Title: Lignin peroxidase functionalities and prospective applications
Abstract: MicrobiologyOpenVolume 6, Issue 1 e00394 REVIEWOpen Access Lignin peroxidase functionalities and prospective applications Ayodeji O. Falade, Ayodeji O. Falade SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorUchechukwu U. Nwodo, Corresponding Author Uchechukwu U. Nwodo [email protected] SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South Africa Correspondence Uchechukwu U. Nwodo SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa. Email: [email protected] for more papers by this authorBenson C. Iweriebor, Benson C. Iweriebor SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorEzekiel Green, Ezekiel Green SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorLeonard V. Mabinya, Leonard V. Mabinya SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorAnthony I. Okoh, Anthony I. Okoh SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this author Ayodeji O. Falade, Ayodeji O. Falade SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorUchechukwu U. Nwodo, Corresponding Author Uchechukwu U. Nwodo [email protected] SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South Africa Correspondence Uchechukwu U. Nwodo SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa. Email: [email protected] for more papers by this authorBenson C. Iweriebor, Benson C. Iweriebor SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorEzekiel Green, Ezekiel Green SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorLeonard V. Mabinya, Leonard V. Mabinya SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this authorAnthony I. Okoh, Anthony I. Okoh SAMRC Microbial Water Quality Monitoring Centre, University of Fort Hare, Alice, South Africa Applied and Environmental Microbiology Research Group (AEMREG), Department of Biochemistry and Microbiology, University of Fort Hare, Alice, South AfricaSearch for more papers by this author https://doi.org/10.1002/mbo3.394Citations: 149AboutSectionsPDF ToolsExport citationAdd to favoritesTrack citationReprints ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinkedInRedditWechat Graphical Abstract Ligninolytic extracellular enzymes are topical owing to their high redox potential and prospective industrial applicability. The applications of peroxidases spans through several sectors including cosmetology, dermatology and bioremediation industries. The litany of potentials attributed to lignin peroxidases is consequent to their versatility in the decolourization of melanin (synthetic or otherwise) and as well, the degradation of xenobiotic with phenolic and non-phenolic constituents. Abstract Ligninolytic extracellular enzymes, including lignin peroxidase, are topical owing to their high redox potential and prospective industrial applications. The prospective applications of lignin peroxidase span through sectors such as biorefinery, textile, energy, bioremediation, cosmetology, and dermatology industries. The litany of potentials attributed to lignin peroxidase is occasioned by its versatility in the degradation of xenobiotics and compounds with both phenolic and non-phenolic constituents. Over the years, ligninolytic enzymes have been studied however; research on lignin peroxidase seems to have been lagging when compared to other ligninolytic enzymes which are extracellular in nature including laccase and manganese peroxidase. This assertion becomes more pronounced when the application of lignin peroxidase is put into perspective. Consequently, a succinct documentation of the contemporary functionalities of lignin peroxidase and, some prospective applications of futuristic relevance has been advanced in this review. Some articulated applications include delignification of feedstock for ethanol production, textile effluent treatment and dye decolourization, coal depolymerization, treatment of hyperpigmentation, and skin-lightening through melanin oxidation. Prospective application of lignin peroxidase in skin-lightening functions through novel mechanisms, hence, it holds high value for the cosmetics sector where it may serve as suitable alternative to hydroquinone; a potent skin-lightening agent whose safety has generated lots of controversy and concern. 1 Introduction The espousal for the utilization and perhaps, the utilization of lignocellulosic biomass for the production of value-added products is on the increase world over, partly, due to the abundance and renewable nature of lignocellulosic biomass. Woody and nonwoody plants possess lignocellulose as major structural components and two carbohydrate polymers viz. cellulose (~30–50%) and hemicelluloses (~15–30%) as well as some non-carbohydrate aromatic polymers (~15–30%) constitute lignocellulose (Foyle, Jennings, & Mulcahy, 2007; Harris & Debolt, 2010; Menon & Rao, 2012). In woody or herbaceous plants, the lignocellulose constituent varies in accordance with the species and in tandem with the biotic and abiotic stressor factors including environmental distress syndromes. Several categorizations has applied to lignocellulosic biomass including waste biomass, virgin biomass, and energy crops. Waste biomass are thought to be low-value by-products largely generated from industrialized forestry activities (sawdust, wood waste, pulp mill waste), agricultural practices (corn stover/cob, sugarcane bagasse, wheat straw, rice husks, animal droppings), and municipal solid waste. On the other hand, terrestrial plants are classified as virgin biomass while energy crops include those generating large amount of lignocellulosic biomass as feedstock for second-generation biofuel production. Increased generation of lignocellulosic wastes from both the industrial and agricultural sectors have continued to pose environmental challenge globally due to, in part, poor waste management. However, the prospect of the valorization of lignocellulosic wastes for value-added products shall suffice as effective waste management strategies. Nonetheless, this proposition is at the moment underexplored. Besides, valorization of lignocellulosic wastes avails the right set of circumstance for the harnessing of value-added products from the compositional structures of the lignocellulosic biomass. The valorized may be, in effect, the products of interest for end users, otherwise, they may serve as raw materials for the production of commercially viable products. On the converse, microbial activities on lignocellulosic biomass during valorization process may also generate industrially important products including enzymes, ethanol, organic acids, microbial polysaccharides, and vitamins. The interest in enzymes is informed by their enormous industrial and biotechnological applications. However, the cost of enzyme production has continued to increase; thus, the need for a cost-effective means of production is imperative. Nonetheless, an important cost reduction strategy for enzyme production includes the exploration of alternative cheap carbon sources for fermentation, and lignocellulosic biomass may serve the purpose of cheap carbon source. Abundance, availability, and renewable nature bestow lignocellulosic biomass the status of near-perfect candidature of a cheap carbon source. Consequently, a variety of lignocellulosic materials have been utilized for different enzyme production processes (Asgher, Iqbal, & Asad, 2012; Asgher, Iqbal, & Irshad, 2012; Kang, Park, Lee, Hong, & Kim, 2004; Knezevic, Milovanovic, Stajic, & Vakojevic, 2013; Reddy, Babu, Komaraiah, Roy, & Kothari, 2003), and a conspectus of some of these processes has been articulated in Table 1. Table 1. Valorization of some lignocellulosic biomass for ligninolytic and cellulolytic enzyme production Lignocellulosic Biomass Microorganism Enzymes Produced References Rice straw P. chrysosporium; T. versicolor; Trichoderma reesei; Aspergillus niger KK2 LiP, MnP, Laccase, Cellulases, Hemicellulases Kang et al. (2004), Iqbal, Ahmed, Zia, and Irfan (2011), Asgher, Ahmad, and Iqbal (2011), Saratale et al. (2014) Sugarcane bagasse Thermoascus aurantiacus; Bacillus circulans; Trametes villosa Xylanase, MnP Milagres, Santos, Piovan, and Roberto (2004), Bocchini, Oliveira, Gomes, and Da Silva (2005), Silva et al. (2014) Wheat straw Phlebia radiata; Trichoderma viride; Trametes suaveolens LiP, MnP, Laccase, Cellulase Vares, Kalsi, and Hatakka (1995), Iqbal et al. (2011), Knezevic et al. (2013) Banana waste P. ostreatus; P. sajor-caju; Schizophyllum commune IBL-06 LiP, MnP, Laccase, Xylanase, Endoglucanase, Exoglucanase Reddy et al. (2003), Irshad and Asgher (2011), Asgher, Irshad, and Iqbal (2012) Corn cobs Trametes versicolor LiP, MnP, Laccase Asgher, Iqbal, & Asad (2012); Asgher, Iqbal, & Irshad (2012) Sawdust Trametes suaveolens MnP, Laccase Knezevic et al. (2013) Pea pods Aspergillus niger HN-1 Filter Paper Cellulase (FPase)β- glucosidase (BGL) Sharma, Rawat, Bhogal, and Oberoi (2015) LiP, Lignin Peroxidase; MnP, Manganese Peroxidase. Irrespective of any delineated path that may lead to products of interest, lignin recalcitrance to degradation has remained a major bottleneck to various industrial operations. Besides the conferral of hydrolytic stability and structural rigidity to plant's cell walls, lignin traps and renders unavailable the saccharides constituting the mono-, di-, oligo- and poly-meric units of cellulose necessary for fermentation. Lignin is imperative for the survival of plants and its recalcitrance to degradation has been attributed to its cross linkages with polysaccharides (cellulose and hemicellulose) via ester and ether linkages and as well as, its molecular architecture, in which various non-phenolic phenylpropanoid uints produce a complicated three-dimensional network joined by an array of ether and carbon–carbon bonds (Ruiz-Dueñas & Martinez, 2009). In a bid to address the challenge of lignin recalcitrance to degradation, several physicochemical pretreatment technologies have been developed to disrupt the non-cellulosic matrix and render cellulose and hemicellulose more accessible for enzymatic hydrolysis (Mosier et al., 2005). Some of these methods include steam explosion, ammonia fiber explosion, acid hydrolysis, alkaline hydrolysis, ozonolysis, organosolvation, and oxidative delignification (Chaturvedi & Verma, 2013). These pretreatment technologies are generally expensive, require high energy inputs, generate compounds inhibitory to fermentation, releases toxic chemicals which leads to corrosion problems and may also lead to material loss (Chaturvedi & Verma, 2013; Huang, Santhanam, Badri, & Hunte, 2013). Nonetheless, biological method of delignification may serve as an alternative pretreatment process as it is saddled with fewer limitations (Huang et al., 2013; Kuhar, Nair, & Kuhad, 2008). Biological pretreatment involves the use of microorganisms or immobilized microbial submolecules such as enzymes. The method may be thought of as cheap and environmental-friendly. However, it is not without demerits which include utilization of, part of, the fermentable sugars as carbon source consequently, lowering product yield (Potumarthi, Baadhe, Nayak, & Jetty, 2013; Wan & Li, 2011). Lignin degradation has been extensively studied in wood-rotting organisms, especially white-rot basidiomycetes (Hatakka, 1994; Leonowicz et al., 1999; Martinez et al., 2004; Wan & Li, 2012), and most of these studies established white-rot fungi as the most effective “delignifyer” partly as a result of the potent ligninolytic extracellular oxidative enzymes (ligninases) produced (Glenn, Morgan, Mayfield, Kuwahara, & Gold, 1983; Tien & Kirk, 1983). The ligninolytic extracellular oxidative enzymes have been classified into phenol oxidases and heme peroxidases. Enzymes in the phenol oxidases include laccases (EC 1.10.3.2) while the heme peroxidases include lignin peroxidase (EC 1.11.1.14), manganese peroxidase (EC 1.11.1.13), versatile peroxidase (EC 1.11.1.16), and dyP-type peroxidases (EC 1.11.1.19). Also implicated in the degradation of lignin are some accessory enzymes such as aryl-alcohol oxidase (EC 1.1.3.7), glyoxal oxidase (EC 1.2.3.5), and glucose 1-oxidase (EC 1.1.3.4) which generate the hydrogen peroxide (H2O2) required by the peroxidases (Ander & Marzullo, 1997; Guillen, Martinez, & Martinez, 1992; Kersten & Kirk, 1987). Lignin-modifying enzymes (LMEs) have also shown capability toward the degradation of various xenobiotics including dyes, chlorophenols, polycyclic aromatic hydrocarbons (PAHs), organophosphorous compounds, and phenols (Wesenberg, Kyriakides, & Agathos, 2003). The high redox potentials of ligninases and their ability to oxidize materials recalcitrant to degradation motivates for their prospects in biopulping and biobleaching (Call & Call, 2005), bioremediation through textile dye transformation (Husain, 2010; Mehta, 2012), decolourization of distillery effluent and other waste effluent treatment (Rajasundari & Murugesan, 2011) and as well, the degradation of herbicides (Pizzul, Castillo, & Stenström, 2009). Consequently, the interest in the application of ligninases for biotechnological purposes continues and the imperativeness of the industrial potential is an indication of the value of these enzymes. This review, therefore, succinctly presents the functionalities of lignin peroxidase and also prospects into futuristic applications. 2 Peroxidases (EC 1.11.1) Peroxidases catalyze the oxidation of various organic and inorganic substrates in the presence of hydrogen peroxide as electron acceptor; typical a reaction is as shown below. S, substrate (electron donor), Sox, oxidized substrate. Peroxidases are distributed widely in nature; vast presence in plants, animals, and microbes have been documented (Battistuzzi, Bellei, Bortolotti, & Sola, 2010). They are grouped as heme and non-heme peroxidases. The heme peroxidases contain a protoporphyrin IX (heme) as prosthetic group while the non-heme peroxidases lack such prosthetic group. A recent classification phylogenetically divides heme peroxidases into two superfamilies (peroxidase-cyclooxygenase superfamily and peroxidase-catalase superfamily) and three families including di-heme peroxidases, dyP-type peroxidases (DyPPrx), and haloperoxidases (HalPrx), respectively (Zamocky & Obinger, 2010). The Peroxidase-cyclooxygenase superfamily is made up of members from all domains of life (Zamocky et al., 2015) as against the old nomenclature “animal heme-dependent peroxidases” which formerly restricted classification to only peroxidases of animal origin. They seem to dominantly catalyze halide oxidation (Zamocky et al., 2015). Several representatives of peroxidase-cyclooxygenase superfamily are involved in the innate immune system (Söderhall, 1999). This function is not restricted to mammalian peroxidases alone as several peroxidases of bacterial origin (Dick et al., 2008) are suspected to be involved in unspecific defense mechanisms (Zamocky & Obinger, 2010). The involvement of the peroxidase-cyclooxygenase superfamily in immunology would be of clinical significance. However, this is out of scope of this present review. The peroxidase-catalase superfamily may be further subdivided into three classes (Welinder, 1992). Class I involve intracellular peroxidases such as yeast cytochrome c peroxidase (CcP) which protects against toxic peroxide in the electron transport chain (Dunford, 1999), and recent evidences also suggest that this peroxidase functions as a mitochondrial peroxide sensing and signaling protein in Saccharomyces cerevisiae (Martins, Kathiresan, & English, 2013). Ascorbate peroxidase (APx) comes next, and it is associated with the removal of hydrogen peroxide in the chloroplast and cytosol of higher plants (Battistuzzi et al., 2010; Dunford, 1999) and lastly, the bacterial catalase-peroxidase (KatG) which is known to exhibit hybrid catalytic activities of both peroxidase and catalase and, they are thought to have cell protective fender under oxidative stress (Battistuzzi et al., 2010; Smulevich, Jakopitsch, Droghetti, & Obinger, 2006; Welinder, 1991). Class II, of the peroxidase-catalase superfamily, are extracellular fungal peroxidases including lignin peroxidase (LiP), manganese peroxidase (MnP), and versatile peroxidase (VP) which are involved in lignin degradation while class III includes peroxidases secreted by plants such as horseradish peroxidase (HRP) which have been implicated in cell wall biosynthesis, Indole-3-acetic acid catabolism and oxidation of poisonous compounds (Battistuzzi et al., 2010; Veitch & Smith, 2001). The pertinent features of class 11 peroxidases motivate for the extensive inroad into the activities of heme peroxidases as have been synopsized in this review. 3 Class II Heme-Peroxidases Class II heme-peroxidases are reported as fungal or bacterial in nature. They are extracellular enzymes associated with lignin degradation and, perhaps, portend vital roles in the valorization of lignocellulosic biomass to commercializable products. Ligninolytic heme-peroxidases including MnP, LiP, and VP play central role in delignification (Ruiz-Dueñas & Martinez, 2009). These peroxidases; MnP, LiP, and VP, oxidize specific components of the lignin structure and may act in synergy if they are produced by same organism. While MnP oxidizes the phenolic structures of lignin and LiP targets the non-phenolic components, VP has the capability of oxidizing both phenolic and non-phenolic structures. MnP (manganese-dependent peroxidase) was discovered by Kuwahara, Glenn, Morgan, and Gold (1984) and has been described as the most common lignin-modifying peroxidase secreted by most white-rot fungi and litter decomposers (Hofrichter, 2002). Its involvement in lignin degradation has been reported and well-studied in fungi (Hofrichter, 2002), however, paucity of information exists on MnP-producing bacteria. The mechanism of action of MnP includes the catalytic oxidation of Mn2+ to Mn3+, which is highly reactive and in turn oxidizes a wide range of phenolic substrates including lignin phenolic structures (Tuor, Wariishi, Schoemaker, & Gold, 1992). Nonetheless, MnP also possesses the capability to oxidize or cleave non-phenolic structures with the contributions of mediators including thiyl or lipid radicals (Abdel-Hamid, Solbiati, & Cann, 2013; Reddy, Sridhar, & Gold, 2003). Moreso, the ability of MnP to oxidize and depolymerize natural and synthetic lignin and as well, recalcitrant compounds has been reported (Bogan, Lamar, & Hammel, 1996; Dehorter & Blondeau, 1993; Hofrichter, 2002; Hofrichter, Steffen, & Hatakka, 2001; Hofrichter, Ullrich, Pecyna, Liers, & Lundell, 2010). MnPs possess two or three residues corresponding to Glu-35, Glu-39 and Asp-175 of Phanerochaete chrysosporium MnP 1 that binds Mn (Floudas et al., 2012; Ruiz-Dueñas et al., 2009). LiP possesses high redox potential for the oxidation of non-phenolic structures which constitute up to 90% of lignin (Martinez et al., 2005). It is also characterized with the ability to oxidize a wide range of aromatic compounds, hence, its role in the enzymatic degradation of lignin. Besides the characteristic oxidation of non-phenolic substrates, LiP has also shown the capability to oxidize a variety of phenolic compounds (Baciocchi et al., 2001). Furthermore, VP, also known as hybrid peroxidase or manganese-lignin peroxidase is characterized with a versatile ligninolytic ability as it combines the catalytic properties of both MnP (oxidation of phenolic substrates by oxidizing Mn2+ to Mn3+) and LiP (oxidation of non-phenolic aromatic compounds). The unique molecular architecture of VP, which is characterized by the presence of different oxidation-active sites, perhaps has been adduced to its versatility in activity. The ability of VP to oxidize high redox potential compounds is linked to an exposed catalytic tryptophan which forms a radical on the surface of the enzyme through a long-range electron transfer to the heme (Ruiz-Dueñas et al., 2009). This has recently been shown in a study by Saez-Jimenez et al. (2015) where direct electron transfer from lignin polymer to a surface tryptophanyl radical of VP was responsible for lignin oxidation. Hence, the significance of VP Trp-164 in delignification is pivotal. The focus on LiPs, in this review, is motivated by their high redox potential for the oxidation of non-phenolic substrates and their emerging application in the treatment of hyperpigmentation and skin lightening. 4 Lignin Peroxidase (EC 1.11.1.14) Lignin peroxidase (LiP) is also referred to as diaryl propane oxygenase and it is a heme-containing enzyme that catalyzes hydrogen peroxide-dependent oxidative degradation of lignin (Fig. 1). Ligninase I similarly serve same function as diaryl propane peroxidase. These enzymes are inclusive of the peroxidase-catalase superfamily (Zamocky & Obinger, 2010). Structurally, LiP is a monomeric hemoprotein. The nonplanarity of the heme cofactor of LiP and those in the other class-II peroxidases has been well documented (Piontek, Glumoff, & Winterhatter, 1993), and observable in the structures of the different ligninolytic peroxidases deposited in the Protein Data Bank (PDB). Figure 1Open in figure viewerPowerPoint Oxidative cleavage of β-1 linkage in lignin structure by lignin peroxidase (LiP) After the discovery of LiP in extracellular medium of white-rot fungus; P. chrysosporium (Glenn et al., 1983; Tien & Kirk, 1983), various isozymes have been identified in the following organisms: P. chrysosporium (Farrell, Murtagh, Tien, Mozuch, & Kirk, 1989), Tramates versicolor (Johansson, Welinder, & Nyman, 1993), Phlebia radiata (Moilanen, Lundell, Vares, & Hatakka, 1996), and Phanerochaete sordida (Sugiura et al., 2009). Farrell et al. (1989) demonstrated the existence of six (6) isozymes of LiP designated H1, H2, H6, H7, H8, and H10 in the extracellular fluid of cultures of P. chrysosporium BKM-F-1767. Another isozyme of lignin peroxidase, Ha was later identified by Dass and Reddy (1990). In the same vein, Glumoff et al. (1990) characterized five (5) isozymes of LiP also from P. chrysosporium. The study reported that the purified isozymes had different isoelectric point, sugar content, substrate specificity, and stability. The N-terminal sequences of their amino acids were also reported to be different which suggested that they were encoded by different genes. Gene sequencing of a lignocellulose degrading fungus; P. chrysosporium strain RP78, revealed about 10 lip genes, thus confirming the existence of isozymes of LiP (Martinez et al., 2004). Furthermore, Morgenstern, Klopman, and Hibbett (2008), in consistence with previous studies, reported P. chrysosporium genome to harbor 10 lip genes designated lip A-J and they, respectively, encode different isoforms of lignin peroxidase. At the moment, the US Department of Energy Joint Genome Institute (http://www.jgi.doe.gov) is consistently updated with lignocellulolytic fungi genome sequences. As recent as 2015, new genes coding for ligninolytic peroxidases, including different LiP isoforms, have been identified (Couturier et al., 2015; Hori et al., 2014; Ruiz-Dueñas et al., 2013). Besides the white-rot and brown-rot fungi, some bacteria have also been reported to possess ligninolytic abilities with the potential of producing ligninases. This group of bacteria has been classified as actinomycetes, α-proteobacteria, ___-proteobacteria (Bugg, Ahmad, Hardiman, & Singh, 2011; Paliwal, Rawat, Rawat, & Rai, 2012). However, bacterial lignin peroxidase has been less studied, thus espousing the need to bioprospect for bacteria species with the ability to produce novel lignin peroxidase with broad spectrum and high enzyme activity. 5 Lignin Peroxidase Structure LiP folds to form a globular shape with a size of about 50 × 40 × 40 Å (Piontek et al., 1993). It is segregated into proximal and distal domains by the heme which is completely fixed in the protein but made accessible through two small channels. The LiP folding motif contains eight major α-helices, eight minor helices, and three short antiparallel β sheets (Choinowski, Blodig, Winterhalter, & Piontek, 1999). Overall, the catalytic cycle of LiP is comparable to that of typical heme-peroxidases. However, structurally some differences between lignin peroxidase and other heme-peroxidases exist. Similarly, the molecular weight range of lignin peroxidase has been documented as 38 kDa to 43 kDa, isoelectric point range of 3.3 to 4.7 (Glumoff et al., 1990; Kirk, Croan, Tien, Murtagh, & Farrell, 1986), and a very low optimum pH of around pH 3.0 with veratryl alcohol as the substrate (Furukawa, Bello, & Horsfall, 2014; Tien & Kirk, 1988). The low optimum pH of LiP distinguishes it from other peroxidases. Crystallographic studies of cytochrome c peroxidase (CcP) and LiP revealed some structural differences; LiP possesses four disulfide bonds while CcP has none. LiP is larger in size and contains about 343 amino acid residues while CcP is made up of 294 residues (Edwards, Raag, Wariishi, Gold, & Poulos, 1993). However, CcP is thought to be abundantly endowed with oxidizable amino acids (seven tryptophans, 14 tyrosine residues, five methionines, and one cysteine) and in contrast, LiP has three tryptophans and eight methionines. Tyrosine is absent in LiP and it also does not have free cysteine. Nonetheless, a very notable difference between LiP and CcP includes the presence of phenylalanines at the contact point between the distal and proximal heme surfaces in LiP and the replacement of phenylalanines with tryptophans in the case of CcP. Similarly, Asp-183 is hydrogen bonded to heme propionate in LiP while Asn-184 plays this role in CcP. This has been suggested to partly account for the low pH optimum of lignin peroxidase as the disruption of the aspartic acid-propionate hydrogen bond would be expected to result in the destabilization of the heme pocket. The works of Choinowski et al. (1999) similarly revealed that the bond between the heme iron and the Nɛ2 atom of the proximal histidine residue in LiP is longer than that in CcP. The weaker iron-nitrogen bond in LiP makes the heme more electron deficient thereby destabilizing the high oxidation states which has been suggested as a reasonable explanation for the higher redox potential of lignin peroxidase when compared to cytochrome c peroxidase. 6 Lignin Peroxidase Catalytic Reactions LiP oxidizes different non-phenolic lignin model compounds including β-O-4 linkage-type arylglycerol-aryl ethers. LiP oxidative properties involve the formation of radical cation through one electron oxidation and this action leads to side-chain cleavage, demethylation, intramolecular addition, and rearrangements (Kirk, Tien, & Kersten, 1986; Miki, Renganathan, & Gold, 1986; Wong, 2009). Hydroxylation of benzylic methylene groups, oxidation of benzyl alcohols to their corresponding aldehydes or ketones, and phenol oxidation are other mechanistic oxidative process assoc