopharyngeal gland with an age-dependent
role change of the worker honeybee (Apis mellifera L.). Eur. J. Biochem. 1999, 265, 127–133. [CrossRef]
64.
Tang, Q.; Hu, Y.; Kang, L.; Wang, C.Z. Characterization of glucose-induced glucose oxidase gene and protein expression in
Helicoverpa armigera larvae. Arch. Insect Biochem. Physiol. 2012, 79, 104–119. [CrossRef]
65.
Afshar, K.; Dufresne, P.J.; Pan, L.; Merkx-Jacques, M.; Bede, J.C. Diet-specific salivary gene expression and glucose oxidase
activity in Spodoptera exigua (Lepidoptera: Noctuidae) larvae. J. Insect Physiol. 2010, 56, 1798–1806. [CrossRef] [PubMed]
66.
Caridis, K.A.; Christakopoulos, P.; Macris, B.J. Simultaneous production of glucose oxidase and catalase by Alternaria alternata.
Appl. Microbiol. Biotechnol. 1991, 34, 794–797. [CrossRef]
67.
Yuivar, Y.; Barahona, S.; Alcaíno, J.; Cifuentes, V.; Baeza, M. Biochemical and Thermodynamical Characterization of Glucose
Oxidase, Invertase, and Alkaline Phosphatase Secreted by Antarctic Yeasts. Front. Mol. Biosci.
2017
, 4, 86. [CrossRef] [PubMed]
68.
Raghukumar, C.; Mohandass, C.; Kamat, S.; Shailaja, M.S. Simultaneous detoxification and decolorization of molasses spent wash
by the immobilized white-rot fungus Flavodon flavus isolated from the marine habitat. Enzym. Microb. Technol.
2004
, 35, 197–202.
[CrossRef]
69.
Macías-Sánchez, K.; García-Soto, J.; López-Ramírez, A.; Martínez-Cadena, G. Rho1 and other GTP-binding proteins are associated
with vesicles carrying glucose oxidase activity from Fusarium oxysporum f. sp. lycopersici. Antonie Leeuwenhoek
2011
, 99, 671–680.
[CrossRef] [PubMed]
70.
Moreno-Jiménez, R.; García-Soto, J.; Martínez-Cadena, G. Small GTP-binding proteins are associated with chitosomes and
vesicles carrying glucose oxidase from Mucor circinelloides. Microbiology (Reading) 2008, 154, 842–851. [CrossRef]
71.
Eremin, A.N.; Makarenko, M.V.; Zhukovskaia, L.A.; Mikhailova, R.V. Isolation and characterization of extracellular glucose
oxidase from Penicillium adametzii LF F-2044.1. Prikl. Biokhim. Mikrobiol. 2006, 42, 345–352. [CrossRef]
72.
Kiess, M.; Hecht, H.J.; Kalisz, H.M. Glucose oxidase from Penicillium amagasakiense. Primary structure and comparisonwith other
glucose-methanol-choline (GMC) oxidoreductases. Eur. J. Biochem. 1998, 252, 90–99. [CrossRef]
73.
Johnstone-Robertson, M.; Clarke, K.G.; Harrison, S.T.L. Characterization of the distribution of glucose oxidase in Penicillium
sp. CBS 120262 and Aspergillus niger NRRL-3 cultures and its effect on integrated product recovery. Biotechnol. Bioeng.
2008
,
99, 910–918. [CrossRef]
74.
Gao, Z.; Li, Z.; Zhang, Y.; Huang, H.; Li, M.; Zhou, L.; Tang, Y.; Yao, B.; Zhang, W. High-level expression of the Penicillium notatum
glucose oxidase gene in Pichia pastoris using codon optimization. Biotechnol. Lett. 2012, 34, 507–514. [CrossRef]
75.
Kim, H.W.; Kimura, S.; Ohno, N.; Okadome, M.; Takahashi, H.; Amachi, S.; Shinoyama, H.; Fujii, T. Purification of Glucose
Oxidase and Catalase Produced by the Apple Blue Mold, Penicillium expansum O-385-10, and Their Characteristics Including the
Browning of Apple Fruit. Jpn. J. Food Microbiol. 2005, 22, 10–16. [CrossRef]
76.
Abalikhina, T.A.; Morozkin, A.D.; Bogdanov, V.P.; Kaverznera, E. Composition and structure of glucose oxidase from Penicillium
vitale. Biokhimiya 1971, 36, 191–198.
77.
Chi, B.B.; Lu, Y.N.; Yin, P.C.; Liu, H.Y.; Chen, H.Y.; Shan, Y. Sequencing and Comparative Genomic Analysis of a Highly
Metal-Tolerant Penicillium janthinellum P1 Provide Insights Into Its Metal Tolerance. Front. Microbiol.
2021
, 12, 663217. [CrossRef]
78.
Zhao, J.; Janse, B.J.H. Comparison of H
2
O
2
-producing enzymes in selected white rot fungi. FEMS Microbiol. Lett.
1996
,
139, 215–221. [CrossRef]
79.
Shin, K.S.; Youn, H.D.; Han, Y.H.; Kang, S.O.; Hah, Y.C. Purification and characterization of D-glucose oxidase from white-rot
fungus Pleurotus ostreatus. Eur. J. Biochem. 1993, 215, 747–752. [CrossRef] [PubMed]
80.
Levasseur, A.; Lomascolo, A.; Chabrol, O.; Ruiz-Dueñas, F.J.; Boukhris-Uzan, E.; Piumi, F.; Kües, U.; Ram, A.F.J.; Murat, C.;
Haon, M.; et al. The genome of the white-rot fungus Pycnoporus cinnabarinus: A basidiomycete model with a versatile arsenal for
lignocellulosic biomass breakdown. BMC Genom. 2014, 15, 486. [CrossRef]
81.
Guimarães, L.H.S.; Peixoto-Nogueira, S.C.; Michelin, M.; Rizzatti, A.C.S.; Sandrim, V.C.; Zanoelo, F.F.; Aquino, A.C.M.M.; Junior,
A.B.; de Lourdes T. M. Polizeli, M. Screening of filamentous fungi for production of enzymes of biotechnological interest. Braz. J.
Microbiol. 2006, 37, 474–480. [CrossRef]
82.
Ohm, R.A.; de Jong, J.F.; Lugones, L.G.; Aerts, A.; Kothe, E.; Stajich, J.E.; de Vries, R.P.; Record, E.; Levasseur, A.; Baker, S.E.; et al.
Genome sequence of the model mushroom Schizophyllum commune. Nat. Biotechnol. 2010, 28, 957–963. [CrossRef]
83.
Kim, K.K.; Fravel, D.R.; Papavizas, G.C. Production, purification, and properties of glucose oxidase from the biocontrol fungus
Talaromyces flavus. Can. J. Microbiol. 1990, 36, 199–205. [CrossRef]
84.
Semashko, T.V.; Mikhailova, R.V.; Eremin, A.N. Extracellular Glucose Oxidase of Penicillium funiculosum 46.1. Appl. Biochem.
Microbiol. 2003, 39, 368–374. [CrossRef]
85.
Rando, D.; Kohring, G.W.; Giffhorn, F. Production, purification and characterization of glucose oxidase from a newly isolated
strain of Penicillium pinophilum. Appl. Microbiol. Biotechnol. 1997, 48, 34–40. [CrossRef]
Biomolecules 2022, 12, 472 22 of 25
86.
Nakamatsu, T.; Akamatsu, T.; Miyajima, R.; Simo, I. Microbial Production of Glucose Oxidase. Agric. Biol. Chem.
1975
,
39, 1803–1811. [CrossRef]
87.
Nierman, W.C.; Fedorova-Abrams, N.D.; Andrianopoulos, A. Genome Sequence of the AIDS-Associated Pathogen Penicillium
marneffei (ATCC18224) and Its Near Taxonomic Relative Talaromyces stipitatus (ATCC10500). Genome Announc.
2015
, 3, e01559-14.
[CrossRef] [PubMed]
88.
Pulci, V.; D’Ovidio, R.; Petruccioli, M.; Federici, F. The glucose oxidase of Penicillium variabile P16: Gene cloning, sequencing and
expression. Lett. Appl. Microbiol. 2004, 38, 233–238. [CrossRef]
89.
Wibberg, D.; Jelonek, L.; Rupp, O.; Hennig, M.; Eikmeyer, F.; Goesmann, A.; Hartmann, A.; Borriss, R.; Grosch, R.; Pühler, A.; et al.
Establishment and interpretation of the genome sequence of the phytopathogenic fungus Rhizoctonia solani AG1-IB isolate 7/3/14.
J. Biotechnol. 2013, 167, 142–155. [CrossRef] [PubMed]
90.
Gazis, R.; Kuo, A.; Riley, R.; LaButti, K.; Lipzen, A.; Lin, J.; Amirebrahimi, M.; Hesse, C.N.; Spatafora, J.W.; Henrissat, B.; et al.
The genome of Xylona heveae provides a window into fungal endophytism. Fungal Biol. 2016, 120, 26–42. [CrossRef]
91.
Guindon, S.; Dufayard, J.F.; Lefort, V.; Anisimova, M.; Hordijk, W.; Gascuel, O. New Algorithms and Methods to Estimate
Maximum-Likelihood Phylogenies: Assessing the Performance of PhyML 3.0. Syst. Biol. 2010, 59, 307–321. [CrossRef]
92.
Sievers, F.; Higgins, D.G. Clustal Omega for making accurate alignments of many protein sequences. Protein Sci.
2018
, 27, 135–145.
[CrossRef]
93.
Di Tommaso, P.; Moretti, S.; Xenarios, I.; Orobitg, M.; Montanyola, A.; Chang, J.M.; Taly, J.F.; Notredame, C. T-Coffee: A web
server for the multiple sequence alignment of protein and RNA sequences using structural information and homology extension.
Nucleic Acids Res. 2011, 39, W13–W17. [CrossRef]
94.
Allameh, S.; Rabbani, M. A Distance-Based Microfluidic Paper-Based Biosensor for Glucose Measurements in Tear Range. Appl.
Biochem. Biotechnol. 2022. [CrossRef]
95.
Asrami, P.N.; Azar, P.A.; Tehrani, M.S.; Mozaffari, S.A. Glucose Oxidase/Nano-ZnO/Thin Film Deposit FTO as an Innovative
Clinical Transducer: A Sensitive Glucose Biosensor. Front. Chem. 2020, 8, 503. [CrossRef]
96.
Devasenathipathy, R.; Mani, V.; Chen, S.M.; Huang, S.T.; Huang, T.T.; Lind, C.M.; Hwae, K.Y.; Chen, T.Y.; Chen, B.J. Glucose
biosensor based on glucose oxidase immobilized at gold nanoparticles decorated graphene-carbon nanotubes. Enzym. Microb.
Technol. 2015, 78, 40–45. [CrossRef] [PubMed]
97.
J˛edrzak, A.; R˛ebi´s, T.; Kuznowicz, M.; Jesionowski, T. Bio-inspired magnetite/lignin/polydopamine-glucose oxidase biosensing
nanoplatform. From synthesis, via sensing assays to comparison with others glucose testing techniques. Int. J. Biol. Macromol.
2019, 127, 677–682. [CrossRef] [PubMed]
98.
Kalaivani, G.J.; Suja, S.K. Nanomolar level sensing of glucose in food samples using glucose oxidase confined MWCNT-Inulin-
TiO
2
bio-nanocomposite. Food Chem. 2019, 298, 124981. [CrossRef] [PubMed]
99.
He, T.; Xu, H.; Zhang, Y.; Yi, S.; Cui, R.; Xing, S.; Wei, C.; Lin, J.; Huang, P. Glucose Oxidase-Instructed Traceable Self-
Oxygenation/Hyperthermia Dually Enhanced Cancer Starvation Therapy. Theranostics
2020
, 10, 1544–1554. [CrossRef] [PubMed]
100.
Chai, Z.; Dong, H.; Sun, X.; Fan, Y.; Wang, Y.; Huang, F. Development of glucose oxidase-immobilized alginate nanoparticles for
enhanced glucose-triggered insulin delivery in diabetic mice. Int. J. Biol. Macromol. 2020, 159, 640–647. [CrossRef]
101.
Shao, L.; Li, Y.; Huang, F.; Wang, X.; Lu, J.; Jia, F.; Pan, Z.; Cui, X.; Ge, G.; Deng, X.; et al. Complementary autophagy inhibition
and glucose metabolism with rattle-structured polydopamine@mesoporous silica nanoparticles for augmented low-temperature
photothermal therapy and in vivo photoacoustic imaging. Theranostics 2020, 10, 7273–7286. [CrossRef]
102.
German, N.; Ramanaviciene, A.; Ramanavicius, A. Formation of Polyaniline and Polypyrrole Nanocomposites with Embedded
Glucose Oxidase and Gold Nanoparticles. Polymers 2019, 11, 377. [CrossRef]
103.
Wei, X.; Chen, J.; Ali, M.C.; Munyemana, J.C.; Qiu, H. Cadmium cobaltite nanosheets synthesized in basic deep eutectic solvents
with oxidase-like, peroxidase-like, and catalase-like activities and application in the colorimetric assay of glucose. Mikrochim.
Acta 2020, 187, 314. [CrossRef]
104.
Garcia-Hernandez, C.; Garcia-Cabezon, C.; Martin-Pedrosa, F.; Rodriguez-Mendez, M.L. Analysis of musts and wines by means
of a bio-electronic tongue based on tyrosinase and glucose oxidase using polypyrrole/gold nanoparticles as the electron mediator.
Food Chem. 2019, 289, 751–756. [CrossRef] [PubMed]
105.
Steffolani, M.E.; Ribotta, P.D.; Pérez, G.T.; León, A.E. Combinations of glucose oxidase,
α
-amylase and xylanase affect dough
properties and bread quality. Int. J. Food Sci. Technol. 2012, 47, 525–534. [CrossRef]
106.
Mohammadnejad, P.; Asl, S.S.; Aminzadeh, S.; Haghbeen, K. A new sensitive spectrophotometric method for determination of
saliva and blood glucose. Spectrochim. Acta A Mol. Biomol. Spectrosc. 2020, 229, 117897. [CrossRef]
107.
Yu, J.; Zhang, Y.; Ye, Y.; DiSanto, R.; Sun, W.; Ranson, D.; Ligler, F.S.; Buse, J.B.; Gu, Z. Microneedle-array patches loaded with
hypoxia-sensitive vesicles provide fast glucose-responsive insulin delivery. Proc. Natl. Acad. Sci. USA
2015
, 112, 8260–8265.
[CrossRef]
108.
Yu, J.; Qian, C.; Zhang, Y.; Cui, Z.; Zhu, Y.; Shen, Q.; Ligler, F.S.; Buse, J.B.; Gu, Z. Hypoxia and H
2
O
2
Dual-Sensitive Vesicles for
Enhanced Glucose-Responsive Insulin Delivery. Nano Lett. 2017, 17, 733–739. [CrossRef] [PubMed]
109.
Guo, Y.; Jia, H.R.; Zhang, X.; Zhang, X.; Sun, Q.; Wang, S.Z.; Zhao, J.; Wu, F.G. A Glucose/Oxygen-Exhausting Nanoreactor
for Starvation- and Hypoxia-Activated Sustainable and Cascade Chemo-Chemodynamic Therapy. Small
2020
, 16, e2000897.
[CrossRef] [PubMed]
Biomolecules 2022, 12, 472 23 of 25
110.
Kornecki, J.F.; Carballares, D.; Tardioli, P.W.; Rodrigues, R.C.; Berenguer-Murcia, Á.; Alcántarae, A.R.; Fernandez-Lafuente, R.
Enzyme production of D-gluconic acid and glucose oxidase: Successful tales of cascade reactions. Catal. Sci. Technol.
2020
,
10, 5740. [CrossRef]
111.
Khatami, S.H.; Vakili, O.; Ahmadi, N.; Fard, E.S.; Mousavi, P.; Khalvati, B.; Maleksabet, A.; Savardashtaki, A.; Taheri-Anganeh,
M.; Movahedpour, A. Glucose oxidase: Applications, sources, and recombinant production. Biotechnol. Appl. Biochem.
2021
,
Online ahead of print. [CrossRef]
112.
FDA. Enzyme Preparations Used in Food (Partial List). 2018. Available online: https://www.fda.gov/food/generally-recognized-
safe-gras/enzyme-preparations-used-food-partial-list (accessed 27 February 2022).
113.
Kalisz, H.M.; Hendle, J.; Schmid, R.D. Structural and biochemical properties of glycosylated and deglycosylated glucose oxidase
from Penicillium amagasakiense. Appl. Microbiol. Biotechnol. 1997, 47, 502–507. [CrossRef]
114.
Kusai, K.; Sekuzu, I.; Hagihara, B.; Okunuki, K.; Yamauchi, S.; Nakai, M. Crystallization of glucose oxidase from Penicillium
amagasakiense. Biochim. Biophys. Acta 1960, 40, 555–557. [CrossRef]
115.
Holland, J.T.; Harper, J.C.; Dolan, P.L.; Manginell, M.M.; Arango, D.C.; Rawlings, J.A.; Apblett, C.A.; Brozik, S.M. Rational
redesign of glucose oxidase for improved catalytic function and stability. PLoS ONE 2012, 7, e37924. [CrossRef] [PubMed]
116.
Song, H.T.; Xiao, W.J.; Yang, Y.M.; Zhao, Y.; Gao, Y.; Liu, S.H.; Liu, Z.L.; Xia, W.C.; Li, R.; Li, N.N.; et al. Improving the
anti-oxidation of glucose oxidase with computer-aided structure optimization. J. Adv. Biotechnol. 2016, 53, 736–740. [CrossRef]
117.
Marín-Navarro, J.; Roupain, N.; Talens-Perales, D.; Polaina, J. Identification and Structural Analysis of Amino Acid Substitutions
that Increase the Stability and Activity of Aspergillus niger Glucose Oxidase. PLoS ONE
2015
, 10, e0144289. [CrossRef] [PubMed]
118.
Mirón, J.; González, M.P.; Pastrana, L.; Murado, M.A. Diauxic production of glucose oxidase by Aspergillus niger in submerged
culture: A dynamic model. Enzym. Microb. Technol. 2002, 31, 615–620. [CrossRef]
119.
Mirón, J.; Vázquez, J.A.; González, P.; Murado, M.A. Enhancement glucose oxidase production by solid-state fermentation of
Aspergillus niger on polyurethane foams using mussel processing wastewaters. Enzym. Microb. Technol.
2010
, 46, 21–27. [CrossRef]
120.
Kriaa, M.; Kammoun, R. Producing Aspergillus tubingensis CTM507 glucose oxidase by solid state fermentation versus submerged
fermentation: Process optimization and enzyme stability by an intermediary metabolite in relation with diauxic growth. J. Chem.
Technol. Biotechnol. 2016, 91, 1540–1550. [CrossRef]
121.
Hatzinikolaou, D.G.; Macris, B.J. Factors regulating production of glucose oxidase by Aspergillus niger. Enzym. Microb. Technol.
1995, 17, 530–534. [CrossRef]
122.
Hatzinikolaou, D.G.; Hansen, O.C.; Macris, B.J.; Tingey, A.; Kekos, D.; Goodenough, P.; Stougaard, P. A new glucose oxidase
from Aspergillus niger: Characterization and regulation studies of enzyme and gene. Appl. Microb. Biotechnol.
1996
, 46, 371–381.
[CrossRef]
123.
Liu, J.Z.; Huang, Y.Y.; Liu, J.; Weng, L.P.; Ji, L.N. Effects of metal ions on simultaneous production of glucose oxidase and catalase
by Aspergillus niger. Lett. Appl. Microbiol. 2001, 32, 16–19. [CrossRef]
124.
Toren, E.C., Jr.; Burger, F.J. Trace determination of metal ion inhibitors of the glucose-glucose oxidase system. Mikrochim. Acta
1968, 56, 538–545. [CrossRef]
125.
Nakamura, S.; Ogura, Y. Mode of Inhibition of Glucose Oxidase by Metal Ions. J. Biochem.
1968
, 64, 439–447. [CrossRef] [PubMed]
126.
Khurshid, S.; Kashmiri, M.A.; Quershi, Z.; Ahmad, W. Optimization of glucose oxidase production by Aspergillus niger. Afr. J.
Biotechnol. 2011, 10, 1674–1678. [CrossRef]
127.
Ramzan, M.; Mehmood, T. Enhanced production of glucose oxidase from UV-mutant of Aspergillus niger. Afr. J. Biotechnol.
2009
,
8, 288–290. [CrossRef]
128.
Park, E.H.; Shin, Y.M.; Lim, Y.Y.; Kwon, T.H.; Kim, D.H.; Yang, M.S. Expression of glucose oxidase by using recombinant yeast. J.
Biotechnol. 2000, 81, 35–44. [CrossRef]
129.
Kapat, A.; Jung, J.K.; Park, Y.H. Enhancement of glucose oxidase production in batch cultivation of recombinant Saccharomyces
cerevisiae: Optimization of oxygen transfer condition. J. Appl. Microbiol. 2001, 90, 216–222. [CrossRef] [PubMed]
130.
Malherbe, D.F.; du Toit, M.; Otero, R.R.C.; van Rensburg, P.; Pretorius, I.S. Expression of the Aspergillus niger glucose oxidase
gene in Saccharomyces cerevisiae and its potential applications in wine production. Appl. Microbiol. Biotechnol.
2003
, 61, 502–511.
[CrossRef]
131.
Demain, A.L.; Vaishnav, P. Production of recombinant proteins by microbes and higher organisms. Biotechnol. Adv.
2009
,
27, 297–306. [CrossRef]
132.
Courjean, O.; Mano, N. Recombinant glucose oxidase from Penicillium amagasakiense for efficient bioelectrochemical applications
in physiological conditions. J. Biotechnol. 2011, 151, 122–129. [CrossRef]
133. Romanos, M.A.; Scorer, C.A.; Clare, J.J. Foreign Gene Expression in Yeast: A Review. Yeast 1992, 8, 423–488. [CrossRef]
134.
Guo, Y.; Lu, F.; Zhao, H.; Tang, Y.; Lu, Z. Cloning and heterologous expression of glucose oxidase gene from Aspergillus niger Z-25
in Pichia pastoris. Appl. Biochem. Biotechnol. 2010, 162, 498–509. [CrossRef]
135.
Qiu, Z.; Guo, Y.; Bao, X.; Hao, J.; Sun, G.; Peng, B.; Bi, W. Expression of Aspergillus niger glucose oxidase in yeast Pichia pastoris
SMD1168. Biotechnol. Biotechnol. Equip. 2016, 30, 998–1005. [CrossRef]
136.
Rasiah, I.A.; Sutton, K.H.; Low, F.L.; Lin, H.M.; Gerrard, J.A. Crosslinking of wheat dough proteins by glucose oxidase and the
resulting effects on bread and croissants. Food Chem. 2005, 89, 325–332. [CrossRef]
137.
Wong, C.M.; Wong, K.H.; Chen, X.D. Glucose oxidase: Natural occurrence, function, properties and industrial applications. Appl.
Microbiol. Biotechnol. 2008, 78, 927–938. [CrossRef] [PubMed]
Biomolecules 2022, 12, 472 24 of 25
138.
Steffolani, M.E.; Ribotta, P.D.; Pérez, G.T.; León, A.E. Effect of glucose oxidase, transglutaminase, and pentosanase on wheat
proteins: Relationship with dough properties and bread-making quality. J. Cereal Sci. 2010, 51, 366–373. [CrossRef]
139.
Vemulapalli, V.; Miller, K.A.; Hoseney, R.C. Glucose Oxidase in Breadmaking Systems. Cereal Chem.
1998
, 75, 439–442. [CrossRef]
140.
El-Rashidy, L.A.; Bahlol, H.E.M.; El-Desoky, A.A. Improving Quality of Pan Bread by Using Glucose Oxidase and Lipase Enzymes.
Middle East J. Appl. Sci. 2015, 5, 1035–1043.
141.
Dagdelen, A.F.; Gocmen, D. Effects of glucose oxidase, hemicellulase and ascorbic acid on dough and bread quality. J. Food Qual.
2007, 30, 1009–1022. [CrossRef]
142.
Decamps, K.; Joye, I.J.; Rakotozafy, L.; Nicolas, J.; Courtin, C.M.; Delcour, J.A. The bread dough stability improving effect of
pyranose oxidase from Trametes multicolour and glucose oxidase from Aspergillus niger: Unraveling the molecular mechanism. J.
Agric. Food Chem. 2013, 61, 7848–7854. [CrossRef]
143.
Kriaa, M.; Ouhibi, R.; Graba, H.; Besbes, S.; Jardak, M.; Kammoun, R. Synergistic effect of Aspergillus tubingensis CTM 507 glucose
oxidase in presence of ascorbic acid and alpha amylase on dough properties, baking quality and shelf life of bread. J. Food Sci.
Technol. 2016, 53, 1259–1268. [CrossRef]
144.
Tozatti, P.; Hopkins, E.J.; Briggs, C.; Hucl, P.; Nickerson, M.T. Effect of chemical oxidizers and enzymatic treatments on the baking
quality of doughs formulated with five Canadian spring wheat cultivars. Food Sci. Technol. Int. 2020, 26, 614–628. [CrossRef]
145.
Röcker, J.; Schmitt, M.; Pasch, L.; Ebert, K.; Grossmann, M. The use of glucose oxidase and catalase for the enzymatic reduction of
the potential ethanol content in wine. Food Chem. 2016, 210, 660–670. [CrossRef] [PubMed]
146.
Valencia, P.; Espinoza, K.; Ramirez, C.; Franco, W.; Urtubia, A. Technical Feasibility of Glucose Oxidase as a Prefermentation
Treatment for Lowering the Alcoholic Degree of Red Wine. Am. J. Enol. Vitic. 2017, 68, 386–389. [CrossRef]
147. B˘anic˘a, F.G. Chemical Sensors and Biosensors: Fundamentals and Applications; John Wiley & Sons: Chichester, UK, 2012.
148.
Lopes, F.M.; de Aleluia Batista, K.; Batista, G.L.A.; Fernandes, K.F. Biosensor for determination of glucose in real samples of
beverages. Food Sci. Technol. 2012, 32, 65–69. [CrossRef]
149.
Mason, M.; Longo, E.; Scampicchio, M. Monitoring of glucose in beer brewing by a carbon nanotubes based nylon nanofibrous
biosensor. J. Nanomater. 2016, 2016, 5217023. [CrossRef]
150. Zeeb, B.; Fischer, L.; Weiss, J. Stabilization of food dispersions by enzymes. Food Funct. 2014, 5, 198–213. [CrossRef]
151.
Golikova, E.P.; Lakina, N.V.; Grebennikova, O.V.; Matveeva, V.G.; Sulman, E.M. A study of biocatalysts based on glucose oxidase.
Faraday Discuss. 2017, 202, 303–314. [CrossRef]
152.
Sisak, C.; Csanádi, Z.; Rónay, E.; Szajáni, B. Elimination of glucose in egg white using immobilized glucose oxidase. Enzym.
Microb. Technol. 2006, 39, 1002–1007. [CrossRef]
153.
El-Hariri, M.; Al-Yazeed, H.A.; Samir, A.; Elhelw, R.; Soliman, R. Genetic and phenotypic diversity of naturally isolated wild
strains of Aspergillus niger with hyper glucose oxidase production. J. Biosci. Biotechnol. 2015, 4, 245–253.
154. Kirk, O.; Borchert, T.V.; Fuglsang, C.C. Industrial enzyme applications. Curr. Opin. Biotechnol. 2002, 13, 345–351. [CrossRef]
155.
Karimi, A.; Mahdizadeh, F.; Salari, D.; Vahabzadeh, F.; Khataee, A. Enzymatic scavenging of oxygen dissolved in water:
Application of response surface methodology in optimization of conditions. Chem. Ind. Chem. Eng. Q.
2012
, 18, 431–439.
[CrossRef]
156.
Crueger, A.; Crueger, W. Glucose transforming enzymes. In Microbial Enzymes and Biotechnology; Fogarty, W.M., Kelly, C.T., Eds.;
Elsevier: New York, NY, USA, 1990; pp. 177–226.
157.
Bhat, S.V.; Swathi, B.R.; Rosy, M.; Govindappa, M. Isolation and characterization of glucose oxidase (GOD) from Aspergillus flavus
and Penicillium sp. Int. J. Curr. Microbiol. Appl. Sci. 2013, 2, 153–161.
158.
Isaksen, A.; Adler-Nissen, J. Antioxidative Effect of Glucose Oxidase and Catalase in Mayonnaises of Different Oxidative
Susceptibility. I. Product Trials. Lebensm. Wiss. Technol. 1997, 30, 841–846. [CrossRef]
159. Cichello, S.A. Oxygen absorbers in food preservation: A review. J. Food Sci. Technol. 2015, 52, 1889–1895. [CrossRef] [PubMed]
160.
Vartiainen, J.; Rättö, M.; Paulussen, S. Antimicrobial Activity of Glucose Oxidase-immobilized Plasma-activated Polypropylene
Films. Packag. Technol. Sci. 2005, 18, 243–251. [CrossRef]
161.
Hanušová, K.; Vápenka, L.; Dobiáš, J.; Mišková, L. Development of antimicrobial packaging materials with immobilized glucose
oxidase and lysozyme. Cent. Eur. J. Chem. 2013, 11, 1066–1078. [CrossRef]
162.
Yuan, H.; Bai, H.; Liu, L.; Lv, F.; Wang, S. A glucose-powered antimicrobial system using organic-inorganic assembled network
materials. Chem. Commun. (Camb.) 2015, 51, 722–724. [CrossRef]
163.
Xu, M.; Wanga, R.; Li, Y. An electrochemical biosensor for rapid detection of E. coli O157:H7 with highly efficient bifunctional
glucose oxidase-polydopamine nanocomposites and Prussian blue modified screen-printed interdigitated electrode. Analyst
2016, 141, 5441. [CrossRef]
164.
Wang, M.; Wang, D.; Chen, Q.; Li, C.; Li, Z.; Lin, J. Recent Advances in Glucose-Oxidase-Based Nanocomposites for Tumor
Therapy. Small 2019, 15, e1903895. [CrossRef]
165.
Chu, K.F.; Dupuy, D.E. Thermal ablation of tumours: Biological mechanisms and advances in therapy. Nat. Rev. Cancer
2014
,
14, 199–208. [CrossRef]
166.
Cheng, L.; Wang, C.; Feng, L.; Yang, K.; Liu, Z. Functional nanomaterials for phototherapies of cancer. Chem. Rev.
2014
,
114, 10869–10939. [CrossRef]
Biomolecules 2022, 12, 472 25 of 25
167.
Wang, T.; Zhang, H.; Liu, H.; Yuan, Q.; Ren, F.; Han, Y.; Sun, Q.; Li, Z.; Gao, M. Boosting H
2
O
2
-Guided Chemodynamic Therapy
of Cancer by Enhancing Reaction Kinetics through Versatile Biomimetic Fenton Nanocatalysts and the Second Near-Infrared
Light Irradiation. Adv. Funct. Mater. 2020, 30, 1906128. [CrossRef]
168.
Clark, L.C., Jr.; Lyons, C. Electrode systems for continuous monitoring in cardiovascular surgery. Ann. N. Y. Acad. Sci.
1962
,
102, 29–45. [CrossRef] [PubMed]
169.
Pullano, S.A.; Greco, M.; Bianco, M.G.; Foti, D.; Brunetti, A.; Fiorillo, A.S. Glucose biosensors in clinical practice: Principles, limits
and perspectives of currently used devices. Theranostics 2022, 12, 493–511. [CrossRef] [PubMed]
170.
Klonoff, D.C.; Ahn, D.; Drincic, A. Continuous glucose monitoring: A review of the technology and clinical use. Diabetes Res.
Clin. Pract. 2017, 133, 178–192. [CrossRef]
171.
Fokkert, M.; van Dijk, P.R.; Edens, M.A.; Hernández, A.D.; Slingerland, R.; Gans, R.; Álvarez, E.D.; Bilo, H. Performance of the
Eversense versus the Free Style Libre Flash glucose monitor during exercise and normal daily activities in subjects with type 1
diabetes mellitus. BMJ Open Diabetes Res. Care 2020, 8, e001193. [CrossRef]
172.
Gutierrez, E.A.; Mundhada, H.; Meier, T.; Duefel, H.; Bocola, M.; Schwaneberg, U. Reengineered glucose oxidase for amperometric
glucose determination in diabetes analytics. Biosens. Bioelectron. 2020, 50, 84–90. [CrossRef]
173.
Yang, F.; Li, M.; Liu, Y.; Wang, T.; Feng, Z.; Cui, H.; Gu, N. Glucose and magnetic-responsive approach toward in situ nitric oxide
bubbles controlled generation for hyperglycemia theranostics. J. Control. Release 2016, 228, 87–95. [CrossRef]
174.
Spinas, G.A.; Laffranchi, R.; Francoys, I.; David, I.; Richter, C.; Reinecke, M. The early phase of glucose-stimulated insulin
secretion requires nitric oxide. Diabetologia 1998, 41, 292–299. [CrossRef]
175.
Ma, R.; Shi, L. Phenylboronic acid-based glucose-responsive polymeric nanoparticles: Synthesis and applications in drug delivery.
Polym. Chem. 2014, 5, 1503–1518. [CrossRef]
176.
Molan, P.C. The evidence supporting the use of honey as a wound dressing. Int. J. Low. Extrem. Wounds
2006
, 5, 40–54. [CrossRef]
177.
Bucekova, M.; Valachova, I.; Kohutova, L.; Prochazka, E.; Klaudiny, J.; Majtan, J. Honeybee glucose oxidase–its expression
in honeybee workers and comparative analyses of its content and H
2
O
2
-mediated antibacterial activity in natural honeys.
Naturwissenschaften 2014, 101, 661–670. [CrossRef] [PubMed]
178.
Bucekova, M.; Jardekova, L.; Juricova, V.; Bugarova, V.; Marco, G.D.; Gismondi, A.; Leonardi, D.; Farkasovska, J.; Godocikova, J.;
Laho, M.; et al. Antibacterial Activity of Different Blossom Honeys: New Findings. Molecules
2019
, 24, 1573. [CrossRef] [PubMed]
179.
Bucekova, M.; Buriova, M.; Pekarik, L.; Majtan, V.; Majtan, J. Phytochemicals-mediated production of hydrogen peroxide is
crucial for high antibacterial activity of honeydew honey. Sci. Rep. 2018, 8, 9061. [CrossRef] [PubMed]
180.
Zhao, Y.; Du, X.; Jiang, L.; Luo, H.; Wang, F.; Wang, J.; Qiu, L.; Liu, L.; Liu, X.; Wang, X.; et al. Glucose Oxidase-Loaded
Antimicrobial Peptide Hydrogels: Potential Dressings for Diabetic Wound. J. Nanosci. Nanotechnol.
2020
, 20, 2087–2094.
[CrossRef]
181.
Chen, Y.; Li, Y.; Yang, X.; Cao, Z.; Nie, H.; Bian, Y.; Yang, G. Glucose-triggered in situ forming keratin hydrogel for the treatment
of diabetic wounds. Acta Biomater. 2021, 125, 208–218. [CrossRef]
182.
Vasquez, J.M.; Idrees, A.; Carmagnola, I.; Sigen, A.; McMahon, S.; Marlinghaus, L.; Ciardelli, G.; Greiser, U.; Tai, H.; Wang, W.;
et al. In situ Forming Hyperbranched PEG—Thiolated Hyaluronic Acid Hydrogels with Honey-Mimetic Antibacterial Properties.
Front. Bioeng. Biotechnol. 2021, 9, 742135. [CrossRef]
183.
Davis, P.; Wood, L.; Wood, Z.; Eaton, A.; Wilkins, J. Clinical experience with a glucose oxidase-containing dressing on recalcitrant
wounds. J. Wound Care 2009, 18, 116–121. [CrossRef]
184.
Rashaan, Z.M.; Krijnen, P.; Kwa, K.A.; van Baar, M.E.; Breederveld, R.S.; van den Akker-van Marle, M.E. Long-term quality of life
and cost-effectiveness of treatment of partial thickness burns: A randomized controlled trial comparing enzyme alginogel vs
silver sulfadiazine (FLAM study). Wound Repair Regen. 2020, 28, 375–384. [CrossRef]
185.
Huang, T.; Yuan, B.; Jiang, W.; Ding, Y.; Jiang, L.; Rena, H.; Tang, J. Glucose oxidase and Fe
3
O
4
/TiO
2
/Ag
3
PO
4
co-embedded
biomimetic mineralization hydrogels as controllable ROS generators for accelerating diabetic wound healing. J. Mater. Chem. B
2021, 9, 6190–6200. [CrossRef]
186.
Zhang, S.; Yang, Z.; Hao, J.; Ding, F.; Li, Z.; Ren, X. Hollow nanosphere-doped bacterial cellulose and polypropylene wound
dressings: Biomimetic nanocatalyst mediated antibacterial therapy. Chem. Eng. J. 2022, 432, 134309. [CrossRef]
187.
Du, X.; Jia, B.; Wang, W.; Zhang, C.; Liu, X.; Qu, Y.; Zhao, M.; Li, W.; Yang, Y.; Li, Y.Q. pH-switchable nanozyme cascade catalysis:
A strategy for spatial-temporal modulation of pathological wound microenvironment to rescue stalled healing in diabetic ulcer. J.
Nanobiotechnol. 2022, 20, 12. [CrossRef] [PubMed]
188.
Simões, D.; Miguel, S.P.; Ribeiro, M.P.; Coutinho, P.; Mendonça, A.G.; Correia, I.J. Recent advances on antimicrobial wound
dressing: A review. Eur. J. Pharm. Biopharm. 2018, 127, 130–141. [CrossRef] [PubMed]
