**3. Major enzymes used in bioremediation of synthetic dyes**

Over the last two decades, there has been increasing interest in the use of formulated enzymes rather than live bacteria as bioremediation agents owing to their specificity, and the ease with which their robustness can be enhanced with engineering [36]. Other advantages of using enzymes include increased enzyme production, enhanced stability and/or activity, and lower costs by using recombinant DNA technology. However, the high structural complexity of dye molecules means that only a few enzymes can degrade these compounds. These dye-degrading enzymes share one common mechanistic feature—they are all redox-active molecules that exhibit relatively wide substrate specificities. The most important dye-degrading enzymes are azoreductases, laccases, and peroxidases.

In this chapter, the accumulated research data on the potential of the oxidoreductive enzymes —high redox potential peroxidases, polyphenol oxidases, and azoreductases—that have been exploited in the decolorization and degradation of dyes are presented. The current initiatives and future prospects for the potential use of genetic engineering techniques to develop novel enzyme variants that are more durable and versatile biocatalyst, with respect to both the varieties of xenobiotics degraded and the operative conditions of dye bioremediation tech‐ nologies are also discussed.

#### **3.1. Peroxidases**

such as biological processes capable of reducing and even eliminating synthetic dye com‐

Biological decolorization and degradation are an environmentally friendly, cost-competitive, and efficient alternative to physical/chemical decomposition [3, 6, 7, 10]. Decolorization by biological means may take place in either one of three ways: (1) adsorption (or biosorption) on the microbial biomass, (2) biodegradation by cells, and (3) biodegradation by enzymes. Biosorption involves the entrapment of dyes in the matrix of the adsorbent (microbial biomass) without destruction of the pollutant. In contrast, biodegradation involves the fragmentation of the original dye structure into smaller compounds, resulting in the decolorization of synthetic dyes. Several studies have described the use of microorganisms as biosorption agents in the removal of dye pollutants from wastewater [11, 12]. However, relative to the operational simplicity and adaptability of microorganisms to a given set of conditions, the biodegradation mechanism is considered efficacious and hence preferable to biosorption for treatment of dye

There are numerous reports of microorganisms capable of decolorizing synthetic dyes: bacteria [14-16], fungi [17-19], yeasts [20, 21], actinomycetes [22, 23], and algae [24, 25]. Several fungi are capable of mineralizing pollutant compounds by action of their highly oxidative and nonspecific ligninolytic enzymes, which are also responsible for the decolorization and degradation of many different dyes. There are reports that white rot fungi (WRF), members of the basidiomycetes such as *Funalia trogii* [26], *Phanerochaete chrysosporium* [17, 18], *Trametes versicolor* [19], *Trametes hirsuta* [27], *Irpex lacteus* [28], and *Lentinula edodes* [29], can efficiently degrade xenobiotic textile dye compounds. Lignin-degrading fungi also degrades a wide range of aromatics owing to the relatively nonspecific activity of the extracellular ligninolytic enzymes, such as lignin peroxidase (LiP), manganese peroxidase (MnP), versatile peroxidase

Bacterial strains that can aerobically decolorize azo dyes have also been isolated during the past few years. However, there are only very few bacteria that are able to grow on azo compounds as the sole carbon source. The degradation of various azo dyes by mixed aerobic and facultative anaerobic microbial consortia under anoxic conditions has also been reported [30]. *Pseudomonas luteola*, *Aeromonas hydrophila*, *Bacillus subtilis*, *Pseudomonas* sp., and *Proteus mirabilis* can also decolorized azo dyes under anoxic conditions [30-32]. These bacteria are specific toward their substrate, cleaving R–N=N–R bonds reductively and using the resultant

The efficiency of dye degradation is mainly dependent on the following: (1) type of microor‐ ganisms used; (2) process conditions such as temperature, oxygen level, pH, available carbon, and nitrogen sources that affects microbial growth; and (3) dye concentration and chemical structure [10, 33]. Dependence on microbial growth to metabolize the toxicants makes the bioremediation of dyes by these organisms a relatively slow process. Another drawback to the

pounds from the environment.

114 Advances in Bioremediation of Wastewater and Polluted Soil

wastewater [13].

(VP), and laccase.

**2. Microbial bioremediation of synthetic dyes**

amines as a source of carbon and energy for their growth.

Peroxidases (oxidoreductases, EC 1.11.1.x) are a diverse group of versatile heme-containing enzymes that utilize hydrogen peroxide or organic hydroperoxides (R-OOH) electron acceptor to catalyze oxidation of numerous substrates. Due to their catalytic versatility and enzymatic stability, peroxidases are one of the most studied groups of enzymes, more so as potential industrial/environmental biocatalysts [37]. Several good reviews have summarized years of peroxidase research and described their potential applications [37-40].

Generally, peroxidases are ubiquitous in nature; found in fungi, plants, animals, and eubac‐ teria; and are classified within different superfamilies on the basis of their sequence homolo‐ gies; animal and nonanimal peroxidases (former plant peroxidases) form the largest groups. The plant superfamily is further grouped into three subclasses according to cellular localiza‐ tion: class I—intracellular, organelle-associated, and bacterial peroxidases (e.g., cytochrome *c* peroxidase [CCP]); class II—secretory fungal peroxidases, including ligninolytic peroxidases such as LiPs, MnPs, and VPs [41–43]; and class III—secreted plant peroxidases (horseradish peroxidase [HRP]) [37]. Among peroxidases, a new superfamily—"dye-decolorizing peroxi‐ dases"—has arisen. These enzymes are known to successfully oxidize a wide range of substrates, but most importantly, they effectively degrade high redox synthetic dyes such as anthraquinone and azo dyes. DyPs form a novel superfamily of peroxidases that are generally characterized by "atypical" molecular architecture and divergent mechanistic behavior that is not fully understood yet but different from the classical peroxidases [37, 40, 41].

Structurally, peroxidases share several features, including the overall protein fold and the general architecture of the heme pocket with the high-spin ferric iron (FeIII) coordinated to the proximal histidine and the conserved distal histidine and arginine residues [38, 39]. These highly conserved residues in the heme catalytic pocket are critical for peroxidase activity, i.e., in the generation and stabilization of compound I and II. The functional and catalytic diversity of heme peroxidases is thereby attributable to specific structural differences around the hemebinding site, including the nature of the axial ligand, and the environment of the substratebinding site.

Catalytically, peroxidases share a general reaction mechanism by using hydrogen peroxide (H2O2) as the final electron acceptor in the oxidation of a broad range of substrates (AH) to radicals (AH⋅+), which involves a three-step sequential mechanism via compound I and compound II intermediates [37, 38, 40, 42]. Initially, the native enzyme, which is in the ferric form FeIII, undergoes two-electron oxidation by H2O2 to produce compound I (oxyferryl porphyrinyl radical [FeIV=O⋅+]). One electron is removed from the ferric iron FeIII to form the ferryl FeIV, whereas the second electron is withdrawn from the porphyrin ring to form a porphyrin cation radical [43]. During this reaction step, H2O2 is reduced to water. Next, compound I oxidizes substrates by one electron and is reduced to compound II. In this step, the porphyrin ring gains an electron. Thus, compound I is able to oxidize substrates with a higher redox potential than compound II [38, 44]. The reduction of compound II, by a second electron of substrate, brings the enzyme back to the native state and completes the catalytic cycle.

In the absence of substrate and in the presence of excess H2O2, compound II is converted to compound III, a ferrous-oxy or ferric superoxide species. In this process, H2O2 reduces compound II by one electron to produce a ferric enzyme and a superoxide radical. The latter readily combines with the ferric peroxidase to produce compound III. In the peroxidase catalytic cycle, the generation of highly reactive radicals that undergo a complex series of spontaneous cleavage reactions accounts for the degradation ability of peroxidase toward numerous substrates, e.g., phenolic and nonphenolic aromatics, metal ions, and complex dyestuff molecules [45].

#### *3.1.1. Ligninolytic peroxidases*

Lignin-modifying peroxidases (LMPs; LIP, MnP, and VP) refer to a group of glycosylated, heme-containing ligninolytic enzymes produced by the fungi during secondary metabolism in nutrient starved cultures [46]. These enzymes are produced in multiple isoforms and are affected by many external factors, such as nutrient level, mediator compounds, and metal ions. Phylogenetically, they belong to class II extracellular fungal peroxidases in the so-called "plant peroxidase superfamily."

Lignin peroxidases (diarylpropane: oxygen, hydrogen peroxide oxidoreductase, EC 1.11.1.14) were first described in the basidiomycete *P. chrysosporium* Burdsall in 1983 [47] and have become the most studied peroxidase since then. They have also been reported in several species of white rot basidiomycetes [48-51], actinomycetes [52, 53], and some bacteria, such as *Brevibacillus laterosporus* MTCC 2298 [54] and *Streptomyces viridosporus* T7A [55]. LiPs are generally dependent on H2O2 and have very high redox potential and low optimum pH [56, 57]. Both these characteristics are important for their ability to oxidize a variety of reducing substrates, including polymeric substrates such as complex dye compounds. LiPs have a typical enzymatic cycle, characteristic of other peroxidases. LiP, MnP, and VP share an almost identical heme environment, which is responsible, among other factors for their high redox potential. However, they differ in the substrates that they can oxidize because of the presence of different catalytic sites in their molecular structures. LiP oxidizes nonphenolic lignin model compounds in direct contact with a tryptophan radical exposed to the solvent [58].

characterized by "atypical" molecular architecture and divergent mechanistic behavior that is

Structurally, peroxidases share several features, including the overall protein fold and the general architecture of the heme pocket with the high-spin ferric iron (FeIII) coordinated to the proximal histidine and the conserved distal histidine and arginine residues [38, 39]. These highly conserved residues in the heme catalytic pocket are critical for peroxidase activity, i.e., in the generation and stabilization of compound I and II. The functional and catalytic diversity of heme peroxidases is thereby attributable to specific structural differences around the hemebinding site, including the nature of the axial ligand, and the environment of the substrate-

Catalytically, peroxidases share a general reaction mechanism by using hydrogen peroxide (H2O2) as the final electron acceptor in the oxidation of a broad range of substrates (AH) to radicals (AH⋅+), which involves a three-step sequential mechanism via compound I and compound II intermediates [37, 38, 40, 42]. Initially, the native enzyme, which is in the ferric form FeIII, undergoes two-electron oxidation by H2O2 to produce compound I (oxyferryl porphyrinyl radical [FeIV=O⋅+]). One electron is removed from the ferric iron FeIII to form the ferryl FeIV, whereas the second electron is withdrawn from the porphyrin ring to form a porphyrin cation radical [43]. During this reaction step, H2O2 is reduced to water. Next, compound I oxidizes substrates by one electron and is reduced to compound II. In this step, the porphyrin ring gains an electron. Thus, compound I is able to oxidize substrates with a higher redox potential than compound II [38, 44]. The reduction of compound II, by a second electron of substrate, brings the enzyme back to the native state and completes the catalytic

In the absence of substrate and in the presence of excess H2O2, compound II is converted to compound III, a ferrous-oxy or ferric superoxide species. In this process, H2O2 reduces compound II by one electron to produce a ferric enzyme and a superoxide radical. The latter readily combines with the ferric peroxidase to produce compound III. In the peroxidase catalytic cycle, the generation of highly reactive radicals that undergo a complex series of spontaneous cleavage reactions accounts for the degradation ability of peroxidase toward numerous substrates, e.g., phenolic and nonphenolic aromatics, metal ions, and complex

Lignin-modifying peroxidases (LMPs; LIP, MnP, and VP) refer to a group of glycosylated, heme-containing ligninolytic enzymes produced by the fungi during secondary metabolism in nutrient starved cultures [46]. These enzymes are produced in multiple isoforms and are affected by many external factors, such as nutrient level, mediator compounds, and metal ions. Phylogenetically, they belong to class II extracellular fungal peroxidases in the so-called "plant

Lignin peroxidases (diarylpropane: oxygen, hydrogen peroxide oxidoreductase, EC 1.11.1.14) were first described in the basidiomycete *P. chrysosporium* Burdsall in 1983 [47] and have

not fully understood yet but different from the classical peroxidases [37, 40, 41].

116 Advances in Bioremediation of Wastewater and Polluted Soil

binding site.

cycle.

dyestuff molecules [45].

*3.1.1. Ligninolytic peroxidases*

peroxidase superfamily."

Biochemically, LiPs utilize veratryl alcohol (VA) to complete the catalytic cycle by reducing compound II to a resting enzyme to avoid inactivation. VA (3,4-dimethoxybenzyl alcohol) is a secondary metabolite, concomitantly produced by basidiomycetous fungi, along with LiPs usually synthesized from glucose in liquid cultures. Moreover, VA acts as a cation radical redox mediator of remote substrates [58, 59]. Similar to other peroxidases, LiP shows little substrate specificity, and, due to its high redox potential, reacts with a wide variety of lignin model compounds and even unrelated molecules. Interestingly, LiP has the distinction of being able to oxidize methoxylated aromatic rings without a free phenolic group, generating cation radicals that can react further by a variety of pathways, including ring opening, demethylation, and phenol dimerization [60], a typical mechanism also employed in the degradation of dye compounds. Numerous studies have reported the application of LiPs in the decolorization of various synthetic dyes and industrial dye wastewaters [20, 43, 48, 54, 61-63].

MnP (MnII:hydrogen peroxide oxidoreductase, EC 1.11.1.13), like LiPs, is a heme-containing peroxidase and is the most common ligninolytic peroxidase produced by almost all white rot basidiomycetes [37]. MnP is a glycoprotein with a heme (ferric protoporphyrin) group that shares the mechanistic properties of other peroxidases and the formation of oxidized inter‐ mediates, compound I and compound II, in the presence of H2O2 for aromatic and nonphenolic substrates oxidation [44]. Catalytically, MnP requires MnII as an electron donor; MnII is oxidized to MnIII [45], a deviation from other typical peroxidase reaction mechanisms. This specific oxidation of MnII by MnP generally occurs in a small channel formed by three acidic residues located directly on the heme internal propionate [64]. MnP depends on MnII as a substrate for compound I and II formation. In this reaction scheme, MnP oxidizes MnII to MnIII in the presence of H2O2, and the MnIII formed oxidizes a variety of compounds [44]. The chelation of MnII to MnIII by organic acids, such as oxalate, is necessary for MnP activity [53]. Its unique catalytic activity enables MnPs to biodegrade dyes, as well as decolorize various types of synthetic dyes, indicating their potential application in the environmental bioreme‐ diation of dye industry wastewater [65-67].

VPs (EC 1.11.1.16) are also known as hybrid peroxidases or lignin–manganese peroxidases because of their dual LiP and MnP catalytic properties. They are relatively new fungal peroxidases that were first thought to be MnPs but have since been isolated and thoroughly characterized in *Pleurotus* and *Bjerkandera* [68-72]. In contrast to other ligninolytic peroxidases, VPs possess two catalytic sites, one for the direct oxidation of low- and high-redox potential compounds and the other for oxidation of MnII in a preferred manner [70-72]. *Pleurotus eryngii* VP (PeVP) possesses three acidic amino acid residues for MnII binding and a catalytic efficiency (*k*cat/*K*m) for MnII oxidation that is typical of MnPs. In addition, PeVP has a tryptophan residue, Trp164, which is analogous to the PeLiP Trp171 that participates in electron transfer from aromatic donors and consequently enables the enzyme to oxidize nonphenolic ligninrelated structures [64]. The dual catalytic mode of action observed accounts for their ability to catalyze the direct degradation/oxidation of a broad spectrum of persistent substrates (e.g., nonphenolic lignin compounds, dyes, such as RB5 and others) in the absence of mediators [71, 72], an important feature as a potential catalyst for a variety of biotechnological applications.

#### *3.1.2. Dye decolorizing peroxidases*

Dye decolorizing peroxidases (DyPs) comprise a novel group of heme-containing enzymes, named for their ability to efficiently oxidize high redox potential trichromatic anthraquinoic (AQ) dyes. They were first reported in the extracellular secretions of a plant pathogenic fungus, *Bjerkandera adusta* Dec 1 (wrongly annotated previously as *Thanetophorus cucumeris*) [73]. The main features of DyP from *B. adusta* Dec1 include the following: (1) a monomeric 60-kDa glycosylated enzyme having higher specificity for AQ than for azo dyes, and different degradation spectra for phenolic compounds such as 2,6-dimethoxy-phenol, guaiacol, and VA; (2) a low pH optima (pH < 3.0); (3) lack of a conserved active site for distal histidine; and (4) structural divergence from classical plant and animal peroxidases (Figure 2) [73, 74]. To date, these enzymes, the physiological function of which is still unclear, have been identified from the genomes of fungi, bacteria, and archaea (http://peroxibase.toulouse.inra.fr/index.php). Interestingly, there is increasing evidence for the key role that microbial DyP peroxidases play in the degradation of lignin (see [37, 40, 41] up-to-date reviews on DyP-type peroxidases and their known biological, chemical, and structural features).

Similar to other peroxidases, DyPs are catalytically bifunctional enzymes displaying both oxidative and hydrolytic activity. They exhibit significant catalytic versatility arising from their ability to oxidize a variety of organic compounds, some of which, including dyes, phenols, βcarotene, lignin model compounds, and aromatic sulfides, are poorly degraded by conven‐ tional peroxidases [75-78]. Moreover, studies have demonstrated that DyPs are very robust enzymes in terms of pH [76], temperature, and pressure [78]. Several mediators can also improve DyPs substrate range. For example, we observed a drastic enhancement of azo dye oxidation in presence of a natural mediator syringaldehydeby AnaPX, a bacterial DyP [75], whereas MnII activates *Rhodococcus jostii* DyP2 activity [79]. The potential utility, as industrial/ environmental biocatalysts in the bioremediation of wastewater contaminated with synthetic dyes specifically—the recalcitrant and xenobiotic AQ dyes that are generally not substrates of peroxidases such as HRP—has made DyPs the focus of significant interest.

Biochemically, the physicochemical properties of DyPs such as UV–vis spectral characteristics, molecular masses, or isoelectric points resemble those of classical heme proteins [76, 80, 81]. DyPs are structurally divergent from typical peroxidase; the basic architecture has a dominant α+β-helical secondary structure with extended loop regions (Figure 2). The N-terminal and C- terminal domains contain an antiparallel β-sheet that is arranged into a characteristic ferre‐ doxin-like motif on the distal side of the heme moiety [41, 82, 83]. Obviously, structural peculiarities, including the nature of the axial ligands, the environment of the substratebinding site, and the involvement of intramolecular electron transfer, appear to account for the novel and the varied catalytic differences between DyPs and other peroxidases. Although DyPs possess a heme iron prosthetic group with a conserved proximal Fe–His–Asp triad found in most other peroxidases, the generally conserved distal His is absent in DyPs. Instead, an Asp residue forming the absolutely conserved novel GXXDG motif and an Arg in the distal position of DyPs are present (Figure 2). The distal Asp residue assumes the part of the catalytic base, mediating peroxide cleavage and thus the formation of compound I [84]. Moreover, the distal Asp's "swinging movement" determines heme cavity access for small organic molecules, making it function as the heme cavity's gatekeeper [75, 85]. The observed smaller heme-access channel for DyPs indicates that the oxidation of bulky substrates such as bulkier AQ dyes requires another substrate interactions sites. The existence of substrate interaction sites, involving long-range electron transport (LRET) via either surface-exposed tyrosyl or trypto‐ phan residues, has been demonstrated in several DyPs [77, 82]. A MnII binding site, which accounts for the oxidase activity of DyP2, in the absence of peroxide, has also been reported in *R. jostii* DyP2 [79].

#### **3.2. Laccases**

VPs possess two catalytic sites, one for the direct oxidation of low- and high-redox potential compounds and the other for oxidation of MnII in a preferred manner [70-72]. *Pleurotus eryngii* VP (PeVP) possesses three acidic amino acid residues for MnII binding and a catalytic efficiency (*k*cat/*K*m) for MnII oxidation that is typical of MnPs. In addition, PeVP has a tryptophan residue, Trp164, which is analogous to the PeLiP Trp171 that participates in electron transfer from aromatic donors and consequently enables the enzyme to oxidize nonphenolic ligninrelated structures [64]. The dual catalytic mode of action observed accounts for their ability to catalyze the direct degradation/oxidation of a broad spectrum of persistent substrates (e.g., nonphenolic lignin compounds, dyes, such as RB5 and others) in the absence of mediators [71, 72], an important feature as a potential catalyst for a variety of biotechnological applications.

Dye decolorizing peroxidases (DyPs) comprise a novel group of heme-containing enzymes, named for their ability to efficiently oxidize high redox potential trichromatic anthraquinoic (AQ) dyes. They were first reported in the extracellular secretions of a plant pathogenic fungus, *Bjerkandera adusta* Dec 1 (wrongly annotated previously as *Thanetophorus cucumeris*) [73]. The main features of DyP from *B. adusta* Dec1 include the following: (1) a monomeric 60-kDa glycosylated enzyme having higher specificity for AQ than for azo dyes, and different degradation spectra for phenolic compounds such as 2,6-dimethoxy-phenol, guaiacol, and VA; (2) a low pH optima (pH < 3.0); (3) lack of a conserved active site for distal histidine; and (4) structural divergence from classical plant and animal peroxidases (Figure 2) [73, 74]. To date, these enzymes, the physiological function of which is still unclear, have been identified from the genomes of fungi, bacteria, and archaea (http://peroxibase.toulouse.inra.fr/index.php). Interestingly, there is increasing evidence for the key role that microbial DyP peroxidases play in the degradation of lignin (see [37, 40, 41] up-to-date reviews on DyP-type peroxidases and

Similar to other peroxidases, DyPs are catalytically bifunctional enzymes displaying both oxidative and hydrolytic activity. They exhibit significant catalytic versatility arising from their ability to oxidize a variety of organic compounds, some of which, including dyes, phenols, βcarotene, lignin model compounds, and aromatic sulfides, are poorly degraded by conven‐ tional peroxidases [75-78]. Moreover, studies have demonstrated that DyPs are very robust enzymes in terms of pH [76], temperature, and pressure [78]. Several mediators can also improve DyPs substrate range. For example, we observed a drastic enhancement of azo dye oxidation in presence of a natural mediator syringaldehydeby AnaPX, a bacterial DyP [75], whereas MnII activates *Rhodococcus jostii* DyP2 activity [79]. The potential utility, as industrial/ environmental biocatalysts in the bioremediation of wastewater contaminated with synthetic dyes specifically—the recalcitrant and xenobiotic AQ dyes that are generally not substrates of

Biochemically, the physicochemical properties of DyPs such as UV–vis spectral characteristics, molecular masses, or isoelectric points resemble those of classical heme proteins [76, 80, 81]. DyPs are structurally divergent from typical peroxidase; the basic architecture has a dominant α+β-helical secondary structure with extended loop regions (Figure 2). The N-terminal and C-

peroxidases such as HRP—has made DyPs the focus of significant interest.

*3.1.2. Dye decolorizing peroxidases*

118 Advances in Bioremediation of Wastewater and Polluted Soil

their known biological, chemical, and structural features).

Laccases belong to the multicopper oxidase family of enzymes that catalyze the oxidation of various substrates with the simultaneous reduction of molecular oxygen to water, through a radical-catalyzed reaction mechanism [86]. They are mainly of fungal or plant origin, although a few representatives have been identified and isolated in bacteria and insects [87, 88]. The most studied laccases are fungal in origin, mainly in phyla Ascomycota, Zygomycota, and Basidiomycota (see reviews [87, 88]). The most biotechnologically useful laccases are also of fungal origin. Physiologically, the functions of laccases are diverse, ranging from lignolysis, pigment formation, detoxification, to pathogenesis. All these functions are attributed to the enzymes' ability to oxidize a wide range of aromatic substrates such as polyphenols and diamines and even some inorganic compounds [87, 88].

Compared with fungal laccases, bacterial laccases are generally more stable at high pH and temperatures [89]. Although fungal laccases can be both intra- and extracellular, bacterial laccases are predominantly intracellular. Lacasse have been isolated from the rhizospheric bacterium *Azospirillum lipoferum* [90], the melanogenic marine bacterium *Marinomonas mediterranea*, and the endospore coat component (CotA) of *B. subtilis* [89, 91]. In addition, the optimum pH of bacterial laccases is higher than that of fungal laccases [89, 91], whose optimum pH is acidic [89, 91]. The optimum temperature for most laccases is between 50°C and 70°C, with thermal stability dependent on the microbial source.

The structure of an active holoenzyme laccase molecule is a dimeric or tetrameric glycoprotein containing four copper atoms per monomer, bound to three redox sites (T1, T2, and T3 Cu pair) [87, 88]. The four Cu atoms differ from each other in their specific properties, such as the characteristic electronic paramagnetic resonance (EPR) signals that allow them to play an

**Figure 2.** The structural model of DyP showing the heme active site architecture. (a) Typical α+β protein structure of DyP and the heme active site showing the conservation of key catalytic proximal Fe–His–Asp triad residues and sub‐ stitution of His with Asp in the distal heme active. (b) Possible coordination of distal Asp and other active site residues with a water molecule involving hydrogen-bond network in the heme active site as implicated by the resolved crystal structures of ferric DyP enzymes [82-84].

important role in the catalytic mechanism of the enzyme. For catalytic activity, a minimum of four Cu atoms per active protein unit is needed. The T2 and T3 Cu atoms form a trinuclear cluster site, which is responsible for the binding and reduction of oxygen to water. T2 Cu is coordinated by two His and T3 Cu pair by six His. The strong antiferromagnetic coupling between the two T3 Cu atoms is maintained by a hydroxyl bridge [87, 88]. The function of the T1 site, in this type of enzyme, involves electron abstraction from reducing substrates (electron donors), with a subsequent electron transfer to the T2/T3 Cu cluster. Generally, laccase catalyzes the four-electron reduction of oxygen to water (at the T2–T3 trinuclear Cu centers) by the sequential one-electron uptake from a suitable reducing substrate (at the T1 mononuclear Cu center) [86]. Lacasses are categorized into high potential lacasses (HPLs) and low potential lacasses (LPLs) on the basis of T1–T3 Cu having a redox potential of 0.6–0.8 V and 0.4–0.6 V, respectively,

The catalytic efficiency of laccases (*k*cat/*K*m) depends on the redox potential of T1 site; conse‐ quently, laccases with high redox potential at the T1 site are of special interest in biotechnology for their potential application in bleaching and bioremediation processes [92]. Compared with LiP, MnP, and VP that exhibit higher redox potentials (1.15–1.25 V), most laccases typically have low redox potential (0.5–0.8 V). Consequently, most laccases lack the ability to degrade nonphenolic aromatic substrates due to their low redox potential. However, in the presence of mediators, particularly small chemical compounds with redox potential higher than 0.9 V, the substrate range of laccases can be expanded to include the oxidation of nonphenolic compounds such as lignin and complex dye compounds [27, 86, 87]. A mediator is a low molecular weight (LMW) chemical compound that is continuously oxidized by the laccase enzyme and subsequently reduced by the substrate. Due to its bulky size, high molecular weight (HMW) substrates cannot enter the laccase active site, and the mediator acts as a carrier of electrons between the enzyme and the substrate, thereby overcoming the steric hindrances that exist between them [86].

Laccases and lacasse-mediator systems (LMSs) have been intensively studied with regard to their degradation of various recalcitrant compounds, such as chlorophenols, polyaromatic hydrocarbons (PAHs), lignin-related structures, organophosphorous compounds, phenols, and synthetic dyes [27, 87]. These enzymes have great potential in various biotechnological processes mainly because of their high nonspecific oxidation capacity, the lack of a requirement for cofactors, and the use of readily available oxygen as an electron acceptor. Laccases and LMSs have found various biotechnological and environmental applications, including as analytical tools/biosensors for phenols, and in the development of oxygen cathodes in biofuel cells, textile dye degradation, organic synthesis, immunoassay labeling, delignification, demethylation, and in bleaching of craft pulp [87, 88, 92].

#### **3.3. Azoreductases**

important role in the catalytic mechanism of the enzyme. For catalytic activity, a minimum of four Cu atoms per active protein unit is needed. The T2 and T3 Cu atoms form a trinuclear cluster site, which is responsible for the binding and reduction of oxygen to water. T2 Cu is coordinated by two His and T3 Cu pair by six His. The strong antiferromagnetic coupling between the two T3 Cu atoms is maintained by a hydroxyl bridge [87, 88]. The function of the T1 site, in this type of enzyme, involves electron abstraction from reducing substrates (electron

structures of ferric DyP enzymes [82-84].

120 Advances in Bioremediation of Wastewater and Polluted Soil

**Figure 2.** The structural model of DyP showing the heme active site architecture. (a) Typical α+β protein structure of DyP and the heme active site showing the conservation of key catalytic proximal Fe–His–Asp triad residues and sub‐ stitution of His with Asp in the distal heme active. (b) Possible coordination of distal Asp and other active site residues with a water molecule involving hydrogen-bond network in the heme active site as implicated by the resolved crystal

Azoreductases are a group oxidoreductive enzymes that catalyze the NAD(P)H-dependent reduction of azo compounds to the corresponding amines, via cleavage of the azo linkages (R-N=N-R), resulting in azo dye degradation [93]. They are a varied family of enzymes that have been identified in almost all species except in viruses. Physiologically, these enzymes partici‐ pate in enzymatic detoxifications systems, involving the reduction of quinones, quinone imines, azo dyes, and nitro groups, and protect cells against the toxic effects of free radicals and reactive oxygen species arising from electron reductions [93].

Azoreductase activity has been characterized from a wide variety of bacteria, including *Pigmentiphaga kullae* K24, *Xenophilus azovorans* KF46F, *Enterococcus faecalis*, *Staphylococcus aureus*, *Escherichia coli*, *Bacillus* sp. strain OY1-2, *Pseudomonas aeruginosa*, and *Rhodobacter* *sphaeroides* [93-101]. There are at least two different types of bacterial azoreductases: those that require flavin and those that do not [94]. Flavin-dependent azoreductases can be further classified into two families according to their amino acid sequences. Azoreductases from *E. coli* and *Bacillus* sp. strain OY1-2 are representative of the two flavin-dependent azoreductases, respectively. *Bacillus* sp. strain OY1-2 azoreductase is a 23-kDa protein with the ability to reduce the azo dyes Rocceline, Sumifix Red B, and Methyl red, producing dimethyl pphenylenediamine and o-aminobenzoic acid in the presence of β-NADPH [102]. On the other hand, *Xenophilus azovarans* KF46 and *P. kullae* K24 azoreductases are monomeric flavin-free enzymes that use NADPH as a cofactor to degrade the azo dyes carboxy-Orange II and I. These two enzymes exhibit different substrate specificities and sizes (21 and 30 kDa) and require the presence of hydroxyl groups in the aromatic ring of the substrate [95].

#### **3.4. Other dye-degrading enzymes**

In addition to peroxidases, laccases, and azoreductases, the bioremediation of synthetic dye compounds with other enzymes such as tyrosinases [103], aryl alcohol oxidases [104], and biosulfidogenic hydrogenases [105] have been demonstrated. Similar to laccases, tyrosinases (monophenol monooxygenase, EC1.14.18.1) are oxidoreductases that can catalyze the oxida‐ tion of phenolic and other aromatic compounds, without the use of a cofactor in presence of oxygen. Catalytically, these enzymes possess both cresolase activity (ortho-hydroxylation of monophenols to *o*-diphenols) and catecholase activity (the oxidation of *o*-diphenols to *o*quinones) [103]. Tyrosinases also degrade aromatic amines and *o*-aminophenols via the similar ortho-hydroxylation and oxidation reactions. The resultant products undergo subsequent polymerization, giving rise to oligomeric products.

Biotechnologically, tyrosinases are used as markers of the oxidative enzymes involved in the degradation of azo dyes. For example, the involvement of tyrosinase in the degradation of Direct Blue-6 by *Pseudomonas desmolyticum* NCIM 2112 [106], disperse dye brown 3REL by a microbial consortium consisting of *Galactomyces geotrichum* MTCC 1360, and sulfonated azo dyes by *Brevibacterium* sp. strain VN-15 [103] and *Bacillus* sp. VUS [62] has been demonstrated. The activity of aryl alcohol oxidase (AAO) has been reported in *B. adusta* Dec 1; it oxidizes VA to veratraldehyde producing H2O2, which is important for supporting the *in vivo* dye-decol‐ orizing activity of fungi [104]. The involvement of a similar enzyme in *Comamonas* sp. UVS during the decolorization of Red HE7B and Direct Blue GL has also been reported [107].
