Enzymes for Lignin Depolymerization: Comparison
View Latest Version
Please note this is a comparison between Version 2 by Jason Zhu and Version 1 by Zheyi Wang.

Lignin, a rigid polymer composed of phenolic subunits with high molecular weight and complex structure, ranks behind only cellulose in the contribution to the biomass of plants. Therefore, lignin can be used as a new environmentally friendly resource for the industrial production of a variety of polymers, dyes and adhesives. Since laccase was found to be able to degrade lignin, increasing attention had been paid to the valorization of lignin. Research has mainly focused on the identification of lignin-degrading enzymes, which play a key role in lignin biodegradation, and the potential application of lignin degradation products. The enzymes involved in lignin depolymerization are mainly divided into four categories: laccase, lignin peroxidase (LiP), manganese peroxidase (MnP) and versatile peroxidase (VP).

  • Lignin
  • Laccase
  • Enzymes

1. Lignin

Coniferyl (G), sinapyl (S) and p-coumaryl (H) alcohols are the three categories of monolignols in lignin, which are divided into guaiacyl (G-type), guaiacyl-syringyl (GS-type) and grassy or guaiacyl-syringylp-coumaric (GSH-type). However, it needs to be noted that, in actual examples, other aromatic units are occasionally included in lignin, which is a common phenomenon. For example, Stewart et al. described the structure of the lignin polymers from poplar. The lignin from wild-type poplar contains p-hydroxybenzoate [7][1]. C-lignin, found in plants such as Vanilla planifolia and Cleome hassleriana, is wholly derived from caffeyl alcohol [8][2]. For sure, the GSH-type proportion and linkages proportion in these practical examples also varied from each other.
As a concept different from lignins in vivo, technical lignins (e.g. kraft lignin, soda lignin, lignosulfonates and organosolv lignin) are isolated lignins, most of which are byproducts from the paper and pulp industry [1,9][3][4]. Kraft lignin, which is recovered from the black liquor, is generated from the paper and pulp industry as a by-product that is generated by treatment with Na2S and NaOH [10][5]. However, the extraction of lignin from black liquor limits the commercial development of lignin until the emergence of the LignoBoost and LignoForce methods, which could also be applied for lignosulfonate and soda lignin. An economical benefit could be achieved as the extracted lignin has a high proportion of phenolic hydroxy groups [11][6].
Lignosulfonate is obtained as a by-product in the spent liquor of the pulping process by treating plant fiber as raw materials with an aqueous solution of sulfur dioxide and a calcium, magnesium, ammonium or sodium base [12][7]. The kraft lignins could also be used as raw metarials for lignosulfonate production. In the revisewsarch provided by Ruwoldt [13][8] and Kienberger et al. [14][9], the lignin extraction processes and their dominant products, including commercialized lignosulfonates, were well-summarized. Membrane processes for the isolation and purification of lignosulfonates are also commercially available. In the meanwhile, fractionation of the lignosulfonates could obtain products with high quality. Recently, Demuner et al. elucidated two different processes for lignosulfonate production from eucalypt kraft lignin by sulfomethylation and acid sulfonation [15][10]. Varied pH levels can be applied in this process, for example, lignosulfonate production through neutral sulphite pulping was investigated [16][11]. It has the higher sulfur content and the lower price at the market when compared with kraft lignin [17][12].
Soda lignin is obtained from the soda or soda-anthraquinone pulping processes, in which 13–16% sodium hydroxide (pH 11–13) is used to extract lignin [18][13]. The chemical composition of soda lignin is closer to the natural lignin when compared with kraft lignin and lignosulphonate [19][14]. The major advantage of soda lignin is that it is obtained from wheat straw, an unwanted agricultural waste derived from the cropping of wheat.
Organosolv lignin is attained from the organosolv process, in which organic solvents are applied. Organosolv lignins, which are hydrophobic and water-insoluble, have several important characteristics: low molecular weight, high quality and high purity [20][15]. Nonetheless, the application of organic solvents may face the problem of environmental pollution [15][10]. However, organosolv lignin is sulfur-free, as is soda lignin, thus relieving the concern of environmental pollution from another perspective. In a revisewarch given by Wang et al., they well-summarized the chemical extraction methods for technical lignin [6][16].
Common derivatives such as vanillin, lipid and PHA could be produced via the lignin degradation process [21][17]. Multiple high value-added small molecules have been linked with lignin. Lignin, with a great amount of phenols, is considered to be a good candidate for producing phenolic resins. At present, products from lignin depend on sources besides the operating conditions [2][18]. Producing lignin-based resin with a high substitution rate of lignin to phenols is still a challenge. Recently, lignin with more S units was converted to desirable phenolic resins with a suitable viscosity and high substitution rate using chemical means with the beginning of methylolation [22][19]. Like phenolic resins, bio-oil generated by lignin can also broaden the application of lignin and thereby meet the new concept of carbon neutrality. Lignin can be transited to bio-oil via thermochemical methods (pyrolysis, gasification and liquefaction) [23][20]. However, these processes are usually energy-consuming and labor-intensive [24][21]. Problems like high energy demand and environment pollution encourage uspeople to explore new ways for lignin degradation and valorization. The biodegradation of lignin could be an ideal choice. By exploiting enzymes by means such as cell-free systems, and taking advantage of the pathway of microorganisms to produce high value-added compounds, the biodegradation and valorization of lignin has a broad prospect.

2. Laccase (EC 1.10.3.2)

As a typical lignin oxidase, increasing attention has been paid to lignin depolymerization by laccase (EC 1.10.3.2). Laccase, a type of multi-copper oxidase (MCO), generally has four copper atoms (one type-1 copper, one type-2 copper and two type-3 coppers) in the catalytic site and oxidizes a variety of substrates by performing the single-electron transfer reaction four times. A O2 molecule is reduced to two molecules of H2O through the electron transfer pathway in laccase [25,26][22][23]. The laccase-like activity could be detected by applying agar plates containing standard laccase substrates guaiacol or syringaldazine and then monitoring the dark colors coming from the oxidation of these substrates [27][24]. Chen et al. analyzed the binding mechanism of laccase from Trametes versicolor (formerly Coriolus versicolor or Polyporus versicolor) with lignin model compounds, and it turned out that hydrophobic contacts were necessary between laccase and the model compounds, but H-bonds were alternative [28][25]. In the reaction between laccases and lignins, the phenoxy radicals are first to be generated. The enzyme kinetics of lignin radical formation can be achieved by electron paramagnetic resonance (EPR) [29][26]. The formation of radicals could result in (1) depolymerization of the lignin [30][27], (2) polymerization of the lignin [31][28] or (3) the change of the functional group [32][29]. These potential outcomes could be effected by factors such as reaction conditions or whether the Laccase-Mediator System is involved in the reaction system. The depolymerization of lignin requiring mediators is a common opinion [30,33][27][30]. Due to the relatively low redox potential of laccase for aromatic compounds, the Laccase–Mediator System (LMS) has been widely studied in order to expand the applications of laccases [34,35,36,37,38][31][32][33][34][35]. The mediators can be divided into three categories: natural mediators, synthetic mediators and other mediators. Some effective mediators have been widely used in industries, such as 1-hydroxybenzotriazole (HBT), methyl syringate and 2,2′-azino-bis(3-ethylbenzo thiazoline-6-sulfonic acid) (ABTS). In the LMS, the mediator is firstly oxidized by laccase and then undergoes a redox reaction with the substrate. This mechanism also implies that the laccase is a type of environment-friendly biocatalyst due to its lower requirements for factors such as heat for pyrolysis. Although mediators can tremendously accelerate the process of substrate oxidation by laccase [39][36], it also has significant influences on the stability of laccase. The free radicals generated from the oxidized mediator could attack laccase and then lead to its inactivation. To avoid the radical attack and improve the stability of laccase in LMS, it could be a promising strategy to find mediators that generate free radicals with lower activities. According to the evolutionary tree created in this review, laccases from different organisms are highly homologous to each other, which suggests that their protein sequences are highly conserved. The identification of the specific amino acids responsible for the increase of the oxidation potential of laccase will contribute to the industrial application of laccase. Laccases from fungi, plants, insects and bacteria have been characterized. Laccases from white-rot fungi Pycnoporus cinnabarinus and T. versicolor have attracted wide attention since the last century [40][37]. P. cinnabarinus produces laccase in one isoform, and neither LiP nor MnP are detected. Thus by comparison between wild-type strains and laccase-less mutants, laccases from P. cinnabarinus are proven to be efficient for lignin degradation [41][38]. Kawai et al. reported that laccase from T. versicolor can catalyze various reactions of nonphenolic-O-4 lignin in the presence of HBT, such as β-ether cleavage, Cα-Cβ cleavage, aromatic ring cleavage and Cα-oxidation [42][39]. Compared with fungal laccases, laccases from bacteria have a higher thermostability and wider effective pH range [43,44,45][40][41][42]. Moreover, they are efficiently expressed in heterologous hosts such as E. coli with relatively high yields. All of these advantages render bacterial laccases great potential in energy and environmental biotechnology. Fungal laccases and bacterial laccases also tend to cause different effects on lignin; a study showed that fungi mineralized kraft lignin, while bacteria partially degraded and modified lignin [46][43]. In addition, due to their low substrate specificity, laccases have also been applied in many fields such as bioremediation, wood and paper industries [47,48][44][45]. The application of laccase is also reflected in the potential of small lignin production. With Trametes villosa laccase combined with HBT, the yield of glucose and ethanol were increased from lignocellulosic by Gutiérrez et al. [30][27] Through catalyzation of the above-mentioned enzymes, lignin enters the degradative pathways. To better understand the degradative pathways for lignin-derived aromatic compounds, it is necessary to summarize the products and effect of the enzymes in lignin depolymerization. The biodegradation of alkaline lignin by Bacillus ligniniphilus L1 resulted in the production of phenylacetic acid, 4-hydroxy-benzoicacid and vanillic acid [49][46]. Based on the whole-genome sequencing (with no LiP and MnP) and the decolorization experiment, laccase is expected to be responsible for the depolymerization of alkaline lignin in B. ligniniphilus L1.

3. Lignin Peroxidase (EC 1.11.1.14)

Lignin peroxidase (LiP) (EC 1.11.1.14), which belongs to the peroxidase-catalase superfamily, was firstly discovered in white-rot fungus Phanerochaete chrysosporium during the 1980s [50,51][47][48]. LiP is a heme peroxidase, which contains a protoporphyrin IX (heme) as the prosthetic group. Heme iron is responsible for the electron transfer and the high redox potential of LiP. Hydrogen peroxide (H2O2) functions as the electron acceptor in the catalytic site of LiP and drives the oxidation step. Two steps are required for the reduction of LiP. Compared with laccase, LiP has a higher reduction potential and requires no mediators [52][49]. However, previous studies found that the addition of veratryl alcohol (3,4-dime-thoxybenzyl alcohol; VA) can enhance the activity of LiP by transforming the electron between LiP and the substrates, thus promoting the rate of lignin depolymerization [53][50]. LiP exhibits oxidative activity on a variety of compounds, such as phenolic compounds, amines, aromatic ethers and polycyclic aromatics [54,55][51][52]. Only two single-electron transfers are required for substrate oxidation. Though H2O2 plays an important role in the typical peroxidase catalytic cycle, excess H2O2 inhibits the activity of LiP [56][53]. To exploit the lignin degradation ability of LiP, explorations from different strategies were carried out. Majeke et al. reported the synergistic application of quinone reductase (QR) and LiP; this combination decreased the molecular weight (MW) of four different lignins [57][54] and showed the potential to benefit from the cooperation of different enzymes. On the other hand, considering LiP could be utilized in cell-free systems, the production of LiP can be improved. The effort for the large-scale production of LiP could boost the application of LiP for reasons such as cost reduction. Liu et al. achieved a maximum LiP activity of 1645 mU/L via multipulse-fed batch production [58][55]. Based on these paradigms, LiP will have larger application prospects.

4. Manganese Peroxidase (EC 1.11.1.13)

Manganese peroxidase (MnP) (EC 1.11.1.13) was also discovered in P. chrysosporium [59][56]. MnP could be secreted by ligninolytic fungi into their microenvironment and, thus, be an ideal choice for cell-free systems. Similar to LiP, MnP is also a heme-containing enzyme, with disulfide bonds and Ca2+ maintaining the conformation of the active site [60][57]. When H2O2 binds to the ferric enzyme in a resting state, the reactions will be initiated by forming an iron–peroxide complex. MnP compound III, an inactive oxidation state, could be formed under excess H2O2 condition [61][58]. Therefore, first, Mn2+ is oxidized to Mn3+; then, Mn3+ acts as the redox mediator to attack phenolic lignin structures and results in the formation of instable free radicals, causing lignin degradation. One remarkable difference in the catalytic mechanism between LiP and MnP is that MnP, but not LiP, requires the conversion between Mn2+ and Mn3+ as an intermediate of the reaction. In the absence of mediators, one molecule of water will be produced in LiP’s catalytic cycle, but the number of water is two for MnP’s catalytic cycle [62][59]. By the treatment of milled wood with the MnP–, Mn3+– or Fe2+–linoleic acid system, MnP exhibited the most intense reactions, including transformation in the lignin structure and water-soluble low molar mass aromatic fragments releasing [63][60]. Base on the increase of the carboxyl content, the degradation of lignin was carried out via β-O-4 and C𝛼–C𝛽 cleavage and side-chain oxidation. In addition, besides the degradation of lignins, attention has been paid to the versatility of MnP. For example, MnP from P. chrysosporium could improve the efficiecy of cellulosic decomposing [64][61]. A manganese peroxidase-producing yeast consortium (MnP-YC4) has been developed to withstand lignin degradation inhibitors while degrading and detoxifying azo dye [65][62].

5. Versatile Peroxidase (EC 1.11.1.16)

Versatile peroxidase (VP) (EC 1.11.1.16), also belonging to the peroxidase-catalase superfamily, was firstly found in Pleurotus eryngji. Studies have revealed that VP is a "hybrid peroxidase", because its gene sequence contains the partial sequences of LiP and MnP. It integrates the substrate specificity and catalytic properties of LiP and MnP, respectively [66][63]. Therefore, VP can oxidize phenolic, nonphenolic and lignin derivatives without manganese ion or any mediator because of the high redox potential and broader substrate spectrum. The high redox potential of VP results in a wide substrate specificity. For eample, VP can oxidize the LiP substrate veratryl alcohol; five molecules of water will be produced in the presence of veratryl alcohol (VA) [39,67][36][64]. In a previous restudyearch, VP showed oxidizing activity on hydroquinone in the absence of exogenous H2O2 but in the presence of Mn (Ⅱ) [68][65]. When oxidizing phenolic compounds, VP uses a similar mechanism to MnP.

References

  1. Stewart, J.J.; Akiyama, T.; Chapple, C.; Ralph, J.; Mansfield, S.D. The effects on lignin structure of overexpression of ferulate 5-hydroxylase in hybrid poplar1. Plant Physiol. 2009, 150, 621–635.
  2. Wang, X.; Zhuo, C.; Xiao, X.; Wang, X.; Docampo-Palacios, M.; Chen, F.; Dixon, R.A. Substrate specificity of LACCASE8 facilitates polymerization of caffeyl alcohol for C-lignin biosynthesis in the seed coat of Cleome hassleriana. Plant Cell 2020, 32, 3825–3845.
  3. Abdelaziz, O.Y.; Brink, D.P.; Prothmann, J.; Ravi, K.; Sun, M.; García-Hidalgo, J.; Sandahl, M.; Hulteberg, C.P.; Turner, C.; Lidén, G. Biological valorization of low molecular weight lignin. Biotechnol. Adv. 2016, 34, 1318–1346.
  4. Ayuso-Fernández, I.; Rencoret, J.; Gutiérrez, A.; Ruiz-Dueñas, F.J.; Martínez, A.T. Peroxidase evolution in white-rot fungi follows wood lignin evolution in plants. Proc. Natl. Acad. Sci. USA 2019, 116, 17900–17905.
  5. Wang, C.; Kelley, S.S.; Venditti, R.A. Lignin-based thermoplastic materials. ChemSusChem 2016, 9, 770–783.
  6. Kazzaz, A.E.; Feizi, Z.H.; Fatehi, P. Grafting strategies for hydroxy groups of lignin for producing materials. Green Chem. 2019, 21, 5714–5752.
  7. Meister, J.J. Modification of lignin. J. Macromol. Sci. Part C Polym. Rev. 2002, 42, 235–289.
  8. Ruwoldt, J. A critical review of the physicochemical properties of lignosulfonates: Chemical structure and behavior in aqueous solution, at surfaces and interfaces. Surfaces 2020, 3, 622–648.
  9. Kienberger, M.; Maitz, S.; Pichler, T.; Demmelmayer, P. Systematic review on isolation processes for technical lignin. Processes 2021, 9, 804.
  10. Demuner, I.F.; Gomes, F.J.B.; Gomes, J.S.; Coura, M.R.; Borges, F.P.; Carvalho, A.M.M.L.; Silva, C.M. Improving kraft pulp mill sustainability by lignosulfonates production from processes residues. J. Clean. Prod. 2021, 317, 128286.
  11. Hanhikoski, S.; Niemelä, K.; Vuorinen, T. Biorefining of Scots pine using neutral sodium sulphite pulping: Investigation of fibre and spent liquor compositions. Ind. Crops Prod. 2019, 129, 135–141.
  12. Tribot, A.; Amer, G.; Alio, M.A.; de Baynast, H.; Delattre, C.; Pons, A.; Mathias, J.-D.; Callois, J.-M.; Vial, C.; Michaud, P. Wood-lignin: Supply, extraction processes and use as bio-based material. Eur. Polym. J. 2019, 112, 228–240.
  13. Guadix-Montero, S.; Sankar, M. Review on catalytic cleavage of C–C inter-unit linkages in lignin model compounds: Towards lignin depolymerisation. Top. Catal. 2018, 61, 183–198.
  14. Wörmeyer, K.; Ingram, T.; Saake, B.; Brunner, G.; Smirnova, I. Comparison of different pretreatment methods for lignocellulosic materials. Part II: Influence of pretreatment on the properties of rye straw lignin. Bioresour. Technol. 2011, 102, 4157–4164.
  15. Vishtal, A.G.; Kraslawski, A. Challenges in industrial applications of technical lignins. BioResources 2011, 6, 3547–3568.
  16. Wang, H.; Pu, Y.; Ragauskas, A.; Yang, B. From lignin to valuable products–strategies, challenges, and prospects. Bioresour. Technol. 2019, 271, 449–461.
  17. Chen, Z.; Wan, C. Biological valorization strategies for converting lignin into fuels and chemicals. Renew. Sustain. Energy Rev. 2017, 73, 610–621.
  18. Ren, T.; Qi, W.; Su, R.; He, Z. Promising techniques for depolymerization of lignin into value-added chemicals. ChemCatChem 2019, 11, 639–654.
  19. Pang, B.; Lam, S.S.; Shen, X.J.; Cao, X.F.; Liu, S.J.; Yuan, T.Q.; Sun, R.C. Valorization of technical lignin for the production of desirable resins with high substitution rate and controllable viscosity. ChemSusChem 2020, 13, 4446–4454.
  20. Qian, Y.; Zuo, C.; Tan, J.; He, J. Structural analysis of bio-oils from sub-and supercritical water liquefaction of woody biomass. Energy 2007, 32, 196–202.
  21. Kloekhorst, A.; Heeres, H.J. Catalytic hydrotreatment of alcell lignin using supported Ru, Pd, and Cu catalysts. ACS Sustain. Chem. Eng. 2015, 3, 1905–1914.
  22. Morpurgo, L.; Graziani, M.; Finazzi-Agro, A.; Rotilio, G.; Mondovi, B. Optical properties of japanese-lacquer-tree (Rhus vernicifera) laccase depleted of type 2 copper (II). Involvement of type-2 copper (II) in the 330nm chromophore. Biochem. J. 1980, 187, 361–366.
  23. Claus, H. Laccases and their occurrence in prokaryotes. Arch. Microbiol. 2003, 179, 145–150.
  24. Mandic, M.; Djokic, L.; Nikolaivits, E.; Prodanovic, R.; O’Connor, K.; Jeremic, S.; Topakas, E.; Nikodinovic-Runic, J. Identification and characterization of new laccase biocatalysts from Pseudomonas species suitable for degradation of synthetic textile dyes. Catalysts 2019, 9, 629.
  25. Chen, M.; Zeng, G.; Lai, C.; Li, J.; Xu, P.; Wu, H. Molecular basis of laccase bound to lignin: Insight from comparative studies on the interaction of Trametes versicolor laccase with various lignin model compounds. RSC Adv. 2015, 5, 52307–52313.
  26. Perna, V.; Agger, J.W.; Andersen, M.L.; Holck, J.; Meyer, A.S. Laccase induced lignin radical formation kinetics evaluated by electron paramagnetic resonance spectroscopy. ACS Sustain. Chem. Eng. 2019, 7, 10425–10434.
  27. Gutiérrez, A.; Rencoret, J.; Cadena, E.M.; Rico, A.; Barth, D.; José, C.; Martínez, Á.T. Demonstration of laccase-based removal of lignin from wood and non-wood plant feedstocks. Bioresour. Technol. 2012, 119, 114–122.
  28. Braunschmid, V.; Binder, K.; Fuerst, S.; Subagia, R.; Danner, C.; Weber, H.; Schwaiger, N.; Nyanhongo, G.S.; Ribitsch, D.; Guebitz, G.M. Comparison of a fungal and a bacterial laccase for lignosulfonate polymerization. Process Biochem. 2021, 109, 207–213.
  29. Mattinen, M.-L.; Maijala, P.; Nousiainen, P.; Smeds, A.; Kontro, J.; Sipilä, J.; Tamminen, T.; Willför, S.; Viikari, L. Oxidation of lignans and lignin model compounds by laccase in aqueous solvent systems. J. Mol. Catal. B: Enzym. 2011, 72, 122–129.
  30. Kawai, S.; Iwatsuki, M.; Nakagawa, M.; Inagaki, M.; Hamabe, A.; Ohashi, H. An alternative β-ether cleavage pathway for a non-phenolic β-O-4 lignin model dimer catalyzed by a laccase-mediator system. Enzym. Microb. Technol. 2004, 35, 154–160.
  31. Hilgers, R.; Vincken, J.-P.; Gruppen, H.; Kabel, M.A. Laccase/mediator systems: Their reactivity toward phenolic lignin structures. ACS Sustain. Chem. Eng. 2018, 6, 2037–2046.
  32. Bujanovic, B.; Ralph, S.; Reiner, R.; Hirth, K.; Atalla, R. Polyoxometalates in oxidative delignification of chemical pulps: Effect on lignin. Materials 2010, 3, 1888–1903.
  33. Rencoret, J.; Pereira, A.; del Río, J.C.; Martínez, A.N.T.; Gutiérrez, A. Delignification and saccharification enhancement of sugarcane byproducts by a laccase-based pretreatment. ACS Sustain. Chem. Eng. 2017, 5, 7145–7154.
  34. Rencoret, J.; Pereira, A.; Del Río, J.C.; Martínez, A.T.; Gutiérrez, A. Laccase-mediator pretreatment of wheat straw degrades lignin and improves saccharification. BioEnergy Res. 2016, 9, 917–930.
  35. Xie, B.; Tobimatsu, Y.; Kamitakahara, H.; Takano, T. Reaction Selectivity in Electro-oxidation of Lignin Dimer Model Compounds and Synthetic Lignin with Different Mediators for the Laccase Mediator System (PZH, NHPI, ABTS). ACS Sustain. Chem. Eng. 2022, 10, 6633–6641.
  36. Kumar, A.; Chandra, R. Ligninolytic enzymes and its mechanisms for degradation of lignocellulosic waste in environment. Heliyon 2020, 6, e03170.
  37. Guo, M.; Lu, F.; Du, L.; Pu, J.; Bai, D. Optimization of the expression of a laccase gene from Trametes versicolor in Pichia methanolica. Appl. Microbiol. Biotechnol. 2006, 71, 848–852.
  38. Eggert, C.; Temp, U.; Eriksson, K.-E.L. Laccase is essential for lignin degradation by the white-rot fungus Pycnoporus cinnabarinus. Febs Lett. 1997, 407, 89–92.
  39. Kawai, S.; Nakagawa, M.; Ohashi, H. Degradation mechanisms of a nonphenolic β-O-4 lignin model dimer by Trametes versicolor laccase in the presence of 1-hydroxybenzotriazole. Enzym. Microb. Technol. 2002, 30, 482–489.
  40. Arias, M.; Blánquez, A.; Hernandez, M.; Rodriguez, J.; Ball, A.; Jiménez-Morillo, N.; González-Vila, F.; González-Pérez, J. Role of a thermostable laccase produced by Streptomyces ipomoeae in the degradation of wheat straw lignin in solid state fermentation. J. Anal. Appl. Pyrolysis 2016, 122, 202–208.
  41. Guan, Z.-B.; Shui, Y.; Song, C.-M.; Zhang, N.; Cai, Y.-J.; Liao, X.-R. Efficient secretory production of CotA-laccase and its application in the decolorization and detoxification of industrial textile wastewater. Environ. Sci. Pollut. Res. 2015, 22, 9515–9523.
  42. Shi, X.; Liu, Q.; Ma, J.; Liao, H.; Xiong, X.; Zhang, K.; Wang, T.; Liu, X.; Xu, T.; Yuan, S. An acid-stable bacterial laccase identified from the endophyte Pantoea ananatis Sd-1 genome exhibiting lignin degradation and dye decolorization abilities. Biotechnol. Lett. 2015, 37, 2279–2288.
  43. Asina, F.; Brzonova, I.; Voeller, K.; Kozliak, E.; Kubátová, A.; Yao, B.; Ji, Y. Biodegradation of lignin by fungi, bacteria and laccases. Bioresour. Technol. 2016, 220, 414–424.
  44. Colao, M.C.; Lupino, S.; Garzillo, A.M.; Buonocore, V.; Ruzzi, M. Heterologous expression of lcc1 gene from Trametes trogii in Pichia pastoris and characterization of the recombinant enzyme. Microb. Cell Factories 2006, 5, 31.
  45. Bo, W.; Yan, Y.; Xu, J.; Fu, X.; Han, H.; Gao, J.; Li, Z.; Wang, L.; Tian, Y.; Peng, R. Heterologous expression and characterization of a laccase from Laccaria bicolor in Pichia pastoris and Arabidopsis thaliana. J. Microbiol. Biotechnol. 2018, 28, 2057–2063.
  46. Zhu, D.; Zhang, P.; Xie, C.; Zhang, W.; Sun, J.; Qian, W.-J.; Yang, B. Biodegradation of alkaline lignin by Bacillus ligniniphilus L1. Biotechnol. Biofuels 2017, 10, 44.
  47. Kirk, T.K.; Croan, S.; Tien, M.; Murtagh, K.E.; Farrell, R.L. Production of multiple ligninases by Phanerochaete chrysosporium: Effect of selected growth conditions and use of a mutant strain. Enzym. Microb. Technol. 1986, 8, 27–32.
  48. Leisola, M.; Kozulic, B.; Meussdoerffer, F.; Fiechter, A. Homology among multiple extracellular peroxidases from Phanerochaete chrysosporium. J. Biol. Chem. 1987, 262, 419–424.
  49. Fujii, K.; Uemura, M.; Hayakawa, C.; Funakawa, S.; Kosaki, T. Environmental control of lignin peroxidase, manganese peroxidase, and laccase activities in forest floor layers in humid Asia. Soil Biol. Biochem. 2013, 57, 109–115.
  50. Sayadi, S.; Ellouz, R. Roles of lignin peroxidase and manganese peroxidase from Phanerochaete chrysosporium in the decolorization of olive mill wastewaters. Appl. Environ. Microbiol. 1995, 61, 1098–1103.
  51. Wang, X.; Yao, B.; Su, X. Linking enzymatic oxidative degradation of lignin to organics detoxification. Int. J. Mol. Sci. 2018, 19, 3373.
  52. Calvo-Flores, F.G.; Dobado, J.A. Lignin as renewable raw material. ChemSusChem 2010, 3, 1227–1235.
  53. Sung, H.J.; Khan, M.F.; Kim, Y.H. Recombinant lignin peroxidase-catalyzed decolorization of melanin using in-situ generated H2O2 for application in whitening cosmetics. Int. J. Biol. Macromol. 2019, 136, 20–26.
  54. Majeke, B.; Collard, F.-X.; Tyhoda, L.; Görgens, J. The synergistic application of quinone reductase and lignin peroxidase for the deconstruction of industrial (technical) lignins and analysis of the degraded lignin products. Bioresour. Technol. 2021, 319, 124152.
  55. Liu, E.; Segato, F.; Wilkins, M.R. Fed-batch production of Thermothelomyces thermophilus lignin peroxidase using a recombinant Aspergillus nidulans strain in stirred-tank bioreactor. Bioresour. Technol. 2021, 325, 124700.
  56. Kuwahara, M.; Glenn, J.K.; Morgan, M.A.; Gold, M.H. Separation and characterization of two extracelluar H2O2-dependent oxidases from ligninolytic cultures of Phanerochaete chrysosporium. FEBS Lett. 1984, 169, 247–250.
  57. Wariishi, H.; Valli, K.; Gold, M.H. Oxidative cleavage of a phenolic diarylpropane lignin model dimer by manganese peroxidase from Phanerochaete chrysosporium. Biochemistry 1989, 28, 6017–6023.
  58. Wang, Y.; Vazquez-Duhalt, R.; Pickard, M.A. Purification, characterization, and chemical modification of manganese peroxidase from Bjerkandera adusta UAMH 8258. Curr. Microbiol. 2002, 45, 77–87.
  59. Hofrichter, M. Lignin conversion by manganese peroxidase (MnP). Enzym. Microb. Technol. 2002, 30, 454–466.
  60. Masarin, F.; Norambuena, M.; Ramires, H.O.; Demuner, B.J.; Pavan, P.C.; Ferraz, A. Manganese peroxidase and biomimetic systems applied to in vitro lignin degradation in Eucalyptus grandis milled wood and kraft pulps. J. Chem. Technol. Biotechnol. 2016, 91, 1422–1430.
  61. Min, K.; Kim, Y.H.; Kim, J.; Kim, Y.; Gong, G.; Um, Y. Effect of manganese peroxidase on the decomposition of cellulosic components: Direct cellulolytic activity and synergistic effect with cellulase. Bioresour. Technol. 2022, 343, 126138.
  62. Ali, S.S.; Al-Tohamy, R.; Khalil, M.A.; Ho, S.-H.; Fu, Y.; Sun, J. Exploring the potential of a newly constructed manganese peroxidase-producing yeast consortium for tolerating lignin degradation inhibitors while simultaneously decolorizing and detoxifying textile azo dye wastewater. Bioresour. Technol. 2022, 351, 126861.
  63. Camarero, S.; Sarkar, S.; Ruiz-Dueñas, F.J.; Martínez, M.J.; Martínez, A.T. Description of a versatile peroxidase involved in the natural degradation of lignin that has both manganese peroxidase and lignin peroxidase substrate interaction sites. J. Biol. Chem. 1999, 274, 10324–10330.
  64. Sáez-Jiménez, V.; Acebes, S.; Guallar, V.; Martínez, A.T.; Ruiz-Dueñas, F.J. Improving the oxidative stability of a high redox potential fungal peroxidase by rational design. PLoS ONE 2015, 10, e0124750.
  65. Chan, J.C.; Paice, M.; Zhang, X. Enzymatic oxidation of lignin: Challenges and barriers toward practical applications. ChemCatChem 2020, 12, 401–425.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
Feedback