172Scientific Bulletin. Series F. Biotechnologies, V ol. XXII, 2018 [626753]

172Scientific Bulletin. Series F. Biotechnologies, V ol. XXII, 2018
ISSN 2285-1364, CD-ROM ISSN 2285-5521, ISSN Online 2285-1372, ISSN-L 2285-1364

DEPOLYMERIZATION OF KRAFT LIGNIN WITH LACCASE
AND PEROXIDASE: A REVIEW

Aglaia BURLACU, Florentina ISRAEL-ROMING , Călina Petruța C ORNEA

University of Agronomic Sciences and Veterinary Medicine of Bucharest,
59 M ărăști Blvd., District 1, Bucharest, Romania

Corresponding author email: [anonimizat]

Abstract
Lignin is a complex aromatic polymer of phenyl propene units non-linear and randomly linked. The main building
blocks are p-coumaryl alcohol, coniferyl alcohol and si napyl alcohol. Lignin is the third most abundant biopolymer on
earth and usually accounts for 15-35% of the lignocellulose biomass. The degradation of lign in is extremely difficult
due to the complexity of the chemical structure (variable upon the source) and t he high molecular weight. Two major
types of enzymes involved in the depol ymerization of lignin are oxidored uctase: laccase and peroxidase, the main
microbial producers being fungi and some ba cteria. Due to its highly branched structure, lignin is considered to be the
most r ecalcitrant component of lignoc ellulose, most of it not being recovered. Therefore, there’s a demand for more
effective methods for depolymerization of lignin in order to obtain value-added products. This review underlines the
importance of valorization of lignin through enzymati c depolymerization with laccase and peroxidase.

Key words : kraft, laccase, lignin, peroxidase.

INTRODUCTION

Lignocellulose presents a speci al interest due to
its structure and composition, consisting of
complex biopolymers, such as: cellulose,
hemicellulose and lignin, materials with potential applicability for energy production (Mitache et al., 2015). Although cellulose and hemicellulose are relatively easy to hyd rolyse to obtain their
subunits, lignin depolymerization is difficult mainly because of its amorphic and complex three-dimensional structure and of its characteristics that make itself a binding polymer of the cells, fibres and vessels in wood or lignified parts of the plant. Lignin plays an important role in plant’s resistance, providing defence against pathogen attack, mechanical support, stress response and water transport (Li et al., 2016; Boerjan et al., 2003; Kilpeläinen et al., 2007). Therefore, there’s an imperative need to
remove lignin from the biomass in order to
have access to cellulose and hemicellulose.
Initially, lignocellulose biomass was delignified
through chemical pathways, but since lignin accounts for 15 – 30% of the biomass it was n’t
an economical process, so scientists started to search for ways to valorise lignin with enzymatic degrading systems that will not affect cellulose and hemicellulose. This review is focused on the importance of
degradation of kraft lignin and highlights the
main enzymes involved in the depolyme-
risation: laccase, lign in peroxidase, and
manganese peroxidase.
LIGNIN
Native lignins have certain variations in their
chemical composition based upon their source, thereby making it difficult to define the precise
structure of lignin.
Generally, lignin is considered to be a network
polymer containing building blocks of p-
coumaryl alcohol, coniferyl alcohol and sinapyl alcohol (as observed in Figure 1), that are non-
linear and randomly linked.

Figure 1. Lignin precursors

Kraft-lignin is a by-product obtained in the pulp and paper industries , through a Kraft

173
process. Kraft lignin differs considerably from
natural lignin in its structure and chemical composition, but given the fact that native lignin ’s structure is so variable and complex,
kraft lignin is often used as a substitute model
for native lignin.
The content of lignin in the cell wall structure is the most significant contributor for the biomass recalcitrance to microbial and enzymatic deconstruction (Li et al., 2016). Over the years, several biological and chemical methods were conducted in order to convert lignin in value added products. One of the
major challenge in using chemical methods for
lignin degradation is related to catalyst selectivity (Xu et al. , 2014), being difficult to
develop a general catalyst that can specifically work on native lignins structures that are so randomly variable. On the other hand, biological methods cannot compete with chemical processes involved in lignin depolymerization because in the end they can’t
produce the desired products more economically. But, biological methods are still preferred due to their selectivity and mild
reaction conditions. Therefore, enzymatic
deconstruction of lignin is somewhat a less
studied field that req uires more research.
Being a large heterogeneous polymer and not containing hydrolysable linkages, lignin degradation requires the act ion of extracellular
enzymes and more important oxidative enzymes (Hatakka, 2005).

Figure 2. Main en zymes used for lignin
depolymerization
Source: de Cassia Pereira et al., 2017

The main enzymes involved in lignin
depolymerization are: lignin peroxidase, manganese peroxidase and laccase (Figure2). Other enzymes include: horseradish peroxidase and dioxygenases (protocatechuate 3,4-dioxygenase; 1,2,3-trihydroxybenzene 1,2-dioxygenase and catechol 1,2-dioxygenase)
(Octavio et al., 2006).
These ligninolytic enzymes have an immense
potential for several industrial and
biotechnological processes, such as: food
industry, textiles, pulp and paper industry, bioremediation, medical, pharmaceutical cosmetic applications etc. (Maciel and Ribeiro,
2010; dos Santos Barbosa et al., 2008; Kunamneni et al., 2008; Maijala et al., 2007).

LACCASE
Laccase (benzenediol: oxygen oxidoreductases,
E.C. 1.10.3.2) is one of the most studied enzyme (Desai and Nityanand, 2011).
Laccases are multi-copper containing enzymes
belonging to blue oxidases group and are able to catalyse one-electron oxidation of phenolic compounds with concomitant reduction of oxygen to water (Gochev and Krastanov, 2007).
Unlike most enzymes, laccases have the ability
to display their activity on a wide range of
substrates like monophenols, diphenols,
polyphenols, methoxyphenols, aromatic
amines, benzenethiols and even some inorganic compounds such as iodine (Desai and Nityanand, 2011; Ai et al., 2015). Laccases from fungi are identified by their capacity to oxidize differe nt substrates such as
guaiacol, remazol bri lliant Blue R, tannic acid,
Poly R-478 etc. to specific coloured products (Desai and Nityanand, 2011). Even though laccases have a broad substrate specificity on phenolic compounds, they cannot work on non-phenolic sub-units, unless in the presence of mediators – low molecular – weight
organic compounds that act as „electron
shuttles” (Desai and Nityanand, 2011).
When laccase, cannot oxidize alone a substrate, it will first oxidize a m ediator that will form
highly reactive and unstable cationic radicals,
which will diffuse away from the enzymatic
pocket and will oxidize more complex substrates that could not enter into the active site due to their size . After that, the co-mediator
(oxidized mediator) will return to its original
state and the electrons taken by laccases are

174
finally transferred back to oxygen to form
water (as shown in Figure 3) (Desai and Nityanand, 2011; Gochev and Krastanov, 2007).

Figure 3. Mechanism of substrate oxidation
by laccase with a mediator
Source: Christopher et al., 2014

The most often used mediators are: 2,2`-azinobis (3-ethylbenzthiazoline-6-sulfonate) (ABTS), 1-hydroxybenzotriazole (HBT), ben-zotriazole (BT), remazol brilliant blue (RBB), chlorpromazine (CPZ), promazine (PZ), 1-nitroso-2-naphtol-3,6-disulfonic acid (NNDS),
N-hydroxyphtalimide (NHPI), 4-hydroxy-3-
nitroso-1-naphthalenesulfonic acid (HNNS), 3-hydroxyanthranilic acid, N-hydroxyacetanilide (NHA), violuric acid (Octavio et al., 2006; Desai and Nityanand, 2011; Gochev and Krastanov, 2007; Lange et al., 2013; Brijwani et al., 2010).
Laccases can display their activity on a wide
range of temperatures and pH. Optimum pH value is variable based on the reactions caused by the substrate employed, molecular oxygen
or the enzyme itself (Desai and Nityanand,
2011). Laccases have a significant biotech-nological potential due to their broad substrate
specificity being used in: environmental biore-mediation (removal of pollutants, such as alkenes, chlorophenols, dyes, herbicides, polycyclic aromatic hydrocarbons and benzo-pyrene) (Gochev and Krastanov, 2007), food and beverages, biosensors, pulp and paper industry (pulp delignificati on), transformation
of antibiotics and ste roids, detergent
manufacturing bioethanol (Octavio et al., 2006; Brijwani et al., 2010).
Mechanism of action
Laccases contain 4 copper atoms, that are
classified into three typ es, referred to as type 1
(T1), type 2 (T2) and type 3 (T3). The copper
atoms differ from each other in their electron paramagnetic resonance (EPR) signals (Gochev and Krastanov, 2007). The type 1 Cu is responsible for the blue colour of the protein, type 2 Cu does not confer colour and the type 3 Cu atoms consists of a pair of Cu atoms in a binuclear co nformation (Desai and
Nityanand, 2011). Type 2 and Type 3 copper sites forms a
trinuclear centre responsibl e for the catalytic
mechanism of the enzyme (Desai and
Nityanand, 2011). Laccase catalysis pathway implies three major steps: the type 1 Cu is reduced by a reducing substrate, the electron is transferred from the type 1 Cu to the trinuclear cluster of type 2 Cu and type 3 Cu and at the trinuclear centre will take place the reduction of oxygen to water
(Brijwani et al., 2010).
Laccase is considered to act as a battery, that stores electrons from individual oxidation reactions in order to reduce molecular oxygen. Therefore, there are required four molecules of reducing substrate for the complete reduction of molecular oxygen to water (Desai and Nityanand, 2011).
Sources
Amongst all of the large blue copper containing
proteins, laccases are the most widely
distributed in sources such as: bacteria, fungi,
higher plants and insects (Desai and Nityanand,
2011; Gochev and Krastanov, 2007).
Laccase was first characterized when it was
extracted from the Japanese lacquer tree Rhus
vernicifera in 1883. Later, in 1896, it was demonstrated that laccases were also present in
fungi (Desai and Nityanand, 2011). In higher plants, laccases can be found in Rhus
vernicifera, Rhus succedanea , Lactarius
piperatus , Prunus persica (Octavio et al.,
2006), Acer pseudoplatanus, Chaetomiaceae sp. (Christopher et al., 2014).
Laccase activity has been reported only in a few bacteria such as: Azospirillum lipoferum,
Marinomonas mediterranea, Streptomyces
griseus , Bacillus subtilis (Octavio et al., 2006),
Streptomyces lavendulae , Streptomyces
maltophilia, Streptomyces coelicolor , Bacillus
licheniformis (Desai and Nityanand, 2011;
Christopher et al., 2014).
The most studied laccases are the ones from
fungal sources, including genera of Ascomycetes , Deuteromycetes , Basidiomycetes
and cellulolytic fungi (Christopher et al., 2014). Amongst these, the most frequently described were the laccases from the white-rot

175
basidiomycetes such as: Trametes versicolor,
T. hirsuta, T. ochracea, T. villosa, T. gallica,
Phlebia radiata, Coriolopsis polyzona,
Lentinus edodes, Pleurotus ostreatus (Desai
and Nityanand, 2011; Brijwani et al., 2010),
Pycnoporus cinnabarinus, Coprinus cinereus
(Christopher et al., 2014). Other fungal strains
include: Agaricus blazei, Melanocarpus
albomycea (Christopher et al., 2014), Stereum
ostrea, Lentinus tigrinus, Ganoderma spp.,
Polyporus versicolor, Pholiata spp., Podospora
anserine, Neurospora crassa, Aspergillus
nidulans, Pyricularia oryzae (Octavio et al.,
2006), Trichoderma harzianum, Trichoderma
atroviride, Trichoderma longibrachiatum,
Aspergillus niger, Phanerochaete
chrysosporium, Theliophora terristrus, Stereum
ostrea (Gochev and Krastanov, 2007). Marine
derived fungi that display laccase activity were:
Coriolopsis byrsina, Cerrena unicolor,
Diaporthe phaseolorum, Pestalotiopsis uvicola
(Desai and Nityanand, 2011).
Bacterial laccases are more stable to high pH
and temperature compared with the fungal ones (acidic optimum pH). The optimal temperature
for laccases is usually between 50 – 70 °C
(Christopher et al., 2014).

Depolymerization of Kraft lignin
At first, some genera of basidiomycetes involved in lignin depolymerisation were found to lack lignin peroxidases, indicating that different enzymes were responsible for the degradation. After some research, it was
suggested that laccases could play a key role in lignin depolymerisation (Gochev and Krastanov, 2007).
Laccases importance in this degradation is due
to their capacity to wor k on both phenolic and
non-phenolic compounds (Desai and Nityanand, 2011).
Regarding the depolymerization of lignin,
laccase will first attac k the phenolic lignin
moiety (<20% of total lignin), releasing phenolic residues (as shown in Figure 4) with
oxidized side chains (phenolic aldehydes, keto-
nes and acids). After that, through a mediator
facilitated process, laccase will oxidize the non-phenolic benzylic structures (Christopher et al., 2014). The phenolic fragments resulted in the first oxidation are able to infiltrate in the bulk
lignin polymer and act as a natural mediator, thus helping the enzyme to oxidize more recalcitrant non-phenolic lignin (Christopher et al., 2014; Reddy et al., 2003).

Figure 4. Simplified reaction of lignin depolymerisation
with laccase
Source: Hatakka, 2005

An important factor in lignin degradation with laccase is the molecular weight of lignin and its phenolic content (Niku-Paavola et al., 2002). It was implied that during laccase action, both
polymerization and depolymerization reactions
can take place, as the phenoxy radicals produced can either lead to oxidation or polymerization (Tamminen et al., 2003). The most common products obtained after lignin depolymerization with laccase and mediators are: 2,6-dimethoxy-4-
methylbenzaldehyde, 4-ethyl-2,6-
dimethoxybenzaldehyde and 2,6-dimethoxy-4-
((E)-prop-1-enyl) benzaldehyde) (Du et al., 2013). White rot fungi are the main microbial strains
involved in the lignin degrada tion due to their
extracellular high laccase activity (Christopher
et al., 2014). In comparison to peroxidases, laccases have a broad substrate specificity and are able to display their activity using only atmospheric
oxygen as electron donor, instead of hydrogen peroxide used by peroxidases (Christopher et al., 2014). By not using hydrogen peroxide, laccases show a greater stability which allows
them to be used more efficiently in an
immobilised way (Octavio et al., 2006). The ability of laccase to work with mediators in
order to oxidize both phenolic and non-phenolic compound of lignin is considered to
be a significant participant in lignin valorise-
tion (Christopher et al., 2014).
PEROXIDASE
Peroxidases (E.C. 1.11.1.7) involved in lignin
degradation are heme- containing enzymes that

176
can oxidize a variety of organic and inorganic
substrates in the presence of hydrogen peroxide as electron acceptor (O’Brien, 2000 ; Falade et
al., 2017).
These enzymes are a group of oxidoreductases
that catalyses the reduction of peroxides such
as hydrogen peroxide and the oxidation of a variety of organic and inorganic compounds. Heme-peroxidases are extracellular enzymes associated with lignin depolymerization. They include three types of enzymes: lignin peroxidase (LiP), manganese peroxidase (MnP) and versatile peroxidase (VP).
Lignin-peroxidase (diaryl propane oxygenase;
LiP, E.C. 1.11.1.14) is a monomeric hemo-
protein. This enzyme has the capacity to catalyse hydrogen peroxide dependent oxidative depolymerization of lignin (Falade et al., 2017).
Manganese peroxidases (Manganese-dependent
peroxidases; MnP; E.C. 1.11.1.13) are
extracellular glycoproteins that are considered
to be the most common ligninolytic enzymes produced by white-rot fungi (Falade et al.,
2017). The production of these enzymes is
regulated by nutrients availability and
environmental factors (Silva, 2014).
Versatile peroxidases (VP; E.C. 1.11.1.16) are
a group of enzymes belonging to the class II
subfamily of peroxidases. Its name derives from its versatile nature, being able to oxidize directly diverse substrates from hydroquinones, substituted phenols to bulky lignin, without redox mediators (Ravichandran and Sridhar, 2016).
VP was first mistaken with MnP in 1996 by
Martínez et al. and in 2000 by Giardina et al. (Ruiz-Dueñas et al., 2009). VP is also known as a hybrid peroxidase or manganese-lignin peroxidase, because of its ability to combine the catalytic properties of both MnP and LiP, being able to oxidize both
phenolic and non-phenolic compounds. These three types of peroxidases can work together on lignin degradation if they are produced by the same organism. While LiP
oxidize the non-phenolic components of lignin
and MnP targets the phenolic ones, VP has the ability to oxidize both phenolic and non-phenolic structures (Falade et al., 2017). Ligninolytic peroxidases are used in a variety
of applications such as: removal of contaminants, organic and polymer synthesis, pulp and paper industry, biosensors, analysis
and diagnostic kits, enzyme immunoassays,
biofuel production (Hamid, 2009).
Mechanism of action
Peroxidases catalyse the oxidation of a wide
variety of substrates, using H
2O2 or other
peroxides. In general, the per oxidase catalytic
cycle involves distinct intermediate enzyme forms and the activation of molecular oxygen is achieved in a two-step p rocess. First, the native
ferric enzyme is oxidised by hydrogen peroxide to form an unstable intermediate called
compound I (Co I), which has a heme structure
of Fe IV=O-porphyrin p-cation radical (Hamid,
2009), with consequent reduction of peroxide to water. Then Co I oxidise electron donor substrate to give compound II (Co II), releasing a free radical. Co II is further reduced by a second substrate molecule, regenerating the iron (III) state and producing another free radical. Lignin peroxidase oxidize various non-phenolic
structures of lignin including β -O-4 linkage-
type arylglycerol-aryl ethers. The oxidation
mechanism involves the formation of radical
cation through one electron oxidation and this
action leads to side-chain cleavage, demethyla-tion, intramolecular addition and nonetheless rearrangements (Falade et al., 2017). Although, LiP mainly oxidize non-phenolic structures, it can also act on a variety of phenolic compounds such as: guaiacol, acetosyringone, catechol, vanillyl alcohol,
syringic acid etc. (Falade et al., 2017).
An important redox mediator for LiP activity in lignin depolymerization is veratryl alcohol, a
non-phenolic metabolite and a high redox potential substrate (Falade et al., 2017). The catalytic capacity of LiP has been attributed to its exposed tryptophan residues, that forms a tryptophanyl radical on the surface of the enzyme through long-range electron transfer to the heme (Falade et al., 2017).
The variation in the tryptophan environment
can influence the enzyme activity, stability and
substrate specificity (Falade et al., 2017). The mechanism of action of LiP involves three steps: oxidation of the resting ferric enzyme by hydrogen peroxide resulting in formation of compound I (oxo-ferryl intermediate), transfer

177
of one electron from the substrate to the
compound I, to form compound II and subsequent donation of a second electron from the reduced substrate to compound II (Abdel-
Hamid et al., 2013).
Similar to laccase, manganese peroxidase can
have the capacity to oxidize non-phenolic compounds in the presence of mediators such as thiol or lipid radicals.
Sources
Peroxidases are widely distributed in nature in
plants, animals and microorganisms.
LiP was first discovered in the extracellular
medium of white-rot fungus Phanerochaete
chrysosporium in 1983 (Falade et al., 2017).
After that, several sources have been reported
such as: Trametes versicolor, Phanerochaete
sordida, Phlebia radiata (Falade et al., 2017),
Trametes villosa, Trametes trogii, Phlebia tremellosa, Phlebia ochraceofulva, Junghuhnia
separabilima (Hatakka, 2005).
Several white-rot basidiomycetes have
displayed exclusively MnP activity as
extracellular peroxidase, such as: Ceriporiopsis
subvermispora, Dichomitus squalens, Lentinula
(Lentinus) edodes, Phanerochaete sordida,
Pleurotus ostreatus (Gasser et al., 2012),
Pleurotus eryngii (Camarero et al., 1999),
Abortiporus biennis, Agaricus bisporus, Bjerkandera sp., Cyathis stercoreus,
Heterobasidion annosum, Nematoloma
frowardii, Panus tigrinus, Rigidoporus lignosus
(Hatakka, 2005).
Pleurotus eryngii is considered to be a model
organism for studies regarding biodegradation
of lignin, due to its sele ctivity in removing
lignin when cultivate d on natural substrates
(Camarero et al., 1999). Some reports indicate that some white-rot fungi can produce both LiP and MnP: Phanerochaete
chryosporium, Phanerochaete flavido-alba,
Phlebia radiata, Bjerkandera adusta and
Trametes versicolor (Gasser et al., 2012;
Hatakka, 2005).
VPs have been detected mainly in Pleurotus
and Bjerkandera species (Gasser et al., 2012).
Some reports suggest that VP’s can also be
found in Panus, Calocybe, Trametes, Lepista,
Dichomitous and Spongipellis species
(Ravichandran and Sridhar, 2016).
Depolymerization of Kraft lignin
In nature, lignin is efficiently mineralized by
multiple enzymes produced mainly by white-
rot fungi, which are found in forest litter and fallen trees (Gasser et al., 2012).
Initially, LiP was considered as the main
enzyme connected to the oxidative breakdown of lignin due to the highly amount of non-phenolic units in lignin str ucture (Camarero et
al., 1999).
The discovery of lignin peroxidase was a major
step in understanding the mechanism of lignin
depolymerization (O’Brien, 2000) .
Lignin peroxidase has a high redox potential for the oxidation of non-phenolic structures which represent up to 90% of lignin (Figure 5) (Martinez, 2005).

Figure 5. Simplified lignin de polymerization with lignin
peroxidase
Source: Falade et al., 2017

Besides the involvement in degradation of non-phenolic structures of li gnin, LiP can also
oxidize the aromatic rings of lignin via long-range electron transfer (Gasser et al., 2012), which results in formation of unstable cation radicals, that will undertake different non-
enzymatic reactions (Falade et al., 2017).
In comparison with laccase, LiP doesn’t require
mediators to degrade high redox potential
compounds, but it does need hydrogen peroxide
to initiate the catalys is (Maciel and Ribeiro,
2010). There are many reports on depolymerisation of both native and synthetic lignins with
manganese peroxidase (Falade et al., 2017).
The action of manganese peroxidase is started by the activation of hyd rogen peroxide with the
iron protoporphyrin IX, that in return will oxidize the manganese co-factor from Mn
2+ to
the highly reactive Mn3+. The Mn3+ centre is
chelated by carboxylic acid anions that will

178
produce small, freely diffusible species, which
will act as redox mediators, oxidizing phenolic lignin structures, as seen in Figure 6 (Falade et al., 2017; Gasser et al., 2012; Lange et al.,
2013).

Figure 6. Simplified lignin de polymerisation with MnP
Source: Hatakka, 2005

Additionally, it was rep orted that lipid
peroxidation by MnP might be an important factor in degradation of non-phenolic structures of lignin, due to the fact that the formed
peroxyl radicals (Figure 6) can act as agents
that will promote the oxidation of non-phenolic
β-O-4-linked lignin compounds (Gasser et al.,
2012). In comparison with LiP, MnP has displayed a
preference to oxidize in vitro phenolic
substrates, due to its lower redox potential. On
the other hand, some reports suggest that unlike LiP, MnP may be able to oxidize Mn
2+ without
hydrogen peroxide with decomposition of acids and concomitant production of peroxyl radicals (Maciel and Ribeiro, 2010; Hofrichter et al., 1999). The interesting part abo ut versatile peroxidase
is that due to its unique molecular structure given by the presence of different oxidation-active sites (Falade et al., 2017; Ruiz-Dueñas et al., 2008 ), it’s able to oxidize t he substrate
without redox mediators, unlike MnP and LiP (Ravichandran and Sridhar, 2016). Therefore, VP can have a great potential for future biotechnological applications (Busse et al., 2013). The non-enzymatic reactions include aromatic
ring cleavage, hydroxylation, demethoxylation,
ether bond cleavage, side chain cleavage and
phenol formation (Busse et al., 2013).

CONCLUSIONS
Lignin is the second most abundant biopolymer
that represents 15-30% of lignocellulosic biomass. There’s an imperative need to find new and
improved methods to valorise lignin, without
affecting the other major components of lingo-
cellulosic biomass. The difficulty in degrading lignin is reflected
by lignin complex and variable structure.
The most important enzymes involved in lignin depolymerisation are: lacca se, lignin pero-
xidase and manganese peroxidase.
While lignin peroxidase attacks the non-
phenolic structures of lignin, laccase and man-
ganese peroxidase catalyse, without mediators,
the oxidation of phenolic fragments of lignin. These enzymes can work together if they are produced by the same microorganism. Therefore, an improvement on this subject could be the exploration and optimization of novel microbial sources that can produce these
enzymes capable of depolymerizing lignin.

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