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Cultivation of Mushrooms and Their Lignocellulolytic Enzyme Production Through the Utilization of Agro-Industrial Waste

Research Center of Microbial Diversity and Sustainable Utilization, Chiang Mai University, Chiang Mai 50200, Thailand
Department of Biology, Faculty of Science, Chiang Mai University, Chiang Mai 50200, Thailand
Division of Biology, Faculty of Science and Technology, Rajamangala University of Technology Thanyaburi, Thanyaburi, Pathumthani 12110, Thailand
School of Preclinic, Institute of Science, Suranaree University of Technology, Nakhon Ratchasima 30000, Thailand
Center of Excellence in Microbial Technology for Agricultural Industry, Suranaree University of Technology, Nakhon Ratchasima 30000, Thailand
School of Science, Mae Fah Luang University, Chiang Rai 57100, Thailand
Academy of Science, The Royal Society of Thailand, Bangkok 10300, Thailand
Author to whom correspondence should be addressed.
Molecules 2020, 25(12), 2811;
Received: 29 May 2020 / Revised: 13 June 2020 / Accepted: 15 June 2020 / Published: 18 June 2020
(This article belongs to the Special Issue Mushrooms:The Versatile Roles)


A large amount of agro-industrial waste is produced worldwide in various agricultural sectors and by different food industries. The disposal and burning of this waste have created major global environmental problems. Agro-industrial waste mainly consists of cellulose, hemicellulose and lignin, all of which are collectively defined as lignocellulosic materials. This waste can serve as a suitable substrate in the solid-state fermentation process involving mushrooms. Mushrooms degrade lignocellulosic substrates through lignocellulosic enzyme production and utilize the degraded products to produce their fruiting bodies. Therefore, mushroom cultivation can be considered a prominent biotechnological process for the reduction and valorization of agro-industrial waste. Such waste is generated as a result of the eco-friendly conversion of low-value by-products into new resources that can be used to produce value-added products. Here, we have produced a brief review of the current findings through an overview of recently published literature. This overview has focused on the use of agro-industrial waste as a growth substrate for mushroom cultivation and lignocellulolytic enzyme production.

1. Introduction

The rapidly growing global population and expansion in the agriculture sector and food industries have resulted in the generation of a large amount of agro-industrial waste annually. Agro-industrial waste is defined as the waste that is generated during the industrial processing of agricultural or animal products or the waste obtained from agricultural activities [1,2]. The waste can further be divided into two types, agricultural residues and industrial residues, respectively [2,3,4,5]. Agricultural residues consist of field residues and process residues. Field residues are generated during the crop harvesting process and are made up of leaves, roots, stalks, straw, seed pods and stems. Process residues are generated during the further processing of the crops and are made up of husks, peels, pulp and shells. Asia is the largest producer of agricultural residues at 47%, followed by the United States (29%), Europe (16%), Africa (6%) and Oceania (2%) [6]. Industrial residues are residues that are produced by the food, fruit and vegetable processing industries and include bran, peels, pomace and bagasse. Generally, most agro-industrial waste is disposed of in landfills or burned, leading to various environmental problems and pose potential harm to the health of humans and wildlife [5,7,8]. However, agro-industrial waste can potentially be converted into different high-value products, including biofuels, value-added fine chemicals and cheap energy sources for microbial fermentation and enzyme production [7,8,9]. These waste products can represent a source of energy, as well as sources of carbon. Additionally, this form of waste is a source of the nutrients that are required for mushroom growth and lignocellulolytic enzyme production via solid state fermentation [9,10,11]. Therefore, in this study, we have summarized the current findings on the use of agro-industrial waste as growth substrates for mushroom cultivation and lignocellulolytic enzyme production.

2. The Composition of Agro-Industrial Wastes

Agro-industrial waste is a major lignocellulosic component. This form of waste includes cellulose, hemicelluloses and lignin, which are normally referred to as “lignocellulosic materials”. Generally, cellulose is the most abundant component, followed by hemicellulose and lignin (Figure 1).
Cellulose is a homopolymer consisting of a linear chain of several hundred to many thousands of β-anhydroglucose units (β-1,4 linked d-glucose units). Each of the β-anhydroglucose units consists of three hydroxyl groups (OH), one primary (C6 position) and two secondary (C2 and C3 positions) hydroxyl groups, each of which exhibits different polarities and is capable of being involved in the intra- and intermolecular hydrogen bonds [12,13]. The intra- and inter-chain hydrogen bonding network makes cellulose a relatively stable polymer and gives the cellulose fibrils high axial stiffness [14].
Hemicellulose is a heteropolymer consisting of a polysaccharide backbone. Its structure greatly varies depending on the sugar units, chain length and the branching of the chain molecules. Typical binding sugars in hemicelluloses are pentoses (xylose and arabinose), hexoses (mannose, glucose, and galactose), hexuronic acids (4-O-methyl-d-glucuronic acid, galacturonic acid, and glucuronic acid), small amounts of rhamnose and fucose, and an acetyl group [12]. These binding sugars can assemble into a range of various hemicellulose polysaccharides, such as galactan mannans, xylans, xyloglucan and β-1,3/1,4-glucans [12,15].
Lignin is a rigid aromatic, amorphous and hydrophobic polymer that has been recognized as a highly branched polymer with a variety of functional groups, such as aliphatic, phenolic hydroxyls, carboxylic, carbonyl, and methoxyl groups. These functional groups give lignin a unique and very complex structure [16,17,18]. The nature of the lignin polymerization reactions results in the formation of a three dimensional, highly-branched, interlocking network of essentially infinite molecular weight. Lignin composition and content are influenced by plant species and the environment [17,18].
The composition of cellulose, hemicellulose and lignin in agro-industrial waste depends upon the species, tissue and maturity of the plant [2,4,5,12]. The values of the main components in some agro-industrial waste are shown in Table 1.

3. Mushroom Cultivation on Agro-Industrial Wastes

Mushroom cultivation is widespread throughout the world and its global production has significantly increased since 2010 (Figure 2). The Food and Agriculture Organization Statistical Database (FAOSTAT) reported that China is the largest mushroom producer, followed by the United States of America and the Netherlands, with global production in 2018 reaching almost 8.99 million tons. The trend to increase mushroom production is expected to continue in the future.
Edible mushrooms are also considered a healthy food because they are rich in proteins, carbohydrates, fiber, vitamins and minerals while being low in fat [57,58]. Normally, the range of protein, carbohydrate and fat contents in mushrooms is 15–35%, 35–70% and less than 5%, respectively [58]. Notably, several species of edible mushrooms are important because of their medicinal properties. Some edible mushrooms appear to be active against human pathogens, cancer, diabetes, hypertension, hypercholesterolemia conditions and tumors [57,58,59]. Today, more than 50 species of edible mushrooms have been commercially cultivated throughout the world. Most commercial edible mushrooms belong to the genera Agaricus, Agrocybe, Auricularia, Flammulina, Ganoderma, Hericium, Lentinula, Lentinus, Pleurotus, Tremella, and Volvariella (Figure 3). The top four globally cultivated edible mushrooms include the genera Lentinula (shiitake and relatives), Pleurotus (oyster mushroom), Auricularia (wood ear mushroom) and Agaricus (button mushroom and relatives) [54,60]. In 2017, world mushroom production was divided among several genera: Lentinula (22%), Pleurotus (19%), Auricularia (18%), Agaricus (15%), Flammulina (11%), Volvariella (5%) and others (10%) [60]. Most of the cultivated edible mushrooms are saprophytic fungi (decomposers) and able to degrade lignocellulosic materials by producing extensive enzymes (especially lignocellulolytic enzymes). They are then able to use these materials as nutrients for their growth. Thus, mushroom cultivation is often associated with the recycling of vast amounts of agro-industrial waste [2,3,4,54].
Agro-industrial wastes (both agricultural residue and industrial residue) have been used as substrates in mushroom cultivation. Most agro-industrial waste is defined as low nitrogen content materials. The carbon/nitrogen (C/N) ratio in agro-industrial waste is varied among different types (Table 1), and it is an important factor in mushroom cultivation. This ratio has a critical influence on mycelium growth, mushroom weight, yields and protein content in the fruiting body of mushrooms [11,61,62]. Therefore, low-level nitrogen substrates for mushroom cultivation are necessary in that they add organic (cereal bran, cereal shell, soybean meal and manure) or inorganic (ammonium chloride and urea) nitrogen supplements [63,64]. Several previous studies have found that the protein content in the fruiting body of mushrooms depends upon both the chemical composition and the C/N ratio of substrates, as well as the species of mushroom being cultivated [1,64,65,66]. Different mushroom species require different C/N ratios in the cultivation substrate in order to obtain the highest production yield, as is shown in Table 2. Moreover, the addition of various supplements, e.g., epsom salts (MgSO4∙7H2O), gypsum (CaSO4·2H2O) and limestone (calcium carbonate, CaCO3), in the substrates also support the mycelia growth and fruiting body production of mushrooms [11,61,67].
Biological efficiency (BE), which is used to evaluate the efficiency of substrate conversion in mushroom cultivation, is calculated as the percentage ratio of the fresh weight of harvested mushrooms over the dry weight of the cultivation substrate [67]. A high BE value ensures a high possibility of utilizing substrates for mushroom cultivation [67,82]. In considering the profitability of mushroom cultivation, the BE value must be over 50%. Utilization of agro-industrial waste for the cultivation of mushrooms has resulted in the production of edible proteins for human consumption [2,7,11]. Cultivation methods for edible mushrooms vary considerably around the world and a variation in the chemical composition of a particular cultivated mushroom has been observed in various studies. This may be related to the specific mushroom species, the growing substrate and the relevant environmental conditions [1,2,11]. Many studies have been conducted to test the ability of mushrooms to grow on different agro-industrial forms of waste, such as wheat straw, barley straw, oat straw, rice straw, corn straw, corn cob, banana leaves, sawdust, sugarcane bagasse, soya stalk and sunflower stalk. A combination of agro-industrial waste can be used in mushroom cultivation. The main results regarding the cultivation of edible mushrooms on different agro-industrial waste, and their proximate composition values, are shown in Table 3 and Table 4.

4. Lignocellulolytic Enzyme Production by Mushroom Using Agro-Industrial Wastes

The decomposition of lignocellulosic materials is carried out by decomposers such as bacteria, microfungi, mushrooms, earthworms, and woodlice, all of which play an important role in the terrestrial carbon cycle [130,131,132]. Lignocellulose is a composite of three main biopolymers: cellulose, hemicellulose and lignin. Due to the different bonding functions that exist among these polymers, lignocellulose degradation requires the synergistic action of multiple carbohydrate-active enzymes. These are involved in the assembly and breakdown of glycosidic bonds [132,133,134]. The degradation of lignocellulosic biomass is achieved through cooperative activities of hydrolytic and oxidative enzymes [134,135,136], as is shown in Figure 4. The hydrolytic system is responsible for cellulose and hemicellulose degradations, whereas the oxidative system is known to participate in lignin degradation.

4.1. Cellulose Degradation Enzymes

Commonly, cellulose hydrolysis requires a combination of three main types of cellulase: endo-1,4-β-d-glucanase (endoglucanase, EC, exo-1,4-β-d-glucanase or cellobiohydrolases (exoglucanase, EC and β-glucosidase (β-d-glucoside glucanhydrolase, EC, in order to convert cellulose into oligosaccharides, cellobiose, and glucose [137,138]. The degradation of cellulose by various cellulase enzymes is diagrammed in Figure 5. Endoglucanases preferentially hydrolyze internal β-1,4-glucosidic linkages in the cellulose chains, generating a number of reducing ends [138,139]. This enzyme also acts on cellodextrins, which are the intermediate product of cellulose hydrolysis, and converts them to cellobiose and glucose. Exoglucanases release cellobiose from the reducing end or the nonreducing end of the cellulose chain, facilitating the production of mostly cellobiose which can readily be converted to glucose by β-glucosidases [136,140,141]. These enzymes may also act on cellodextrins and larger cello-oligosaccharides, in which case they are commonly named cellodextrinases [142]. Oligosaccharides released as a result of these activities are converted to glucose by the action of cellodextrinases (EC, whereas the cellobiose released mainly by the action of cellobiohydrolases is converted to glucose by β-glucosidases [139].
Cellulases are produced in a wide range of organisms such as plants, some animals, and certain microorganisms including protozoans, bacteria, and fungi. Among these organisms, fungi have been studied extensively for their cellulase producing capabilities, such as the genera Aspergillus, Penicillium, Rhizopus and Trichoderma [143,144,145,146]. However, mushrooms are the most potent degraders of natural lignocellulosic waste. They are mostly grown on litter, dead wood, or in soil and nature-rich cellulose [9]. Several previous reports have found that various mushrooms species can produce cellulase via solid state fermentation (SSF) of agricultural or natural lignocellulosic waste [147,148,149]. Many agricultural or natural lignocellulosic solid waste, especially different kinds of straw (wheat, sorghum, rice) and sawdust (oak and pine), were used as a substrate or source for mushroom growth and cellulases production [150,151,152,153]. Furthermore, other forms of lignocellulosic waste, such as peanut hulls, mandarin peels, cotton waste, corn stovers and tree leaves (Fagus sylvatica), have also been used as substrates to determine cellulase activity [150,154,155]. The high-value potential of these forms of waste is encouraging as they can be sources that support the growth and cellulases production of different mushroom species, namely Ganoderma, Grifola, Lentinula, Lentinus, Pleurotus, Piptoporus and Trametes by SSF [152,156,157,158,159]. Different agricultural or natural lignocellulosic forms of waste that have been fermented by various mushroom species are summarized in Table 5.
Cellulase activity is mainly tested using a reducing sugar assay to determine cellulase hydrolysis activity at the end of the production process [169]. The common enzyme activity assays consist of total cellulase assays, endoglucanase assays, exoglucanase assays and β-glucosidase assays [140]. Filter paper assay (FPA) is widely used to determine total cellulase activity. The degree of filter paper activity is determined as the micromole of glucose equivalent liberated per minute of culture filtrate under assay conditions [170]. Endoglucanase activity can be measured using the carboxymethyl cellulose (CMC) as a substrate. This carboxymethyl cellulase (CMCase) is mainly measured by examining the reducing sugars of enzymatic reactions with CMC based on the procedure described by [171]. The exoglucanase activity mainly uses commercial Avicel as a substrate for measuring the activity [169]. The β-glucosidase assay can be measured based on the procedure of Kubicek [172] using chromogenic and nonchromogenic substrates such as p-nitrophenol-β-glucoside (pNPG) and cellobiose, respectively [173,174]. Moreover, various reducing sugar assays, for instance, 3,5-dinitrosalicylic acid (DNS), glucose oxidase (GOD) and high-performance liquid chromatography were also used.

4.2. Hemicellulose Degradation Enzymes

Hemicelluloses are usually classified based on the backbone sugars present in the structural polymer with typical glucose galactose, xylose, mannose, and arabinose. The principal hemicelluloses are comprised of xyloglucans, xylans, mannans, glucomannans, and mixed linkage β-glucans [175,176]. In order to digest hemicellulose, microorganisms need to be able to produce a variety of enzymes to hydrolyze complex substrates with a synergistic action. Hemicellulolytic enzymes or hemicellulases are glycoside hydrolases or carbohydrate esterases that are responsible for polysaccharide degradation. The enzymes include xylanase (EC, β-xylosidase (EC, α-arabinofuranosidase (EC α-glucuronidase (EC, and β-mannosidases (EC [134,138].

4.2.1. Xylanases

Xylan is a heteropolysaccharide and a major hemicellulose. The main chain of xylan consists of β-1,4-linked d-xylopyranosyl residues, which are partially replaced with O-acetyl, l-arabinosyl and 4-O-methyl-d-glucuronic acid. The xylan backbone is substituted by different side chains with l-arabinose, d-galactose, d-mannoses, and glucouronic acid linked by glysosidic bonds and ester bonds with ferulic acid [177,178,179,180]. Biodegradation of xylan requires diverse modes of action of hydrolytic enzymes. Xylanases are a group of glycoside hydrolase enzymes that breakdown hemicelluloses through the degradation of the linear polysaccharide xylan into xylose by catalyzing the hydrolysis of the glycosidic linkage (β-1,4) of xylosides. The xylanolytic enzyme system includes a mixture of endo-1,4-β-xylanases also called endo-xylanases, β-xylosidases, α-arabino- furanosidases, α-glucuronidases and acetylxylanases, which attach to the specific site of xylan as is displayed in Figure 6 [181,182]. Endo-xylanases randomly hydrolyze β-1,4-xylanopyranosyl linkages of xylan to form xylo-oligosaccharides, xylotriose, xylobiose and xylose. The hydrolysis of xylans is not attacked randomly but depends upon the degree of branching, chain length, and presence of substituents in the substrate molecule [183].
β-Xylosidase attacks from the non-reducing end of xylo-oligosaccharides, xylotriose or xylobiose, that are generated by the action of endo-xylanase and ultimately liberate xylose sugar (Figure 6). Biomass can be used as a substrate for this enzyme production process. However, a limitation of the commercial application of this substance is related to various factors such as their physical limitations, the limited hydrolysis of xylans due to their diverged branched nature, the fact that their enzymes are associated with a narrow pH range and thermal instability, their end product inhibition levels and the cost of enzyme production. These comprise the unreachability of substrates to xylanase enzymes [178]. The use of substrates with agricultural or industrial biomass for enzyme production serves as an alternative way to overcome the limitations of the costs of enzyme production; however, biomass pretreatment is sometimes needed to improve efficiency in the practical hydrolysis of biomass.
Many microorganisms, such as fungi, bacteria and yeast, can degrade hemicellulose by producing xylanases. A determination of xylanase activities can be analyzed by several methods. The plate assay has been used for decades as a primary screening method to select xylanase producing strains. The screening strains are cultured on agar medium containing xylan as their carbon source until clear zones are observed (the xylan hydrolysis area) after being stained with Congo red dye [183] or Gram’s iodine solution [184]. Plate assay methods rely on interactions between a dye and a polymeric substrate for the indirect detection of hydrolysis but require the use of relevant controls and independent confirmation of the relevant enzymatic activities. Xylans, such as oat spelt, beech wood [185], and birch-wood xylans [186,187], was used as a substrate to determine endo-xylanase activity. The enzyme activities were determined from the presence of reducing sugars as xylose equivalents liberated from the enzymatic hydrolysis by the DNS method [188] or the Nelson [189] and Somogyi [190] methods. However, xylans obtained from natural sources contain not only xylose residues but also arabinose and glucuronic acid residues. Thus, comparisons of xylanase activity in various studies have been difficult. Xylanase activity varies according to the source of the xylans. Other types of substrates can be applied. Specifically, p-nitrophenyl-glycoside substrate (p-nitrophenyl esters with substrate) can be used as a chromogenic substrate for the calorimetric assay of β-xylosidase activity. The substrate is colorless in neutral or alkaline solution. After enzymatic hydrolysis, p-nitrophenol is liberated as alkaline pH develops a yellow color that is suitable for the quantitative measurement of the enzyme activity.
Multifunctional xylanolytic enzyme system is relatively common in fungi, actinomycetes and bacteria [190,191]. A large variety of industrial xylanase enzymes are produced from various kind of microorganisms [192]. SSF with batch processing has been used for the utilization of agro-industrial waste [193]. However, very few studies have reported on the xylanolytic enzymes obtained from mushroom on SSF (Table 6). These potential outcomes provide opportunities for scientists to explore the hydrolytic potential of xylanase for the efficient saccharifcation of lignocellulosic biomass from mushroom cultivation.

4.2.2. Mananases

The two most important and representative hemicelluloses are xylans and mannans. Mannans are polysaccharides that consist of mannose-based backbones linked by β-1,4-linkage with variable degrees of side substitutions. These polysaccharides are renewable resources and their enzymatic conversion is of great interest in the field of lignocellulose biotechnology [194]. The enzyme breakdown of mannans is accomplished with β-mannanase (β-1,4-d-mannan mannohydrolase, EC as it randomly attacks the internal β-1,4-d-mannopyranosyl linkage within the main chain of various mannan-based polysaccharides, such as galactomannans, glucomannans, and galactoglucomannans, to release mannooligosaccharides (MOS), manotetrose, manotriose and manobiose [176].
The degradation of the mannan backbone is performed by the action of β-mannanases, and the further degradation requires β-mannosidase (β-1,4-d-mannopyranoside hydrolase, EC to hydrolyze the terminal ends (non-reducing ends) of MOS into sugar-based mannose. Subsequently, β-glucosidases remove 1,4-glucopyranose units at the non-reducing ends of the oligomers derived from the degradation of glucomannan and galactoglucomannan [171,205] as is shown in Figure 7. Xylanases and mannanases are important enzymes for the hydrolysis of hemicelluloses. β-mannan is found in many feedstuffs including soybean meal, palm kernel meal, copra meal, and sesame meal and other leguminous feeds [206]. β-Mannanases are widely applied to randomly hydrolyze the β-1,4-mannopyranoside linkage of mannan-based polysaccharides in many industries.

4.2.3. Arabinanases

Arabinanases are a group of hydrolytic enzymes that include endo-arabinanases (EC, arabinosidases (EC, and α-L-arabinofuranosidase. These work synergistically to generate l-arabinose from arabinan as is shown in Figure 8 [207,208,209,210,211]. The biodegradation of xylan requires the cooperation of xylanases, β-xylosidase, α-l-arabinofuranosidase, α-glucuronidase, and acetylxylanases [181,182]. The removal of the side groups of xylans is catalyzed by α-l-arabinofuranosidases (E.C., α-d-glucuronidases and acetylxylan esterases, which remove acetyl and phenolic side branches and act synergistically on the complex polymer [178]. Fungi produce extracellular arabinanases, a group of hydrolytic enzymes that include α-l-arabinofuranosidases and endo-arabinanases to specifically release l-arabinose from polysaccharides including xylans and pectin [212]. Importantly, α-l-arabinofuranosidases catalyze the hydrolysis of α-l-arabinofuranosidic linkage at terminal non-reducing- α-l-1,2-, α-l-1,3- and α-l-1,5-arabinofuranosyl residues obtained from different oligosaccharides and polysaccharides (α-l-arabinosides, arabinans, arabinoxylans, and arabinogalactans) and act synergistically with other hemicellulases to completely breakdown hemicellulose [212,213]. The l-arabinofuranoside substitutions on xylan strongly inhibit the action of xylan-degrading enzymes, thus preventing the complete degradation of xylan to xylose units [213]. The α-l-arabinofuranosidases can be found in plants, bacteria and fungi [186].
The colorimetric method is used to determine α-l-arabinofuranosidases activity. Notably, the p-nitrophenol-linked substrate, 4-nitrophenyl α-l-arabinofuranoside, is used for the enzyme assay by determining the amount of p-nitrophenol released from the enzyme-substrate reaction [186,214,215]. Arabinoxylans, such as wheat four arabinoxylan and sugar beet arabinan, is also used for the determination of enzyme activity [180] by monitoring the generation of arabinose from polysaccharide substrates. Liberated arabinose can be determined by the DNS method [187].

4.3. Lignin Degradation Enzymes

Lignin degradation is the primordial step in lignocellulose degradation enabling the accessibility of cellulose and hemicellulose [216,217]. Ligninolytic microorganisms can degrade lignins via the secretion of oxidative enzymes, such as peroxidases and laccases, or by producing a source of heterogeneous aromatics. Ligninolytic enzymes or ligninases are mainly comprised of laccases (Lac, EC, lignin peroxidases (LiPs, EC, manganese peroxidases (MnPs, EC, versatile peroxidases (VPs) and dye decolorizing peroxidases (DyPs, EC [116,218]. These enzymes display less substrate specificity than cellulases and hemicellulases [124,218,219]. Additionally, Lac, LiP and MnP, and many other enzymes, such as aromatic acid reductase, aryl alcohol dehydrogenase, catalase aromatic aldehyde oxidase, dioxygenase, quinone oxidoreductase, vanillate hydroxylase, veratryl alcohol oxidase and versatile peroxidase, are also involved in lignin digestion [219].
Mushroom species are most frequently reported as Lac and MnP producers and least frequently reported as LiP and VP producers. Previous publications have reported that T. versicolor [220] and Bjerkandera adusta [221] produce both oxidase (Lac) and peroxidase (MnP and LiP). Lentinula edodes [222], P. eryngii [223] and Ceripotiopsis subvermispora [224] are lignin-degrading mushrooms that use Lac and at least one of the peroxidases. Only Lac was produced from S. commune and only peroxidases were produced from Phanerochaete chrysosporium [225,226]. Several publications have reported that Ph. chrysosporium is an excellent lignin decomposer, and it has been suggested for its commercial use. The ligninolytic enzymes were fermented in SSF using different agro-industrial waste, as is shown in Table 7.

4.3.1. Laccases

Laccases are a group of multicopper containing enzymes belonging to the blue multicopper oxidase family. The enzymes are also known as polyphenol oxidases, among which laccases oxidize one-electron of phenolic compounds with an associated reduction of oxygen to water as a by-product [240,241]. The enzymes do not require H2O2 for substrate oxidation. Lac can oxidize both phenolic aromatic compounds such as methylated phenol, aromatic amine and non-phenolic aromatic compounds such as veratryl alcohol in lignin to form phenoxy-free radicals. In this way, lignin degradation and lignin structural conversion can occur [242], as is shown in Figure 9. This oxidation process produces phenoxy radicals that can be converted to quinine by a second enzyme catalyzed reaction [166,243].
Laccases contain four copper ions except for the laccase that is obtained from Phlebia radiata, which has only two copper ions [245]. There are three types of Lac depending on the copper number at the active site [246]. Type I: copper does not bind O2 but functions only as an electron transfer site. The type I copper center consists of a single copper atom that is coordinated with two histidine residues and one cysteine residue. In some cases, a methionine motif serves as a ligand with a trinuclear center. The Type II copper center has two histidines and a water molecule that serves as a ligand. The type III copper center contains two copper atoms that each possess three histidine ligands and are linked to one another via a hydroxide bridging ligand. Most of the studies on Lac have reported that the fungi and mushrooms present in basidiomycetes, deuteromycetes and ascomycetes act as Lac producers [247]. Among these fungi, the major Lac producers are white-rot fungi in basidiomycetes [246]. White-rot fungi Pycnoporus cinnabarinus, Phlebia radiate, P. ostreatus, and T. versicolour are also known to produce one isoform of Lac [248]. Cotton stalks, aromatic compounds, wood, and plant extracts were found to be inducers for Lac production [249]. For Lac production, extracted 3-hydroxyanthranilic acid (3-HAA) obtained from wheat straw was found to be a potential Lac stimulator [250]. The mixture of coffee pulp and urea was also able to enhance the Lac activity in Py. sanguineus culture. Some researchers have found a novel Lac obtained from T. orientalis, which has a molecular mass of 44.0 kDa. The enzyme contains a typical copper II binding domain and shares three N-glycosylation sites. But it has no copper I binding domain [251] Dias and colleagues [252] have reported a new zymogram dried 2,2’-azino-bis(3-ethylbenzo- thiazoline-6-sulfonic acid) (ABTS)-impregnated discs assay for laccase activity detection, which is associated with easy assay and rapid screening. The laccase activity was determined at a wavelength of 420 nm by measuring the oxidation of ABTS in phosphate citrate buffer at a pH value of 4.0 [253]. The other guaiacol assay has been reported for laccase assay by Kalra et al. [254] to measure the reddish-brown color development at 450 nm as a consequence of the oxidation of guaiacol by Lac.

4.3.2. Lignin Peroxidases

Lignin peroxidase (LiP) belongs to the family of oxidoreductases. LiP has ferric heme as an electron donor which is able to reduce oxygen molecules to hydrogen peroxidase and superoxides. LiP-Fe(III) uses H2O2 to oxidize aryl cation radicals as the initial substrate. The resulting amount of the lacked electron LiP is not stable and draws electrons from the substrate for stability of the electron condition. Finally, the oxidation cycle ends when LiP-Fe(IV) is turned to the resting ferric state [255]. This reaction exhibits a degree of stoichiometry of one H2O2 compound consumed per the amount of aldehyde formed. LiP is a strong oxidant and is non-specific with a substrate. It can degrade both structures of phenolic aromatic and non-phenolic aromatic compounds. Veratryl alcohol was found to be an inducer of LiP that was produced from white-rot fungi. The molecular weight of LiP was approximately 41 kDa and contains one mole of Fe protoporphyrin IX. It is a glycoprotein with isoelectric point (pI) as 3.2–4.0 that displays high redox potential activity and an optimum pH value at 3.0 [250].
There are two methods for lignin peroxidase detection [250]. One involves the measurement of veratraldehyde from veratryl alcohol oxidation using a UV spectrophotometer at 310 nm. One unit of activity is defined as one micromole of veratryl alcohol oxidized in one min, while the activities are reported in units/L (U/L). The 1,2-bis(3,4-dimethoxyphenyl) propane-1,3-diol is a substrate of this enzyme, whereas 3,4-dimethoxybenzaldehyde, 1-(3,4-dimethoxyphenyl) ethane-1,2-diol, and H2O, are its products, as is displayed in Figure 10.
The other method is the Azure B assay. In this method, the relevant reaction assay contains Azure B dye, H2O2, and sodium tartrate buffer (pH 4.5). The activity is measured at a 615 nm wavelength [256]. This method has been identified as a good assay to reduce the turbidity caused by organic materials under the UV range. Mushrooms have been found as the first LiP producers, namely T. versicolor, P. ostreatus, G. lucidum, and Bjerkandera spices [232,257].

4.3.3. Manganese Peroxidase

Manganese peroxidase (MnP) belongs to the family of oxidoreductases and cannot react directly with the lignin structure [250]. There are two groups: (1) Manganese dependent peroxidase is an extracellular enzyme that requires both H2O2 for lignin oxidation, Mn2+ as a co-factor and (2) Manganese independent peroxidase is an extracellular enzyme that requires H2O2 in lignin oxidation but does not need Mn2+ (Figure 11) [258]. The major substrates of manganese peroxidase are low molecular weight substances and organic acid compounds. In the mechanism cycle of lignin degradation, Mn2+ is an electron donor and MnP is oxidized by H2O2 as follows:
“MnP + H2O2 → MnP compound I + H2O”
“MnP compound I + Mn2+ → MnP compound II + Mn3+
“MnP compound II + Mn2+ → MnP + Mn3+ + H2O”
The electron-lacking MnP is nonstable and accepts an electron from Mn2+ to Mn3+ that then reacts with certain organic acid chelators such as oxalate, malonate, and lactate. The chelated-Mn3+ will act as a mediator to oxidize simple phenols, amines, and phenolic lignins. The enzyme can oxidize both phenolic and non-phenolic lignins [260]. The 3,3’-diaminobenzidine (DAB) assay [261] and manganese peroxidase (MnP) assay [262] are the methods used for identification of peroxidase using 0.01% phenol red or 2 mM 2,6-dimethoxyphenol (DMP) as a substrate.
Many mushroom species have been identified as MnP-producing fungi, especially P. ostreatus and Ph. chrysosporium [263]. Manganese dependent peroxidase is produced from P. pulmonarius, which can oxidize both non-phenolic and phenolic compounds for xenobiotic compound degradation. Kuhar and co-workers [264] have reported that MnCl2 can induce MnP activity and has a high specificity for Mn2+ binding sites.

4.3.4. Versatile Peroxidase

Versatile peroxidase (VP) is also known as a hybrid peroxidase or polyvalent peroxidase for Mn2+ oxidation. VP includes both LiP and MnP activities. Consequently, VP is able to degrade a wider range of substrates than non-hybrid enzymes. VP requires H2O2 as an electron acceptor to catalyze the oxidative reaction at the heme center with the release of a water molecule [250]. VP is a heme-containing glycoprotein that has a two-channel structure: the wider channel for access to H2O2 and the narrow channel for access to manganese. Low molecular substrates will be oxidized at the heme center by H2O2-ferric state binding (heme forming iron peroxide complex). This activated heme complex is able to oxidize the aromatic substrate using Mn2+, and then secretes Mn3+ and water [265] (Figure 12). VP has been produced by SSF of P. eryngii and P. ostreatus on wheat straw, sawdust, and banana peels [223,266]. Pleurotus ostreatus and Bjerkandera sp. were cultured in glucose-peptone broth and glucose ammonium medium using submerged fermentation for VP production [267]. The molecular weight and pI of VP obtained from P. eryngii were approximately 40 kDa and 4.1, respectively [268]. The VP activity can be determined by monitoring manganese oxidation and Reactive clack (RB5) decolorization [267].

4.3.5. Dye Decolorizing Peroxidases

Dye decolorizing peroxidases (DyPs) are a new family of glycoproteins that have one heme as a cofactor occurring in basidiomycetous fungi and eubacteria. DyPs require H2O2 as an electron acceptor and are similar to VP; however, DyPs can oxidize the high-redox potential anthraquinone dyes in addition to typical peroxidase substrates such as RBs, phenols, veratryl alcohol [269,270]. There are four types of DyPs from A to D based on their primary sequences [271]. However, type A DyPs has been reported as the potential type that is most effective in lignin depolymerization. The important characteristic of DyPs is the degradation of hydroxyl-free anthraquinone, which is not a substrate of other peroxidases [270]. DyPs can oxidize certain phenolic compounds such as 2,6-dimethoxyphenol and guaiacol. Only a few types of fungi can produce DyPs, especially type d-DyP, whereas they are mostly present in bacteria (types A, B, and C). The first DyP was discovered in B. adusta [272]. The wood-rotting fungi A. auricula-jadae, Mycetinis scorodonius, Exidia glandulosa, P. sapidus DSM8266 and Mycena epipterygia have also been reported as DyPs producers [273,274]. White-rot fungus, Irpex lacteus CD2, exhibited DyPs activity when it was grown in Kirk’s medium containing lignins [275]. Many previous publications have reported that DyPs might be important for the ligninolytic system in white-rot fungi despite the fact that the biological roles of DyPs are unknown in terms of different substrate specificities. The mechanism of DyPs is similar to that of plant peroxidase, which is known to generate transient intermediates (compound I and compound II). The reaction of compound I with 1 eq electrons from a reducing substrate generates the [FeIV = O]+ intermediate compound II [271]. The optimum pH value of DyPs is acidic [276]. DyPs activity was assayed by the decolorization of an anthraquinone dye RB19 at 595 nm [275].

4.4. Application of Lignocellulolytic Enzymes in Bioprocessing

Enzyme technology possesses great potential to reduce environmental pollution and offers potential benefits in the comprehensive utilization of lignocellulosic biomass. Lignocellulolytic enzymes have received attention because of their potential applications in various agro-industrial bioprocesses, such as the conversion of hemicellulosic biomass to fuels and chemical production, the clarification of juices, the green processing of certain foods and beverages, the enhancement of animal digestibility in feedstock, the delignification of paper and pulp, the improvement of fabric properties in the textile industry and waste utilization [277,278,279]. Cellulase is widely used in the textile and laundry detergent industries as it can play a part in the hydrolysis of cellulose and improve fabric properties for the textile industry and for cleaning textiles in the laundry detergent industry [154,280]. The food and beverage processing industries have used cellulase for the hydrolysis of cellulose during the drying of coffee beans and for the extraction of fruits and vegetables in juice production [281,282]. Cellulase, α-l-arabinofuranosidases and other glycosidases have also been used in brewery and wine production [213,277]. The enzymatic hydrolysis of grapes utilizes α-l-arabinofuranosidases and other glycosidases to enhance the flavor of wine by the release of free terpenols, an important aspect in the development of the aroma in wine. The enzyme treatment by α-l-arabinofuranosidases during sourdough preparation in the bread industry delays the staling process of bread and increases the shelf life of bread [213]. This results in economic benefits in terms of the preservation of bread and bread storage issues. Enzyme technology has a significant potential to improve the properties of pulp. Cellulases, xylanase and other hemicellulases are commonly used enzymes to assist in pulp bleaching for the reduction of environmental pollution loads [283]. Cellulases are used to improve the performance of dissolved pulp [277]. Additionally, α-l-arabinofuranosidases enhance the delignification of pulp in the bleaching process as it can cleave the arabionose side chain that inhibits the action of xylanase [213]. Laccase can be used for lignin removal in prehydrolysis of lignocellulosic biomass [284]. Xylanolytic enzymes have potential applications across food and feed industries [278]. A combination of α-l-arabinofuranosidases with cellulases, pectinases and xylanases enhance the feed digestibility and utilization of polysaccharides in feedstuffs [186,213]. Arabinoxylans are the major non-starch polysaccharide fractions in wheat, which increase digesta viscosity, reduce the digestibility of nutrients and decrease the feed efficiency and growth performance when fed to poultry, especially in broiler chickens [278]. Various reports have revealed the positive effects of MOS on intestinal microflora, along with efficient intestinal structure and function. MOS-based nutrition supplements are widely used in nutrition as a natural additive [279]. The treatment of copra meal rich in β-mannan with mannanase has been reported to reduce the population of Salmonella and Escherichia coli, increase the level of metabolizable energy and improve the nutrient digestibility in broilers [285]. Olaniyi et al. [207] reported that the treatment of cassava peels and corn cobs with mannanase increased the degradation of the complex carbohydrate fractions in the samples and resulted in increasing the amount of crude protein and certain mineral contents. Kim et al. [273] reported that the supplementation of β-mannanase for diet feeds does not mitigate the heat stress of aged laying hens raised under hot climatic conditions. Saeed et al. [206] describes the promising beneficial effects of β-mannanase in the poultry feed industry as the supplementation of β-mannanase in poultry diets that positively improved blood glucose and anabolic hormone homeostasis, digestible energy, and digestible amino acids. These enzymes have been used as food additives in the poultry raising industry and have been employed in the improvement of nutritional properties of agricultural silage and grain feed.
Manganese peroxidase is an important enzyme associated with the lignin and organic pollutant degradation systems, for instance bioremediation, dye decolorization, pulp bleaching, biomechanical pulping and in the production of a range of highly valuable products that have been obtained from residual lignins [286]. DyPs can be applied in the treatment of wastewater that contain synthetic dyes which are used in the manufacture of textiles, cosmetics, food, and pharmaceuticals. In the food industry, DyPs obtained from M. scorodonius, namely the MaxiBright® brand, are used to whiten whey in cheese making [274]. Enzymes have been extensively used in various industries as well as in a lot of the resulting products. Thus, genetic engineering is a powerful tool for the enhancement of ligninolytic enzyme production. White-rot fungus, Ph. chrysosporium, is a good model for the study of lignin degradation using DNA technology. The genome sequence encoded several genes such as ten lignin peroxidases, five manganese peroxidases, and several other lignocellulolytic enzymes [287,288]. Laser mutagenesis of Phellinus igniarius SJZ2 (mutant) overexpressed Lac activity during 4 h of fermentation and was increased by 36.84% in comparison with the wild type [242]. In addition to the overexpression of Lac in Saccharomyces cerevisiae using the laccase III (cvl3) gene obtained from T. versicolor, IFO1030 was secreted in the culture (45 U/L) [289]. Lignocellulosic enzymes are obtained from mushrooms, especially white-rot basidiomycetes, which are interesting tools in the biotechnological process that is used in a wide range of lignin substrates.

5. Conclusions

The utilization of agro-industrial waste in mushroom cultivation and the production of lignocellulolytic enzymes can facilitate the reduction of some global waste management problems. The cultivation of edible mushrooms using agro-industrial waste represents the bioconversion of that waste into edible protein. Different types of agro-industrial waste can be used for the cultivation of substrates for mushroom cultivation. However, the composition and availability of agro-industrial waste in each area has been considered for the support of mushroom cultivation. Different mushroom species and C/N ratios in substrates are the crucial factors that affect the production and chemical composition of mushrooms. The nitrogen content of agro-industrial waste is low; therefore, this waste is generally associated with other nitrogen sources. The selected suitable substrate and mushroom species are important in obtaining the maximum yields.
Mushrooms seem to be the most important players in lignocellulose degradation by producing both hydrolytic and oxidative enzymes. Hydrolytic enzymes (cellulases and hemicellulases) are known to be responsible for polysaccharide degradation, while oxidative enzymes (ligninases) are responsible for lignin modification and degradation. Current results indicate that agro-industrial waste has been evaluated for its potential use in lignocellulosic enzyme production by mushrooms. However, the variability of waste composition and mushroom species are influential in enzyme production. Therefore, further studies are needed to demine the suitable conditions (substrates, mushroom species and fermentation process) for effective lignocellulosic enzyme production in the pilot study and on the industrial scale.

Author Contributions

The project approach was conceptually designed by J.K., N.S., S.L.; writing and original draft preparation, J.K., N.S., W.P., K.S., P.K., K.J., S.V.; chemical structure drawing, K.S.; the research was supervised by J.K., N.S., S.L.; All authors have read and agreed to the published version of the manuscript.


This research work was partially supported by Chiang Mai University.


We are grateful to Russell K. Hollis for the English proofreading of this manuscript.

Conflicts of Interest

The authors declare that they have no conflicts of interest.


  1. Mirabella, N.; Castellani, V.; Sala, S. Current options for the valorization of food manufacturing waste: A review. J. Clean. Prod. 2014, 65, 28–41. [Google Scholar] [CrossRef][Green Version]
  2. Sadh, P.K.; Duhan, S.; Duhan, J.S. Agro-industrial wastes and their utilization using solid state fermentation: A review. Bioresour. Bioprocess 2018, 5, 1. [Google Scholar] [CrossRef][Green Version]
  3. Panesar, P.S.; Kaur, R.; Singla, G.; Sangwan, R.S. Bio-processing of agro-industrial wastes for production of food-grade enzymes: Progress and prospects. Appl. Food Biotechnol. 2016, 3, 4. [Google Scholar]
  4. Ravindran, R.; Jaiswal, A.K. Exploitation of food industry waste for high-value products. Trends Biotechnol. 2016, 34, 58–69. [Google Scholar] [CrossRef] [PubMed][Green Version]
  5. Anwar, Z.; Gulfraz, M.; Irshad, M. Agro-industrial lignocellulosic biomass a key to unlock the future bio-energy: A brief review. J. Radiat. Res. Appl. Sc. 2014, 7, 163–173. [Google Scholar] [CrossRef]
  6. Cherubin, M.R.; Oliveira, D.M.D.S.; Feigl, B.J.; Pimentel, L.G.; Lisboa, I.P.; Gmach, M.R.; Varanda, L.L.; Morais, M.C.; Satiro, L.S.; Popin, G.V.; et al. Crop residue harvest for bioenergy production and its implications on soil functioning and plant growth: A review. Sci. Agricola 2018, 75, 255–272. [Google Scholar] [CrossRef][Green Version]
  7. da Silva, L.L. Adding value to agro-Industrial wastes. Ind. Chem. 2016, 2, e103. [Google Scholar] [CrossRef]
  8. Hongzhang, C. Biotechnology of Lignocellulose: Theory and Practice; Springer: New York, NY, USA, 2016. [Google Scholar]
  9. Sánchez, C. Lignocellulosic residues: Biodegradation and bioconversion by fungi. Biotechnol. Adv. 2009, 27, 185–194. [Google Scholar] [CrossRef]
  10. Knob, A.; Forthamp, D.; Prolo, T.; Izidoro, S.C.; Almeida, J.M. Agro-residues as alternative for xylanase production by filamentous fungi. BioResources 2014, 9, 5738–5773. [Google Scholar]
  11. Grimm, D.; Wösten, H.A.B. Mushroom cultivation in the circular economy. Appl. Microbiol. Biotechnol. 2018, 102, 7795–7803. [Google Scholar] [CrossRef][Green Version]
  12. Zhou, X.; Broadbelt, L.J.; Vinu, R. Mechanistic understanding of thermochemical conversion of polymers and lignocellulosic biomass. Adv. Chem. Eng. 2016, 49, 95–198. [Google Scholar]
  13. Heinze, T. Cellulose: Structure and properties. In Cellulose Chemistry and Properties: Fibers, Nanocelluloses and Advanced Materials; Rojas, O., Ed.; Springer: Cham, Switzerland, 2016; Volume 271, pp. 1–52. [Google Scholar]
  14. Jedvert, K.; Heinze, T. Cellulose modification and shaping—A review. J. Polym. Eng. 2017, 37, 845–860. [Google Scholar] [CrossRef]
  15. Ebringerová, A.; Hromádková, Z.; Heinze, T. Hemicellulose. Adv. Polym. Sci. 2005, 186, 1–67. [Google Scholar]
  16. Geneau-Sbartai, C.; Leyris, J.; Slivestre, F.; Rigal, L. Sunflower cake as a natural composite: Composition and plastic properties. J. Agric. Food Chem. 2008, 56, 11198–11208. [Google Scholar] [CrossRef]
  17. Rico-García, D.; Ruiz-Rubio, L.; Pérez-Alvarez, L.; Hernández-Olmos, S.L.; Guerrero-Ramírez, G.L.; Vilas-Vilela, J.L. Lignin-based hydrogels: Synthesis and applications. Polymers 2020, 12, 18. [Google Scholar] [CrossRef][Green Version]
  18. Davin, L.B.; Lewis, N.G. Lignin primary structures and diligent sites. Curr. Opin. Biotechnol. 2005, 16, 407–415. [Google Scholar] [CrossRef]
  19. Nawirska, A.; Kwaśniewska, M. Dietary fibre fractions from fruit and vegetable processing waste. Food Chem. 2005, 91, 221–225. [Google Scholar] [CrossRef]
  20. Silveira, M.L.L.; Furlan, S.A.; Ninow, J.L. Development of an alternative technology for the oyster mushroom production using liquid inoculum. Cienc. Technol. Aliment. 2008, 28, 858–862. [Google Scholar] [CrossRef][Green Version]
  21. Tarrés, Q.; Espinosa, E.; Domínguez-Robles, J.; Rodríguez, A.; Mutjé, P.; Aguilar, M.D. The suitability of banana leaf residue as raw material for the production of high lignin content micro/nano fibers: From residue to value-added products. Ind. Crop. Prod. 2017, 99, 27–33. [Google Scholar] [CrossRef]
  22. Nigam, P.S.; Gupta, N.; Anthwal, A. Pre-treatment of agro-industrial residues. In Biotechnology for Agro-Industrial Residues Utilization; Nigam, P.S., Pandey, A., Eds.; Springer: Dordrecht, The Nederlands, 2009; pp. 13–33. [Google Scholar]
  23. Adapa, P.K.; Tabil, L.G.; Schoenau, G.J.; Canam, T.; Dumonceaux, T. Quantitative ananlysis of lignocellulosic companents of non-treated and stream exploded barley, canola, oat and wheat straw using fourier transform infrared spectroscopy. J. Agric. Sci. Technol. 2011, B1, 177–188. [Google Scholar]
  24. Carrijo, O.A.; Liz, R.S.; Makishima, N. Fiber of green coconut shell as an agricultural substrate. Hortic. Bras. 2002, 20, 533–535. [Google Scholar] [CrossRef]
  25. Graminha, E.B.N.; Gonçalvez, A.Z.L.; Pirota, R.D.P.B.; Balsalobre, M.A.A.; da Silva, R.; Gomes, E. Enzyme production by solid-state fermentation: Application to animal nutrition. Anim. Feed Sci. Technol. 2008, 144, 1–22. [Google Scholar] [CrossRef]
  26. Gouvea, B.M.; Torres, C.; Franca, A.S.; Oliveira, L.S.; Oliveira, E.S. Feasibility of ethanol production from coffee husks. Biotechnol. Lett. 2009, 31, 1315–1319. [Google Scholar] [CrossRef] [PubMed]
  27. Rofiqah, U.; Kurniawan, A.; Aji, R.W.N. Effect of temperature in ionic liquids pretreatment on structure of lignocellulose from corncob. J. Phys. Conf. Ser. 2019, 1373, 1–7. [Google Scholar] [CrossRef]
  28. Pointner, M.; Kuttner, P.; Obrlik, T.; Jager, A.; Kahr, H. Composition of corncobs as a substrate for fermentation of biofuels. Agron. Res. 2014, 12, 391–396. [Google Scholar]
  29. El-Tayeb, T.S.; Abdelhafez, A.A.; Ali, S.H.; Ramadan, E.M. Effect of acid hydrolysis and fungal biotreatment on agro-industrial wastes for obtainment of free sugars for bioethanol production. Braz. J. Microbiol. 2012, 43, 1523–1535. [Google Scholar] [CrossRef][Green Version]
  30. Sun, Y.; Chen, J. Hydrolysis of lignocellulosic material for ethanol production: A review. Bioresour. Technol. 2002, 83, 1–11. [Google Scholar] [CrossRef]
  31. Abbas, A.; Ansumali, S. Global potential of rice husk as a renewable feedstock for ethanol biofuel production. Bioenerg. Res. 2010, 3, 328–334. [Google Scholar] [CrossRef]
  32. Limayema, A.; Ricke, S.C. Lignocellulosic biomass for bioethanol production: Current perspectives, potential issues and future prospects. Prog. Energy Comb. Sci. 2012, 38, 449–467. [Google Scholar] [CrossRef]
  33. Buzala, K.; Przybysz, P.; Rosicka-Kaczmarek, J.; Kalinowska, H. Comparison of digestibility of wood pulps produced by the sulfate and TMP methods and woodchips of various botanical origins and sizes. Cellulose 2015, 22, 2737–2747. [Google Scholar] [CrossRef][Green Version]
  34. Chuayplod, P.; Aht-ong, D. A study of microcrystalline cellulose prepared from parawood (Hevea brasiliensis) sawdust waste using different acid types. J. Met. Mater. Miner. 2018, 28, 106–114. [Google Scholar]
  35. Da Silva Neta, J.M.; Oliveira, L.S.C.; da Silva Flavio, L.H.; Tabosa, J.N.; Pacheco, J.G.A.; da Silva, M.J.V. Use of sweet sorghum bagasse (Sorghum bicolor (L.) Moench) for cellulose acetate synthesis. BioResources 2019, 14, 3534–3553. [Google Scholar]
  36. Dong, M.; Wang, S.; Xu, F.; Wang, J.; Yang, N.; Li, Q.; Chen, J.; Li, W. Pretreatment of sweet sorghum straw and its enzymatic digestion: Insight into the structural changes and visualization of hydrolysis process. Biotechnol. Biofuels. 2019, 12, 276. [Google Scholar] [CrossRef][Green Version]
  37. Cardoso, W.S.; Tardin, F.D.; Tavares, G.; Queiroz, P.V.; Mota, S.S.; Kasuya, M.C.M.; de Queiroz, J.H. Use of sorghum straw (Sorghum bicolor) for second generation ethanol production: Pretreatment and enzymatic hydrolysis. Quim. Nova 2013, 36, 623–627. [Google Scholar] [CrossRef][Green Version]
  38. Dos Santos, R.M.; Neto, W.P.F.; Silverio, H.A.; Martins, D.F. Cellulose nanocrystals from pineapple leaf, a new approach for the reuse of this agro-waste. Ind. Crops Prod. 2013, 50, 707–714. [Google Scholar] [CrossRef]
  39. Choonut, A.; Saejong, M.; Sangkharak, K. The Production of ethanol and hydrogen from pineapple peel by Saccharomyces cerevisiae and Enterobacter aerogenes. Energy Procedia 2014, 52, 242–249. [Google Scholar] [CrossRef][Green Version]
  40. Taher, I.B.; Fickers, P.; Chnitit, S.; Hassouna, M. Optimization of enzymatic hydrolysis and fermentation conditions for improved bioethanol production from potato peel residues. Biotechnol. Prog. 2017, 33, 397–406. [Google Scholar] [CrossRef]
  41. Rivas, B.; Torrado, A.; Torre, P.; Converti, A.; Domínguez, J.M. Submerged citric acid fermentation on orange peel autohydrolysate. J. Agric. Food. Chem. 2008, 56, 2380–2387. [Google Scholar] [CrossRef]
  42. Ververis, C.; Georghiou, K.; Danielidis, D.; Hatzinikolaou, D.G.; Santas, P.; Santas, R.; Corleti, V. Cellulose, hemicelluloses, lignin and ash content of some organic materials and their suitability for use as paper pulp supplements. Bioresour. Technol. 2007, 98, 296–301. [Google Scholar] [CrossRef] [PubMed]
  43. Szymánska-Chargot, M.; Chylińska, M.; Gdula, K.; Koziol, A.; Zdunek, A. Isolation and characterization of cellulose from different fruit and vegetable pomaces. Polymers 2017, 9, 495. [Google Scholar] [CrossRef] [PubMed]
  44. Motte, J.C.; Trably, E.; Escudié, R.; Hamelin, J.; Steyer, J.P.; Bernet, N.; Delgenes, J.P.; Dumas, C. Total solids content: A key parameter of metabolic pathways in dry anaerobic digestion. Biotechnol. Biofuels 2013, 6, 164. [Google Scholar] [CrossRef][Green Version]
  45. Zhao, X.; Peng, F.; Cheng, K.; Liu, D. Enhancement of enzymatic digestibility of sugarcane bagasse by alkali-peracetic acid pretreatment. Enzyme Microbial. Technol. 2009, 44, 17–23. [Google Scholar] [CrossRef]
  46. Moutta, R.O.; Chandel, A.K.; Rodrigues, R.C.L.B.; Silva, M.B.; Rocha, G.J.M.; da Silva, S.S. Statistical optimization of sugarcane leaves hydrolysis into simple sugars by dilute sulfuric acid catalyzed process. Sugar Technol. 2012, 13, 53–60. [Google Scholar] [CrossRef]
  47. Saad, M.B.W.; Oliveira, L.R.M.; Candido, R.G.; Quintana, G.; Rocha, G.J.M.; Goncalves, A.R. Preliminary studies on fungal treatment of sugarcane straw for organosolv pulping. Enzyme Microbial. Technol. 2008, 45, 220–225. [Google Scholar] [CrossRef]
  48. Ariffin, H.; Hassan, M.A.; Umi Kalsom, M.S.; Abdullah, N.; Ghazali, F.M.; Shirai, Y. Production of bacterial endoglucanase from oil palm empty fruit bunch by Bacillus pumilus EB3. J. Biosci. Bioeng. 2008, 3, 231–2236. [Google Scholar] [CrossRef] [PubMed]
  49. Zainudin, M.H.M.; Rahman, N.A.; Abd-Aziz, S.; Funaoka, M.; Shinano, T.; Shirai, Y. Utilization of glucose recovered by phase separation system from acid-hydrolysed oil palm empty fruit bunch for bioethanol production. Sci. Pertanika J. Trop. Agric. 2012, 35, 117–126. [Google Scholar]
  50. Tufail, T.; Saeed, F.; Imran, M.; Arshammad, M.U.; Anjum, F.M.; Afzaal, M.; Ain, H.B.U.; Shahbaz, M.; Gondal, T.A.; Hussain, S. Biochemical characterization of wheat straw cell wall with special reference to bioactive profile. Int. J. Food Prop. 2018, 21, 1303–1310. [Google Scholar] [CrossRef][Green Version]
  51. Li, X.; Liu, Y.; Hao, J.; Wang, W. Study of almond shell characteristics. Materials 2018, 11, 1782. [Google Scholar] [CrossRef][Green Version]
  52. Xinyuan, J.; Yuanyuan, L.; Zhong, G.; An, M.; Zecai, H.; Suwen, Y. Pyrolysis characteristics and correlation analysis with the major components of seven kinds of nutshell. Sci. Silvae Sin. 2015, 51, 79–86. [Google Scholar]
  53. Akgül, M.; Korkut, S.; Camlibel, O.; Ayata, Ü. Some chemical properties of Luffa and its suitability for medium density fiberboard (MDF) production. Bioresurse 2013, 8, 1709–1717. [Google Scholar] [CrossRef]
  54. Rodríguez, G.; Lama, A.; Rodríguez, R.; Jiménez, A.; Guillén, R.; Fernandez-Bolanos, J. Olive stone an attractive source of bioactive and valuable compounds. Bioresour. Technol. 2008, 99, 5261–5269. [Google Scholar] [CrossRef] [PubMed]
  55. Ndika, E.V.; Chidozie, U.S.; Ikechukwu, U.K. Chemical modification of cellulose from palm kernel de-oiled cake to microcrystalline cellulose and its evaluation as a pharmaceutical excipient. Afr. J. Pure Appl. Chem. 2019, 13, 49–57. [Google Scholar]
  56. FAOSTAT. Food and Agriculture Data. Available online: (accessed on 20 May 2020).
  57. Kalač, P. A review of chemical composition and nutritional value of wild-growing and cultivated mushrooms. J. Sci. Food Agric. 2013, 93, 209–218. [Google Scholar] [CrossRef]
  58. Valverde, M.E.; Hernándea-Pérez, T.; Paredes-López, O. Edible mushrooms: Improving human health and promoting quality life. Int. J. Microbial. 2015, 376–387. [Google Scholar] [CrossRef]
  59. Cheung, P.C.K. The nutritional and health benefits of mushrooms. Nutr. Bull. 2010, 35, 292–299. [Google Scholar] [CrossRef]
  60. Ma, G.; Yang, W.; Zhao, L.; Pei, F.; Fang, D.; Hu, Q. A critical review on the health promoting effects of mushrooms nutraceuticals. Food Sci. Hum. Wellness 2018, 7, 125–133. [Google Scholar] [CrossRef]
  61. Royse, D.J.; Baars, J.; Tan, Q. Current overview of mushroom production in the world. In Edible and Medicinal Mushrooms: Technology and Applications; Zied, D.C., Pardo-Gimenez, A., Eds.; Wiley-Blackwell: West Sussex, UK, 2007; pp. 5–13. [Google Scholar]
  62. Hoa, H.T.; Wang, C.; Wang, C. The effects of different substrates on the growth, yield, and nutritional composition of two oyster mushrooms (Pleurotus ostreatus and Pleurotus cystidiosus). Mycobiology 2015, 43, 423–434. [Google Scholar] [CrossRef][Green Version]
  63. Cueva, M.B.R.; Hernáadez, A.; Niňo-Ruiz, Z. Influence of C/N ratio on productivity and the protein contents of Pleurotus ostreatus grown in differents residue mixtures. Rev. FCA Uncuyo. 2017, 49, 331–334. [Google Scholar]
  64. Ragunathan, R.; Swaminathan, K. Nutritional status of Pleurotus spp. grown on various agro-wastes. Food Chem. 2003, 80, 371–375. [Google Scholar] [CrossRef]
  65. Wang, D.; Sakoda, A.; Suzuki, M. Biological efficiency and nutritional value of Pleurotus ostreatus cultivated on spent beer grain. Bioresour. Technol. 2001, 78, 293–300. [Google Scholar] [CrossRef]
  66. Carrasco, J.; Zied, D.C.; Pardo, J.E.; Preston, G.M.; Pardo-Gimenez, A. Supplementation in mushroom crops and its impact on yield and quality. AMB Expr. 2018, 8, 146. [Google Scholar] [CrossRef] [PubMed][Green Version]
  67. Moonmoon, M.; Shelly, N.J.; Khan, M.A.; Uddin, M.N.; Hossain, K.; Tania, M.; Ahmed, S. Effects of different levels of wheat bran, rice bran and maize powder supplementation with saw dust on the production of shiitake mushroom (Lentinus edodes (Berk.) Singer). Saudi J. Biol. Sci. 2011, 18, 323–328. [Google Scholar] [CrossRef] [PubMed][Green Version]
  68. Philippoussis, A. Production of mushrooms using agro-industrial residues as substrates. In Biotechnology for Agro-Industrial Residues Processing; Nigam, P.S., Pandey, A., Eds.; Springer: Dordrecht, The Netherlands, 2009; pp. 163–196. [Google Scholar]
  69. Shekhar, H.S.; Kilpatrick, M. Mushroom (Agaricus bisporus) compost quality factor for predicting potential yield of fruiting bodies. Can. J. Microbiol. 2000, 46, 515–519. [Google Scholar]
  70. Oei, P. Mushroom Cultivation, 3rd ed.; Backhuys Publishers: Leiden, The Netherlands, 2003; 429. [Google Scholar]
  71. Lisiecka, J.; Sobieralski, K.; Siwulski, M.; Jasinska, A. Almond mushroom Agaricus brasiliensis (Wasser et al.)–properties and culture condition. Acta Sci. Pol. Hortorum Cultus 2013, 12, 27–40. [Google Scholar]
  72. Cies, L. Resultados de dos ciclos de cultivo de champiñón Portobello. El Champiñón en Castilla-La Mancha 2009, 29, 1. [Google Scholar]
  73. Kopytowski, F.J.; Minhoni, M.T.A. C/N ratio on yield of Agaricus blazei Murrill ss. Heinemann. Mushroom Sci. 2004, 16, 213–220. [Google Scholar]
  74. Zied, D.C.; Savoie, J.; Pardo-Giménez, A. Soybean the main nitrogen source in cultivation substrates of edible and medicinal mushrooms. In Soybean and Nutrition; El-Shemy, H., Ed.; Janeza Trdine: Rijeka, Croatia, 2011; pp. 434–452. [Google Scholar]
  75. Poppe, J.; Höfte, M. Twenty wastes for twenty cultivated mushroom. Mushroom Sci. 1995, 14, 171–179. [Google Scholar]
  76. Chang, S.T.; Milles, P.G. Edible Mushroom and Their Cultivation; CRC Press: Florida, FL, USA, 1989; p. 345. [Google Scholar]
  77. Chang-Ho, Y.; Ho, T.M. Effect of nitrogen amendment on the growth of Volvariella volvacea. Mushroom Sci. 1979, 10, 619–625. [Google Scholar]
  78. Kaul, T.; Khurana, M.; Kachroo, J. Chemical composition of cereal straw of the Kashmir valley. Mushroom Sci. 1981, 11, 19–25. [Google Scholar]
  79. Heltay, I.; Zavodi, I. Rice straw compost. Mushroom Sci. 1960, 4, 393–399. [Google Scholar]
  80. Shi, L.; Chen, D.; Xu, C.; Ren, A.; Yu, H.; Zhao, M. Highly-efficient liposome-mediated transformation system for the basidiomycetous fungus Flammulina velutipes. J. Gen. Appl. Microbiol. 2017, 63, 179–185. [Google Scholar] [CrossRef] [PubMed][Green Version]
  81. Hsieh, C.; Yang, F.C. Reusing soy residue for the solid-state fermentation of Ganoderma lucidum. Bioresur. Technol. 2004, 91, 105–109. [Google Scholar] [CrossRef]
  82. Wakchaure, G.C. Production and marketing of mushrooms: Global and national scenario. In Mushrooms Cultivation, Marketing and Consumption; Singh, M., Vijay, B., Kamal, S., Wakchaure, G.C., Eds.; ICAR Publishing: Solan, India, 2011; pp. 15–22. [Google Scholar]
  83. Girmay, Z.; Gorems, W.; Birhanu, G.; Zewdie, S. Growth and yield performance of Pleurotus ostreatus (Jacq. Fr.) Kumm (oyster mushroom) on different substrates. AMB Expr. 2016, 6, 87. [Google Scholar] [CrossRef][Green Version]
  84. Toker, H.; Baysal, E.; Yigibasi, O.N.; Colak, M.; Perker, H.; Simsek, H.; Yilmaz, F. Cultivation of Agaricus bisporus on wheat straw and waste tea leaves based composts using poplar leaves as activator material. Afr. J. Biotechnol. 2007, 6, 204–212. [Google Scholar]
  85. Tsai, S.Y.; Wu, T.P.; Huang, S.J.; Mau, J.L. Nonvolatile taste components of Agaricus bisporus harvested at different stages of maturity. Food Chem. 2007, 103, 1457–1464. [Google Scholar] [CrossRef]
  86. Pardo-Giménez, A.; Pardo, J.E.; Dias, E.S.; Rinker, D.L.; Caitano, C.E.C.; Zied, D.C. Optimization of cultivation techniques improves the agronomic behavior of Agaricus subrufescens. Sci. Rep. 2020, 10, 8154. [Google Scholar] [CrossRef] [PubMed]
  87. Koutrotsios, G.; Mountzouris, K.C.; Chatzipavlidis, I.; Zervakis, G.I. Bioconversion of lignocellulosic residues by Agrocybe cylindracea and Pleurotus ostreatus mushroom fungi–assessment of their effect on the final product and spent substrate properties. Food Chem. 2014, 161, 127–135. [Google Scholar] [CrossRef] [PubMed]
  88. Hassan, F.R.H. Cultivation of the monkey head mushroom (Hericium erinaceus) in Egypt. J. App. Sci. Res. 2007, 3, 1229–1233. [Google Scholar]
  89. Gaitán-Hernández, R.; Cortés, N.; Mata, G. Improvement of yield of the edible and medicinal mushroom Lentinula edodes on wheat straw by use of supplemented spawn. Braz. J. Microbiol. 2014, 45, 467–474. [Google Scholar] [CrossRef][Green Version]
  90. Patil, S.S. Productivity and proximate content of Pleurotus sajor-caju. Biosci. Discov. 2013, 4, 169–172. [Google Scholar]
  91. Medany, G.M. Cultivation possibility of golden oyster mushroom (Pleurotus citrinopileatus) under the Egyptian conditions. Egypt. J. Agric. Res. 2014, 92, 749–761. [Google Scholar]
  92. Ana, S.; Aak, A.; Aa, H.; Ea, S. Effect of residues agricultural wastes on the productivity and quality of Pleurotus colombinus l. by using polyethylene bags wall technique. Adv. Plants Agric. Res. 2016, 5, 528–536. [Google Scholar]
  93. Telang, S.M.; Patil, S.S.; Baig, M.M.V. Comparative study on yield and nutritional aspect of Pleurotus eous mushroom cultivated on different substrate. Food Sci. Res. J. 2010, 1, 60–63. [Google Scholar]
  94. Sardar, H.; Ali, M.A.; Anjum, M.A.; Nawaz, F.; Hussain, S.; Naz, S.; Karimi, S.M. Agro-industrial residues influence mineral elements accumulation and nutritional composition of king oyster mushroom (Pleurotus eryngii). Sci. Hort. 2017, 225, 327–334. [Google Scholar] [CrossRef]
  95. Prasad, S.; Rathore, H.; Sharma, S.; Tiwari, G. Yield and proximate composition of Pleurotus florida cultivated on wheat straw supplemented with perennial grasses. Indian J. Agric. Sci. 2018, 88, 91–94. [Google Scholar]
  96. Nasreen, Z.; Ali, S.; Usman, S.; Nazir, S.; Yasmeen, A. Comparative study on the growth and yield of Pleurotus ostreatus mushroom on lignocellulosic by-products. Int. J. Adv. Res. Bot. 2016, 2, 42–49. [Google Scholar]
  97. Telang, S.M.; Patil, S.S.; Baig, M.M.V. Comparative study on yield and nutritional aspect of Pleurotus sapidus mushroom cultivated on different substrate. Food Sci. Res. J. 2010, 1, 127–129. [Google Scholar]
  98. De Andrade, M.C.N.; Zied, D.C.; Minhoni, M.T.A.; Filho, J.K. Yield of four Agaricus bisporus strains in three compost formulations and chemical composition analyses of the mushrooms. Braz. J. Microbial. 2008, 39, 593–598. [Google Scholar] [CrossRef][Green Version]
  99. De Carvalho, C.S.M.; Sales-Campos, C.; de Carvalho, L.P.; Minhoni, M.T.A.; Saad, A.L.M.; Alquati, G.P.; de Andrade, M.C.N. Cultivation and bromatological analysis of medicinal mushroom Ganoderma lucidum (Curt.: Fr.) P. Karst cultivated in agricultural waste. Afr. J. Biotechnol. 2015, 14, 412–418. [Google Scholar]
  100. Gao, S.; Huang, Z.; Feng, X.; Bian, Y.; Huang, W.; Lui, Y. Bioconversion of rice straw agroresidues by Lentinula edodes and evaluation of non-volatile taste compounds in mushrooms. Sci. Rep. 2020, 10, 1814. [Google Scholar] [CrossRef] [PubMed]
  101. Adenipekun, C.O.; Omolaso, P.O. Comparative study on cultivation, yield performance and proximate composition of Pleurotus pulmonarius Fries. (Quelet) on rice straw and banana leaves. World J. Agric. Sci. 2015, 11, 151–158. [Google Scholar]
  102. Emriru, B.; Zenebech, K.; Kebede, F. Effect of substrates on the yield, yield attribute and dietary values of oyster mushroom (Pleurotus ostreatus) in the pastoral regions of northern Ethiopia. Afr. J. Food Agric. Nutr. Dev. 2016, 16, 11199–11218. [Google Scholar]
  103. Ashraf, J.; Ali, M.A.; Ahmad, M.; Ayyub, C.M.; Shafi, J. Effect of different substrate supplements on oyster mushroom (Pleurotus spp.) production. Food Sci. Technol. 2013, 1, 44–51. [Google Scholar]
  104. Biswas, M.K.; Layak, M. Techniques for increasing the biological efficiency of paddy straw mushroom (Volvariella volvacea) in eastern India. Food Sci. Technol. 2014, 2, 52–57. [Google Scholar]
  105. Ahlawat, O.P.; Ahlawat, K.; Dhar, B.L. Influence of lignocellulolytic enzymes on substrate colonization and yield in monosporous isolates and parent strains of Volvariella volvacea (Bull. Fr.). Sing. India J. Microbiol. 2005, 45, 205–210. [Google Scholar]
  106. Reyes, R.G.; Lopez, L.L.M.A.; Kumakura, K.; Kalaw, S.P.; Kikukawa, T.; Eguchi, F. Coprinus comatus, a newly domesticated wild nutriceutical mushroom in the Philippines. J. Argic. Tecnhol. 2009, 5, 299–316. [Google Scholar]
  107. Stojkovic, D.; Reis, F.S.; Barros, L.; Glamočlija, J.; Ćirić, A.; van Griensven, L.J.I.; Sokovic, M.; Ferreira, I.C.F.R. Nutrients and non-nutrients composition and bioactivity of wild and cultivated Coprinus comatus (O.F.Müll.). Pers. Food Chem. Toxicol. 2013, 59, 289–296. [Google Scholar] [CrossRef]
  108. Salami, A.O.; Bankole, F.A.; Salako, Y.A. Nutrient and mineral content of oyster mushroom (Pleurotus florida) grown on selected lignocellulosic agro-waste substrates. Virol. Mycol. 2016, 5, 2. [Google Scholar]
  109. Adedokun, O.M.; Akuma, A.H. Maximizing agricultural residues: Nutritional properties of straw mushroom on maize husk, wastes cotton and plantain leaves. Nat. Res. 2013, 4, 534–537. [Google Scholar] [CrossRef][Green Version]
  110. Ahmad, W.; Iqdal, J.; Salim, M.; Ahmad, I.; Sarwar, M.A.; Shehzad, M.A.; Rafiq, M.A. Performance of oyster mushroom (Pleurotus ostreatus) on cotton waste amended with maize and banana leaves. Pak. J. Nutr. 2011, 10, 509–513. [Google Scholar] [CrossRef][Green Version]
  111. Garuba, T.; Abdukkareem, K.A.; Ibrahim, I.A.; Oyebamiji, O.I.; Shoyooye, O.A.; Ajibade, T.D. Influence of substrates on the nutritional quality of Pleurotus pulmonarius and Pleurotus ostreatus. Ceylon J. Sci. 2017, 46, 67–74. [Google Scholar] [CrossRef][Green Version]
  112. Haq, I.U.; Khan, M.A.; Khan, S.A.; Ahmad, M. Biochemical analysis of fruiting bodies of Volvariella volvacea strain Vv pk, grown on six different substrates. Soil Environ. 2011, 30, 146–150. [Google Scholar]
  113. Ahmed, S.A.; Kadam, J.A.; Mane, V.P.; Patil, S.S.; Baig, M.M.V. Biological efficiency and nutritional contents of Pleurotus florida (Mont.) Singer cultivated on different agro-wastes. Nat. Sci. 2009, 7, 44–48. [Google Scholar]
  114. Herawati, E.; Arung, E.T.; Amirta, R. Domestication and nutrient analysis of Schizopyllum commune, alternative natural food sources in East Kalimantan. Agric. Agric. Sci. Procedia 2016, 9, 291–296. [Google Scholar] [CrossRef][Green Version]
  115. Triyono, S.; Haryanto, A.; Telaumbanua, M.; Lumbanraja, D.J.; To, F. Cultivation of straw mushroom (Volvariella volvacea) on oil palm empty fruit bunch growth medium. Int. J. Recycl. Org. Waste Agric. 2019, 8, 381–392. [Google Scholar] [CrossRef][Green Version]
  116. Familoni, T.V.; Ogidi, C.O.; Akinyele, B.J.; Onifade, A.K. Evaluation of yield, biological efficiency and proximate composition of Pleurotus species cultivated on different wood dusts. Czech Mycol. 2018, 70, 33–45. [Google Scholar] [CrossRef][Green Version]
  117. Salmones, D.; Mata, G.; Ramos, L.M.; Waliszewski, K.N. Cultivation of shiitake mushroom, Lentinula edodes, in several lignocellulosic materials originating from the subtropics. Agron. EDP Sci. 1999, 19, 13–19. [Google Scholar] [CrossRef]
  118. Selvakumar, P.; Rajasekar, S.; Babu, A.G.; Periasamy, K.; Raaman, N.; Reddy, M.S. Improving biological efficiency of Pleurotus strain through protoplast fusion between P. ostreatus var. florida and P. djamor var. roseus. Food Sci. Biotechnol. 2015, 24, 1741–1748. [Google Scholar]
  119. Iqbal, B.; Khan, H.; Saifullah, L.; Khan, I.; Shah, B.; Naeem, A.; Ullah, W.; Khan, N.; Adnan, M.; Shah, S.R.A.; et al. Substrates evaluation for the quality, production and growth of oyster mushroom (Pleurotus florida Cetto). J. Entomol. Zool. Stud. 2016, 4, 98–107. [Google Scholar]
  120. Sardar, A.; Satankar, V.; Jagajanantha, P.; Mageshwaran, V. Effect of substrates (cotton stalks and cotton seed hulls) on growth, yield and nutritional composition of two oyster mushrooms (Pleurotus ostreatus and Pleurotus florida). J. Cotton Res. Dev. 2020, 34, 135–145. [Google Scholar]
  121. Kortei, N.K.; Dzogbefia, V.P.; Obodai, M. Assessing the effect of composting cassava peel based substrates on the yield, nutritional quality, and physical characteristics of Pleurotus ostreatus (Jacq. ex Fr.) Kummer. Biotechnol. Res. Int. 2014, 571520, 1–9. [Google Scholar] [CrossRef] [PubMed][Green Version]
  122. Apetorgbor, M.M.; Apetorgbor, A.K. Comparative studies on yield of Volvariella volvacea using root and tuber peels for improved livelihood of communities. J. Ghana Sci. Assoc. 2015, 16, 35–43. [Google Scholar]
  123. Koutrotsios, G.; Patsou, M.; Mitsou, E.K.; Bekiaris, G.; Kotsou, M.; Tarantilis, P.A. Valorization of olive by-products as substrates for the cultivation of Ganoderma lucidum and Pleurotus ostreatus mushrooms with enhanced functional and prebiotic properties. Catalysts 2019, 9, 537. [Google Scholar] [CrossRef][Green Version]
  124. Liang, C.H.; Wu, C.Y.; Lu, P.L.; Kuo, Y.C.; Liang, Z.C. Biological efficiency and nutritional value of the culinary-medicinal mushroom Auricularia cultivated on a sawdust basal substrate supplement with different proportions of grass plants. Saudi J. Biol. Sci. 2019, 26, 263–269. [Google Scholar] [CrossRef][Green Version]
  125. Pati, S.S.; Ahmed, S.A.; Telang, S.M.; Baig, M.M.V. The nutritional value of Pleurotus ostreatus (Jacq.; Fr.) Kumm cultivated on different lignocellulosis agro-wastes. Innov. Rom. Food Biotechnol. 2010, 7, 66–76. [Google Scholar]
  126. De Siqueira, F.G.; Martos, E.T.; da Silva, G.; da Silva, R.; Dias, E.S. Biological efficiency of Agaricus brasiliensis cultivated in compost with nitrogen concentrations. Hortic. Bras. 2011, 29, 157–161. [Google Scholar] [CrossRef][Green Version]
  127. Harith, N.; Abdullah, N.; Sabaratnam, V. Cultivation of Flammulina velutipes mushroom using various agro-residues as a fruiting substrate. Pesq. Agropec. Bras. 2014, 49, 181–188. [Google Scholar] [CrossRef][Green Version]
  128. Wiafe-Kwagyan, M.; Obadai, M.; Odamtten, G.T.; Kortei, N.K. The potential use of rice waste lignocellulose and its amendments as substrate for the cultivation of Pleurotus eous strain P-3 in Ghana. Int. J. Adv. Phar. Biol. Chem. 2016, 5, 116–130. [Google Scholar]
  129. Bernardi, E.; Volcão, L.M.; Melo, L.G.; Nascimento, J.S. Productivity, biological efficiency and bromatological composition of Pleurotus sajor-caju growth on different substrates in Brazil. Agric. Nat. Resour. 2019, 53, 99–105. [Google Scholar]
  130. Cragg, S.M.; Beckham, G.T.; Bruce, N.C.; Bugg, T.D.H.; Daniel, D.L.; Dupree, P.; Etxabe, A.G.; Goodell, B.S.; Jellison, J.; McGeehan, J.E.; et al. Lignocellulose degration mechanisms across the tree of life. Curr. Opin. Chem. Biol. 2015, 29, 108–119. [Google Scholar] [CrossRef] [PubMed][Green Version]
  131. Bredon, M.; Dittmer, J.; Noël, C.; Moumen, B.; Bouchon, D. Lignocellulose degradation at the holobiont level: Teamwork in a keystone soil invertebrate. Microbiome 2018, 6, 162. [Google Scholar] [CrossRef] [PubMed][Green Version]
  132. Eichorst, S.A.; Kuske, C.R. Identification of cellulose-responsive bacterial and fungal communities in geographically and edaphically different soils by using stable isotope probing. Appl. Environ. Microbiol. 2012, 78, 2316–2327. [Google Scholar] [CrossRef] [PubMed][Green Version]
  133. Andlar, M.; Rezić, T.; Marđetko, N.; Kracher, D.; Ludwig, R.; Santek, B. Lignocellulose degradation: An overview of fungi and fungal enzymes involved in lignocellulose degradation. Eng. Life Sci. 2018, 18, 768–778. [Google Scholar] [CrossRef]
  134. Lombard, V.; Golaconda, R.H.; Drula, E.; Coutinho, P.M.; Henrissat, B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 2014, 42, 490–495. [Google Scholar] [CrossRef] [PubMed][Green Version]
  135. López-Mondéjar, R.; Zühlke, D.; Becher, D.; Riedel, K.; Baldrian, P. Cellulose and hemicellulose decomposition by forest soil bacteria proceeds by the action of structurally variable enzymatic systems. Sci. Rep. 2016, 6, 25279. [Google Scholar] [CrossRef] [PubMed]
  136. Madeira, J.V., Jr.; Contesini, F.J.; Calzado, F.; Rubio, M.V.; Zubieta, M.P.; Lopes, D.B.; de Melo, R.R. Agro-industrial residues and microbial enzymes: An overview on the eco-friendly bioconversion into high value-added products. In Biotechnology of Microbial Enzymes; Brahmachari, G., Ed.; Elsevier: Amsterdam, The Netherlands, 2017; pp. 475–511. [Google Scholar]
  137. Ritota, M.; Manzi, P. Pleurotus spp. cultivation on different agri-food by-products: Example of biotechnological application. Sustainability 2019, 11, 5049. [Google Scholar] [CrossRef][Green Version]
  138. Horn, S.J.; Vaaje-Kolstad, G.; Westereng, B.; Eijsink, V. Novel enzymes for the degradation of cellulose. Biotechnol. Biofuels 2012, 5, 45. [Google Scholar] [CrossRef][Green Version]
  139. Sajith, S.; Priji, P.; Sreedevi, S.; Benjamin, S. An overview on fungal cellulases with an industrial perspective. J. Nutr. Food. Sci. 2016, 6, 461. [Google Scholar]
  140. Yeoman, C.J.; Han, Y.; Dodd, D.; Schroeder, C.M.; Mackie, R.I.; Cann, I.K. Thermostable enzymes as biocatalysts in the biofuel industry. Adv. Appl. Microbiol. 2010, 70, 1–55. [Google Scholar]
  141. Zhang, Y.H.P.; Himmel, M.E.; Mielenz, J.R. Outlook for cellulase improvement, screening and selection strategies. Biotechnol. Adv. 2006, 24, 452–481. [Google Scholar] [CrossRef] [PubMed]
  142. Saini, K.J.; Saini, R.; Lakshmi Tewari, L. 2015. Lignocellulosic agriculture wastes as biomass feedstocks for second-generation bioethanol production: Concepts and recent developments. 3 Biotech 2015, 5, 337–353. [Google Scholar] [CrossRef][Green Version]
  143. Qi, M.; Jun, H.S.; Forsberg, C.W. Cel9D, an atypical 1,4-β-d-glucan glucohydrolase from Fibrobacter succinogenes: Characteristics, catalytic residues and synergistic interactions with other cellulases. J. Bacteriol. 2008, 190, 1976–1984. [Google Scholar] [CrossRef] [PubMed][Green Version]
  144. Pothiraj, C.; Balaji, P.; Eyini, M. Enhanced production of cellulases by various fungal cultures in solid state fermentation of cassava waste. Afr. J. Biotechnol. 2006, 5, 1882–1885. [Google Scholar]
  145. Bansal, N.; Tewari, R.; Soni, R.; Soni, S.K. Production of cellulases from Aspergillus niger NS-2 in solid state fermentation on agricultural and kitchen waste residues. Waste Manag. 2012, 32, 1341–1346. [Google Scholar] [CrossRef] [PubMed]
  146. Prasanna, H.N.; Ramanjaneyulu, G.; Rajasekhar Reddy, B. Optimization of cellulase production by Penicillium sp. 3 Biotech 2016, 6, 1–11. [Google Scholar] [CrossRef] [PubMed][Green Version]
  147. Ellilä, S.; Fonseca, L.; Uchima, C.; Cota, J.; Goldman, G.H.; Saloheimo, M.; Sacon, V.; Siika-aho, M. Development of a low-cost cellulase production process using Trichoderma reesei for Brazilian biorefineries. Biotechnol. Biofuels 2017, 10, 1–17. [Google Scholar] [CrossRef]
  148. Reddy, G.V.; Babu, P.R.; Komaraiah, P.; Roy, K.R.R.M.; Kothari, I.L. Utilization of banana waste for the production of lignolytic and cellulolytic enzymes by solid substrate fermentation using two Pleurotus species (P. ostreatus and P. sajor-caju). Process. Biochem. 2003, 38, 1457–1462. [Google Scholar] [CrossRef]
  149. Balaraju, K.; Park, K.; Jahagirdar, S.; Kaviyarasan, V. Production of cellulase and laccase enzymes by Oudemansiella radicata using agro wastes under solid-state and submerged conditions. Res. Biotechnol. 2010, 1, 21–28. [Google Scholar]
  150. Pandey, V.K.; Singh, M.P. Biodegradation of wheat straw by Pleurotus ostreatus. Cell. Mol. Biol. 2014, 60, 29–34. [Google Scholar]
  151. Elisashvili, V.; Chichua, D.; Kachlishvili, E.; Tsiklauri, N.; Khardziani, T. Lignocellulolytic enzyme activity during growth and fruiting of the edible and medicinal mushroom Pleurotus ostreatus (Jacq.: Fr.) Kumm. (Agaricomycetideae). Int. J. Med. Mushrooms 2003, 5, 193–198. [Google Scholar] [CrossRef]
  152. Elisashvili, V.; Penninckx, M.; Kachlishvili, E.; Tsiklauri, N.; Metreveli, E.; Kharziani, T.; Kvesitadze, G. Lentinus edodes and Pleurotus species lignocellulolytic enzymes activity in submerged and solid state fermentation of lignocellulosic wastes of different composition. Bioresour. Technol. 2008, 99, 457–462. [Google Scholar] [CrossRef] [PubMed]
  153. Montoya, S.; Orrego, C.E.; Levin, L. Growth, fruiting and lignocellulolytic enzyme production by the edible mushroom Grifola frondosa (maitake). World J. Microbiol. Biotechnol. 2012, 28, 1533–1541. [Google Scholar] [CrossRef] [PubMed]
  154. Cardoso, W.S.; Queiroz, P.V.; Tavares, G.P.; Santos, F.A.; Soares, F.E.D.F.; Kasuya, M.C.M.; Queiroz, J.H.D. Multi-enzyme complex of white rot fungi in saccharification of lignocellulosic material. Braz. J. Microbiol. 2018, 49, 879–884. [Google Scholar] [CrossRef] [PubMed]
  155. Elisashvili, V.; Kachlishvili, E.; Tsiklauri, N.; Metreveli, E.; Khardziani, T.; Agathos, S.N. Lignocellulose-degrading enzyme production by white-rot basidiomycetes isolated from the forests of Georgia. World J. Microbiol. Biotechnol. 2009, 25, 331–339. [Google Scholar] [CrossRef]
  156. Chuwech, M.; Rakariyatham, N. Potential of peanut hulls as substrates for fungal cellulase bioproduction through solid state fermentation. Asia. Pac. J. Sci. Technol. 2014, 19, 235–343. [Google Scholar]
  157. Lechner, B.E.; Papinutti, V.L. Production of lignocellulosic enzymes during growth and fruiting of the edible fungus Lentinus tigrinus on wheat straw. Process. Biochem. 2006, 41, 594–598. [Google Scholar] [CrossRef]
  158. Valášková, V.; Baldrian, P. Estimation of bound and free fractions of lignocellulose-degrading enzymes of wood-rotting fungi Pleurotus ostreatus, Trametes versicolor and Piptoporus betulinus. Res. Microbiol. 2006, 157, 119–124. [Google Scholar] [CrossRef]
  159. Wu, Y.; Shin, H. Cellulase from the fruiting bodies and mycelia of edible mushrooms: A review. J. Mushrooms 2016, 14, 127–135. [Google Scholar] [CrossRef]
  160. Deswal, D.; Khasa, Y.P.; Kuhad, R. Optimization of cellulose production by a brown rot fungus Fomitopsis sp. RCK2010 under solid state fermentation. Bioresour. Technol. 2011, 102, 6065–6072. [Google Scholar] [CrossRef]
  161. Kachlishvili, E.; Penninckx, M.J.; Tsiklauri, N.; Elisashvili, V. Effect of nitrogen source on lignocellulolytic enzyme production by white rot basidiomycetes under solid state cultivation. World J. Microbial. Biotechnol. 2005, 224, 391–397. [Google Scholar]
  162. Machuca, A.; Ferraz, A. Hydrolytic and oxidative enzymes produced by white- and brown-rot fungi during Eucalyptus grandis decay in solid medium. Enzym. Microb. Technol. 2001, 29, 386–391. [Google Scholar] [CrossRef]
  163. Pandit, N.P.; Maheshwari, S.K. Optimization of cellulase enzyme production from sugarcane pressmud using oyster mushroom-Pleurotus sajor-caju by solid state fermentation. J. Bioremed. Biodegrad. 2012, 3, 1–5. [Google Scholar] [CrossRef]
  164. Khalil, M.I.; Hoque, M.M.; Basunia, M.A.; Alam, N.; Khan, M.A. Production of cellulase by Pleurotus ostreatus and Pleurotus sajor-caju in solid state fermentation of lignocellulosic biomass. Turk. J. Agric. For. 2011, 35, 333–341. [Google Scholar]
  165. Levin, L.; Herrmann, C.; Papinutti, V.L. Optimization of lignocellulolytic enzyme production by the white-rot fungus Trametes trogii in solid-state fermentation using response surface methodology. Biochem. Eng. J. 2008, 39, 207–214. [Google Scholar] [CrossRef]
  166. Giorgio, E.M.; Fonseca, M.I.; Tejerina, M.R.; Ramos-Hryb, A.B.; Sanabria, N.; Zapata, P.D.; Villalba, L.L. Chips and sawdust substrates application for lignocellulolytic enzymes production by solid state fermentation. Int. Res. J. Microbiol. 2012, 3, 120–127. [Google Scholar]
  167. Nguyen, K.A.; Kumla, J.; Suwannarach, N.; Penkhrue, W.; Lumyong, S. Optimization of high endoglucanase yields production from polypore fungus, Microporus xanthopus strain KA038 under solid-state fermentation using green tea waste. Bio 2019, 8, bio047183. [Google Scholar] [CrossRef][Green Version]
  168. Xu, C.; Ma, F.; Zhang, X. Lignocellulose degradation and enzyme production by Irpex lacteus CD2 during solid-state fermentation of corn stover. J. Biosci. Bioeng. 2009, 108, 372–375. [Google Scholar] [CrossRef] [PubMed]
  169. Philippoussis, A.; Diamantopoulou, P. Agro-food industry wastes and agricultural residues conversion into high value products by mushroom cultivation. In Proceedings of the 7th International Conference on Mushroom Biology and Mushroom Products (ICMBMP7), Institute National de la Recherche Agronomique (INRA), Arcachon, France, 4–7 October 2011; pp. 339–351. [Google Scholar]
  170. Dashtban, M.; Maki, M.; Leung, K.T.; Mao, C.; Qin, W. Cellulase activities in biomass conversion: Measurement methods and comparison. Critical. Rev. Biotechnol. 2010, 30, 302–309. [Google Scholar] [CrossRef] [PubMed]
  171. Ghose, T.K. Measurement of cellulase activities. Pure Appl. Chem. 1987, 59, 257–268. [Google Scholar] [CrossRef]
  172. Mandels, M.; Andreotti, R.; Roche, C. Measurement of saccharifying cellulase. Biotechnol. Bioeng. Symp. 1976, 6, 21–33. [Google Scholar]
  173. Kubicek, C.P. Release of carboxymethyl-cellulase and β-glucosidase from cell walls of Trichoderma reesei. Eur. J. Appl. Biotechnol. 1981, 13, 226–231. [Google Scholar] [CrossRef]
  174. Korotkova, O.G.; Semenova, M.V.; Morozova, V.V.; Zorov, I.N.; Sokolova, L.M.; Bubnova, T.M.; Okunev, O.N.; Sinitsyn, A.P. Isolation and properties of fungal beta-glucosidases. Biochemistry 2009, 74, 569–577. [Google Scholar] [PubMed]
  175. Sørensen, A.; Lübeck, M.; Lübeck, P.S.; Ahring, B.K. Fungal beta-glucosidases: A bottleneck in industrial use of lignocellulosic materials. Biomolecules 2013, 3, 612–631. [Google Scholar] [CrossRef] [PubMed][Green Version]
  176. Scheller, H.V.; Ulvskov, P. Hemicelluloses. Annu. Rev. Plant Biol. 2010, 61, 263–289. [Google Scholar] [CrossRef] [PubMed]
  177. De Souza, W.R. Microbial Degradation of Lignocellulosic Biomass. In Sustainable Degradation of Lignocellulosic Biomass—Techniques, Applications and Commercialization; Chandel, A.K., da Silva, S.S., Eds.; IntechOpen: London, UK, 2012; pp. 207–247. [Google Scholar]
  178. Ahmed, S.; Jabeen, A.; Jamil, A. Xylanase from Trichoderma harzianum: Enzyme characterization and gene isolation. J. Chem. Soc. Pak. 2011, 29, 176. [Google Scholar]
  179. Walia, A.; Guleria, S.; Mehta, P.; Chauhan, A.; Parkash, J. Microbial xylanases and their industrial application in pulp and paper biobleaching: A review. 3 Biotech 2017, 7, 11. [Google Scholar] [CrossRef] [PubMed][Green Version]
  180. Vos, A.M.; Jurak, E.; de Gijsel, P.; Ohm, R.A.; Henrissat, B.; Lugones, L.G.; Kabel, M.A.; Wosten, H.A.B. Production of α-1,3-l-arabinofuranosidase active on substituted xylan does not improve compost degradation by Agaricus bisporus. PLoS ONE 2018, 13, e0201090. [Google Scholar] [CrossRef]
  181. Dos Santos, C.R.; de Giuseppe, P.O.; de Souza, F.H.M.; Zanphorlin, L.M.; Domingues, M.N.; Pirolla, R.A.S.; Honorato, R.V.; Tonoli, C.C.C.; de Morais, M.A.B.; Martins, V.P.M.; et al. Murakami, M.T. The mechanism by which a distinguishing arabinofuranosidase can cope with internal di-substitutions in arabinoxylans. Biotechnol. Biofuels 2018, 11, 223. [Google Scholar] [CrossRef][Green Version]
  182. Gómez, S.; Payne, A.M.; Savko, M.; Fox, G.C.; Shepard, W.E.; Fernandez, F.J.; Vega, M.C. Structural and functional characterization of a highly stable endo-β-1,4-xylanase from Fusarium oxysporum and its development as an efficient immobilized biocatalyst. Biotechnol. Biofuels 2016, 9, 191. [Google Scholar] [CrossRef][Green Version]
  183. Bajaj, P.; Mahajan, R. Cellulase and xylanase synergism in industrial biotechnology. Appl. Microbiol. Biot. 2019, 103, 8711–8724. [Google Scholar] [CrossRef]
  184. Burlacu, A.; Cornea, C.P.; Israel-Roming, F. Screening of xylanase producing microorganisms. Res. J. Agric. Sci. 2016, 48, 8–15. [Google Scholar]
  185. Meddeb-Mouelhi, F.; Moisan, J.K.; Beauregard, M. A comparison of plate assay methods for detecting extracellular cellulase and xylanase activity. Enzyme Microb. Technol. 2014, 66, 16–19. [Google Scholar] [CrossRef]
  186. Lim, S.H.; Lee, Y.H.; Kang, H.W. Efficient recovery of lignocellulolytic enzymes of spent mushroom compost from oyster Mushrooms, Pleurotus spp., and potential use in dye decolorization. Mycobiology 2013, 41, 214–220. [Google Scholar] [CrossRef] [PubMed][Green Version]
  187. Amore, A.; Amoresano, A.; Birolo, L.; Henrissat, B.; Leo, G.; Palmese, A.; Faraco, V. A family GH51 α-l-arabinofuranosidase from Pleurotus ostreatus: Identification, recombinant expression and characterization. Appl. Microbiol. Biotechnol. 2011, 94, 995–1006. [Google Scholar] [CrossRef] [PubMed]
  188. Miller, G.L. Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal. Chem. 1959, 31, 426–428. [Google Scholar] [CrossRef]
  189. Nelson, N. A photometric adaptation of Somogyi methods for determination of glucose. J. Biol. Chem. 1994, 153, 375–380. [Google Scholar]
  190. Somogyi, M. Notes on sugar determination. J. Biol. Chem. 1952, 195, 19–23. [Google Scholar]
  191. Azeri, C.; Tamer, A.U.; Oskay, M. Thermoactive cellulase-free xylanase production from alkaliphilic Bacillus strains using various agro-residues and their potential in biobleaching of kraft pulp. Afr. J. Biotechnol. 2010, 9, 63–72. [Google Scholar]
  192. Driss, D.; Bhiri, F.; Elleuch, L.; Bouly, N.; Stals, I.; Miled, N.; Blibech, M.; Ghorbel, R.; Chaabouni, S.E. Purification and properties of an extracellular acidophilic endo-1,4-β-xylanase, naturally deleted in the ‘‘thumb’’, from Penicillium occitanis Pol6. Proc. Biochem. 2012, 46, 1299–1306. [Google Scholar] [CrossRef]
  193. Hatanaka, K. Incorporation of fluorous glycosides to cell membrane and saccharide chain elongation by cellular enzymes. Top. Curr. Chem. 2012, 308, 291–306. [Google Scholar]
  194. Kuhad, R.C.; Sing, A. Lignocellulose biotechnology: Current and future prospects. Crit. Rev. Biotechnol. 1993, 13, 151–172. [Google Scholar] [CrossRef]
  195. Soni, H.; Rawat, H.K.; Pletschke, B.I.; Kango, N. Purification and characterization of β-mannanase from Aspergillus terreus and its applicability in depolymerization of mannans and saccharification of lignocellulosic biomass. Biotech 2016, 6, 136. [Google Scholar] [CrossRef][Green Version]
  196. Sherief, A.A.; El-Tanash, A.B.; Temraz, A.M. Lignocellulolytic enzymes and substrate utilization during growth and fruiting of Pleurotus ostreatus on some solid wastes. J. Environ. Sci. Technol. 2010, 3, 18–34. [Google Scholar] [CrossRef][Green Version]
  197. Manavalan, T.; Manavalan, A.; Thangavelu, K.P.; Heese, K. Secretome analysis of Ganoderma lucidum cultivated in sugarcane bagasse. J. Proteom. 2012, 77, 298–309. [Google Scholar] [CrossRef]
  198. Iandolo, D.; Piscitelli, A.; Sannia, G.; Faraco, V. Enzyme production by solid substrate fermentation of Pleurotus ostreatus and Trametes versicolor on tomato pomace. Appl. Biochem. Biotechnol. 2010, 163, 40–51. [Google Scholar] [CrossRef][Green Version]
  199. Zhu, N.; Liu, J.; Yang, J.; Lin, Y.; Yang, Y.; Ji, L.; Li, M.; Yuan, H. Comparative analysis of the secretomes of Schizophyllum commune and other wood-decay basidiomycetes during solid-state fermentation reveals its unique lignocellulose-degrading enzyme system. Biotechnol. Biofuels 2016, 9, 42. [Google Scholar] [CrossRef] [PubMed][Green Version]
  200. Steffen, K.T.; Cajthaml, T.; Snajdr, J.; Baldrian, P. Differential degradation of oak (Quercus petraea) leaf litter by litter-decomposing basidiomycetes. Res. Microbiol. 2007, 158, 447–455. [Google Scholar] [CrossRef] [PubMed]
  201. Carabajal, M.; Levin, L.; Albertó, E.; Lechner, B. Effect of co-cultivation of two Pleurotus species on lignocellulolytic enzyme production and mushroom fructification. Int. Biodeterior. 2012, 66, 71–76. [Google Scholar] [CrossRef]
  202. Heidorne, F.O.; Magalhães, P.O.; Ferraz, A.L.; Milagres, A.M.F. Characterization of hemicellulases and cellulases produced by Ceriporiopsis subvermispora grown on wood under biopulping conditions. Enzyme Microb. Technol. 2006, 38, 436–442. [Google Scholar] [CrossRef]
  203. Papinutti, V.L.; Forchiassin, F. Lignocellulolytic enzymes from Fomes sclerodermeus growing in solid-state fermentation. J. Food Eng. 2007, 81, 54–59. [Google Scholar] [CrossRef]
  204. Boonrung, S.; Mongkolthanaruk, W.; Aimi, T.; Boonlue, S. Cellulase and xylanase acting at alkaline pH from mushroom, Leucoagaricus meleagris KKU-C1. Chiang Mai J. Sci. 2014, 41, 84–96. [Google Scholar]
  205. De Oliveira Rodrigues, P.; Gurgel, L.V.A.; Pasquini, D.; Badotti, F.; Góes-Neto, A.; Baffi, M.A. Lignocellulose-degrading enzymes production by solid-state fermentation through fungal consortium among ascomycetes and basidiomycetes. Renew. Energy 2020, 145, 2683–2693. [Google Scholar] [CrossRef]
  206. Chauhan, P.S.; Puri, N.; Sharma, P.; Gupta, N. Mannanases: Microbial sources, production, properties and potential biotechnological applications. Appl. Microbiol. Biot. 2012, 93, 1817–1830. [Google Scholar] [CrossRef] [PubMed]
  207. Saeed, M.; Ayaşan, T.; Alagawany, M.; El-Hack, M.E.A.; Abdel-Latif, M.A.; Patra, A.K. The role of β-mannanase (Hemicell) in improving poultry productivity, health and environment. Braz. J. Poultry Sci. 2019, 21, 1–8. [Google Scholar] [CrossRef]
  208. Olaniyi, O.O.; Bankefa, E.O.; Folasade, I.O.; Familoni, T.V. Nutrient enrichment of mannanase-treated cassava peels and corn cob. Res. J. Microbiol. 2015, 10, 533–541. [Google Scholar] [CrossRef][Green Version]
  209. Pinho, G.P.; Matoso, J.R.M.; Silvério, F.O.; Mota, W.C.; Lopes, P.S.N.; Ribeiro, L.M. A new spectrophotometric method for determining the enzymatic activity of endo-β-mannanase in seeds. J. Braz. Chem. Soc. 2014, 25, 1246–1252. [Google Scholar] [CrossRef]
  210. Titapoka, S.; Keawsompong, S.; Haltric, D.; Nitisinprasert, S. Selection and characterization of mannanase-producing bacteria useful for the formation of prebiotic manno-ligosaccharides from copra meal. World J. Microbiol. Biotechnol. 2008, 24, 1425–1433. [Google Scholar] [CrossRef]
  211. Maijala, P.; Kango, N.; Szijarto, N.; Viikari, L. Characterization of hemicellulases from thermophilic fungi. Anton. Leeuw. 2012, 101, 905–917. [Google Scholar] [CrossRef]
  212. Wang, S.; Yang, Y.; Zhang, J.; Sun, J.; Matsukawa, S.; Xie, J.; Wei, D. Characterization of abnZ2 (yxiA1) and abnZ3 (yxiA3) in Paenibacillus polymyxa, encoding two novel endo-1,5-α-l-arabinanases. Bioresour. Bioprocess. 2014, 1, 14. [Google Scholar] [CrossRef][Green Version]
  213. Seiboth, B.; Metz, B. Fungal arabinan and l-arabinose metabolism. Appl. Microbiol. Biotechnol. 2011, 89, 1665–1673. [Google Scholar] [CrossRef] [PubMed][Green Version]
  214. Numan, M.T.; Bhosle, N.B. α-l-Arabinofuranosidases: The potential applications in biotechnology. J. Ind. Microbiol. Biotechnol. 2006, 33, 247–260. [Google Scholar] [CrossRef] [PubMed]
  215. Yanay, T.; Sato, M. Purification and characterization of a novel α-l-arabinofuranosidase from Pichia capsulata X91. Biosci. Biotechnol. Biochem. 2000, 64, 1181–1188. [Google Scholar] [CrossRef] [PubMed][Green Version]
  216. Jurak, E.; Patyshakuliyeva, A.; de Vries, R.P.; Gruppen, H.; Kabel, M.A. Compost grown Agaricus bisporus lacks the ability to degrade and consume highly substituted xylan fragments. PLoS ONE 2015, 10, e0134169. [Google Scholar] [CrossRef] [PubMed]
  217. Anderson, W.F.; Akin, D.E. Structural and chemical properties of grass lignocelluloses related to conversion for biofuels. J. Ind. Microbiol. Biotechnol. 2008, 35, 355–366. [Google Scholar] [CrossRef]
  218. Scharf, M.E.; Tartar, A. Termite digestomes as sources for novel lignocellulases. Biofuels Bioprod. Biorefin. 2008, 2, 540–552. [Google Scholar] [CrossRef]
  219. Pollegioni, L.; Tonin, F.; Rosini, E. Lignin-degrading enzymes. FEBS J. 2015, 282, 1190–1213. [Google Scholar] [CrossRef]
  220. Niladevi, K.N. Ligninolytic enzymes. In Biotechnology for Agro-Industrial Residues Utilisation; Nigam, P.S., Pandey, A., Eds.; Springer: Amsterdam, The Netherlands, 2009; pp. 397–414. [Google Scholar]
  221. Rogalski, J.; Lundell, T.; Leonowicz, A.; Hatakka, A. Production of laccase, lignin peroxidase and manganese-dependent peroxidase by various strains of Trametes versicolor depending on culture conditions. Acta Microbiol. Pol. 1991, 40, 221–234. [Google Scholar]
  222. Tripathi, A.; Upadhyay, R.C.; Singh, S. Extracellular Ligninolytic Enzymes in Bjerkandera adusta and Lentinus squarrosulus. Indian J. Microbiol. 2012, 52, 381–387. [Google Scholar] [CrossRef][Green Version]
  223. Saeki, N.; Takeda, H.; Tanesaka, E.; Yoshida, M. Induction of manganese peroxidase and laccase by Lentinula edodes under liquid culture conditions and their isozyme detection by enzymatic staining on native-PAGE. Mycoscience 2011, 52, 132–136. [Google Scholar] [CrossRef][Green Version]
  224. Caramelo, L.; Martinez, M.J.; Martinez, A.T. A search for ligninolytic peroxidases in the fungus Pleurotus eryngii involving alpha-keto-gamma-thiomethylbutyric acid and lignin model dimers. Appl. Environ. Microbiol. 1999, 65, 916–922. [Google Scholar] [CrossRef][Green Version]
  225. Chmelová, D.; Ondrejovič, M. Effect of potential inductors on laccase production by white-rot fungus Ceriporiopsis subvermispora. J. Microbiol. Biotechnol. Food Sci. 2014, 3, 84–87. [Google Scholar]
  226. Tovar-Herrera, O.E.; Martha-Paz, A.M.; Pérez-LLano, Y.; Aranda, E.; Tacoronte-Morales, J.E.; Pedroso-Cabrera, M.T.; Arévalo-Niño, K.; Folch-Mallol, J.L.; Batista-García, R.A. Schizophyllum commune: An unexploited source for lignocellulose degrading enzymes. MicrobiologyOpen 2018, 7, e00637. [Google Scholar] [CrossRef] [PubMed]
  227. Kalra, K.; Chauhan, R.; Shaves, M.; Sachdeva, S. Isolation of laccase producing Trichoderma spp. and effect of pH and temperature on its activity. Int. J. Chem. Environ. Technol. 2013, 5, 2229–2235. [Google Scholar]
  228. Zeng, S.; Zhao, J.; Xia, L. Simultaneous production of laccase and degradation of bisphenol A with Trametes versicolor cultivated on agricultural wastes. Bioprocess Biosyst. Eng. 2017, 40, 1237–1245. [Google Scholar] [CrossRef] [PubMed][Green Version]
  229. Adekunle, A.E.; Zeng, C.; Guo, C.; Liu, C. Laccase production from Trametes versicolor in solid-state fermentation of steam-exploded pretreated cornstalk. Waste Biomass. Valori. 2017, 8, 153–159. [Google Scholar] [CrossRef]
  230. Aâssi, D.; Zouari-Mechichi, H.; Frikha, F.; Rodriguez-Couto, S.; Mechichi, T. Sawdust waste as a low-cost support- substrate for laccases production and adsorbent for azo dyes decolorization. J. Environ. Health Sci. 2016, 14, 1–12. [Google Scholar]
  231. Karp, S.G.; Faraco, V.; Amore, A.; Letti, L.A.J.; Soccol, V.T.; Soccol, C.R. Statistical optimization of laccase production and delignification of sugarcane bagasse by Pleurotus ostreatus in solid-state fermentation. Biomed. Res. Int. 2015, 2015, 181–204. [Google Scholar] [CrossRef][Green Version]
  232. Moilanen, U.; Winquist, E.; Mattila, T.; Hatakka, A.; Eerikäinen, T. Production of manganese peroxidase and laccase in a solid-state bioreactor and modeling of enzyme production kinetics. Bioprocess Biosyst. Eng. 2015, 38, 57–68. [Google Scholar] [CrossRef]
  233. Hariharan, S.; Padma, N. Optimization of lignin peroxidase, manganese peroxidase, and Lac production from Ganoderma lucidum under solid state fermentation of pineapple leaf. Bioresouyces 2013, 8, 250–271. [Google Scholar] [CrossRef][Green Version]
  234. Usha, K.Y.; Praveen, K.; Rajasekhar Reddy, B. Enhanced production of ligninolytic enzymes by a mushroom Stereum ostrea. Biotechnol. Res. Int. 2014, 2014, 815495. [Google Scholar] [CrossRef]
  235. Asgher, M.; Asad, M.J.; Legge, R.L. Enhanced lignin peroxidase synthesis by Phanerochaete chrysosporium in solid state bioprocessing of a lignocellulosic substrate. World J. Microbiol. Biot. 2006, 22, 449–453. [Google Scholar] [CrossRef]
  236. Silva, E.M.; Martins, S.F.; Milagres, A.M.F. Extraction of manganese peroxidase produced by Lentinula edodes. Bioresour. Technol. 2008, 99, 2471–2475. [Google Scholar] [CrossRef] [PubMed]
  237. Robinson, T.; Nigam, P.S. Remediation of textile dye waste water using a white-rot fungus Bjerkandera adusta through solid-state fermentation (SSF). Appl. Biochem. Biotechnol. 2008, 151, 618–628. [Google Scholar] [CrossRef] [PubMed]
  238. Ferreira da Silva, I.; Rodrigues da Luz, J.M.; Oliveira, S.F.; Humberto de Queiroz, J.; Kasuya, M.C.M. High-yield cellulase and LiP production after SSF of agricultural wastes by Pleurotus ostreatus using different surfactants. Biocatal. Agric. Biotechnol. 2019, 22, 101428. [Google Scholar] [CrossRef]
  239. Mehboob, N.; Asad, M.; Imran, M.; Gulfraz, M.; Wattoo, F.H.; Hadri, S.H.; Asghar, M. Production of lignin peroxidase by Ganoderma leucidum using solid state fermentation. Afr. J. Biotechnol. 2011, 10, 9880–9887. [Google Scholar]
  240. Coconi-Linares, N.; Magaña-Ortíz, D.; Guzmán-Ortiz, D.A.; Fernández, F.; Loske, A.M.; Gómez-Lim, M.A. High-yield production of manganese peroxidase, lignin peroxidase, and versatile peroxidase in Phanerochaete chrysosporium. Appl. Microbiol. Biotechnol. 2014, 98, 9283–9294. [Google Scholar] [CrossRef]
  241. Gochev, V.K.; Krastanov, A.I. Fungal laccases. Bulg. J. Agric. Sci. 2007, 13, 75–83. [Google Scholar]
  242. Zheng, Y.; Guo, M.; Zhou, Q.; Liu, H. Effect of lignin degradation product sinapyl alcohol on laccase catalysis during lignin degradation. Ind. Crops Prod. 2019, 139, 111544. [Google Scholar] [CrossRef]
  243. Zhu, Z.; Li, N.; Li, W.; Li, J.; Li, Z.; Wang, J.; Tang, X. Laser mutagenesis of Phellinus igniarius protoplasts for the selective breeding of strains with high laccase activity. Appl. Biochem. Biotechnol. 2020, 190, 584–600. [Google Scholar] [CrossRef]
  244. Palma, C.; Lloret, L.; Sepúlveda, L.; Contreras, E. Production of versatile peroxidase from Pleurotus eryngii by solid-state fermentation using agricultural residues and evaluation of its catalytic properties. Prep. Biochem. Biotechnol. 2016, 46, 200–207. [Google Scholar] [CrossRef]
  245. Rich, J.O.; Anderson, A.M.; Berhow, M.A. Laccase-mediator catalyzed conversion of model lignin compounds. Biocat. Agric. Biotechnol. 2016, 5, 111–115. [Google Scholar] [CrossRef]
  246. Song, Q.; Deng, X.; Song, R. Expression of Pleurotus ostreatus laccase gene in Pichia pastoris and Its degradation of corn stover lignin. Microorganisms 2020, 8, 601. [Google Scholar] [CrossRef] [PubMed][Green Version]
  247. Barrios-Estrada, C.; de Jesus Rostro-Alanis, M.; Munoz-Gutierrez, B.D.; Iqbal, H.M.N.; Kannan, S.; Parra-Saldivar, R. Emergent contaminants: Endocrine disruptors and their laccase-assisted degradation - A review. Sci. Total Environ. 2018, 612, 1516–1531. [Google Scholar] [CrossRef] [PubMed]
  248. Leonowicz, A.; Cho, N.; Luterek, J.; Wilkolazka, A.; Wojtas-Wasilewska, M.; Matuszewska, A.; Hofrichter, M.; Wesenberg, D.; Rogalski, J. Fungal laccase: Properties and activity on lignin. J. Basic Microbiol. 2001, 41, 185–227. [Google Scholar] [CrossRef]
  249. Shraddha Shekher, R.; Sehgal, S.; Kamthania, M.; Kumar, A. Laccase: Microbial sources, production, purification, and potential biotechnological applications. Enzyme Res. 2011, 217861. [Google Scholar] [CrossRef][Green Version]
  250. Ardon, O.; Kerem, Z.; Hadar, Y. Enhancement of lignin degradation and laccase activity in Pleurotus ostreatus by cotton stalk extract. Can. J. Microbiol. 1998, 44, 676–680. [Google Scholar] [CrossRef]
  251. Gunjal, A.B.; Patil, N.N.; Shinde, S.S. Ligninase in Degradation of Lignocellulosic Wastes. In Enzymes in Degradation of the Lignocellulosic Wastes; Springer International Publishing: Cham, Switzerland, 2020. [Google Scholar]
  252. Zheng, F.; An, Q.; Meng, G.; Wu, X.; Dai, Y.; Si, J.; Cui, B. A novel laccase from white rot fungus Trametes orientalis: Purification, characterization, and application. Int. J. Biol. Macromol. 2017, 102, 758–770. [Google Scholar] [CrossRef]
  253. Dias, A.A.; Matos, A.J.S.; Fraga, I.; Sampaio, A.; Bezerra, R.M.F. An easy method for screening and detection of laccase activity. Open Biotechnol. J. 2017, 11, 89–93. [Google Scholar] [CrossRef]
  254. Dias, A.A.; Bezerra, R.M.; Pereira, A.N. Activity and elution profile of laccase during biological decolorization of olive mill wastewater. Bioresour. Technol. 2004, 92, 7–13. [Google Scholar] [CrossRef]
  255. Minussi, R.C.; Pastore, G.M.; Duran, N. Potential applications of laccase in the food industry. Trends Food Sci. Technol. 2002, 13, 205–216. [Google Scholar] [CrossRef]
  256. Datta, R.; Kelkar, A.; Baraniya, D.; Molaei, A.; Moulick, A.; Meena, R.S.; Formanek, P. Enzymatic Degradation of Lignin in Soil: A Review. Sustainability 2017, 9, 1163. [Google Scholar] [CrossRef][Green Version]
  257. Arora, D.S.; Gill, P.K. Comparison of two assay procedures for lignin peroxidase. Enzyme Microb. Technol. 2001, 28, 602–605. [Google Scholar] [CrossRef]
  258. Zhao, M.; Zhang, C.; Zeng, G.; Huang, D.; Xu, P.; Cheng, M. Growth, metabolism of Phanerochaete chrysosporium and route of lignin degradation in response to cadmium stress in solid-state fermentation. Chemosphere 2015, 138, 560–567. [Google Scholar] [CrossRef] [PubMed]
  259. Kong, W.; Chen, H.; Lyu, S.; Ma, F.; Yu, H.; Zhang, X. Characterization of a novel manganese peroxidase from white-rot fungus Echinodontium taxodii 2538, and its use for the degradation of lignin-related compounds. Process Biochem. 2016, 51, 1776–1783. [Google Scholar] [CrossRef]
  260. Burlacu, A.; Israel-Roming, F.; Cornea, C.P. Depolymerization of kraft lignin with laccase and peroxidase: A review. Sci. Bull. Ser. F Biotechnol. 2018, 22, 172–179. [Google Scholar]
  261. Brink, D.P.; Ravi, K.; Lidén, G.; Gorwa-Grauslund, M.F. Mapping the diversity of microbial lignin catabolism: Experiences from the eLignin database. Appl. Microbiol. Biotechnol. 2019, 103, 3979–4002. [Google Scholar] [CrossRef] [PubMed][Green Version]
  262. Herzog, V.; Fahimi, H.D. A new sensitive colorimetric assay for peroxidase using 3,3′-diaminobenzidine as hydrogen donor. Anal. Biochem. 1973, 55, 554–562. [Google Scholar] [CrossRef]
  263. De Jong, E.; Field, J.A.; de Bont, J.A. Evidence for a new extracellular peroxidase manganese-inhibited peroxidase from the white-rot fungus Bjerkandera sp. BOS 55. FEBS Lett. 1992, 299, 107–110. [Google Scholar] [CrossRef][Green Version]
  264. Rajan, A.; Kurup, J.G.; Abraham, T.E. Solid state production of manganese peroxidases using arecanut husk as substrate. Braz. Arch. Biol. Technol. 2010, 53, 555–562. [Google Scholar] [CrossRef][Green Version]
  265. Kuhar, F.; Castiglia, V.C.; Zamora, J.C. Detection of manganese peroxidase and other exoenzymes in four isolates of Geastrum (Geastrales) in pure culture. Rev. Argent. Microbiol. 2016, 48, 274–278. [Google Scholar] [CrossRef]
  266. Busse, N.; Wagner, D.; Kraume, M.; Czermak, P. Reaction kinetics of versatile peroxidase for the degradation of lignin compounds. Am. J. Biochem. Biotechnol. 2013, 9, 365–394. [Google Scholar] [CrossRef]
  267. Giardina, P.; Palmieri, G.; Fontanella, B.; Rivieccio, V.; Sannia, G. Manganese peroxidase isoenzymes produced by Pleurotus ostreatus grown on wood sawdust. Arch. Biochem. Biophys. 2000, 376, 171–179. [Google Scholar] [CrossRef] [PubMed]
  268. Ravichandran, A.; Sridhar, M. Versatile peroxidases: Super peroxidases with potential biotechnological applications-A mini review. J. Dairy Vet. Anim. Res. 2016, 4, 277–280. [Google Scholar]
  269. Chen, M.; Yao, S.; Zhang, H.; Liang, X. Purification and characterization of a versatile peroxidase from edible mushroom Pleurotus eryngii. Chin. J. Chem. Eng. 2010, 18, 824–829. [Google Scholar] [CrossRef]
  270. Fisher, A.B.; Fong, S.S. Lignin biodegradation and industrial implications. AIMS Bioeng. 2014, 1, 92–112. [Google Scholar] [CrossRef]
  271. Sugano, Y.; Muramatsu, R.; Ichiyanagi, A.; Sato, T.; Shoda, M. DyP, a unique dye-decolorizing peroxidase, represents a novel heme peroxidase family: ASP171 replaces the distal histidine of classical peroxidases. J. Biol. Chem. 2007, 282, 36652–36658. [Google Scholar] [CrossRef] [PubMed][Green Version]
  272. Chen, C.; Shrestha, R.; Jia, K.; Gao, P.F.; Geisbrecht, B.V.; Bossmann, S.H.; Shi, J.; Li, P. Characterization of dye-decolorizing peroxidase (DyP) from Thermomonospora curvata reveals unique catalytic properties of A-type DyPs. J. Biol. Chem. 2015, 290, 23447–23463. [Google Scholar] [CrossRef][Green Version]
  273. Kim, S.J.; Shoda, M. Purification and characterization of a novel peroxidase from Geotrichum candidum Dec 1 involved in decolorization of dyes. Appl. Environ. Microbiol. 1999, 65, 1029–1035. [Google Scholar] [CrossRef][Green Version]
  274. Liers, C.; Pecyna, M.J.; Kellner, H.; Worrich, A.; Zorn, H.; Steffen, K.T.; Hofrichter, M.; Ullrich, R. Substrate oxidation by dye-decolorizing peroxidases (DyPs) from wood- and litter-degrading agricomycetes compared to other fungal and heme-proteins. Appl. Microbiol. Biotechnol. 2013, 97, 5839–5849. [Google Scholar] [CrossRef]
  275. Lauber, C.; Schwarz, T.; Nguyen, Q.K.; Lorenz, P.; Lochnit, G.; Zorn, H. Identification, heterologous expression and characterization of a dye-decolorizing peroxidase of Pleurotus sapidus. AMB Express 2017, 7, 164. [Google Scholar] [CrossRef][Green Version]
  276. Qin, X.; Luo, H.; Zhang, X.; Yao, B.; Ma, F.; Su, X. Dye-decolorizing peroxidases in Irpex lacteus combining the catalytic properties of heme peroxidases and laccase play important roles in ligninolytic system. Biotechnol. Biofuels 2018, 11, 302. [Google Scholar] [CrossRef] [PubMed]
  277. Lončar, N.; Draškovic, N.; Božić, N.; Romero, E.; Simić, S.; Opsenica, I.; Vujcic, Z.; Fraaije, M.W. Expreesion and characterization of a dye-decolorizing peroxidase from Pseudomonas fluorescens Pf0-1. Catalysts 2019, 9, 463. [Google Scholar] [CrossRef][Green Version]
  278. Karmakar, M.; Ray, R.R. Current trends in research and application of microbial cellulases. Res. J. Microbiol. 2011, 6, 41–53. [Google Scholar] [CrossRef][Green Version]
  279. Zhang, L.; Xu, J.; Lei, L.; Jiang, Y.; Gao, F.; Zhou, G.H. Effects of xylanase supplementation on growth performance, nutrient digestibility and non-starch polysaccharide degradation in different Sections of the gastrointestinal tract of broilers fed wheat-based diets. Asian Aust. J. Anim. Sci. 2014, 27, 855–861. [Google Scholar] [CrossRef][Green Version]
  280. Van Zyl, W.H.; Rosea, S.H.; Trollopeb, K.; Gorgensb, J.F. Fungal β-mannanases: Mannan hydrolysis, heterologous production and biotechnological applications. Process Biochem. 2010, 45, 1203–1213. [Google Scholar] [CrossRef]
  281. Jayasekara, S.; Ratnayake, R. Microbial cellulases: An overview and applications. In Cellulose; Pascual, A.R., Martin, M.E.E., Eds.; Intechopen: London, UK, 2019; pp. 1–21. [Google Scholar]
  282. Daba, A.S.; Youssef, G.A.; Kabeil, S.S.; Hafez, E.E. Production of recombinant cellulase enzyme from Pleurotus ostreatus (Jacq.) P. Kumm. (type NRRL-0366). Afr. J. Microbiol. Res. 2011, 5, 1197–1202. [Google Scholar]
  283. Sharma, H.P.; Patel, H.; Sharma, S. Enzymatic extraction and clarification of juice from various fruits—A review. Trends Post Harvest Technol. 2014, 2, 1–14. [Google Scholar]
  284. Shi, H.; Ding, H.; Huang, Y.; Wang, L.; Zhang, Y.; Li, X.; Wang, F. Expression and characterization of a GH43 endo-arabinanase from Thermotoga thermarum. BMC Biotechnol. 2014, 14, 35. [Google Scholar] [CrossRef][Green Version]
  285. Saleem, F.; Ahmed, S.; Jamil, A. Isolation of a xylan degrading gene from genomic DNA library of a thermophilic fungus Chaetomium thermophile ATCC 28076. Pak. J. Bot. 2008, 40, 1225–1230. [Google Scholar]
  286. Khanongnuch, C.; Sanguansook, C.; Lumyong, S. Nutritive quality of β-mannanase treated copra meal in broiler diets and effectiveness on some fecal bacteria. Int. J. Poult. Sci. 2006, 5, 1087–1091. [Google Scholar]
  287. Järvinen, J.; Taskila, S.; Isomäki, R.; Ojamo, H. Screening of white-rot fungi manganese peroxidases: A comparison between the specific activities of the enzyme from different native producers. AMB Express 2012, 2, 62. [Google Scholar] [CrossRef] [PubMed][Green Version]
  288. Martinez, D.; Larrondo, L.F.; Putnam, N.; Gelpke, M.D.S.; Huang, K.; Chapman, J.; Helfenbein, K.G.; Ramaiya, P.; Detter, C.J.; Larimerm, F.; et al. Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nature 2004, 22, 695–700. [Google Scholar]
  289. Iimura, Y.; Sonoki, T.; Habe, H. Heterologous expression of Trametes versicolor laccase in Saccharomyces cerevisiae. Protein Expr. Purif. 2018, 141, 39–43. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Main composition of agro-industrial wastes.
Figure 1. Main composition of agro-industrial wastes.
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Figure 2. Data of global mushroom production during 2004–2018 from FAOSTAT [56].
Figure 2. Data of global mushroom production during 2004–2018 from FAOSTAT [56].
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Figure 3. Examples of some commercially important cultivated mushrooms.
Figure 3. Examples of some commercially important cultivated mushrooms.
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Figure 4. Scheme of the main enzymes involved in the lignocellulosic degradation process.
Figure 4. Scheme of the main enzymes involved in the lignocellulosic degradation process.
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Figure 5. Enzymes involved in cellulose degradation.
Figure 5. Enzymes involved in cellulose degradation.
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Figure 6. Enzymes involved in xylan degradation.
Figure 6. Enzymes involved in xylan degradation.
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Figure 7. Enzymes involved in mannan degradation.
Figure 7. Enzymes involved in mannan degradation.
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Figure 8. Enzymes involved in arabinan degradation.
Figure 8. Enzymes involved in arabinan degradation.
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Figure 9. Typical reaction of laccase on phenols oxidation modifled from Minussi et al. [244].
Figure 9. Typical reaction of laccase on phenols oxidation modifled from Minussi et al. [244].
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Figure 10. General reaction catalyzed by lignin peroxidase. (A) cleavage of C-C of lignin, (B) oxidation of veratryl alcohol is generally used to estimate the lignin peroxidase activity.
Figure 10. General reaction catalyzed by lignin peroxidase. (A) cleavage of C-C of lignin, (B) oxidation of veratryl alcohol is generally used to estimate the lignin peroxidase activity.
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Figure 11. Lignin depolymerisation with manganese peroxidase [259].
Figure 11. Lignin depolymerisation with manganese peroxidase [259].
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Figure 12. Scheme of the versatile peroxidase catalytic cycle [265].
Figure 12. Scheme of the versatile peroxidase catalytic cycle [265].
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Table 1. Main composition and carbon/nitrogen (C/N) ratio of some agro-industrial wastes.
Table 1. Main composition and carbon/nitrogen (C/N) ratio of some agro-industrial wastes.
Agro-Industrial WastesComposition (% Dry Weight Basis)C/N RatioReference
Apple pomace43242048/1[19]
Banana straw53291540/1[20]
Banana leaves55202538/1[21]
Barley straw23–3321–2214–1982–120/1[22,23]
Canola straw22171833–45/1[23]
Coconut husk24–433–1225–4575–186/1[24,25]
Coffee husk437940/1[26]
Corn bran343949ND[25]
Corn cob35–4535–4411–1550–123/1[27,28]
Corn stalk34–6119–247–957–80/1[25,29]
Corn straw3025850/1[25]
Cotton stalk58142270–78/1[22]
Oat bran49251812/1[25]
Oat straw25–4021–2717–1848–83/1[22,23]
Rice bran35251712–48/1[25]
Rice husk35252030–80/1[31]
Rice straw32–3923–2418–3635–72/1[29,32]
Rye straw38311982/1[22]
Beech sawdust413322100–331/1[33]
Birch sawdust403620700/1[33]
Oak sawdust25–3818–2918–25162–200/1[31,33]
Pine sawdust422528724–1070/1[33]
Poplar sawdust44322146–71/1[33]
Rubber tree sawdust382515177/1[34]
Spruce sawdust422628763–1000/1[33]
Sorghum stalk17251145/1[25]
Sorghum straw3626820–46/1[35,36]
Pineapple leaf36232749/1[37]
Pineapple peel2275377/1[38]
Potato peel355425/1[39]
Orange peel9–146–111–2102/1[40,41]
Lemon peel1252ND[41]
Tomato pomace955ND[42]
Banana peel1210318–29/1[22]
Soya stalk35252020–40/1[43]
Sugarcane bagasse30–4526–3611–2350/1[22,29,44]
Sugarcane straw36–4121–3116–2670–120/1[45,46]
Sunflower stalk42301397/1[43]
Oil palm empty fruit bunch45–5128–2912–1577/1[47,48]
Water hyacinth2134711/1[10]
Wheat bran30501519/1[25]
Wheat straw27–3821–2918–2150–80/1[22,25,49]
Walnut shell362843175/1[50]
Almond shell38293061/1[51]
Chestnut shell2116368/1[51]
Pistachio shell43251643/1[51]
Hazelnut shell55343550–58/1[52]
Olive oil cake31212614–17/1[53]
Oil palm cake64155ND[54]
Sunflower oil cake25128ND[54]
Cotton seed hull31201859–67/1[55]
“ND” = not determined.
Table 2. The carbon/nitrogen ratio in substrate to obtain the highest yield of some mushroom species.
Table 2. The carbon/nitrogen ratio in substrate to obtain the highest yield of some mushroom species.
Mushroom SpeciesC/N Ratio (%)Reference
Agaricus bisporus16/119/122/1[68]
Agaricus bitorquis16/119/122/1[69]
Agaricus brasiliensis10/126–28/150/1[70]
Agaricus brunescens16/119/121/1[71]
Agaricus subrufescens16/127/133/1[72]
Lentinula edodes25/130–35/155/1[73]
Lentinus sajor-caju40/145–55/190/1[74]
Pleurotus cornucopiae40/145–55/197/1[75]
Pleurotus eryngii40/145–55/170/1[75]
Pleurotus flabellatus40/145–60/1100/1[76]
Pleurotus florida40/145–60/1150/1[77,78]
Pleurotus ostreatus40/145–60/190/1[78]
Flammulina velutipesND30/1ND[79]
Ganoderma lucidumND70–80/1ND[80]
Volvariella volvaceaND40–60/1ND[81]
“ND” = not determined.
Table 3. Biological efficiency and chemical composition of some mushrooms grown on the non-combination of agro-industrial wastes.
Table 3. Biological efficiency and chemical composition of some mushrooms grown on the non-combination of agro-industrial wastes.
Agro-Industrial WastesMushroom SpeciesBiological Efficacy (%)Chemical Composition (% Dry Weight)Reference
Crude ProteinCarbohydrateFatFiberAsh
Wheat strawAgaricus bisporus47.2–51.121.0–27.038.0–48.03.0–4.017.0–23.38.0–11.0[83,84]
Agaricus subrufescens53.728.463.[85]
Agrocybe cylindracea61.41.589.60.340.48.6[86]
Hericium erinaceus39.4–43.526.858.93.7ND10.5[87]
Lentinula edodes66.0–93.115.2–15.463.7–65.71.1–1.5ND3.8–4.4[88]
Lentinus sajor-caju74.922.956.[89]
Pleurotus citrinopileatus98.3–105.625.364.02.7ND8.1[90]
Pleurotus columbinus69.22.925.90.425.48.5[91]
Pleurotus eous75.119.550.[92]
Pleurotus eryngii48.221.556.02.413.57.6[93]
Pleurotus florida66.427.951.22.412.28.7[94]
Pleurotus ostreatus22.6–52.611.6–14.647.5–74.41.8–2.519.1–27.18.6–12.0[86,95]
Barley strawLentinula edodes64.1–88.615.1–16.875.1–77.71.9–2.2ND5.2–5.8[88]
Pleurotus ostreatus21.312.854.729.90.91.2[95]
Oat strawAgaricus bisporus47.2–52.926.8–36.2ND2.3–3.16.6–10.39.8–11.3[97]
Ganoderma lucidum2.39.9NDNDND1.0[98]
Rice strawLentinula edodes48.716.[99]
Lentinus sajor-caju78.323.455.[89]
Hericium erinaceus33.924.160.54.2ND11.3[87]
Pleurotus citrinopileatus76.5–89.222.864.93.2ND91[90]
Pleurotus columbinus71.44.827.[91]
Pleurotus eous79.829.348.[92]
Pleurotus eryngii45.921.853.01.913.88.7[93]
Pleurotus pulmonarius23.521.1ND5.27.06.9[100]
Pleurotus ostreatus25.6–84.612.5–23.455.3–57.42.8–16.27.7–0.76.3–13.6[95,101]
Pleurotus djamor82.724.837.[102]
Volvariella volvacea10.2–15.036.9–38.142.8–42.30.8–1.04.4–6.09.0–10.3[103,104,105]
Corprinus comatus18.010.976.61.9ND20.5[106]
Corn strawPleurotus florida31.626.331.30.519.65.2[107]
Volvariella volvaceaND23.013.91.436.611.9[108]
Corn cobAgrocybe cylindracea33.514.872.42.917.010.1[86]
Pleurotus columbinus79.11.928.[91]
Pleurotus cystidiosus50.124.540.[109]
Pleurotus eryngii51.823.854.[93]
Pleurotus florida55.[107]
Pleurotus ostreatus31.7–66.115.4–29.730.8–73.42.7–3.413.8–29.87.1—8.0[86,109]
Banana leavesPleurotus ostreatusND15.[62,110]
Pleurotus pulmonarius17.916.9–23.526.21.9–5.55.8–7.26.4–10.3[62,100]
Volvariella volvacea15.223.9NDND8.16.1[111]
Soya stalkLentinus sajor-caju83.025.852.[89]
Pleurotus eous82.330.550.[92]
Pleurotus ostreatus85.224.753.[101]
Pleurotus columbinus90.67.433.[91]
Pleurotus florida87.623.557.[112]
Pleurotus sapidus72.726.824.[96]
Sunflower stalkLentinus sajor-caju63.[89]
Pleurotus eous61.527.452.[92]
Pleurotus sapidus45.920.[96]
Oil palm empty fruit bunchSchizopyllum commune3.[113]
Volvariella volvacea3.6–6.533.5–41.027.9–45.73.7–5.17.7–16.09.4–9.9[114]
Cotton stalkPleurotus florida25.129.837.[115]
Pleurotus pulmonarius42.329.344.[115]
Pleurotus ostreatus44.330.[115]
Rice huskPleurotus ostreatus9.55.948.530.90.314.3[95]
Sugarcane bagasseLentinula edodes130.0–133.013.1–13.873.0–78.90.9–1.0ND6.2–7.1[116]
Pleurotus cystidiosus49.522.145.22.322.87.5[109]
Pleurotus djmor101.725.[117]
Pleurotus eryngii41.320.549.[93]
Pleurotus florida75.68.7ND4.02.50.3[118]
Pleurotus ostreatus65.727.[109]
Sugarcane strawLentinula edodes83.0–98.014.472.5–78.20.7–0.9NR6.4–6.5[116]
Cottonseed hullPleurotus florida13.620.061.211.911.95.5[119]
Pleurotus ostreatus8.917.565.[119]
Cassava peelPleurotus ostreatus24.0–26.110.5–10.773.0–74.62.1–2.28.5–8.97.5–7.7[120]
Volvariella volvacea0.6-2.311.5–14.351.4–53.42.4–2.60.4–0.55.0–6.2[121]
Hardwood sawdustHericium erinaceus47.5–50.324.860.93.6ND10.6[87]
Acacia sawdustPleurotus cystidiosus36.315.755.[109]
Pleurotus ostreatus46.419.551.31.322.05.9[109]
Beech sawdustAgrocybe cylindracea38.318.470.33.415.08.2[86]
Ganoderma lucidum61.216.877.[122]
Pleurotus ostreatus46.816.173.63.515.86.2[86]
SawdustAuricularia polytricha13.9–44.610.278.40.9ND4.2[123]
Pleurotus columbinus89.11.725.[91]
Pleurotus citrinopileatus38.4–51.624.165.62.6ND7.8[90]
Pleurotus eryngii35.519.552.[93]
“ND” = not determined.
Table 4. Biological efficiency and chemical composition of some mushrooms grown on the combination of agro-industrial wastes.
Table 4. Biological efficiency and chemical composition of some mushrooms grown on the combination of agro-industrial wastes.
Agro-Industrial WastesMushroom SpeciesBiological Efficacy (%)Chemical Composition (% Dry Weight)Reference
Crude ProteinCarbohydrateFatFiberAsh
Soya stalk (50%) + rice straw (50%)Pleurotus florida85.222.754.[111]
Pleurotus ostreatus81.723.[124]
Soya stalk (50%) + wheat straw (50%)Pleurotus florida78.222.457.[111]
Pleurotus ostreatus77.721.[124]
Wheat straw (50%) + Rice straw (50%)Hericium erinaceus32.5–37.225.660.63.9ND9.7[87]
Pleurotus florida72.320.[111]
Pleurotus ostreatus71.820.356.[124]
Oat straw (80%) + wheat bran (20%)Ganoderma lucidum2.0–2.510.6–12.5NDND47.8–57.71.3–1.5[98]
Cotton stalk (50%) + Cottonseed hull (50%)Pleurotus florida17.324.552.[119]
Pleurotus ostreatus20.222.858.02.910.85.5[119]
Acacia sawdust (50%) + corn cob (50%)Pleurotus cystidiosus43.621.444.82.823.67.3[109]
Pleurotus ostreatus58.818.746.93.324.56.7[109]
Acacia sawdust (50%) + sugarcane bagasse (50%)Pleurotus cystidiosus41.125.637.51.828.56.8[109]
Pleurotus ostreatus58.924.237.82.528.86.7[109]
Sugarcane bagasse (50%) + grasses (50%)Agaricus brasiliensis44.328.3ND1.65.86.7[125]
Rubber tree sawdust (50%) + rice straw (50%)Flammulina velutipes123.917.0–27.058.0–87.01.8–7.3ND7.3–10.4[126]
Beech sawdust (50%) + olive pruning residues (50%)Ganoderma lucidum20.515.379.[122]
Wheat straw (50%) + olive pruning residues (50%)Pleurotus ostreatus56.819.971.71.916.56.5[122]
Sawdust (90%) + rice bran (10%)Pleurotus eous48.4–68.127.828.65.617.34.9[127]
Sugarcane bagasse (50%) + rice straw (50%)Lentinus sajor-caju83.930.933.8ND24.56.9[128]
Cassava peel (50%) + corn cobs (50%)Pleurotus ostreatus31.1–33.710.6–10.873.6–74.82.1–2.28.6–8.97.3–7.8[120]
Hard wood sawdust (50%) + rice straw (50%)Hericium erinaceus36.5–[87]
Hard wood sawdust (50%) + wheat straw (50%)Hericium erinaceus41.4–46.524.760.84.2ND10.3[87]
Hardwood sawdust (30%) + corn stalk (60%)
+ rice bran (10%)
Auricularia polytricha27.3–[129]
“ND” = not determined.
Table 5. Production of enzymes in solid state fermentation of cellulose degradation by some mushrooms using agro-industrial wastes.
Table 5. Production of enzymes in solid state fermentation of cellulose degradation by some mushrooms using agro-industrial wastes.
EnzymeAgro-Industrial WastesMushroom SpeciesActivityReference
Total cellulaseWheat strawLentinula edodes45–60 U/mL[151]
Pleurotus dryinus41–120 U/mL[151,160]
Pleurotus ostreatus665–1185 U/mL[151]
Pleurotus tuber-regium505 U/mL[151]
Fomitopsis sp.3.5 U/gds[159]
Tree leaves (Fagus sylvatica)Lentinula edodes40–45 U/mL[151]
Pleurotus dryinus205 U/mL[151]
Pleurotus ostreatus14–15 U/mL[151]
Pleurotus tuber-regium20 U/mL[151]
Sorghum strawPycnosporus sanguineus0.8 U/gds[153]
Pleurotus ostreatus1.3 U/gds[153]
Pleurotus eryngii0.7 U/gds[153]
Phanerochaete chrysosporium1.1 U/gds[153]
Trametes versicolor1.0 U/gds[153]
Eucalyptus wood chipWolfiporia cocos1.4–8.3 U/gds[161]
Laetiporeus sulfureus0.8–15.4 U/gds[161]
Poria medulla-panis0.4–3.4 U/gds[161]
Pycnoporus coccineus1.8–3.7 U/gds[161]
Phlebia tremellosa0.6–2.0 U/gds[161]
Trametes versicolor0.8–4.0 U/gds[161]
EndoglucanaseWheat strawLentinus tigrinus1230 U/gds[156]
Lentinula edodes180–345 U/mL[151]
Pleurotus dryinus910 U/mL[151]
Pleurotus ostreatus185-245 U/mL[151]
Pleurotus tuber-regium150 U/mL[151]
Piptoporus betulinus83.5 U/gds[157]
Tree leaves (Fagus sylvatica)Lentinula edodes40–65 U/mL[151]
Pleurotus dryinus1130 U/mL[151]
Pleurotus ostreatus25–1300 U/mL[151]
Pleurotus tuber-regium150 U/mL[151]
Sorghum strawPycnosporus sanguineus2.0 U/gds[153]
Pleurotus ostreatus2.3 U/gds[153]
Pleurotus eryngii1.4 U/gds[153]
Phanerochaete chrysosporium4.0 U/gds[153]
Trametes versicolor2.2 U/gds[153]
Sugarcane bagasseLentinus sajor-caju13.9–18.9 U/gds[162]
Pleurotus ostreatus3.0 U/gds[163]
SawdustTrametes trogii504 U/gds[164]
Coriolus versicolor0.6 U/gds[165]
Ganoderma applanatum0.1 U/gds[165]
Pycnoporus sanguineus0.6 U/gds[165]
Trametes villosa0.2 U/gds[165]
Pleurotus ostreatus3.0 U/gds[163]
Lentinus sajor-caju0.9 U/gds[163]
Rice strawPleurotus ostreatus7.1 U/gds[163]
Lentinus sajor-caju1.9 U/gds[163]
Oak sawdustGrifola frondosa12.3 U/gds[152]
Pine chipCoriolus versicolor2.4 U/gds[165]
Ganoderma applanatum2.8 U/gds[165]
Pycnoporus sanguineus4.8 U/gds[165]
Trametes villosa3.9 U/gds[165]
Green tea wasteMicroporus xanthopus38.6 U/gds[166]
Wheat strawFomitopsis sp.53.6 U/gds[159]
ExoglucanaseOak sawdustGrifola frondosa16.2 U/gds[152]
Rice strawPleurotus ostreatus2.0 U/gds[163]
Lentinus sajor-caju1.8 U/gds[163]
Sugarcane bagassePleurotus ostreatus7.0 U/gds[163]
Lentinus sajor-caju2.0 U/gds[163]
SawdustPleurotus ostreatus2.8 U/gds[163]
Lentinus sajor-caju0.6 U/gds[163]
Corn stoverIrpex lacteus69.3 U/gds[167]
β -GlucosidaseSorghum strawPycnosporus sanguineus0.4 U/gds[153]
Pleurotus ostreatus0.2 U/gds[153]
Pleurotus eryngii0.2 U/gds[153]
Phanerochaete chrysosporium1.1 U/gds[153]
Trametes versicolor1.9 U/gds[153]
Eucalyptus wood chipWolfiporia cocos8.3–42.0 U/gds[161]
Laetiporeus sulfureus7.6–37 U/gds[161]
Poria medulla-panis2.7–10.5 U/gds[161]
Pycnoporus coccineus8.0–22.0 U/gds[161]
Phlebia tremellosa3.8–15.6 U/gds[161]
Trametes versicolor3.8–20.0 U/gds[161]
Oak sawdustGrifola frondosa2.3 U/gds[152]
Rice strawPleurotus ostreatus2.5 U/gds[163]
Lentinus sajor-caju1.2 U/gds[163]
Sugarcane bagassePleurotus ostreatus3.5 U/gds[163]
Lentinus sajor-caju2.6–12.3 U/gds[162,163]
SawdustPleurotus ostreatus2.2 U/gds[163]
Lentinus sajor-caju0.2 U/gds[163]
Coriolus versicolor0.5 U/gds[165]
Ganoderma applanatum0.4 U/gds[165]
Pycnoporus sanguineus0.4 U/gds[165]
Trametes villosa0.5 U/gds[165]
Trametes trogii0.89 U/gds[164]
Pine chipCoriolus versicolor0.3 U/gds[165]
Ganoderma applanatum0.1 U/gds[165]
Pycnoporus sanguineus0.8 U/gds[165]
Trametes villosa0.5 U/gds[165]
Wheat strawPiptoporus betulinus78.8 U/gds[157]
Pleurotus dryinus401 U/gds[160]
Lentinula edodes0.1 U/gds[168]
Sorghum strawPleurotus eryngii0.23 U/gds[153]
Table 6. Production of enzymes in solid state fermentation of hemicellulose degradation by some mushrooms using agro-industrial wastes.
Table 6. Production of enzymes in solid state fermentation of hemicellulose degradation by some mushrooms using agro-industrial wastes.
EnzymeAgro-Industrial WastesMushroom SpeciesActivityReference
Total xylanaseTree leaves
(Fagus sylvatica)
Lentinula edodes85–200 U/mL[151]
Pleurotus dryinus2145 U/mL[151]
Pleurotus ostreatus160–1400 U/mL[151]
Pleurotus tuber-regium155 U/mL[151]
Wheat strawLentinula edodes195–275 U/mL[151]
Pleurotus dryinus1450 U/mL[151]
Pleurotus ostreatus260–735 U/mL[151]
Pleurotus tuber-regium260 U/mL[151]
Wheat branFomes fomentarius7–63 U/mL[154]
Ganoderma applanatum3 U/mL[154]
Pleurotus ostreatus16 U/mL[154]
Trametes hirsuta8 U/mL[154]
Trametes ochracea35 U/mL[154]
Trametes versicolor21 U/mL[154]
Trametes pubescens32 U/mL[154]
Trametes biforme19 U/mL[154]
Endo-1,4-β-xylanaseRice strawPleurotus ostreatus21 U/gds[195]
SawdustPleurotus ostreatus9 U/gds[195]
Sugarcane bagasseGanoderma lucidum33 U/gds[196]
Tomato pomacePleurotus ostreatus9 U/gds[197]
Trametes versicolor50 U/gds[197]
Jerusalem artichoke stalkSchizophyllum commune106 U/gds[198]
Oak leavesMarasmius quercophilus73 U/gds[199]
Mycena inclinata105 U/gds[199]
Pholiota lenta83.2 U/gds[199]
Wheat strawPleurotus citrinopileatus0.12 U/gds[200]
Pleurotus ostreatus0.14 U/gds[200]
Pine wood chipCeriporiopsis subvermispora0.25 U/gds[201]
Eucalyptus wood chipCeriporiopsis subvermispora0.12 U/gds[201]
Soya branFomes sclerodermeus31 U/gds[202]
Rice bran mixed rice huskLeucoagaricus meleagris0.8 U/gds[203]
Sugarcane bagasse
mixed wheat bran
Pleurotus ostreatus8.7 U/gds[204]
Ganoderma lucidum16.3 U/gds[204]
Trametes versicolor36.7 U/gds[204]
1,4-β-XylosidaseOak leavesMarasmius quercophilus1.7 U/gds[199]
Mycena inclinata5.8 U/gds[199]
Pholiota lenta1.6 U/gds[199]
Pine wood chipCeriporiopsis subvermispora4.4 U/gds[201]
Eucalyptus wood chipCeriporiopsis subvermispora2.6 U/gds[201]
Wheat strawPleurotus citrinopileatus11.5 U/gds[200]
Pleurotus ostreatus14.3 U/gds[200]
Sugarcane bagasse mixed wheat branGanoderma lucidum0.4 U/gds[204]
Trametes versicolor1.5 U/gds[204]
Endo-1,4-β-mannanaseOak leavesMarasmius quercophilus3.4 U/gds[199]
Mycena inclinata3.2 U/gds[199]
Pholiota lenta11.8 U/gds[199]
Pine wood chipCeriporiopsis subvermispora90.4 U/gds[201]
Eucalyptus wood chipCeriporiopsis subvermispora52.2 U/gds[201]
1,4-β-MannosidaseOak leavesMarasmius quercophilus5.9 U/gds[199]
Mycena inclinata4.2 U/gds[199]
Table 7. Production of enzymes in solid state fermentation of lignin degradation by some mushrooms using agro-industrial wastes.
Table 7. Production of enzymes in solid state fermentation of lignin degradation by some mushrooms using agro-industrial wastes.
EnzymeAgro-Industrial WastesMushroom SpeciesActivityReference
LaccaseTree leaves (Fagus sylvatica)Lentinula edodes7–52 U/L[151]
Pleurotus dryinus16 U/L[151]
Pleurotus ostreatus6.3–8.0 U/L[151]
Pleurotus tuber-regium2.1 U/L[151]
Wheat strawLentinula edodes3.6–5.2 U/L[151]
Pleurotus dryinus5.7 U/L[151]
Pleurotus ostreatus1.1–10.1 U/L[151]
Pleurotus tuber-regium10 U/L[151]
Pleurotus citrinopileatus1.2–3.7 U/gds[200]
Wheat branFomes fomentarius7430–17510 U/L[154]
Ganoderma applanatum1910 U/L[154]
Pleurotus ostreatus9210 U/L[154]
Trametes hirsuta7350 U/L[154]
Trametes ochracea3930 U/L[154]
Trametes versicolor17860 U/L[154]
Trametes pubescens5319 U/L[154]
Trametes biforme4960 U/L[154]
Wheat bran mixed corn strawTrametes versicolor32.1 U/gds[227]
LaccaseCorn stalkTrametes versicolor2,765.81 U/L[228]
SawdustCoriolopsis gallica200 U/gds[229]
Sugarcane bagassePleurotus ostreatus151.6 U/gds[230]
Oat huskCerrena unicolor28.2 U/gds[231]
Pineapple leavesGanoderma lucidum42.7 U/gds[232]
Rice bran mixed wheat branStereum ostrea24962 U/L[233]
Rice strawSchizophyllum commune431.2 U/gsd[234]
Sugarcane bagasse mixed
wheat bran
Ganoderma lucidum9.4 U/gds[204]
Pleurotus ostreatus2.1 U/gds[204]
Trametes versicolor1.9 U/gds[204]
Soya branFomes sclerodermeus14.5 U/gds[202]
Manganese peroxidaseTree leaves (Fagus sylvatica)Lentinula edodes1.0–6.7 U/L[151]
Pleurotus dryinus5.7 U/L[151]
Pleurotus ostreatus7–15 U/L[151]
Pleurotus tuber-regium20 U/L[151]
Wheat strawLentinula edodes20–55 U/L[151]
Pleurotus dryinus13 U/L[151]
Pleurotus ostreatus7–12 U/L[151]
Pleurotus tuber-regium2.2 U/L[151]
Pleurotus citrinopileatus4.9 U/gds[200]
Wheat branFomes fomentarius350 U/L[154]
Pleurotus ostreatus20 U/L[154]
Trametes versicolor20–50 U/L[154]
Trametes biforme570 U/L[154]
Oat huskCerrena unicolor20.4 U/gds[231]
Pineapple leavesGanoderma lucidum82.7 U/gds[232]
Rice bran mixed wheat branStereum ostrea3895 U/L[233]
Rice strawSchizophyllum commune1964 U/gsd[234]
Eucalyptus sawdustLentinula edodes700 U/gds[235]
Soya branFomes sclerodermeus14.5 U/gds[202]
Sugarcane bagasse mixed
wheat bran
Ganoderma lucidum1.9 U/gds[204]
Pleurotus ostreatus2.3 U/gds[204]
Tramets versicolor2.1 U/gds[204]
Barley huskBjerkandera adusta510 U/kgds[236]
Lignin peroxidasesJatropha wastePleurotus ostreatus49916 U/L[237]
Corn cobGanoderma lucidum561.4 U/gds[238]
Pineapple leavesGanoderma lucidum287.5 U/gds[232]
Rice bran mixed wheat branStereum ostrea72.8 U/L[233]
Rice strawSchizophyllum commune1467.3 U/gsd[234]
Barley huskBjerkandera adusta1700 U/kgds[236]
Sugarcane bagasse mixed
wheat bran
Ganoderma lucidum0.6 U/gds[204]
Pleurotus ostreatus0.5 U/gds[204]
Trametes versicolor0.7 U/gds[204]
Versatile peroxidaseBanana peelPleurotus eryngii36 U/gds[239]

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MDPI and ACS Style

Kumla, J.; Suwannarach, N.; Sujarit, K.; Penkhrue, W.; Kakumyan, P.; Jatuwong, K.; Vadthanarat, S.; Lumyong, S. Cultivation of Mushrooms and Their Lignocellulolytic Enzyme Production Through the Utilization of Agro-Industrial Waste. Molecules 2020, 25, 2811.

AMA Style

Kumla J, Suwannarach N, Sujarit K, Penkhrue W, Kakumyan P, Jatuwong K, Vadthanarat S, Lumyong S. Cultivation of Mushrooms and Their Lignocellulolytic Enzyme Production Through the Utilization of Agro-Industrial Waste. Molecules. 2020; 25(12):2811.

Chicago/Turabian Style

Kumla, Jaturong, Nakarin Suwannarach, Kanaporn Sujarit, Watsana Penkhrue, Pattana Kakumyan, Kritsana Jatuwong, Santhiti Vadthanarat, and Saisamorn Lumyong. 2020. "Cultivation of Mushrooms and Their Lignocellulolytic Enzyme Production Through the Utilization of Agro-Industrial Waste" Molecules 25, no. 12: 2811.

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