Biotechnology Advances 65 (2023) 108153
Available online 11 April 2023
0734-9750/© 2023 The Author(s). Published by Elsevier Inc. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).
Research review paper
Biocatalysis for bioreneries: The case of dye-decolorizing peroxidases
Diogo Silva, Carolina F. Rodrigues, Constança Lorena, Patrícia T. Borges, Lígia O. Martins
Instituto de Tecnologia Química e Biol´
onio Xavier, Universidade NOVA de Lisboa, Av. da República, 2780-157 Oeiras, Portugal
Dye-decolorizing Peroxidases (DyPs) are heme-containing enzymes in fungi and bacteria that catalyze the
reduction of hydrogen peroxide to water with concomitant oxidation of various substrates, including anthra-
quinone dyes, lignin-related phenolic and non-phenolic compounds, and metal ions. Investigation of DyPs has
shed new light on peroxidases, one of the most extensively studied families of oxidoreductases; still, details of
their microbial physiological role and catalytic mechanisms remain to be fully disclosed. They display a
distinctive ferredoxin-like fold encompassing anti-parallel β-sheets and
-helices, and long conserved loops
surround the heme pocket with a role in catalysis and stability. A tunnel routes H
to the heme pocket,
whereas binding sites for the reducing substrates are in cavities near the heme or close to distal aromatic residues
at the surface. Variations in reactions, the role of catalytic residues, and mechanisms were observed among
different classes of DyP. They were hypothetically related to the presence or absence of distal H
O molecules in
the heme pocket. The engineering of DyPs for improved properties directed their biotechnological applications,
primarily centered on treating textile efuents and degradation of other hazardous pollutants, to elds such as
biosensors and valorization of lignin, the most abundant renewable aromatic polymer. In this review, we track
recent research contributions that furthered our understanding of the activity, stability, and structural properties
of DyPs and their biotechnological applications. Overall, the study of DyP-type peroxidases has signicant im-
plications for environmental sustainability and the development of new bio-based products and materials with
improved end-of-life options via biodegradation and chemical recyclability, fostering the transition to a sus-
tainable bio-based industry in the circular economy realm.
Lignocellulose bioreneries are promising alternative sources of
renewable bulk and ne chemicals, materials, energy, and fuels for
sustainable development. The pulp and paper industry produces about
50 million tons of lignin annually, but most are burned for power; only 1
million tons reach the chemicals market (Sun et al., 2018). Lignin is the
second most abundant carbon source and the largest source of natural
phenolic compounds on Earth. It is currently used for low- and medium-
value applications (e.g., binding and dispersing agents), with energy
capturing about 90% of the market. However, lignin is expected to
become an alternative feedstock for aromatic chemicals, most currently
derived from the BTX process in the petroleum industry (Chen and Wan,
2017; Hamalainen et al., 2018). The recent implementation of novel
strategies for lignin depolymerization involving electrochemistry, pho-
tocatalysis, heterogeneous catalysis, and ionic liquids allowed to derive
of well-dened compounds from lignin in acceptable quantities and
implement a lignin-derived platform of chemicals (Fig. 1) (Renders
et al., 2017; Sun et al., 2018; Van den Bosch et al., 2018). The challenge
at present is the set-up of atom-economic and waste-free processes that
allow the full implementation of lignin as sustainable starting material
for the production of drop-in chemicals, polymers, or emerging func-
tional materials (Llevot et al., 2016; Natte et al., 2020; Runeberg et al.,
2019; von Vacano et al., 2022; Zalesak et al., 2019).
Biocatalysis offers an environmentally friendly tool for lignin valo-
rization. In nature, fungal laccases and peroxidases, such as lignin (LiP),
manganese (MnP), and versatile (VP) peroxidases, are well recognized
for playing a critical role in lignin depolymerization (Fernandez-Fueyo
et al., 2012; Floudas et al., 2012; Hammel and Cullen, 2008; Ruiz-
Duenas and Martinez, 2009), whereas, in comparison, bacterial systems
are far less understood (Kamimura et al., 2019; Kamimura et al., 2017).
Microbial lignin conversion comprises peroxidases and laccases that
randomly depolymerize lignin into small fragments, an array of auxil-
iary enzymes (aryl alcohol oxidases, pyranose oxidase, galactose
* Corresponding author.
E-mail address: email@example.com (L.O. Martins).
Contents lists available at ScienceDirect
journal homepage: www.elsevier.com/locate/biotechadv
Received 30 January 2023; Received in revised form 6 April 2023; Accepted 9 April 2023
Biotechnology Advances 65 (2023) 108153
oxidase, vanillyl alcohol oxidase) that produce compounds that are
substrates (e.g., hydrogen peroxide required by peroxidases), metabo-
lites (acids) that stabilize/chelate enzyme-generated radicals, and other
enzymes that indirectly participate by generating reactive species that
Dye-decolorizing peroxidases (DyPs) are a family of bacterial and
fungal heme peroxidases that show attractive catalytic properties for
biotechnological purposes. In the last two decades, numerous microbial
DyP-like peroxidases have been identied and characterized (Linde
et al., 2015b; Singh and Eltis, 2015; Sugano and Yoshida, 2021; Yoshida
and Sugano, 2015). DyPs have received signicant attention due to their
potential for use in the treatment of textile efuent and for the degra-
dation of other hazardous pollutants in the food and cosmetic industries,
where they can be used to remove the colour from natural food in-
gredients or cosmetics. Additionally, these enzymes have potential ap-
plications in the lignocellulose bioreneries; DyPs are versatile enzymes
that can oxidize kraft lignin, syringyl, and guaiacyl-type phenolics and
several lignin-phenolic models (Linde et al., 2015b; Silva et al., 2022)
and non-phenolic methoxylated aromatics (e.g., veratryl alcohol), and
metal ions (Kimani et al., 2021; Min et al., 2015; Santos et al., 2014).
Their eco-physiological role seems diverse; DyPs are present in the
genome of several lignin-degrading wood decay basidiomycetes (Flou-
das et al., 2012; Ruiz-Duenas et al., 2013) and are widely expressed
among lignin-degrading fungi from forest habitats (Kellner et al., 2014).
Fungal DyPs can oxidize nonphenolic model dimers, representatives of
the main moiety of the lignin macromolecule, even if at signicantly
lower catalytic efciencies than those observed for LiPs and VPs (Linde
et al., 2021), whereas bacterial DyPs act synergistically by oxidizing less
recalcitrant phenolic dimeric and simple aromatics, catabolic products
from the fungal enzymes. The oxidation of Mn(II) to diffusible Mn(III) an
essential aspect of the synergistic microbial attack on lignin in nature,
characteristic of basidiomycete MnPs and VPs (Ruiz-Duenas et al.,
2009), has also been reported for bacterial and fungal DyPs (Ahmad
et al., 2011; Fernandez-Fueyo et al., 2015; Roberts et al., 2011b; Santos
et al., 2014; Singh et al., 2012).
A possible role as a virulence factor in accelerating plant parasitism
was suggested since alizarin is an anti-fungicide anthraquinone pro-
duced by plants which is degraded by the DyP of the plant pathogen
Bjerkandera adusta Dec1(Sugawara et al., 2019). In Streptomyces lividans,
a morphogenetic pathway involves the cooperative action of a DyP-type
peroxidase, DtpA, the radical copper oxidase GlxA, and a cellulose
synthase-like protein, CslA, trigging the developmental switch between
vegetative mycelium and aerial hyphae (Petrus et al., 2016). DyP acti-
vates the GlxA enzyme, creating a cross-linked tyrosyl-cysteine cofactor
essential for enzymatic activity (Chaplin et al., 2015). A few DyPs were
identied as cargo of encapsulins, prokaryotic proteinaceous nano-
compartments with a stabilizing role by increasing the local concen-
tration of functionally related enzymes and conning unstable reaction
intermediates (Sutter et al., 2008). DyPs and encapsulins are encoded in
a two-gene operon indicating a tight translational coupling (Putri et al.,
2017; Sutter et al., 2008). The DypB from Rhodococcus jostii, a widely
distributed bacterium able to degrade and transform polychlorinated
biphenyls and other aromatics, assembles in vitro with encapsulin
(Rahmanpour and Bugg, 2013). One possibility is that this nano-
compartment can disassemble and localize DypB on the surface of
lignocellulose, generating short-lived oxidants (Mn(II) or phenolic rad-
icals) that diffuse and decompose the bulky recalcitrant lignin polymer.
In the human pathogens Mycobacterium tuberculosis and Mycobacterium
smegmatis, a DyP-encapsulin association is putatively involved in pro-
tecting cells against oxidative stress at low pH, conditions that resemble
the host lysosome and helping to evade the host immune system (Con-
treras et al., 2014; Lien et al., 2021). Recently, the structure of the
M. smegmatis DyP-loaded encapsulin was solved by cryo-EM at 3.7 Å
resolution (PDB 7BOK) (Tang et al., 2021). The encapsulin comprises an
icosahedral shell that protects a dodecameric DyP cargo. It is believed
that understanding the assembly and physiological role of the encap-
sulin systems can provide a rational framework for drug design.
2. The Hallmark features of DyP-type peroxidases
2.1. Structural insights
DyPs overall fold is signicantly different from the
structures present in the peroxidase-catalase superfamily with a role in
lignin decomposition, such as LiP, MnP, and VP (Hammel and Cullen,
2008; Lundell et al., 2010; Martinez et al., 2009; Veitch, 2004; Zamocky
et al., 2008). DyP belongs to the dimeric
+β barrel superfamily (Fig. 2)
along with chlorite dismutases (Clds), involved in chlorite detoxica-
tion, and other related proteins such as Escherichia coli EfeB, forming the
CDE superfamily with a common evolutionary origin (Hofbauer et al.,
Fig. 1. Lignin-derived chemical platform. The depolymerization of lignin leads to a set of aromatic chemicals that can be used as DyPs substrates. These molecules
can be functionalized, resulting in added-value compounds such as drugs, food additives, and polymers, and defunctionalized, resulting in bulk chemicals such as
benzene, toluene, and xylene. Adapted from (Sun et al., 2018).
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
2021). The heme pocket is buried within the ferredoxin-like fold. It is
lined by the conserved proximal His, and at the distal site, by a nega-
tively charged residue, usually an aspartate and a positively charged
arginine. Bacterial DyPs are mainly assembled in dimers (Li et al., 2012;
Liu et al., 2011; Rodrigues et al., 2021), whereas known fungal DyPs are
exclusively monomers (Chen and Li, 2016; Johjima et al., 2003; Sugano
et al., 2007; Zitare et al., 2021). High-order (hexameric) oligomerization
states were observed in Bacteroides thetaiotaomicron BtDyP (Zubieta
et al., 2007), R. jostii RHA1 DyPB (Roberts et al., 2011a), and Pseudo-
monas putida MET94 PpDyP (Borges et al., 2022).
A classication based on structure-based sequence alignments
(Yoshida and Sugano, 2015) was proposed: Classes P (primitive) and I
(intermediate), and Class V (advanced). Class P members have the
smallest molecular size among the three classes, with members of class V
having the largest size and higher catalytic efciencies (Catucci et al.,
2020). DyPs from class I harbor a twin-arginine translocation (TAT)
system, and BsDyP (also known as YwnN) from Bacillus subtilis is a well-
recognized extracellularly secreted enzyme by the TAT system (van der
Ploeg et al., 2011). The catalytic efciency of DyPs varies depending on
the classes; with class V exhibiting the highest catalytic efciencies, for
example, to oxidize anthraquinone dyes, the k
is between 10
followed by Class I, ranging from 10
Class P DyPs ranging from 10
Recently, a new comprehensive phylogenetic analysis was performed
using sequences in the InterPro DyP-type Peroxidases family
(IPR006314) (Yoshida and Sugano, 2023). Although DyPs are located in
Fig. 2. Structural domains of DyP-type peroxi-
dases. Schematic representation of a subunit from
+β barrel superfamily, consisting of
two ferredoxin folds where the N-, the C-terminal
and linker are colored in green, purple, and yellow
(a). Cartoon representation of a ferredoxin-like
2 colored in red and pink,
respectively, and β1, β2, β3, and β4 colored in
dark green, dark blue, light green, and cyan (b).
Cartoon representation of Bacillus subtilis BsDyP
monomer (c). Adapted from (Hofbauer et al.,
2021). (For interpretation of the references to
colour in this gure legend, the reader is referred
to the web version of this article.)
Fig. 3. Tunnels and cavities in DyPs from different classes. Loops, molecular tunnels, and cavities that give access to the heme pocket in Class P K. pneumoniae DyP
(PDB 6FKS) (a, b), class I B. subtilis BsDyP (PDB 7PKX) (c,d), and class V A. auricula-judae DyP (PDB 4AU9) (e,f). The regions lining the cavities are colored light pink
(loop 1), green (loop 2), dark red (small
-helix), and orange (loop 3). The tunnels (T) and cavities (C) corresponding to panels (a,c,e) are represented as gray
accessible surface area in panels (b,d,f), respectively. From (Rodrigues et al., 2021). (For interpretation of the references to colour in this gure legend, the reader is
referred to the web version of this article.)
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
most clades of the phylogenetic tree, there are new unexplored sub-
classes, P1, P2, I2, V4, and I1; the subclasses I1 and P1 include members
from Archaea and of Metazoa origins. Interestingly, the authors found
extra domains in unexplored classes, such as the pyruvate formate lyase
domain in subclass P2 and the cytochrome P450 domain in subclass V4.
The conserved catalytic distal arginine residue is present in all sub-
classes. In contrast, the distal aspartate residue is substituted with
different residues, alanine, serine, asparagine, and glutamate in DyPs
from I1, I2, I3, and P2 subclasses, revealing a less crucial role.
2.2. Active site structure and catalytic mechanism
Three structurally conserved loops delimiting the heme pocket pro-
mote local exibility around the DyPs heme pocket and modulate en-
zymes’ activity, specicity, and stability (Borges et al., 2022; Rodrigues
et al., 2021). They dene the access of the solvent to the heme through
two main molecular pathways (Fig. 3): one tunnel giving access of H
to the distal side (Strittmatter et al., 2013a; Yoshida and Sugano, 2015)
and one cavity that allows electron transfer from reduced substrates to
the heme propionate (Catucci et al., 2020; Perez-Boada et al., 2005;
Pfanzagl et al., 2019). In X-ray crystal structures of DyPs from class V,
this cavity seems to be occluded by longer (10 to 30 residues) loops
(Fig. 3 e,f); however, the expected high exibility in this region may
promote solvent access and oxidation of small substrates at the heme
The catalytic mechanism have been investigated in several DyPs
along with the identication of intermediates Compound I (Cpd I) and
Compound II (Cpd II) and the determination of rate constants (Ahmad
et al., 2011; Brissos et al., 2017; Chen et al., 2015; Lucic et al., 2020a;
Lucic et al., 2020b; Lucic et al., 2021; Lucic et al., 2022; Mendes et al.,
2015a; Pfanzagl et al., 2018; Shrestha et al., 2017; Shrestha et al., 2021;
Sugano et al., 2007). Interestingly, the intermediate precursor of Cpd I,
Compound 0, was readily observed in the reaction of several DyPs
(Brissos et al., 2017; Mendes et al., 2015b; Pfanzagl et al., 2018). Which
residue, Asp or Arg, is selected to facilitate proton transfer and O
ssion remains a topic of debate. The distal aspartate has been reported
to have an essential catalytic role in some DyPs (Chen et al., 2015; Lucic
et al., 2020a; Pfanzagl et al., 2019; Shrestha et al., 2017; Sugano et al.,
2007). In other DyPs, in contrast, the distal arginine is reportedly the
critical residue in Cpd I formation (Lucic et al., 2020b; Mendes et al.,
2015a; Shrestha et al., 2021; Singh et al., 2012). These ndings suggest
functional diversity within this family despite close structural resem-
blance. It was suggested that the presence or absence of a distal bound
water molecule in the heme pocket, a “dry” or “wet” distal site, affects
the spatial stereochemistry of the active site and can determine which of
the two residues, Asp or Arg, facilitate Cpd I formation (Lucic et al.,
2021). The role of a water molecule in the distal heme pocket has been
previously investigated in classical peroxidases, with the hypothesis that
a “dry” (non-H
O-containing) and “wet” (H
O-containing) heme pocket
may promote different reactions (Jones, 2001). More recently, it was
reported that catalytic arginine allows Cpd I formation, whether the
distal heme site is ‘wet’ or ‘dry’ (Lucic et al., 2022). Interestingly, a ‘wet’
site leads to a one-electron reduction pathway, i.e., a faster reduction of
Cpd I resulting in a high concentration of Cpd II, which is then reduced
to the ferric state (Fig. 4, left) (Lucic et al., 2022). This pathway is
compatible with the catalytic mechanism described in R. jostii DypB
(Shrestha et al., 2021), Thermomonospora curvata TcDyP (Chen et al.,
2015), B. subtilis BsDyP (Mendes et al., 2015b) and P. putida PpDyP
(Brissos et al., 2017; Mendes et al., 2015a). Instead, a ‘dry’ distal pocket
favors the Cpd I two-electron reduction to a protonated Cpd II (Fe(IV) −
OH) that is promptly reduced to the ferric state (Fig. 4 right), similar to
the mechanistic pathway observed in the P-class ElDyP from Enterobacter
lignolyticus and Dictyostelium discoideum DyPA (Rai et al., 2021; Shrestha
et al., 2017).
2.3. Long-range electron transfer
DyP-type peroxidases can oxidize bulky substrates at distal sites.
Electron transfer occurs via surface-exposed oxidation sites, usually
redox-active tryptophan and tyrosine residues, to the heme through
long-range electron transfer (LRET) pathways (Gray and Winkler, 2021;
Linde et al., 2015a; Linde et al., 2015b; Shrestha et al., 2016; Sugano and
Yoshida, 2021; Uchida et al., 2015). These distal oxidation sites were
previously identied in fungal LiP, VP, and MnP (Gelpke et al., 2002;
Fig. 4. Proposed catalytic mechanisms for DyPs. Left: In a wet site, the reduction of Compound I has no deuterium effect, and yields highly populated Compound II,
which is subsequently reduced to the ferric form. Right: Compound I reduction proceeds through a low Compound II concentration in a dry distal heme site.
Reduction is coupled with proton uptake, and the resulting protonated Compound II (Fe(IV)–OH) is reduced to the ferric state. Adapted from (Lucic et al., 2022).
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
Ruiz-Duenas et al., 2009; Ruiz-Duenas et al., 2007; Saez-Jimenez et al.,
2016). The DyPs subclasses P1–4 could contain different Tyr/Trp radical
sites, while it is suggested that subclasses I1–4 and V1–4 uses conserved
tryptophan residues to oxidize bulky substrates (Yoshida and Sugano,
Generally, the surface substrate binding sites in DyPs seem to fall
under two categories: (i) radical-forming aromatic residues that allow
for the oxidation of, e.g., veratryl alcohol, bulky substrates, and dyes,
and (ii) negatively charged patches involved in the oxidation of Mn(II).
Radical sites in surface-exposed aromatic residues and LRET pathways
were predicted in, e.g., Auricularia auricula-judae DyP (AauDyP) (Baratto
et al., 2015; Linde et al., 2015a; Strittmatter et al., 2013a; Strittmatter
et al., 2015; Strittmatter et al., 2013b), S. lividans DtpA (Chaplin et al.,
2019), Klebsiella pneumoniae DyP (KpDyP) (Nys et al., 2021), and
D. discoideum DyPA (Rai et al., 2021). Surface carboxylic residues
constitute the binding site for Mn(II) and were identied in R. jostii
RHA1 DypB (Roberts et al., 2011a), Amycolatopsis sp. 75iv2 DyP2
(Brown et al., 2011) and Pleurotus ostreatus PosDyP4 (Fig. 5) (Fern´
Fueyo et al., 2018).
3. Exploring the industrial potential of DyPs
3.1. Engineering for improved properties
The large substrate scope of DyP-type peroxidases makes these en-
zymes an excellent starting point for generating interesting biocatalysts
for biorenery elds. Generating tailor-made enzymes represents a
procient way to design efcient and robust biocatalysts to support their
industrial application’s major limiting factors: thermostability, resis-
tance to organic solvents, extremes of pH (acid or alkaline), and in-
hibitors. A summary of reported engineering studies is in Table 1.
Rational design and gene recombination was followed in Coprinus
cinereus DyP to resist laundry detergent conditions and improve overall
enzyme stability and resistance towards hydrogen peroxide (Cherry
et al., 1999). Similarly, the replacement of sensitive methionine residues
in the Anabaena sp. DyP heme cavity resulted in ~10-fold higher H
Fig. 5. Long-range electron transfer pathways in PosDyP4.The acidic residues
part of the Mn(II) site are represented as sticks with carbon atoms colored in
light pink. The aromatic residues Y339 and W405, T338, and H334, are rep-
resented as sticks with carbon atoms colored in cyan. The LRET paths from the
Mn(II) and aromatics oxidation are shown as arrows. The heme is shown as
sticks with carbon, oxygen, and nitrogen atoms colored yellow, red, and blue.
The manganese and iron ions are shown as purple and orange spheres,
respectively. Adapted from (Fernandez-Fueyo et al., 2018). (For interpretation
of the references to colour in this gure legend, the reader is referred to the web
version of this article.)
Tailor enzymes using enzyme engineering for improved properties.
DyP-type peroxidase Enzyme
R. jostii RHA1
DyPB Rational Design
activity for Mn
Singh et al.
up-shift of pH
optimum to pH
Brissos et al.
Barbosa et al.
PfDyP Rational Design
efciency for 2-
and Mn (II)
et al. (2019)
VcDyP Rational Design
Optimum pH for
to pH 7
Uchida et al.
Dyp1B Rational Design
et al. (2023)
~ 10-fold higher
Rodrigues et al.
ABTS and higher
stability at 40 ◦C
car et al.
at pH 10.5
and 50 ◦C
Cherry et al.
AnaPX Rational Design 8-fold enhanced
Ogola et al.
Linde et al.
Higher yields of
Alessa et al.
Krahe et al.
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
resistance (Ogola et al., 2010). A different strategy was followed using
P. ostreatus DyP4 for higher protein yields, improved catalytic efciency,
and enhanced hydrogen peroxide resistance upon fusion with the E. coli
osmotically-inducible protein Y for extracellular secretion and engi-
neered by directed evolution and saturation mutagenesis (Alessa et al.,
Lignin-oxidizing Pseudomonas uorescens Dyp1B was expressed in
the periplasm of P. putida KT2440, using a TAT fusion construct, leading
to enhanced activity for oxidation of 2,4-dichlorophenol (DCP) and
polymeric lignin substrates (Ehibhatiomhan et al., 2023). Moreover,
amino acid replacements at the manganese ion binding site of
P. uorescens DyP1 resulted in increased k
compared with wild-type Dyp1B. This study also identies a correlation
between the efciency for Mn
oxidation and the activity of low mo-
lecular weight phenols released from lignin, and alkali Kraft lignin. The
authors claim that polymeric lignin oxidation by DyP-type peroxidases is
performed primarily via the formation of Mn
, which acts as a diffus-
ible oxidant to attack polymeric lignin. Site-saturation mutagenesis was
also used to change the active site of P. uorescens Dyp1B, resulting in
variants capable of ~10-fold higher efciency for 2-chlorophenol and
Mn(II) and exhibiting higher thermostability (Rahman Pour et al.,
2019). R. jostii RHA1 DyPB displayed 80-fold improved activity for Mn
(II) after the replacement to alanine of an asparagine found within a
pocket of negatively charged residues at the heme edge (Singh et al.,
2013). Higher activity and stability of variants of Pleurotus sapidus Psa-
POX, which were traced to a single mutation, were selected from a
basidiospore-derived monokaryons library (Krahe et al., 2021). In
contrast, an engineered BsDyP variant after three rounds of directed
evolution revealed ~10-fold higher activity for 2,6-dimethoxyphenol
(DMP) but decreased enzyme’s overall stability by 2 kcal mol
(Rodrigues et al., 2021). Furthermore, the evolved variant exhibits
slightly reduced thermal stability than the wild-type indicating a trade-
off between functionality and stability. The analysis of X-ray structures
showed that one loop in the proximal side of the heme pocket becomes
more exible in the evolved variant, the size of the active site cavity is
increased, as well as the width of its mouth, resulting in an enhanced
exposure of the heme to solvent. These conformational changes were
pointed out to facilitate electron transfer from the substrate to the
enzyme but to decrease thermodynamic and kinetic stability.
As previously mentioned, DyPs can function as physiologic cargo
proteins for encapsulin, a bacterial nanocompartment protein. Taking
advantage of this, Lonˇ
car and colleagues packaged SviDyP from Sac-
charomonospora viridis DSM43017 with the encapsulin from meso-
thermophile bacterium Mycolicibacterium hassiacum (EncMh) (Lonˇ
et al., 2020). They showed that the packaged DyP showed signicantly
higher operational stability.
The biosynthesis of molecules containing stereogenic centers has a
high industrial interest; for example, chiral sulfoxides, containing a
sulfur atom as a chiral center, have a wide range of applications from
chiral auxiliaries to pharmaceuticals, e.g., S-omeprazole is a multi-
billion-dollar drug produced by a modied cyclohexanone mono-
oxygenase. A-type TfuDyP from Thermobida fusca reportedly converts
aromatic sulde to chiral sulfoxide products (van Bloois et al., 2010). A
structural-based design was followed to engineer the heme pocket of
AauDyP to perform sulfoxidation better (Linde et al., 2016); the entrance
to the heme was enlarged for better accommodation of sulde sub-
strates, which became positioned closer to the reactive cofactor with
more favorable interaction energies.
Taking myoglobin (Mb) as a starting point, Lin and colleagues
engineered a so-called articial DyP by inserting a Tyr/Trp chain near
the heme pocket, which resulted in ~100-fold enhanced dye-
decolorizing activity when compared to Mb (Li et al., 2017); An addi-
tional round of rational design resulted in an articial enzyme with
catalytic efciency for Reactive Blue 19 (1.2 ×10
to native DyPs (Zhang et al., 2019). These enzymes could oxidize
phenolic and aromatic amine substrates, as well as Kraft lignin and the
model lignin dimer guaiacylglycerol-β-guaiacyl ether (GGE), at mildly
acidic pH (Guo et al., 2021). Likewise, a cytochrome c was converted
into a DyP-type peroxidase after the replacement of residues at the
protein surface; the articial enzymes showed improved activity to-
wards Reactive Blue 19 (Omura et al., 2022).
The optimal acidic pH (3–4) for enzymatic activity in DyPs repre-
sents, for some applications, a severe drawback. Uchida et al. designed,
using site-directed mutagenesis, an articial Vibrio cholerae VcDyP with
the optimal pH shifted from 4.5 to 7 (Uchida et al., 2021), showing that
DyPs pH dependence is mainly affected by hydrogen bonds between His
and Asp residues placed at the heme second shell. Directed evolution
was used to improve the efciency of the bacterial PpDyP from Pseu-
domonas putida MET94 for phenolic compounds. Three rounds of
random mutagenesis by error-prone PCR followed by high-throughput
screening allow identifying the 6E10 variant showing a 100-fold
enhanced catalytic efciency (k
) for DMP, similar to those
exhibited by fungal lignin peroxidases (~10
). The evolved
variant showed improved efciency for several syringyl-type phenolics,
guaiacol, aromatic amines, and the lignin phenolic model dimer, GGE.
Importantly, variant 6E10 displayed optimal pH at 8.5, an upshift of 4
units compared to the wild-type, showed resistance to hydrogen
peroxide inactivation and was produced at 2-fold higher yields (Brissos
et al., 2017). Constant-pH MD simulations revealed a more positively
charged microenvironment near the heme pocket of variant 6E10
(Borges et al., 2022).
3.2. Biotechnological applications
DyPs display a vast repertoire of biotechnological applications
related to their versatility and multi-functionality (Table 2). Azo and
anthraquinone dyes are stable xenobiotics and abundant chemicals in
dye-contaminated wastewater that must be treated before being dis-
charged into the environment (Mendes et al., 2011; Rawat et al., 2016).
DyPs oxidize a wide range of synthetic dyes and are obvious candidates
for the bioremediation of dye-containing wastewaters from textile,
cosmetic, food, feed, and pharmaceutical industries, among others
(Habib et al., 2019; Ogola et al., 2009; Sugano et al., 2009; Sugawara
et al., 2017; Uchida et al., 2015). Furthermore, the identied interme-
diate of the decolorization of RB19 by the TfuDyP gene from T. fusca
inhibited the growth of B. subtilis (Pi et al., 2022). Enzymatic decolor-
ization methods (with laccases, azoreductases, and other oxidoreduc-
tases) are highly sought because, unlike chemical catalysts, enzymes
successfully convert specic complex chemical structures under mild
environmental conditions with high efciency (Mendes et al., 2015c).
DyPs are potentially interesting for developing H
devices for sensing and degrading substrates of interest, for example, in
biomedical, environmental, and food industry elds or for ecological
degradation of persistent dye pollutants. PpDyP, electronically coupled
to Ag electrodes, was shown to preserve its structural integrity and
catalytic efciency (Sezer et al., 2012). More recently, P. putida MET94
PpDyP and variants generated by directed evolution were investigated in
the design of biosensors for H
detection and showed superior sta-
bility, sensitivity, and shorter response times compared to H
sensors reported in the literature, making them excellent candidates for
developing biosensors in disposable and single-use congurations for
various biological applications (Barbosa et al., 2020). In additional
work, the potential use of CboDyP, ScoDyP, and TfuDyP enzymes as
biosensors after immobilization on biocompatible silver electrodes
and functionalization with alkanethiols were also tested but showed less
promising results (Zuccarello et al., 2021). Interestingly, PpDyP and
B. subtilis BsDyP form long-lived oxyferryl species by applying a reduc-
tive electrode potential under aerobic conditions that can oxidize ABTS
over a broad pH range with similar efciencies (Scocozza et al., 2021).
These promising ndings pave the way for designing DyP-based elec-
trocatalytic reactors that operate in an extended pH range without
harmful reagents such as H
. The use of functional fused biocatalysts
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
to bacterial SviDyP, such as alditol oxidase (HotAldO) and chitooligo-
saccharide oxidase (ChitO) couple the oxidase-peroxidase activity,
where the oxidase originates hydrogen peroxide that fuels the peroxi-
dase. The potential of these fusion enzymes as sugar detectors/bio-
sensors was demonstrated: P-HotAldO can detect the alditols, xylitol,
and sorbitol, whereas P-ChitO can detect mono, di-, and oligosaccha-
rides, such as glucose, lactose, cellobiose, and maltose, with a detection
limit in the low-micromolar range (Colpa et al., 2017).
DyPs have promising applications in the food industry, namely in
producing divanillin, a avor whose production is challenging and in
high demand. Divanillin, was identied with LC-MS as the main product
resulting from the oxidation of vanillin mediated by TfuDyP (Lonˇ
et al., 2016). The fusion of the bacterial peroxidase SviDyP with eugenol
oxidase and 5-hydroxymethylfurfural oxidase in a cascade reaction that
includes the oxidation of vanillyl alcohol to vanillin by the oxidases
followed by dimerization into divanillin catalyzed by SviDyP (Colpa
et al., 2017). The one-pot reaction catalyzed by an enzymatic process
involving FtrDyP and the laccase FtrLcc, from the basidiomycete Funalia
trogii originated the high-value grapefruit avor constituent (+)-noot-
katone from (+)-valencene in the presence of Mn(II) and p-coumaric
acid (Kolwek et al., 2018). In the same line, the generation of aromatic
aldehydes, such as p-anisaldehyde after cleavage of trans-anethole, an
aryl alkene, is a crucial activity for synthesizing veratraldehyde, and
acetophenone, which is used as fragrance and avor in the food industry
by the DyP from the basidiomycete P. sapidus PsaPOX (Krahe et al.,
2020). In the area of animal feed, it was recently reported that DyPs
from B. subtilis SCK6 and Streptomyces thermocarboxydus 4129 degrade
different mycotoxins such as aatoxin B
), zearalenone and
deoxynivalenol in the presence of Mn(II), combined with laccases and
manganese peroxidases (Qin et al., 2021). These mycotoxins are widely
present in contaminated agricultural products and feed, resulting in
signicant economic loss and a health threat to humans and animals.
Biobleaching of kraft pulp is a process of high biotechnological in-
terest for the pulp and paper industry. Increased pulp brightness of
eucalyptus kraft pulp was observed upon the addition of SviDyP during
bleaching processes (Yu et al., 2014). Additionally, scanning electron
microscopy (SEM) highlighted the existence of local fractures and large
areas containing holes that generated a looser ber structure with the
lignin internal structure exposed. This structure breakdown helps the
permeation of bleaching agents and facilitates delignication to obtain
molecules of interest. The effect of DyPs encoded by Streptomyces coeli-
color A3(2) (ScDyPs) was assessed in pre-treated organosolv lignins from
Miscanthus x giganteus and aspen. Treatment with type-A DyPs,
ScDyP1A, and ScDyP2A resulted in lignin depolymerization, whereas
treatment with ScDyPB increased lignin’s molecular weight (Pupart
et al., 2023). The DyP2 from Amycolatopsis sp. 75iv2 alone or in com-
bination with small laccase transforms a variety of lignins, including
native and modied organosolv hardwood lignin, generating product
proles that depend on the enzyme used and the presence or absence of
redox mediators (Vuong et al., 2021).
Reported enzymatic activities ground the role of DyP in the valori-
zation of lignocellulose towards, e.g., β-aryl ether lignin (Ahmad et al.,
2011; Rai et al., 2021), guaiacylglycerol-β-guaiacyl ether (Brissos et al.,
2017; Brown et al., 2012), β-O-4 dilignols (Huang et al., 2017), and
veratryl glycerol-β-guaiacyl (Min et al., 2015). Furthermore, the oxida-
tion of phenolic and non-phenolic aromatic substrates such as guaiacol,
4-amino antipyrine, and pyrogallol (Ogola et al., 2009), veratryl alcohol
(Kimani et al., 2021) was reported. A new DyP discovered from the
Biotechnological applications of DyPs.
DyP-type peroxidase Enzyme Biotechnological Application Ref.
R. jostii RHA1 DyPB Wild-type Bioreneries: oxidation of β-aryl ether lignin models and Mn(II) Ahmad et al. (2011)
P. uorescens PfDyP Wild-type Oxidation of Mn (II) and Kraft lignin; bioreneries; lignin depolymerization to
Rahmanpour and Bugg,
P. putida MET94 PpDyP
Wild-type Decolorization of antraquinone and azo dyes, oxidation of phenolic lignin-derived
compounds, Mn(II) and Fe(II) Santos et al. (2014)
Bioreneries: conversion of aromatic amines to phenazines and bio-dyes; oxidation of
a wide array of syringyl, guaiacyl and hydroxibenzene phenolics; synthesis of
syringaresinol, divanillin and diapocynin dimers
Brissos et al. (2017);
Silva et al. (2022)
Wild-type and variant
Biosensors with higher sensitivity and shorter response times compared to other H
biosensors reported Barbosa et al. (2020)
S. coelicolor ScDyPs Wild-type Bioreneries: combined use of ScDyP1A and ScDyP2A to depolymerize lignin Pupart et al. (2023)
B. subtilis BsDyP Wild-type Decolorization of antraquinone and azo dyes, oxidation of phenolic lignin-derived
compounds, and Mn(II) Santos et al. (2014)
T. fusca TfuDyP Wild-type
Decolorization of Eosin Y, Acid Blue 129 and Cocetin dyes; bioremediation of
contaminated wastewater; divanillin production for food industry Lonˇ
car et al. (2016)
Production of anti-bacterial compounds via the oxidation of RB19, an anthraquinone
dye Pi et al. (2022)
S. viridis DSM43017 SviDyP
Wild-type Biobleaching for paper and pulp industry Yu et al. (2014)
Fusion proteins P-
HotAldO and P-ChitO Sugar detection Colpa et al. (2017)
T. curvata TcDyP Wild-type Lignin valorization: enzymatic activity towards stereospecic β-O-4 dilignolçs Huang et al. (2017)
Wild-type Degradation of mycotoxins B
), ZEN and DON in the presence of Mn(II);
decontamination of agricultural products Qin et al. (2021)
C. cinereus CIP Variants 972 and 974 Adaptation to laundry detergent environment Cherry et al. (1999)
Anabaena sp. AnaPX Wild-type
Decolorization of antraquinone and azo dyes; bioremediation of contaminated
wastewater; lignin degradation: active towards several aromatic substrates such as
guaiacol, 4-aminoantipyrine and pyrogallol
Ogola et al. (2009)
F. trogii FtrDyP Wild-type combined
with FtrLcc Food industry: grapefruit avor constituent (+)-nootkatone Kolwek et al. (2018)
Amycolatopsis sp. 75iv2
DyP 2 Wild-type Modication of native and organosolv hardwood lignin to different extents;Lignin
Brown et al. (2012);
Vuong et al. (2021)
P. sapidus PsaPOX Wild-type Fragrance and avor industry: generation of aromatic aldehydes; food industry:
β-carotene bleaching Krahe et al. (2020)
Agrobacterium sp. B1
Lignin valorization: 2-methoxyhydroquinone, hydroxyquinol, vanillin and vanillic
acid Rashid and Bugg, 2021
Xylaria grammica XgrDyP Wild-type Lignin depolymerization: Oxidation of DMP, veratryl alcohol and Mn (II);
Decolorization of Reactive Blue 5; Bioremediation Kimani et al. (2021)
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
Actinobacteria genome can oxidize sinapic and caffeic acid, 3-methylca-
techol, dopamine hydrochloride, and tannic acid at low temperatures
(25 ◦C) (Cordas et al., 2022). DyPs potential was tested in converting
lignin into low molecular weight aromatic products in the presence of
the accessory enzymes dihydrolipoamide dehydrogenases, peroxir-
edoxin, LigE, and arylsulfotransferase (Rashid and Bugg, 2021; Bugg
et al., 2020). This association prevented the repolymerization or
recondensation of the phenoxy radicals that originated from lignin
oxidation and promoted the formation of monomeric products,
including the monocyclic aromatics 2-methoxyhydroquinone, hydrox-
yquinol, vanillin, and vanillic acid. Recently, the catalytic potential of
the engineered PpDyP variant 6E10 was assessed for the oxidation of 24
syringyl, guaiacyl, and hydroxybenzene lignin-phenolic derivatives
(Silva et al., 2022). Variant 6E10 exhibited up to 100-fold higher
oxidation rates at pH 8 for all substrates. The main products of the re-
actions were dimeric isomers with molecular weights of (2 ×MW sub-
strate - 2H), and syringaresinol, divanillin, and diapocynin were among
the dimers identied. These are building blocks exploitable in medicinal
chemistry, food additives, and polymers. Combined, these results show
that DyPs has the potential to be included in the portfolio of enzymes
that can be used in processes of the functionalization and valorization of
lignin monomers in biorenery setups.
Current studies on biological lignin degradation and conversion have
opened exciting opportunities and a bright future for biological lignin
valorization. A deep understanding of the ligninolytic and auxiliary
enzymes is critical to design biocatalyst systems for value-added pro-
duction from lignin. DyP-type peroxidases have been identied as po-
tential catalysts in emerging lignocellulose bioreneries due to their
ability to oxidize and valorize lignin-related compounds, particularly
lignin-related phenolics. Rational design or direct evolution engineering
approaches are needed to engineer enzymes and pathways to improve
catalytic efciency and robustness further to enable cost-effective
complete conversion of heterogeneous lignin renewable feedstock.
Bacteria that are easy to culture, grow fast, and for which molecular
biology tools are well established can be valuable for exploring enzy-
matic lignin valorization strategies. The curtain behind DyP biological
functions and mechanisms has been opening in recent years, revealing
interesting properties. Still clearly, further studies are needed to have a
clear picture of how these enzymes operate. In this review, we thor-
oughly report recent reports on the investigation involving DyP-type
peroxidases, including those more on the protein science, such as
structural aspects and mechanisms of reaction, but also on the technical
side describing recent engineering attempts and applications to tackle
crucial global concerns.
Declaration of Competing Interest
This work was supported by the Fundaç˜
ao para a Ciˆ
encia e Tecno-
logia, Portugal, grants PTDC/BBBEBB/0122/2014, PTDC/BII-BBF/
29564/2017, EXPL/BIA-BQM/0473/2021, FCT 2022.02027.PTDC,
MOSTMICRO-ITQB (UIDB/04612/2020 and UIDP/04612/2020),
LS4FUTURE Associated Laboratory (LA/P/0087/2020). DS and CFR
acknowledge FCT Ph.D. Fellowships SFRH/BD/132702/2017 and UI/
Ahmad, M., Roberts, J.N., Hardiman, E.M., Singh, R., Eltis, L.D., Bugg, T.D., 2011.
Identication of DypB from Rhodococcus jostii RHA1 as a lignin peroxidase.
Biochemistry 50 (23), 5096–5107.
Alessa, A.H.A., Tee, K.L., Gonzalez-Perez, D., Omar Ali, H.E.M., Evans, C.A.,
Trevaskis, A., Xu, J.H., Wong, T.S., 2019. Accelerated directed evolution of dye-
decolorizing peroxidase using a bacterial extracellular protein secretion system
(BENNY). Bioresour. Bioproc. 6, 20.
Baratto, M.C., Sinicropi, A., Linde, D., Saez-Jimenez, V., Sorace, L., Ruiz-Duenas, F.J.,
Martinez, A.T., Basosi, R., Pogni, R., 2015. Redox-active sites in Auricularia auricula-
judae dye-decolorizing peroxidase and several directed variants: a multifrequency
EPR study. J. Phys. Chem. B 119, 13583–13592.
Barbosa, C., Silveira, C.M., Silva, D., Brissos, V., Hildebrandt, P., Martins, L.O.,
Todorovic, S., 2020. Immobilized dye-decolorizing peroxidase (DyP) and directed
evolution variants for hydrogen peroxide biosensing. Biosens. Bioelectron. 153,
van Bloois, E., Torres Pazmino, D.E., Winter, R.T., Fraaije, M.W., 2010. A robust and
extracellular heme-containing peroxidase from Thermobida fusca as prototype of a
bacterial peroxidase superfamily. Appl. Microniol. Biotechnol. 86 (5), 1419–1430.
Borges, P.T., Silva, D., Silva, T.F.D., Brissos, V., Canellas, M., Lucas, M.F., Masgrau, L.,
Melo, E.P., Machuqueiro, M., Frazao, C., Martins, L.O., 2022. Unveiling molecular
details behind improved activity at neutral to alkaline pH of an engineered DyP-type
peroxidase. Comp. Strut. Biotechnol. J. 20, 3899–3910.
Brissos, V., Tavares, D., Sousa, A.C., Robalo, M.P., Martins, L.O., 2017. Engineering a
bacterial DyP-type peroxidase for enhanced oxidation of lignin-related Phenolics at
alkaline pH. ACS Catal. 7, 3454–3465.
Brown, M.E., Walker, M.C., Nakashige, T.G., Iavarone, A.T., Chang, M.C., 2011.
Discovery and characterization of heme enzymes from unsequenced bacteria:
application to microbial lignin degradation. J. Am. Chem. Soc. 133, 18006–18009.
Brown, M.E., Barros, T., Chang, M.C., 2012. Identication and characterization of a
multifunctional dye peroxidase from a lignin-reactive bacterium. ACS Chem. Biol. 7,
Bugg, T.D.H., Williamson, J.J., Rashid, G.M.M., 2020. Bacterial enzymes for lignin
depolymerisation: new biocatalysts for generation of renewable chemicals from
biomass. Curr. Opin. Chem. Biol. 55, 26–33.
Catucci, G., Valetti, F., Sadeghi, S.J., Gilardi, G., 2020. Biochemical features of dye-
decolorizing peroxidases: current impact on lignin degradation. Biotechnol. Appl.
Biochem. 67, 751–759.
Chaplin, A.K., Petrus, M.L., Mangiameli, G., Hough, M.A., Svistunenko, D.A., Nicholls, P.,
Claessen, D., Vijgenboom, E., Worrall, J.A., 2015. GlxA is a new structural member
of the radical copper oxidase family and is required for glycan deposition at hyphal
tips and morphogenesis of Streptomyces lividans. Biochem. J. 469, 433–444.
Chaplin, A.K., Chicano, T.M., Hampshire, B.V., Wilson, M.T., Hough, M.A.,
Svistunenko, D.A., Worrall, J.A.R., 2019. An aromatic dyad motif in dye
decolourising peroxidases has implications for free radical formation and catalysis.
Chemistry 25, 6141–6153.
Chen, C., Li, T., 2016. Bacterial dye-decolorizing peroxidases: biochemical properties and
biotechnological opportunities. Phys. Sci. Rev. 1 (9).
Chen, C., Shrestha, R., Jia, K., Gao, P.F., Geisbrecht, B.V., Bossmann, S.H., Shi, J., Li, P.,
2015. Characterization of dye-decolorizing peroxidase (DyP) from Thermomonospora
curvata reveals unique catalytic properties of A-type DyPs. J. Biol. Chem. 290 (38),
Chen, Z., Wan, C., 2017. Biological valorization strategies for converting lignin into fuels
and chemicals. Renew. Sust. Energ. Rev. 73, 610–621.
Cherry, J.R., Lamsa, M.H., Scheider, P., Vind, J., Svendsen, A., Jones, A., Pedersen, A.H.,
1999. Directed evolution of a fungal peroxidase. Nat. Biotechnol. 17, 5.
Colpa, D.I., Lonˇ
car, N., Schmidt, M., Fraaije, M.W., 2017. Creating oxidase-peroxidase
fusion enzymes as a toolbox for Cascade reactions. ChemBioChem 18 (22),
Contreras, H., Joens, M.S., McMath, L.M., Le, V.P., Tullius, M.V., Kimmey, J.M.,
Bionghi, N., Horwitz, M.A., Fitzpatrick, J.A., Goulding, C.W., 2014. Characterization
of a Mycobacterium tuberculosis nanocompartment and its potential cargo proteins.
J. Biol. Chem. 289 (26), 18279–18289.
Cordas, C.M., Nguyen, G.S., Valerio, G.N., Jonsson, M., Sollner, K., Aune, I.H.,
Wentzel, A., Moura, J.J.G., 2022. Discovery and characterization of a novel Dyp-type
peroxidase from a marine actinobacterium isolated from Trondheim fjord, Norway.
J. Inorg. Biochem. 226, 111651.
Ehibhatiomhan, A.O., Pour, R.R., Farnaud, S., Bugg, T.D.H., Mendel-Williams, S., 2023.
Periplasmic expression of Pseudomonas uorescens peroxidase Dyp1B and site-
directed mutant Dyp1B enzymes enhances polymeric lignin degradation activity in
pseudomonas putida KT2440. Enzym. Microb. Technol. 162, 110147.
Fernandez-Fueyo, E., Ruiz-Duenas, F.J., Ferreira, P., Floudas, D., Hibbett, D.S.,
Canessa, P., Larrondo, L.F., James, T.Y., Seelenfreund, D., Lobos, S., Polanco, R.,
Tello, M., Honda, Y., Watanabe, T., Watanabe, T., Ryu, J.S., Kubicek, C.P.,
Schmoll, M., Gaskell, J., Hammel, K.E., St John, F.J., Vanden Wymelenberg, A.,
Sabat, G., Splinter BonDurant, S., Syed, K., Yadav, J.S., Doddapaneni, H.,
Subramanian, V., Lavin, J.L., Oguiza, J.A., Perez, G., Pisabarro, A.G., Ramirez, L.,
Santoyo, F., Master, E., Coutinho, P.M., Henrissat, B., Lombard, V., Magnuson, J.K.,
Kues, U., Hori, C., Igarashi, K., Samejima, M., Held, B.W., Barry, K.W., LaButti, K.M.,
Lapidus, A., Lindquist, E.A., Lucas, S.M., Riley, R., Salamov, A.A., Hoffmeister, D.,
Schwenk, D., Hadar, Y., Yarden, O., de Vries, R.P., Wiebenga, A., Stenlid, J.,
Eastwood, D., Grigoriev, I.V., Berka, R.M., Blanchette, R.A., Kersten, P., Martinez, A.
T., Vicuna, R., Cullen, D., 2012. Comparative genomics of Ceriporiopsis
subvermispora and Phanerochaete chrysosporium provide insight into selective
ligninolysis. Proc. Natl. Acad. Sci. U. S. A. 109 (14), 5458–5463.
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
Fernandez-Fueyo, E., Linde, D., Almendral, D., Lopez-Lucendo, M.F., Ruiz-Duenas, F.J.,
Martinez, A.T., 2015. Description of the rst fungal dye-decolorizing peroxidase
oxidizing manganese(II). Appl. Microbiol. Biotechnol. 99 (21), 8927–8942.
andez-Fueyo, E., Dav´
o-Siguero, I., Almendral, D., Linde, D., Baratto, M.C., Pogni, R.,
Romero, A., Guallar, V., Martínez, A.T., 2018. Description of a non-canonical Mn(II)-
oxidation site in peroxidases. ACS Catal. 8, 8386–8395.
Floudas, D., Binder, M., Riley, R., Barry, K., Blanchette, R.A., Henrissat, B., Martinez, A.
T., Otillar, R., Spatafora, J.W., Yadav, J.S., Aerts, A., Benoit, I., Boyd, A., Carlson, A.,
Copeland, A., Coutinho, P.M., de Vries, R.P., Ferreira, P., Findley, K., Foster, B.,
Gaskell, J., Glotzer, D., Gorecki, P., Heitman, J., Hesse, C., Hori, C., Igarashi, K.,
Jurgens, J.A., Kallen, N., Kersten, P., Kohler, A., Kues, U., Kumar, T.K., Kuo, A.,
LaButti, K., Larrondo, L.F., Lindquist, E., Ling, A., Lombard, V., Lucas, S., Lundell, T.,
Martin, R., McLaughlin, D.J., Morgenstern, I., Morin, E., Murat, C., Nagy, L.G.,
Nolan, M., Ohm, R.A., Patyshakuliyeva, A., Rokas, A., Ruiz-Duenas, F.J., Sabat, G.,
Salamov, A., Samejima, M., Schmutz, J., Slot, J.C., St John, F., Stenlid, J., Sun, H.,
Sun, S., Syed, K., Tsang, A., Wiebenga, A., Young, D., Pisabarro, A., Eastwood, D.C.,
Martin, F., Cullen, D., Grigoriev, I.V., Hibbett, D.S., 2012. The Paleozoic origin of
enzymatic lignin decomposition reconstructed from 31 fungal genomes. Science 336
Gelpke, M.D.S., Lee, J., Gold, M.H., 2002. Lignin peroxidase oxidation of Veratryl
alcohol: effects of the mutants H82A, Q222A, W171A, and F267L. Biochemistry 41,
Gray, H.B., Winkler, J.R., 2021. Functional and protective hole hopping in
metalloenzymes. Chem. Sci. 12, 13988–14003.
Guo, W.J., Xu, J.K., Liu, J.J., Lang, J.J., Gao, S.Q., Wen, G.B., Lin, Y.W., 2021.
Biotransformation of lignin by an articial Heme enzyme designed in myoglobin
with a covalently linked Heme group. Front. Bioeng. Biotechnol. 9, 664388.
Habib, M.H., Rozeboom, H.J., Fraaije, M.W., 2019. Characterization of a new DyP-
peroxidase from the Alkaliphilic Cellulomonad, Cellulomonas bogoriensis. Molecules
Hamalainen, V., Gronroos, T., Suonpaa, A., Hekkila, M.W., Romein, B., Ihalainen, P.,
Malandra, S., Birikh, K.R., 2018. Enzymatic processes to unlock the lignin value.
Front. Bioeng. Biotechnol. 6, 20.
Hammel, K.E., Cullen, D., 2008. Role of fungal peroxidases in biological ligninolysis.
Curr. Opin. Plant Biol. 11 (3), 349–355.
Hofbauer, S., Pfanzagl, V., Michlits, H., Schmidt, D., Obinger, C., Furtmuller, P.G., 2021.
Understanding molecular enzymology of porphyrin-binding alpha +beta barrel
proteins - one fold, multiple functions. Biochim. Biophys. Acta, Proteins Proteomics
Huang, G., Shrestha, R., Jia, K., Geisbrecht, B.V., Li, P., 2017. Enantioselective synthesis
of Dilignol model compounds and their Stereodiscrimination study with a dye-
decolorizing peroxidase. Org. Lett. 19, 3.
Johjima, T., Ohkuma, M., Kudo, T., 2003. Isolation and cDNA cloning of novel hydrogen
peroxide-dependent phenol oxidase from the basidiomycete Termitomyces
albuminosus. Appl. Microbiol. Biotechnol. 61, 220–225.
Jones, P., 2001. Roles of water in Heme peroxidase and catalase mechanisms. J. Biol.
Chem. 276, 13791–13796.
Kamimura, N., Takahashi, K., Mori, K., Araki, T., Fujita, M., Higuchi, Y., Masai, E., 2017.
Bacterial catabolism of lignin-derived aromatics: new ndings in a recent decade:
update on bacterial lignin catabolism. Environ. Microbiol. Rep. 9 (6), 679–705.
Kamimura, N., Sakamoto, S., Mitsuda, N., Masai, E., Kajita, S., 2019. Advances in
microbial lignin degradation and its applications. Curr. Opin. Biotechnol. 56,
Kellner, H., Luis, P., Pecyna, M.J., Barbi, F., Kapturska, D., Kruger, D., Zak, D.R.,
Marmeisse, R., Vandenbol, M., Hofrichter, M., 2014. Widespread occurrence of
expressed fungal secretory peroxidases in forest soils. PLoS One 9 (4), e95557.
Kimani, V., Ullrich, R., Buttner, E., Herzog, R., Kellner, H., Jehmlich, N., Hofrichter, M.,
Liers, C., 2021. First dye-decolorizing peroxidase from an Ascomycetous fungus
secreted by Xylaria grammica. Biomolecules 11 (9).
Kolwek, J., Behrens, C., Linke, D., Krings, U., Berger, R.G., 2018. Cell-free one-pot
conversion of (+)-valencene to (+)-nootkatone by a unique dye-decolorizing
peroxidase combined with a laccase from Funalia trogii. J. Ind. Microbiol. Biotechnol.
Krahe, N.K., Berger, R.G., Ersoy, F., 2020. A DyP-type peroxidase of Pleurotus sapidus
with alkene cleaving activity. MOLEFW 25.
Krahe, N.K., Berger, R.G., Witt, M., Zorn, H., Omarini, A.B., Ersoy, F., 2021.
Monokaryotic Pleurotus sapidus strains with intraspecic variability of an alkene
cleaving DyP-type peroxidase activity as a result of gene mutation and differential
gene expression. Int. J. Mol. Sci. 22.
Li, J., Liu, C., Li, B., Yuan, H., Yang, J., Zheng, B., 2012. Identication and molecular
characterization of a novel DyP-type peroxidase from Pseudomonas aeruginosa
PKE117. Appl. Biochem. Biotechnol. 166, 774–785.
Li, L.L., Yuan, H., Liao, F., He, B., Gao, S.Q., Wen, G.B., Tan, X., Lin, Y.W., 2017. Rational
design of articial dye-decolorizing peroxidases using myoglobin by engineering
Tyr/Trp in the heme center. Dalton Trans. 46 (34), 11230–11238.
Lien, K.A., Dinshaw, K., Nichols, R.J., Cassidy-Amstutz, C., Knight, M., Singh, R., Eltis, L.
D., Savage, D.F., Stanley, S.A., 2021. A nanocompartment system contributes to
defense against oxidative stress in Mycobacterium tuberculosis. E-Life 10.
Linde, D., Pogni, R., Canellas, M., Lucas, F., Guallar, V., Baratto, M.C., Sinicropi, A., Saez-
Jimenez, V., Coscolin, C., Romero, A., Medrano, F.J., Ruiz-Duenas, F.J., Martinez, A.
T., 2015a. Catalytic surface radical in dye-decolorizing peroxidase: a computational,
spectroscopic and site-directed mutagenesis study. Biochem. J. 466, 253–262.
Linde, D., Ruiz-Duenas, F.J., Fernandez-Fueyo, E., Guallar, V., Hammel, K.E., Pogni, R.,
Martinez, A.T., 2015b. Basidiomycete DyPs: genomic diversity, structural-functional
aspects, reaction mechanism and environmental signicance. Arch. Biochem.
Biophys. 574, 66–74.
Linde, D., Ca˜
nellas, M., Coscolín, C., Dav´
o-Siguero, I., Romero, A., Lucas, F., Ruiz-
nas, F.J., Guallar, V., Martínez, A.T., 2016. Asymmetric sulfoxidation by
engineering the heme pocket of a dye-decolorizing peroxidase. Catal. Sci. Technol. 6,
Linde, D., Ayuso-Fernandez, I., Laloux, M., Aguiar-Cervera, J.E., de Lacey, A.L., Ruiz-
Duenas, F.J., Martinez, A.T., 2021. Comparing Ligninolytic capabilities of bacterial
and fungal dye-decolorizing peroxidases and class-II peroxidase-catalases. Int. J.
Mol. Sci. 22 (5).
Liu, X., Du, Q., Wang, Z., Zhu, D., Huang, Y., Li, N., Wei, T., Xu, S., Gu, L., 2011. Crystal
structure and biochemical features of EfeB/YcdB from Escherichia coli O157: ASP235
plays divergent roles in different enzyme-catalyzed processes. J. Biol. Chem. 286,
Llevot, A., Grau, E., Carlotti, S., Grelier, S., Cramail, H., 2016. From lignin-derived
aromatic compounds to novel biobased polymers. Macromol. Rapid Commun. 37
car, N., Colpa, D.I., Fraaije, M.W., 2016. Exploring the biocatalytic potential of a
DyP-type peroxidase by proling the substrate acceptance of Thermobida fusca DyP
peroxidase. Tetrahedron 72, 7276–7281.
car, N., Rozeboom, H.J., Franken, L.E., Stuart, M.C.A., Fraaije, M.W., 2020. Structure
of a robust bacterial protein cage and its application as a versatile biocatalytic
platform through enzyme encapsulation. Biochem. Biophys. Res. Commun. 529,
Lucic, M., Chaplin, A.K., Moreno-Chicano, T., Dworkowski, F.S.N., Wilson, M.T.,
Svistunenko, D.A., Hough, M.A., Worrall, J.A.R., 2020a. A subtle structural change
in the distal haem pocket has a remarkable effect on tuning hydrogen peroxide
reactivity in dye decolourising peroxidases from Streptomyces lividans. Dalton Trans.
49 (5), 1620–1636.
Lucic, M., Svistunenko, D.A., Wilson, M.T., Chaplin, A.K., Davy, B., Ebrahim, A.,
Axford, D., Tosha, T., Sugimoto, H., Owada, S., Dworkowski, F.S.N., Tews, I.,
Owen, R.L., Hough, M.A., Worrall, J.A.R., 2020b. Serial femtosecond zero dose
crystallography captures a water-free distal Heme site in a dye-Decolorising
peroxidase to reveal a catalytic role for an arginine in Fe(IV) =O formation. Angew.
Chem. Int. Ed. Eng. 59 (48), 21656–21662.
Lucic, M., Wilson, M.T., Svistunenko, D.A., Owen, R.L., Hough, M.A., Worrall, J.A.R.,
2021. Aspartate or arginine? Validated redox state X-ray structures elucidate
mechanistic subtleties of Fe(IV) =O formation in bacterial dye-decolorizing
peroxidases. J. Biol. Inorg. Chem. 26 (7), 743–761.
Lucic, M., Wilson, M.T., Tosha, T., Sugimoto, H., Shilova, A., Axford, D., Owen, R.L.,
Hough, M.A., Worrall, J.A.R., 2022. Serial femtosecond crystallography reveals the
role of water in the one- or two-Electron redox chemistry of compound I in the
catalytic cycle of the B-type dye-decolorizing peroxidase DtpB. ACS Catal. 12 (21),
Lundell, T.K., Makela, M.R., Hilden, K., 2010. Lignin-modifying enzymes in lamentous
basidiomycetes–ecological, functional and phylogenetic review. J. Basic Microbiol.
Martinez, A.T., Ruiz-Duenas, F.J., Martinez, M.J., Del Rio, J.C., Gutierrez, A., 2009.
Enzymatic delignication of plant cell wall: from nature to mill. Curr. Opin.
Biotechnol. 20, 348–357.
Mendes, S., Farinha, A., Ramos, C.G., Leitao, J.H., Viegas, C.A., Martins, L.O., 2011.
Synergistic action of azoreductase and laccase leads to maximal decolourization and
detoxication of model dye-containing wastewaters. Bioresour. Technol. 102 (21),
Mendes, S., Brissos, V., Gabriel, A., Catarino, T., Turner, D.L., Todorovic, S., Martins, L.
O., 2015a. An integrated view of redox and catalytic properties of B-type PpDyP
from Pseudomonas putida MET94 and its distal variants. Arch. Biochem. Biophys.
Mendes, S., Catarino, T., Silveira, C., Todorovic, S., Martins, L.O., 2015b. The catalytic
mechanism of A-type dye-decolourising peroxidase BsDyP: neither aspartate nor
arginine is individually essential for peroxidase activity. Catal. Sci. Technol. 5,
Mendes, S., Robalo, M.P., Martins, L.O., 2015c. Bacterial enzymes and multi-enzymatic
systems for cleaning-up dyes from the environment. Microb. Degrad. Synth. Dyes
Min, K., Gong, G., Woo, H.M., Kim, Y., Um, Y., 2015. A dye-decolorizing peroxidase from
Bacillus subtilis exhibiting substrate-dependent optimum temperature for dyes and
beta-ether lignin dimer. Sci. Rep. 5, 8245.
Natte, K., Narani, A., Goyal, V., Sarki, N., Jagadeesh, R.V., 2020. Synthesis of functional
chemicals from lignin-derived monomers by selective organic transformations. Adv.
Synth. Catal. 362, 5143–5169.
Nys, K., Furtmuller, P.G., Obinger, C., Van Doorslaer, S., Pfanzagl, V., 2021. On the track
of long-range electron transfer in B-type dye-decolorizing peroxidases: identication
of a Tyrosyl radical by computational prediction and electron paramagnetic
resonance spectroscopy. Biochemistry 60, 1226–1241.
Ogola, H.J., Kamiike, T., Hashimoto, N., Ashida, H., Ishikawa, T., Shibata, H., Sawa, Y.,
2009. Molecular characterization of a novel peroxidase from the cyanobacterium
Anabaena sp. strain PCC 7120. Appl. Environ. Microbiol. 75, 7509–7518.
Ogola, H.J., Hashimoto, N., Miyabe, S., Ashida, H., Ishikawa, T., Shibata, H., Sawa, Y.,
2010. Enhancement of hydrogen peroxide stability of a novel Anabaena sp. DyP-type
peroxidase by site-directed mutagenesis of methionine residues. Appl. Microbiol.
Biotechnol. 87, 1727–1736.
Omura, I., Ishimori, K., Uchida, T., 2022. Converting cytochrome c into a DyP-like
metalloenzyme. Dalton Trans. 51 (33), 12641–12649.
Perez-Boada, M., Ruiz-Duenas, F.J., Pogni, R., Basosi, R., Choinowski, T., Martinez, M.J.,
Piontek, K., Martinez, A.T., 2005. Versatile peroxidase oxidation of high redox
potential aromatic compounds: site-directed mutagenesis, spectroscopic and
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
crystallographic investigation of three long-range electron transfer pathways. J. Mol.
Biol. 354, 385–402.
Petrus, M.L., Vijgenboom, E., Chaplin, A.K., Worrall, J.A., van Wezel, G.P., Claessen, D.,
2016. The DyP-type peroxidase DtpA is a tat-substrate required for GlxA maturation
and morphogenesis in Streptomyces. Open Biol. 6, 150149.
Pfanzagl, V., Nys, K., Bellei, M., Michlits, H., Mlynek, G., Battistuzzi, G., Djinovic-
Carugo, K., Van Doorslaer, S., Furtmuller, P.G., Hofbauer, S., Obinger, C., 2018.
Roles of distal aspartate and arginine of B-class dye-decolorizing peroxidase in
heterolytic hydrogen peroxide cleavage. J. Biol. Chem. 293, 14823–14838.
Pfanzagl, V., Bellei, M., Hofbauer, S., Laurent, C.V.F.P., Furtmuller, P.G., Oostenbrink, C.,
Battistuzzi, G., Obinger, C., 2019. Redox thermodynamics of B-class dye-decolorizing
peroxidases. J. Inorg. Biochem. 199.
Pi, Q., Zhu, Z., Tang, L., 2022. Transformation of reactive blue 19 by a recombinant
peroxidase DyP. Bioprocess Biosyst. Eng. 45 (2), 425–429.
van der Ploeg, R., Mader, U., Homuth, G., Schaffer, M., Denham, E.L., Monteferrante, C.
G., Miethke, M., Marahiel, M.A., Harwood, C.R., Winter, T., Hecker, M.,
Antelmann, H., van Dijl, J.M., 2011. Environmental salinity determines the
specicity and need for TAT-dependent secretion of the YwbN protein in Bacillus
subtilis. PLoS One 6 (3), e18140.
Pupart, H., Joul, P., Bramanis, M.I., Lukk, T., 2023. Characterization of the Ensemble of
Lignin-Remodeling DyP-type peroxidases from Streptomyces coelicolor A3(2).
Energies 16 (3).
Putri, R.M., Allende-Ballestero, C., Luque, D., Klem, R., Rousou, K.A., Liu, A., Traulsen, C.
H., Rurup, W.F., Koay, M.S.T., Caston, J.R., Cornelissen, J., 2017. Structural
characterization of native and modied Encapsulins as Nanoplatforms for in vitro
catalysis and cellular uptake. ACS Nano 11, 12796–12804.
Qin, X., Xin, Y., Su, X., Wang, X., Wang, Y., Zhang, J., Tu, T., Yao, B., Luo, H., Huang, H.,
2021. Efcient degradation of Zearalenone by dye-decolorizing peroxidase from
Streptomyces thermocarboxydus combining catalytic properties of manganese
peroxidase and laccase. Toxins 13 (9).
Rahman Pour, R., Ehibhatiomhan, A., Huang, Y., Ashley, B., Rashid, G.M., Mendel-
Williams, S., Bugg, T.D.H., 2019. Protein engineering of Pseudomonas uorescens
peroxidase Dyp1B for oxidation of phenolic and polymeric lignin substrates. Enzym.
Microb. Technol. 123, 21–29.
Rahmanpour, R., Bugg, T.D., 2013. Assembly in vitro of Rhodococcus jostii RHA1
encapsulin and peroxidase DypB to form a nanocompartment. FEBS J. 280,
Rahmanpour, R., Bugg, T.D., 2015. Characterisation of DyP-type peroxidases from
Pseudomonas uorescens Pf-5: oxidation of Mn(II) and polymeric lignin by Dyp1B.
Arch. Biochem. Biophys. 574, 93–98.
Rai, A., Klare, J.P., Reinke, P.Y.A., Englmaier, F., Fohrer, J., Fedorov, R., Taft, M.H.,
Chizhov, I., Curth, U., Plettenburg, O., Manstein, D.J., 2021. Structural and
biochemical characterization of a dye-decolorizing peroxidase from Dictyostelium
discoideum. Int. J. Mol. Sci. 22.
Rashid, G.M.M., Bugg, T.D.H., 2021. Enhanced biocatalytic degradation of lignin using
combinations of lignin-degrading enzymes and accessory enzymes. Catal. Sci.
Technol. 11, 3568–3577.
Rawat, D., Mishra, V., Sharma, R.S., 2016. Detoxication of azo dyes in the context of
environmental processes. Chemosphere 155, 591–605.
Renders, T., Van den Bosch, S., Koelewijn, S.F., Schutyser, W., Sels, B.F., 2017. Lignin-
rst biomass fractionation: the advent of active stabilisation strategies. Energy
Environ. Sci. 10, 1551.
Roberts, J.N., Hardiman, E.M., Singh, R., Eltis, L.D., Bugg, T.D., 2011a. Identication of
DypB from Rhodococcus jostii RHA1 as a lignin peroxidase. Biochemistry 50,
Roberts, J.N., Singh, R., Grigg, J.C., Murphy, M.E., Bugg, T.D., Eltis, L.D., 2011b.
Characterization of dye-decolorizing peroxidases from Rhodococcus jostii RHA1.
Biochemistry 50 (23), 5108–5119.
Rodrigues, C.F., Borges, P.T., Scocozza, M.F., Silva, D., Taborda, A., Brissos, V.,
Frazao, C., Martins, L.O., 2021. Loops around the Heme pocket have a critical role in
the function and stability of BsDyP from Bacillus subtilis. Int. J. Mol. Sci. 22 (19),
Ruiz-Duenas, F.J., Martinez, A.T., 2009. Microbial degradation of lignin: how a bulky
recalcitrant polymer is efciently recycled in nature and how we can take advantage
of this. Microb. Biotechnol. 2 (2), 164–177.
Ruiz-Duenas, F.J., Morales, M., Perez-Boada, M., Choinowski, T., Martinez, M.J.,
Piontek, K., Martinez, A.T., 2007. Manganese oxidation site in Pleurotus eryngii
versatile peroxidase: a site-directed mutagenesis, kinetic, and crystallographic study.
Biochemistry 46, 11.
Ruiz-Duenas, F.J., Morales, M., Garcia, E., Miki, Y., Martinez, M.J., Martinez, A.T., 2009.
Substrate oxidation sites in versatile peroxidase and other basidiomycete
peroxidases. J. Exp. Bot. 60 (2), 441–452.
Ruiz-Duenas, F.J., Lundell, T., Floudas, D., Nagy, L.G., Barrasa, J.M., Hibbett, D.S.,
Martinez, A.T., 2013. Lignin-degrading peroxidases in Polyporales: an evolutionary
survey based on 10 sequenced genomes. Mycologia 105, 1428–1444.
Runeberg, P.A., Brusentsev, Y., Rendon, S.M.K., Eklund, P.C., 2019. Oxidative
transformations of Lignans. Molecules 24 (2), 300.
Saez-Jimenez, V., Acebes, S., Garcia-Ruiz, E., Romero, A., Guallar, V., Alcalde, M.,
Medrano, F.J., Martinez, A.T., Ruiz-Duenas, F.J., 2016. Unveiling the basis of
alkaline stability of an evolved versatile peroxidase. Biochem. J. 473, 1917–1928.
Santos, A., Mendes, S., Brissos, V., Martins, L.O., 2014. New dye-decolorizing peroxidases
from Bacillus subtilis and Pseudomonas putida MET94: towards biotechnological
applications. Appl. Microbiol. Biotechnol. 98 (5), 2053–2065.
Scocozza, M.F., Martins, L.O., Murgida, D.H., 2021. Direct electrochemical generation of
catalytically competent Oxyferryl species of classes I and P dye decolorizing
peroxidases. Int. J. Mol. Sci. 22.
Sezer, M., Genebra, T., Mendes, S., Martins, L.O., Todorovic, S., 2012. A DyP-type
peroxidase at a bio-compatible interface: structural and mechanistic insights. Soft
Matter 8 (40).
Shrestha, R., 2017. Molecular Mechanism and Enzymological Studies of Dye-
Decolorizing Peroxidases (DyPs) from Thermomonospora Curvata and Enterobacter
Lignolyticus, Department of Chemistry. Kansas State University.
Shrestha, R., Chen, X., Ramyar, K.X., Hayati, Z., Carlson, E.A., Bossmann, S.H., Song, L.,
Geisbrecht, B.V., Li, P., 2016. Identication of surface-exposed protein radicals and a
substrate oxidation site in A-class dye-decolorizing peroxidase from
Thermomonospora curvata. ACS Catal. 6, 8036–8047.
Shrestha, R., Huang, G.C., Meekins, D.A., Geisbrecht, B.V., Li, P., 2017. Mechanistic
insights into dye-decolorizing peroxidase revealed by solvent isotope and viscosity
effects. ACS Catal. 7 (9), 6352–6364.
Shrestha, R., Jia, K., Khadka, S., Eltis, L.D., Li, P., 2021. Mechanistic insights into DyPB
from Rhodococcus jostii RHA1 via kinetic characterization. ACS Catal. 11,
Silva, D., Sousa, A.C., Robalo, M.P., Martins, L.O., 2022. A wide array of lignin-related
phenolics are oxidized by an evolved bacterial dye-decolourising peroxidase. New
Singh, R., Eltis, L.D., 2015. The multihued palette of dye-decolorizing peroxidases. Arch.
Biochem. Biophys. 574, 56–65.
Singh, R., Grigg, J.C., Armstrong, Z., Murphy, M.E., Eltis, L.D., 2012. Distal heme pocket
residues of B-type dye-decolorizing peroxidase: arginine but not aspartate is essential
for peroxidase activity. J. Biol. Chem. 287 (13), 10623–10630.
Singh, R., Grigg, J.C., Qin, W., Kadla, J.F., Murphy, M.E., Eltis, L.D., 2013. Improved
manganese-oxidizing activity of DyPB, a peroxidase from a lignolytic bacterium. ACS
Chem. Biol. 8, 700–706.
Strittmatter, E., Liers, C., Ullrich, R., Wachter, S., Hofrichter, M., Plattner, D.A.,
Piontek, K., 2013a. First crystal structure of a fungal high-redox potential dye-
decolorizing peroxidase: substrate interaction sites and long-range electron transfer.
J. Biol. Chem. 288, 4095–4102.
Strittmatter, E., Wachter, S., Liers, C., Ullrich, R., Hofrichter, M., Plattner, D.A.,
Piontek, K., 2013b. Radical formation on a conserved tyrosine residue is crucial for
DyP activity. Arch. Biochem. Biophys. 537, 161–167.
Strittmatter, E., Serrer, K., Liers, C., Ullrich, R., Hofrichter, M., Piontek, K., Schleicher, E.,
Plattner, D.A., 2015. The toolbox of Auricularia auricula-judae dye-decolorizing
peroxidase - identication of three new potential substrate-interaction sites. Arch.
Biochem. Biophys. 574, 75–85.
Sugano, Y., Yoshida, T., 2021. DyP-type peroxidases: recent advances and perspectives.
Int. J. Mol. Sci. 22.
Sugano, Y., Muramatsu, R., Ichiyanagi, A., Sato, T., Shoda, M., 2007. DyP, a unique dye-
decolorizing peroxidase, represents a novel heme peroxidase family: ASP171
replaces the distal histidine of classical peroxidases. J. Biol. Chem. 282 (50),
Sugano, Y., Matsushima, Y., Tsuchiya, K., Aoki, H., Hirai, M., Shoda, M., 2009.
Degradation pathway of an anthraquinone dye catalyzed by a unique peroxidase DyP
from Thanatephorus cucumeris Dec 1. Biodegradation 20 (3), 433–440.
Sugawara, K., Nishihashi, Y., Narioka, T., Yoshida, T., Morita, M., Sugano, Y., 2017.
Characterization of a novel DyP-type peroxidase from Streptomyces avermitilis.
J. Biosci. Bioeng. 123, 425–430.
Sugawara, K., Igeta, E., Amano, Y., Hyuga, M., Sugano, Y., 2019. Degradation of
antifungal anthraquinone compounds is a probable physiological role of DyP
secreted by Bjerkandera adusta. AMB Express 9 (1), 56.
Sun, Z., Fridrich, B., de Santi, A., Elangovan, S., Barta, K., 2018. Bright side of lignin
depolymerization: toward new platform chemicals. Chem. Rev. 118 (2), 614–678.
Sutter, M., Boehringer, D., Gutmann, S., Gunther, S., Prangishvili, D., Loessner, M.J.,
Stetter, K.O., Weber-Ban, E., Ban, N., 2008. Structural basis of enzyme encapsulation
into a bacterial nanocompartment. Nat. Struct. Mol. Biol. 15, 939–947.
Tang, Y., Mu, A., Zhang, Y., Zhou, S., Wang, W., Lai, Y., Zhou, X., Liu, F., Yang, X.,
Gong, H., Wang, Q., Rao, Z., 2021. Cryo-EM structure of Mycobacterium smegmatis
DyP-loaded encapsulin. Proc. Natl. Acad. Sci. U. S. A. 118.
Uchida, T., Sasaki, M., Tanaka, Y., Ishimorit, K., 2015. A dye-decolorizing peroxidase
from vibrio cholerae. Biochemistry 54 (43), 6610–6621.
Uchida, T., Omura, I., Umetsu, S., Ishimori, K., 2021. Radical transfer but not heme distal
residues is essential for pH dependence of dye-decolorizing activity of peroxidase
from vibrio cholerae. J. Inorg. Biochem. 219, 111422.
von Vacano, B., Mangold, H., Vandermeulen, G.W.M., Battagliarin, G., Hofmann, M.,
Bean, J., Kunkel, A., 2022. Sustainable design of structural and functional polymers
for a circular economy. Angew. Chem. Int. Ed. 62, e202210823.
Van den Bosch, S., Koelewijn, S.F., Renders, T., Van den Bossche, G., Vangeel, T.,
Schutyser, W., Sels, B.F., 2018. Catalytic strategies towards lignin-derived chemicals.
Top. Curr. Chem. (Cham.) 376 (5), 36.
Veitch, N.C., 2004. Horseradish peroxidase: a modern view of a classic enzyme.
Phytochemistry 65, 249–259.
Vuong, T.V., Singh, R., Eltis, L.D., Master, E.R., 2021. The comparative abilities of a small
laccase and a dye-decoloring peroxidase from the same bacterium to transform
natural and technical Lignins. Front. Microbiol. 12, 723524.
Yoshida, T., Sugano, Y., 2015. A structural and functional perspective of DyP-type
peroxidase family. Arch. Biochem. Biophys. 574, 49–55.
Yoshida, T., Sugano, Y., 2023. Unexpected diversity of dye-decolorizing peroxidases.
Biochem. Biophys. Rep. 33, 101401.
Yu, W., Liu, W., Huang, H., Zheng, F., Wang, X., Wu, Y., Li, K., Xie, X., Jin, Y., 2014.
Application of a novel alkali-tolerant thermostable DyP-type peroxidase from
Saccharomonospora viridis DSM 43017 in biobleaching of eucalyptus Kraft pulp. PLoS
One 9, e110319.
D. Silva et al.
Biotechnology Advances 65 (2023) 108153
Zalesak, F., Bon, D.J.D., Pospisil, J., 2019. Lignans and Neolignans: plant secondary
metabolites as a reservoir of biologically active substances. Pharmacol. Res. 146,
Zamocky, M., Jakopitsch, C., Furtmuller, P.G., Dunand, C., Obinger, C., 2008. The
peroxidase-cyclooxygenase superfamily: reconstructed evolution of critical enzymes
of the innate immune system. Proteins 72, 589–605.
Zhang, P., Xu, J., Wang, X.-J., He, B., Gao, S.-Q., Lin, Y.-W., 2019. The third generation of
articial dye-decolorizing peroxidase rationally designed in myoglobin. ACS Catal.
Zitare, U.A., Habib, M.H., Rozeboom, H., Mascotti, M.L., Todorovic, S., Fraaije, M.W.,
2021. Mutational and structural analysis of an ancestral fungal dye-decolorizing
peroxidase. FEBS J. 288, 3602–3618.
Zubieta, C., Krishna, S.S., Kapoor, M., Kozbial, P., McMullan, D., Axelrod, H.L., Miller, M.
D., Abdubek, P., Ambing, E., Astakhova, T., Carlton, D., Chiu, H.J., Clayton, T.,
Deller, M.C., Duan, L., Elsliger, M.A., Feuerhelm, J., Grzechnik, S.K., Hale, J.,
Hampton, E., Han, G.W., Jaroszewski, L., Jin, K.K., Klock, H.E., Knuth, M.W.,
Kumar, A., Marciano, D., Morse, A.T., Nigoghossian, E., Okach, L., Oommachen, S.,
Reyes, R., Rife, C.L., Schimmel, P., van den Bedem, H., Weekes, D., White, A., Xu, Q.,
Hodgson, K.O., Wooley, J., Deacon, A.M., Godzik, A., Lesley, S.A., Wilson, I.A., 2007.
Crystal structures of two novel dye-decolorizing peroxidases reveal a beta-barrel fold
with a conserved heme-binding motif. Proteins 69, 223–233.
Zuccarello, L., Barbosa, C., Galdino, E., Lonˇ
car, N., Silveira, C.M., Fraaije, M.W.,
Todorovic, S., 2021. SERR Spectroelectrochemistry as a guide for rational design of
DyP-based bioelectronics devices. Int. J. Mol. Sci. 22.
D. Silva et al.