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    Topic review

    Textile Dye Biodecolorization by MnP

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    Submitted by: Tao Wang
    (This entry belongs to Entry Collection "Environmental Sciences ")

    Definition

    Manganese peroxidase (MnP) is an oxidoreductase with ligninolytic activity and is a promising biocatalyst for the biodegradation of hazardous environmental contaminants, and especially for dye wastewater decolorization.

    1. Introduction

    The textile industry produces large quantities of wastewater containing different types of dyes used during the dyeing process, which cause great harm to the environment [1][2]. Many dyes and their intermediate metabolites have been identified as mutagenic, teratogenic, or carcinogenic, and represent serious health threats to living ecosystems [3].
    At present, the treatment of dye wastewater mainly relies on physical or chemical management techniques, including chemical reduction, adsorption, ionizing radiation, precipitation, flocculation and flotation, membrane filtration, electric coagulation, electrochemical destruction, and ion exchange ozonation [4][5]. These technologies have obvious shortcomings such as the excessive use of chemicals, sludge production, expensive factory requirements or high operating expenses, low decolorization efficiencies, and the inability to handle large numbers of dyes with different structures, so they are not economically suitable for large-scale wastewater decolorization [6].
    The current focus is to reduce toxicity and develop an efficient, economical, and green dye detoxification and decolorization technology. Compared with physical and chemical methods, biological methods offer beneficial and effective prospects due to their economical and environmentally friendly advantages, as well as being simple to use, safe, and efficient, with no secondary pollution [7][8]. Therefore, biotechnology is considered the best choice to degrade and remove these pollutants effectively. In the biotechnology field, enzyme biocatalysis is currently the main research area due to its broad application prospects [9][10].
    Manganese peroxidases (EC 1.11.1.13; MnPs) are a family of heme-containing glycoproteins belonging to the oxidoreductase group. It was discovered in Phanerochaete chrysosporium and is also found in many bacteria and white-rot fungi (WRF) [11][12][13][14]. There are different MnPs in nature with differentiated properties. For example, long and short MnPs were reported in WRF associated with the presence/absence of the C-terminal tail extension, and these showed different catalytic and stability properties [15]. According to the residues of the Mn2+-binding site, three novel subfamilies of MnP were described in Agaricales including MnP-ESD (Glu/Ser/Asp Mn2+-oxidation site), MnP-DGD (Asp/Gly/Asp Mn2+-oxidation site), and MnP-DED (Asp/Glu/Asp Mn2+-oxidation site) [16]. However, the Mn2+-binding site is not the unique feature of MnPs, because versatile peroxidases (VPs), which evolved directly from MnPs, also possess such a site and can oxidize Mn2+ to Mn3+ [17].
    For enzyme applications, MnPs can catalyze the peroxide-dependent degradation of a variety of toxic dye pollutants, phenolic compounds, antibiotics, and polycyclic aromatic hydrocarbons, so are promising biocatalysts for hazardous environmental contaminants biodegradation [18][19]. Moreover, the use of MnPs is suitable for dye wastewater decolorization as the process is simple and the enzyme can be recycled, thus reducing operating costs [20][21][22].

    2. The Crystal Structure of MnPs

    The crystal structure of an enzyme provides information on the catalytic mechanism and for potential in-depth design and transformation, and for realizing the green biotechnological use of enzymes [23][24][25].
    The heme conformation of MnP is similar to that of lignin peroxidase (LiP) and is evolutionarily conserved [26]. In its resting-state form, MnP is a strongly helical protein containing a Fe3+ penta-coordinated structure with the porphyrin ring of the heme cofactor and a proximal histidine, with the sixth coordination position open for H2O2 [27].
    To date, several crystal structures of MnP from different sources have been reported, and the highest-resolution crystal structures (~0.93Å) of MnP complexed with Mn2+ (Mn-MnP) are shown in Figure 1 [28]. The conserved Ca2+ ions are important for the stability of the protein [29]; these are indicated as gold yellow spheres and the position of the Mn2+ substrate is shown in violet. The active site is composed of three highly conserved amino acids (Glu35, Glu39, and Asp179) and one heme propionate. The Mn2+ substrate binds in the center of the active site, and the heme propionate (HEM) is located in the internal hydrophobic cavity of the enzyme. The spatial structure of HEM is further stabilized by four hydrogen bonds (green dashed line), two electrostatic interactions (orange dashed line), and some other weak interactions. The catalytic site of heme peroxidases is strongly conserved, with only minor variations occurring in the replacement of Phe with Trp in several enzymes such as ascorbate peroxidase and cytochrome c. The Asp–His pair (242 and 173, respectively) is also conserved.
    Figure 1. The overall structure (A), active site structure (B,C), and interaction mode (D) of Mn–MnP refined at 0.93 Å resolution [28]. PDB ID: 3M5Q.

    3. MnP Catalysis

    At the beginning of the catalytic cycle, H2O2 or organic peroxide binds to the enzyme in resting state in ferric (Fe3+) form (Figure 2). This process releases one molecule of H2O and forms MnP–compound I (Fe4+-oxo-porphyrin radical complex), with two oxidation equivalents. This oxidizes Mn2+ to Mn3+, forming MnP–compound II (Fe4+-oxo-porphyrin complex). Immediately afterwards, the MnP–compound II combines with Mn2+ in a similar manner to generate Mn3+, releasing one molecule of H2O, and is reduced to the original state of ferric MnP, completing the catalytic cycle [30].
    Figure 2. The MnP catalytic cycle [30].
    The MnP catalytic cycle resembles that of other lignin and heme peroxidases in the presence of native Fe3+ enzymes and two reactive intermediates [31]. However, in contrast to other peroxidases, MnP preferentially uses Mn2+ as the substrate, converting it to the strong oxidation state of Mn3+ through a series of redox reactions [32].

    4. Application of Unmodified MnPs in the Decolorization of Dye Wastewater

    Table 1 contains a summary of recent studies on the breakdown and decolorization of textile-derived dye compounds by microbial MnPs.
    Table 1. Recent applications of unmodified MnPs in dye decolorization.

    Source

    Types of Dyes

    Initial Concentration of Dyes

    Removal Rate

    Time Cost

    Reference

    Microbial consortium SR

    Crystal Violet

    20 mg/L

    63%

    6 days

    [20]

    Cresol Red

    100 mg/L

    93%

    CBB G250

    100 mg/L

    96%

    Trametes pubescens strain i8

    Acid Blue 158

    50 μM

    95%

    24 h

    [22]

    Poly R-478

    88%

    Remazol Brilliant Violet 5R

    76%

    Direct Red 5B

    66%

    Indigo Carmine

    64%

    Methyl Green

    50%

    Cibacet Brilliant Blue BG

    46%

    Remazol Brilliant Blue Reactif

    42%

    Aspergillus terreus GS28

    Direct Blue-1

    100 mg/L

    98.4%

    168 h

    [33]

    Bjerkandera adusta strain CX-9

    Acid Blue 158

    50 μM

    91%

    12 h

    [34]

    Poly R-478

    80%

    Cibacet Brilliant Blue BG

    77%

    Remazol Brilliant Violet 5R

    70%

    Trametes sp.48424

    Indigo Carmine

    100 mg/L

    94.6%

    18 h

    [35]

    Remazol Brilliant Blue R

    85.0%

    Remazol Brilliant Violet 5R

    88.4%

    Methyl Green

    93.1%

    Microbial consortium ZSY

    Metanil Yellow G

    100 mg/L

    93.39%

    48 h

    [36]

    Microbial Consortium ZW1

    Methanil Yellow G

    100 mg/L

    93.3%

    16 h

    [37]

    Trichoderma harzianum

    Alizarin Blue Black B

    0.03%

    92.34%

    14 days

    [38]

    Phanerochaete chrysosporium CDBB 686

    Congo Red

    50 ppm

    41.84%

    36 h

    [39]

    Poly R-478

    56.86%

    Methyl Green

    69.79%

    Bjerkandera adusta CCBAS 930

    Alizarin Blue Black B

    0.01%

    86.5%

    20 days

    [40]

    Acid Blue 129

    89.22%

    Cerrena unicolor BBP6

    Congo Red

    100 mg/L

    53.9%

    12 h

    [41]

    Methyl Orange

    77.6%

    12 h

    Remazol Brilliant Blue R

    81.0%

    5 h

    Bromophenol Blue

    62.2%

    12 h

    Crystal Violet

    80.9%

    12 h

    Azure Blue

    63.1%

    24 h

    Phanerochaete chrysosporium

    Indigo Carmine

    30 mg/L

    90.18%

    6 h

    [42]

    Trametes versicolor

    Dye mixture

    (Brilliant Blue FCF

    and

    Allura Red AC)

    100 mg/L

    80.45%

    14 days

    [43]

    Irpex lacteus

    86.04%

    19 days

    Bjerkandera adusta

    82.83%

    9 days

    Ceriporia lacerata ZJSY

    Congo Red

    100 mg/L

    90%

    48 h

    [44]

    Bacillus cohnni RKS9

    Congo Red

    100 mg/L

    99%

    12 h

    [45]

    Schizophyllum commune IBL-06

    Solar Brilliant Red 80

    0.01%

    100%

    3 days

    [46]

    Irpex lacteus CD2

    Remazol Brilliant Violet 5R

    50 mg/L

    92.8%

    5 h

    [47]

    Remazol Brilliant Blue R

    87.1%

    5 h

    Indigo Carmine

    91.5%

    5 h

    Direct Red 5B

    82.4%

    36 h

    The entry is from 10.3390/molecules26154403

    References

    1. Verma, A.K.; Dash, R.R.; Bhunia, P. A review on chemical coagulation/flocculation technologies for removal of colour from textile wastewaters. J. Environ. Manag. 2012, 93, 154–168.
    2. Kishor, R.; Purchase, D.; Saratale, G.D.; Saratale, R.G.; Ferreira, L.F.R.; Bilal, M.; Chandra, R.; Bharagava, R.N. Ecotoxicological and health concerns of persistent coloring pollutants of textile industry wastewater and treatment approaches for environmental safety. J. Environ. Chem. Eng. 2021, 9, 105012.
    3. Tkaczyk, A.; Mitrowska, K.; Posyniak, A. Synthetic organic dyes as contaminants of the aquatic environment and their implications for ecosystems: A review. Sci. Total. Environ. 2020, 717, 137222.
    4. Selvaraj, V.; Karthika, T.S.; Mansiya, C.; Alagar, M. An over review on recently developed techniques, mechanisms and intermediate involved in the advanced azo dye degradation for industrial applications. J. Mol. Struct. 2021, 1224, 129195.
    5. Ihsanullah, I.; Jamal, A.; Ilyas, M.; Zubair, M.; Khan, G.; Atieh, M.A. Bioremediation of dyes: Current status and prospects. J. Water Process. Eng. 2020, 38, 101680.
    6. Katheresan, V.; Kansedo, J.; Lau, S.Y. Efficiency of various recent wastewater dye removal methods: A review. J. Environ. Chem. Eng. 2018, 6, 4676–4697.
    7. Ghosh, A.; Dastidar, M.G.; Sreekrishnan, T. Bioremediation of chromium complex dyes and treatment of sludge generated during the process. Int. Biodeterior. Biodegrad. 2017, 119, 448–460.
    8. Kishor, R.; Saratale, G.D.; Saratale, R.G.; Ferreira, L.F.R.; Bilal, M.; Iqbal, H.M.; Bharagava, R.N. Efficient degradation and detoxification of methylene blue dye by a newly isolated ligninolytic enzyme producing bacterium Bacillus Albus MW407057. Colloids Surf. B Biointerfaces 2021, 206, 111947.
    9. Gurung, N.; Ray, S.; Bose, S.; Rai, V. A Broader View: Microbial Enzymes and Their Relevance in Industries, Medicine, and Beyond. BioMed Res. Int. 2013, 2013, 329121.
    10. Vilar, D.D.S.; Bilal, M.; Bharagava, R.N.; Kumar, A.; Nadda, A.K.; Salazar-Banda, G.R.; Eguiluz, K.I.B.; Ferreira, L.F.R. Lignin-modifying enzymes: A green and environmental responsive technology for organic compound degradation. J. Chem. Technol. Biotechnol. 2021, 96, 6751.
    11. 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.
    12. Glenn, J.K.; Gold, M.H. Purification and characterization of an extracellular Mn(II)-dependent peroxidase from the lignin-degrading basidiomycete, Phanerochaete chrysosporium. Arch. Biochem. Biophys. 1985, 242, 329–341.
    13. Paszczyński, A.; Huynh, V.B.; Crawford, R. Enzymatic activities of an extracellular, manganese-dependent peroxidase from Phanerochaete chrysosporium. FEMS Microbiol. Lett. 1985, 29, 37–41.
    14. Paszczyński, A.; Huynh, V.B.; Crawford, R. Comparison of ligninase-I and peroxidase-M2 from the white-rot fungus Phanerochaete chrysosporium. Arch. Biochem. Biophys. 1986, 244, 750–765.
    15. Fernández-Fueyo, E.; Acebes, S.; Ruiz-Dueñas, F.J.; Martínez, M.J.; Romero, A.; Medrano, F.J.; Guallar, V.; Martínez, A.T. Structural implications of the C-terminal tail in the catalytic and stability properties of manganese peroxidases from ligninolytic fungi. Acta Crystallogr. Sect. D Biol. Crystallogr. 2014, 70, 3253–3265.
    16. Ruiz-Dueñas, F.J.; Barrasa, J.M.; Sánchez-García, M.; Camarero, S.; Miyauchi, S.; Serrano, A.; Linde, D.; Babiker, R.; Drula, E.; Ayuso-Fernández, I.; et al. Genomic analysis enlightens Agaricales lifestyle evolution and increasing peroxidase diversity. Mol. Biol. Evol. 2021, 38, 1428–1446.
    17. Ayuso-Fernández, I.; Ruiz-Dueñas, F.J.; Martínez, A.T. Evolutionary convergence in lignin-degrading enzymes. Proc. Natl. Acad. Sci. USA 2018, 115, 6428–6433.
    18. Singh, A.K.; Bilal, M.; Iqbal, H.M.; Meyer, A.S.; Raj, A. Bioremediation of lignin derivatives and phenolics in wastewater with lignin modifying enzymes: Status, opportunities and challenges. Sci. Total. Environ. 2021, 777, 145988.
    19. Bilal, M.; Bagheri, A.R.; Vilar, D.S.; Aramesh, N.; Eguiluz, K.I.B.; Ferreira, L.F.R.; Ashraf, S.S.; Iqbal, H.M. Oxidoreductases as a versatile biocatalytic tool to tackle pollutants for clean environment—A review. J. Chem. Technol. Biotechnol. 2021, 96, 1–44.
    20. Yang, X.; Wang, J.; Zhao, X.; Wang, Q.; Xue, R. Increasing manganese peroxidase production and biodecolorization of triphenylmethane dyes by novel fungal consortium. Bioresour. Technol. 2011, 102, 10535–10541.
    21. Bilal, M.; Asgher, M. Sandal reactive dyes decolorization and cytotoxicity reduction using manganese peroxidase immobilized onto polyvinyl alcohol-alginate beads. Chem. Cent. J. 2015, 9, 1–14.
    22. Rekik, H.; Jaouadi, N.Z.; Bouacem, K.; Zenati, B.; Kourdali, S.; Badis, A.; Annane, R.; Bouanane-Darenfed, A.; Bejar, S.; Jaouadi, B. Physical and enzymatic properties of a new manganese peroxidase from the white-rot fungus Trametes pubescens strain i8 for lignin biodegradation and textile-dyes biodecolorization. Int. J. Biol. Macromol. 2019, 125, 514–525.
    23. Engel, M.; Hoffmann, T.; Wagner, L.; Wermann, M.; Heiser, U.; Kiefersauer, R.; Huber, R.; Bode, W.; Demuth, H.-U.; Brandstetter, H. The crystal structure of dipeptidyl peptidase IV (CD26) reveals its functional regulation and enzymatic mechanism. Proc. Natl. Acad. Sci. USA 2003, 100, 5063–5068.
    24. Qiu, J.; Wilkens, C.; Barrett, K.; Meyer, A.S. Microbial enzymes catalyzing keratin degradation: Classification, structure, function. Biotechnol. Adv. 2020, 44, 107607.
    25. Chang, M.; Zhou, Y.; Wang, H.; Liu, Z.; Zhang, Y.; Feng, Y. Crystal structure of the multifunctional SAM-dependent enzyme LepI provides insights into its catalytic mechanism. Biochem. Biophys. Res. Commun. 2019, 515, 255–260.
    26. Ayuso-Fernández, I.; Martínez, A.T.; Ruiz-Dueñas, F.J. Experimental recreation of the evolution of lignin-degrading enzymes from the Jurassic to date. Biotechnol. Biofuels 2017, 10, 1–13.
    27. Blodig, W.; Smith, A.; Doyle, W.A.; Piontek, K. Crystal structures of pristine and oxidatively processed lignin peroxidase expressed in Escherichia coli and of the W171F variant that eliminates the redox active tryptophan 171. Implications for the reaction mechanism. J. Mol. Biol. 2001, 305, 851–861.
    28. Sundaramoorthy, M.; Gold, M.H.; Poulos, T.L. Ultrahigh (0.93A) resolution structure of manganese peroxidase from Phanerochaete chrysosporium: Implications for the catalytic mechanism. J. Inorg. Biochem. 2010, 104, 683–690.
    29. George, S.J.; Kvaratskhelia, M.; Dilworth, M.J.; Thorneley, R.N.F. Reversible alkaline inactivation of lignin peroxidase involves the release of both the distal and proximal site calcium ions and bishistidine co-ordination of the haem. Biochem. J. 1999, 344, 237–244.
    30. Hofrichter, M. Review: Lignin conversion by manganese peroxidase (MnP). Enzym. Microb. Technol. 2002, 30, 454–466.
    31. Wariishi, H.; Huang, J.; Dunford, H.; Gold, M. Reactions of lignin peroxidase compounds I and II with veratryl alcohol. Transient-state kinetic characterization. J. Biol. Chem. 1991, 266, 20694.
    32. Wariishi, H.; Valli, K.; Gold, M.H.; Wariishi, H.; Valli, K.; Gold, M.H. Manganese(II) oxidation by manganese peroxidase from the basidiomycete Phanerochaete chrysosporium. Kinetic mechanism and role of chelators. J. Biol. Chem. 1992, 267, 23688–23695.
    33. Singh, G.; Dwivedi, S. Decolorization and degradation of Direct Blue-1 (Azo dye) by newly isolated fungus Aspergillus terreus GS28, from sludge of carpet industry. Environ. Technol. Innov. 2020, 18, 100751.
    34. Bouacem, K.; Rekik, H.; Jaouadi, N.Z.; Zenati, B.; Kourdali, S.; El Hattab, M.; Badis, A.; Annane, R.; Bejar, S.; Hacene, H.; et al. Purification and characterization of two novel peroxidases from the dye-decolorizing fungus Bjerkandera adusta strain CX-9. Int. J. Biol. Macromol. 2018, 106, 636–646.
    35. Zhang, H.; Zhang, S.; He, F.; Qin, X.; Zhang, X.; Yang, Y. Characterization of a manganese peroxidase from white-rot fungus Trametes sp.48424 with strong ability of degrading different types of dyes and polycyclic aromatic hydrocarbons. J. Hazard. Mater. 2016, 320, 265–277.
    36. Guo, G.; Hao, J.; Tian, F.; Liu, C.; Ding, K.; Zhang, C.; Yang, F.; Xu, J. Decolorization of Metanil Yellow G by a halophilic alkalithermophilic bacterial consortium. Bioresour. Technol. 2020, 316, 123923.
    37. Guo, G.; Hao, J.; Tian, F.; Liu, C.; Ding, K.; Xu, J.; Zhou, W.; Guan, Z. Decolorization and detoxification of azo dye by halo-alkaliphilic bacterial consortium: Systematic investigations of performance, pathway and metagenome. Ecotoxicol. Environ. Saf. 2020, 204, 111073.
    38. Rybczyńska-Tkaczyk, K.; Święciło, A.; Szychowski, K.; Korniłłowicz-Kowalska, T. Comparative study of eco- and cytotoxicity during biotransformation of anthraquinone dye Alizarin Blue Black B in optimized cultures of microscopic fungi. Ecotoxicol. Environ. Saf. 2018, 147, 776–787.
    39. Sosa-Martínez, J.D.; Balagurusamy, N.; Montañez, J.; Peralta, R.A.; Moreira, R.D.F.P.M.; Bracht, A.; Peralta, R.M.; Morales-Oyervides, L. Synthetic dyes biodegradation by fungal ligninolytic enzymes: Process optimization, metabolites evaluation and toxicity assessment. J. Hazard. Mater. 2020, 400, 123254.
    40. Rybczyńska-Tkaczyk, K.; Korniłłowicz-Kowalska, T.; Szychowski, K.; Gmiński, J. Biotransformation and toxicity effect of monoanthraquinone dyes during Bjerkandera adusta CCBAS 930 cultures. Ecotoxicol. Environ. Saf. 2020, 191, 110203.
    41. Zhang, H.; Zhang, J.; Zhang, X.; Geng, A. Purification and characterization of a novel manganese peroxidase from white-rot fungus Cerrena unicolor BBP6 and its application in dye decolorization and denim bleaching. Process. Biochem. 2018, 66, 222–229.
    42. Li, H.; Zhang, R.; Tang, L.; Zhang, J.; Mao, Z. Manganese peroxidase production from cassava residue by Phanerochaete chrysosporium in solid state fermentation and its decolorization of indigo carmine. Chin. J. Chem. Eng. 2015, 23, 227–233.
    43. Merino-Restrepo, A.; Mejía, F.; Velásquez-Quintero, C.; Hormaza-Anaguano, A. Evaluation of several white-rot fungi for the decolorization of a binary mixture of anionic dyes and characterization of the residual biomass as potential organic soil amendment. J. Environ. Manag. 2020, 254, 109805.
    44. Wang, N.; Chu, Y.; Wu, F.; Zhao, Z.; Xu, X. Decolorization and degradation of Congo red by a newly isolated white rot fungus, Ceriporia lacerata, from decayed mulberry branches. Int. Biodeterior. Biodegrad. 2017, 117, 236–244.
    45. Kishor, R.; Purchase, D.; Saratale, G.D.; Ferreira, L.F.R.; Bilal, M.; Iqbal, H.M.; Bharagava, R.N. Environment friendly degradation and detoxification of Congo red dye and textile industry wastewater by a newly isolated Bacillus cohnni (RKS9). Environ. Technol. Innov. 2021, 22, 101425.
    46. Asgher, M.; Yasmeen, Q.; Iqbal, H.M.N. Enhanced decolorization of Solar brilliant red 80 textile dye by an indigenous white rot fungus Schizophyllum commune IBL-06. Saudi J. Biol. Sci. 2013, 20, 347–352.
    47. Xing, Q.; Zhang, J.; Zhang, X.; Yang, Y. Induction, Purification and Characterization of a Novel Manganese Peroxidase from Irpex lacteus CD2 and Its Application in the Decolorization of Different Types of Dye. PLoS ONE 2014, 9, e113282.
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