Open access peer-reviewed chapter

Effect of Metal Ions, Chemical Agents and Organic Compounds on Lignocellulolytic Enzymes Activities

Written By

Josiani de Cassia Pereira, Ellen Cristine Giese, Marcia Maria de Souza Moretti, Ana Carolina dos Santos Gomes, Olavo Micali Perrone, Maurício Boscolo, Roberto da Silva, Eleni Gomes and Daniela Alonso Bocchini Martins

Reviewed: 22 September 2016 Published: 29 March 2017

DOI: 10.5772/65934

From the Edited Volume

Enzyme Inhibitors and Activators

Edited by Murat Senturk

Chapter metrics overview

3,252 Chapter Downloads

View Full Metrics


Lignocellulolytic enzymes have been extensively studied due to their potential for industrial applications such as food, textile, pharmaceutical, paper, and, more recently, energy. The influence of metal ions, chemical agents, and organic compounds on these enzyme activities are addressed in this chapter, based on data available in the scientific literature.


  • cellulases
  • hemicellulases
  • ligninases
  • enzymatic activities
  • metal ions
  • chemical agents
  • organic compounds

1. Introduction

Lignocellulolytic enzymes comprise cellulases, hemicellulases, and ligninases, which respectively degrade cellulose, hemicellulose, and lignin, the main constituents of plant cell wall, which collectively are named lignocellulose. Cellulases are employed in many industrial sectors, such as textile [1], detergents [2], animal feed, and vinification [25]. In the last years, the potential of these enzymes to saccharify cellulose from lignocellulosic residues has been extensively studied aiming the use of glucose for cellulosic ethanol production [6]. Hemicellulases are used in biobleaching of Kraft pulp for paper production [7, 8], bioclarification of fruit juices [9], and obtainment of C5 and C6 sugars from lignocellulosic residues, in the context of second-generation ethanol production [10]. Finally, ligninases are used in paper, textile, cosmetic, and pharmaceutical industries, in bioremediation and wastewater treatment [11, 12], in organic synthesis, and in biological pretreatment of lignocellulosic residues [13] to be used for cellulosic ethanol production.

Many studies have elucidated how cellulases bind to their substrates, as well as their catalytic mechanisms [1417]. The modes of action of hemicellulases and ligninases have also been explored [18, 19]. The knowledge about these enzymes activators and inhibitors is also relevant, mainly in the context of industrial applications. Metal ions, for example, influence these enzymes activities and may be present in water and/or other reagents employed in industrial processes or may result from equipment corrosion [20]. However, the interference mechanisms are not well understood. There is also a lack of data to corroborate if the inhibition or activation occurs via allosteric or nonallosteric mechanism. So, this chapter presents a brief review of the main activators and inhibitors of lignocellulolytic enzymes, which are summarized in Figure 1.

Figure 1.

General distribution of activators and inhibitors of lignocellulases. HMF furfural: hydroxymethyl furfural; LPMOs: lytic polysaccharide monooxygenases; XEGIP: xyloglucan-endo-β-glucanase inhibitor proteins; XOS: xylooligosaccharides; SDS: sodium dodecyl sulfate; TAXI: T. xylanase inhibitor; XIP: xylanase inhibitor protein; TLXI: thaumatin-like xylanase inhibitor.


2. Cellulases

Cellulases are glycoside hydrolases produced mainly by microorganisms, especially filamentous fungi. Microbial cellulases include endoglucanases, exoglucanases, and β-glucosidases, which synergistically degrade cellulose.

The glycosidic bonds in cellulose molecule are not easily accessible to the active site of cellulases. So, many of these enzymes are modular, consisting of one or more noncatalytic carbohydrate binding modules (CBMs). CBMs associate the enzyme with the insoluble substrate and are connected to the catalytic module by linker peptides varying in length and structure [21, 22].

Endoglucanases (EG, endo-1,4-β-endoglucanases, E.C. hydrolyze the amorphous fraction of cellulose, releasing cellodextrins and cello-oligosaccharides [22] decreasing the substrate polymerization degree. They are classified into 11 families of glycosil-hydrolases: GH 5, 6, 7, 8, 9, 12, 44, 45, 48, 51, and 74 [23]. Some endoglucanases have affinity with others substrates, besides cellulose, such as xyloglucan, xylan, and mannan [24].

Exoglucanases or cellobiohydrolases (CBH, exo-1,4-β-exoglucanases, E.C. degrade the crystalline fraction of cellulose, releasing cellobiose, and are named Type I or II (action in nonreducing or reducing ends, respectively). Exoglucanases are clustered in two families of glycosil-hydrolases: GH 7 (CBH I) and GH 6 (CBH II) [22].

β-Glucosidases or cellobiases (beta-D-glucosideglucohydrolase, BG, E.C. hydrolyze cellobiose to glucose and also remove the nonreducing terminal β-D-glucosyl residue from glycoconjugates [25].

2.1. Metal ions associate to cellulases activities

Metal ions can be associated to proteins and can also form complexes with other molecules linked to enzymes acting as electron donors or acceptors as Lewis´s acids, or as structural regulators [26]. These ions can either activate or inhibit the enzymatic activity by interacting with amine or carboxylic acid group of the amino acids [27].

Several studies have reported the activation or inactivation of microbial cellulases by metal ions (Table 1).

Enzyme Microorganism Activator metals Inactivating metals Reference
Endoglucanase Aspergillus fumigatus Co2+ and Mg2+ K+, Mn2+, Na+, Cu2+, Fe2+, Fe3+, Pb2+, Ni2+, Cd2+, Hg2+ [28]
Endoglucanase Penicillium simplicissimum H-11 Mg2+ and Sn2+ Cu2+, Co2+, Li2+, Fe2+, Mn2+ [29]
Endoglucanase Aspergillus niger Ca2+ and Mn2+ Co2+, Fe2+, Cu2+ [30]
Endoglucanase AspergillusnigerANL301 Mn2+, Fe2+, Mg2+, Ca2+, Cu2+, Zn2+, Hg2+ [31]
Endoglucanase exoglucanase Aspergillus niger NRRL 567 Zn2+, Ca2+, Mn2+, Co2+ Mg2+, Fe2+, Hg2+ [32]
Endoglucanase Cu2+
Exoglucanase Cu2+
Endoglucanase Daldiniaeschscholzii (Ehrenb.:Fr.) Ca2+, Co2+ Hg2+, Cu2+, Fe2+ [33]
β-Glucosidase Melanocarpus sp. Na+, K+, Ca2+, Mg2+, Zn2+ Cu2+ [34]
Cellobiohydrolase Trichoderma reesei Mn2+, Ba2+, Ca2+ Hg2+ [35]
β-Glucosidase Aspergillus niger322 Pb2+, Hg2+, Mn2+, Fe2+ [36]
Endoglucanase Penicillium pinophilim MS20 Co2+, Zn2+, Mg2+ Na+, Cu2+, Hg2+, Fe2+, Pb2+, Ni2+, Mn2+, Cd2+ [37]
Endoglucanase Mucor circinelloides Ca2+, Mg2+, Co2+, Cu2+ Mn2+ [38]
β-Glucosidase Penicillium citrinum YS40-5 Na+ Zn2+, Cu2+ [39]
β-Glucosidase Fusarium oxysporum Mn2+, Fe2+, Ca2+, Mg2+, Cu2+ Hg2+ [40]
β-Glucosidase Monascus sanguineus - Ca2+, K+ [41]
Exoglucanase Aspergillus fumigatus Ca2+, Mg2+, Zn2+ - [42]
Cellobiohydrolase Penicillium purpurogenumKJS506 - Fe2+, Hg2+ [43]
Cellobiohydrolase Agaricus arvencis Ca2+, Cu2+, Mg2+ Zn2+ [44]
Endoglucanase Aspergillus terreus Cu2+, Mg2+, Ca2+, Na+ Fe2+, Mn2+, Zn2+, K+ [45]

Table 1.

Effect of metal ions on microbial cellulases activities.

Mono-, di-, and trivalent metal ions such as Na+, K+, Ca2+, Mg2+, Mn2+, Fe2+, Co2+, Cu2+, Ni2+, Zn2+, Hg2+, and Fe3+ are commonly studied in the characterization assays of cellulases [46]. Besides ionic charge, ion radius size has a great influence on the activity and stability of the enzyme. It was demonstrated that larger radius has less influence on catalytic amino acids, while the smaller radius can more intensely attract charged amino acids changing the enzyme’s overall conformation with damage on the catalytic site [47, 48].

The studies reported inhibitory effects of Fe2+ and Cu2+ on endoglucanases, exoglucanases, and β-glucosidases activities. However, the effect of other divalent ions on cellulases activities seems to be variable among the enzymes secreted by different microorganisms (e.g., Table 1). The effect of divalent ions on cellulases is not well elucidated, and possibly occurs by redox effects on the amino acids, increasing or decreasing their activities [49].

Inhibition of cellulases by Hg2+ is related to the interaction with catalytic amino acid residues containing sulfur, leading to oxidation and irregular formation of disulfide bonds [45, 46, 49]. Fe2+ can complex with D/L-lysine and L-methionine [50], Cu2+ with histidine [51], and Ba2+ with arginine, glutamine, proline, serine, and valine [52].

Sajadi [53] evaluated the interaction of amino acids, such as arginine and glutamine, with metal ions and established the following order of interaction degree: Ca2+ < Mg2+ < Mn2+ < Co2+ < Cu2+ > Zn2+.

2.2. Chemical agents and organic compounds associate to cellulases activities

Cellulases activities may also be affected by drugs (2,3-dichloride-1,4-nafthoquinone, for example), fungicides (such as phenylmercury acetate and ethylen-bis-dithiocarbamate), antibiotics and disinfectants (Phenylmercury nitrate and 8-hydroxiquinoline, among others), sugars (final product inhibition), protein (such as those secreted by plant as defense mechanism), CBM-binding organic compounds, products from sugar and lignin degradation (such as phenolic compounds) [54], food additives (such as Octyl gallate), plant hormones (auxins, such as indoleacetic  acid), and ionic solids (Sodium azide) [5558].

Cellulose degradation products such as cello-oligosaccharides and cellobiose can inhibit endo- and exoglucanase activities, respectively. Endoglucanases that act on xyloglucan and xylan can be inhibited by the xylooligomers released [59]. The addition of xylanase to the reaction media is an alternative to remove these products [60]. The inhibition of β-glycosidases activities by glucose is frequently observed [6, 61]. Disaccharides such as cellobiose and xylobiose, and monosaccharides such as mannose and galactose can inhibit some exoglucanases activities [22, 59, 62].

Gluconolactone, resulting from cellulose oxidation by lytic polysaccharide monooxygenases (LPMOs) activities, can act as β-glycosidases inhibitor. Cellobiose and also other substrates of β-glycosidases compete with gluconolactone and other LPMO-degrading products [6365]. On the other hand, β-glycosidases can be activated by soforose and lactose [66, 67].

It is relevant to consider that sugars released by enzymatic hydrolysis of lignocellulose can be degraded and converted into inhibitory compounds. Under acidic conditions, glucose, mannose, and galactose can be converted into furan aldehydes such as hydroxymethylfurfurals (HMF). HMF, in turn, can be converted into levulinic and formic acids [68].

Lignin degradation during the hydrolysis of some lignocellulosic materials such as alkali or acid pretreatment, or else during enzymatic hydrolysis (by laccases action) can release phenolic compounds [68] such as vanillin, syringaldehyde, trans-cinnamic acid, and hydroxybenzoic acid. These compounds are potential inhibitors of endo/exoglucanases and β-glycosidases activities due to the presence of hydroxyl, carbonyl, and methoxyl groups [69, 70].

As mentioned above, another class of cellulolytic inhibitors has a proteic origin. Specific xyloglucan endo-β-glucanase inhibitor proteins (XEGIPs) are presented in the cell walls of some vegetables such as tomatoes, tobacco, and wheat and inhibit endoglucanases that act on xyloglucan [7173]. These proteins are part of the plant protecting mechanism against pathogens and act by forming high-affinity complexes with the enzyme [73].

Another factor that affects the catalysis by cellulases is the enzymes interaction with lignin, the phenomenon called “nonproductive adsorption” or “nonspecific binding.” Cellulases can adsorb lignin through their CBMs [21, 7477], more specifically by its alanine residues [76]. Some cellulases show higher catalytic activity when CBMs are removed by decreasing nonproductive adsorption on lignin [74].

Nonproductive adsorption of cellulases on lignin can also be decreased by adding surfactants to the reaction media, which increases the efficiency of enzymatic catalysis [7881]. Tween 20, 40, 60, 80, and 100, Triton X-100, polyethylene glycol (PEG), among others surfactants, tend to decrease the surface tension of aqueous systems, which may alter the properties of liquids such as detergency, emulsification, greasing, and solubilization. Surfactant properties can decrease the nonproductive adsorption of cellulases on lignin, acting as “activators agents” of these enzymes [78].

Chelating agents such as EDTA (ethylene diamine tetra acetic acid), ethylene glycol (or β-mercaptoethanol), and DPPE (1,2-bis diphenylphosphino-ethylene) may activate some enzymes activities, especially cellulases, by sequestering inhibitors’ metal ions from the aqueous system [82]. When chelating agents complex with metals in the reaction media, the active site of enzyme is available to react with the substrate, which represents the positive effect of these compounds on cellulases activities. In contrast, the negative effect of chelating agents on enzymatic activity suggests that enzyme activities depend on the inorganic ion that was sequestered [20, 33, 45].


3. Hemicellulases

Since hemicellulose is very heterogeneous, its complete degradation requires the synergic action of several enzymes, mainly endoxylanases and β-xylosidases as well as a variety of accessory enzymes that act in substituted xylans and include α-D-glucuronidases, acetyl xylan esterases, ferulic acid esterases, α-galactosidases, acetyl mannan esterases, and α-L-arabinofuranosidases [83].

α-L-Arabinofuranosidases (EC; AFases) are exopolysaccharide hydrolases which remove side chains containing arabinose residues linked by α-1,2, α-1,3, and α-1,5 glycosidic bonds to the main chain of arabinananas or arabinoxylans [84]. AFases are grouped into six families of glycoside hydrolases: GH 3, 10, 43, 51, 54, and 62 [85]. A variety of AFases have been purified from fungi, bacteria, and plants [8688]. These enzymes’ activities can be affected by metal ions, ionic and nonionic detergents, and by chelating and reducing agents [85].

Xylans with acetyl and methyl glucuronic acid (MeGlcA) as substituents groups are named O-acetyl-4-O-methylglucuronoxylans. On the other hand, when α-4-0-methylglucuronic acid and α-arabinofuranose are the substituent groups, xylans are named as arabino 4-O-methylglucuronoxylan [89]. α-glucuronidases (EC hydrolyze α-1,2-glycosidic bond of MeGlcA in the side chain [90]. Among xylan-degrading enzymes, α-glucuronidases are the less studied and characterized ones. They are grouped into three families of glycosyl-hydrolases: GH 4, 67, and 115 [91].

Endoxylanases (E.C.; endo-β-1,4-xylanases) hydrolyze β-1,4 glycosidic linkages in the backbone of xylans that are composed of xylose residues [92]. According to the similarities of amino acid sequences, the majority of xylanases are grouped into glycoside hydrolases (GH) families 10 and 11 and are also classified into families GH 5, 7, 8, and 43 [93].

β-Xylosidases (E.C.; β-1,4-xylosidases) release β-D-xylopyranosyl residues from the nonreducing end of xylobiose and some small 4-β-D-xylooligosaccharides [92]. These enzymes have been classified into 10 families: GH 1, 3, 30, 39, 43, 51, 52, 54, 116, and 120, based on the predicted structural motifs of the enzyme’s catalytic domain. β-Xylosidases play a crucial role in endoxylanases activities, since their substrates, such as xylobiose, can inhibit endoxylanases action [94, 95].

3.1. Metal ions associate to hemicellulases activities

The inhibitory effect of Hg2+ on AFases activities has been reported [9699]. Besides Hg2+, Ag2+, and Pb2+ are mixed inhibitors, which do not bind to the active site, but to another region of the enzyme, and thus do not interfere with substrate binding to the catalytic site. In addition, Hg2+ is known to react with histidine and tryptophan residues, reducing the enzyme availability to metabolic function [100]. Zn2+, Cd2+, and Co2+ have also been described as potential inhibitors of AFases [88, 99, 101].

Most scientific works about α-glucuronidases purification and characterization report that these enzymes do not require metal ions for their activities [102106]. On the other hand, various metal ions exert inhibitory effects on α-glucuronidases activities, such as Ag2+, Zn2+, Cd2+, Hg2+, Mn2+, Fe2+, and Fe3+ (e.g., Table 2).

Metal ions Microorganism Referees
Ag2+ Bacillus stearothermophilus [105]
Saccharophagus degradans 2-40 [106]
Zn2+ Bacillus stearothermophilus [105]
Cd2+ Thermotoga maritime [104]
Hg2+ Thermotoga maritime [104]
Bacillus stearothermophilus [105]
Aspergillus niger [102]
Helix pomatia [103]
Saccharophagus degradans 2-40 [106]
Mn2+ Thermotoga maritime [104]
Bacillus stearothermophilus [105]
Aspergillus niger [102]
Fe2+ and Fe3+ Aspergillus niger [102]
Ni2+ Bacillus stearothermophilus [105]
Saccharophagus degradans 2-40 [106]
Cu2+ Thermotoga maritima [104]
Bacillus stearothermophilus [105]
K+ Geobacillus stearothermophilus [105]

Table 2.

Metal ions that exert inhibitory effects on α-glucuronidases activities.

Some GH 10 family enzymes require metal ions for their stability and activities. For example, Pseudomonas fluorescens sub sp. produces a xylanase that is one of the first GH 10 enzymes found to contain a calcium-binding site [93]. On the other hand, there are many GH 43 enzymes with crystal structures that showed tightly bound metal ions such as Ca2+, with structural roles [107]. Besides, many studies have reported the apparent activation of fungal β-xylosidases by Mn2+ and Ca2+, suggesting that these ions activate and protect the active site [95].

The negative effect of heavy metals, such as Hg2+, Fe2+, Co2+, Mn2+, Ag2+, Cu2+, and Pb2+ on xylanases activities have been reported [108]. Inhibition by heavy metal ions (such as Zn2+, Pb2+, and Hg2+) may occur due to the formation of a complex with the reactive groups of the enzyme. Metals from group Ilb exhibit high affinity for SH, CONH2, NH2, COOH, and PO4 [109]. Furthermore, inhibition of xylanase by Hg2+ has been reported as related to the presence of tryptophan residues, which oxidize indole ring, thereby inhibiting the enzyme activity [110]. Xylanase from Bacillus halodurans TSEV1 was strongly inhibited by Hg2+, Cu2+, and Pb2+, probably due to the catalysis of the cysteine thiol group autooxidation, which leads to the formation of intra- and intermolecular disulfide bonds or to the formation of sulfenic acid [111].

3.2. Chemical agents and organic compounds associate to hemicellulases activities

Some authors have reported that the addition of chelating agents such as EDTA and reducing agents such as β-mercaptoethanol and DTT (dithiothreitol) does not affect AFases activity [85, 99, 112]. Such agents are well known as inhibitors of thiol groups, and these data suggest that sulfhydryl groups are not related to the active site of AFases.

There are few studies reporting the action of ionic detergents in AFases activities. At low concentrations (1–2 mM), ionic detergents such as SDS can stimulate the enzyme activity, whereas in higher concentrations (20 mM) they can cause an inhibitory effect [113]. Since SDS interferes in hydrophobic regions of the enzyme, it alters its three-dimensional structure [114], indicating that these concentrations may be critical and cause enzyme denaturation.

Among the compounds that significantly activate the enzyme activity there are 2-mercaptoethanol, DTT (dithiothreitol), L-cysteine, and NAD+ indicating that these reducing agents are required for maximal activities of α-glucuronidases [115]. Some of the family 4 enzymes are known to be NAD+ dependent. The role of NAD+ for the activity of the hydrolytic GHF4 is not well known. The pyridine nucleotide cofactor could have structural and/or catalytic function and, in addition, could also be important for the regulation of enzyme activity [116].

Xylanases have received great attention in recent years, mainly due to their potential for the application in the processes of xylooligosaccharides (XOs) production, pulp bleaching, removal of antinutritional factors of animal feeds, bread making (improving the separation of wheat or other cereal gluten from starch), juice extraction from fruits or vegetables, clarification of fruit juices and wines, and extraction of more fermentable sugar from barley to produce beer [111, 117].

Xylanase proteic inhibitors might hamper their efficacy when used in industrial application. Two distinct types of xylanase inhibitors have been identified in barley, wheat, and rye: XIP (xylanase inhibitor protein), a monomeric and glycosylated protein (XIP-I most widely studied in the XIP class), that can inhibit all GH 10 and GH 11 fungal xylanases, except that from Aspergillus aculeatus. The other type of xylanase inhibitor, TAXI (Triticumaestivum xylanase inhibitor) is a mixture of two proteins, TAXI I and TAXI II, which differ according to xylanase specificities and pI. TAXI inhibitors seem to be specific for GH 11 bacterial and fungal xylanases. More recently, a third class of inhibitor called TLXI (thaumatin-like xylanase inhibitor) also purified from wheat, showed variable activities against most of GH 11 xylanases and does not inhibit GH 10 microbial xylanases [117, 118].

Many other substances, such as EDTA (a chelating reagent), β-mercaptoethanol, and DTT (both disulfide bonds reducing agents) have been extensively investigated regarding their influence on xylanases activities. Xylanase from Talaromyces thermophile is inhibited by EDTA and DTT, suggesting that disulfide bonds are essential to maintain the enzyme conformation [119]. On the other hand, the activation of xylanases in the presence of β-mercaptoethanol and DTT was reported and indicates the presence of a reduced thiol group of cysteine in these enzymes [120].

The effect of different modulators on the activity of xylanase from B. halodurans TSEV1 has been investigated. These modulators include N-bromosuccinimide (N-BS), ethyl-3-(3-dimethyl aminopropyl) carbodiimide (EDAC), iodoacetate (IAA), and Woodward’s reagent K (WRK). The inhibition of xylanase activity in the presence of NBS suggests the presence of tryptophan residues in their active site. EDAC and WRK inhibited the enzyme activity, which indicates the importance of carboxylic groups in enzyme catalysis [111].

Treatments for deconstruction of the lignocellulosic structure are frequently employed in the use of biomass as sugar’s source for ethanol production and can generate besides soluble sugars, other sources such as furan derivatives, organic acids, and phenolic compounds that can act as xylanases inhibitors, as described for cellulase [121].

Significant inhibition of xylanase activity by vanillic acid, syringic acid, acetosyringone, and syringaldehyde has been observed [121]. Boukari et al. [122] reported that endoxylanase from Thermobacillus xylanilyticus was inhibited by phenolic compounds, including cinnamic acid, p-coumaricacid, caffeic acid, ferulic acid, and 3, 4, 5-trimethoxy-cinnamic acid by the noncompetitive multisite inhibition mechanism.

Studies on the inhibitory effect of sugars on xylanases (mainly β-xylosidases) are essential for a better understanding about the decrease in the enzyme activity during biomass conversion. This kind of inhibition was subject of research for a long time, bringing up many different opinions about its mechanism. Jordan et al. [123] studied the active site of the GH 43 β-xylosidase from Selenomonas ruminantium and reported that it comprises of two subsites and a single access route for ligands. The authors classified the inhibitors into two groups: I, single binding inhibitors including cellobiose (4-O-β-D-glucopyranosyl D-glucose), D-glucose, maltose (4-O-a-D-glucopyranosyl-D-glucose), D-xylose, and L-xylose; II, double binding inhibitors including D-arabinose, L-arabinose, D-erythrose, and D-ribose. Both groups have presented competitive or noncompetitive inhibition.


4. Ligninolytic enzymes

Microorganisms that colonize on living and decaying wood are capable of producing oxidative extracellular enzymes which together play a fundamental role in lignin biodegradation. The ligninases, or lignin-degrading enzymes, can oxidize lignin and several related compounds, e.g., environmental pollutants containing polycyclic aromatic hydrocarbons, dyes, and chlorophenols [124].

Lignin-peroxidase (LiP, E.C., manganese-peroxidase (MnP, E.C., and laccase (E.C. are the major lignin-modifying enzyme systems of white-rot fungi and have also been described in actinomycetes and bacteria. These enzymes oxidize phenolic compounds and reduce molecular oxygen to water, generating intermediary radicals as illustrated in Figure 2 [125, 126].

Figure 2.

Simplified reactions of lignin peroxidase, manganese peroxidase, and laccase.

Accessory enzymes involved in the main reactions of degradation of lignin have also been described and comprise the following: cellobiose-quinoneoxireductase (E.C., aryl alcohol oxidase (E.C., glyoxal oxidase (GO, E.C., manganese-independent peroxidase (E.C., versatile peroxidase (VP, E.C., and cellobiose dehydrogenase (E.C. [127, 128].

Besides ligninolytic enzymes have been used to reduce the lignin content in several feedstock and to degrade recalcitrant aromatic compounds, due to the high chemical similarity of these compounds with lignin [13, 130, 132], the lignin-degrading enzymes have been applied in various industries such as textile dye bleaching, pulp and paper delignification, food, brewery, animal feed, laundry detergents, and xenobiotic compound degradation. Phenol oxidases such as laccases, particularly, have been applied in immunoassay, biosensors, biocatalysts, and oxygen cathode manufacturing [127, 132].

The performance of these enzymes is easily affected by environmental factors including metal ions and other chemical compounds usually found in the aforementioned industries. Ligninases with stronger tolerance to metal ions and organic solvents exhibit high potential for the application in the recalcitrant xenobiotics biodegradation and also improve the effectiveness of biotechnological and industrial enzymatic process [133, 134].

4.1. Laccases (E.C.

Laccases are multicopper blue oxidases that catalyze the one-electron oxidation of a wide range of substrates with a concomitant four-electron reduction of molecular oxygen to water [126]. The active site of laccase comprises four copper atoms in three groups: T1 (mononuclear copper), T2 (normal copper), and T3 (coupled binuclear copper). The T1 and T2 Cu2+-sites contribute as the primary electron acceptors while T3 is reduced by an intramolecular two-electron transfer from T1 and T2 Cu2+ sites [126, 135].

4.1.1. Metal ions associate to laccase activity

Although laccases are efficient on a wide range of substrates without cofactors, in most cases, the addition of Cu2+, Cd2+, Ni2+, Mo2+, and Mn2+ ions increases the activity of laccases, whereas Ag2+, Hg2+, Pb2+, Zn2+, NaN3, NaCl, and H2O2 inhibit their activity [126].

Apart from the inhibition problem, the influence of metal ions on the performance of enzyme-catalyzed reaction is also important, in addition to the study of effects of single metal ions on the enzyme activity. Lu et al. [136] observed that monovalent and trivalent metal ions inhibited the 4-nitrophenol degradation by laccase-Cu2+, as well as the addition of low concentrations of divalent ions. The suppressive effects of cations on laccase activity comprised Mg2+ > Na+ > Al3+ > K+ > Mn2+ > Hg2+ > Co2+.

4.1.2. Chemical agents and organic compounds associate to laccase activity

The Michaelis-Menten equation has been suitably used to describe the laccase kinetics and apparent binding constant (Km) and maximal reaction rate (Vmax) values. In water-miscible solvents, these kinetic parameters can be affected by the changes in water thermodynamic activity. In the case of laccase from the white-rot fungus Phlebiaradiata, e. g., pKI values show the linear dependence on solvent hydrophobicity (logP) in a system of 2,6 dimethoxyphenol as substrate in the presence of methanol, ethanol, n-propanol, acetonitrile, acetone, and DMSO [137].

Previously, the changes in Vmax by the addition of solvents have been compared to free and immobilized laccases. The activity of laccase from P. radiata was rather similar to both forms of the enzyme in the presence of 10% of ethanol, methanol, acetone, DMSO, and dioxane. The immobilized laccase was less vulnerable to Cu-chelatorthioglycolic acid, 2,6-dimethoxy-1,4-benzoquinone [128].

In the conditions of low water content, which is the case of water/organic mixtures, the values of the apparent Km tend to grow exponentially with water concentration. The apparent Vmax of immobilized laccase from Coriolusversicolor increased two orders of magnitude values with a linear increase in water content [139].

4.2. LiP (E.C.

Lignin-peroxidases are heme-containing glycoproteins that contain Fe3+ in their active site. LiP catalyzes the H2O2-dependent oxidative depolymerization of nonphenolic lignin and lignin-model compounds as well as a variety of phenolic compounds [140].

4.2.1. Metal ions, chemical agents, and organic compounds associate to LiP activity

The decrease in LiP activity is described as inhibition or denaturation according to the concentration of inhibitor compounds in an aqueous reaction system. The hydrogen bonding and anion stabilization are important characteristics to describe the effect of compounds on the active sites of enzymes, as well as water activity (aw), logP, and solvation [141].

The addition of Cu2+, Mn2+, and Fe2+ ions increases the activity of LiP, whereas Ag2+ inhibit their activity [142]. On the other hand, different solvents and organic compounds have been described as LiP potential inhibitors: alcohols, aldehydes, ketones, esters, ethers, amines, acids, amides, acetonitrile, cysteine, DMSO, EDTA, DMF, TEMED, CTAB, sodium azide, and H2O2 [141145].

Vazquez-Duhalt et al. [146] chemically modified a LiP from the white-rot fungus Phanerochaete chrysosporium by reductive alkylation with benzyl, naphthyl, and anthracyl moieties, thereby increasing its superficial hydrophobicity. These modifications altered the kinetics and increased the yield of oxidation of pyrroles, pyridines, and aromatic amines in 10% acetonitrile.

4.3. MnP (E.C.

Manganese-peroxidases catalyze the H2O2-dependent oxidation of Mn2+ into Mn3+, which is stabilized by fungal chelators such as oxalic acid or different organic acids. Then, the oxidation of various phenolic substrates (e.g., amines, dyes, lignin related compounds) occurs under the action of chelated Mn3+ ions that comprise a diffusible charge-transfer mediator in these reactions [142, 147].

4.3.1. Metal ions associate to MnP activity

MnP activity is completely inhibited by Hg2+, Pb2+, Ag+, lactate, NaN3, CaCl2, TEMED, ascorbic acid, β-mercaptoethanol, and dithreitol [148, 149]. Partial inhibition of MnP activity was observed with EDTA, a metal chelating compound that complexes with inorganic cofactors and prosthetic groups of enzymes. High concentrations of Cu2+ and Fe2+ (~4 mM) could enhance MnP activities [149]. Youngs et al. [150] related that Cd2+ is a reversible competitive inhibitor of Mn2+ to MnP activity. The inhibition was not observed in reaction systems containing 2,6-dimethoxyphenol or guaiacol in the absence of Mn2+.





This work was supported by “Programa de Internacionalização da Pesquisa da UNESP” (PROINTER, Edital 06/2016), Pró-reitoria de Pesquisa (PROPe) of the Universidade Estadual Paulista “Júlio de Mesquita Filho” (UNESP), Brazil. Josiani de Cassia Pereira and Ana Carolina dos Santos Gomes thank to “Comissão de Aperfeiçoamento de Pessoal do Nível Superior (CAPES)”, Brazil, for their PhD scholarships. Olavo Micali Perrone and Marcia Maria de Souza Moretti thank to ‘‘Fundação de Amparo à Pesquisa do Estado de São Paulo – FAPESP”, Brazil for their PhD and Post-doc Scholarships (Processes 2015/11588-4 and 2014/20521-8).


  1. 1. Simic, K., Soljacic, I., Pusic, T., 2015. Application of cellulases in the process of fiishing [Uporaba celulaz v procesu plemenitenja]. Tekstilec , 58, 47–56. doi:10.14502/Tekstilec2015.58.47
  2. 2. Podrepsek, G.H., Primozic, M., Knez, Z., Habulin, M., 2012. Immobilization of cellulase for industrial production. Chem. Eng. Transact. 27, 235–240. doi:10.3303/CET1227040
  3. 3. Karmakar, M., Ray, R.R., 2011. Current trends in research and applications of microbial cellulases. Res. J. Microbiol. 6, 41–53. doi:10.3023/jm.2011.41.53
  4. 4. Leite, R.S.R., Alves-Prado, H.F., Cabral, H., Pagnocca, F.C., Gomes, E., Da-Silva, R., 2008. Production and characteristics comparison of crude β-glucosidases produced by microorganisms Thermoascus aurantiacus e Aureobasidium pullulans in agricultural wastes. Enzyme Microb. Technol. 43, 391–395. doi:10.1016/j.enzmictec.2008.07.006
  5. 5. Graminha, E.B.N., Gonçalves, A.Z.L., Pirota, R.D.P.B., Balsalobre, M.A.A., Da Silva, R., Gomes, E., 2008. Enzyme production by solid-state fermentation: Application to animal nutrition. Anim. Feed Sci. Technol. 144, 1–22. doi:10.1016/j.anifeedsci.2007.09.029
  6. 6. de Cassia Pereira, J., Travaini, R., Paganini Marques, N., Bolado-Rodríguez, S., Bocchini Martins, D.A., 2016. Saccharification of ozonated sugarcane bagasse using enzymes from Myceliophthora thermophila JCP 1-4 for sugars release and ethanol production. Bioresour. Technol. 204, 122–129. doi:10.1016/j.biortech.2015.12.064
  7. 7. Bajpai, P., 2004. Biological bleaching of chemical pulps. Crit. Rev. Biotechnol, 24, 1-58.doi:10.1080/07388550490465817
  8. 8. Tavares, V.B., Gomes, E., Silva, R., 1997. Characterization of a cellulase-free xylanase producing Bacillus sp for biobleaching of kraft pulp. Rev. Microbiol. 28, 179–182.
  9. 9. Rakesh Sharma (2012). Enzyme Inhibition: Mechanisms and Scope, Enzyme Inhibition and Bioapplications, Prof. Rakesh Sharma (Ed.), InTech, DOI: 10.5772/39273. Available from:
  10. 10. Sues, A., Millati, R., Edebo, L., Taherzadeh, M.J., 2005. Ethanol production from hexoses, pentoses, and dilute-acid hydrolyzate by Mucor indicus. FEMS Yeast Res. 5, 669–676. doi:10.1016/j.femsyr.2004.10.013
  11. 11. Gomes, E., Aguiar, A.P., Boscolo, M., Carvalho, C.C., da Silva, R., Bonfá, M.R.B., 2007. Ligninases production by basidiomicetes strains on lignocellulosic agricultural residues and decolorization of synthetic dyes. J. Biotechnol. 131, S228. doi:10.1016/j.jbiotec.2007.07.414
  12. 12. Sahadevan, L., Misra, C., Thankamani, V., 2013. Ligninolytic enzymes for application in treatment of effluent from pulp and paper industries. Univ. J. Environ. Res. Technol. 3, 14–26.
  13. 13. Plácido, J., Capareda, S., 2015. Ligninolytic enzymes: A biotechnological alternative for bioethanol production. Bioresour. Bioprocess. 2, 23. doi:10.1186/s40643-015-0049-5
  14. 14. Fong, M., Berrin, J.-G., Paës, G., 2016. Investigation of the binding properties of a multi-modular GH45 cellulase using bioinspired model assemblies. Biotechnol. Biofuels 9, 12. doi:10.1186/s13068-016-0428-y
  15. 15. Gao, D., Chundawat, S.P.S., Sethi, A., Balan, V., Gnanakaran, S., Dale, B.E., 2013. Increased enzyme binding to substrate is not necessary for more efficient cellulose hydrolysis. Proc. Natl. Acad. Sci. 110, 10922–10927. doi:10.1073/pnas.1213426110
  16. 16. Jervis, E.J., Haynes, C.A., Kilburn, D.G., 1997. Surface diffusion of cellulases and their isolated binding domains on cellulose. J. Biol. Chem. 272, 24016–24023. doi:10.1074/jbc.272.38.24016
  17. 17. Wahlström, R., Rahikainen, J., Kruus, K., Suurnäkki, A., 2014. Cellulose hydrolysis and binding with Trichoderma reesei Cel5A and Cel7A and their core domains in ionic liquid solutions. Biotechnol. Bioeng. 111, 726–733. doi:10.1002/bit.25144
  18. 18. Nonaka, H., Kobayashi, A., Funaoka, M., 2013. Behavior of lignin-binding cellulase in the presence of fresh cellulosic substrate. Bioresour. Technol. 135, 53–57. doi:10.1016/j.biortech.2012.10.065
  19. 19. Tenkanen, M., Buchert, J., Viikari, L., 1995. Binding of hemicellulases on isolated polysaccharide substrates. Enzyme Microb. Technol. 17, 499–505. doi:10.1016/0141-0229(94)00050-2
  20. 20. Oviedo, C., Rodríguez, J., 2003. EDTA: The chelating agent under environmental scrutiny. Qum. Nova 26, 901–905.
  21. 21. Boraston, A.B., Bolam, D.N., Gilbert, H.J., Davies, G.J., 2004. Carbohydrate-binding modules: Fine-tuning polysaccharide recognition. Biochem. J. 382, 769–81. doi:10.1042/BJ20040892
  22. 22. Payne, C.M., Knott B.C., Mayes, H.B., Hansson, H., Himmel, M.E., Sandgren, M., Ståhlberg, J., Beckham, G.T., 2015. Fungal cellulases. Chem. Rev., 115, 1308-1448. doi:10.1021/cr500351c
  23. 23. Vlasenko, E., Sch??lein, M., Cherry, J., Xu, F., 2010. Substrate specificity of family 5, 6, 7, 9, 12, and 45 endoglucanases. Bioresour. Technol. 101, 2405–2411. doi:10.1016/j.biortech.2009.11.057
  24. 24. Nakazawa, H., Okada, K., Kobayashi, R., Kubota, T., Onodera, T., Ochiai, N., Omata, N., Ogasawara, W., Okada, H., Morikawa, Y., 2008. Characterization of the catalytic domains of Trichoderma reesei endoglucanase I, II, and III, expressed in Escherichia coli. Appl. Microbiol. Biotechnol. 81, 681–689. doi:10.1007/s00253-008-1667-z
  25. 25. Sørensen, A., Lübeck, M., Lübeck, P.S., Ahring, B.K., 2013. Fungal beta-glucosidases: A bottleneck in industrial use of lignocellulosic materials. Biomolecules. 3, 612–631. doi:10.3390/biom3030612
  26. 26. Riordan, J.F., 1977. The role of metals in enzyme activity. Ann. Clin. Lab. Sci. 7, 119–129.
  27. 27. Ishida, N., Okubo, A., Kawai, H., Yamazaki, S., Toda, S., 1980. Interaction of amino acids with transition metal ions in solution (I) solution structure of L-lysine with Co(II) and Cu(II) ions as studied by nuclear magnetic resonance spectroscopy. Agric. Biol. Chem. 44, 263–270. doi:10.1080/00021369.1980.10863934
  28. 28. Bagewadi, Z.K., Ninnekar, H.Z., 2015. Production, purifiation and characterization of endoglucanase from Aspergillus fumigatus and enzymatic hydrolysis of lignocellulosic waste. International Journal of Biotechnology and Biomedical Sciences, 1, 25–32.
  29. 29. Bai, H., Wang, H., Sun, J., Irfan, M., Han, M., Huang, Y., Han, X., Yang, Q., 2013. Purification and characterization of beta 1,4-glucanases from Penicillium simplicissimum H-11. BioResources 8, 3657–3671.
  30. 30. Bhavsar, N.H., Raol, B.V., Amin, S.S., Raol, G.G., 2015. Original research article production, optimization and characterization of fungal cellulase for enzymatic saccharifiation of lignocellosic agro-waste. Internation Journal of Current Microbiology and Applied Sciences, 4, 30–46.
  31. 31. Nwodo, S., Obinna, C., Uzoma, C.N., Veronica, A.O., 2011. Kinetic study and characterization of 1,4-E-endoglucanase of Aspergillus niger ANL301. Dyn. Biochem. Process Biotechnol. Mol. Biol. 5, 2–7.
  32. 32. Ghori, M.I., Ahmed, S., Malana, M.A., Jamil, A., 2012. Kinetics of exoglucanase and endoglucanase produced by Aspergillus niger NRRL 567. Afr. J. Biotechnol. 11, 7227–7231. doi:10.5897/AJB12.329
  33. 33. Karnchanatat, A., Petsom, A., Sangvanich, P., Piapukiew, J., Whalley, A.J.S., Reynolds, C.D., Gadd, G.M., Sihanonth, P., 2008. A novel thermostable endoglucanase from the wood-decaying fungus Daldinia eschscholzii (Ehrenb.:Fr.) Rehm. Enzyme Microb. Technol. 42, 404–413. doi:10.1016/j.enzmictec.2007.11.009
  34. 34. Kaur, J., Chadha, B.S., Kumar, B.A., Kaur, G.S., Saini, H.S., 2007. Purification and characterization of ??-glucosidase from Melanocarpus sp. MTCC 3922. Electr. J. Biotechnol. 10, 260–270. doi:10.2225/vol10-issue2-fulltext-4
  35. 35. Kim, D.W., Jang, Y.H., Kim, C.S., Lee, N.S., 2001. Effect of metal ions on the degradation and adsorption of two cellobiohydrolases on microcrystalline cellulose. Bull. Korean Chem. Soc. 22, 716–720.
  36. 36. Peshin, a., Mathur, J.M.S., 1999. Purification and characterization of β-glycosidase from Aspergillus niger strain 322. Lett. Appl. Microbiol. 28, 401–404. doi:10.1046/j.1365-2672.1999.00533.x
  37. 37. Pol, D., Laxman, R.S., Rao, M., 2012. Purification and biochemical characterization of endoglucanase from penicillium pinophilum MS 20. Indian J. Biochem. Biophys. 49, 189–194.
  38. 38. Saha, B.C., 2004. Production, purification and properties of endoglucanase from a newly isolated strain of Mucor circinelloides. Process Biochem. 39, 1871–1876. doi: 10.1016/j.procbio.2003.09.013
  39. 39. Ng, I.S., Li, C.W., Chan, S.P., Chir, J.L., Chen, P.T., Tong, C.G., Yu, S.M., Ho, T.H.D., 2010. High-level production of a thermoacidophilic β-glucosidase from Penicillium citrinum YS40-5 by solid-state fermentation with rice bran. Bioresour. Technol. 101, 1310–1317. doi:10.1016/j.biortech.2009.08.049
  40. 40. Olajuyigbe, F.M., Nlekerem, C.M., Ogunyewo, O.A., 2016. Production and characterization of highly thermostable β-glucosidase during the biodegradation of methyl cellulose by Fusarium oxysporum. Biochem. Res. Int., 2016, 1-8. doi:10.1155/2016/3978124
  41. 41. Dikshit, R., Tallapragada, P., 2015. Partial purification and characterization of β-glucosidase from Monascus sanguineus. Brazil. Arch. Biol. Technol. 58, 185–191.
  42. 42. Mahmood, R.T., Asad, M.J., Mehboob, N., Mushtaq, M., Gulfraz, M., Asgher, M., Minhas, N.M., Hadri, S.H., 2013. Production, purification, and characterization of exoglucanase by Aspergillus fumigatus. Appl. Biochem. Biotechnol. 170, 895–908. doi:10.1007/s12010-013-0227-x
  43. 43. Lee, K.M., Joo, A.R., Jeya, M., Lee, K.M., Moon, H.J., Lee, J.K., 2011. Production and characterization of cellobiohydrolase from a novel strain of penicillium purpurogenum KJS506. Appl. Biochem. Biotechnol. 163, 25–39. doi:10.1007/s12010-010-9013-1
  44. 44. Lee, K.M., Moon, H.J., Kalyani, D., Kim, H., Kim, I.W., Jeya, M., Lee, J.K., 2011. Characterization of cellobiohydrolase from a newly isolated strain of Agaricus arvencis. J. Microbiol. Biotechnol. 21, 711–718. doi:10.4014/jmb.1102.02001
  45. 45. Nazir, A., Soni, R., Saini, H.S., Manhas, R.K., Chadha, B.S., 2009. Purification and characterization of an endoglucanase from Aspergillus terreus highly active against barley β-glucan and xyloglucan. World J. Microbiol. Biotechnol. 25, 1189–1197. doi:10.1007/s11274-009-0001-y
  46. 46. Mandels, M., Reese, E.T., 1965. Inhibition of cellulases. Annu. Rev. Phytopathol. 3, 85–102. doi:10.1146/
  47. 47. Coolbear, T., Whittaker, J.M., Daniel, R.M., 1992. The effect of metal ions on the activity and thermostability of the extracellular proteinase from a thermophilic Bacillus, strain EA.1. Biochem. J. 287 ( Pt 2, 367–374.
  48. 48. Zeng, J., Gao, X., Dai, Z., Tang, B., Tang, X.F., 2014. Effects of metal ions on stability and activity of hyperthermophilic pyrolysin and further stabilization of this enzyme by modification of a Ca2+-binding site. Appl. Environ. Microbiol. 80, 2763–2772. doi:10.1128/AEM.00006-14
  49. 49. Tejirian, A., Xu, F., 2010. Inhibition of cellulase-catalyzed lignocellulosic hydrolysis by iron and oxidative metal ions and complexes. Appl. Environ. Microbiol. 76, 7673–7682. doi:10.1128/AEM.01376-10
  50. 50. Vassilev, K., 2013. Catalytic activity of amino acids-metal complexes in oxidation reactions. J. Biomater. Nanobiotechnol. 04, 28–36. doi:10.4236/jbnb.2013.42A004
  51. 51. Kryukova, N.P., Bolotin, S.N., Panyushkin, V.T., 2003. Increased information leakage from text. Ceur Workshop Proc. 52, 1–4. doi:10.1023/A
  52. 52. Bush, M.F., Oomens, J., Saykally, R.J., Williams, E.R., 2008. Effects of alkaline earth metal ion complexation on amino acid zwitterion stability: Results from infrared action spectroscopy. J. Am. Chem. Soc. 130, 6463–6471. doi:10.1021/ja711343q
  53. 53. Sajadi, S.A.A., 2010. Metal ion-binding properties of L-glutamic acid and L-aspartic acid, a comparative investigation. Nat.Sci. 2, 85–90.doi 10.4236/ns.2010.22013 Natural.
  54. 54. Kim, Y., Ximenes, E., Mosier, N.S., Ladisch, M.R., 2011. Soluble inhibitors/deactivators of cellulase enzymes from lignocellulosic biomass. Enzyme Microb. Technol. 48, 408–415. doi:10.1016/j.enzmictec.2011.01.007
  55. 55. Reese, E.T., Mandels, M., 1957. Chemical inhibition of cellulases and B-glucosidases. Res. Rept. Pioneering Res. Div., QMRE Center, Natick, Mass., Microbial. Ser. 17, 60.
  56. 56. Jermyn, M. A., 1952. Fungal cellulases. I. General properties of unpurified enzyme preparations from Aspergillus oryzae. Aust. J. of Sci. Res. Ser. B., 5, 409–432.
  57. 57. Sison, B., Schubert, W., Nord, F.F., 1958. On the mechanism of enzyme action. LXV. A cellulolytic enzyme from the mold Poria vaillantii. Arch. Biochem. Biophys., 75, 260–272.
  58. 58. Li, P., Zhu, J., Kong, Q., Jiang, B., Wan, X., Yue, J., Li, M., Jiang, H., Li, J., Gao, Z., 2013. The ethylene bis-dithiocarbamate fungicide mancozeb activates voltage-gated KCNQ2 potassium channel. Toxicol. Lett. 219, 211–217. doi:10.1016/j.toxlet.2013.03.020
  59. 59. Qing, Q., Yang, B., Wyman, C.E., 2010. Xylooligomers are strong inhibitors of cellulose hydrolysis by enzymes. Bioresour. Technol. 101, 9624–9630. doi:10.1016/j.biortech.2010.06.137
  60. 60. Qing, Q., Wyman, C.E., 2011. Supplementation with xylanase and β-xylosidase to reduce xylo-oligomer and xylan inhibition of enzymatic hydrolysis of cellulose and pretreated corn stover. Biotechnol. Biofuels 4, 18. doi:10.1186/1754-6834-4-18
  61. 61. Pereira, J.C., Alonso, D., Martins, B., Gomes, E., Silva, R., 2015. Production and characterization of β-glucosidase obtained by the solid-state cultivation of the thermophilic fungus thermomucor indicae-seudaticae, N31. Appl. Biochem. Biotechnol., 175, 723-732. Doi: 10.1007/s12010-014-1332-1
  62. 62. Hsieh, C.W.C., Cannella, D., Jørgensen, H., Felby, C., Thygesen, L.G., 2014. Cellulase inhibition by high concentrations of monosaccharides. J. Agric. Food Chem. 62, 3800–3805. doi:10.1021/jf5012962
  63. 63. Kou, Y., Xu, J., Cao, Y., Lv, X., Zhao, G., Chen, G., Zhang, W., Liu, W., 2014. Gluconolactone induces cellulase gene expression in cellulolytic filamentous fungus Trichoderma reesei. RSC Adv. 4, 36057. doi:10.1039/C4RA06731B
  64. 64. Mishra, S.K., Sangwan, N.S., Sangwan, R.S., 2013. Purification and physicokinetic characterization of a gluconolactone inhibition-insensitive β-glucosidase from Andrographis paniculata NEES. Leaf. Prep. Biochem. Biotechnol. 43, 481–499. doi:10.1080/10826068.2012.759966
  65. 65. Tiwari, P., Misra, B.N., Sangwan, N.S., 2013. β-Glucosidases from the fungus Trichoderma: An effient cellulase machinery in biotechnological applications-glucosidases from the fungus trichoderma: An effient cellulase machinery in biotechnological applicationsglucosidases from the fungus Trichode. Biomed Res. Int., 2013, 1-10. doi:10.1155/2013/203735
  66. 66. Dhillon, G.S., Kaur, S., Brar, S.K., Verma, M., 2012. Potential of apple pomace as a solid substrate for fungal cellulase and hemicellulase bioproduction through solid-state fermentation. Ind. Crops Prod. 38, 6–13. doi:10.1016/j.indcrop.2011.12.036
  67. 67. Nisizawa, T., Suzuki, H., Nakayama, M., Nisizawa, K., 1971. Inductive formation of cellulase by sophorose in Trichoderma viride. J. Biochem. 70, 375–385.
  68. 68. Jönsson, L.J., Alriksson, B., Nilvebrant, N.-O., 2013. Bioconversion of lignocellulose: Inhibitors and detoxification. Biotechnol. Biofuels 6, 16. doi:10.1186/1754-6834-6-16
  69. 69. Qin, L., Li, W.-C., Liu, L., Zhu, J.-Q., Li, X., Li, B.-Z., Yuan, Y.-J., 2016. Inhibition of lignin-derived phenolic compounds to cellulase. Biotechnol. Biofuels. 9, 70. doi:10.1186/s13068-016-0485-2
  70. 70. Ximenes, E., Kim, Y., Mosier, N., Dien, B., Ladisch, M., 2011. Deactivation of cellulases by phenols. Enzyme Microb. Technol. 48, 54–60. doi:10.1016/j.enzmictec.2010.09.006
  71. 71. Juge, N., 2006. Plant protein inhibitors of cell wall degrading enzymes. Trends Plant Sci. 11, 359–367. doi:10.1016/j.tplants.2006.05.006
  72. 72. Misas-Villamil, J.C., van der Hoorn, R. AL, 2008. Enzyme-inhibitor interactions at the plant-pathogen interface. Curr. Opin. Plant Biol. 11, 380–388. doi:10.1016/j.pbi.2008.04.007
  73. 73. York, W.S., Qin, Q., Rose, J.K.C., 2004. Proteinaceous inhibitors of endo-β-glucanases. Biochim. Biophys. Acta: Proteins Proteom. 1696, 223–233. doi:10.1016/j.bbapap.2003.07.003
  74. 74. Le Costaouëc, T., Pakarinen, A., Várnai, A., Puranen, T., Viikari, L., 2013. The role of carbohydrate binding module (CBM) at high substrate consistency: Comparison of Trichoderma reesei and Thermoascus aurantiacus Cel7A (CBHI) and Cel5A (EGII). Bioresour. Technol. 143, 196–203. doi:10.1016/j.biortech.2013.05.079
  75. 75. Rahikainen, J.L., Evans, J.D., Mikander, S., Kalliola, A., Puranen, T., Tamminen, T., Marjamaa, K., Kruus, K., 2013. Cellulase-lignin interactions: The role of carbohydrate-binding module and pH in non-productive binding. Enzyme Microb. Technol. 53, 315–321. doi:10.1016/j.enzmictec.2013.07.003
  76. 76. Strobel, K.L., Pfeiffer, K.A., Blanch, H.W., Clark, D.S., 2015. Structural insights into the affinity of Cel7A carbohydratebinding module for lignin. J. Biol. Chem. 290, 22818–22826. doi:10.1074/jbc.M115.673467
  77. 77. Li, Y., Sun, Z., Ge, X., Zhang, J., 2016. Effects of lignin and surfactant on adsorption and hydrolysis of cellulases on cellulose. Biotechnol. Biofuels 9, 20. doi:10.1186/s13068-016-0434-0
  78. 78. Hsieh, C.-W.C., Cannella, D., Jørgensen, H., Felby, C., Thygesen, L.G., 2015. Cellobiohydrolase and endoglucanase respond differently to surfactants during the hydrolysis of cellulose. Biotechnol. Biofuels 8, 52. doi:10.1186/s13068-015-0242-y
  79. 79. Pardo, A.G., 1996. Effect of surfactants on cellulase production by Nectria catalinensis. Curr. Microbiol. 33, 275–278. doi:10.1007/s002849900113
  80. 80. Reese, E.T., Maguire, A., 1969. Surfactants as stimulants of enzyme production by microorganisms. Appl. Microbiol. 17, 242–245.
  81. 81. Trinh, D.K., Quyen, D.T., Do, T.T., Nghiem, N.M., 2013. Purification and characterization of a novel detergent- and organic solvent-resistant endo-beta-1,4-glucanase from a newly isolated basidiomycete Peniophora sp. NDVN01. Turk. J. Biol. 37, 377–384. doi:10.3906/biy-1207-37
  82. 82. Miyano, H., Toyo’oka, T., Imai, K., Nakajima, T., 1985. Influences of metal ions on the reaction of amino and imino acids with fluorogenic reagents. Anal. Biochem. 150, 125–130. doi:10.1016/0003-2697(85)90450-6
  83. 83. Shallom, D., Shoham, Y., 2003. Microbial hemicellulases. Curr. Opin. Microbiol. 6, 219–228. doi:10.1016/S1369-5274(03)00056-0
  84. 84. Koseki, T., Okuda, M., Sudoh, S., Kizaki, Y., Iwano, K., Aramaki, I., Matsuzawa, H., 2003. Role of two α-L-arabinofuranosidases in arabinoxylan degradation and characteristics of the encoding genes from shochu koji molds, Aspergillus kawachii and Aspergillus awamori. J. Biosci. Bioeng. 96, 232–241. doi:10.1016/S1389-1723(03)80187-1
  85. 85. Numan, M.T., Bhosle, N.B., 2006. α-L-Arabinofuranosidases: The potential applications in biotechnology. J. Ind. Microbiol. Biotechnol. 33, 247–260. doi:10.1007/s10295-005-00721
  86. 86. Bezalel, L., Shoham, Y., Rosenberg, E., Aviv, R., 1993. Applied microbiology biotechnology characterization and delignifiation activity of a thermostable α-L-arabinofuranosidase from Bacillus stearothermophilus. Appl. Microbiol. Biotechnol., 57–62.
  87. 87. Hong, M.R., Park, C.S., Oh, D.K., 2009. Characterization of a thermostable endo-1,5-α-L-arabinanase from Caldicellulorsiruptor saccharolyticus. Biotechnol. Lett. 31, 1439–1443. doi:10.1007/s10529-009-0019-0
  88. 88. Sakamoto, T., Kawasaki, H., 2003. Purification and properties of two type-B α-L-arabinofuranosidases produced by Penicillium chrysogenum. Biochim. Biophys. Acta: Gen. Subj. 1621, 204–210. doi:10.1016/S0304-4165(03)00058-8
  89. 89. Sunna, a, Antranikian, G., 1997. Xylanolytic enzymes from fungi and bacteria. Crit. Rev. Biotechnol. 17, 39–67. doi:10.3109/07388559709146606
  90. 90. Roy, N., Timell, T.E., 1964. The acid hydrolysis of glycosides. Can. J. Chem. 42, 1456–1472. doi:10.1016/S0008-6215(00)82576-8
  91. 91. Henrissat, B., Vegetales, M., Grenoble, F., 1991. A classification of glycosyl hydrolases based sequence similarities amino acid. Biochem. J. 280, 309–316. doi:10.1007/s007920050009
  92. 92. Daniela Alonso Bocchini Martins, Heloiza Ferreira Alves do Prado, Rodrigo Simões Ribeiro Leite, Henrique Ferreira, Márcia Maria de Souza Moretti, Roberto da Silva and Eleni Gomes (2011). Agroindustrial Wastes as Substrates for Microbial Enzymes Production and Source of Sugar for Bioethanol Production, Integrated Waste Management - Volume II, Mr. Sunil Kumar (Ed.), InTech, DOI: 10.5772/23377. Available from:
  93. 93. Bhradwaj, A., Mahanta, P., Ramakumar, S., Ghosh, A., Leelavathi, S., Reddy, V.S., 2012. Emerging role of N-and C-terminal interactions in stabilizing (β/α)8 fold with special emphasis on family 10 xylanases. Comput. Struct. Biotechnol. J. 2, 1–10. doi:10.5936/csbj.201209014
  94. 94. Lee, C.C., Braker, J.D., Grigorescu, A.A., Wagschal, K., Jordan, D.B., 2013. Divalent metal activation of a GH43 β-xylosidase. Enzyme Microb. Technol. 52, 84–90. doi:10.1016/j.enzmictec.2012.10.010
  95. 95. Yang, X., Shi, P., Huang, H., Luo, H., Wang, Y., Zhang, W., Yao, B., 2014. Two xylose-tolerant GH43 bifunctional β-xylosidase/α- arabinosidases and one GH11 xylanase from Humicola insolens and their synergy in the degradation of xylan. Food Chem. 148, 381–387. doi:10.1016/j.foodchem.2013.10.062
  96. 96. Gilead, S., Shoham, Y., 1995. Purification and characterization of alpha-L-arabinofuranosidase from Bacillus stearothermophilus T-6. Appl. Environ. Microbiol. 61, 170–174.
  97. 97. Raweesri, P., Riangrungrojana, P., Pinphanichakarn, P., 2008. α-L-Arabinofuranosidase from Streptomyces sp. PC22: Purification, characterization and its synergistic action with xylanolytic enzymes in the degradation of xylan and agricultural residues. Bioresour. Technol. 99, 8981–8986. doi:10.1016/j.biortech.2008.05.016
  98. 98. Khandeparker, R., Numan, M.T.H., Mukherjee, B., Satwekar, A., Bhosle, N.B., 2008. Purification and characterization of α-L-arabinofuranosidase from Arthrobacter sp. MTCC 5214 in solid-state fermentation. Process Biochem. 43, 707–712. doi:10.1016/j.procbio.2008.02.014
  99. 99. Margolles, A., de los Reyes-Gavilán, C.G., 2003. Purification and functional characterization of a novel alpha-L-arabinofuranosidase from Bifidobacterium longum B667. Appl. Environ. Microbiol. 69, 5096–103. doi:10.1128/AEM.69.9.5096
  100. 100. Volkin, D.B., Klibanov A.M.,. 1989. Minimizing protein inactivation, in: T. E. Creighton (ed.), Protein Function, A Practical Approach. IRL Press, Oxford, England. pp. 1–24.
  101. 101. Tsujibo, H., Takada, C., Wakamatsu, Y., Kosaka, M., Tsuji, A., Miyamoto, K., Inamori, Y., 2002. Cloning and expression of an alpha-L-arabinofuranosidase gene (stxIV) from Streptomyces thermoviolaceus OPC-520, and characterization of the enzyme. Biosci. Biotechnol. Biochem. 66, 434–438. doi:10.1271/bbb.66.434
  102. 102. Uchida, H., Nanri, T., Kawabata, Y., Kusakabe, I., Murakami, K., 1992. Purification and characterization of intracellular α-glucuronidase from Aspergillus niger. Biosci. Biotechnol. Biochem. 56, 1608–1615.
  103. 103. Kawabata, Y., Ono, K., Gama, Y., Yoshida, S., Kobayashi, H., Kusakabe, I., 1995. Purification and characterization of alpha-glucuronidase from snail acetone powder. Biosci. Biotechnol. Biochem. 59, 1086–1090. doi:10.1271/bbb.59.1086
  104. 104. Ruile, P., Winterhalter, C., Liebl, W., 1997. Isolation and analysis of a gene encoding alpha-glucuronidase, an enzyme with a novel primary structure involved in the breakdown of xylan. Mol. Microbiol. 23, 267–279.
  105. 105. Zaide, G., Shallom, D., Shulami, S., Zolotnitsky, G., Golan, G., Baasov, T., Shoham, G., Shoham, Y., 2001. Biochemical characterization and identification of catalytic residues in alpha-glucuronidase from Bacillus stearothermophilus T-6. Eur. J. Biochem. 268, 3006–3016. doi:ejb2193 [pii]
  106. 106. Wang, W., Yan, R., Nocek, B.P., Voung, T.V., Leo, R. Di, Xu, X., Cui, H., Gatenholm, P., Toriz, G., Tenkanen, M., Savchenko, A., Master, E.R., 2016. Biochemical and structural characterization of a fie-domain GH115 alpha-glucuronidase from the marine bacterium Saccharophagus degradans 2-40T. J. Biol. Chem., 291, 14120-14133. doi:10.1074/jbc.M115.702944
  107. 107. Jordan, D.B., Lee, C.C., Wagschal, K., Braker, J.D., 2013. Activation of a GH43 β-xylosidase by divalent metal cations: Slow binding of divalent metal and high substrate specificity. Arch. Biochem. Biophys. 533, 79–87.
  108. 108. Juturu, V., Wu, J.C., 2012. Microbial xylanases: Engineering, production and industrial applications. Biotechnol. Adv. 30, 1219–1227. doi:10.1016/j.biotechadv.2011.11.006
  109. 109. Heinen, P.R., Henn, C., Peralta, R.M., Bracht, A., De Cássia, R., Simão, G., Luís, J., Silva, C., De Lourdes, M., Polizeli, T.M., Kadowaki, M.K., 2014. African journal of biotechnology xylanase from Fusarium heterosporum: Properties and influence of thiol compounds on xylanase activity. 13, 1047–1055. doi:10.5897/AJB2013.13282
  110. 110. Verma, D., Kawarabayasi, Y., Miyazaki, K., Satyanarayana, T., 2013. Cloning, expression and characteristics of a novel alkalistable and thermostable xylanase encoding gene (Mxyl) retrieved from compost-soil metagenome. PLoS One 8, 1-8. doi:10.1371/journal.pone.0052459
  111. 111. Kumar, V., Satyanarayana, T., 2013. Biochemical and thermodynamic characteristics of thermo-alkali-stable xylanase from a novel polyextremophilic Bacillus halodurans TSEV1. Extremophiles 17, 797–808. doi:10.1007/s00792-013-0565-1
  112. 112. Tuncer, M., 2000. Characterization of β-xylosidase and α-L-arabinofuranosidase activities from Thermomonospora fusca BD25. Turk. J. Biol. 24, 753–767.
  113. 113. Debeche, T., Cummings, N., Connerton, I., Debeire, P., O’Donohue, M.J., 2000. Genetic and biochemical characterization of a highly thermostable alpha-L-arabinofuranosidase from Thermobacillus xylanilyticus. Appl. Environ. Microbiol. 66, 1734–1736. doi:10.1128/AEM.66.4.1734-1736.2000
  114. 114. Nelson, D.L., Cox, M.M. Princípios de bioquímica de Lehninger. 5. ed. Artmed, Porto Alegre, 2011.
  115. 115. Suresh, C., Kitaoka, M., Hayashi, K., 2003. A thermostable non-xylanolytic α-glucuronidase of Thermotoga maritima MSB8. Biosci. Biotechnol. Biochem. 67, 2359–2364. doi:10.1271/bbb.67.2359
  116. 116. Raasch, C., Armbrecht, M., Streit, W., Höcker, B., Sträter, N., Liebl, W., 2002. Identification of residues important for NAD+ binding by the Thermotoga maritima α-glucosidase AglA, a member of glycoside hydrolase family 4. FEBS Lett. 517, 267–271. doi:10.1016/S0014-5793(02)02641-8
  117. 117. Brutus, A., Villard, C., Durand, A., Tahir, T., Furniss, C., Puigserver, A., Juge, N., Giardina, T., 2004. The inhibition specificity of recombinant Penicillium funiculosum xylanase B towards wheat proteinaceous inhibitors. Biochim. Biophys. Acta: Proteins Proteom. 1701, 121–128. doi:10.1016/j.bbapap.2004.06.010
  118. 118. Paës, G., Berrin, J.G., Beaugrand, J., 2012. GH11 xylanases: Structure/function/properties relationships and applications. Biotechnol. Adv. 30, 564–592. doi:10.1016/j.biotechadv.2011.10.003
  119. 119. Maalej, I., Belhaj, I., Masmoudi, N.F., Belghith, H., 2009. Highly thermostable xylanase of the thermophilic fungus talaromyces thermophilus: Purification and characterization. Appl. Biochem. Biotechnol. 158, 200–212. doi:10.1007/s12010-008-8317-x
  120. 120. Fialho, M.B., Carmona, E.C., 2004. Purification and characterization of xylanases from Aspergillus giganteus. Folia Microbiol. (Praha). 49, 13–18.
  121. 121. Duarte, C.L., Ribeiro, M.a., Oikawa, H., Mori, M.N., 2013. Study of thermal treatment combined with radiation on the decomposition of polysaccharides in sugarcane bagasse. Radiat. Phys. Chem. 84, 191–195. doi:10.1016/j.radphyschem.2012.06.019
  122. 122. Boukari, I., O’Donohue, M., Remond, C., Chabbert, B., 2011. Probing a family GH11 endo-β-1,4-xylanase inhibition mechanism by phenolic compounds: Role of functional phenolic groups. J. Mol. Catal. B: Enzym. 72, 130–138. doi:10.1016/j.molcatb.2011.05.010
  123. 123. Jordan, D.B., Braker, J.D., 2007. Inhibition of the two-subsite β-D-xylosidase from Selenomonas ruminantium by sugars: Competitive, noncompetitive, double binding, and slow binding modes. Arch. Biochem. Biophys. 465, 231–246. doi:10.1016/
  124. 124. Eriksson, K-E.L., Bermek, H.L., Lignocellulose, Ligninase. Schaechter, M. Encyclopedia of Microbiology. 3. ed. Elsevier, San Diego, Volume 1, p. 373, 2009.
  125. 125. Mester, T., Ming, T., 2000. Oxidation mechanism of ligninolytic enzymes involved in the degradation of environmental pollutants. Int. Biodeterior. Biodegrad. 46, 51–59.
  126. 126. Manavalan, T., Manavalan, A., Heese, K., 2015. Characterization of lignocellulolytic enzymes from white-rot. Fungi Curr. Microbiol. 70, 485–498.
  127. 127. Maciel, M.J.M., Castro e Silva, A., Ribeiro, H.C.T., 2010. Industrial and biotechnological applications of ligninolytic enzymes of the basidiomycota: A review. Electr. J. Biotechnol. 13, 1–13. doi: 10.2225/vol13-issue6-fulltext-2
  128. 128. Pollegioni, L., Tonin, F., Rosini, E., 2015. Lignin-degrading enzymes. FEBS J. 282, 1190–1213. doi: 10.1111/febs.13224
  129. 129. Ruiz-Dueñas, F.J., Martínez, A.T., 2009. Microbial degradation of lignin: How a bulky recalcitrant polymer is efficiently recycled in nature and how we can take advantage of this. Microbiol. Biotechnol. 2, 164–177. doi: 10.1111/j.1751-7915.2008.00078.x
  130. 130. Coelho-Moreira, J.S., Maciel, G.M., Castoldi, R., Mariano, S.S., Inácio, F.D., Bracht, A., Peralta, R.M. Involvement of lignin-modifying enzymes in the degradation of herbicides, in: Price, A.J., Kelton, J.A. Herbicides—Advances in Research. Intech, 2013.
  131. 131. Abdel-Hamid, A.M., Solbiati, J.O., Cann, I.K., 2013. Insights into lignin degradation and its potential industrial applications. Adv. Appl. Microbiol. 82, 1–28. doi:10.1016/B978-0-12-407679-2.00001-6
  132. 132. Karigar, C.S., Rao, S.S., 2011. Role of microbial enzymes in the bioremediation of pollutants: A review. Enzyme Res. 2011, 1–11. doi:10.4061/2011/805187
  133. 133. Viswanath, B., Rajesh, B., Janardhan, A., Kumar, A.P., Narasimha, G., 2014. Fungal laccases and their applications in bioremediation. Enzyme Res. 2014, 1–21. doi:10.1155/2014/163242
  134. 134. Yaropolov, A.I., Skorobogat'ko, O.V., Vartanov, S.S., Varfolomeyev, S.D., 1994. Laccase: Properties, catalytic mechanism, and applicability. Appl. Biochem. Biotechnol. 49, 257–280. doi:10.1007/BF02783061
  135. 135. Lu, C., Cao, L., Liu, R., Lei, Y., Ding, G., 2012. Effect of common metal ions on the rate of degradation of 4-nitrophenol by a laccase-Cu2+ synergistic system. J. Environ. Manage. 113, 1–6. doi: 10.1016/j.jenvman.2012.08.023
  136. 136. Rodakiewicz-Nowak, J., Kasture, S.M, Dudek, B., Haber, J., 2000. Effect of various water-miscible solvents on enzymatic activity of fungal laccases. J. Mol. Catal. B: Enzym. 11, 1–11.
  137. 137. Rogalski, J., Jozwik, E., Hatakka, A., Leonowicz, A., 1995. Immobilization of laccase from Phlebiaradiata on controlled porosity glass. J. Mol. Catal. A: Chem. 95, 99–108.
  138. 138. vanErp, S.H., Kamenskaya, E.O., Khmelnitsky, Y.L., 1991. The effect of water content and nature of organic solvent on enzyme activity in low-water media. A quantitative description. Eur. J. Biochem. 2, 379–384.
  139. 139. Dasthban, M., Schraft, H., Syed, T.A., Qin, W., 2010. Fungal biodegradation and enzymatic modification of lignin. Int. J. Biochem. Mol. Biol. 1, 36–50.
  140. 140. Gorjup, B., Lampic, N., Pencaa, R., Perdiha, A., Perdih, M., 1999. Solvent effects on ligninases. Enzyme Microb. Technol. 25, 15–22.
  141. 141. Asgher, M., Iqbal, H.M.N., Irshad, M., 2012. Characterization of purified and xerogel immobilized novel lignin peroxidase produced from trametes versicolor IBL-04 using solid state medium of corncobs. BMC Biotechnol. 12, 46. doi:10.1186/1472-6750-12-46
  142. 142. Tuisel, H., Grover, T.A., Lancaster Jr., J.R., Bumpus, J.A., Aust, S.D., 1991. Inhibition of lignin peroxidase H2 by sodium azide. Arch. Biochem. Biophys. 288, 456–462.
  143. 143. Chang, H.C., Bumpus, J.A., 2001. Inhibition of lignin peroxidase mediated oxidation activity by ethylenediamine tetra acetic acid and N-N-N’-N’-tetramethylenediamine. Proc. Natl. Sci. Counc. Repub. China B 25, 26–33.
  144. 144. Liu, A., Huang, X., Song, S., Wang, D., Lu, X., Qu, Y., Gao, P., 2003. Kinetics of the H2O2-dependent ligninase-catalyzed oxidation of veratryl alcohol in the presence of cationic surfactant studied by spectrophotometric technique.Spectrochim. Acta A: Mol. Biomol. Spectrosc. 59, 2547–2551.
  145. 145. Vazquez-Duhalt, R., Westlake, D.W.S., Fedorak, P.M., 1994. Lignin peroxidase oxidation of aromatic compounds in systems containing organic solvents. Appl. Environ. Microbiol. 60, 459–466
  146. 146. Wesenberg, D., Kyriakides, I., Agatho, S.N., 2003. White-rot fungi and their enzymes for the treatment of industrial dye effluents. Biotechnol. Adv. 22, 161–187.
  147. 147. Kanayama, N., Tohru, S., Keiichi, K., 2002. Purification and characterization of an alkaline manganese peroxidase from Aspergillus terreus LD-1. J. Biosci. Bioeng. 93, 405–410.
  148. 148. Asgher, M., Iqbal, H.M.N., 2011. Characterization of a novel manganese peroxidase purified from solid state culture of Trametes versicolor IBL-04. BioResources 6, 4302–4315.
  149. 149. Youngs, H.L., Sundaramoorthy, M., Gold, M.H., 2000.Effects of cadmium on manganese peroxidase competitive inhibition of MnII oxidation and thermal stabilization of the enzyme. Eur. J. Biochem. 267, 1761–1769.

Written By

Josiani de Cassia Pereira, Ellen Cristine Giese, Marcia Maria de Souza Moretti, Ana Carolina dos Santos Gomes, Olavo Micali Perrone, Maurício Boscolo, Roberto da Silva, Eleni Gomes and Daniela Alonso Bocchini Martins

Reviewed: 22 September 2016 Published: 29 March 2017