literature review xylanase

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• Published: 18 October 2019

A detailed overview of xylanases: an emerging biomolecule for current and future prospective

• Nisha Bhardwaj 1   na1 ,
• Bikash Kumar 1   na1 &
• Pradeep Verma   ORCID: orcid.org/0000-0003-2266-9437 1  

Bioresources and Bioprocessing volume  6 , Article number:  40 ( 2019 ) Cite this article

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Xylan is the second most abundant naturally occurring renewable polysaccharide available on earth. It is a complex heteropolysaccharide consisting of different monosaccharides such as l -arabinose, d -galactose, d -mannoses and organic acids such as acetic acid, ferulic acid, glucuronic acid interwoven together with help of glycosidic and ester bonds. The breakdown of xylan is restricted due to its heterogeneous nature and it can be overcome by xylanases which are capable of cleaving the heterogeneous β-1,4-glycoside linkage. Xylanases are abundantly present in nature (e.g., molluscs, insects and microorganisms) and several microorganisms such as bacteria, fungi, yeast, and algae are used extensively for its production. Microbial xylanases show varying substrate specificities and biochemical properties which makes it suitable for various applications in industrial and biotechnological sectors. The suitability of xylanases for its application in food and feed, paper and pulp, textile, pharmaceuticals, and lignocellulosic biorefinery has led to an increase in demand of xylanases globally. The present review gives an insight of using microbial xylanases as an “Emerging Green Tool” along with its current status and future prospective.

Introduction

The major constituent of the plant cell wall is “lignocelluloses”, as the name suggests it consists of lignin (15–20%), hemicellulose (25–30%) and cellulose (40–50%) (Gray et al. 2006 ; Singla et al. 2012 ). These components together form a three-dimensional complex network with the help of covalent and non-covalent interactions (Sánchez 2009 ). Hemicelluloses consist of xylan, a heteropolysaccharide substituted with monosaccharides such as l -arabinose, d -galactose, d -mannoses and organic acids such as acetic acid, ferulic acid, glucuronic acid interwoven together with help of glycosidic and ester bonds (Collins et al. 2005 ; Ahmed et al. 2007 ; Motta et al. 2013 ; Sharma 2017 ). Xylan is readily available in nature, followed by cellulose the second most abundant polysaccharide which covers 33% of total lignocellulosic biomass found on the globe (Collins et al. 2005 ; Polizeli et al. 2005 ; Chávez et al. 2006 ; Walia et al. 2017 ). It accounts for 15–30% in hardwoods and 7–10% in softwood (Walia et al. 2017 ). There is a need for depolymerization of this complex polymer for its efficient utilization in different industrial application. Xylanase is a group of enzymes consisting of endo -1,4-β- d -xylanases (EC 3.2.1.8), β- d -xylosidases (E.C. 3.2.1.37), α-glucuronidase (EC 3.2.1.139) acetylxylan esterase (EC 3.1.1.72), α- l -arabinofuranosidases (E.C. 3.2.1.55), p -coumaric esterase (3.1.1.B10) and ferulic acid esterase (EC 3.1.1.73) involved in the depolymerization of xylan into simple monosaccharide and xylooligosaccharides (Gomez et al. 2008 ; Juturu and Wu 2014 ; Walia et al. 2017 ; Romero-Fernández et al. 2018 ).

Xylanases are produced by different living organisms such as microorganisms, protozoans, and molluscs, and also found in the rumen of higher animals (Beg et al. 2001 ). The xylanases are mainly produced by microorganisms, e.g., bacteria, fungi, and actinomycetes at industrial scale (Motta et al. 2013 ). The utilization of lignocellulosic biomass (LCB) for production of different biochemicals such as bioethanol, enzymes, and value-added compounds has tremendously improved in recent years. It results in providing opportunities for scientists to explore the hydrolytic potential of xylanase for efficient saccharification of LCB for ethanol and xylooligosaccharides generation. Xylanase also finds application in several industries like pulp and paper bleaching, food, feed, and pharmaceuticals.

Xylanase is required in huge amount for industrial level application with characteristic properties to survive the harsh industrial level processing’s (Qiu et al. 2010 ). Therefore, there is a need to select potent microorganisms for xylanase production, followed by optimization of media components for enhanced production. The understanding of the genetic constituents of the microbe will help in deducing the mode of action of the enzyme. This will help in regulating the enzyme action for employment in desired industrial application. The microorganisms also produce other protein and metabolites with desired xylanase enzyme. Therefore, purification of the crude enzyme is a prerequisite to obtain purified enzymes. The characterization of purified xylanase will help in elucidating its stability and specificity toward different substrates. This will help in selecting the suitable industrial process in which it can be utilized. With the advent of advanced biotechnological techniques such as recombinant DNA technology, several attempts have been made to identify, isolate and clone the gene encoding for xylanase in a suitable system. This approach helps in the engineering of efficient microorganisms for enhanced xylanase production with desired properties. This review gives a comprehensive insight into xylanase classification, its mode of action, different xylanase sources with available production methods and its optimization strategies for enhanced production. The review also gives a brief idea about different strategies employed for xylanase purification and characterization, biotechnological approach for enhanced xylanase production with desired properties which are further used for different industrial applications.

Structure of xylan and role of xylanolytic enzymes in its breakdown

Xylan consists of d -xylose backbone linked with β-1,4-glycosidic bonds and l -arabinose traces forming into a complex heteropolymeric structure. Xylan is present in various biomasses that have several forms such as in hardwoods as O -acetyl-4- O -methylglucuronoxylan, in softwoods as arabino-4- O -methylglucuronoxylan and in grasses and annual plants as arabinoxylans. These residues can be substituted with acetyl, feruloyl, glucopyranosyl, 4- O -methyl- d -glucuronopyranosyl, p -coumaroyl or α- l -arabinofuranosyl side-chain groups with varying degrees. Xylanolytic enzymes play a key role in the breakdown of the complex structure of xylan. Hence, for complete and efficient hydrolysis of xylan into its constituent sugars requires synergistic action of various enzymes with specifically targeting appropriate bonds of xylan.

The multifunction xylanolytic system exists in bacteria (Zhang et al. 2016a , 2016b ), fungi (Driss et al. 2011 ; Bhardwaj et al. 2018 ) and actinomycetes (Hunt et al. 2016 ) where xylan backbone is randomly cleaved by the action of endo -1,4-β- d -xylanases; xylose polymer is broken down to its monomeric form by action of β- d -xylosidases. Acetyl and phenolic side branches were removed by the action of α-glucuronidase and acetylxylan esterase. α- l -Arabinofuranosidases catalyze the removal of the side groups. The ester bonds present on the xylan are cleaved by the action of p -coumaric esterase and ferulic acid esterase (Beg et al. 2001 ; Collins et al. 2005 ; Chakdar et al. 2016 ; Walia et al. 2017 ). The schematic structure of xylan showing bonds which are attacked by a specific xylanolytic enzyme for complete hydrolysis of xylan to its constituent monomeric units is represented in Fig.  1 .

(Adapted from Beg et al. 2001 , Lange 2017 )

Structure of xylan showing bonds which are attacked by specific xylanolytic enzyme for complete hydrolysis of xylan to its constituents

Classification of xylanase

Xylanase can be broadly classified into three types on the basis of (a) molecular mass and isoelectric point, (b) crystal structure and (c) catalytic/kinetic property (Wong et al. 1988 ; Jeffries 1996 ; Biely et al. 1997 ; Liu and Kokare 2017 ). On basis of molecular mass and isoelectric point, the xylanase was classified into two groups, i.e., (a) high-molecular weight with low isoelectric (acidic) point (HMWLI) and (b) low-molecular weight with high isoelectric (basic) point (LMWHI). However, several exceptions to this classification have been observed where not all xylanases fall in the category of HMWLI (above 30 kDa) or LMWHI (below 30 kDa) (Collins et al. 2002 , 2005 ). Therefore, a more appropriate system including primary structure (crystal), comparison of catalytic domain with mechanistic features such as kinetic, catalytic property, substrate specificity, and product description was introduced (Henrissat and Coutinho 2001 ; Collins et al. 2005 ). The genomic, structural (3D crystal structure) and functional information of xylanase is available under glycoside hydrolase (GH) families available on carbohydrate-active enzyme (CAZy) database.

The CAZy is knowledge-based, highly curated database on enzymes that play a key role in breakdown, modification, and assembly of glycosidic bonds in carbohydrates and glycoconjugates. It consists of genomic, sequence annotation, family classifications, structural (3D crystal) and functional (biochemical) information on carbohydrate-active enzyme from publicly available resources such as National Center for Biotechnology Information, NCBI (Lombard et al. 2014 ).

The major GH families associated with xylanase are 5, 7, 8, 9, 10, 11, 12, 16, 26, 30, 43, 44, 51, and 62. The GH families 5, 7, 8, 10, 11, and 43 have a single distinct catalytic domain, whereas enzymes grouped under GH families 16, 51, and 62 have two catalytic domains with bi-functional property (Collins et al. 2005 ). The enzyme grouped under GH families 9, 12, 26, 30, and 44 has secondary xylanase activity. Based on the hydrophobic cluster analysis of the catalytic domains along with similarities studies of amino acid sequences, xylanases have been primarily classified as GH 10 and GH 11 (Verma and Satyanarayana 2012a ). The catalytic properties of GH 10 and GH 11 have been studied extensively, whereas the information on GH families 5, 7, 8 and 43 is very limited (Taibi et al. 2012 ). Different structural and functional properties of different GH families are tabulated in Table  1 .

Mode of action of xylanases grouped under various GH families

There is the difference in structure, physicochemical properties, substrate specificities and mode of action of members of GH families 5, 7, 8, 10, 11 and 43 (Collins et al. 2005 ). The hydrolysis of xylan by xylanase may occur by two different mechanisms, i.e., retention or inversion (Subramaniyan and Prema 2002 ; Lombard et al. 2014 ).

This process is represented by double displacement mechanism with α-glycosyl and oxo -carbonium intermediate formation followed by its subsequent hydrolysis. Glutamate residues play a vital role in the catalytic mechanism. First, two carboxylic acid residues present in the active site result in α-glycosyl enzyme intermediate formation. The intermediate formation occurs via protonation of the substrate by a carboxylic acid residue acting as an acid catalyst and departure of the leaving group due to nucleophilic attack caused by another carboxylic acid. This collectively results in β to α inversion due to the α-glycosyl enzyme intermediate formation. Second, the first carboxylate group abstracts a proton from a nucleophilic water molecule and attacks the anomeric carbon resulting in second substitution, where the anomeric carbon gives rise to product with the β configuration (α to β inversion) via a transition state of oxo-carbonium ions (Fig.  2 ) (Collins et al. 2005 ; Lombard et al. 2014 ). Enzymes of families 5, 7, 10, and 11 mostly work on the principle of retention.

Mode of action of xylanase: retention

The enzymes of families 8 and 43 act via inversion of the anomeric center with glutamate and aspartate as the major catalytic residue. This is a single displacement mechanism, in which only one carboxylate ion offers for overall acid catalyzed group departure (Fig.  3 ). This enzyme also acts as the base for activating a nucleophilic water molecule to attack the anomeric carbon (depending on the distance between two molecules) for breaking the glycosidic bonds and causing inversion of anomeric carbon configuration (Collins et al. 2005 ; Motta et al. 2013 ; Lombard et al. 2014 ).

Mode of action of xylanase: inversion

Several attempts have been made to understand the mode of action of xylanase obtained for different organisms. An unusual mode of action of GH8 xylanase (β-xylosidase, an α-arabinofuranosidase, and an acetylesterase activity) was observed in Pseudoalteromonas atlantica , which showed the presence of a long tail of unsubstituted xylose residue on the reducing end of oligosaccharides produced (Ray et al. 2019 ). Thermophilic xylanase obtained from Bacillus licheniformis DMS has novel hydrolysis properties similar to GH30. It breaks the linear β-(1-4) linkage of beech wood and birchwood xylan along with glucuronoxylan and arabinoxylan. B. licheniformis DMS xylanase had both the properties of endoxylanase and appendage dependent xylanase activity. It showed equal production of both xylobiose and xylotriose by hydrolysis of the commercial substrate and agro-waste such as corn cob (Ghosh et al. 2019 ).

Thermothelomyces thermophila ( Tt Xyn30A) that hydrolyzes xylan into xylose and two acidic xylooligosaccharides, namely xylotriose (MeGlcA 2 Xyl3) and xylobiose, i.e., MeGlcA 2 Xyl 2 , was studied. TtXyn30A catalyzed the release of the disaccharide xylobiose from the non-reducing end of xylooligosaccharides, thus exhibiting an exo-acting catalytic behavior. TtXyn30A also showed the capability to cleave linear parts of xylan and uronic xylooligosaccharides as well as resulting in the formation of aldotriuronic and aldotetrauronic acid (Katsimpouras et al. 2019 ). Puchart et al. ( 2018 ) have reported the mode of action of hydrolysis of eucalyptus plant using endoxylanase belonging to GH10, GH11, and GH30 family. All the endoxylanse resulted in the formation of acetylated XOS. The GH10 endoxylanase results in short xylooligosaccharides, whereas GH30 endoxylanase results in longer xylooligosaccharides. An acetyl esterase (AcXEs) played a key role in understanding the plant decay or depolymerization mechanism and also showed efficiency in plant biomass bioconversion (Rytioja et al. 2014 ).

A novel modular endoxylanase with transglycosylation activity was reported from Cellulosimicrobium sp. HY-13 belonging to GH6 family (Ham et al. 2012 ). A GH30 family xylanase XynA was reported from Erwinia chrysanthemi belonging to subfamily 8 with the special property of hydrolyzing 4- O -methyl-glucuronoxylan (Urbániková et al. 2011 ). Xyn11B from thermophilic fungus Humicola insolens Y1 encoding multi-cellular xylanase belonged to GH11 reported by Shi et al. ( 2015 ). Bacteroides intestinalis DSM17393, a xylan degrading human gut bacterium, reported the presence of two putative GH8 xylanases which hydrolyze both xylopentose and xylohexose (Hong et al. 2014 ). Endo xylanase XynB from marine bacterium Glaciecola mesophila KMM241 with xylan binding ability and GH8 catalytic domain was reported by Guo et al. ( 2013 ).

Mechanism for glycosidic hydrolase family 10 (GH10)

Among all the above-mentioned GH families, GH 10 consists of endoxylanase, e.g., endo -1,4-β-xylanases, endo -1,3-β-xylanases and cellobiohydrolases (Collins et al. 2005 ). Endo -1,4-β-xylanases or xylanase mainly comes under this GH10 family. It usually consists of high-molecular weight xylanase with low isoelectric points and displays an (α/β) 8 -barrel fold. This structure mimics the shape of a ‘Salad Bowl’, because of an enlarged loop architecture, one face of the molecule is having ~ 45 Å large radius and the other face is having ~ 30 Å radius because of simple (α/β) turns (Zhang et al. 2016a , 2016b ). However, these two categories are relatively the same because along with sharing similar fold also shares some common residues and has similar catalytic mechanisms. The xylanase belonging to GH 10 family has low substrate specificity and, however, exhibits high catalytic versatility than that of GH 11 family. Xylanase belonging to GH10 family exhibits greater catalytic versatility and lower substrate specificity as compared to those belonging to GH11 (Biely et al. 1997 ; Faulds et al. 2006 ; Motta et al. 2013 ). GH10 xylanase attacks the xylose linkages which are closer to the side-chain residues (Dodd and Cann 2009 ). This could be explained by fact that the xylose residues bind at subsites (Fig.  4 ) on xylanase that causes cleavage of the bond between the monomeric residues at the non-reducing (− 1) and the reducing end (+ 1) of the polysaccharide substrate (Davies et al. 1997 ).

Schematic representation of site for attack of GH10 xylanase on xylan

Maslen et al. ( 2007 ) demonstrated that when arabinoxylan was hydrolyzed by GH10 and GH11 xylanase, the products generated have arabinose residues substituted on xylose at the + 1 subsite and + 2 subsites, respectively. Therefore, xylanases from family 11 and 10 preferentially cleave the unsubstituted regions of the arabinoxylan backbone and the unhampered substituted regions along the xylan backbone (Biely et al. 1997 ; Motta et al. 2013 ). The degree of side-chain decorations of xylan influences the specificity of the enzyme toward substrates and, thus, has an important implication on the hydrolytic product formation by xylan deconstruction (Dodd and Cann 2009 ). Yang and Han ( 2018 ) demonstrated the positional binding and substrate interaction of GH10 xylanase of Thermotoga maritime using molecular docking approach.

Researchers have reported in their previous studies that GH10 endoxylanase had better performance than GH11 in synergy with cellulase enzyme for pretreated lignocellulosic biomass hydrolysis. The reason behind this may be because GH11 endoxylanase has the lower accessibility toward acetylated xylan backbone (Faulds et al. 2006 ). Hu et al. ( 2011 ) proposed a model holocellulosic substrate, i.e., mixture of pure cellulose and 10% pre-deacetylated commercial birchwood xylan to understand the synergism between two family xylanase and cellulase (during the xylan extraction process). This study showed that substrate deacetylation has increased the hydrolytic performance of GH11 as the acetyl group restricted the accessibility of xylan more for GH11 than GH10. Thermostability is the second factor for better performance of GH10 endoxylanase over GH11 because lignocellulosic biomass hydrolysis occurs better at high temperature (50 °C) and 2–3 days long-time duration.

Source for xylanase production

The xylanase is ubiquitous in nature and its presence is observed diversely in a wide range of living organisms, such as marine, terrestrial and rumen bacteria (Chakdar et al. 2016 ), thermophilic and mesophilic fungi (Chadha et al. 2019 ; Singh et al. 2019 ), protozoa (Devillard et al. 1999 ; Béra-Maillet et al. 2005 ), crustaceans (Izumi et al. 1997 ), snails (Suzuki et al. 1991 ), insects (Brennan et al. 2004 ), algae (Jensen et al. 2018 ), plants and seeds (immature cucumber seeds and germinating barley) (Bae et al. 2008 ; Sizova et al. 2011 ). Bacteria and fungus are widely used for industrial production of xylanase. Several microbial sources of xylanase are classified in Table  2 .

Bacterial sources of xylanase

Among bacteria, Bacillus species have been reported widely as the most potent xylanolytic enzyme producers such as Bacillus sp., B. halodurans (Gupta et al. 2015 ) , B. pumilus (Thomas et al. 2014 ), B. subtilis (Banka et al. 2014 ), B. amyloliquefaciens , B. circulans , and B. stearothermophilus (Chakdar et al. 2016 ). Xylanase with high temperature stability, acid/alkali stability, and cold adaptability have been isolated and purified from a wide range of bacteria found in extreme environment. Thermotolerant xylanase active at a very high temperature of 60–70 °C has been reported from Bacillus spp. (Thomas et al. 2014 ), Bacillus Halodurans TSEV1 (Kumar and Satyanarayana 2014 ), Clostridium thermocellum (Fernandes et al. 2015 ), Rhodothermus marinus (Karlsson et al. 2004 ), Streptomyces sp. (Sukhumsirichart et al. 2014 ), Stenotrophomonas maltophila (Raj et al. 2013 ), Thermotoga thermarum (Shi et al. 2013 ). Psychrophilic xylanases are not very common but found to be isolated from several bacteria such as Clostridium sp. PXLY1 (Akila and Chandra 2003 ), Flavobacterium sp. MSY-2 and Flavobacterium frigidarium (Humphry et al. 2001 ; Dornez et al. 2011 ) Pseudoalteromonas haloplanktis TAH3A (Van Petegem et al. 2002 ).

Several alkali stable xylanases have been isolated from firmicutes such as B. pumilus (Thomas et al. 2014 ), B. halodurans TSEV1 (Kumar and Satyanarayana 2014 ) and Geobacillus thermoleovorans (Verma and Satyanarayana 2012b ) and actinomycetes such as Actinomadura sp. Cpt20 (Taibi et al. 2012 ) and Streptomyces althioticus LMZM (Luo et al. 2016 ).

Fungal sources of xylanases

The mesophilic fungi of genera Aspergillus and Trichoderma are well known to be potent xylanase producer and most widely used for commercial production. Thielavia terrestris, (Garcia-Huante et al. 2017 ) , Talaromyces thermophilus (Maalej et al. 2009 ), Paecilomyces thermophile (Fan et al. 2012 ), Achaetomium sp. X2-8 (Chadha et al. 2019 ), Rhizomucor pusillus (Hüttner et al. 2018 ), Rasamsonia emersonii, (Martínez et al. 2016 ) T. Leycettanus (Wang et al. 2017 ) , Melanocarpus albomyces (Gupta et al. 2013 ) and Aspergillus oryzae LC1 (Bhardwaj et al. 2019 ) were found to be producer of hyper-thermophilic active xylanase. Several alkali stable xylanases were obtained from different fungal strains such as Paenibacillus barcinonensis (Valenzuela et al. 2010 ) , Aspergillus fumigatus MA28 (Bajaj and Abbass 2011 ), Cladosporium oxysporum (Guan et al. 2016 ) and Aspergillus oryzae LC1 (Bhardwaj et al. 2019 ).

Strategies employed for xylanase production from different microbial sources

The production of xylanase from microorganisms is affected by the fermentation process employed, choice of substrate and different media components. These components are often regulated by different process optimization for enhanced production of the enzyme for its application at large scale.

Different fermentation process employed for xylanase production: submerged and solid-state fermentation

Xylanases are produced by a different fermentation process using various microorganisms. The better understanding of the physiology and different metabolic processes of the microbial system has led to an improvement in the fermentation process. However, there is still an opportunity to improve the yield of enzymes. The optimization of the xylanase production will be discussed in a later section.

The xylanase production has been carried out under submerged fermentation (SmF) and solid-state fermentation (SSF) (Motta et al. 2013 ). The choice of the fermentation process usually depends on the type of microorganisms used (Table  3 ). Bacteria require a high amount of water during growth; therefore, SmF is preferred whereas fungi due to its mycelia nature require less moisture and can be grown under SSF (Walia et al. 2017 ). Several reports suggest that submerged fermentation using bacteria and fungi is the most preferred method for xylanase production. Statistically speaking approximately 90% of total xylanase is produced globally through SmF. During SmF, the synergistic effect of different xylan degrading enzymes can be observed and even result in better biomass utilization for enhanced xylanase production (Polizeli et al. 2005 ; Bajpai 2014 ). Xylanase production utilizes soybean residues and rice straw as a substrate under SmF by Aspergillus oryzae LC1 and Aspergillus foetidus (Bhardwaj et al. 2017 ; De Queiroz Brito Cunha et al. 2018a , b ). Similarly, Irfan et al. ( 2016 ) suggested the production of xylanase under SmF by B. subtilis  BS04 and B. megaterium  BM07. Different advantages of the SmF are homogenous condition throughout medium; method is well characterized and can be easily scaled up (Guleria et al. 2013 ). There are some disadvantages to SmF as well which limit its industrial application, i.e., high maintenance cost, energy intensive and complex downstream (Virupakshi et al. 2005 ; Walia et al. 2017 ).

Recent trends suggest that xylanase production by SSF is also gaining popularity (Walia et al. 2014 ). Bacillus sp. was used for the production of thermo-alkalophillic extracellular xylanase under SSF using wheat bran as substrate (Kamble and Jadhav  2012 ). Similarly, SSF of Trichoderma koeningi  using corn cob supplemented with pineapple peel powder showed enhanced production of xylanase (Bandikari et al. 2014 ). It has several advantages such as low cultivation, operation and capital cost, a lower rate of contamination, easy enzyme recovery, and high productivity per reactor volume. The disadvantages associated with SSF are not suitable for all microorganisms (preferred for the fungal system) and require proper aeration and humidity control and up-scaling is a tedious process (Mienda et al. 2011 ).

Selection of suitable substrate for xylanase production

Quantity and quality of the fermentation product vary with different substrates. There are various commercially available substrates, i.e., xylan, carboxymethyl cellulose (CMC), pectin, and starch for, i.e., xylanase, cellulase, pectinase, and amylase, respectively (Barman et al. 2015 ; Bhardwaj et al. 2017 ; Kumar et al. 2018a ). Due to the high cost of commercial substrates and considering the economic feasibility of the process, scientists are working from past several years to find alternative substrates for the production of these enzymes.

Agrowastes and other organic wastes (domestic and industrial) are used as a carbon source for the production of xylanase with the focus on sustainability and best utilization of these wastes (Table  3 ). Some of the most commonly used agro-residues for xylanase production are wheat bran, wheat husk (Kumar et al. 2018a , b , c , d ), rice straw (Bhardwaj et al. 2017 ), rice husk, sugarcane bagasse (Suleman and Aujla 2016 ), coconut coir, coconut oil cake (Rosmine et al. 2019 ), groundnut shell (Namasivayam et al. 2015 ), wood pulp (Kalpana and Rajeswari 2015 ), sawdust, chilli post-harvest (Sindhu et al. 2017 ), corncobs, molasses, sugar beet pulp fruit, and vegetable waste (Bandikari et al. 2014 ). Recent studies also showed that wastewater from pulp industry was reused as media for xylanase production (de Queiroz-Fernandes et al. 2017 ).

Role of important media components used for xylanase production

Naturally, xylanolytic enzymes are induced by the different intermediate products generated by their own action. Xylan is found to be best xylanase inducer (Taibi et al. 2012 ; Guleria et al. 2013 ; Walia et al. 2013 , 2014 ). However, xylan being a high-molecular weight polymer cannot stimulate xylanase as it cannot enter the microbial cells. Therefore, a small amount of constitutive enzyme produced in the media results in the generation of low-molecular weight fragments, i.e., xylobiose, xylotriose, xylotetraose, xylose from the breakdown of xylan and further induces the xylanolytic enzymes for enhanced enzyme production (Walia et al. 2017 ). Cellulose, synthetic alkyl, aryl β- d xylosides, and methyl β- d -xyloside also act as an inducer for xylanolytic enzyme production (Thomas et al. 2013 ). Busk and Lange ( 2013 ) observed that poor quality paper can efficiently induce the xylanase production in Thermoascus aurantiacus even in the absence of xylan and xylooligosaccharides.

Nitrogen is an important structural element required for the metabolic processes in the microbial system. Therefore, the choice of nitrogen source is important for the growth of microorganisms that subsequently affect the overall enzyme yield. Peptone, tryptone, soymeal, yeast extract, etc. have found to be suitable nitrogen source. The requirement of these nitrogen sources varies for different microorganisms; therefore, optimizing the type and level of nitrogen source in the media is an important parameter (Seyis and Aksoz 2005 ; Naveen et al. 2014 ; Irfan et al. 2016 ). Trace elements, amino acids, and vitamins are also important parameters for the growth of different microorganisms (Simair et al. 2010 ; Bibra et al. 2018 ). Therefore, regulating their levels in the media is important for regulating the production of xylanase. Also, the addition of biosurfactant such as Tween 80 affected the level of xylanase production (Liu et al. 2006 ; Kumar et al. 2013 ).

Strategies employed for the selection of the method of xylanase production and its optimization

Intially a common minimal media providing essential nutrients to the growth of microorganisms are used. This will allow to check the strains are capable of producing required enzymes/metabolite of desired interest. Then, the process is further optimized for higher production of enzymes from the strain (Walia et al. 2017 ). For the production of desired product, different strategies are used for improving yield such as optimization of media components, regulating physical growth parameters, and improving the strain by use of the different biotechnological tool (Sharma 2017 ). The schematic representation of the methodology adapted for production, purification and characterization of xylanase are shown in Fig. 5 . In this section, the focus will be on the optimization of media and growth parameters and biotechnological tool approach will be discussed in a later section. During SmF for enzyme production, different components which need to be optimized are selection of substrate and microorganisms, regulation of nutrients concentration in media, i.e., carbon, nitrogen, trace elements, vitamins and amino acids, and physical parameters, i.e., temperature, pH, agitation, aeration, inoculum sizes, and incubation period (Motta et al. 2013 ; Walia et al. 2015a , 2017 ).

Schematic representation of the methodology for production, purification and characterization of xylanase

During optimization of the SSF, there is requirement of regulating particle size, pretreatment, humidity, water content and water activity (a w ) of substrate, type and size of the inoculums, removal of extra heat generated during microbial metabolism and most importantly maintaining the uniform environment (temperature) and evolution of CO 2 and consumption of O 2 , i.e., gaseous system (Murugan et al. 2011 ; Behera and Ray 2016 ; Behnam et al. 2016 ; Leite et al. 2016 ; Walia et al. 2017 ).

Approach for enhanced xylanase production: one factor at a time (OFAT)

To proceed for the optimization of the xylanase production, one factor at a time (OFAT) approach is used for the selection of important factors affecting the xylanase yield. In the OFAT approach, one factor is kept variable keeping other factors at constant (Bhardwaj et al. 2018 ). The factor may be important physical or nutritional parameters regulating the growth of microorganisms and its enzyme yield. Ramanjaneyulu et al. ( 2017 ) have evaluated several operating parameters for nutritional (different substrates and their concentrations, additional carbon and nitrogen sources) and physical factors (incubation temperature, pH, agitation) along with inoculum size of Fusarium  sp. BVKT R2 in a shake flask culture (SmF) by OFAT approach. The high xylanase yield of 4200 U/mL was obtained with birch wood xylan in mineral salt medium with 1.5% sorbitol (additional carbon source), 1.5% yeast extract (nitrogen source) at temperature of 30 °C, pH of 5.0, agitation of 200 rpm and inoculum of agar plugs (6) for only 5 days incubation. Under unoptimized condition, xylanase yield was only 1290 U/mL after 7 days of incubation, thus improving by 3.2-fold. Bhardwaj et al. ( 2018 ) also optimized xylanase production using Aspergillus oryzae LC1 using OFAT approach. The physical parameters (liquid to solid ratio, pH, inoculums size, incubation time and temperature) and nutrient parameters (substrate concentration) were optimized using OFAT approach for enhanced production of xylanase by T. viride -IR05 under SSF (Irfan et al. 2014 ).

Statistical approach for enhanced xylanase production

The OFAT approach is tedious and requires a large set of experiments for optimization. The recent trend suggested the application of the statistical approach to design experiments considering different factors as variable and performing the interaction studies among several physical and nutritional parameters. The statistical-based approach has shown satisfactory results for optimization of xylanase production using fungal and bacterial strains with the minimum number of experimental sets (Guleria et al. 2015 , 2016a ; Walia et al. 2015b ; Bhardwaj et al. 2017 ).

Response surface methodology (RSM) was employed to optimize the fermentation medium constituents and the physical factors affecting xylanase production using Bacillus tequilensis strain ARMATI under SmF (Khusro et al. 2016 ). The experimental design consists of central composite design (CCD) with four (4) independent variable (carbon and nitrogen source, temperature and time) resulting in 30 experimental runs. The central composite design gave an optimum parameter for studied variable (1.5% w/v birchwood xylan, 1% w/v yeast extract, temperature 40 °C, time 24 h) showing 3.7-fold enhanced xylanase production as compared to OFAT. High coefficient of determination (R 2 ) of 0.9978 with p  < 0.05 as obtained by analysis of variance (ANOVA) analysis suggested the accuracy of the overall process at a significant level. The R 2 value of 0.9978 represents that sample variation of 99.78% and only 0.21% of the total variation in the response cannot be explained by the model. The xylanase obtained has shown high thermal (60 °C) and alkali stability (pH 9). Bhardwaj et al. ( 2017 ) optimized nutritional components (rice straw, MgSO 4 , and CaCl 2 concentration) and physical parameters (temperature and pH) for enhanced xylanase production with an Aspergillus oryzae LC1 under submerged fermentation using CCD-RSM. The statistical design suggested optimum condition of 1% rice straw (w/v), 1.0 g/L CaCl 2 , and 0.3 g/L MgSO 4 , with pH 5 and 25 °C. It resulted in maximum xylanase activity of 935 ± 2.3 IU/mL which is 3.8-fold higher than the un-optimized Mendel’s Stenberg Basal Salt medium (245 ± 1.9 IU/mL). The enzyme showed thermal (25–60 °C) and pH (3–10) stability. The xylanase also showed potential for efficient enzymatic hydrolysis of different lignocellulosic agro-residues.

Similarly, Tai et al. ( 2019 ) reported the optimization of five physical and two nutritional parameters using the RSM approach for enhanced xylanase production. Indigenous fungus Aspergillus niger DWA8 was grown under SSF on an oil palm frond. One physical (moisture content 75%) and one nutritional parameters (substrate concentration 2.5 g) have significant effect on xylanase production. Under optimum condition, an increase in xylanase yield by 78.5% was observed as compared to an un-optimized condition. The xylanase was efficiently used for saccharification of biomass. The statistical optimization method for enhanced xylanase production has been applied and widely accepted for SSF and SmF that helped in overcoming several limitations of classical empirical (OFAT) methods.

Biotechnological approach for enhanced xylanase production

There is a need of high yield of the enzyme with specific properties such as high stability over a wide range of temperature and pH, high substrate specificity and strong resistance to metal cations and chemicals for the industrial application (Garg et al. 2010 ; Qiu et al. 2010 ). The native enzyme is usually produced in low quantity and also lacks all the characteristics to meet the industrial needs (Ahmed et al. 2009 ). Therefore, different biotechnological approaches are used for improving the yield and imparting characteristic properties to the desired enzyme. These approaches involve genetic manipulation involving mutation and recombinant DNA technology.

Mutagenesis of microorganisms for enhanced xylanase production

Several researchers suggested that the application of physical mutagens such as UV radiation (Rahim et al. 2009 ; Abdel-Aziz et al. 2011 ) and chemical mutagens such as N -methyl N -nitro N -nitroso guanidine (MNNG) (Haq et al. 2004 , 2008 ) resulted in enhanced xylanase production. Burlacu et al. ( 2017 ) demonstrated the improvement of xylanase production in fungal strains, i.e., Aspergillus brasiliensis and Penicillium digitatum by physical mutagenesis (5–50 min, exposure to UV light) and chemical mutagenesis (150 µg/mL of N -methyl- N ′-nitro- N -nitrosoguanidine or ethyl methane sulfonate). The exposure to physical and chemical mutagens has resulted in significant changes in the mutant strain as compared to the wild type. Han et al. ( 2017 ) demonstrated the site-directed mutagenesis of XynCDBFV gene of ruminal fungus  Neocallimastix patriciarum for improving the thermostability of XynCDBFV, a glycoside hydrolase (GH) family 11 xylanase. Similar work has also been carried out in different bacterial strains, a rifampin-resistant mutant of  Cellulomonas biazotea , designated 7Rf, resulting in elevated levels of xylanases production as compared to the parental strain. After mutation, maximum xylanase and β-xylosidase production of 493 IU/L/h and 30.7 IU/L/h of β-xylosidase were obtained respectively. This increase in xylanase and β-xylosidase yield were 1.21- and 2.29-fold higher respectively as compared to the parental strain (Rajoka et al. 1997 ). Bacillus mojavensis PTCC 1723 when subjected to UV light exposure (280 nm, 30 s) resulted in the xylanase yield 330.6 IU/mL which is 3.45 times higher as compared to 95.7 IU/mL for wild strain (Ghazi et al. 2014 ). Lu et al. ( 2016 ) demonstrated mutation of XynHB, alkaline stable xylanase from  Bacillus pumilus  HBP8 at N188A. The mutant XynHBN188A is expressed in E. coli and Pichia pastoris with improved xylanase yield by 1.5- and 7.5-fold, respectively. The codon-based optimization and high-density fermentation using Pichia pastoris system were utilized for improving the xylanase yield.

Gene cloning and expression of xylanase genes using recombinant DNA

The recombinant xylanases are designed to have equivalent or better properties than the wild-type enzymes with high yield in the expression system which can be employed in the fermentation industry. The highly thermo-alkalophilic xylanase producing strains can be directly employed during simultaneous saccharification and fermentation for ethanol generation using lignocellulosic biomass. The inherited stability will enable the xylanase to work efficiently even at high temperature and varying pH range of the fermentation system.

Several reports suggests that desired xylanase gene was cloned into the suitable vector followed by its expression in the suitable microbial systems such as bacteria, yeasts, and fungus (Belancic et al. 1995 ; Goswami et al. 2014 ; Jhamb and Sahoo 2012 ; Juturu and Wu 2012 ; Motta et al. 2013 ; Nevalainen and Peterson 2014 ; Verma et al. 2013 ).

Expression in bacteria

Goswami et al. ( 2014 ) demonstrated the expression of a xylanase gene from Bacillus brevis in E. coli BL21. The recombinant strain predominantly secreted xylanase in the culture medium with 30 IU/mL xylanase activity. The culture filtrate is free from cellulase activity and found to be active in a wide range of pH and temperature. A thermo-alkali stable xylanase encoding gene (Mxyl) was retrieved from compost-soil metagenome library construct and cloned into pET28a vector expressed in E. coli BL21(DE3). The recombinant xylanase has shown half-life of 2 h and 15 min at 80 °C and 90 °C, respectively. The recombinant xylanase has pH and temperature optima of 9.0 and 90 °C, respectively (Verma et al. 2013 ).

Escherichia coli is preferred and most widely used expression host due to its inexpensive growth conditions, easy manipulation, simple transformation techniques requirement, high level of product accumulation in the cell cytoplasm (Jhamb and Sahoo 2012 ). However, efficient and functional expression of many xylanase genes is not possible with E. coli which may be due to repetitive appearance of rare codons and the requirement for specific translational modifications (disulfide-bond formation and glycosylation) (Belancic et al. 1995 ; Jhamb and Sahoo 2012 ; Juturu and Wu 2012 ; Motta et al. 2013 ). One of the other important concerns associated with E. coli is the presence of endotoxins (lipopolysaccharide) which makes the protein purification process very tedious. Lactobacillus and Bacillus species are used for heterologous expression of xylanase than in E. coli . It is capable of performing N-glycosylation, generally regarded as safe (GRAS) due to the absence of endotoxins and their secretory production is beneficial in industries (Bron et al. 1998 ; Subramaniyan and Prema 2002 ; Upreti et al. 2003 ; Juturu and Wu 2012 ). Zhang et al. ( 2010a , b ) demonstrated the expression and characterization of the xylanase gene (xynB) from Dictyoglomus thermophilum Rt46B.1 in Bacillus subtilis system. The pH and temperature optima for the purified recombinant enzyme were 6.5 and 85 °C, respectively. The xylanase was stable up to 95 °C and retained its activity in surfactants such as EDTA, DTT, Tween-20 and Triton X-100.

Expression in yeast

The heterologous protein expression in yeast system is highly attractive due to its ability to perform eukaryotic post-translational modifications and can grow to very higher cell densities with the ability to secrete enzyme into the fermentation system. Most of the yeasts are considered as GRAS organisms and do not produce toxins (Juturu and Wu 2012 ). Saccharomyces cerevisiae already established as an industrial microorganism, thus, can be conveniently used for xylanase production (Ahmed et al. 2009 ).

The application of Pichia pastoris as expression system has gained impetuous because it can promote the expression of the protein on their own using alcohol oxidase as promoter using methanol utilization pathway (Ahmed et al. 2009 ; Juturu and Wu 2012 ; Motta et al. 2013 ). Pichia pastoris as expression system is preferred as it can grow to very high cell densities, inherit strong and tightly regulated promoters, and produce high titer of recombinant protein (g/L) both intracellularly and in the secretory manner (Ahmad et al. 2014 ). Basit et al. ( 2018 ) demonstrated the cloning of two GH11 xylanase genes, MYCTH_56237  and  MYCTH_49824 , from thermophilic fungus  Myceliophthora thermophila  and its expression in  Pichia pastoris . The specific activities of purified recombinant xylanase were observed at 1533.7 U/mg and 1412.5 U/mg for MYCTH_56237  and  MYCTH_49824 , respectively. The recombinant xylanase showed stability under harsh condition (high pH and temperature) and high efficiency for biomass saccharification. However, the application of Pichia pastoris at large scale is limited due to health and fire hazards of methanol (Ahmed et al. 2009 ). In the case of P. pastoris as expression system, lower protein yield was obtained while expressing membrane-attached protein or proteolytic degradation prone protein and complex protein such as hetero-oligomers (Ahmad et al. 2014 ).

Expression in filamentous fungi

Filamentous fungi can be efficiently used for heterologous and homologous gene expression resulting in high yield of recombinant gene products (Su et al. 2012 ; Motta et al. 2013 ; Nevalainen and Peterson 2014 ; Nevalainen et al. 2018 ). Similar to yeast, it can regulate expression yields with their own promoters and can provide eukaryotic style post-translational modification of proteins such as N-glycosylation, proteolytic processing, or formation of multiple disulfide bonds (Ahmed et al. 2009 ; Fleissner and Dersch 2010 ; Landowski et al. 2015 ).

The application of fungi as an expression system also has advantages associated with cost-effectiveness of the overall process due to low-cost substrate and downstream processing. Further, fungi have already been subjected to many strain improvement procedures for enhanced production of xylanase. Therefore, the native xylanase expressing machinery can be efficiently used for functional expression of a foreign xylanase gene from other sources. Xyn2 xylanase gene was expressed in T. reesei by homologous expression resulting in the 1.61 g/L of xylanase 2 on glucose-containing medium (Li et al. 2012 ). Godlewski et al. ( 2009 ) demonstrated xylanase B(XynB) gene expression in  T. reesei. Similarly, the expression of xylanase 2 (XYN2) and xylanase gene from the thermophilic fungus  Humicola grisea var. thermoidea and P. griseofulvum was expressed in Trichoderma reesei and Aspergillus oryzae , respectively (De Faria et al. 2002 ; Motta et al. 2013 ). Nevalainen and Peterson ( 2014 ) presented a comprehensive review on application of filamentous fungus as expression system and suggested that research is now focused on understanding the cellular mechanisms for better internal protein quality control and secretion stress. The better utilization of “omics” tools can help in improving the regulation of xylanase production using filamentous fungus as an expression system.

Strategies for enhanced purification and characterization of xylanase for industrial application

The microbial system produces a wide range of biochemical’s during different growth and development of the microorganisms. These biochemical’s are enzymes, secondary metabolites, etc. which are of great importance to human applications. Similarly, enzymes are produced by microorganisms along with other enzymes or metabolites. Therefore, purification is prerequisites for obtaining pure enzyme with minimum or no impurities (Zhang et al. 2012 ). The characterization of the purified enzyme such as evaluation of temperature and pH optimum, thermal and acid/alkali stability, role of metal ions and inhibitors in regulation of enzyme activity, and substrates specificity was performed for selecting the suitable industrial process (Bhardwaj et al. 2019 ). There are different enzyme purification strategies for the xylanase such as ammonium sulfate precipitation (salting in) followed by dialysis (salting out), gel permeation chromatography, ion exchange chromatography, recently developed techniques aqueous phase chromatography and ultrafiltration (Walia et al. 2014 ; Guleria et al. 2016b ; Bhardwaj et al. 2019 ).

Ammonium sulfate precipitation followed by dialysis

The crude xylanase preparation is subjected to different ranges of ammonium sulfate concentration (30–90%) for selection of suitable salt concentration for precipitation of the enzyme. The precipitated enzyme is then subjected to dialysis for removal of the salt. The crude xylanase obtained from Streptomyces P12-137 was subjected to ammonium sulfate precipitation (40–90%) followed by dialysis. The purification fold of 4.18 was observed with two different endoxylanase observed as F5 (65%) and F6 (80%) with the specific activity of 45.4 U/mg and 36.5 U/mg, respectively. This was also confirmed by HPLC analysis. The purified enzyme was further characterized by incubating at different temperature and pH followed by analyzing the enzyme for xylanase activity. The optimum pH and temperature of pH 7.0, 60 °C and 6.5, 60 °C, for F5 and F6 xylanase, respectively, were obtained (Coman et al. 2013 ).

Bhardwaj et al. ( 2017 ) performed partial purification of the crude xylanase obtained from Aspergillus oryzae LC1 using ammonium sulfate (60%) precipitation followed by dialysis against 50 mM acetate buffer (pH 5.0). The partially purified enzyme was further characterized which showed stability over a wide pH range of 3 to 10 and thermal stability over the temperature range of 25 to 60 °C. Similarly, Kumar et al. ( 2018d ) have demonstrated the purification and characterization of xylanase obtained from sea sediment bacteria using a combination of ammonium sulfate precipitation and dialysis. The improvement in specific activity and characteristic properties of xylanase was observed. The major limitations of the precipitation are needed to remove salt from protein sample so further processing in the form of dialysis or chromatography is required. Further, for dialysis, there is a need to have a better understanding of the protein solubility. It is also stated that ammonium precipitation concentrates the protein rather than purifying it. Thus, contaminant present in the crude sample may also be present along with the protein sample even after precipitation and dialysis (Biosciences 2019 ). The xylanase is also concentrated or precipitated using trichloroacetic acid (TCA) and acetone. However, the TCA may denature the protein; therefore, it is not advisable to use TCA when the protein is required in the folded state (for activity assay) and the toxicity of TCA also limits its applications (Koontz 2014 ).

Chromatography techniques for enhanced xylanase yield employed for purification

Usually, it has been observed that xylanase purification was performed by the multi-step process where the concentration of protein using ammonium acetate/TCA/acetone precipitation or ultracentrifugation was followed by a single step or series of chromatography techniques. Yadav et al. ( 2018 ) demonstrated the purification and characterization of extracellular xylanase obtained from  A. kamchatkensis  NASTPD13 cultures. The crude xylanase was subjected to ammonium sulfate (80%) precipitation followed by dialysis. The dialyzed sample was further subjected to Sephadex G100 column chromatography. The fractions collected showing maximum xylanase activity were concentrated and analyzed by SDS-PAGE (MW obtained was 37 kDa). The two-step purification has led to increased xylanase activity by 11-fold with a 33 U/mg specific activity. The characterization of purified protein showed pH and temperature optimum of 9.0 and 65 °C, respectively, and also retained more than 50% of its activity over a wide range of 6–9 pH and 30–65 °C temperature. An insight into several purification strategies employed for xylanase from different microorganisms along with the process efficiency in terms of recovery potential and kinetics property is tabulated in Table  4 .

Purification of endoxylanase obtained from Bacillus pumilus B20 was performed in three steps (Geetha and Gunasekaran 2017 ). The first step was ammonium sulfate precipitation (60–80%) followed by FPLC using DEAE Sepharose column as the second step and further subjecting the eluted sample onto a Sephacryl S-200 column as the third purification step. At each step, the specific activity was improved as compared to the crude enzyme by 5 to 14.8-fold with maximum 755.8 U/mg specific activity at the end of all the three purification steps. After the purification, the fractions showing maximum xylanase activity were subjected to xylanase assay and other characterization studies such as SDS-PAGE, zymography, temperature and pH stability. The SDS-PAGE and zymography analysis showed the purified enzyme of ~ 85 kDa, i.e., endoxylanase (XylB). The purified enzyme was stable in a pH range of pH 6.5 to 7.5 and the temperature range of 20 to 50 °C. The purified enzyme was highly specific to different commercial and natural xylan substrate and has the potential to generate xylooligosaccharides.

Aqueous two-phase system employed for purification of xylanase

The conventional multistep purification techniques are time consuming, which increases the cost of the overall process and also results in loss of protein at each step (Iqbal et al. 2016 ; Ramakrishnan et al. 2016 ). The 60–70% of total processing cost in enzyme downstream process comes from the purification step (Loureiro et al. 2017 ; Bhardwaj et al. 2019 ). Therefore, several scientists suggested a single step liquid–liquid fractionation technique, i.e., aqueous two-phase system (Naganagouda and Mulimani 2008 ; Yasinok et al. 2010 ; Glyk et al. 2015 ).

Garai and Kumar ( 2013 ) purified alkaline xylanase from Aspergillus candidus using aqueous two-phase system (ATPS) composed of PEG 4000/NaH 2 PO 4 system. The critical factors of ATPS such as PEG molecular weight, PEG and phosphate salt concentration using Box–Behnken design approach were used for the optimization of enhanced xylanase purification. The optimum condition was PEG 4000 at 8.66% w/w with a high salt concentration of 22.4 w/w that resulted in 8.41% purification fold. The enzyme was stable at alkaline pH and activity is enhanced with Mn 2+ ions. Ng et al. ( 2018 ) demonstrated the recovery of xylanase from Bacillus subtilis fermentation broth with an alcohol/salt ATPS. The ATPS system consists of 26% (w/w) 1-propanol and 18% (w/w) ammonium sulfate resulting in 5.74 ± 0.33 purification fold and yield of 71.88% ± 0.15.

Gómez-garcía et al. ( 2018 ) demonstrated purification of xylanase by Trichoderma harzianum using ATPS with PEG/salt system. The PEG molecular weight, PEG, phosphate salt concentration, and salt conditions were optimized. The best  enzyme recovery and purification fold of 62.5% and 10% respectively was obtained using 20.2% PEG 8000, 14.8% K 2 HPO 4 , and tie to a length of 45% w/w. Bhardwaj et al. ( 2019 ) subjected crude xylanase from Aspergillus oryzae LC1 to four different single-step purification by ammonium sulfate precipitation, ion exchange, gel filtration chromatography and ATPS PEG/Salt system. The xylanase purification using single-step ATPS system resulted in highest purification yield (PY) of 86.8% and 13-fold purification fold (PF) which was much higher than other purification strategy, i.e., ammonium precipitation (PY-21%, PF-4.1), anion exchange (PY-31.9%, PF-3) and gel filtration (PY-78.7%, PF-6.6).

Therefore, ATPS exhibits several advantages over traditional purification techniques, i.e., it requires low-cost materials, low energy consumption with high yield and better resolution (Naganagouda and Mulimani 2008 ; Yasinok et al. 2010 ; Glyk et al. 2015 ). The ATPS method is independent of protein concentration and does not affect the native property of protein (Iqbal et al. 2016 ; Ramakrishnan et al. 2016 ).

Structural properties of xylanase responsible for thermal and pH stability required for industrial application

The high stability of xylanase was due to the presence of intrinsic structural properties. The presence of extra disulfide and salt bridges, hydrophobic side chains, and N-terminal proline residues helps in reduction of conformational freedom of the protein structure. Thus, it help in providing more stability to protein at the higher temperature (Turunen et al. 2001 ; Chen et al. 2015 ). Different structural modifications such as high Thr/Ser ratio and high charged residues, i.e., Arg, cause enhanced polar interaction and improved stabilization of the alpha-helix region and secondary structures (Hakulinen et al. 2003 ). The xylanase protein has a large number of ion pairs/aromatic residues on the surface of protein resulting in enhanced interactions (Polizeli et al. 2005 ; Chen et al. 2015 ). The low average protein rigidity i.e. low B factor, low flexibility results in high rigidity at extreme physical conditions (Xie et al. 2014 ). The presence of divalent metal ions and removal of N or C terminal disordered residues protect xylanase from heat and protease inactivation (Andrews et al. 2004 ; Chen et al. 2015 ). The presence of carbohydrate-binding modules (CBM22 and CBM9) at N or C terminal often imparts heat stability to xylanase. The pH stability of the xylanases is often affected by the presence of several amino acids near the catalytic residues (Singh et al. 2019 ).

Cost estimation of the xylanase production

Polizeli et al. ( 2005 ) suggested that 20% of the total global enzyme production is from biomass hydrolysis enzymes, i.e., xylanase, cellulase and pectinases. An extensive study on cost involved in each step of xylanase production at industrial scale is unavailable on public domain. Klein-Marcuschamer et al. ( 2012 ) performed a study on the cost analysis of application of enzymes during the lignocellulosic biomass based biofuel production and suggested breakdown of the operating cost (annual) in their enzyme production facility. They suggested percentage of cost involved for each component, i.e., raw materials (28%), labor (7%), transportation (1%), consumables (4%), utilities (10%), facility dependent (48%), and waste treatment (2%). This clearly shows that maximum contribution of 48% comes from the capital investment followed by cost of substrate (28%). Klein-Marcuschamer et al. ( 2012 ) also suggested the baseline production cost of hydrolysis enzyme as $10.14/kg.

Da Gama Ferreira et al. ( 2018 ) performed techno-economic analysis of the β-glucosidase enzyme production from E. coli on industrial scale. They showed major cost during industrial production are facility dependent (45%) followed by raw materials (25%) and consumables (23%), that are similar to observations made by Klein-Marcuschamer et al. ( 2012 ). Capital investment/facility-dependent cost is required for development of infrastructure (i.e., equipments), insurance, maintenance and depreciation. This upstream and downstream process during enzyme production involves the cost on part of capital investment along with the cost of consumables and utilities. da Gama Ferreira et al. ( 2018 ) performed sensitivity analyses of process scale, inoculation volume with respect to volumetric productivity, which suggested that remarkable reduction in cost of enzyme production may be observed under optimized conditions. Thus, based on the above studies, we can suggest that xylanase production can be based on the cost of substrate and consumable, along with the cost of each step involved in upstream and downstream processing. Therefore, utilizing cheap raw materials, less number of steps during upstream and downstream process (such as single step purification instead of multistep process) can help in keeping the enzyme production cost as low as possible.

Xylanase employed as a greener tool in different industries

Xylanase with such unique characteristics of thermo-alkali tolerant nature has a diverse range of application in different industries such as paper and pulp, deinking, biomass utilization and food feed industries (Fig. 6 ).

Xylanase as a greener tool in different industries

Xylanase employed in the food and feed industry

The xylanase finds application in food industries such as bakery. The bread is made up of wheat consisting of hemicelluloses such as arabinoxylan. The xylanase can solubilize the water unextractable arabinoxylan into water-extractable arabinoxylan (Courtin and Delcour 2002 ). This help in uniform water distribution and improvement in gluten network formation throughout the dough. The addition of xylanase improves the rheological properties of dough such as softness, extensibility, and elasticity along with bread-specific volume and crumb firmness (Harbak and Thygesen 2002 ; Camacho and Aguilar 2003 ; Butt et al. 2008 ). The breakdown products of arabinoxylan, i.e., arabino-xylooligosaccharides in bread have its health benefits (Polizeli et al. 2005 ; Bajpai 2014 ).

Butt et al. ( 2008 ) demonstrated the role of GH11 endoxylanases from B. subtilis in solubilizing the arabinoxylan. This increases the viscosity and volume of dough and decreases gluten agglomeration and dough firmness resulting in the development of uniform and fine crumbs. GH11 xylanase (0.12 U/g flour) from Penicillium occitanis Pol6 resulted in improvement of bread-making process such as the decrease in water absorption (8%) and an increase in dough rising (36.8%), volume (17.8%), specific volume (34.9%) and cohesiveness. The bread has improved rheological and sensory properties (texture, taste, flavor, softness, and overall acceptability). Low springiness and gumminess were observed in the bread prepared using xylanase (Driss et al. 2013 ). Partially purified microbial xylanase was used by Ghoshal et al. ( 2013 ) to produce whole‐wheat bread with better sensory properties (brighter color). The addition of xylanase also resulted in increased specific volume, and shelf life, with lower firmness and reduced staling during storage. Panzea, new generation xylanase obtained from Bacillus Licheniformis , can help in improving dough properties at low enzyme dosage. It helps in achieving the desired texture, appearance, loaf volume and crumb structure (Bajpai 2014 ). Similarly, recombinant xylanase (r-XynBS27) obtained from Pichia pastoris ( xynBS27 gene from Streptomyces sp. S27) used as an additive during bread-making process. The recombinant xylanase resulted in improvement in a specific volume and reducing sugar content with a decrease in firmness, consistency, and stiffness (De Queiroz Brito Cunha et al. 2018a , b ).

Fruit juice clarification

The enzymatic process in fruit juice extraction and clarification is widely used. Raw juices of fruit contain polysaccharides such as cellulose, hemicellulose, starch pectin and surface-bound lignin and decrease the quality of the juice, e.g., hazy color and high viscosity (Danalache et al. 2018 ). The use of enzymes decreases the viscosity and avoids the formation of clusters, by removing the suspended and undissolved solid using centrifugation and filtration methods. This increases the clarity, aroma, and color of the juice (Danalache et al. 2018 ). Xylanase from Streptomyces sp was used for the clarification of orange, mousambi, and pineapple with 20.9%, 23.6% and 27.9% clarity, respectively (Rosmine et al. 2017 ). Immobilized xylanase obtained from Bacillus pumilus VLK-1 was used for orange (29%) and grape juice (26%) enrichment (Kumar et al. 2014 ). Xylanase immobilized on 1,3,5-triazine-functionalized silica-encapsulated magnetic nanoparticles was reported to clarify the three different types of fruit juices after five hours of incubation at 50 °C (Shahrestani et al. 2016 ). Partially purified xylanase from Streptomyces sp AOA40 was used in fruit juice industry for increased clarity of juices from apple (17.8%), orange (18.4%) and grape (17.9%) (Adigüzel and Tunçer 2016 ). Glutaraldehyde-activated immobilized xylanase was used for the clarification of tomato juice. Xylanase from P. acidilactici GC25 was used to treat the kiwi, apple, peach, orange, apricot, grape, and pomegranate in which increase in the amount of reducing sugar was observed along with the decrease in turbidity of the juice (Adiguzel et al. 2019 ).

Animal feed

Xylanases plays an important role in animal feed by breaking the feed ingredient arabinoxylan and reduces the raw material viscosity. Aspergillus japonicus C03 with good endoxylanase and cellulase production ability with high stability in the presence of goat ruminal environment showed ruminant feed applications (Facchini et al. 2011 ). A number of studies reported the availability of distillers dried grains with soluble (DDGS) to be utilized in animal feeds and use of exogenous xylanase in poultry diets to treat the higher fiber content (Pirgozliev et al. 2016 ; Whiting et al. 2019 ). The exogenous enzymes effectively improved the nutritional value of co-products of bioethanol as reported previously with DDGS obtained from corn (Liu et al. 2011 ). Xylanases have been involved in animal feed over decades, as it reduces the viscosity of digesta in poultry. Xylanase addition showed the weight gain improvement and enhanced feed conversion ratio because of the improvement in the arabinoxylan digestibility in monogastric animal diets (Paloheimo et al. 2010 ; Van Dorn et al. 2018 ). Xylanase utilized as a dietary supplement for the nutrients digestibility, digesta viscosity growing pigs fed corn intestinal morphology diet based on soybean meal was reported by Passos et al. ( 2015 ). ECONASE XT a well-known commercial endo -1,4-β-xylanase which has been used as feed additives for chicken fattening, weaned piglets and fattening for pigs (Rychen et al. 2018 ).

Xylanase in paper and pulp industries

Bio-bleaching.

The process of removal of lignin from wood pulp to produce bright and completely white finished paper is known as bleaching (Beg et al. 2001 ). Traditionally, chemical bleaching agents (such as chlorine) were used for bleaching (Subramaniyan and Prema 2002 ). The use of ligno-hemicellulolytic enzymes for bleaching has gained impetuous all over the world. Xylanases are capable of hydrolyzing xylan which is linked to the cellulose and lignin of the pulp fiber. Thus, xylan disruption will eventually lead to the separation between these components, enhance swelling in the fiber wall, and improve lignin extraction from the pulp (Thomas et al. 2015 ). Thus, xylanase in combination with lignin-degrading enzyme help in increasing the brightness of pulp (Viikari et al. 1994 ; Sunna and Antranikian 1997 ; Pérez et al. 2002 ; Motta et al. 2013 ). The exposures of the cellulose fiber to enzymatic pulping enhance the bonding forces of paper and improve paper strength via degradation of xylan and removal of lignin during enzymatic treatment (Lin et al. 2018 ). The enzymatic system has been highly selective, non-toxic, environmentally friendly approach for bio-bleaching (Bajpai 2012 ).

Paper and biomass pulp processing takes place at varying pH and temperature. Therefore, thermo-alkali stable xylanases are required for the bio-bleaching. An alkaliphilic Bacillus strain produced thermoactive cellulase-free xylanase using agro-residues active at high temperature 60 °C and pH 6–10 and was utilized for bio-bleaching of kraft pulp (Azeri et al. 2010 ). Paenibacillus campinasensis BL11 xylanase pretreatments showed the increased brightness and viscosity of hardwood kraft pulp (Ko et al. 2010 ). S. thermophilum xylanase active at high temperature (50–70 °C) was used for the bleaching of bagasse pulp (Joshi and Khare 2011 ). T. lanuginosus VAPS24 xylanase was stable at wide range pH that can be useful in both alkaline and acidic bioprocesses (Kumar and Shukla 2018 ). An alkaliphilic Bacillus liceniformis Alk-1 xylanase was utilized in a purified form for the enzymatic pretreatment on eucalyptus kraft pulp bleaching (Raj et al. 2018 ). The xylanase preparation obtained from white-rot fungi, S. commune ARC-11 was capable of ethanol soda pulp pre-bleaching from Eulaliopsisbinata (Gautam et al. 2018 ). The paper manufacturing units of various countries, i.e., Japan. South America, North America and Europe are slowly replacing chemical pulp bleaching by xylanase mediated pulp bleaching. Canada is known to be the leading producer of pulp and they are bleaching more than 10% its pulp via xylanase (Dhiman et al. 2008a ). In addition to that, several reports also suggest that the xylanase enzyme-mediated pretreatment can help in generation of cellulosic nanofibres (CNF) with improved crystallinity from unbleached bagasse and eucalyptus pulp (Nie et al. 2018 ; Zhang et al. 2018 ; Tao et al. 2019 ). Zhang et al. ( 2018 ) suggested that application of commercial Novozyme X2753 can simplify the CNF’s production and purification process. Tao et al. ( 2019 ) demonstrated that xylanase can directly act on the unbleached pulp, where it acts on the covalent bond between hemicellulose molecule and hydrogen bond between hemicelluloses and cellulose. The presence of a small amount of hemicelluloses in cellulose nanofibrils increases light blockage efficiency and subsequently the energy storage capacity of solar cells. Thus, xylanase-mediated bio-bleached pulp acts as a potential substrate for flexible solar cells.

Deinking of waste paper

The dislodgement of ink from the waste used paper is required for its recycling and reuse. Chemical-based methods involving chlorine or chlorine-based derivatives, ClO − , NaOH, NaCO 3 , H 2 O 2 , Na 2 SiO 2 , have been used for removing ink from the paper. This resulted in generation of hazardous effluents and required tedious treatment before disposal to the environment (Maity et al. 2012 ). The enzyme-based methods utilizing xylanase and laccase have been suggested for the removal of ink from paper and pulp industries effluents (Chandra and Singh 2012 ; Dhiman et al. 2014 ).

Virk et al. ( 2013 ) explored the deinking efficiencies of bacterial alkalophilic laccase and xylanase along with physical deinking methods such as microwave and ultrasound for recycling of waste paper. The combination of xylanase and laccase enzymes showed an increase in brightness of different waste pulp old newsprint pulp (21.6%), inkjet print pulp (4.1%), laser print pulp (3.1%), magazine pulp (8.3%), and xerox paper pulp (1.9%) only. Gupta et al. ( 2015 ) reported that synergistic action of xylanase and laccase enzyme (co-cultivation of Bacillus sp. and B. halodurans FNP135) resulted in improvement of physical properties like freeness, breaking length, burst factor and tear factor by 17.8%, 34.8%, 2.77%, and 2.4%, respectively, of old newspaper. The appearance of the newspaper was also improved with an increase in 11.8% brightness and 39% whiteness. The effective dose of commercial cellulase and xylanase from Bacillus halodurans TSEV1 for removal of ink was determined at 1.2. U/mg (each enzyme) by Kumar and Satyanarayana ( 2014 ). The cellulase and xylanase complex obtained from Escherichia coli SD5 facilitated the reduction in hexenuronic acid (Hex A) and kappa number, increase in brightness (10%) and tear strength of recycled paper (Kumar et al. 2018c ).

Xylanase employed in textile industries

The textile processing can be broadly divided into desizing, scouring and bleaching. Desizing involves removal of adhesive material from plant fibers and scouring to remove the inhibitory material from desized fibers (Hartzell et al. 1998 ; Dhiman et al. 2008b ). The conventional method used for desizing and scouring involves the application of high temperature under the influence of oxidizing agents in the alkaline system. This method is not only chemical intensive but also non-specific that causes hamper to the useful cellulosic fractions compromising the overall strength of the textile fibers. Therefore, application of highly thermo-alkali stable cellulase-free xylanolytic enzyme can efficiently be used for desizing and scouring (Csiszár et al. 2001 ; Losonczi et al. 2005 ; Dhiman et al. 2008b ; Bajpai 2014 ).

Dhiman et al. ( 2008b ) demonstrated the application of alkalo-thermophilic xylanase from Bacillus stearothermophilus SDX for processing of cotton and microply fabrics. The desizing and bioscouring treatments were performed using 5 g/L of xylanase at 70 °C, pH 9.5, for 90 min. This resulted in weight loss for 0.91% in microply and 0.83% in cotton with overall whiteness index of 11.81% for cotton and 52.15% for micropoly. The processed fabric has increased tensile strength (1.1–1.2%) and tearness value (1.6–2.4%) as compared to control.

Garg et al. ( 2013 ) demonstrated the application of alkalo-thermostable xylanase from Bacillus pumilus ASH in bioscouring of jute fabric. The oven-dried jute fabric when incubated with a small dose of 5 IU/g xylanase at 55 °C for 2 h resulted in an increase in 4.3% whiteness and 10.7% brightness of fabric. Further, it also helped in decreasing in yellowness of fabric by 5.57%. Similarly, xylanase from Bacillus pumilus was studied for enzymatic desizing of cotton and micropoly fabrics (Battan et al. 2012 ). The enzymatic desizing with enzyme load of 5 IU/g at pH 7.0, temperature 60 °C for 90 min resulted in improved whiteness of 0.9% with respect to the chemical process. The addition of surfactant such as EDTA improved the desizing and bioscouring efficiency (Losonczi et al. 2005 ; Battan et al. 2012 ; Garg et al. 2013 ).

The synergistic action of xylanase and pectinase enzyme was used for scouring of cotton fabrics. The bioscouring was performed with 5.0 IU xylanase and 4.0 IU pectinase from Bacillus pumilus  strain AJK (MTCC 10414) along with surfactants such as 1.0 mM EDTA and 1% Tween-80 at high pH 8.5 for 1 h at 50 °C. They observed improvement in whiteness, brightness, and reduction in yellowness by 1.2%, 3.2%, and 4.2% respectively that is better in comparison to chemical-based alkaline scouring method (Singh et al. 2018 ). El et al. ( 2018 ) reported improvement in desizing, bioscouring and bio-finishing efficiency using xylanase obtained from T. longibrachiatum KT693225 without any requirement of additives.

Xylanase employed in chemical and pharmaceutical industries

The non-digestible sugar molecules together form oligomers known as xylooligosaccharides, which are made up of xylose monomers (Vazquez et al. 2000 ). XOS has various applications in biotechnology, pharmaceutical, food and feed industries (Chang et al. 2017 ). XOS plays a vital role as prebiotic as it is not hydrolyzed or absorbed in the gastrointestinal tract. Thus, XOS selectively stimulates the growth of important gastrointestinal microorganisms regulating the human digestive health (Roberfroid 1997 ; Collins and Gibson 1999 ; Vazquez et al. 2000 ). The potential of XOS as an efficient feed alternative is established by the fact that it help in cholesterol reduction, inhibit starch retro-gradation, improve the bioavailability of calcium thus improving the nutritional and sensory properties of food (Voragen 1998 ; Motta et al. 2013 ). XOS has shown the application in pharmaceutical sectors due to its immunomodulatory (Chen et al. 2012 ), anti-cancerous (Gupta et al. 2018 ), anti-microbial, antioxidant (Kallel et al. 2015b ), anti-allergy, anti-inflammatory (Aachary and Prapulla 2011 ), and anti-hyperlipidemic activity (Li et al. 2010 ). XOS have also shown phyto-pharmaceutical and feed applications such as growth regulatory activity in aquaculture and poultry. These properties may be due to the presence of uronic substituents in acidic oligosaccharides.

The process of XOS synthesis involves the physical (autohydrolysis), chemical (hydronium ions generated by water autoionization and in situ organic acids) or enzymatic hydrolysis (xylanase or β-xylosidase) of hemicellulose-rich agricultural wastes (Aachary and Prapulla 2011 ). Several reports suggested that XOS can be enzymatically produced from different agro-residues such as hardwoods (Huang et al. 2016 ), straws (Gullón et al. 2008 ; Kallel et al. 2015a ; Moniz et al. 2016 ) corn cobs (Chapla et al. 2013 ; Gowdhaman and Ponnusami 2015 ), bran (Otieno and Ahring 2012 ), sugarcane bagasse (Jayapal et al. 2013 ) and bamboo (Xiao et al. 2013 ) using microbial xylanases.

Alkaline xylanase from Bacillus mojavensis A21 utilized corncob xylan for the release of xylotriose and xylobiose (Haddar et al. 2012 ). Bacillus aerophilus KGJ2 xylanase showed efficiency toward XOS synthesis, e.g., xylobiose, xylotriose, and xylose after hydrolysis of xylan (Gowdhaman et al. 2014 ). A cellulase free xylanase (EX624) from Streptomyces sp. CS624 produced xylose, xylobiose and xylotriose with commercial beech wood xylan and wheat bran (Mander et al. 2014 ). Using deoiled Jatropha curcas seed cake as substrate, Sporotrichum thermophile xylanase was produced which showed the efficiency to produce XOS by the hydrolysis of oat spelt xylan, with 73% xylotetraose, 15.4% xylotriose and 10% xylobiose (Sadaf and Khare 2014 ). Xylanase obtained from the mixed microbial culture of Cellulomonas uda NCIM 2523 and Acetobacter xylinum NCIM 2526 using Prosopis juliflora showed the potential to produce XOS with probiotic activity from beech wood xylan (Anthony et al. 2016 ). A xylanase gene PbXyn10A isolated from Paenibacillus barengoltaii cloned in E. coli showed 75% XOS yield from xylan extracted from raw corncobs (Liu et al. 2018 ). The hydrolysis of xylan using xylanase from Pichia stipitis produced 2% XOS consisting of xylotetraose 14%, xylotriose 49% and xylobiose 29% (Ding et al. 2018 ). Bhardwaj et al. ( 2019 ) demonstrated the partially purified xylanase obtained from Aspergillus oryzae LC1 resulted in the generation of xylobiose, xylotriose, and xylotetraose.

Xylanase employed in biorefinery

Efficient conversion of lignocellulosic biomass (LCB) into fuel-grade ethanol has become a world priority for producing environmentally friendly renewable energy at a reasonable price for the transportation sector. The process of bioconversion of lignocellulosic biomass requires hydrolysis of the pretreated biomass for the conversion of complex carbohydrate polymer of LCB to the simple monomers which will be further converted to ethanol by fermentation. The xylanolytic enzyme in combination with cellulolytic enzyme plays an important role in the hydrolysis process.

Several reports suggest that xylanase obtained from several microorganisms plays an important role in saccharification of LCB for lignocellulosic-based biorefinery (Hu et al. 2011 ; Choudhary et al. 2014 ; Ramanjaneyulu et al. 2017 ; Basit et al. 2018 ). Hydrolysis and fermentation are important steps in biomass to bioethanol generation. Initially, several groups demonstrated separate hydrolysis of biomass followed by fermentation (SHF). SHF is a time-consuming process and thus increases the overall cost of the process. Later on, different integrated process (combined hydrolysis and fermentation) have been developed such as simultaneous saccharification and co-fermentation (SSCF), simultaneous saccharification and fermentation (SSF), and consolidated bioprocessing (CB) (Malhotra and Chapadgaonkar 2018 ). These strategies resulted in an enhancement in reaction rates and ethanol yields (Eklund and Zacchi 1995 ; Sun and Cheng 2002 ). Bibra et al. ( 2018 ) showed thermostable xylanase production using Geobacillus sp. DUSELR13, which is applied further for ethanol generation from LCB. SSF Geobacillus sp. DUSELR13 and Geobacillus thermoglucosidasius are co-cultured for SSF of prairie cord grass (PCG), and corn stover (CS). The SSF resulted in 3.53 and 3.72 g/L ethanol from PCG and CS, respectively.

Hu et al. ( 2011 ) suggested that xylanase causes fiber swelling improving porosity that helps in improving the accessibility of cellulose. To ferment both cellulose-derived hexoses (C6) and xylan-derived pentoses (C5), simultaneous saccharification and co-fermentation (SSCF) was introduced which causes ethanol production using single microorganisms co-cultured with cellulase and xylanase producing strain. Yasuda et al. ( 2014 ) demonstrated bioethanol generation by SSCF of anhydrous ammonia-pretreated  Pennisetum purpureum Schumach (Napier grass) using Escherichia coli KO11 and Saccharomyces cerevisiae cellulase, and xylanase. SSCF for 96 h was reported to have a maximum 74% ethanol yield as compared to theoretical yield calculated based on glucan and xylan yield of 397 mg/g and 214 mg/g, respectively.

Bondesson and Galbe ( 2016 ) designed experimental setup of SSCF for ethanol production from steam-pretreated, acetic acid-impregnated wheat straw using a pentose fermenting S. cerevisiae KE6-12b strain. The highest 37.5 g/L ethanol concentration with 0.32 g/g ethanol yield was obtained based on the glucose–xylose available in the pretreated wheat straw. Shariq and Sohail ( 2018 ) demonstrated that yeast strain Candida tropicalis MK-160 can help in xylanase and endoglucanase production as well as ethanol production. Therefore, it can be potentially used for SSCF involves single microorganism.

The consolidated processing or simultaneous delignification, saccharification, and fermentation involve the cultivation of ligno-hemicellulolytic enzyme-producing strains along with ethanol-producing strain in a reactor. It may be monoculture or co-culture of different microorganisms. It will help to decrease the overall process cost required for bioreactor and operation of different enzyme production and ethanol generation (Chadha et al. 1995 ).

To design a monoculture-based consolidated processing, different engineered microorganisms are used having ligno-hemicellulolytic enzymes having the capability along with ethanol generating potential. Shen et al. ( 2012 ) engineered a thermostable self-splicing bacterial intein-modified xylanase for consolidated lignocellulosic biomass processing. Sun et al. ( 2012 ) demonstrated expression of recombinant Saccharomyces cerevisiae strain having an engineered minihemicellulosome. It has the capability of converting xylan directly to ethanol. Horisawa et al. ( 2019 ) suggested direct ethanol production from lignocellulosic materials by consolidated bioprocessing using the mixed culture of wood rot fungi, i.e., Schizophyllum commune, Bjerkandera adusta , and Fomitopsis palustris .

Commercial xylanase enzyme and their application

The xylanase enzyme is commercially employed in several industries such as pulp bleaching, food, feed, and brewing. The major application of the xylanase is in pulp bleaching and is produced by different companies around the world with various trade name such as Bleachzyme (Biocon, India), Cartzyme (Sandoz, US), Cartzyme MP (Clarient, UK), Ecozyme (Thomas Swan, UK), Irgazyme 10 A & Irgazyme 40–4× (Genercor, Finland), Ecopulp (Alko Rajamaki, Finland), VAI Xylanase (Voest Alpine, Austria), and Rholase 7118 (Rohm, Germany). Xylanase is commercially produced by several international industries for its application in food and feed industries. Sankyo from Japan, Ciba Giegy from Switzerland produces xylanase as trade name Sanzyme, and Albazyme-10A, respectively, and has been used commercially in food and baking industries. A Danish firm Novo Nordisk, produces three commercial xylanases namely Pulpzyme (HA, HB, HC), Biofeed (Beta, plus) and Ceremix and used in pulp bleaching, feed and brewing industries, respectively (Walia et al. 2017 ). An American firm Alltech, Inc., commercially produces xylanase with trade name Allzym PT and Fibrozyme and has been used for the upgrading the animal feed. A Japanese firm named Amano Pharmaceutical Co, Ltd. produced xylanase enzyme named Amano 90 and it has been used in food, feed and pharmaceutical industries. Most of the commercial xylanases are produced by fungal source due to its high production potential.

Challenges and future trends in commercial production, purification and application of xylanase

The search of super xylanase is still on, therefore, searching for new microbial source with the ability to produce highly active and stable xylanase is going on around the world. The strains isolated from different extreme habitat can be of potential applications as these strains already possesses the ability to withstand different stress such as high temperature and pH variations. The selection of such thermal and pH tolerant strains and subjecting them to different optimization strategies for enhanced xylanase production can be one alternative. The advancement in biotechnological tools and techniques (Recombinant DNA technology or genetic engineering) provides an opportunity to select the gene responsible for xylanase production that can be isolated and efficiently transferred to the expression system. These expression systems can be regulated for enhanced production of xylanase with desired property for specific industrial applications. The availability of a high amount of genomics, proteomics and metabolomics data can be used via different bioinformatics tools to develop different approaches for enhanced xylanase production. The combination of new technology such as synthetic biology (DNA oligo-synthesis) and conventional recombinant DNA technology can be used for attaining the objective of high xylanase production with desired industrial properties (Fig.  7 ). However, limitations associated with mimicking the natural system into synthetic system need to be taken care before full-scale applications. Also, the ethical, socio-economic and health concerns need to taken care before commercial exploitation of the developed strategies.

Future prospect for development in area of xylanase production using conventional and advanced approaches

The enzymatic breakdown of the xylan into its constituent component requires the synergistic action of xylanases and other debranching enzymes. The key enzyme used for xylan hydrolysis is endo -1,4-β-xylanase that cleaves β-1,4-glycoside linkage of xylan. The xylanase can be grouped under different GH families with major xylanases from GH10 or GH11 families, followed by GH5, GH7, GH8 and GH43. The xylanase enzyme acts as a “Green” alternative to already existing industrial processes for processing of xylan to different industrially important product such as paper, textile, food, feed, pharmaceuticals, and biofuels. The application of xylanases in the production of the above-mentioned products can regulate the overall economics of the process. Therefore, already existing method can be further improved or new strategies may be developed for enhanced and cost-efficient production of xylanase with desired characteristics. Further, it is often observed that native enzyme cannot meet the industrial process requirement; thus, the combination of already existing technology with new technology such as synthetic biology (DNA oligo-synthesis), rational engineering and directed evolution can be used for attaining the objective of high xylanase production with desired industrial properties.

Availability of data and materials

Not applicable.

Abbreviations

lignocellulosic biomass

enzyme commission

high-molecular weight with low isoelectric point

low-molecular weight with high isoelectric point

carbohydrate-active enzyme database

National Centre for Biotechnological Information

glycoside hydrolase

xylooligosaccharides

xylotetraose

catalytic nucleophile/base

catalystic proton donor

aspartic acid

gluctamic acid

submerged fermentation

solid-state fermentation

carboxymethyl cellulose

one factor at a time

central composite design

analysis of variance

response surface methodology

generally regarded as safe

trichloroacetic acid

sodium dodecyl sulfate-polyacrylamide gel electrophoresis

diethylaminoethyl

fast protein liquid chromatography

aqueous two-phase system

polyethylene glycol

distillers dried grains with soluble

simultaneous saccharification and co-fermentation

simultaneous saccharification and fermentation

consolidated bioprocessing

prairie cord grass

corn stover

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Acknowledgements

The authors are thankful to the Department of Biotechnology, Government of India for providing the financial support (Grant Nos. BT/304/NE/TBP/2012 and BT/PR7333/PBD/26/373/2012). NB is thankful to the University Grants Commission for providing fellowship for doctoral studies. BK is thankful to Jawaharlal Nehru memorial Fund and CSIR for providing scholarship for doctoral studies.

The authors thank Department of Biotechnology, Government of India (Grant Nos. BT/304/NE/TBP/2012 and BT/PR7333/PBD/26/373/2012).

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Nisha Bhardwaj, Bikash Kumar & Pradeep Verma

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Bhardwaj, N., Kumar, B. & Verma, P. A detailed overview of xylanases: an emerging biomolecule for current and future prospective. Bioresour. Bioprocess. 6 , 40 (2019). https://doi.org/10.1186/s40643-019-0276-2

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Optimization, purification, and characterization of xylanase production by a newly isolated Trichoderma harzianum strain by a two-step statistical experimental design strategy

• Priyashini Dhaver 1 ,
• Brett Pletschke 2 ,
• Bruce Sithole 3 , 4 &
• Roshini Govinden 1  

Scientific Reports volume  12 , Article number:  17791 ( 2022 ) Cite this article

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Xylanases are hydrolytic enzymes with a wide range of applications in several industries such as biofuels, paper and pulp, food, and feed. The objective of this study was to optimize the culture conditions and medium components for maximal xylanase production from a newly isolated Trichoderma harzianum strain using the Plackett–Burman Design (PBD) and Box Behnken Design (BBD) experimental strategies. Xylanase production was enhanced 4.16-fold to 153.80 U/ml by BBD compared to a preliminary one-factor-at-a-time (OFAT) activity of 37.01 U/ml and 2.24-fold compared to the PBD (68.70 U/ml). The optimal conditions for xylanase production were: 6 days of fermentation, incubation temperature of 70 °C, pH 5.0, agitation of 160 rpm, and 1.2% wheat bran and ammonium sulphate. The experimental design effectively provided conditions for the production of an acidic-thermostable enzyme with exciting potential for application in animal feed improvement. The acidic-thermostable xylanase was purified from the submerged culture and SDS-PAGE analysis revealed a molecular weight of 72 kDa. This protein had maximum xylanolytic activity at pH 6.0 and 65 °C and was stable for 4 h retaining > 70% activity and exhibited substrate specificity for beechwood xylan with a K m of 5.56 mg/ml and V max of 1052.63 µmol/min/mg. Enzyme activity was enhanced by Fe 2+ , Mg 2+ , and Zn 2+ . There was an absence of strong inhibitors of xylanase activity. Overall, these characteristics indicate the potential for at least two industrial applications.

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Introduction.

After cellulose, hemicellulose is the second most abundant terrestrial polysaccharide composed of ß-1,4- d -xylopyranoside residues and ß-1,4-xylan as main constituents with arabinosyl and acetyl side chains 1 . Hemicellulose is a short crosslinked polymer compared to cellulose, which is a long straight-chain homopolymer. Xylans have a β-(1,4) linked backbone made of d -xylose and there are three subtypes of xylan based on the side chain. The subtypes are homoxylan, glucuronoxylan, and arabinoxylan. Homoxylan is only found in two or three types of plants and is mostly cross-linked by β-(1,4)-glycosidic bonds. Xylan is a renewable biomass resource that has potential as a substrate in many production processes. However, it must be hydrolysed to xylose and xylo-oligosaccharides which can be accomplished by xylanolytic enzymes. Among them, xylanases deserve special attention as they degrade the major hemicellulosic polysaccharides by catalyzing the hydrolysis of xylopyranosyl linkages of ß-1,4-xylan 2 . The main enzymes involved are endo-1,4-β-xylanases which make random cuts in the xylan backbone and β-xylosidases which are exoglycosidases with the ability to degrade the non-reducing ends of xylooligosaccharides into xylose. The side groups in xylans are cleaved by α- l -arabinofuranosidases (EC 3.2.1.55), α-glucuronidases (EC 3.2.1.139), and acetyl xylan esterases (EC 3.1.1.72) 3 . The most common microbial xylanases that would hydrolyze all types of xylan are grouped based on amino acid similarities and structural characteristics, into glycoside hydrolase (GH) families 10 or 11 in the Carbohydrate-Active enzyme (CAZy) database ( http://www.cazy.org ) 4 . GH10 families are able to catalyze the hydrolysis of a wide range of xylans, while GH11 families are known to cleave unsubstituted regions of arabinoxylan 4 . However, both GH10 and GH11 xylanases have applications in various industries such as food and feed 5 , biofuel production 6 , paper and pulp 7 , and medical and pharmaceutical 8 . Xylanases are also found in GH families 30, 8, and 5. The complete degradation of heterogeneous xylan into simple sugars requires the synergistic action of several inducible hemicellulases 9 .

Microbes such as bacteria, fungi, and actinomycetes are ubiquitous in nature 10 and several endogenous and exogenous microbial enzymes have been widely explored, resulting in a variety of microorganisms commonly regarded as the most significant and convenient producers of large quantities of enzymes in a short period on inexpensive feedstock. Xylanases are produced by microbial biosynthesis for industrial applications 11 , 12 . Thermophilic fungi in particular are promising candidates for biotechnological applications due to their strong ability to degrade plant polysaccharide components and their robustness under harsh environmental conditions 2 . Many of the filamentous fungi that have been studied produce several xylanases and have high xylan-degrading ability. Trichoderma sp. and Aspergillus sp. are most frequently employed for industrial applications 9 including the bioconversion of plant biomass into animal feed 5 , plant fertilizers, and chemicals for the food industry 9 .

The production of enzymes is costly, thus, to meet industrial demand, a low-cost growth medium is required for microbial growth and enzyme production. There are two possible cultivation methods for microbial xylanase production: solid-state and submerged fermentation 13 . Submerged fermentation technology has the advantage of short production periods to achieve high yields at low costs. Both the nutrient medium composition and culture conditions have a strong influence on xylanase production. The physical and chemical factors known to influence xylanase production are temperature, pH, incubation period, carbon and nitrogen sources and concentration, and agitation speed 14 . Temperature effects on enzyme production are predominantly related to the growth of the organism (mesophilic, psychrophilic, or thermophilic). The pH is one of the most important factors governing microbial growth due to their sensitivity to the hydrogen ion concentration in the medium 15 . It is also key to enzyme activity as it can alter the ionic charges on the molecule, which could cause changes to the enzyme’s shape (they may denature), and that usually leads to a reduction or loss of the catalytic properties of the enzymes and cessation of metabolic activity 16 .

Supplementation of the growth medium with carbon and nitrogen sources usually increases enzyme production as this provides an enriched environment for microbial growth 17 . Therefore, screening of the most influential factors and optimization of the growth conditions are essential to ensure maximal enzyme production, potentially significantly reducing production costs for xylanases 18 . There are two approaches to optimize the fermentation process classical and statistical. The classical approach is based on the testing of “one-factor-at-a-time (OFAT)” and the statistical approach includes the Plackett–Burman design (PBD) and response surface methodologies (RSM).

The OFAT approach is a conventional single-dimensional investigation that involves changing one independent variable at a time while the others remain at their optimal level 19 , 20 . This is the main strategy used for selecting optimal conditions, which continues to be widely used in preliminary optimization studies 21 . The main disadvantages of OFAT are the partial explanation regarding the effect of the factors on the response and the absence of the interaction effects between the variables 22 . This method also involves a relatively high number of experiments, which makes it laborious and time-consuming 21 . Moreover, it may lead to unreliable results and inaccurate conclusions.

To resolve this problem, optimization studies can be carried out by using multivariate statistical methods 21 , PBD and RSM can potentially eliminate the limitations of the OFAT optimization process 23 . PBD is a powerful statistical technique for screening medium components in shake flask fermentation and reduces the total number of experiments 24 . This technique is useful and has been widely used as the first step of an optimization procedure, however, it cannot determine the interaction effects 23 but allows the evaluation of the importance of each factor in moderately few experiments 25 . RSM using a Box–Behnken Design (BBD) is an effective optimization tool. The RSM design can provide the dependence of enzyme production on independent variables, predicted results for responses, and levels for independent variables in the form of mathematical models 26 , 27 .

Hydrolysis of xylan and hemicellulosic materials to various xylooligosaccharides has been accomplished using crude xylanases. However, to meet the desired requirements of some applications, robust xylanases (resistant to metal ions and chemicals, displaying pH, and thermostability) with specific biochemical properties for pH and temperature optima as well as, high specific activity are required, which would require purification of appropriate candidate enzymes 28 . Purification of xylanases is necessary to remove contamination by proteins and other enzymes in the culture medium, such as cellulases, as well as compounds derived from hydrolysis of the substrate. These contaminants can complicate activity assays, protein quantification, and physicochemical assays 29 . Purification and physicochemical characterization (activity and stability at various pH and temperatures) of pure xylanases provide information on the enzyme's structural and functional features, which may be used to assess its application potential 29 . Purification should be centred on attaining the highest yield while retaining the highest possible enzymatic activity and purity 30 .

Xylanases are required in large quantities for industrial-level applications because they not only have application in several processes but possess the necessary characteristics to withstand harsh conditions during these industrial processes. As a result, there is a need to select microorganisms that produce high levels of xylanases with appropriate properties, followed by optimization of growth which would lead to higher levels of enzymes 19 . There are several reports available on the optimization of media components and the physical growth parameters for the production and purification of xylanases for various applications using different substrates 8 , 12 , 21 . Xylanase production by Trichoderma reesei SAF3 31 and Trichoderma stromaticum AM7 32 was increased by optimization. While there are reports on multiple other species, there were very few reports on thermostable Trichoderma harzianum xylanases in literature.

Therefore, in the present study, a recently isolated and characterized fungal strain, T. harzianum , producing a thermophilic and acidic xylanase was the subject of study. The major focus of this study was to employ statistical design strategies to optimize xylanase production. Although this is not a novel approach, its application to a novel thermophilic and acidic xylanase is. We also report on the purification and characterization of the T. harzianum xylanase, to determine its applicability for future studies in animal feed improvement.

Materials and methods

Microbial strain.

The T. harzianum strain was selected from a previous screening study as the candidate for xylanase production 33 . Fungal cultures were streaked on the PDA plates and slants, grown for 5 days at 30 °C followed by the addition of sterile mineral oil to cover the fungal mycelium and storage at 4 °C. Long-term stocks were prepared by washing fungal spores from the 5-day PDA plates with distilled water and adding 50% glycerol in a 1:1 ratio to the spore suspension and storing at − 20 °C and − 80 °C.

Medium and cultivation

Nutrient salt solution (NSS) used for xylanase production comprised [(g/L): (0.005 g) CaCl, (0.23 g) KH 2 PO 4 , (0.05 g) MgSO 4 , (0.005 g) NaNO 3 , (0.002 g) ZnSO 4 , (0.009 g) FeSO 4 , (0.23 g) KCl, (7 g) peptone, and (20 g) wheat bran]. Erlenmeyer flasks (250 ml) containing 50 ml of the medium were each inoculated with two 5 mm fungal plugs from a 5-day-old plate culture and incubated at 30 °C at 200 rpm for 7 days in a shaker (New Brunswick Innova 44, Germany). Cultured media were removed after the incubation period and the cell-free supernatant was recovered by centrifuging samples at 16,873 ×  g for 10 min (Eppendorf centrifuge 5418, Germany). Xylanase activity was determined as described below in the xylanase assay method (“ Xylanase assay ”).

Xylanase assay

Xylanase activity was quantified using the 3,5-dintrosalicylic acid (DNS) assay for reducing sugars according to the method by Miller 34 . The reaction included 600 µl of 1% (w/v) of beechwood xylan (1 g in 100 ml of 50 mM citrate buffer pH 5.0) in 15 ml test tubes to which 66.67 µl of the culture supernatant was added. The reaction mixture was incubated in a water bath at 55 °C for 15 min and terminated by the addition of 1 ml DNS reagent followed by heating for 5 min at 100 °C in a water bath. One unit (U) of xylanase was defined as the amount of enzyme that released 1 µmol xylose as reducing sugar equivalents per min under the specified assay conditions.

Optimization of xylanase production: one factor at a time (OFAT)

To optimize the growth parameters, OFAT was used to evaluate the effect of a single parameter at a time performed in earlier study 33 . The enzyme activity was obtained to determine the optimal yield and was reported in previous studies 33 .

Statistical optimization, experimental design, and data analysis

Plackett–burman design (pbd).

Six variables were selected for this study: Incubation temperature (X 1 ), Incubation time (X 2 ), pH (X 3 ), Agitation (X 4 ), Wheat bran (X 5 ), and Ammonium sulphate (X 6 ) (Table 1 ). The total number of experimental runs carried out for the six variables was twelve 35 . Each variable was represented by a high level denoted by ‘+’ and a low level denoted by ‘−’. The high level of each variable was sufficiently far from the low level so that any significant effect would be observed. The experimental runs were performed in duplicate and an average of the results was reported Table 2 represents the PBD based on the first-order polynomial model Eq. ( 1 ):

where Y is defined as the response (peak area and retention factor), β 0 is the model intercept, β i is the linear coefficient and Χ i is the level of the independent variable. The PBD was analyzed using R studio software 36 to estimate the significant factors. ANOVA was performed to determine the p -values as well as the R coefficients to check the significance and fit of the regression model. Screened parameters were represented on a Pareto chart of standardized effects. The effect of each variable was analyzed and the ones with the highest influence on the production of xylanase were selected for the second level optimization by BBD of RSM.

Optimization of the significant variables using response surface methodology (RSM)

The BBD was used to elucidate the main interaction and quadratic effects of the three significant variables arising from the PBD, with replicated centre points 21 . The experimental design and statistical analysis were performed using R Studio 36 . A three-level three-factor BBD was used to evaluate the combined effect of the three significant independent variables, Incubation time (X 2 ), pH (X 3 ), and wheat bran (X 5 ) (Table 3 ). The design consisted of 16 combinations, including three replicates of the centre point as shown in Table 4 . After the experimental runs were completed, the average xylanase activities were taken as the response (Y). A multiple regression analysis of the data was carried out to obtain an empirical model that relates the response measured to the independent variables 37 . The second-order polynomial Eq. ( 2 ) is shown below:

where Y represents the response variable (peak area), β 0 is the interception coefficient, β 1 is the coefficient for the linear effects, β 2 is the coefficient for the quadratic effect, β 12 are interaction coefficient and Χ 1 Χ 2 is the coded independent variables that influence the response variable Y. The response in each run was the average of duplicates. In this experimental design, data were analyzed by one-way ANOVA with Tukey’s multiple comparison test (p ≤ 0.05) using R studio 36 , and ggplot2 was used for the generation of 3D response surface and contour plots 38 .

Scaled-up production in the optimized medium

The optimized parameters for each factor from the statistical design experiments were implemented for the scaled-up production of the xylanases. The nutrient salt solution was prepared as previously described (“ Medium and cultivation ”) and supplemented with the optimized wheat bran and ammonium sulphate concentrations. Erlenmeyer flasks (2 l) containing 400 ml of the medium were each inoculated with two 5 mm fungal plugs from a 5-day-old plate culture and incubated at the optimized parameters in a shaker (New Brunswick Innova 44, Germany). Cultured media were removed after the incubation period and the cell-free supernatant was recovered by centrifuging samples at 16,873 ×  g for 10 min (Eppendorf centrifuge 5418, Germany). Xylanase activity was determined as described in “ Xylanase assay ”.

Purification of xylanase

All purification steps were carried out at 4 °C. Partial purification of the xylanase was carried out by ammonium sulphate precipitation (20–80%). The pellets were dissolved in 50 mM citrate buffer pH 5.0 and subjected to dialysis overnight in the same buffer. The fraction that resulted in the highest activity, was concentrated in 3 kDa Amicon centrifugal tubes, the protein precipitate dissolved in 50 mM citrate buffer pH 5.0 buffer and loaded onto an anion exchange column (HiTrap Q FF 5 ml) which was connected to the AKTA Purifier (AKTA Purifier, GE Healthcare Bio-Science, AB75184, Uppsala Sweden). Before loading the sample, the column was equilibrated with 20 mM Tris buffer, (pH 8.0). The enzyme was eluted using a 0–2 M sodium chloride gradient at a flow rate of 1.5 ml/min. Fractions were collected and those displaying xylanase activity were pooled, concentrated, and dialyzed against a 50 mM citrate buffer (pH 5.0), to be used for further characterization of the enzyme. Protein concentration was measured by the Bradford method 39 using bovine serum albumin as the standard. The samples were separated on a 12% SDS-polyacrylamide gel according to Laemmli 40 . Native PAGE was performed with 1% xylan as the substrate. Once electrophoresis was completed, the gel was incubated in 50 mM citrate buffer pH 5.0 at the optimum temperature (70 °C) for 20 min and thereafter stained with 0.1% Congo red solution for 30 min and destained in 1 M NaCl until clearance bands representing xylanase activity were obtained.

Characterization of purified xylanase

Effect of ph and temperature on xylanase activity.

The pH optimum was determined by measuring enzyme activity between pH 4.0 and 10.0 The following buffers were used: 0.1 M sodium citrate buffer (pH 3.0–5.0), 0.1 M potassium phosphate buffer (pH 6.0–8.0) and 0.1 M Glycine NaOH buffer (pH 9.0–10.0) 41 . Enzyme assays were conducted as previously described (“ Xylanase assay ”). For determination of the optimum temperature, the reactions were carried out at the optimum pH between 40 to 80 °C with intervals of 5 °C.

pH and thermostability

The pH stability of the enzyme was determined by incubating the enzyme in the optimal pH buffer for 4 h at 55 °C with aliquots removed every 30 min. A substrate control was also incubated for 4 h. Thereafter, xylanase activity was assayed using the DNS method and reported as residual activity (%). Temperature stability was determined by incubating the enzyme in the optimal pH buffer at optimal temperature for 4 h with aliquots collected every 30 min. The activity was assayed and reported as residual activity (%).

Effect of metallic ions and different solvents on xylanase activity

The effect of metallic ions (CaCl 2 , CoCl 2 , FeSO 4 , MgSO 4 , MnSO 4 , and ZnSO 4 ) and chemical agents (SDS, DMSO, and EDTA) on enzyme activity was evaluated at two concentrations: 2 mM and 10 mM. The residual activity was measured using the standard assay conditions. The activity in the absence of metallic ions or inhibitors was taken as the control (100%) 42 .

Substrate specificity

The specificity of the purified xylanase was verified by assaying the activity using various substrates, viz., beechwood xylan, birchwood xylan, xylan from Larchwood, wheat arabinoxylan (soluble and insoluble), carboxymethylcellulose (CMC) and Avicel. Substrates (1% w/v) were suspended in 50 mM citrate buffer (pH 6.0) and incubated with the purified enzyme at 65 °C for 15 min and thereafter the xylanase activity was determined by the DNS method as described previously (“ Xylanase assay ”) 43 .

Kinetic parameters

The K m and V max values for the xylanase were determined by measuring the enzymatic activity using different concentrations of the xylan substrate (1–20 mg/ml). The activity was measured under standard assay conditions as described previously. The Michaelis-Menton and Lineweaver- Burk plots were acquired to determine K m and V max .

Equipment and settings

Neither image acquisition tools nor image processing software packages were used for the figures in this study. For Fig.  7 , processing such as changing brightness and contrast was applied equally across the entire image and applied equally to the controls. The contrast does not allow for any data to disappear. There were no excessive manipulations, such as processing to emphasize one region in the image at the expense of others.

Results and discussion

Screening of significant medium constituents for xylanase production.

The rows in Table 2 represent the twelve different experiments. The data obtained from the PBD runs indicate a wide variation in xylanase activity from 9.8 to 68.7 U/ml across the twelve runs. This variation demonstrated that the effect of the medium and culture conditions on the production of xylanase was significant (p < 0.05). The R 2 , or coefficient of determination, is the percentage of response variance that can be ascribed to the model rather than a random error 44 . According to Xie et al. 45 , R 2 should be at least 90% for a model to fit well. The determination coefficient ( R 2 ) indicates that the independent variables were responsible for 97 percent of the sample variance in xylanase output, and just roughly 3% of the overall variation was not explained by the model. The greater the correlation between experimental and anticipated values, the closer R (correlation coefficient) is to 1. The value of R (0.97) indicated that the experimental data and the theoretical values predicted by the model equation were in close agreement. As indicated in Table 5 , the p -value was used to verify the significance of each of the coefficients. The incubation period (X 2 ), pH (X 3 ), and wheat bran (X 5 ) were all shown to have a significant ( p  < 0.05) effect on xylanase activity. The Pareto chart of standardization (Fig.  1 ) confirmed that these three factors significantly influenced xylanase production ( p  < 0.05), as they crossed the p -line. However, the other independent factors ( p  > 0.05) were generally considered insignificant.

Pareto chart of standardized effects for the production of xylanase. Incubation temperature (X 1 ), incubation time (X 2 ), pH (X 3 ), agitation (X 4 ), wheat bran (X 5 ), ammonium sulphate (X 6 ).

There is a 97% chance that the model explains the measured variations in response. The magnitude and direction of the factor coefficient in the equation clarify the influence of the six variables for xylanase production. The higher magnitude indicated a large effect on the response. The corresponding response of xylanase activity was expressed in terms of the following regression Eq. ( 3 ) derived from the Unstandardized Beta values (Table 6 ):

where Y is defined as the peak area, X 1 refers to the incubation temperature, X 2 is the incubation time, X 3 is the pH, X 4 is the agitation, X 5 is the wheat bran and X 6 is the ammonium sulphate.

Optimization of significant variables for xylanase production

Box behnken design.

A total of 16 runs were performed to determine the conditions for optimal xylanase production by T. harzianu m. A matrix was run with the three significant variables that emerged from the PBD experiments. The results for the BBD runs (Table 4 ) show that the lowest activity of 27.38 U/ml was obtained under zero-level conditions (5 days, pH 5.0, and 1% wheat bran) in run 15 while run 8 resulted in the highest xylanase activity of 153.80 U/ml under the following conditions: 6 days of incubation, pH 5.0, and 1.2% wheat bran. This was significantly and markedly (over four-fold)_higher (p ≥ 0.05) than the highest enzyme activities obtained during OFAT (38.50 U/ml). Long et al. 21 confirmed a similar but lower influence of optimized parameters on xylanase production (174.46–266.70 U/ml) by Trichoderma orientalis . Using the quadratic equation, the predicted values were determined (Table 4 ). The R 2 or coefficient of determination (0.9647, close to 1) confirmed the validity of the model, i.e., that 96.47% of the variability of the response can be expressed by the model. The value of the coefficient of adjusted determination, adjusted R 2 , was 0.9112 confirming that the actual values were close to the predicted values 46 , 47 . The correlation was confirmed by plotting the actual value curve as a function of the predicted values (Fig.  2 ) which shows the points distributed around the regression line. Figure  2 shows that the actual response values agreed well with the predicted response values, thus the predicted xylanase production is within the limits of the experimental factors. Therefore, the model is considered of sufficient quality 46 with a 96.47% chance that it explains the measured variations in response.

Graphical representation of the minimal difference between the actual (straight line) and predicted responses (circles) for the Response Surface Methodology Design for optimal xylanase activity.

Maximum xylanase production (153.80 U/ml) by the T. harzianum strain occurred in BBD run 8 under acidic conditions (pH 5.0), the higher wheat bran (1.2%), and a 5-day incubation period. Lightly lower activity can be observed for run 12 (116.74 U/ml) where the incubation period was 5 days, the wheat bran was 1.2% and the pH was 6.0. Even lower but similar enzyme activities were obtained for runs 6 (101.32 U/ml) and 7 (103.37 U/ml) where either the incubation time (4 or 6 days) or wheat bran (0.8 or 1.2%) was at their low or high levels, respectively compared to run 8 where both these parameters were at their high levels (6 days and 1.2%). This may be due to the presence of two isoforms that are maximally produced under acidic conditions. The presence of isoforms requires different periods of incubation for maximal xylanase activity and various wheat bran concentrations. In the presence of xylan, most microorganisms can produce multiple types of xylanases. Fungi are well-known for producing a wide range of xylanases (up to 30 multiple forms) 5 , 48 . Zhang et al. 49 reported that three xylanase isoforms were produced by Aspergillus fumigatus . Multiple forms of xylanases differ in stability, catalytic efficiency, absorption, and activity on substrates 50 . Okafor et al. 51 isolated a strain of Penicillium chrysogenum PCL501 from wood wastes and found that after 4 days of fermentation, wheat bran produced the highest xylanase activity of 6.47 U/ml. Abdel-Sater et al. 52 obtained maximum xylanase production from T. harzianum after 8 days of fermentation whereas, Thomas et al. 53 achieved maximum xylanase production in 4 days of fermentation by an Aspergillus sp.

The production of multiple forms of xylanases can be influenced by many factors, including the presence of various alleles of the same gene, variable mRNA processing, proteolytic digestion post secretion, and post-translational modifications such as glycosylation and autoaggregation 54 . Because xylanases have varying catalytic efficiencies, the production of several xylanases is particularly beneficial for the complete hydrolysis of hemicellulosic substances 55 . Generally, xylanase production is directly proportional to the duration of the fermentation time up to a certain level and then decreases, thus, incubation time affects xylanase production by fungi 56 .

Second-order regression and prediction

The second-order regression equation provides the xylanase activity produced by the T. harzianum strain as a function of incubation time (X 2 ), pH (X 3 ), and wheat bran (X 5 ) which can be presented in the following Eq. ( 4 ):

where Y is the peak area, X 2 is the incubation time, X 3 is the pH and X 5 is the wheat bran concentration. The statistically insignificant parameters ( p  > 0.05) and their interactions were omitted from the equation. The model constants and coefficients were generated using the unstandardized beta values.

ANOVA and Pareto chart

The “Lack of fit p -value” (Table 7 ) was insignificant as the p -value was greater than 0.05, however, literature shows this p -value (> 0.05) is considered acceptable 57 . According to Bezerra et al. 58 , significant regression and a non-significant lack of fit present in the model were well-fitted to the experiments. Based on this, the regression equation can be validated 59 . ANOVA was performed to determine the p -values (Table 7 ). This showed the model, the linear and square terms for X 2 (Incubation time), and the interaction between X 3 (pH) and X 5 (Wheat bran) as well as the linear terms of X 3 (pH) to be significant as the p -values were 0.00001, 0.0001, 0.00005, 0.02042 and 0.01137, respectively. The Pareto chart of standardization histogram graph (Fig.  3 ) also showed that Incubation time (X 2 , X 2 2 ), the interaction between pH and wheat bran (X 3, X 5 ), and pH (X 3 ) was significant ( p  < 0.05), as it crosses the p -line.

Pareto chart standardized effects of nine interactive factors affecting the production of xylanase optimization. Incubation temperature (X 1 ), incubation time (X 2 ), pH (X 3 ), agitation (X 4 ), wheat bran (X 5 ), ammonium sulphate (X 6 ).

Interaction of variables

The relationship between the parameters and the responses can be understood by studying the three-dimensional (3D) response surface plots for xylanase activity generated from the quadratic model. The 3D response surface plot can also be used to determine the optimum level of each variable for xylanase activity (Figs. 4 , 5 , 6 ). While maintaining other variables at their optimal level, the Z-axis (referring to xylanase activity) versus any two variables was constructed in the response surface plot.

3D-response surface plots ( a ) and contour plots ( b ) of the combined effects of Incubation time (X 2 ) and pH (X 3 ) on xylanase production by Trichoderma harzianum strain.

3D-response surface plots ( a ) and contour plots ( b ) of the combined effects of Incubation time (X 2 ) and wheat bran (X 5 ) on xylanase production by Trichoderma harzianum strain.

3D-response surface plots ( a ) and contour plots ( b ) of the combined effects of pH (X 3 ) and wheat bran (X 5 ) on xylanase production by Trichoderma harzianum strain.

Figure  4 a,b illustrate the combined effects of incubation time and pH xylanase activity increases at a high pH and shorter incubation time. Figure  4 b illustrates the contour plot, which shows that high enzyme activity was obtained at the shortest (4 days) and longest period (6 days) of incubation in acidic (4.0–6.0) conditions. Yadav et al. 29 , reported similar pH conditions for optimization of xylanse production from Anoxybacillus kamchatkensis NASTPD13.

Figure  5 a,b show that xylanase production is directly proportional to incubation time and wheat bran. This could be due to higher levels of degradation of xylan present in the wheat bran by T. harzianum . In Fig.  5 b, it is apparent that the xylanase activity is highest at high concentrations of wheat bran with the shortest (4 days) and longest period (6 days) of incubation. Previous studies showed the time course during the OFAT approach, being favourable at 4 days and 6 days of incubation with the optimum being at 5 days 33 . The RSM plots correspond with the OFAT results, as the plots show the highest xylanase activity obtained at high wheat bran between 4 and 6 days. Simultaneously, it was highlighted by Beg et al. 60 that wheat bran could effectively induce higher xylanase production by Aspergillus awamori. Li et al. 61 also reported the importance of the substrate concentrations for xylanase production by A. awamori. The facts mentioned here, correspond to the reports by Cui and Zhao 62 , as they mention that the enzymes, which are involved in substrate degradation, were generally mostly inducible. These were formed only when the substrate it correlates with, was present in the nutrient salt solution.

Figure  6 shows the highest xylanase activity obtained with high wheat bran and the entire pH range tested. However, at lower wheat bran concentrations, higher xylanase activity can be observed at the pH extremes tested (at approximately pH 4 and pH 6). Figure  6 b illustrates, at high wheat bran concentration and over a wide pH range with the highest activity obtained at the highest pH and wheat bran concentration. The interaction between the pH and wheat bran (130 U/ml) and between incubation time and wheat bran (130 U/ml) had the highest effect on xylanase activity compared to the interaction between incubation time and pH (120 U/ml).

The study successfully demonstrated a notable increase in enzyme activity using the statistically designed experiments compared to OFAT. It was also demonstrated that multiple forms of xylanase were produced (isoforms) based on variations in the growth and media conditions. Based on Table 4 , there could potentially be 5 different isoforms. High xylanase activity was observed for runs 4, 6, 7, 8, and 12. Supplementary Fig.  6 representing these RSM runs indicates the presence of isoforms by several zones of clearance on the substrate native PAGE gels.

Multiple forms of xylanases with different pH optima could be beneficial for animal feed improvement 13 . Xylanase is used to reduce the viscosity of the feed and improve the absorption of nutrients in the digestive tract of animals. The enzymes could be applied before the pelleting process, which operates between pH 4.0 and 6.0 thus requiring enzymes that are active within this pH range. Most xylanases reported to date are optimally active in the acidic or neutral pH range. Xylanases with acidic pH optima could potentially also be useful for applications containing waste, as a method of waste management, and as a feedstock for fermentable sugars 63 .

Scaled-up fermentation in optimized conditions for further studies

The xylanase enzyme was produced on a larger scale for further studies. The enzyme was produced at pH 5.0 for 6 days of incubation and with 1.2% wheat bran. The enzyme activity was determined in order to compare the activities to the smaller scale production. The enzyme activity obtained was 152.78 U/ml which was similar to the enzyme production on a smaller scale (153.80 U/ml).

Purification of xylanase from the T. harzianum isolate and zymography

The xylanase from T. harzianum was purified using ammonium sulphate precipitation, dialysis, and chromatographic methods in combination. Table 8 summarizes the purification stages. The enzyme was fractionated with the following ammonium sulphate saturations: (0–19%, 20–29%, 30–39%, 40–49%, 50–59%, 60–69% and 70–79%). The 70–79% saturation fraction resulted in significantly high xylanase activity with a recovery of 20.31% enzyme activity. The 50–59%, 60–69%, and fractions also showed relatively high recovery of enzyme activity (18.73%, and 17.48%, respectively) whereas 10.42% enzyme was recovered in the 40–49% fraction (Table 8 ) therefore these fractions were further studied to confirm if they were isoforms. The active fractions were then dialyzed at 4 °C overnight to remove the salts, and the enzyme was loaded onto DEAE Sephadex for further purification. A 0–2 M sodium chloride concentration gradient was used to elute the bound protein. Xylanase activity was measured in both bound and unbound protein fractions. The primary peak eluted at 0.5 M sodium chloride and the corresponding fraction had a specific activity of 254.62 mol/mg, and a 2.52-fold purity. Furthermore, a single band with a molecular weight of 72 kDa was evident on SDS-PAGE gels of the purified enzyme (50%) (Fig.  7 a). The other ammonium sulphate fractions (60–79%) also have the same molecular weight protein (72 kDa). To assess the activity/purity, the purified xylanase was subjected to zymogram analysis by substrate native-PAGE (1% beechwood xylan). The xylanolytic activity of the enzyme was indicated by clear zones in the gel after Congo-red staining (Fig.  7 b). Purified preparations of enzymes are a requisite for their application as well as elucidating their basic characteristics and mechanisms. Based on the high molecular weight of the purified enzyme, it can be tentatively inferred that it may belong to the GH10 family since enzymes belonging to this family feature a larger molecular weight 63 . Enzymes are also classified based on their catalytic reactions. Based on the sequence similarities of amino acids, xylanases are classified into glycosyl hydrolase (GH) families 10 (GH10) and 11 (GH11) 2 . Family GH10 contains xylanases of high molecular mass (> 30 kDa) with a (β/α) 8  barrel structure and acidic  pI  values, while GH11 include are the low-molecular-weight endoxylanases which are divided into alkaline  pI  and acidic  pI  xylanases 2 .

A 12% SDS PAGE ( a ) and Native substrate-PAGE ( b ) analysis of purified xylanase. 12% SDS PAGE (cropped) represents Lane M: Molecular weight marker (Thermoscientific, USA), 1–4: 50, 60, 70, and 80% ammonium sulphate fractions (Coomassie-stained), and 5–8: Purified xylanase from Trichoderma harzianum (Xyn). Native substrate-PAGE (cropped) represents Lane M: Molecular weight marker (Thermoscientific, USA), 1: 50% Ammonium sulphate fraction showing zone of clearance, and Lane 2: Purified xylanase (Xyn) from Trichoderma harzianum on native substrate gel showing zone of clearance. The original gels are presented in Supplementary Figs.  1 – 5 .

Characterization of xylanase

Ph optimum and stability.

The enzyme activity is greatly affected by pH because substrate binding and catalysis are dependent on the charge distribution of both the substrate and the enzyme molecules. The reaction pH was adjusted to 4.0–10.0 with various buffers as described above. The optimum pH of T. harzianum xylanase is pH 6.0 with an activity of 40 U/ml (Fig.  8 a). The enzyme is fairly stable at pH 6.0 and remains active (Fig.  9 ) retaining > 70% of its activity over 4 h. Souza et al. 64 reported that the xylanase from Thermoascus aurantiacus expressed in E. coli showed optimum activity and stability at a similar pH. Yadav et al. 29 reported that xylanase from Anoxybacillus kamchatkensis NASTPD13 showed high activity between pH 6.0 to 9.0 and at pH 6.0, the enzyme retained 71% of its activity over 24 h. The purified 60–79% ammonium sulphate fraction was further confirmed to contain the same protein as that purified in the 50% fraction as it displayed the same pH optimum and size (pH 6.0) (Fig.  8 a). Thus, the purified fractions of the 50–79% ammonium sulphate fractions can be combined to increase the yield (%).

Effect of pH ( a ) and temperature ( b ) on the activity of purified xylanases (50%, 60%, 70%, and 80% ammonium sulphate fractions). Data points represent the means ± SD (n = 4).

pH and temperature stability of purified xylanases (50% ammonium sulphate fraction) produced by the Trichoderma harzianum isolate. Data points represent the means ± SD (n = 4).

Optimum temperature and thermal stability

The experiment was carried out at different reaction temperatures ranging from 40 to 79 °C to find the optimal temperature of the xylanase. The highest activity of xylanase was observed at 65 °C (Fig.  8 b). Thermal stability data illustrated in Fig.  9 shows that the enzyme retained > 70% activity at 65 °C for 4 h. A similar result was reported by de Oliveira Simões et al. 65 . However, in that study, the enzyme was subjected to treatment for 24 h and was stable for 1 h. The purified 60–79% ammonium sulphate fraction contained the same purified protein as the 50% fraction, with the same molecular weight, pH and temperature optima obtained (65 °C) (Fig.  8 b). This confirms that these fractions are not isoforms of the xylanase produced. However, the shape of the curve for the 50% ammonium sulphate fraction is different from the other fractions, which seem to show an optimum rather than a broad bell shape.

The advantages of enzymes that prefer high temperatures are well known because the solubility of the reagents and products is increased, the viscosity is reduced, and the mass transfer rate is higher 66 . When looking for enzymes for industrial uses, stability, and activity at high temperatures are highly desirable.

Effect of metal ions and inhibitors

The effects of 8 metal ions (Ca 2+ , Co 2+ , Fe 2+ , Mg 2+ , Mn 2+ , Zn 2+ , K + , and Na + ) at a final concentration of 2 mM and 10 mM on xylanase activity were determined (Table 9 ) at the optimal pH and temperature (6.0 and 65 °C). Enzyme activity was slightly increased by 2 mM Mn2 + , K + , and Na + (101.11–101.77 U/ml) whereas the enzyme activity was slightly but significantly higher with 10 mM Ca 2+ , Co 2+ , Fe 2+ , Mg 2+ , Zn 2+ (104.27–110.89 U/ml) (p ≥ 0.05) and thus, these ions act as cofactors for the enzyme. Maximum enhancement was observed for Fe 2+ (10.88%) followed by Mg 2+ (9.43%) and Zn 2+ (8.43%) at 10 mM. Fu et al. 43 reported similar findings for xylanase from Trichoderma sp. TPS-36.

Inhibitory effects were observed for Fe 2+ (15.29%), Mg 2+ (3.44%), Zn 2+ (5.95%) at 2 mM and Na + (1.24%) at 10 mM, however, this inhibition of xylanase was weak (< 50%). Co 2+ and Ca 2+ had no effect on xylanase activity (100%) at either concentrations.

Fu et al . 43 also reported weak (< 50%) inhibition of xylanase with the same ions and that Co 2+ and Ca 2+ had no effect on xylanase activity (100%) at either concentrations.

Substrate specificity of purified xylanase

To determine the substrate specificity of the xylanase for polysaccharide degradation, potential substrates, including birchwood xylan, beechwood xylan, wheat arabinoxylan (soluble and insoluble), xylan from Larchwood, CMC and Avicel were tested under optimal conditions (pH 6.0 and temperature 65 °C) using the purified xylanase. Higher hydrolytic activity was observed for the xylans from beechwood, birchwood, and Larchwood compared to wheat arabinoxylan (Table 10 ). The xylanase most actively degraded birchwood xylan (174.07%), followed by Larchwood xylan (131.03%), and presented the lowest activity towards wheat arabinoxylan (soluble 70.54% and insoluble 46.62%). The purified xylanase exclusively hydrolyzed xylans, with no activity on CMC and Avicel. This suggested that xylanase's substrate-binding domain has a high affinity for xylans from softwood (birchwood and beechwood) 67 . This might be due to differences in xylan polymer structures and the presence of reactive groups on the surface that are more readily bound. The purified xylanase exhibited significant hydrolytic activity on the diverse xylan substrates, indicating that it might be classified as an endo-1,4-xylanase 68 .

Kinetic analysis

The Michaelis constant, K m , may be determined by measuring the substrate concentration at half the maximum velocity. K m is a constant that remains fixed for every given enzyme and substrate combination. As a result, a low K m improves the enzyme's affinity for the substrate 69 The concentration range of the substrate under investigation was 1–20 mg/ml, the study revealed K m and V max were 5.56 mg/ml and 1052.63 µmol/min/mg (Fig.  10 ). The value of K m is within the range of fungal xylanases reported in literature (0.14–14 mg/ml). Raj et al. 70 obtained similar values (4.96 mg/ml and lower V max 402 µmol/mg/ml) for xylanase from alkaliphilic Bacillus licheniformis . Fu et al. 43 reported high V max (1250 µmol/min/mg) similar to this study. Because xylanase has a high V max value and a low K m value, it has a high affinity for the substrate, beechwood xylan, and can catalyze it more efficiently and quickly than other substrates. Xylanases from Caldicoprobacter algeriensis sp. nov. strain TH7C1T were shown to have high selectivity for beechwood xylan 42 .

Double reciprocal plot of the purified (50% ammonium sulphate fraction) xylanase from Trichoderma harzianum on beechwood xylan. Data points represent the means ± SD (n = 3).

The current study describes the successful optimization of xylanase production via statistical modelling using PBD and BBD by a T. harzianum strain in submerged fermentation. The most influential independent variables were identified and optimized—resulting in a 4.16-fold and 2.24-fold increase in enzyme activity with BBD compared to the OFAT and PBD, respectively. PBD allowed for the consideration of various variables and avoided loss of information, which might be essential in the optimization of the fermentation process. The predictions of the mathematical models were validated by experimental results. Quadratic models with three independent variables were shown to accurately define xylanase production, with high R 2 values for correlations between the actual and predicted values of the response variables. Results showed high enzyme activities obtained within a high pH range which indicates the potential of the xylanase for use over a wide range of applications. Acidic-thermostable xylanase was purified with a 10.42% recovery and 2.52-fold purity. The specific activity of purified xylanase was 254.63 U/mg. The acidic-thermostability of T. harzianum xylanase is advantageous for animal feed manufacturing. Future studies will include scaling up the production of xylanase from T. harzianum under optimized conditions, which include the factors and their variables that resulted in the highest xylanase activity (RSM, run 8). Studies will also include sequencing the xylanase protein to understand the structure-guided function of this enzyme.

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author upon reasonable request. Other data generated or analysed during this study are included in this article [and its supplementary information file] .

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Acknowledgements

This work was supported by the Technology Innovation Agency (TIA) managed DST/CON/0177/2018: SIIP: ENZYME AND MICROBIAL TECHNOLOGIES (EMT) grant and the Biorefinery Industry Development Facility (BIDF) at the Council for Scientific and Industrial Research (CSIR), Durban, South Africa.

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Dhaver, P., Pletschke, B., Sithole, B. et al. Optimization, purification, and characterization of xylanase production by a newly isolated Trichoderma harzianum strain by a two-step statistical experimental design strategy. Sci Rep 12 , 17791 (2022). https://doi.org/10.1038/s41598-022-22723-x

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Microbial xylanases and their industrial applications: a review

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• 1 Department of Microbiology, University of Delhi, New Delhi, India. [email protected]
• PMID: 11548999
• DOI: 10.1007/s002530100704

Despite an increased knowledge of microbial xylanolytic systems in the past few years, further studies are required to achieve a complete understanding of the mechanism of xylan degradation by microorganisms and their enzymes. The enzyme system used by microbes for the metabolism of xylan is the most important tool for investigating the use of the second most abundant polysaccharide (xylan) in nature. Recent studies on microbial xylanolytic systems have generally focussed on induction of enzyme production under different conditions, purification, characterization, molecular cloning and expression, and use of enzyme predominantly for pulp bleaching. Rationale approaches to achieve these goals require a detailed knowledge of the regulatory mechanism governing enzyme production. This review will focus on complex xylan structure and the microbial enzyme complex involved in its complete breakdown, studies on xylanase regulation and production and their potential industrial applications, with special reference to biobleaching.

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• DOI: 10.33451/florafauna.v28i2pp231-240
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Microbial Xylanases: A Review

• Jyoti Richhariya , T. Sharma , S. Dassani
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Production and optimization of xylanase enzyme from bacillus subtilis using agricultural wastes by solid state fermentation, xylanases from fungi: properties and industrial applications, production of fungal xylanases, engineering thermostable microbial xylanases toward its industrial applications, microbial xylanases and their industrial application in pulp and paper biobleaching: a review, biotechnology of microbial xylanases: enzymology, molecular biology, and application, production and industrial applications of xylanase: a review, xylanases and their applications in baking industry, production and characterization of xylanase from aspergillus niger using wheat bran, corn cobs and sugar cane bagasse as carbon sources with different concentrations, a review of xylanase production by the fermentation of xylan: classification, characterization and applications, related papers.

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Xylanases: An Overview of its Diverse Function in the Field of Biorefinery

• First Online: 11 August 2021

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• Nisha Bhardwaj 6   na1 ,
• Komal Agrawal 6   na1 &
• Pradeep Verma 6   na1  

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Hemicellulases are enzymatic complex that has its application in biorefinery and can effectively catalyse the hydrolysis process of xylan (a principal hemicellulose), comprising of a linear polymeric chain made up of β-D-xylopyranosyl units linked with β-1,4-glycosidic linkages. Hemicellulases are produced by various microorganisms such as bacteria, fungi, algae, insects, etc. However, xylanase from filamentous fungi such as Aspergillus sp. and Trichoderma sp. is more preferred over other sources. Endo-1,4-β-xylanase and β-xylosidase are found to be most important hemicellulases and have tremendous application in biorefinery as well as other industries. The growing interest of industries such as biorefinery, animal feed, pharmaceuticals, paper and pulp industries, etc., has gained interest of the researchers and focused the research to find new sources of xylanases that utilise economically feasible production methods and be sustainable as well. Thus, the present chapter will describe the application of xylanases in biorefinery as well as different industries along with the general production strategy applied by the researchers in current time.

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Bhardwaj, N., Agrawal, K., Verma, P. (2021). Xylanases: An Overview of its Diverse Function in the Field of Biorefinery. In: Srivastava, M., Srivastava, N., Singh, R. (eds) Bioenergy Research: Commercial Opportunities & Challenges . Clean Energy Production Technologies. Springer, Singapore. https://doi.org/10.1007/978-981-16-1190-2_10

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Home > Books > Sustainable Degradation of Lignocellulosic Biomass - Techniques, Applications and Commercialization

A Review of Xylanase Production by the Fermentation of Xylan: Classification, Characterization and Applications

Submitted: 12 April 2012 Published: 15 May 2013

DOI: 10.5772/53544

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F. l. motta *.

• Development of Biotechnological Processes Laboratory, School of Chemical Engineering, University of Campinas, Campinas, Brazil

C. C. P. Andrade

• Bioprocess Engineering Laboratory, Food Engineering Department, University of Campinas, Campinas, Brazil

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1. Introduction

The enzymatic hydrolysis of xylan, which is the second most abundant natural polysaccharide, is one of the most important industrial applications of this polysaccharide [ 1 , 2 ]. The primary chain of xylan is composed of β-xylopyranose residues, and its complete hydrolysis requires the action of several enzymes, including endo-1,4-β-D-xylanase (EC3.2.1.8), which is crucial for xylan depolymerization [ 2 ]. Due to the diversity in the chemical structures of xylans derived from the cell walls of wood, cereal or other plant materials, a large variety of xylanases with various hydrolytic activities, physicochemical properties and structures are known. Moreover, xylan derivatives are frequently used to induce the production of xylanases [ 3 ] by microorganisms [ 4 ], using either solid-state or submerged fermentation [ 5 ].

Xylanases and the microorganisms that produce them are currently used in the management of waste, to degrade xylan to renewable fuels and chemicals, in addition to their use in food, agro-fiber, and the paper and pulp industries, where the enzymes help to reduce their environmental impact [ 6 ]. Oligosaccharides produced by the action of xylanases are further used as functional food additives or alternative sweeteners with beneficial properties [ 7 ].

To meet the needs of industry, more attention has been focused on the enzyme stability under different processing conditions, such as pH, temperature and inhibitory irons, in addition to its ability to hydrolyze soluble or insoluble xylans. Although many wild-type xylanases contain certain desired characteristics, such as thermostability, pH stability or high activity, no individual xylanase is capable of meeting all of the requirements of the feed and food industries. Moreover, as industrial applications require cheaper enzymes, the elevation of expression levels and the efficient secretion of xylanases are crucial to ensure the viability of the process; therefore, genetic engineering and recombinant DNA technology have an important role in the large-scale expression of xylanases in homologous or heterologous protein-expression hosts.

Considering the future prospects of xylanases in biotechnological applications, the goal of this review chapter is to present an overview of xylanase production via fermentation and to describe some of the characteristics of these enzymes and their primary substrate, xylan. Moreover, this review will discuss the fermentation processes as well as the genetic techniques applied to improve xylanase yields.

The three main components that constitute lignocellulosic substrates are cellulose, hemicellulose and lignin [ 8 ]. Schulze [ 9 ] first introduced the term ‘hemicellulose’ to represent the fractions isolated or extracted from plant materials using a dilute alkali. Hemicelluloses are composed of complex mixtures of xylan, xyloglucan, glucomannan, galactoglucomannan, arabinogalactan or other heteropolymers [ 8 ].

The substrate of xylanase, xylan, is the second most-abundant polysaccharide in nature, accounting for approximately one-third of the renewable organic carbon on Earth [ 10 ], and it constitutes the major component of hemicellulose, a complex of polymeric carbohydrates, including xylan, xyloglucan (heteropolymer of D-xylose and D-glucose), glucomannan (heteropolymer of D-glucose and D-mannose), galactoglucomannan (heteropolymer of D-galactose, D-glucose and D-mannose) and arabinogalactan (heteropolymer of D-galactose and arabinose) [ 11 ]. Xylan is primarily present in the secondary cell wall and together with cellulose (1,4-β-glucan) and lignin (a complex polyphenolic compound) make up the major polymeric constituents of plant cell walls [ 12 ]. Within the cell wall structure, all three constituents interact via covalent and non-covalent linkages, with xylan being found at the interface between lignin and cellulose, where it is believed to be important for fiber cohesion and plant cell wall integrity [ 1 ].

2.1. Structure and distribution

A complex, highly branched heteropolysaccharide, xylan varies in structure between different plant species, and the homopolymeric backbone chain of 1,4-linked β-D-xylopyranosyl units can be substituted to varying degrees with glucuronopyranosyl, 4-O-methyl-D-glucuronopyranosyl, α-L-arabinofuranosyl, acetyl, feruloyl or p-coumaroyl side-chain groups [ 12 , 13 ] ( Figure 1) .

Xylan is distributed in several types of tissues and cells and is present in a variety of plant species [ 12 ], being found in large quantities in hardwoods from angiosperms (15–30% of the cell wall content) and softwoods from gymnosperms (7–10%), as well as in annual plants (<30%) [ 14 ]. Wood xylan exists as O-acetyl-4-O-methylglucuronoxylan in hardwoods and as arabino-4-O-methylglucuronoxylan in softwoods, while xylans in grasses and annual plants are typically arabinoxylans [ 12 ]. Linear unsubstituted xylan has also been reported in esparto grass [ 15 ], tobacco [ 16 ] and certain marine algae [ 17 , 18 ], with the latter containing xylopyranosyl residues linked by both 1,3-β and 1,4-β linkages [ 17 , 19 ].

Similar to other polysaccharides of plant origin, xylan has a large polydiversity and polymolecularity [ 20 ]. The degree of polymerization in xylans is also variable, with, for example, hardwood and softwood xylans generally consisting of 150-200 and 70-130 β-xylopyranose residues, respectively [ 12 ].

Structure of xylan and the xylanolytic enzymes involved in its degradation. Ac: Acetyl group; α-Araf: α-arabinofuranose; α-4-O-Me-GlcA: α-4-O-methylglucuronic acid. Source: Sunna and Antranikian [ 20 ].

Based on the common substituents found on the backbone, xylans are categorized as linear homoxylan, arabinoxylan, glucuronoxylan or glucuronoarabinoxylan. Homoxylans consisting exclusively of xylosyl residues are not widespread in nature; they have been isolated from limited sources, such as esparto grass, tobacco stalks and guar seed husks [ 20 ]. However, based on the nature of its substituents, a broad distinction may therefore be made among xylans, in which the complexity increases from linear to highly substituted xylans. Four main families of xylans can be considered [ 21 ]:

Arabinoxylans, having only side chains of single terminal units of α-L-arabinofuranosyl substituents. In the particular case of cereals, arabinoxylans vary in the degree of arabinosyl substitution, with either 2-O- and 3-O-mono-substituted or double (2-O-, 3-O-) substituted xylosyl residues.

Glucuronoxylans, in which α-D-glucuronic acid or its 4-O-methyl ether derivative represents the only substituent.

Glucuronoarabinoxylan, in which α-D-glucuronic (and 4-O-methyl-α-D-glucuronic) acid and α-L-arabinose are both present.

Galactoglucuronoarabinoxylans, which are characterized by the presence of terminal β-D-galactopyranosyl residues on complex oligosaccharide side chains of xylans and are typically found in perennial plants.

In each category there exists microheterogeneity with respect to the degree and nature of branching. The side chains determine the solubility, physical conformation and reactivity of the xylan molecule with the other hemicellulosic components and hence greatly influence the mode and extent of enzymatic cleavage [ 12 ]. Endospermic arabinoxylans of annual plants, also called pentosans, are more soluble in water and alkaline solutions than xylans of lignocellulosic materials because of their branched structures [ 22 ].

2.2. Enzymatic hydrolysis of xylan

Due to the heterogeneity and complex chemical nature of plant xylan, its complete breakdown requires the action of a complex of several hydrolytic enzymes with diverse specificities and modes of action. Thus, it is not surprising for xylan-degrading cells to produce an arsenal of polymer-degrading proteins [ 1 ]. The xylanolytic enzyme system that carries out the xylan hydrolysis is normally composed of a repertoire of hydrolytic enzymes, including endoxylanase (endo-1,4-β-xylanase, E.C.3.2.1.8), β-xylosidase (xylan-1,4-β-xylosidase, E.C.3.2.1.37), α-glucuronidase (α-glucosiduronase, E.C.3.2.1.139), α-arabinofuranosidase (α-L-arabinofuranosidase, E.C.3.2.1.55) and acetylxylan esterase (E.C.3.1.1.72) [ 23 ]. All of these enzymes act cooperatively to convert xylan into its constituent sugars [ 24 ]. Among all xylanases, endoxylanases are the most important due to their direct involvement in cleaving the glycosidic bonds and in liberating short xylooligosaccharides [ 8 ].

Xylan, being a high molecular mass polymer, cannot penetrate the cell wall. The low molecular mass fragments of xylan play a key role in the regulation of xylanase biosynthesis. These fragments include xylose, xylobiose, xylooligosaccharides, heterodisaccharides of xylose and glucose and their positional isomers. These molecules are liberated from xylan through the action of small amounts of constitutively produced enzymes [ 12 ]. Xylanase catalyzes the random hydrolysis of xylan to xylooligosaccharides, while β-xylosidase releases xylose residues from the nonreducing ends of xylooligosaccharides. However, a complete degradation requires the synergistic action of acetyl esterase to remove the acetyl substituents from the β-1,4-linked D-xylose backbone of xylan [ 25 , 26 ].

3. Xylanases

3.1. classification and mode of action.

Xylanases, as glycoside hydrolase members, are able to catalyze the hydrolysis of the glycosidic linkage (β-1,4) of xylosides, leading to the formation of a sugar hemiacetal and the corresponding free aglycone (nonsugar compound remaining after replacement of the glycoside by a hydrogen atom [ 27 ]). Xylanases have been classified in at least three ways: based on the molecular weight and isoelectric point (pI) [ 28 ], the crystal structure [ 29 ] and kinetic properties, or the substrate specificity and product profile. As the first classification is not sufficient to describe all xylanases, several exceptions have been identified [ 10 ] because not all xylanases have a high molecular mass (above 30 kDa) and low pI or a low molecular mass (less than 30 kDa) and high pI [ 6 ]. Therefore, a more complete system, based on the primary structure and comparison of the catalytic domains, was introduced [ 10 , 30 ], analyzing both the structural and mechanistic features [ 10 ].

Updated information on the characteristics and classification of enzymes may be found in the Carbohydrate-Active Enzyme (CAZy) database. This is a knowledge-based resource specializing on enzymes that build and breakdown complex carbohydrates and glycoconjugates. This database contains information from sequence annotations found in publicly available sources (such as the National Center for Biotechnology Information, NCBI), family classifications and known functional information [ 31 ]. According to the CAZy database (http://www.cazy.org), xylanases (EC3.2.1.8) are related to glycoside hydrolase (GH) families 5, 7, 8, 9, 10, 11, 12, 16, 26, 30, 43, 44, 51 and 62. However, the sequences classified in families 16, 51 and 62 appear to be bifunctional enzymes containing two catalytic domains, unlike families 5, 7, 8, 10, 11 and 43, which have a truly distinct catalytic domain with endo-1,4-β-xylanase activity [ 10 ]. Using the same analysis, families 9, 12, 26, 30 and 44 may have residual or secondary xylanase activity.

Xylanases have been primarily classified as GH 10 and 11 based on the hydrophobic cluster analysis of the catalytic domains and similarities in the amino acid sequences [ 8 ]. Although members of these two families have been thoroughly studied, the catalytic properties of the members of the remaining families (5, 7, 8 and 43) are recent and remain very limited [ 32 ].

Members of GH families 5, 7, 8, 10, 11 and 43 differ in their physicochemical properties, structure, mode of action and substrate specificities [ 10 ]. Several models have been proposed to explain the mechanism of xylanase action. Xylanase activity leads to the hydrolysis of xylan. Generally, this hydrolysis may result either in the retention (GH families 5, 7, 10 and 11) or the inversion (GH families 8 and 43) of the anomeric center of the reducing sugar monomer of the carbohydrate [ 33 , 34 ].

Families 5, 7, 10 and 11 contain enzymes that catalyze the hydrolysis with the retention of the anomeric configuration, with two glutamate residues being implicated in the catalytic mechanism. This indicates a double-displacement mechanism, in which a covalent glycosyl-enzyme intermediate is formed and subsequently hydrolyzed, and two carboxylic acid residues, suitably located in the active site, are involved in the formation of the intermediate; one acts as a general acid catalyst by protonating the substrate, while the second performs a nucleophilic attack, which results in the departure of the leaving group and the formation of the α-glycosyl enzyme intermediate (β to α inversion). In the second step, the first carboxylate group instead functions as a general base, abstracting a proton from a nucleophilic water molecule, which attacks the anomeric carbon. This leads to a second substitution, in which the anomeric carbon again passes via a transition state to give rise to a product with the β configuration (α to β inversion) [ 10 , 34 ].

In contrast to the mechanism mentioned above, the enzymes in families 8 and 43 generally act via an inversion of the anomeric center, and glutamate and aspartate may be the catalytic residues. Inverting enzymes work via a single displacement reaction, in which one carboxylate provides for a general acid-catalyzed leaving group departure. The second function of these enzymes, acting as general base, activates a nucleophilic water molecule to attack the anomeric carbon, thereby cleaving the glycosidic bond and leading to an inversion of the configuration at the anomeric carbon. Generally, the distance between the two residues allows for the accommodation of the water molecule between the anomeric carbon and the general base [ 10 , 34 ].

3.1.1. GH families 10 and 11

Xylanase from the GH10 family (or family G) have a low molecular mass with a pI between 8–9.5, while those from the GH11 family (or family F) have a high molecular mass and lower pI values [ 35 , 36 ].

Glycoside hydrolase family 10 is composed of endo-1,4-β-xylanases and endo-1,3-β-xylanases (EC 3.2.1.32) [ 34 ]. Members of this family are also capable of hydrolyzing the aryl β-glycosides of xylobiose and xylotriose at the aglyconic bond. Furthermore, these enzymes are highly active on short xylooligosaccharides, thereby indicating small substrate-binding sites. Crystal structure analyses, kinetic analyses of the activity on xylooligosaccharides of various sizes and end product analyses have indicated that family 10 xylanases typically have four to five substrate-binding sites [ 37 ]. Members of this family also typically have a high molecular mass, a low pI and display an (α/β)8-barrel fold [ 10 , 34 , 38 ].

Compared to other xylanases, GH11 members display several interesting properties, such as high substrate selectivity and high catalytic efficiency, a small size, and a variety of optimum pH and temperature values, making them suitable in various conditions and in many applications [ 39 ]. Family 11 is composed only of xylanases (EC3.2.1.8), leading to their consideration as “true xylanases,” as they are exclusively active on D-xylose-containing substrates. GH11 enzymes are generally characterized by a high pI, a low molecular weight, a double-displacement catalytic mechanism, two glutamates that act as the catalytic residues and a β-jelly roll fold structure. Additionally, the products of their action can be further hydrolyzed by the family 10 enzymes [ 37 ]. Similar to family 10 xylanases, these enzymes can hydrolyze the aryl β-glycosides of xylobiose and xylotriose at the aglyconic bond, but they are inactive on aryl cellobiosides. Furthermore, in contrast to the family 10 xylanases, but similar to the family 8 cold-adapted xylanases, these enzymes are most active on long-chain xylooligosaccharides, and it has been found that they have larger substrate-binding clefts, containing at least seven subsites [ 10 ].

Xylanases belonging to GH10 exhibit greater catalytic versatility and lower substrate specificity than those belonging to GH11 [ 37 , 40 ]. According to Davies et al . [ 41 ], the binding sites for xylose residues in xylanases are termed subsites, with bond cleavage occurring between the sugar residues at the -1 (non-reducing) and the +1 (reducing) ends of the polysaccharide substrate. As observed in assays using arabinoxylan as the substrate, GH10 products have arabinose residues substituted on xylose at the +1 subsite, whereas GH 11 products have arabinose residues substituted at the +2 subsite [ 42 ]. These results suggest that GH 10 enzymes are able to hydrolyze xylose linkages closer to the side-chain residues [ 43 ]. Therefore, xylanases from family 11 preferentially cleave the unsubstituted regions of the arabinoxylan backbone, whereas GH10 enzymes cleave the decorated regions, being less hampered by the presence of substituents along the xylan backbone [ 37 ]. The xylan side-chain decorations are recognized by xylanases, and the degree of substitution in xylan will influence the hydrolytic products; this difference in substrate specificity has important implications in the deconstruction of xylan [ 43 ].

3.1.2. GH families 5, 7, 8 and 43

GH family 5 (or family A) is the largest glycoside hydrolase family, and only seven amino acid residues, including the nucleophile and the general acid/base residue, are strictly conserved among all members [ 10 ]. Structural alignment among the members of family 5 and 10 showed that these enzymes are as structurally different within family 5 as they are to the family 10 enzymes, therefore both families are classified into clan GH-A. The concept of clan or superfamily demonstrates a broader relationship between GH families, suggesting a more distant common evolutionary ancestor [ 44 , 45 ]. Furthermore, the activity of these enzymes is affected by substituents on the xylan main chain, and it is unable to cleave linkages adjacent to substituted residues. Hydrolysis studies have shown that the shortest substituted fragments formed from glucuronoxylan and arabinoxylan are substituted xylotrioses, with the substitution being found on the internal xylose residue. Therefore, the products produced by family 5 are shorter than those produced by family 7 [ 10 ].

GH family 8 (or family D) is composed of cellulases (EC 3.2.1.4), and also contains chitosanases (EC 3.2.1.132), lichenases (EC 3.2.1.73) and endo-1,4-β-xylanases (EC 3.2.1.8). This family of cold-adapted xylanases was found to hydrolyze xylan to xylotriose and xylotetraose and was most active on long-chain xylooligsaccharides. Similar to family 11 xylanases, a large substrate binding cleft containing at least six xylose-binding residues, with the catalytic site in the middle, was proposed [ 6 ]. However, unlike family 10 and 11 xylanases, enzymes from family 8 were found to catalyze hydrolysis with the inversion of the anomeric configuration and, under the conditions used, were found to be inactive on aryl β-glycosides of xylose, xylobiose and xylotriose [ 10 , 31 ].

GH families 7 and 43 contain only a few enzymes exhibiting xylanase activity that have been identified and studied. Family 7 has characteristics in common with both family 10 and 11 xylanases. Similar to the former family, those in family 7 have a high molecular weight and low pI, as well as a small substrate-binding site, containing approximately four subsites, with the catalytic site in the middle [ 10 ]. The members of family 43 have not been as thoroughly studied, and the structure of only one member has been determined, indicating that members of this family may display a five-blade β-propeller fold. Furthermore, a glutamate and aspartate in the center of a long V-shaped surface groove formed across the face of the propeller have been suggested as the catalytic residues. Family 43 is grouped with family 62 in clan GH-F, and, as also demonstrated in the family 8 enzymes, its members are believed to catalyze hydrolysis via a single displacement mechanism [ 10 , 31 ].

3.2. Properties and applications

The heterogeneity and complexity of xylan have resulted in a diverse range of xylanases, which differ in their physicochemical properties, structure, mode of action and substrate specificities [ 10 ]. As the xylosidic linkages in lignocellulose are neither equivalent nor equally accessible, the production of an enzymatic system with specialized functions is a strategy to achieve superior xylan hydrolysis [ 28 ]. Together with the heterogeneous nature of xylan, the multiplicity of xylanases in microorganisms may be caused by a redundancy in gene expression. Generally, a single xylanase gene encodes multiple xylanases, and xylanase multiplicity may arise from posttranslational modifications, such as differential glycosylation, proteolysis or both [ 23 ].

The potential applications of xylanases also include the bioconversion of lignocellulosic material and agro-wastes into fermentative products, the clarification of juices, the improvement of the consistency of beer and the digestibility of animal feedstocks [ 28 ]. One of the most important biotechnological applications of xylanase is its use in pulp bleaching [ 46 ]. Xylanases may also be applicable to the production of rayon, cellophane and several chemicals such as cellulose esters (acetates, nitrates, propionates and butyrates) and cellulose ethers (carboxymethyl cellulose and methyl and ethyl cellulose), which are all produced by dissolving pulp and purifying fibers from other carbohydrates [ 33 ].

3.2.1. The paper and pulp industries

During the past several years, the use of enzymes in paper and pulp bleaching has caught the attention of researchers and industries all over the world. Xylanase enzymes have proven to be a cost-effective means for mills to take advantage of a variety of bleaching benefits [ 47 ]. Xylanases and other side-cleaving enzymes have been used in pulp bleaching primarily to reduce lignin and increase the brightness of the pulp [ 20 , 46 ]. The importance of xylanase in the pulp and paper industries is related to the hydrolysis of xylan, which facilitates the release of lignin from paper pulp and, consequently, reduces the usage of chlorine as the bleaching agent [ 33 ].

Bleaching is the process of lignin removal from chemical pulps to produce bright or completely white finished pulp [ 1 ]. Thus, the bleaching of pulp using enzymes or ligninolytic microorganisms is called biobleaching [ 48 ]. This process is necessary due to the presence of residual lignin and its derivatives in the pulping process, which causes the resultant pulp to gain a characteristic brown color. The intensity of this pulp color is related to the amount and chemical state of the remaining lignin [ 33 ].

The bleaching of pulp involves the destruction, alteration or solubilization of the lignin, colored organic matter and other undesirable residues on the fibers [ 33 ]. Bleaching of kraft pulp usually requires large amounts of chlorine-based chemicals and sodium hydrosulfite, which cause several effluent-based problems in the pulp and paper industries. The use of these chemicals generates chlorinated organic substances, some of which are toxic, mutagenic, persistent, and highly resistant to biodegradation, in addition to causing numerous harmful disturbances in biological systems and forming one of the major sources of environmental pollution [ 1 , 33 , 49 ].

As hemicellulose is easier to depolymerize than lignin, biobleaching of pulp appears to be more effective with the use of xylanases than with lignin-degrading enzymes. This is due to the fact of the removal of even a small portion of the hemicellulose could be sufficient to open up the polymer, which facilitates removal of the residual lignin by mild oxidants [ 33 , 50 ].

The use of xylanase in bleaching pulp requires the use of enzymes with special characteristics. A key requirement is to be cellulose-free, to avoid damaging the pulp fibers [ 3 ], as cellulose is the primary product in the paper industry [ 33 ]. Other desirable characteristics are stability at high temperatures [ 51 ] and an alkaline optimal pH [ 48 ].

Madlala et al . [ 52 ] used different preparations of commercial Xylanase P and crude xylanase from Thermomyces lanuginosus to evaluate the bleaching process of paper pulp. It was demonstrated that the use of enzymes could increase the pulp brightness (over 5 brightness points over the control) and reduce the amount of bleaching chemicals used (up to 30% for chlorine dioxide). Chipeta et al . [ 53 ] evaluated crude xylanase preparations from Aspergillus oryzae NRRL 3485 and Aspergillus phoenicis ATCC 13157 and found that at a charge of 10 U per gram of pulp it was possible to reduce the usage of chlorine dioxide up to 30% without compromising the pulp brightness.

3.2.2. Bioconversion of lignocellulose in biofuels

Currently, second-generation biofuels are the primary products of the bioconversion of lignocellulosic materials. According to Taherzadech and Karimi [ 54 ], ethanol is the most important renewable fuel in terms of volume and market value, and following the fossil fuel crisis, it has been identified as an alternative fuel [ 48 ]. Despite the primarily first-generation production of ethanol, from sugar and starch, the second-generation production of ethanol has only begun to be tested in pilot plants [ 55 ]. And, unlike first-generation biofuels, second-generation biofuels do not compete with food production and can provide environmental, economic, and strategic benefits for the production of fuels [ 56 ].

Xylanase, together with other hydrolytic enzymes, can be used for the generation of biological fuels, such as ethanol, from lignocellulosic biomass [ 1 , 57 ]. However, enzymatic hydrolysis is still a major cost factor in the conversion of lignocellulosic raw materials to ethanol [ 56 ]. In bioethanol fuel production, the first step is the delignification of lignocellulose, to liberate cellulose and hemicellulose from their complex with lignin. The second step is a depolymerization of the carbohydrate polymers to produce free sugars, followed by the fermentation of mixed pentose and hexose sugars to produce ethanol [ 1 , 58 ]. Simultaneous saccharification and fermentation is an alternative process, in which both hydrolytic enzymes and fermentative microorganism are present in the reaction [ 48 , 59 ].

3.2.3. The pharmaceutical, food and feed industries

Xylanase, together with pectinase, carboxymethylcellulase and amylase, can be used for the clarification of juices because the turbidity observed is due to both pectic materials and other materials suspended in a stable colloidal system [ 60 ]. Xylanase may also improve the extraction of coffee, plant oils, and starch [ 25 ]. The xylose resulting from xylan depolymerization may also be converted to xylitol, a valuable sweetener that has applications in both the pharmaceutical and food industries [ 61 - 63 ].

In the bakery industry, xylanase may improve the quality of bread, by increasing the bread’s specific volume. In combination with amylases, this characteristic was enhanced, as observed upon the introduction of Aspergillus niger var. awamori [ 64 ]. According to Collins et al . [ 65 ], psychrophilic enzymes may be suitable for use in the baking industry as they are generally optimally active at the temperatures most frequently used for dough preparation (at or below 35 °C). These enzymes could also be used as more efficient baking additives than the currently used commercial mesophilic enzymes, which are optimally active at higher temperatures.

Xylanase may also improve the nutritional properties of agricultural silage and grain feed. The use of this enzyme in poultry diets showed that the decrease in weight gain and feed conversion efficiency in rye-fed broiler chicks has been associated with intestinal viscosity [ 66 ]. The incorporation of xylanase from Trichoderma longibrachiatum into the rye-based diet of broiler chickens reduced intestinal viscosity, thus, improving both the weight gain of the chicks and their feed conversion efficiency [ 67 ].

Xylanases can also be used in cereals as a pretreatment for arabinoxylan-containing substrates, as arabinoxylans are partly water soluble and result in a highly viscous aqueous solution. This high viscosity of cereal grain water extract may lead to brewing problems, by decreasing the rate of filtration or haze formation in beer. Additionally, it is unfavorable in the cereal grains used in animal feeding [ 68 , 69 ].

The enzymatic hydrolysis of xylan may also result in oligomers known as xylooligosaccharides (XOs), which may be used in pharmaceutical, agriculture and feed products. XOs have prebiotic effects, as they are neither hydrolyzed nor absorbed in the upper gastrointestinal tract, and they affect the host by selectively stimulating the growth or activity of one or a number of bacteria in the colon, thus improving health [ 70 - 72 ]. Among their key physiological advantages are the reduction of cholesterol, maintenance of gastrointestinal health, and improvement of the biological availability of calcium. They also inhibit starch retrogradation, improving the nutritional and sensory properties of food [ 73 ]. For the production of XOs, the enzyme complex must have low exoxylanase or β-xylosidase activity, to prevent the production of high amounts of xylose, which has inhibitory effects on XO production [ 74 , 75 ].

3.3. Xylanase assays

The xylanase activity is often assayed based on measurement of reducing sugar released during the course of hydrolysis of xylan, by DNS or Nelson-Somogyi methods. Due to absent of standardization, Bailey et al. [ 76 ] compared the measurement of xylanase activity by twenty different laboratories. According to the author, the major source of variation between apparent xylanase activities was probably the substrate chosen, although small differences in protocols were also significant. After standardization of substrate and method, the interlaboratory standard variation of the results decreased from 108% to 17% from the mean. Others researchers use the 4-o-methylglucuronoxylan covalently dyed with Remazol Brilliant Blue (RBB xylan) as substrate, and the xylanase is assayed based on the release of the dyed fragments [ 77 ]. There are also available some commercial methods for xylanase assays, as the fluorescence-based method EnzChek® Ultra Xylanase Assay Kit (Invitrogen, Carlsbad, CA) or the Xylazyme tablet (Megazyme, Bray, Ireland), which employs azurine-crosslinked arabinoxylan (AZCLArabinoxylan) as substrate and its hydrolysis by xylanase produces water soluble dyed fragments.

3.4. Producing microorganisms

Microorganisms, in particular, have been regarded as a good source of useful enzymes because they multiply at extremely high rates and synthesize biologically active products that can be controlled by humans. In recent years, there has been a phenomenal increase in the use of enzymes as industrial catalysts. These enzymes offer advantages over the use of conventional chemical catalysts for numerous reasons: they exhibit high catalytic activity and a high degree of substrate specificity, they can be produced in large amounts, they are highly biodegradable, they pose no threat to the environment and they are economically viable [ 4 ].

In this context, microbial xylanases are the preferred catalysts for xylan hydrolysis, due to their high specificity, mild reaction conditions, negligible substrate loss and side product generation. Xylanases derived from microorganisms have many potential applications in the food, feed, and paper pulp industries [ 10 , 12 , 78 ]. Complete xylanolytic enzyme systems, which including all of these activities, have been found to be widespread among fungi [ 20 , 24 ], actinomycetes [ 79 ] and bacteria [ 12 ], and some of the most important xylanolytic enzyme producers include Aspergillus, Trichoderma, Streptomyces, Phanerochaetes, Chytridiomycetes, Ruminococcus, Fibrobacteres, Clostridia and Bacillus [ 12 , 78 , 80 , 81 ]. The ecological niches of these microorganisms are diverse and widespread and typically include environments where plant materials accumulate and deteriorate, as well as in the rumen of ruminants [ 78 , 82 , 83 ].

Although there have been many reports on microbial xylanases since the 1960s, the prime focus has been on plant pathology related studies [ 84 ]. Only during the 1980’s did the use of xylanases for biobleaching begin to be tested [ 85 ]. Since 1982, several microorganisms, including fungi and bacteria, have been reported to readily hydrolyze xylans by synthesizing 1,4-β-D endoxylanases (E.C. 3.2.18) and β-xylosidases (EC.3.2.1.37) [ 86 ]. Table 1 presents a list of some of the xylanase-producing microorganisms and their activities.

The production of xylanases must be improved by finding more potent fungal or bacterial strains or by inducing mutant strains to excrete greater amounts of the enzymes. Moreover, the level of microbial enzyme production is influenced by a variety of nutritional and physiological factors, such as the supply of carbon and nitrogen, physical circumstances and chemical conditions [ 98 ].

Review of xylanases-producing microorganisms.

3.4.1. Fungi

Filamentous fungi are particularly interesting producers of xylanases and other xylan-degrading enzymes because they excrete the enzymes into the medium and their enzyme levels are much higher than those of yeast and bacteria. In addition to xylanases, fungi produce several auxiliary enzymes required for the degradation of substituted xylan [ 2 ].

The fungal genera Trichoderma , Aspergillus , Fusarium , and Pichia are considered great producers of xylanases [ 99 ]. White-rot fungi have also been shown to produce extracellular xylanases that act on a wide range of hemicellulosic materials, are useful as food sources [ 100 ] and produce metabolites of interest to the pharmaceutical, cosmetic, and food industries [ 78 ]. White-rot basidiomycetes normally secrete large amounts of these enzymes to degrade lignocellulosic materials. For example, Phanerochaete chrysosporium produces high levels of α-glucuronidase [ 101 ], and Coriolus versicolor produces a complex xylanolytic combination of enzymes [ 102 ]. Xylanase is also produced by Cuninghamella subvermispora when growing on plant cell-wall polysaccharides or on wood chips [ 103 ].

Fungal xylanases are generally associated with celluloses [ 104 ]. On cellulose these strains produce both cellulase and xylanase, which may be due to traces of hemicellulose present in the cellulosic substrates [ 105 ]; however, selective production of xylanase may be possible using only xylan as the carbon source. The mechanisms that govern the formation of extracellular enzymes with regards to the carbon sources present in the medium are influenced by the availability of precursors for protein synthesis. Therefore, in some fungi, growing the cells on xylan uncontaminated by cellulose under a lower nitrogen/carbon ratio may be a possible strategy for producing xylanolytic systems free of cellulases [ 106 ]. Another major problem associated with fungi is the reduced xylanase yield in fermenter studies. Agitation is normally used to maintain the medium homogeneity, but the shearing forces in the fermenter can disrupt the fragile fungal biomass, leading to the reported low productivity. Higher rates of agitation may also lead to hyphal disruption, further decreasing the xylanase activity [ 50 ].

3.4.2. Bacteria

Xylanases have been reported in Bacillus , Streptomyces and other bacterial genera that do not have any role related to plant pathogenicity [ 86 ]. The extreme thermophile Rhodothermus marinus has been reported to produce α-L-arabinofuranosidase [ 107 ], and two different polypeptides with α-arabinofuranosidase activity from Bacillus polymyxa were characterized at the gene level for the production of α-arabinofuranosidases [ 108 ].

Bacteria, just like many other industrial enzymes, have fascinated researchers due to their alkaline-thermostable xylanase-producing trait [ 33 ]. The optimum pH of bacterial xylanases are, in general, slightly higher than the optimal pH of fungal xylanases [ 109 ], which is a suitable characteristic in most industrial applications, especially the paper and pulp industries. Noteworthy producers of high levels of xylanase activity at an alkaline pH and high temperature are Bacillus spp. [ 33 ]. When considering only temperature, a handful of xylanases that show optimum activity at higher temperatures have been reported from various microorganisms. These include Geobacillus thermoleovorans , Streptomyces sp. S27, Bacillus firmus , Actinomadura sp. strain Cpt20 and Saccharopolyspora pathunthaniensis S582, all of which produce xylanases that show activity between 65 and 90 °C [ 8 ]. One xylanase, reported from Thermotoga sp. [ 110 ], has been shown to exhibit a temperature optima between 100 and 105 °C.

3.5. Production of xylanases under SSF and SmF

Xylanases are produced by either solid-state or submerged fermentation [ 5 ]. Although most xylanase manufacturers produce these enzymes using submerged fermentation (SmF) techniques (nearly for 90% of the total xylanase sales worldwide) [ 2 ], the enzyme productivity via solid-state fermentation (SSF) is normally much higher than that of submerged fermentation [ 5 ]. The growing interest in using solid-state fermentation (SSF) techniques to produce a wide variety of enzymes, including xylanases from fungal origins, is primarily due to the economic and engineering advantages of this process [ 111 ].

The advantages of SSF processes over SmF include a low cultivation cost for the fermentation, lower risk of contamination [ 1 ], improved enzyme stability, mimicking the natural habit of the fungus, production of enzymes with higher specific activities, generation of a protein-enriched byproduct, and easier downstream processing of the enzymes produced [ 112 ]. SSF conditions are especially suitable for the growth of fungi, as these organisms are able to grow at relatively low water activities, contrary to most bacteria and yeast, which will not proliferate under these culture conditions [ 113 ].

On the contrary, submerged fermentation allows better control of the conditions during fermentation [ 114 ]. The submerged fermentation of aerobic microorganisms is a well-known and widely used method for the production of cellulase and xylanase [ 115 ]. In general, SmF is the preferred method of production when the preparations require more purified enzymes, whereas synergistic effects from a battery of xylan-degrading enzymes can easily be found in preparations obtained by SSF using complex substrates, though the latter is commonly sought in applications aimed at improving animal feed [ 113 ].

The choice of the substrate is of great importance for the selection of the fermentation process and the successful production of xylanases. In this context, purified xylans can be excellent substrates because the low molecular weight compounds derived from them are the best xylanase inducers. The use of such substrates has led to increased yields of xylanase production and a selective induction of xylanases, with concomitantly low cellulase activity in a number of microorganisms. However, for large-scale processes other alternatives have to be considered due to the cost of such substrates. Some lignocellulolytic substrates such as barley husk, corn cobs, hay, wheat bran or straw have been compared in relation to pure substrates, and many have performed significantly better than isolated xylans (or celluloses) with respect to the yields of xylanase in large-scale production processes. Solid-state fermentation processes are practical for complex substrates, including agricultural, forestry and food processing residues and wastes, which are used as inducing carbon sources for the production of xylanases [ 113 ]. The use of abundantly available and cost-effective agricultural residues, such as wheat bran, corn cobs, rice bran, rice husks, and other similar substrates, to achieve higher xylanase yields via SSF allows the reduction of the overall manufacturing cost of biobleached paper. This has facilitated the use of this environmentally friendly technology in the paper industry [ 1 ].

3.6. Cloning and expression of xylanases

To meet specific industrial needs, an ideal xylanase should have specific properties, such as stability over a wide range of pH values and temperatures, high specific activity, and strong resistance to metal cations and chemicals [ 116 ]. Other specifications include cost-effectiveness, eco-friendliness, and ease of use [ 32 ]. Therefore, most of the reported xylanases do not possess all of the characteristics required by industry [ 8 ].

Native enzymes are not sufficient to meet the demand, due to low yields and incompatibility of the standard industrial fermentation processes [ 35 ]. Therefore, molecular approaches must be implemented to design xylanases with the required characteristics [ 8 ]. Heterologous expression is the main tool for the production of xylanases at the industrial level [ 35 ]. Protein engineering (alteration or modification of existing proteins) by recombinant DNA technology could be useful in improving the specific characteristics of existing xylanases [ 8 ]. Genetic engineering and recombinant DNA technology allow the large-scale expression of xylanases in homologous or heterologous protein-expression hosts. As industrial applications require cheaper enzymes, the elevation of expression levels and efficient secretion of xylanases are vital for ensuring the viability of the process [ 23 ].

An increasing number of publications have described numerous xylanases from several sources and the cloning, sequencing, mutagenesis and crystallographic analysis of these enzymes [ 12 ]. The available amino acid sequence data, X-ray crystallographic data, molecular dynamics and computational design of xylanases provide information that authenticates the relationship between the structure and function of xylanases. All of these methods aid in the design of xylanases that are required in industrial processes, such as improvement of the stability of xylanases at higher temperatures and alkaline pHs [ 8 ].

To attempt these processes for commercial purposes, genes encoding several xylanases have been cloned in homologous and heterologous hosts [ 12 , 48 ]. Recombinant xylanases have shown equivalent or better properties than the native enzymes, and the xylanase genes from anaerobic microorganisms have also been expressed successfully in hosts that can be employed in the fermentation industry [ 35 ].

3.6.1. Expression in bacteria

Escherichia coli is known for its ease of manipulation, inexpensive growth conditions, simple techniques required for transformation and accumulation of high levels of product in the cell cytoplasm; therefore, this organism has become the most widely used expression host [ 117 ]. Despite Escherichia coli’s use as a good cloning host for recombinant proteins, it does not provide efficient and functional expression of many xylanases [ 23 , 24 ], and not all genes are easily expressed in E. coli [ 117 ]. This problem may be due to the repetitive appearance of rare codons and the requirement for specific translational modifications, such as disulfide-bond formation and glycosylation [ 23 ]. Therefore, this microorganism is useful for the detailed study of xylanase gene structure and for the improvement of the enzymes via protein engineering [ 35 ].

Lactobacillus species and Bacillus subitilis have been attractive hosts for the production of heterologous proteins, obtaining higher expression levels than E. coli [ 23 , 118 ]. B. subtilis and Lactobacillus are gram-positive and perform N-glycosilation [ 119 ]. Their primary interest in industry and research, is due to the fact that are non-toxic and are generally recognized as safe (GRAS) [ 23 , 118 ]. Members of the genus Bacillus , unlike E. coli , do not contain endotoxins (lipopolysaccharides), which are difficult to remove from many proteins during the purification process. The secretory production could also be advantageous in industrial production [ 33 ].

3.6.2. Expression in yeast

Heterologous protein expression in yeast systems is highly attractive because they provide additional benefits over bacterial expression systems. Among these benefits are the ability to perform eukaryotic post-translational modifications, the ability to grow to very high cell densities and the ability to secrete proteins into the fermentation media. Moreover, yeast are free of toxins and the majority have GRAS status [ 23 ].

Saccharomyces cerevisiae secretes high amounts of xylanases into the culture medium. Because it has already been established as an industrial microorganism, it can be used conveniently for the industrial production of xylanases at low costs [ 35 ]. Pichia pastoris has also emerged as an excellent host for the commercial production of xylanases due to very high expression under its own promoters [ 35 ]. However, the success of this methylotrophic yeast, similar to Hansenula polymorpha , is reached with the promoters of alcohol oxidase, an enzyme involved in the methanol-utilization pathway [ 23 ]. Therefore, these promoters have limited use at the large scale due to the health and fire hazards of methanol [ 35 ].

3.6.3. Expression in filamentous fungi

Filamentous fungi are capable producers of xylanases, via both heterologous and homologous gene expression, and reach high expression yields with their own promoters [ 35 ]. Filamentous fungi have already undergone intricate strain improvement for high-level protein secretion and are feasible when using the native xylanase-expressing machinery for functional expression of foreign xylanases from remote sources. The xylanase gene from P. griseofulvum has been successfully expressed in A. oryzae [ 120 ].

4. Conclusion

Xylan, the major hemicellulose component, requires the synergistic action of several hemicellulase enzymes for its complete hydrolysis to monomer sugars. The principle enzyme in this processes is endo-1,4-β-xylanase, which cleaves the glycosidic bonds between xylosides, generating short xylooligosaccharides. The majority of the studied xylanases have been classified into the GH10 or GH11 families, whereas studies of the xylanases in families 5, 7, 8 and 43 are still emerging.

The conversion of xylan to useful products represents part of our efforts to strengthen the overall economics of the processing of lignocellulosic biomass and to develop new means of energy production from renewable resources. Among these products are xylanases, enzymes that have a wide range of important industrial applications. Therefore, in the future, new methods will be developed for easier and cheaper production of these enzymes to fulfill the demands of various industries. In this context, the use of lignocellulosic agricultural waste for the production of these enzymes by either submerged or solid-sate fermentation has been very attractive, in addition to molecular techniques that are being tested to improve the enzyme’s characteristics and increase its expression rates. Moreover, as the native enzyme does not fulfill all of the process requirements, bioprospecting for new genes, rational engineering and directed evolution of known genes are powerful tools that can be used to improve these enzymes.

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Article Contents

1 introduction, 2 the substrate: xylan, 3 xylanases: multiplicity and multiple-domains, 4 classification of xylanases, 5 glycoside hydrolase families 5, 7, 8, 10, 11 and 43, 6 extremophilic xylanases, 7 application of xylanases: extremophilic xylanases, 8 concluding remarks, acknowledgements.

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Xylanases, xylanase families and extremophilic xylanases

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Tony Collins, Charles Gerday, Georges Feller, Xylanases, xylanase families and extremophilic xylanases, FEMS Microbiology Reviews , Volume 29, Issue 1, January 2005, Pages 3–23, https://doi.org/10.1016/j.femsre.2004.06.005

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Xylanases are hydrolytic enzymes which randomly cleave the β 1,4 backbone of the complex plant cell wall polysaccharide xylan. Diverse forms of these enzymes exist, displaying varying folds, mechanisms of action, substrate specificities, hydrolytic activities (yields, rates and products) and physicochemical characteristics. Research has mainly focused on only two of the xylanase containing glycoside hydrolase families, namely families 10 and 11, yet enzymes with xylanase activity belonging to families 5, 7, 8 and 43 have also been identified and studied, albeit to a lesser extent. Driven by industrial demands for enzymes that can operate under process conditions, a number of extremophilic xylanases have been isolated, in particular those from thermophiles, alkaliphiles and acidiphiles, while little attention has been paid to cold-adapted xylanases. Here, the diverse physicochemical and functional characteristics, as well as the folds and mechanisms of action of all six xylanase containing families will be discussed. The adaptation strategies of the extremophilic xylanases isolated to date and the potential industrial applications of these enzymes will also be presented.

Xylanases are glycosidases ( O -glycoside hydrolases, EC 3.2.1.x) which catalyze the endohydrolysis of 1,4-β- d -xylosidic linkages in xylan. They are a widespread group of enzymes, involved in the production of xylose, a primary carbon source for cell metabolism and in plant cell infection by plant pathogens, and are produced by a plethora of organisms including bacteria, algae, fungi, protozoa, gastropods and anthropods [1] . First reported in 1955 [2] , they were originally termed pentosanases, and were recognized by the International Union of Biochemistry and Molecular Biology (IUBMB) in 1961 when they were assigned the enzyme code EC 3.2.1.8. Their official name is endo-1,4-β-xylanase, but commonly used synonymous terms include xylanase, endoxylanase, 1,4-β- d -xylan-xylanohydrolase, endo-1,4-β- d -xylanase, β-1,4-xylanase and β-xylanase.

In the present review, the diversity of xylanases, their substrate, action and function, their importance in industry, classification into families and adaptation to various extreme environments are discussed. Special emphasis is paid to the ‘new’ xylanase containing families, highlighting their similarities and differences to the better known family 10 and 11 members, as well as to the peculiarities and interests of these hitherto scantily reviewed enzymes. In addition, the adaptation strategies, characteristics and industrial potential of extremophilic xylanases will be discussed.

The substrate of xylanases, xylan, is a major structural polysaccharide in plant cells, and is the second most abundant polysaccharide in nature, accounting for approximately one-third of all renewable organic carbon on earth [1] . Xylan constitutes the major component of hemicellulose; a complex of polymeric carbohydrates including xylan, xyloglucan (heteropolymer of d -xylose and d -glucose), glucomannan (heteropolymer of d -glucose and d -mannose), galactoglucomannan (heteropolymer of d -galactose, d -glucose and d -mannose) and arabinogalactan (heteropolymer of d -galactose and arabinose) [3] . This, together with cellulose (1,4-β-glucan) and lignin (a complex polyphenolic compound) make up the major polymeric constituents of plant cell walls [4] . Within the cell wall structure, all three constituents interact via covalent and non-covalent linkages, with the xylan being found at the interface between the lignin and cellulose where it is believed to be important for fiber cohesion and plant cell wall integrity [5] .

Xylan is found in large quantities in hardwoods from angiosperms (15–30% of the cell wall content) and softwoods from gymnosperms (7–10%), as well as in annual plants (<30%) [6] . It is typically located in the secondary cell wall of plants, but is also found in the primary cell wall, in particular in monocots [7] . A complex, highly branched heteropolysaccharide, it varies in structure between different plant species, and the homopolymeric backbone chain of 1,4-linked β- d -xylopyranosyl units can be substituted to varying degrees with glucuronopyranosyl, 4- O -methyl- d -glucuronopyranosyl, α- l -arabinofuranosyl, acetyl, feruloyl and/or p -coumaroyl side-chain groups [ 4 , 8 ] ( Fig. 1 ). Wood xylan exists as O -acetyl-4- O -methylglucuronoxylan in hardwoods and as arabino-4- O -methylglucuronoxylan in softwoods, while xylans in grasses and annual plants are typically arabinoxylans [4] . Linear unsubstituted xylan has also been reported, e.g., in esparto grass [9] , tobacco [10] and certain marine algae [ 11 , 12 ], with the latter containing xylopyranosyl residues linked by both 1,3-β and 1,4-β linkages [ 12 , 13 ]. The degree of polymerisation in xylans is also variable, with, for example, hardwood and softwood xylans generally consisting of 150–200 and 70–130 β-xylopyranose residues, respectively [4] .

(a) Structure of xylan and the sites of its attack by xylanolytic enzymes. The backbone of the substrate is composed of 1,4- β-linked xylose residues. Ac., Acetyl group; α-araf., α-arabinofuranose; α-4- O -Me-GlcUA, α-4- O -methylglucuronic acid; pcou., p -coumaric acid; fer., ferulic acid. (b) Hydrolysis of xylo-oligosaccharide by β-xylosidase. Adapted from [18] . Figures were prepared with CS Chemdraw Ultra version 6.0.

Due to its heterogeneity and complexity, the complete hydrolysis of xylan requires a large variety of cooperatively acting enzymes [ 14–16 ] ( Fig. 1 ). Endo-1,4-β- d -xylanases (EC 3.2.1.8) randomly cleave the xylan backbone, β- d -xylosidases (EC 3.2.1.37) cleave xylose monomers from the non-reducing end of xylo-oligosaccharides and xylobiose while removal of the side groups is catalysed by α- l -arabinofuranosidases (EC 3.2.1.55), α- d -glucuronidases (EC 3.2.1.139), acetylxylan esterases (EC 3.1.1.72), ferulic acid esterases (EC 3.1.1.73) and p -coumaric acid esterases (EC 3.1.1.-). Indeed, complete xylanolytic enzyme systems, including all of these activities, have been found to be quite widespread among fungi [ 17 , 18 ], actinomycetes [19] and bacteria [18] , and some of the most important xylanolytic enzyme producers include the Aspergilli, Trichodermi, Streptomycetes, Phanerochaetes, Chytridiomycetes, Ruminococci, Fibrobacteres, Clostridia and Bacilli [ 16 , 18 , 20 , 21 ]. The ecological niches of these micro-organisms are diverse and widespread and typically include environments where plant material accumulate and deteriorate, as well as in the rumen of ruminants [ 1 , 16 , 22 ].

In addition to the production of a variety of xylanolytic enzymes, many micro-organisms produce multiple xylanases [ 23–25 ]. These may have diverse physicochemical properties, structures, specific activities and yields, as well as overlapping but dissimilar specificities, thereby increasing the efficiency and extent of hydrolysis, but also the diversity and complexity of the enzymes. Typical examples of micro-organisms which produce xylanase isoenzymes include Aspergillus niger , which produces fifteen extracellular xylanases [26] , and Trichoderma viride , which secretes thirteen [26] . This multiplicity may be the result of genetic redundancy [7] , but cases of differential post-translational processing have also been reported [15] . The isoenzyme genes may be found as polycistronic or non-polycistronic multiple copies within the genome, and in some cases several xylanases are expressed as a distinct gene product. For example, the xylanase, β-xylosidase and acetyl esterase genes of Caldocellum saccharolyticum (now known as Caldocellulosiruptor saccharolyticus ) are polycistronic [27] , while the XynC gene product from Fibrobacter succinogenes S85 encodes two different xylanase catalytic domains [28] . Moreover, as well as multiple catalytic domains, many xylanases are also characterized by the presence of various supplementary domains. Examples include xylan binding domains [29] , cellulose binding domains [ 30 , 31 ], dockerin domains (implicated in binding to multidomain complexes produced by certain micro-organism, e.g., Clostridium thermocellum ) [ 32 , 33 ], thermostabilising domains [34] and domains for which the function has not as yet been elucidated. These domains may fold and function in an independent manner [ 35 , 36 ] and are typically separated by short junction segments enriched in hydroxyl amino acids [4] .

The vast majority of xylanases are excreted into the extracellular environment as the large size of the substrate prevents its penetration into the cell. In fact, the current belief is that xylanase production is induced by means of the products of their own action [ 6 , 15 , 37 ]. It is believed that small amounts of constitutively produced enzymes liberate xylo-oligomers which may be transported into the cell where they are further degraded by β-xylosidases, or indeed by intracellular xylanases [ 38–40 ], and where they induce further xylanase synthesis.

The heterogeneity and complexity of xylan has resulted in an abundance of diverse xylanases with varying specificities, primary sequences and folds, and hence has lead to limitations with the classification of these enzymes by substrate specificity alone. Wong et al. [7] classified xylanases on the basis of their physicochemical properties and proposed two groups: those with a low molecular weight (<30 kDa) and basic p I , and those with a high molecular weight (>30 kDa) and acidic p I . However, several exceptions to this pattern have been found [ 18 , 21 ] and approximately 30% of presently identified xylanases, in particular fungal xylanases, cannot be classified by this system.

Later, a more complete classification system was introduced [41] which allowed the classification of not only xylanases, but glycosidases in general (EC 3.2.1.x), and which has now become the standard means for the classification of these enzymes. This system is based on primary structure comparisons of the catalytic domains only and groups enzymes in families of related sequences [42] . The initial classification grouped cellulases and xylanases into 6 families (A–F) [41] , which was updated to 77 families in 1999 (1–77) [42] and which continues to grow as new glycosidase sequences are identified. At the time of writing, 96 glycoside hydrolase families exist (see the carbohydrate–active enzyme CAZY server at http://afmb.cnrs-mrs.fr/˜cazy/CAZY/ [43] ), with approximately one-third of these families being polyspecific, i.e., contain enzymes with diverse substrate specificities. As the structure and molecular mechanism of an enzyme are related to its primary structure, this classification system reflects both structural and mechanistic features. Enzymes within a particular family have a similar three-dimensional structure [42] and similar molecular mechanism [44] and it has also been suggested that they may have a similar specificity of action on small, soluble, synthetic substrates [45] . Furthermore, divergent evolution has resulted in some of the families having related three-dimensional structures and thus the grouping of families into higher hierarchical levels, known as clans, has been introduced [46] . Presently, 14 different clans have been proposed (GH-A to GH-N), with most clans encompassing two to three families, apart from clan GH-A which currently encompasses 17 families.

Within this classification system, xylanases are normally reported as being confined to families 10 (formerly F) and 11 (formerly G) [ 6 , 16 , 18 , 47–49 ]. Interestingly, a search of the appropriate databases (e.g., CAZY [43] ) using the enzyme classification number EC 3.2.1.8 indicates that enzymes with xylanase activity are also found in families 5, 7, 8, 16, 26, 43, 52 and 62. A closer look at the available literature however, shows that only those sequences classified in families 5, 7, 8, 10, 11 and 43 ( Table 1 ) contain truly distinct catalytic domains with a demonstrated endo-1,4-β-xylanase activity. Those sequences reported for families 16, 52 and 62 appear, in fact, to be bifunctional enzymes containing two catalytic domains; a family 10 or 11 xylanase domain as well as a second glycosidase domain. For example, a Ruminococcus flavefaciens enzyme contains an amino-terminal family 11 xylanase and a carboxy-terminal family 16 lichenase and is thus classified in both families 11 and 16 [50] . In addition, those enzymes classified in family 26 appear not to be endo-1,4-β-xylanases, but endo-1,3-β-xylanases. Thus, the current view that enzymes with xylanase activity are solely restricted to families 10 and 11 is not entirely correct and should be expanded to include families 5, 7, 8 and 43.

Glycoside hydrolase families containing enzymes with a demonstrated activity on xylan. The fold, mechanism of action and catalytic residues characteristic to each family are given

a Putative catalytic residues only, these have not been conclusively confirmed.

5.1 Catalytic mechanisms

Members of families 5, 7, 8, 10, 11 and 43 differ in their physico-chemical properties, structure, mode of action and substrate specificities. Similarities do however exist, for example, families 5 and 10 are both classified in clan GH-A, thus indicating a similar three-dimensional fold. Furthermore, families 5, 7, 10 and 11 contain enzymes which catalyse hydrolysis with retention of anomeric configuration with two glutamate residues being implicated in the catalytic mechanism in all cases [43] . This indicates a double-displacement mechanism, in which a covalent glycosyl-enzyme intermediate is formed and subsequently hydrolysed via oxocarbenium-ion-like transition states [ 51–53 ] ( Fig. 2(a) ). Two carboxylic acid residues suitably located in the active site (approximately 5.5 Å apart) are involved in the formation of the intermediate; one acts as a general acid catalyst by protonating the substrate, while the second performs a nucleophilic attack which results in the departure of the leaving group and the formation of the α-glycosyl enzyme intermediate (inversion β to α). In the second step, the first carboxylate group now functions as a general base, abstracting a proton from a nucleophilic water molecule which attacks the anomeric carbon. This leads to a second substitution in which the anomeric carbon again passes via an oxocarbenium-ion-like transition state to give rise to a product with the β configuration (inversion α to β). Thus the overall result is a retention of the configuration at the anomeric centre.

General mechanisms for (a) retaining and (b) inverting glycosidases. Adapted from [51] . Figures were prepared with CS Chemdraw Ultra version 6.0.

In contrast, enzymes in families 8 and 43 typically operate with inversion of the anomeric centre and a glutamate and aspartate are believed to be the catalytic residues [ 43 , 54 ]. Inverting enzymes function by a single displacement reaction ( Fig. 2(b) ) in which one carboxylate provides for a general acid-catalyzed leaving group departure and the second functions as a general base, activating a nucleophilic water molecule to attack the anomeric carbon, thereby cleaving the glycosidic bond and leading to an inversion of the configuration at the anomeric carbon [ 51–53 ]. Typically the distance between the two residues is around 9.5 Å so as to allow for accommodation of the water molecule between the anomeric carbon and the general base [ 51–53 ]. However, Alzari et al. [55] and Guérin et al. [56] have shown that this distance is 7.5 Å in the inverting endoglucanase CelA and have suggested that the distance between the two catalytic residues is less constrained in inverting than in retaining enzymes.

5.2 Glycoside hydrolase family 5

At the time of writing, family 5 (formerly family A) of the glycoside hydrolases consists of 467 sequences with varying activities, including: endoglycosylceramidase (EC 3.2.1.123), cellulase (EC 3.2.1.4), licheninase (EC 3.2.1.73), β-mannosidase (EC 3.2.1.25), glucan 1,3-β-glucosidase (EC 3.2.1.58), glucan endo-1,6-β-glucosidase (EC 3.2.1.75), mannan endo-1,4-β-mannosidase (EC 3.2.1.58), cellulose 1,4-β-cellobiosidase (EC 3.2.1.91), endo-1,6-β-galactanase (EC 3.2.1.-), 1,3-β-mannanase (EC 3.2.1.-) and endo-1,4-β-xylanase (EC 3.2.1.8) [43] . This is the largest glycoside hydrolase family and only seven amino acid residues, including the nucleophile and the general acid/base residue, are strictly conserved among all members. It is a rather diverse group of enzymes, with structural alignments indicating rms deviations of 1.25 ± 0.12 Å between equivalent residues in its members [57] and a further classification of this family into nine subfamilies has even been suggested [58] .

Eight enzymes with a demonstrated activity on xylan have been reported in this family to date ( Table 2 ) and a number of other putative xylanases, identified by sequence similarity during genome sequencing programs, have been identified in Clostridium acetobutylicum ATCC 824 (2 putative xylanases) [59] , Leptosphaeria maculans [43] , Xanthomonas axonopodis pv. citri str. 306 (2 putative xylanases) [60] , Xanthomonas campestris pv. campestris str. ATCC 33913 [60] , Bacillus subtilis str. 168 [61] and Bacteroides thetaiotaomicron VPI-5482 [62] . Xylanase activity has also been apparently demonstrated for enzymes from Aeromonas puncata ( caviae ) W-61, Erwinia ( Pectobacterium ) chrysanthemi P860219, Meloidogyne incognita and Ruminococcus albus 7 [43] but it appears that no literature has been published in direct relation to these. In total, 20 putative xylanolytic entries can be proposed.

Organisms producing family 5 enzymes with a demonstrated activity on xylan

Although characterisation of these enzymes is incomplete, large variations in the catalytic properties are already evident, this being in keeping with the poor sequence identity between some of these enzymes. Those enzymes from Prevotella ruminicola 23 [63] , Clostridium cellulovorans [64] , F. succinogenes S85 [65] and R. albus SY3 [66] were all found to have carboxymethyl cellulase as well as xylanase activities, while those from Trichoderma reesei (also known as Hypocrea jecorina ) [ 67 , 68 ], Erwinia chrysanthemi D1 [ 57 , 69–71 ] and E. chrysanthemi SR120A [69] appear to be specific for xylan. XynD from Aeromonas punctata ME-1 was found to be active on xylan but was not apparently tested on other substrates [72] . Furthermore, a search of the InterPRO database ( http://www.ebi.ac.uk/interpro , [73] ) indicates that only the four carboxymethyl cellulose active enzymes (i.e., those from P. ruminicola 23, C. cellulovorans, F. succinogenes S85 and R. albus SY3) give hits with family 5 signatures; the remaining 16 sequences give hits with family 30 or with other non-glycoside hydrolase family signatures. Moreover, these four enzymes have relatively high amino acid sequence similarity with family 5 members, while the majority of the remaining sequences have homology with both family 5 and family 30 enzymes. This has already been noted for the E. chrysanthemi and A. punctata ME-1 enzymes where it was suggested that these enzymes be classified into a new family (or subfamily) situated between families 5 and 30 [ 70 , 72 ]. In addition, structural analysis of the family 5 xylanase, XynA from E. chrysanthemi showed that, as would be expected for a family 5 enzyme, the catalytic domain displayed a common (β/α) 8 barrel fold ( Fig. 3(a) ) [57] . However, while the β-barrels aligned well with those of another family 5 enzyme, the α-helices and loops were altered, showing differences in the positioning, orientation and length. Moreover, structural alignment with a number of family 5 and 10 enzymes showed that this enzyme is nearly as structurally different to the family 5 enzymes as are the family 10 enzymes [57] .

Representative structures of enzymes from various glycoside hydrolase families. (a) Structure of the family 5 enzyme, XynA, from Erwinia chrysanthemi . The (β/α) 8 barrel structure of the catalytic domain and the β 9 -barrel of the small domain are shown [57] . (b) Structure of the family 8 xylanase, pXyl, from Pseudoalteromonas haloplanktis TAH3a. The (α/α) 6 barrel structure of the catalytic domain is shown in two perpendicular views [82] . (c) Structure of the Streptomyces lividans xylanase showing the typical family 10-fold [95] . (d) Structure of the Trichoderma reesei family 11 xylanase showing the typical family 11-fold [127] . (e) Structure of the Trichoderma reesei family 7 non-specific EGI [131] . (f) Structure of the Cellvibrio japonicus family 43 α- l -arabinanase [54] . Figures were prepared with Swiss-Pdb Viewer v3.7b2 [205] .

Of the four family 5 enzymes exhibiting carboxymethyl cellulase and xylanase activities, only the P. ruminicola 23 xylanase was found to have highest activity on xylan, with the carboxymethyl cellulase activity being only 18% of this [63] . In contrast, EngB from C. cellulovorans was found to be most active on lichenan, with approximately 15% activity on carboxymethylcellulose and 14% on xylan. No activity was detected on microcrystalline cellulose, avicel, polygalacturonic acid, mannan, laminaran, p -nitrophenyl β-cellobioside, p -nitrophenyl β- d -glucoside or p -nitrophenyl β- d -xyloside [64] . Unfortunately, as with the P. ruminicola 23 xylanase, the endoglucanase isolated from Fibrobacter succinogenes S85 was not tested with lichenan, this enzyme was found to have highest activity on carboxymethyl cellulose, as well as activity on oat spelt xylan (71% as compared to CMC), p -nitrophenyl cellobioside (5.3%), cellobiose (3.5%), p -nitrophenyl lactoside (2.27%) and p -nitrophenyl glucoside (0.27%) [65] .

The action patterns of the xylan specific family 5 enzymes with sequence similarities to family 30 enzymes also appear to vary from one to another. Only those enzymes from A. punctata ME-1 [72] , E. chrysanthemi D1 [ 57 , 70 , 71 ] and T. reesei [ 67 , 68 ] have been characterized in this respect, and while the first does appear to be an endoxylanase, producing xylotriose and higher xylo-oligosaccharides from birchwood xylan, the second appears to be a type of ‘appendage-dependent xylanase’ or glucuronoxylanase [74] and the third an exoxylanase. Substrate specificity studies showed that the xylanase from E. chrysanthemi D1 requires d -glucuronic acid (or its 4- O -methyl derivative) substituents on the xylan backbone to enable cleavage of 1,4-β xylosidic bonds and that it cleaves between the first and second xylose residues on the non-reducing terminal side of the substituted residue [71] . On the other hand, XYN IV from T. reesei is active on unsubstituted and substituted xylans and attacks at the first glycosidic linkage from the reducing end, producing mainly xylose, but also xylobiose and low amount of higher xylo-oligosaccharides [ 67 , 68 ]. Furthermore, the activity of this enzyme is affected by substituents on the xylan main chain and it is unable to cleave linkages next to substituted residues. Hydrolysis studies have shown that the shortest substituted fragments formed from glucuronoxylan and arabinoxylan are substituted xylotrioses, with the substitution being found on the internal xylose residue. Indeed these products are shorter than those produced by the family 7 EGI [75] produced by the same micro-organism as well as the family 11 enzymes, but, in agreement with members of this latter family and with the family 5 XynA from E. chrysanthemi D1 they contain a single xylose at the non-reducing end.

Of the above mentioned enzymes, only the structure of Xyn A from E. chrysanthemi has been determined (1.42 Å resolution) [57] while crystallisation and preliminary X-ray analysis (at 2.2 Å resolution) of the T. reesei XYN IV has also recently been reported [76] . Xyn A is composed of two domains, the larger domain contains the catalytic site and displays a (β/α) 8 barrel fold while the small domain probably functions as a xylan binding domain and has a β 9 -barrel fold ( Fig. 3(a) ). The two domains are connected by two linker peptides as well as 11 hydrogen bonds and hydrophobic interactions [57] . The β-barrel of the catalytic domain of Xyn A is elliptical in shape and the active site is formed by an acidic cleft situated on the carboxy-terminal side of the β-strands near the larger face of the molecule. This 8-fold α/β barrel structure is indeed the most frequently encountered fold and has also been described for members of families 1, 2, 10, 17, 26, 30, 35, 39, 42, 50, 51, 53, 59, 72, 79 and 86 [43] . This type of fold was originally described for triose-phosphate isomerase (TIM barrel) and as a common characteristic of all these structures is the positioning of the glutamate residues implicated in the catalysis on the carboxy-terminal of β-sheets 4 and 7, these were originally termed the 4/7 superfamily. Nowadays they are better known as clan GH-A.

5.3 Glycoside hydrolase family 8

Family 8 (formerly family D) is mainly composed of cellulases (EC 3.2.1.4), but also contains chitosanases (EC 3.2.1.132), lichenases (EC 3.2.1.73) and endo-1,4-β-xylanases (EC 3.2.1.8) [43] . It is a rapidly expanding family; from 18 members, including one xylanase, in 2001, to 61 members, including four xylanases, in 2004 ( Table 3 ). Three of the xylanases have been isolated from Bacillus sp. while the fourth is a cold-adapted enzyme isolated from the Antarctic bacterium Pseudoalteromonas haloplanktis TAH3a. Xylanase Y from the alkaliphile Bacillus halodurans C-125 was identified as part of the sequencing program of the whole genome of this organism [ 77 , 78 ] and the sequence of the xylanase from B. halodurans MIR32 is found to be identical to that for this enzyme [43] . Information is minimal and while we have demonstrated xylanase activity by these enzymes [79] , little literature has been published in direct relation to these two xylanases. Indeed the means used for their initial classification as xylanases is unclear and was probably only based on their weak isology to the cold-adapted xylanase and xylanase Y from Bacillus sp. KK-1. In contrast, the biochemical properties and substrate specificities of these latter two enzymes have been investigated [ 80–84 ]. Both have a high molecular weight (∼46 and ∼45 kDa, respectively) and while the cold-adapted xylanase also has a high p I (∼pH 9.5), this was not determined for the Bacillus sp. KK-1 xylanase. In addition, both were found to be optimally active at pH 6.5 and are solely active on xylan, being more active on birchwood than oat spelt xylan and being inactive on cellulose, carboxymethylcellulose, starch, lichenan and chitosan (this latter substrate was only tested for the cold-adapted xylanase). The cold-adapted xylanase was found to hydrolyse xylan to principally xylotriose and xylotetraose and was most active on long chain xylo-oligsaccharides. Similar to family 11 xylanases, a large substrate binding cleft containing at least six xylose binding residues, with the catalytic site in the middle, was proposed [80] . However, unlike family 10 and 11 xylanases this enzyme was found to catalyse hydrolysis with inversion of the anomeric configuration and, under the conditions used, was found to be inactive on aryl-β-glycosides of xylose, xylobiose and xylotriose.

Organisms producing family 8 enzymes with a demonstrated xylanase activity

This cold-adapted xylanase folds into a distorted (α/α) 6 barrel ( Fig. 3(b) ) formed by six inner and six outer α helices [ 82 , 83 ] and as such can be classified with family 48 in clan GH-M [43] . This topology has also been observed for family 9 endoglucanases, family 15 glucoamylases, family 48 cellobiohydrolases and a family 65 maltose phosphorylase [ 55 , 85 , 86 ]. However, in contrast to the common (α/α) 6 barrel proteins, the cold-adapted enzyme has an extra α-helix near the amino terminus. The globular core has an overall distorted spherical shape with a long acidic cleft running across the molecular surface at the N-terminal end of the inner helices while the proposed catalytic residues (glutamate and aspartate) are located close to each other near the middle of the cleft.

As the three-dimensional structure is believed to be comparable for members of the same family, the (α/α) 6 barrel fold described above can probably be extended to the other family 8 xylanases. Furthermore, one would expect these enzymes to operate with inversion of the anomeric configuration [ 80 , 87 ], with an aspartic and glutamic acid residue as proton acceptor and proton donor, respectively [ 56 , 83 ].

5.4 Glycoside hydrolase family 10

This family consists of endo-1,4-β-xylanases (EC 3.2.1.8), endo-1,3-β-xylanases (EC 3.2.1.32) and cellobiohydrolases (EC 3.2.1.91) [43] . The major enzymes of this family are endo-1,4-β-xylanases, however, substrate specificity studies have revealed that these may not be entirely specific for xylan and may also be active on low molecular mass cellulose substrates [ 88 , 89 ], in particular on aryl-cellobiosides [ 90 , 91 ] and certain cello-oligosaccharides [ 45 , 88 ]. In effect, it has been found that the replacement of one or two xylose residues by glucose is normally tolerated by the xylanases of this family, with this generally resulting in a lowered catalytic efficiency [90] . In consonance with family 11 xylanases, but in contrast to the cold-adapted family 8 xylanase, members of this family are also capable of hydrolyzing aryl β-glycosides of xylobiose and xylotriose at the aglyconic bond [ 90 , 92 , 93 ]. Furthermore, these enzymes are highly active on short xylo-oligosaccharides, thereby indicating small substrate binding sites [90] . In effect, crystal structure analyses, kinetic analyses of activity on xylo-oligosaccharides of various sizes and end product analyses have indicated that family 10 xylanases typically have four to five substrate binding sites [ 90 , 94 , 95 ]. Hydrolysis studies have also shown that most family 10 xylanases can attack the xylosidic linkage on the non-reducing end of a substituted residue or 1,3-β bond, but can only cleave at the third xylosidic linkage after a substituted residue and the second after a 1,3-β bond [90] . This indicates that the subsites on the non-reducing side (i.e., subsites −1, −2) are more specific than those on the reducing side (subsite +1) of the cleavage site.

Members of this family typically have a high molecular mass, a low p I and display an (α/β) 8 barrel fold [ 40 , 43 , 58 , 95–105 ] ( Fig. 3(c) ) ( Table 4 ). The structure has been likened to a ‘salad bowl’, with one face of the molecule having a large radius (approximately 45 Å) due to an elaborate loop architecture, while the opposite face, which consists of simple α/β turns, has a radius of approximately 30 Å. This is similar to the fold described for family 5 enzymes and both are members of clan GH-A. Indeed, these two families are quite closely related and in addition to sharing a common fold they have the same type of catalytic mechanism and share several common residues [ 57 , 106 ]. However, the family 10 xylanases are a more closely related family and have a high percentage of spatially equivalent and identical residues as well as much smaller rms deviations between equivalent residues (0.95 ± 0.11 Å) in its members [57] .

Family 10 xylanases for which structural coordinates are available

5.5 Glycoside hydrolase family 11

In contrast to all other families hitherto discussed, this family is monospecific, it consists solely of xylanases. Moreover, these xylanases are ‘true xylanases’ as they are exclusively active on d -xylose containing substrates. They have a lower catalytic versatility than family 10 xylanases and indeed the products of their action can be further hydrolyzed by the family 10 enzymes [ 90 , 92 ]. Like family 10 xylanases, these enzymes can hydrolyze aryl β-glycosides of xylobiose and xylotriose at the aglyconic bond, but in contrast to this family they are inactive on aryl cellobiosides. Furthermore, substituents or β 1,3 linkages represent a more serious hindrance to their activity, resulting in the production of larger products than family 10 xylanases [90] . Hydrolysis studies indicate that aldopentauronic acid, with an unsubstituted xylose residue at the non-reducing end [ 90 , 107 ], and an isomeric xylotetraose, with the 1,3-β bond at the non-reducing end [108] , are the smallest acidic and mixed linkage fragments liberated from heteroxylans and rhodymenan, respectively. Others have, however, suggested that this isomeric xylotetraose may contain a 1,4-β bond at the non-reducing end, with the 1,3-β linkage occurring subsequent to this bond [ 90 , 109 ]. In further contrast to the family 10 xylanases, but in common with the family 8 cold-adapted xylanase, these enzymes are most active on long chain xylo-oligosaccharides and indeed it has been found that they have larger substrate binding clefts, e.g., family 11 xylanases from Schizophyllum commune and A. niger have at least seven subsites [ 110 , 111 ], while, as has already been stated, family 10 enzymes are reported to have four to five subsites [ 90 , 94 , 95 ]. Further differences between family 10 and 11 xylanases include their stereochemistry of protonation and the effect of ω -epoxyalkyl glycosides of xylose and xylo-oligosaccharides on their activity; family 10 xylanases are anti -protonators and are unaffected by the ω -epoxyalkyl glycosides while family 11 xylanases are syn -protonators and are inactivated by the ω -epoxyalkyl glycosides [ 88 , 112 , 113 ].

Family 11 enzymes are generally characterized by a high p I , a low molecular weight, a double displacement catalytic mechanism, two glutamates acting as the catalytic residues and a β-jelly roll fold structure [ 114–127 ] ( Table 5 ) ( Fig. 3(d) ). The structure consists principally of β-pleated sheets formed into a two-layered trough that surrounds the catalytic site. Two [48] , or perhaps three [128] , β-sheets are present and the hydrophobic faces of these are packed against each other to form the hydrophobic core of the protein. Only one α-helix is present and this is typically packed against the hydrophobic face of the second β-sheet. This type of structure has also been described for family 12 endoglucanases and thus both families have been grouped into the same clan, clan GH-C.

Family 11 xylanases for which structural coordinates are available

5.6 Glycoside hydrolase families 7 and 43

To date, only one enzyme exhibiting xylanase activity has been identified and studied in each of these families and thus their importance as xylanase containing families is unclear. In addition, neither enzyme studied is a true xylanase; the family 7 enzyme, EGI (Cel7B) from T. reesei , is a non-specific endo-β-1,4-glucanase (EC 3.2.1.4) [ 75 , 129–131 ] and the family 43 XYND from Paenibacillus polymyxa has both xylanase and α- l -arabinofuranosidase activities [132] .

The family 7 endoglucanase I (EGI) from T. reesei is not produced during growth on xylan [75] and while its activity on cellulose (hydroxyethylcellulose) is only slightly higher than that on xylan (beechwood and grass), its activity on cello-oligosaccharides (G 3 , G 5 ) is 10-fold higher than that on xylo-oligosaccharides (X 3 , X 5 ) [130] . Hydrolyses of both these substrates takes place in the same active site but a shift in the bond cleavage frequency appears to occur towards the non-reducing end linkages in xylo-oligosaccharides [ 75 , 130 ]. This enzyme has characteristics in common with both family 10 and 11 xylanases. As for the former family, it has a high molecular weight and low p I as well as a small substrate binding site, approximately four subsites, with the catalytic site in the middle. On the other hand, like family 11 xylanases, this enzyme displays a β-jelly roll fold [131] ( Fig. 3(e) ) and produces aldopentauronic acid and isomeric xylotetraose as the shortest acidic and mixed linkage fragments from glucuronoxylan and rhodymenan, respectively [75] . However, even though the core structures of this enzyme and the family 11 xylanases are similar, differences, such as: variations in the location, length and orientation of the structural elements outside of this core, the presence of four short helical segments as opposed to one in the family 11 xylanases and differences in the type and conformation of the amino acid residues lining the active site, results in family 7 enzymes being classified in clan GH-B with family 16 enzymes and not in clan GH-C with the family 11 enzymes.

The family 43 enzyme (XYND) has a molecular weight of 64 kDa and was found clustered with a lichenase gene, with only 155bp separating the two genes, in Paenibacillus polymyxa . Unfortunately however, additional studies of the physicochemical or functional characteristics of this enzyme have not been carried out. Further putative family 43 xylanases in Caldicellulosiruptor sp. [ 133 , 134 ], C. acetobutylicum [59] , Bifidobacterium longum [135] and Bacillus sp. [61] have also been inferred on the basis of sequence homology to the above enzyme but xylanase activity has apparently not been confirmed by functional analysis. Indeed, members of this family have not been as thoroughly studied as some of the other glycoside hydrolase families and the structure of only one member has been determined, indicating that members of this family may display a five-blade β-propeller fold ( Fig. 3(f) ). Furtherrmore, a glutamate and aspartate in the centre of a long V-shaped surface groove formed across the face of the propeller have been suggested as the catalytic residues [54] . The family is grouped with family 62 in clan GH-F [ 43 , 54 ] and, as has also been demonstrated for family 8 enzymes, its members are believed to catalyze hydrolysis via the single displacement mechanism.

The preponderance of xylanases studied are of fungal or bacterial origin and in the majority of cases are found to be optimally active at, or near, mesophilic temperatures (approximately 40–60 °C) [ 16 , 18 ] and neutral (in particular for bacterial xylanases) or slightly acidic (in particular for fungal xylanases) pHs. Nevertheless, xylanases have also been reported which are not only stable, but active, at the extremes of pH and temperature. Indeed, xylanases active at temperatures ranging from 5 to 105 °C [ 4 , 80 , 81 ], pH's from 2 to 11 [ 4 , 114 , 136 ] and NaCl concentrations as high as 30%[ 137 , 138 ] have been reported. These are produced by micro-organisms which have colonized environments that may be said to be extreme from an anthropocentric point of view and which produce enzymes adapted to these extreme habitats. Of the extremophilic xylanases, the thermophiles, alkaliphiles and acidophiles have been the most extensively studied while cold-adapted xylanases have been much less investigated.

6.1 Thermophiles

A number of thermophilic (optimal growth at 50–80 °C) and hyperthermophilic (optimal growth at >80 °C) xylanase producing micro-organisms have been isolated from a variety of sources, including terrestrial and marine solfataric fields, thermal springs, hot pools and self-heating decaying organic debris [ 6 , 128 , 139–142 ]. The majority of the xylanases produced have been found to belong to families 10 and 11, with as yet, no reported studies of thermophilic xylanases belonging to any of the other glycoside hydrolase families. Interestingly, the gene for the thermostable xylanase (half-life of 8 minutes at 100 °C) from the extreme thermophilic archaeon Thermococcus zilligii [143] has thus far proven refractory to cloning with family 10 and 11 consensus primers [141] , suggesting that this enzyme may belong to one of the other less well studied glycoside hydrolase families described in this paper (i.e., families 5, 7, 8 or 43) or indeed to another as yet unknown xylanase family.

Family 10 xylanases have been isolated from various thermophilic and hyperthermophilic organisms, including Thermotoga sp. [ 34 , 144 ], Caldicellulosiruptor sp. [145] , Rhodothermus marinus [146] , Bacillus stearothermophilus [147] , Thermoascus aurantiacus [58] and C. thermocellum [58] . Indeed, a family 10 xylanase, XynA from Thermotoga sp. strain FjSS3-B.1 is one of the most thermostable xylanases reported to date with an apparent optimum temperature for activity of 105 °C and a half-life of 90 minutes at 95 °C [148] . While less frequent, family 11 thermophilic xylanases have also been isolated, with those from Thermomyces lanuginosus [ 6 , 149 ], Paecilomyces varioti [122] , Caldicellulosiruptor sp Rt69B.1. [133] , Dictyoglomus thermophilum [121] , Chaetomium thermophilum [120] , Nonomuraea flexuosa [120] and Bacillus strain D3 [ 128 , 150 ] being the most thoroughly investigated. Those from Nonomuraea flexuosa and Dictyoglomus thermophilum are among the most stable, with apparent temperature optima of 80 and 85 °C, respectively. In addition to the above mentioned xylanase producing bacteria a number of xylanase producing hyperthermophilic archaea have also been recently reported: Thermococcus zilligii [143] , Pyrococcus furiosus [143] , Sulfolobus solfataricus [140] , Pyrodictium abyssi [ 151 , 152 ] and a number of Thermofilum strains [153] .

Crystal structure analyses, sequence alignments and mutagenesis studies have indicated that mesophilic and thermophilic xylanases are very similar and that enhanced stability is probably due to an array of minor modifications, with many xylanases using unique strategies to improve their thermostability. These modifications include: an increase in the number of salt bridges and hydrogen bonds [ 120 , 124 ], an improved internal packing [120] , an increased number of charged surface residues [154] , the presence, often as tandem repeats, of thermostabilising domains [ 34 , 144 , 155 ], and/or the introduction of disulphide bridges, in particular at the N- or C- termini or in the α-helix regions [ 122 , 156 , 157 ]. Recently, the thermostabilising role of calcium on a modular family 10 xylanase was demonstrated [146] while the Bacillus D3 xylanase was also shown to use a very unique adaptation strategy. Here a series of surface aromatic residues form clusters or “sticky patches” between pairs of molecules and these intermolecular hydrophobic interactions are believed to contribute to the thermostability of this enzyme [ 128 , 150 ]. Collectively, or singly, all of the above mentioned modifications could improve the network of interactions within the protein, thereby leading to a more rigid and stable enzyme.

A number of comprehensive structural studies of thermal adaptation for family 10 and 11 xylanases have allowed identification of specific adaptation strategies for each family. For example, a comparison of the thermophilic xylanases from Thermoascus aurantiacus and C.thermocellum with mesophilic family 10 xylanases indicated that the thermostability in this family is a consequence of an improved hydrophobic packing, a favorable interaction of charged side chains with the helix dipoles as well as an increased proline content in the N-termini of helices [58] . In contrast, a recent comparative structural analysis of 5 thermophilic and 7 mesophilic family 11 enzymes suggested that a general thermostabilising adaptation in this family is a higher threonine to serine ratio (threonine has a high β-forming propensity), an increased number of residues in the β-strands and frequently an additional β-strand B1 at the N-terminus [120] . It is evident that the structural differences between the families are the basis for this difference in adaptation strategies; family 10 enzymes have a high α-helix content (approximately 40%) [95] while family 11 enzymes have a high β-sheet content (greater than 50%) [120] .

6.2 Psychrophiles

Even though cold-temperature environments are the most abundant on earth [158] , only a small number of cold-adapted, or psychrophilic, xylanase producers have been identified. These encompass a wide range of organisms; two gram negative bacteria ( Pseudoalteromonas haloplanktis TAH3a [ 80–83 ] and Flavobacterium frigidarium sp. nov. [159] ), a gram positive bacterium ( Clostridium strain PXYL1 [160] ), a yeast isolate ( Cryptococcus adeliae [161] ), krill ( Euphasia superba [162] ), a number of fungi ( Penicillium sp., Alternaria alternata and Phoma sp. 2 [163] ) and a number of basidiomycetes (e.g., Coprinus psychromorbidus [164] ). All have been isolated from the Antarctic environment, but, apart from the bacterial family 8 xylanase from Pseudoalteromonas haloplanktis TAH3a (pXyl) and the Cryptococcus adeliae family 10 xylanase (X B ), studies of the xylanases produced are minimal. Indeed, only these two xylanases and two xylanases, xylanases A and B, from Antarctic krill have been purified and characterized.

In accordance with most other psychrophilic enzymes investigated to date [ 165–167 ], the common features of the psychrophilic xylanases studied are a low temperature optimum, high catalytic activities at low temperatures and poor stability ( Fig. 4 ). Indeed, comparative studies of pXyl and X B with mesophilic xylanases showed that these enzymes have a higher catalytic activity at low and moderate temperatures, having, respectively, 10 and 3 times higher activity at 5 °C and 3 and 2 times higher activity at 30 °C [80] . Moreover, all psychrophilic enzymes studied display high catalytic activity at low temperatures. At 5 °C, activity of pXyl is 60% of the maximum while xylanases A and B from Euphasia superba display, respectively, approximately 30% and 40% of their maximum activity. In comparison, a mesophilic xylanase showed less than 5% of its maximum activity at this temperature [80] . Likewise, the apparent optimal temperatures for activity of pXyl, X B and the microfungal xylanases, which are, respectively, approximately 25, 9 and 10–30 °C lower than that of the mesophilic reference xylanases used, gives further evidence of the adaptation to cold environments of these enzymes. Poor thermal stability of the psychrophilic xylanases studied is indicated by short half-lives (e.g., at 55 °C pXyl has a 12 times shorter half-life of inactivation than a mesophilic xylanase) and low denaturation temperatures (pXyl shows a 10 °C and X B a 14 °C decrease in melting temperature compared to mesophilic reference xylanases) while a lower chemical stability of the cold-adapted family 8 xylanase is demonstrated by short half-lives of guanidine hydrochloride inactivation and unfolding [81] .

(a) Unfolding as monitored by differential scanning calorimetry at a scan rate of 1 K min −1 and (b) thermodependence of activity of the cold-adapted family 8 xylanase pXyl (circles, solid lines), the mesophilic family 11 xylanase Xyl1 [206] (squares, dashed lines) and the thermophilic family 8 endoglucanase CelA [ 55 , 207 ] (triangles, short dashed lines). Baseline subtracted DSC data have been normalized for protein concentration. The lower molecular weight of the mesophilic enzyme gives rise to the observed low calorimetric enthalpy.

Fluorescence monitoring of acrylamide quenching indicated that, in addition to a reduced stability and increased low temperature activity, the family 8 cold-adapted xylanase has an increased flexibility compared to a thermophilic homologous enzyme (CelA from C. thermocellum ) [81] . Indeed, this supports the hypothesis that efficient catalysis at low temperatures is brought about by an increased flexibility of the molecular edifice, thereby allowing the molecular motions necessary for activity in the low temperature, low energy environment but also leading to the observed reduced stability.

At the structural level, it was found that, when compared to a thermophilic and mesophilic homolog, respectively, the family 8 and yeast cold-adapted xylanases are both distinguished by a number of discrete modifications which could give rise to a decrease in the stability, and hence an increase in the flexibility, of the molecular structure. The Pseudoalteromonas haloplanktis TAH3a xylanase is characterised by a reduced number of salt-bridges and an increased exposure of hydrophobic residues [ 82 , 83 ], while the family 10 yeast xylanase is characterized by a less compact hydrophobic packing, the loss of one salt bridge and a destabilization of the helix macrodipoles [161] . Indeed it can be seen that these modifications are an extension of those observed between thermophilic and mesophilic xylanases, however, further analysis of psychrophilic xylanases, especially comparative studies with more closely related and better characterised homologs are necessary to better understand temperature adaptation in these enzymes. In particular, further comparative studies of family 10 and 11 psychrophilic xylanases would be most beneficial.

6.3 Alkaliphiles and acidophiles

While the majority of natural environments on earth are essentially neutral, with pH values of between 5 and 9, habitats with extreme pHs are also common, in particular in geothermal regions, carbonate laden soils, soda deserts and soda lakes such as found in Egypt (Wadi Natrun), the African Rift valley (Lakes Magadi and Nakuru in Kenya), Central Asia, Western USA (Yellowstone National Park) and Southern Europe (Vulcano Island, Italy), Indeed, xylanase producing alkaliphilic micro-organisms, which typically grow optimally at pH values above 9, and acidophiles, which grow optimally between pH 1 and 5, have been isolated from these environments [ 136 , 168 , 169 ] and also from such sources as kraft pulp [170] , pulp and paper industry wastes [171] , decomposing organic matter [172] , faeces [168] , plant sources [173] , soils [ 136 , 174 ] and even from neutral environments where they are found coexisting with neutrophilic micro-organisms [168] .

The first report of a xylanase produced by an alkaliphilic micro-organism was as early as 1973 for a xylanase from Bacillus sp. C-59-2 [175] and since this initial finding a number of xylanases have been isolated from various acidophilic and alkaliphilic micro-organisms. These include family 10 and 11 xylanases from a number of Bacillus sp. [ 116 , 147 , 174 ], Trichoderma sp. [ 126 , 127 , 176 ], Aspergillus sp. [ 114 , 115 ], Penicillium sp. [136] , Acidobacterium sp. [177] and Cryptococcus sp. [178] . In addition, family 8 xylanases have been isolated from alkalophilic B. halodurans C-125 [ 77 , 78 ] and B. halodurans MIR32 [43] , however analysis of the amino-acid sequences indicates that these may be intracellular enzymes and thus may not be adapted to the environment of their hosts.

Many of the alkaliphilic microorganisms studied have been found to produce xylanases with pH optima in the near neutral region but with relatively high activities being retained in alkaline conditions. In addition, a number of xylanases with more alkaline pH optima have also been isolated and one the most alkaliphilic xylanases reported to date is XylB from Bacillus sp. AR-009, which has a pH optimum of pH 9–10 [169] . Other highly alkaliphilic xylanases include xylanase J from Bacillus sp. strain 41M-1 [174] and a xylanase from Bacillus pumilus 13 a [172] , both of which have a pH optimum of 9. Much fewer acidophilic than alkaliphilic xylanases have been studied and the most important of these are the family 10 and 11 members from T. reesei [126] , A. niger [115] , Aspergillus kawachii [ 114 , 179 ], Cryptococcus sp. S-2 [178] and Penicillium sp. 40 [136] . The latter three of these are among the most acidophilic of the studied xylanases with a pH optimum of 2 and stability over a broad pH range; the Penicillium sp. 40 xylanase is stable from pH 2 to pH 5 [136] and the A. kawachii XynC is stable at pH 1–9 [ 114 , 179 ].

The pH activity profiles of enzymes are highly dependent on the p K a s of the catalytic residues which are themselves dependent on the local environment and hence on the nature of the amino-acids in the vicinity of the catalytic residues. A recent study of a family 11 xylanase showed that, in general, residues that contribute positive charges and hydrogen bonds serve to lower the p K a values with shorter bonds having a more pronounced effect. The chemical nature of the donor is also important, with COOH being more effective than OH and CONH 2 [180] . In contrast to this, neighbouring carboxyl groups can either lower or raise the p K a values of the catalytic glutamic acids depending upon the electrostatic linkage of the residues involved in the interaction [180] . In fact it has been noted that family 11 acidophilic xylanases have an aspartic acid residue hydrogen bonded to the general acid/base catalyst which is replaced by an asparagine in the xylanases active under more alkaline conditions [ 114 , 126 , 181 , 182 ]. This residue influences the pH dependence of activity and mutation of this aspartic acid to its amide derivative in the A. kawachii acidophilic xylanase resulted in an upward shift of the pH optimum from pH 2 to a pH of 5. Indeed, tertiary structure analysis of this enzyme [114] as well as of the family 11 xylanases from T. reesei [126] and A. niger [115] indicated that adaptation to low pH is brought about by an increase in negative charge and a substitution and reorientation of residues, in particular aromatic residues, in the active sites. In contrast, a random mutagenesis study of a Neocallimastix patriciarum xylanase indicated that an increased negative charge and increased hydrophobicity increased the pH optimum of this enzyme [183] .

Stability at the extremes of pH appears to be characterized by a spatially biased distribution of charged residues. The acidophilic and acid stable xylanase from A. kawachii , for example, is characterized by a concentration of acidic residues on its surface [114] which are believed to reduce electrostatic repulsion of the positively charged residues at low pHs. In contrast, enzymes stable in alkaline conditions are typically characterized by a decreased number of acidic residues and an increased number of arginines. Furthermore, a recent comparative structural study of family 11 enzymes suggests a correlation between pH activity/stability and the number of salt bridges, with acidophilic xylanases having much less of these interactions than their alkaliphilic homologs [120] . Indeed it was even suggested that adaptation to high pH may occur via a similar mechanism to adaptation to high temperatures [120] .

Global markets for industrial enzymes grew from €1 billion in 1995 [184] to almost €2 billion in 2001 [185] and continue to increase as new enzymes and applications are discovered. In the grain-processing enzymes sector alone (which currently accounts for approximately 25–28% of total enzyme sales) an increase in market value from €510 million in 2001 to €760 million in 2010 has been forecasted [185] . Presently the technical industries, dominated by the detergent, starch, textile and fuel alcohol industries, account for the majority of the total enzymes market, with the feed and food enzymes together totaling only about 35%. Recently however, sales in some of the major technical industries has stagnated (3% drop in 2001) while sales in both the food and feed industries are increasing, with annual growth rates of approximately 4–5% being forecasted [185] .

Hydrolases constitute approximately 75% of the markets for industrial enzymes, with the glycosidases, including cellulases, amylases and hemicellulases, constituting the second largest group after proteases [186] . Xylanases constitute the major commercial proportion of hemicellulases but represent only a small percentage of the total enzyme sales. The sales figures are expected to increase however, as these enzymes have attracted increasing attention due to their potential for use in several applications. In effect, the United States Patent and Trademark Office ( http://www.uspto.gov/ ) lists 468 patents introduced since 2001 with reference to xylanases (search field = all fields).

Xylanases have potential applications in a wide range of industrial processes, covering all three sectors of industrial enzymes markets [186] and some of the most important of these are listed in Table 6 . Other less well documented putative applications include: in brewing, to increase wort filterability and reduce haze in the final product [187] ; in coffee extraction and in the preparation of soluble coffee [7] ; in detergents [188] ; in the protoplastation of plant cells [4] ; in the production of pharmacologically active polysaccharides for use as antimicrobial agents [107] or antioxidants [189] ; in the production of alkyl glycosides for use as surfactants [190] ; and in the washing of precision devices and semiconductors [191] . The xylanases are frequently utilized alone, but are more commonly used in conjunction with other enzymes and in particular with other hydrolases, but also with proteases, oxidases, isomerases etc.

Potential applications for xylanases

Many of the xylanases used in industry today appear to be of mesophilic and/or neutrophilic origin, yet enzymes from extremophilic sources may be of tremendous utility in many biotechnological processes. In particular, thermophilic enzymes could be used in applications where a cooling step would be uneconomical or where high temperatures are required to increase the bioavailability and/or solubility of substrates, to reduce viscosity and/or to reduce the risk of contamination. Acidophilic and alkaliphilic enzymes would obviously be beneficial in processes where extreme pH conditions are required or where adjustment of the pH to neutral conditions is uneconomical. On the other hand, cold-adapted xylanases would be beneficial to those processes where heating is economically counterproductive or where low temperatures are required to avoid alteration of ingredient and/or product quality (e.g., flavour, colour etc.), to avoid microbial development and fermentation and/or to avoid product denaturation. In fact, enzymes which combine a number of extremophilic characteristics may be of the most use in industry.

The major current application of xylanases is in the pulp and paper industries where the high temperature (55–70 °C) and alkaline pH of the pulp substrate requires thermo-alkaliphilic enzymes for efficient biobleaching [ 5 , 192 ]. Thermo-alkaliphilic or even thermo-acidophilic xylanases may also be of use in bioconversion processes where a variety of treatments, including hot water and steam explosion, alkaline, solvent or acidic pretreatments may be used prior to or simultaneous to enzyme treatment [ 193 , 194 ]. Alkaliphilic xylanases would also be required for detergent applications where high pHs are typically used [188] while a thermostable xylanase would be beneficial in animal feeds if added to the feeds before the pelleting process (typically carried out at 70–95 °C). In addition, for this latter application the enzyme must be highly active at the temperature (approximately 40 °C) and pH (approximately pH 4.8) of the digestive tract [195] . Cold adapted xylanases, which are most active at low and intermediate temperatures, could offer advantages over the currently used xylanases in many of the low to moderate temperature processes, in particular in the food industry. For example, they would be most suited for use in the baking industry as dough preparation and proofing is generally carried out at temperatures below 35 °C and indeed it has recently been demonstrated that a cold-adapted family 8 xylanase is more efficient in baking than a commonly used commercial enzyme [79] .

A large variety of enzymes with xylanase activity have been isolated and studied and can be classified in glycoside hydrolase families 5, 7, 8, 10, 11 and 43, with each of these families being characterized by a particular fold and mechanism of action. While extensive studies of family 10 and 11 xylanases have been carried out, leading to an enrichment in the understanding of these enzymes, studies of the xylanase members of the other families are sparse. Only a small number of enzymes with xylanase activity have been identified in families 5, 7, 8 and 43, however, the numbers will increase with further studies and may indeed be extended to additional families as novel enzymes from novel sources are discovered. New approaches, such as genome sequencing programs, functional and/or consensus PCR screening of environmental DNA libraries (known as metagenomics) as well as the study of extremophilic enzymes will, without doubt, further extend the current repertoire, understanding, and applications of xylanases and xylanase families.

The authors thank N. Gerardin and R. Marchand for their skillful technical assistance and Jean-Claude Marx for his help with the figures. We also wish to thank the French Institute for Polar Research for generously accommodating our research fellows at the French Antarctic station in J.S. Dumont d'Urville, Terre Adelie. This work was supported by the ‘Région Wallonne’ (contracts FIRST Europe ‘objective 3’ R0202/215266 and BIOVAL 981/3860), the European Union (network contract CT970131) and the ‘Fonds National de la Recherche Scientifique’ (contracts 2.4515.00 and 2.4536.04).

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Isolation, Purification, and Characterization of Xylanase Produced by a New Species of Bacillus in Solid State Fermentation

Rajashri d. kamble.

1 Department of Biotechnology Engineering, Tatyasaheb Kore Institute of Engineering & Technology, Warananagar Panhala, Kolhapur, Maharashtra, 416113, India

Anandrao R. Jadhav

2 Department of Microbiology, K.R.P. Kanya Mahavidyalaya, Islampur, Walwa, Sangli, Maharashtra, 415414, India

A thermoalkalophilic new species of Bacillus , similar to Bacillus arseniciselenatis DSM 15340, produced extracellular xylanase under solid state fermentation when wheat bran is used as carbon source. The extracellular xylanase was isolated by ammonium sulfate (80%) precipitation and purified using ion exchange chromatography. The molecular weight of xylanase was ~29.8 kDa. The optimum temperature and pH for the enzyme activity were 50°C and pH 8.0. The enzyme was active on birchwood xylan and little active on p -nitrophenyl xylopyranoside but not on Avicel, CMC, cellobiose, and starch, showing its absolute substrate specificity. For birchwood xylan, the enzyme gave a Km 5.26 mg/mL and Vmax 277.7  μ mol/min/mg, respectively. In addition, the xylanase was also capable of producing high-quality xylo-oligosaccharides, which indicated its application potential not only in pulp biobleaching processes but also in the nutraceutical industry.

1. Introduction

Xylan is the most abundant noncellulosic polysaccharide present in both hardwoods and annual plants and accounts for 20–35% of the total dry weight in tropical plant biomass [ 1 – 3 ]. In temperate softwoods, xylans are less abundant and may comprise about 8% of the total dry weight [ 4 ]. Xylan is found mainly in the secondary cell wall and is considered to be forming an interphase between lignin and other polysaccharides. It is likely that xylan molecules covalently link with lignin phenolic residues and also interact with polysaccharides, such as pectin and glucan. In simplest forms, xylans are linear homopolymers that contain D-xylose monomers linked through β -1, 4–glycosyl bonds [ 5 , 6 ]. Xylanase (E.C 3.2.1.8) degrades β -1, 4 xylan by cleaving β -1, 4 glycosidic linkages randomly, and the products are xylose and xylo-oligosaccharides like xylobiose [ 7 , 8 ]. Xylanases are of industrial importance, which can be used in paper manufacturing to bleach paper pulp, increasing the brightness of pulp and improving the digestibility of animal feed and for clarification of fruit juices. Applications of xylanase avoid the use of chemicals that are expensive and cause pollution [ 9 ]. Microorganisms are the rich sources of xylanases, produced by diverse genera and species of bacteria, actinomycetes, and fungi. Several species of Bacillus and filamentous fungi secrete high amounts of extracellular xylanases [ 10 ]. Xylanase secretion often associates with low or high amount of cellulases. To use xylanase for pulp treatment, it is preferable to use cellulose-free xylanases, since the cellulase may adversely affect the quality of the paper pulp [ 11 – 15 ]. The most practical approach is the screening for naturally occurring microbial strains capable of secreting cellulose-free xylanases under optimized fermentation conditions. To use xylanase prominently in bleaching process it should be stable at high temperature and alkaline pH [ 16 , 17 ].

Industrial production of enzymes on large scale is associated mainly with substrate. The use of agriculture residues as low-cost substrates for the production of industrial enzymes is a significant way to reduce production cost. The technique of fermentation using solid state substrate has the great advantage over submerged fermentation due to absence or near absence of aqueous phase that provides natural habitat for growth of microorganisms, economy of the space, simplicity of the media, no complex machinery, equipments and control systems, greater compactness of the fermentation vessel owing to a lower water volume, greater product yields, reduced energy demand, lower capital and recurring expenditures in industry, easier scale-up of processes, lesser volume of solvent needed for product recovery, superior yields, absence of foam build-up, and easier control of contamination due to the low moisture level in the system [ 10 , 18 ]. In consideration with these facts the present study aims to characterize extracellular alkalothermophilic xylanase produced by Bacillus arseniciselenatis DSM 15340 when grown in solid state fermentation. To our knowledge, this is the first report describing the production of thermoalkalophilic cellulase-free xylanase by Bacillus arseniciselenatis DSM 15340. In addition, this xylanase was found to be able to degrade xylan into xylo-oligosaccharides.

2. Materials and Methods

2.1. screening of xylanolytic strains.

Soil samples were collected from coastal areas of Mandovi, Goa, India. Enrichment was done using birchwood xylan (Sigma Chemicals, Germany) as a sole source of carbon. Twenty five bacterial cultures were screened for xylanolytic ability by adding dye-labelled substrate, for example, xylan-brilliant red 3BA in xylan agar medium [ 19 ].

2.2. Phenotypic Characteristics

Prominent selected isolate was identified on the basis of morphological, cultural, biochemical properties [ 20 ] and 16S rRNA sequencing. Culture was deposited at National Centre for Cell Sciences (NCCS), Pune, India.

2.3. Phylogenic Analysis

The partial 16S rRNA sequences were retrieved on NCBI server ( http://blast.ncbi.nlm.nih.gov/Blast.cgi ) using BLAST tool. Top 10 similar sequences were downloaded in FASTA format. Multiple alignment of sequences and calculations of levels of sequence similarity were performed by using ClustalW2 program. A phylogenetic tree obtained was analyzed for closely related organism. The evolutionary history was inferred using the neighbor-joining method [ 14 ].

2.4. Growth Conditions of Culture

The bacterial isolate was maintained in liquid medium as well as solid medium in basal salt solution (BSS) containing 0.5% xylan having pH 8.0 at 45°C and stored at 4°C.

2.5. Xylanase Production in Solid State Fermentation (SSF)

The selected strain was further tested for their abilities to produce extracellular xylanase under solid state fermentation. Wheat bran was used as the substrate. For this the strain was cultured in Erlenmeyer flasks (250 mL) containing 10 g of wheat bran moistened with 18 mL of the basal salt solution (BSS: substrate-to-moisture ratio 1 : 1. 8). After 48 h of fermentation spent; solid substrate was removed and suspended in 50 mM phosphate buffer (pH 8.0), vortexed thoroughly to extract the xylanase. The sample was centrifuged at 5000 ×g for 10 minutes at 4°C. Centrifugation will remove xylanase from substrate. Supernatant was filtered through Whatman No. 1 filter paper and the clear filtrate was used as crude xylanase preparation. Prior to centrifugation, the samples were withdrawn for determining viable number of cells by standard viable plate count technique.

2.6. Xylanase Assay

Xylanase activity was measured according to Bailey et al. [ 21 ]. A 900  μ L of 1% solubilised birchwood xylan solution was added with 100  μ L enzyme solution in a test tube. 1.5 mL DNS reagent was added and incubated at 50°C for 5 min in water bath [ 22 ]. The absorbance was measured at 540 nm. The reaction was terminated at zero time in the control tubes. The standard graph was prepared using 0–500  μ g xylose. An autozero was set in UV-VIS spectrophotometer (Hitachi, Japan) using buffer solution. One unit of xylanase activity was defined as the amount of enzyme that liberates 1 micromole of reducing sugars equivalent to xylose per minute under the assay conditions described. Solubilised xylan was prepared by stirring birchwood xylan with 1 M NaOH for six hours at room temperature followed by centrifugation and freeze drying the supernatant after neutralising the alkali with 1 M HCl.

2.7. Cellulase Assay

Cellulase activity was measured according to Ghose with necessary modifications [ 23 ]. A 900  μ L 1% carboxy methyl cellulose solution was added with 100  μ L enzyme in a test tube. 1.5 mL DNS reagent was added and incubated at 50°C for 5 min in water bath. The absorbance was measured at 540 nm. The reaction was terminated at zero time in control tubes. A standard graph was prepared using 0–500  μ g glucose. An autozero was set in spectrophotometer using buffer solution. One unit of cellulase activity was defined as the amount of enzyme that liberates 1 micromole of glucose equivalents per minute under the assay conditions.

2.8. 1,4- β -xylosidase Assay

1,4- β -xylosidase activity was measured according to Lachke [ 24 ]. A 900  μ L p -nitrophenyl β -xyloside ( ρ -NPX) solution was added with 100  μ L of appropriately diluted enzyme solution in a test tube. The mixture was incubated at 50°C for 30 min. Then 1 mL of 2 M sodium carbonate solution was added. The absorbance was measured at 410 nm. The reaction was terminated at zero time in control tubes. One unit of 1,4- β -xylosidase activity was defined as the amount of enzyme that catalyzes the formation of 1 micromole of ρ -nitrophenol per minute under assay conditions.

2.9. Determination of Total Protein Content

Total soluble protein was measured according to Lowry et al. [ 25 ]. Protein concentration was determined using bovine serum albumin (BSA) as a standard. The protein content of the chromatographic eluant was measured by monitoring the optical density at 280 nm.

2.10. Ammonium Sulphate Precipitation

Protein precipitation by salting out technique using ammonium sulphate (NH 4 (SO 4 ) 2 ) was carried out with constant gentle stirring [ 26 ]. This was left overnight and the precipitate was collected by centrifugation at 10,000 ×g for 10 min. The precipitate obtained was dissolved in phosphate buffer (50 mM, pH 8.0) and dialyzed against the same buffer for 24 h. Dialysis was carried out using cellulose tubing (molecular weight cut-off 13,000 kDa, Himedia LA393-10 MT).

2.11. Ion Exchange Chromatography

Dialyzed enzyme (2 mL) was loaded onto a anion exchange DEAE Cellulose (Sigma-Aldrich Co., USA) column. The column was packed with activated DEAE-cellulose equilibrated with 50 mM phosphate buffer (pH 8.0). The height of column was 20 cm with the 2.5 cm diameter. The protein was eluted with the 0.0 to 0.5 M NaCl gradient. The 50 fractions were collected having 5 mL volume of each fraction with the flow rate of 1 mL/min. All the steps were carried out at 4 to 8°C.

2.12. Molecular Mass Determination by SDS-PAGE

SDS-PAGE of partially purified xylanase was performed in a 12.5% acrylamide gel Laemmli [ 27 ]. Coomassie brilliant blue R-250 was used to stain the gel. The protein molecular weight markers used were of medium range containing 14.4 kDa to 94.0 kDa obtained from Bangalore GeNei, India.

2.13. Substrate Specificity

Substrate specificity of the xylanase was found by using 1% xylan, cellobiose, starch, carboxy methyl cellulose (CMC), and p -nitrophenyl xylopyranoside and Avicel as substrates.

2.14. Kinetic Parameters

Initial reaction rates using birchwood and oat spelt xylan as substrate were determined at substrate concentrations of 0.5–10 mg/mL in 50 mM phosphate buffer (pH 7.0) at 45°C. The kinetic constants, Km and Vmax, were estimated using the linear regression method of Lineweaver and Burk [ 28 ].

2.15. Identification of Hydrolysis Products

To 50 mL of birchwood xylan suspension (1% of birchwood xylan in 50 mM Phosphate buffer pH 7.0), 40  μ g of xylanase enzyme was added and incubated at 45°C. Hydrolysis products were detected by thin layer chromatography (TLC) [ 29 ]. TLC (TLC plates, 0.25 mm layers of silica gel F 254, Merck, India) was performed using the mixture of n -butanol : ethanol : H 2 O (5 : 3 : 2 by vol) as a solvent system. Compounds were detected by spraying with 50% sulphuric acid in ethanol followed by heating at 150°C for 5 min. D-xylose (X 1 ), xylobiose (X 2 ), xylotriose (X 3 ), and xylotetraose (X 4 ) were applied as standard.

2.16. Effect of Temperature on Activity and Stability

The optimum temperature for maximum xylanase activity was determined by varying the reaction temperature from 30 to 80°C. To evaluate thermal stability, 0.5 mL of the enzyme solution was incubated at 30–80°C temperatures for up to 4 h. The relative enzyme activity was recorded at 1 h interval during period of 4 h.

2.17. Effect of pH on Activity and Stability

The effect of pH on enzyme activity was determined by incubating xylanase at various pH ranging from 6.0 to 11.0. The various buffers used were 50 mM sodium phosphate (pH 6, 7), 50 mM Tris HCl (pH 8, 9), 50 mM carbonate bicarbonate buffer (pH 10), and 50 mM glycine-NaOH buffer (pH 11). To evaluate the stability of the enzyme at each pH, the purified enzyme was incubated into the respective buffer over a pH range of 6.0–11.0 for up to 4 h at optimum temperature. The relative enzyme activity was determined at 1 h interval during the 4 h period of incubation.

3. Results and Discussion

3.1. isolation and identification of bacteria.

About 25 bacterial strains, which formed clear halos around their colonies on xylan agar plates, were picked up for further studies, isolated from soil collected at selected study site. The strain that showed 33 mm zone of clearance around the colony proved its xylanolytic ability ( Figure 1 ). It was identified on the basis of various morphological and biochemical characteristics as shown in Table 1 .

Plate showing zone of clearance around colony by isolate.

Morphological, physiological, and biochemical characteristics of the isolate.

+: Positive; −: Negative.

The isolate was confirmed as Bacillus arseniciselenatis strain DSM-15340 with partial 16S rRNA sequencing having a length of 1499 bp nucleotide. The sequence was deposited in Gene Bank (Accession No. {"type":"entrez-nucleotide","attrs":{"text":"AJ865469","term_id":"56266617","term_text":"AJ865469"}} AJ865469 ). The phylogenetic relation of this isolate is as shown in Figure 2 . It is closely associated with Bacillus sp. AMnr. It was also isolated from soil sample collected at coastal areas of Mandovi, Goa. Shivaji et al. isolated Bacillus arseniciselenatis DSM 15340 and Bacillus arsenicus from a bore well located in the chakdah region of West Bengal, India [ 30 ].

The phylogenetic tree of Bacillus arseniciselenatis DSM 15340 (designated as “0”).

3.2. Xylanase Production in SSF

When the strain was grown on wheat bran for 3 days of incubation at pH 8.0 and 45°C, maximum xylanase production was observed, that is, 910.49 U/gram dry substance, which was absolutely free from cellulase. Several workers reported the suitability of wheat bran for xylanase production in SSF [ 31 , 32 ]. Commercial wheat bran consists of 30% cellulose, 27% hemicellulose, 21% lignin, and 8% ash [ 33 ]. Hence there was increase in possibility of cellulase contamination when grown on wheat bran. Haltrich et al. also reported that xylanases were always associated with cellulase [ 34 ]. From twenty selected strains, five were able to produce cellulase along with xylanase in SSF. This was due to the presence of cellulose in substrate wheat bran used in SSF.

3.3. Purification of Xylanase

The culture filtrate was precipitated by fractional (35–80%) ammonium sulphate saturation. Proteins precipitated within this range had maximum xylanase activity and was used for purification. Xylanase was further purified by DEAE cellulose ion exchange column. The enzyme was eluted from DEAE cellulose column at a NaCl concentration of 0.25 M ( Figure 3 ). The fractions (no. 19–25) having maximum specific activity were concentrated. Xylanase was purified 3.06-fold with a specific activity of 299.25 U/mg ( Table 2 ).

Elution profile of xylanase from DEAE-cellulose column chromatography.

Purification steps of xylanase enzyme isolated from Bacillus arseniciselenatis DSM 15340 when grown on wheat bran.

The specific activity of xylanase produced by Bacillus pumilus was previously reported as 298 U/mg by Panbangred et al. [ 35 ].

3.4. Molecular Weight Determination

The purified enzyme showed a single-protein band on SDS-PAGE. The molecular mass of denatured xylanase, estimated from the relative mobility of proteins on SDS-PAGE, was ~29.8 kDa as shown in Figure 4 . The present results were supported by previous work. The enzyme from a fungus Plectosphaerella cucumerina had a molecular weight of 19 kDa reported by Zhang et al. [ 36 ]. Xylanase produced by Bacillus sp . strain BP-23 is of 32 kDa [ 37 ] whereas the second xylanase obtained from Bacillus firmus had a molecular weight of 45 kDa [ 38 ].

SDS-PAGE analysis of purified xylanases from Bacillus arseniciselenatis DSM 15340. Lane M: molecular markers; Lane B: Purified xylanase enzyme.

3.5. Substrate Specificity

The action of the purified xylanase towards various substrates was studied. The enzyme was active on birchwood xylan, little active on p -nitrophenyl xylopyranoside but not on Avicel, CMC, cellobiose, and starch ( Table 3 ). Purified xylanase was not active on Avicel, CMC, cellobiose, and starch even when the enzyme concentration was 5 times greater than used in normal assay and incubation period of 20 minutes rather than 5 minutes. Similarly, xylanase with absolute substrate specificity was purified from Trichoderma viride by Ujiie et al. [ 39 ]. Kanda et al. purified two different xylanases, named xyl I and III that showed no activity towards glycans, other than xylan, such as starch, pachyman, and Avicel (microcrystalline cellulose), except for the almost one twentieth activity of xyl III toward carboxymethyl cellulose (CMC) [ 40 ].

Substrate specificity of purified xylanase.

Each value represents the mean + standard error values.

3.6. Kinetic Parameters

The kinetic parameters Km and V max⁡ of the enzyme were determined from Lineweaver-Burk double-reciprocal plots of xylanase activity at 45°C using various concentrations of birchwood xylan as substrate ( Figure 5 ). The Km and V max⁡ values of xylanase were 5.26 mg/mL and 277.7  μ mol/min/mg, respectively. Wang et al. reported that Km and V max⁡ values of xylanase isolated from Bacillus sp . NTU-06 were 3.45 mg/mL and 387.3  μ mol/min/mg, respectively [ 41 ]. Bansod et al. also reported that Km values of xylanases lie in the range from 0.5 to 19.6 mg/mL [ 42 ]. Xylanases isolated from Aeromonas cavie 171 ME-1 and Bacillus sp. strain 41m-1 showed similar values of V max⁡ 260 to 350  μ mol/min/mg protein [ 43 , 44 ].

Double reciprocal plot for determining the V max⁡ and Km values of xylanase Bacillus arseniciselenatis DSM 15340 when acted on Birchwood xylan.

3.7. Analysis of Hydrolytic Products

After 1 h of incubation of birchwood xylan with xylanase Bacillus arseniciselenatis DSM 15340, xylotriose and xylotetraose were the main products in the hydrolytic mixture along with little amount xylobiose. ( Figure 6 ). The present results indicated that xylanase cleaved the substrate to liberate mainly xylooligosaccharides, but not able to act on resulting oligosaccharides to form xylose, suggesting that it is a endoxylanase.

TLC analysis for hydrolysis products released from birchwood xylan by xylanase from Bacillus arseniciselenatis DSM 15340. S: substrate; 1: sample; X 1 : D-xylose; X 2 : xylobiose; X 3 : xylotriose; X 4 : xylotetraose.

Analysis of hydrolytic products of xylan by the xylanase of Thermoascus aurantiacus showed that xylan was degraded to various xylo-oligosaccharides without a significant accumulation of xylose [ 45 ]. Xylobiose and xylotriose were the main hydrolysis products when xylanase of Bacillus stearothermophilus reacted with oat spelt xylan and resulted oligosaccharides were then cleaved to form xylose by the β -xylosidase action [ 46 ]. The end products were xylobiose, xylotriose, xylotetraose, and higher oligosaccharides when xylan was hydrolyzed with endoxylanase of alkalophilic Bacillus sp. No. C-125. No xylose was found in the hydrolysis products when analysed by HPLC [ 47 ].

3.8. Effect of Temperature on Activity and Stability

For xylanase from Bacillus arseniciselenatis DSM 15340, activity was found to be gradually increased with increasing temperature and found significantly declined at 80°C ( Figure 7 ). 50°C was found to be the most favourable for enzyme activity. Stability of the enzyme was the most important factor in studying characteristics. In case of xylanase purified from Bacillus arseniciselenatis DSM 15340, it was more stable at temperatures 30°C and 40°C for 4 h of incubation and retained almost 93% activity. At higher temperature values xylanase stability was gradually declined ( Figure 8 ). Bernier et al. reported that multiple forms of xylanases were purified from Aeromonas sp. by Ohkoshi et al. and the properties of the three xylanases were well characterized [ 2 , 48 ]. It was found that these xylanases were most active at 50°C to 60°C. Kang et al. purified two xylanases which gave the highest activity at 50°C. They showed relatively high stabilities at 50°C temperature [ 49 ].

Effect of temperature on activity of xylanase from Bacillus arseniciselenatis DSM 15340.

Effect of temperature on stability of xylanase from Bacillus arseniciselenatis DSM 15340.

3.9. Effect of pH on Activity and Stability

pH was the most important factor to characterize the enzyme. Xylanase from Bacillus arseniciselenatis DSM 15340 showed 100% activity at pH 8.0 ( Figure 9 ). At higher pH values also, activity was 95%. With respect to stability, at all tested pH values xylanase activity was 100% activity for 1 h. pH 10.0 was the most favourable for stability and retained 60% activity for 4 h of incubation ( Figure 10 ).

Effect of pH on activity of xylanase from Bacillus arseniciselenatis DSM 15340.

Effect of pH on stability of xylanase from Bacillus arseniciselenatis DSM 15340.

Similarly, Honda et al. purified two xylanases namely xylanases N and xylanase A from Bacillus sp. No. C-125 [ 47 ]. Among these xylanases, N shows maximum activity at pH ranging from 6.0 to 7.0, while xylanase A was active at pH ranging from 6.0 to 10.0 and showed some activity at pH 12.0 also. In the present study, 100% activity was retained by Bacillus arseniciselenatis DSM 15340 xylanase for 2 h of incubation at pH 10.0. Stability at the extreme pH values may be due to charged amino acid residues. The enzymes stable in alkaline conditions were characterized by a decreased number of acidic residues and an increased number of arginines [ 50 ].

4. Conclusions

Bacillus arseniciselenatis DSM 15340 produced a thermoalkalophilic cellulose-free xylanase in higher amount when grown on solid state conditions using cheaply available agroresidual substrate wheat bran. Hence it can be used for large-scale production of xylanase using such agroresidual substrates. The purified xylanase also was capable of producing high-quality xylo-oligosaccharides, indicating its application potential not only in pulp biobleaching processes but also in the nutraceutical industry.

Acknowledgment

The authors are thankful to Dr. V. V. Kajinnii, Principal, Tatyasaheb Kore Institute of Engineering and Technology, Warananagar, India, for financial support.

• Open access
• Published: 17 August 2024

Emergency robotic surgery: the experience of a single center and review of the literature

• Graziano Ceccarelli 1 ,
• Fausto Catena 2 ,
• Pasquale Avella 3 , 4 ,
• Brian WCA Tian 5 ,
• Fabio Rondelli 1 ,
• Germano Guerra 4 ,
• Michele De Rosa 1 &
• Aldo Rocca 3 , 4  

World Journal of Emergency Surgery volume  19 , Article number:  28 ( 2024 ) Cite this article

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Backgrounds

Laparoscopic surgery is widely used in abdominal emergency surgery (AES), and the possibility of extending this approach to the more recent robotic surgery (RS) arouses great interest. The slow diffusion of robotic technology mainly due to high costs and the longer RS operative time when compared to laparoscopy may represent disincentives, especially in AES. This study aims to report our experience in the use of RS in AES assessing its safety and feasibility, with particular focus on intra- and post-operative complications, conversion rate, and surgical learning curve. Our data were also compared to other experiences though an extensive literature review.

We retrospectively analysed a single surgeon series of the last 10 years. From January 2014 to December 2023, 36 patients underwent urgent or emergency RS. The robotic devices used were Da Vinci Si (15 cases) and Xi (21 cases).

36 (4.3%) out of 834 robotic procedures were included in our analysis: 20 (56.56%) females. The mean age was 63 years and 30% of patients were ≥ 70 years. 2 (5.55%) procedures were performed at night. No conversions to open were reported in this series. According to the Clavien-Dindo classification, 2 (5.5%) major complications were collected. Intraoperative and 30-day mortality were 0%.

Conclusions

Our study demonstrates that RS may be a useful and reliable approach also to AES and intraoperative laparoscopic complications when performed in selected hemodynamically stable patients in very well-trained robotic centers. The technology may increase the minimally invasive use and conversion rate in emergent settings in a completely robotic or hybrid approach.

Introduction

Abdominal Emergency Surgery (AES) can be defined as a procedure requiring to deal with an acute threat to life, organ, trauma, acute disease process, acute exacerbation of a chronic disease process, or complication of a surgical or other interventional procedure, normally within hours of decision to operate [ 1 , 2 ].

Further, “expedited surgery” refers to the clinical situation exemplified by a patient in need of prompt treatment but not in imminent danger to life or organ survival; this procedure often takes place a few days after the decision to operate [ 3 ].

Nowadays, minimally invasive laparoscopic approach to urgent abdominal surgery (cholecystitis, acute appendicectomies, bowel perforation or obstruction, etc.) represents the standard of care in many cases and recent guidelines recommend it [ 4 , 5 , 6 , 7 ].

Nevertheless, after more than 20 years from clinical introduction, Robotic Surgery (RS) represents the most important technological evolution and a revolutionary concept of computer-assisted technology in minimally invasive surgery [ 8 ]. It allows to overcome many limits of conventional laparoscopy and to expand the use of minimally invasive approaches.

Its peculiar features include a three-dimensional high-definition view, articulated instruments, tremor eradication, and improved ergonomics for surgeons, enable the performance of extremely accurate procedures (micro-sutures, fine dissections, etc.) with consequently lowering conversion rates and postoperative complications, particularly in case of challenging surgical procedures [ 9 , 10 , 11 ]. In addition, compared to traditional laparoscopic surgery, RS demonstrated shorter learning curves for several complex procedures [ 12 , 13 ]. On the other hand, the main drawbacks of robotic technology are linked to its limited diffusion also due to expensive costs [ 14 , 15 , 16 , 17 , 18 , 19 , 20 ].

Nevertheless, robotic surgical technologies have expanded and evolved over the past 20 years, bringing new devices, and improving the most established ones [ 21 , 22 ].

The spreading of robotic platforms and their easier management led to increased RS applications in all abdominal surgical specialities including upper gastrointestinal surgery [ 15 , 23 , 24 , 25 ], colorectal surgery [ 26 , 27 , 28 ], HBP surgery [ 14 , 18 , 29 , 30 , 31 ], abdominal wall surgery and many others [ 7 ].

Despite the huge diffusion of RS in all surgical fields, its application in urgent scenarios has never been investigated representing a new field of interest, with limited literature experiences [ 32 ].

So considering that our experience in RS has been implemented since 2002 and it raised from general to major complex surgery [ 33 , 34 , 35 , 36 , 37 , 38 ], we aim to set the state of art of Robotic Emergency Surgery sharing our experience through the analysis of our peri-operative outcomes and indications in RS. Furthermore, due to the limited evidence available, we have as a secondary endpoint an extensive analysis of previous literature experiences.

Study design and patient selection

We retrospectively reviewed a prospectively collected database of patients undergoing RS at General and Robotic Surgery Unit of San Giovanni Battista Hospital (Foligno, Italy) and General Surgery Unit of San Donato Hospital (Arezzo, Italy) from January 2014 to December 2023.

The patients’ data were analyzed according to Strengthening the Reporting of Observational Studies in Epidemiology (STROBE) [ 39 ]. All patients signed an informed consent allowing the anonymous scientific use of clinical data and images. The study was carried out according to the Declaration of Helsinki guidelines and was approved by the Institutional Review Board of the University of Molise (protocol number 10/21, approved date: May 12, 2021).

In all participating centres, data were prospectively collected from electronic patient records.

We selected 834 consecutive robotic procedures for abdominal surgery performed by both centres. Patients were divided into two cohorts: elective surgery and urgent or emergency surgery groups.

Urgent surgery was defined as a condition requiring surgery within 72 h in stable patients, but not suitable for discharge. Moreover, emergency surgery was defined as a clinical scenario requiring within 24 h in stable patients, with a low risk of deterioration. All patients < 18 years old and affected by hemodynamical instability were excluded.

Criteria adopted to assess baseline characteristics of patients, surgical issues and technologies that allow to benefit of RS in urgent and emergency settings are summarized in Table  1 .

Furthermore, to analyze the diagnosis and intraoperative data we carried out a specialities classification as reported in Table  2 .

Implementation of the robotic surgery program and learning curve completion

Our experience with RS started in September 2002 with the da Vinci S ® platform (Intuitive Surgical, Sunnyvale, California, USA), and over time, its application in abdominal surgery grew as well as platform technologies. During the study period, the da Vinci Si ® platform (Intuitive Surgical, Sunnyvale, California, USA) and, since 2017, da Vinci Xi ® (Intuitive Surgical, Sunnyvale, California, USA) were available at our institutions.

Beginning from colorectal surgery, hiatal hernia repairs and cholecystectomies, our surgical team have gradually selected more challenging procedures by carrying out liver and pancreatic resections, oesophageal benign and malignant disorders, bariatric surgery, abdominal wall hernia repairs and nephrectomies [ 14 , 15 , 25 , 40 , 41 , 42 , 43 , 44 , 45 , 46 , 47 ].

All procedures were performed by a well-trained surgeon in minimally invasive surgery (G.C.) with 10 years of previous experience in RS.

36 (4.3%) out of 834 robotic procedures were included in our analysis and treated as urgent or emergent procedures.

All patients signed an informed consent allowing the anonymous scientific use of clinical data and images. The study was carried out according to the Declaration of Helsinki guidelines and was approved by the Institutional Review Board of the University of Molise (protocol number 10/21, approved date: 12 May 2021).

The collected data included demographic characteristics such as age, sex, and preoperative comorbidities classified according to the Charlson comorbidity Index (CCI) [ 48 ]. The anesthetic risk assessment was performed thanks to the American Society of Anesthesiologists (ASA) score [ 49 ].

We reviewed pre-operative diagnosis, main and associated surgical procedures, operative time and intraoperative complications, and conversion to open rate. Postoperative complications were stratified according to the Clavien-Dindo classification [ 50 ] and were considered severe when ≥ 3. Patients were monitored until their 30th postoperative day.

Categorical variables were expressed as frequencies and percentages, while quantitative data were collected as means or medians and interquartile ranges.

Literature review

We performed literature research on the PubMed Dataset (US National Library of Medicine, http://www.ncbi.nlm.nih.gov/PubMed ), using the subsequent keywords: “robotic surgery”, “urgent surgery”, and “emergency surgery”. We selected only English studies. Original articles, case reports and case series were included, while editorials, letters, and reviews were excluded. Number of treated patients does not represent an exclusion criterion. Articles were first evaluated by title and abstract examination, then a full-text read was performed. More than 50 studies described RS in AES. An extensive analysis was performed to summarize similarities and differences among RS approaches according to abdominal surgery specialities.

Pre-, intra- e post-operative outcomes

36 (4.3%) out of 834 robotic procedures were included in our analysis. Baseline characteristics of patients are listed in Table  3 .

According to surgical procedures, Fig.  1 shows AES performed thanks to RS compared to elective surgery.

Over the cohort of 36 patients treated, 16 (44.44%) were males while 20 (56.56%) were females. The mean age was 63.20 years (range: 43–88 years): 30.55% [ 11 ] of patients were ≥ 70 years. The mean Body Mass Index was 26.68 kg/m 2 (range: 23–42). According to the ASA score, 8 (22.22%) patients were classified as ASA 3. No ASA 4 was treated. Patients’ CCI are listed in Table  3 . The da Vinci Si platform was used for the first 15 (41.67%) cases, while Xi for the last 21 (58.33%). 2 (5.55%) procedures were performed at night. No conversions to open were reported in this series. According to Clavien-Dindo grade, 2 (5.55%) major complication was collected: 1 after urgent surgery and 1 after emergency setting. Two minor complications requiring conservative treatments were observed: both complications were related to primary disorders and not to RS. Intraoperative and 30-day mortality were 0%. The mean length of stay was 4.92 days (range: 1–21). The mean follow-up was 26.53 months (range: 7–68).

Number of elective and urgent/emergency procedures ( A ) and percentage of overall surgical procedures performed in emergency settings ( B ) according to abdominal surgery specialities. Abbreviations: HPB, Hepatopancreatic and Biliary Surgery;

Our experience demonstrates the safety and feasibility of RS also in urgent and emergency abdominal settings in patients not affected by hemodynamic instability. To date, the minimally invasive approaches in emergency scenarios are mainly validated for laparoscopy, as reported by several literature experiences included in the last WSES review [ 51 ]. Despite the diagnostic role of MIS, authors demonstrated several benefits of laparoscopic approaches in hemodynamically stable patients undergone AES, including trauma. However, patient selection, surgeons’ expertise as well as specific surgical training represent crucial key points [ 51 ].

In literature, RS in AES studies is related to high-volume centres, and well-trained surgeons’ experiences and their optimal outcomes should encourage further applications and Randomized Clinical Trials [ 32 , 52 ]. We further analyzed short- and long-term outcomes of RS in AES according to specialties (Table  4 ).

Robotic surgery in emergency setting

In the literature, RS in the emergency setting is reported by a limited number of experiences, especially case reports and case series.

The urologist experience described by Capibaribe et al. [ 53 ] demonstrated the safety and efficacy of robotic treatment in the case of vesicourethral anastomotic stenosis after open radical prostatectomy, providing better continence results, without pubectomy.

Globally, the major barrier to RS adoption is due to limited device access resulting from a shared use policy by several surgical teams (gynaecologists, general surgeons, thoracic surgeons, and urologists). Furthermore, the lack of dedicated teams (surgeons, nurses, and anaesthesiologists) during the night shift might further limit RS.

On the other hand, in emergency settings, the “time-sparing” concept is largely known. Commonly, to avoid useless costs due to waste disposable instruments, a hybrid approach should be discussed by the whole surgical team: before robotic docking, a laparoscopic exploration could be the first surgical step to verify clinical environments for doing RS.

A crucial issue is represented by frequent operating table position changes, especially during explorative steps (tilting, Trendelenburg, or reverse-Trendelenburg positions) and rapid conversion to open surgery when necessary [ 54 ]. It could be underlined that quick and safe docking and undocking are performed by skilled teams and well-trained surgeons in elective procedures [ 54 , 55 ].

In the last years, the RS technologies have also impacted operative time [ 56 ]: The Xi robot represents a radical evolution from the Si robot. Literature experiences demonstrated better docking ability during Da Vinci Xi surgery when compared to previous robotic systems (Da Vinci S, Si, X) [ 57 , 58 ]. These features were due to laser targeting and improved cannula mounts that resulted in a simplified “linear” port configuration and an abbreviated docking time.

Besides, the ability to exchange the robotic camera from port-to-port increased versatility for multi-quadrant surgeries thanks to the smaller 8 mm camera [ 59 , 60 ]. The multi-quadrant operations represent challenges due to the axis of visualization shift up to 360°. This procedure requires undocking the robot and rotating it on the axis. It is crucial for many colorectal surgeries that require access to the entire abdomen such as subtotal colectomy and total proctocolectomy.

Furthermore, it was reported that the Xi system’s better fluency is also due to thinner robotic arms that reduce their collisions during surgery and synchronous movements with the operating Table [ 56 ]. In addition, Da Vinci Xi integrates the Indocyanine-Green technology that could be easily used to better identify bile duct during cholecystectomy in patients affected by acute cholecystitis, to assess organ vascularization during their resections and anastomosis, as reported in our experience.

Bianchi et al. [ 61 ] performed an extensive comparison of Da Vinci Si and Xi systems to define their advantages and disadvantages. 89 patients (64 in the Si system vs. 25 in the Xi system group) who underwent liver surgery were included. The Si system group experienced a greater total incisional length (+ 8.99 mm; p  < 0.0001) due to a higher number of robotic/laparoscopic ports. Nevertheless, no differences were described regarding operative time, conversion rate, estimated blood loss, postoperative complications, mortality, use of analgesics, and costs. The authors concluded that da Vinci Xi represents an effective technological advancement.

Hill et al. [ 62 ] hypothesized that Da Vinci Xi will allow for greater efficiency and result in shorter operative times if compared to Da Vinci Si. To validate their hypothesis, the authors performed a retrospective review of patients undergoing sigmoid colon resection or Low Anterior Rectal resection. A total of 93 patients underwent sigmoid resection thanks to RS (Si, n  = 52 vs. Xi, n  = 41). The Xi group had significantly shorter surgical times for Low Anterior Rectal and sigmoid resection (162 vs. 238 min, p  = 0.0001). Nowadays, no data are available on the Da Vinci Si and Xi comparison in AES.

However, according to the type of procedures, the mean operative times of Da Vinci Xi were superimposable to the Si group in our experience.

The robotic technology in hemodynamically stable patients could potentially reduce the conversion to open rate (0% in our short series), thanks to high-definition view and accuracy of dissection and fine micro-sutures.

In 2022, the World Society of Emergency Surgery (WSES) published a position paper on RS in AES after the literature evaluation by a steering committee and an international expert panel [ 32 ]. Ten studies (3 case reports, 3 case series, and 4 retrospective comparative cohort articles) were found and 6 statements were proposed. Experts concluded that RS can be considered safe, and feasible in selected cases represented by hemodynamically stable patients. It should be emphasized that the WSES team reported some RS drawbacks: it is mandatory to perform dedicated surgical training, RS showed longer operative times, higher costs and difficult availability and accessibility represent the main issues during night shifts [ 32 ].

These aspects probably may change in the future with RS diffusion and new robotic devices in the health market.

In our experience, the mean age of patients was 63 years. Therefore, more than 30% of patients were older than 70 years (range: 43–88 years) and it is in line with RS literature experiences that showed good outcomes also in the elderly population [ 28 , 41 ]. Nevertheless, operative time represents a crucial point in this frail cohort. Despite RS showing longer operative time when compared to open and laparoscopic surgery, this disadvantage may be offset by lower postoperative complication rates, shorter hospital stays, and lower conversion rates [ 28 , 41 , 63 , 64 , 65 ].

Another key aspect is represented by enhanced vision through near-infrared imaging. It may be useful in AES in case of tissue perfusion evaluations or biliary tree identification in challenging procedures. This feature is not routinely available in laparoscopic surgery [ 66 , 67 ].

Figure  2 shows an emergency scenario due to splenic artery aneurysm repair thanks to RS.

The robotic approach during Emergency Setting for patients affected by splenic artery aneurysms using a vessel resection and end-to-end vascular anastomosis

Robotics in emergency upper-GI and bariatric surgery

One of the earliest studies on RS in AES was published in 2012 [ [ 68 ]]: Sudan et al. experience in complex bariatric surgery involved 2 patients affected by a stomach stricture and an acute abdomen due to perforation with biliary peritonitis after biliopancreatic diversion, respectively. The perforation was treated through an initial laparoscopic investigation followed by a handsewn robotic stitch reparation of duodenal stump dehiscence.

In 2020, Cubas et al. [ 69 ] presented an RS procedure for incarcerated Morgagni Hernia in a 29-year-old male. The hernia defect (reported as 10 × 7 cm) was corrected via mesh placement. Patient discharge was possible on POD 5. No recurrence was detected at 1-year follow-up.

During the same year, Ceccarelli et al. [ 45 ] published a series of 5 patients affected by strangulated Giant Hiatal Hernia: 3 (60%) patients experienced RS while 2 (40%) laparoscopic approach. The authors described an easier incarcerated stomach management thanks to RS, maybe due to better surgeon ergonomic position and more accurate dissection preserving pleural integrity and vagus nerve.

Kim et al. [ 70 ] reported a case of robotic transthoracic repair of a right-sided traumatic diaphragmatic rupture in a 45-year-old male with a history of chronic obstructive pulmonary disease presented as a restrained driver in a low-speed motor vehicle collision. The patient was effectively operated after a 48-hour observation.

In 2021, 300 USA hospitals were involved in retrospective data collection of adult patients affected by Hiatal Hernia and treated in elective and urgent/emergency scenarios from 2015 to 2017 [ 71 ]. Data analysis revealed that laparoscopy (64%) was the most frequent approach used during AES, followed by open surgery (30%). A limited number of patients (6%) experienced RS. After cost evaluations and outcomes analysis, authors declared the technical feasibility of minimally invasive approaches when compared to open surgery due to lower cost, lower length of hospital stay, complications, and mortality.

Robinson et al. [ 72 ], in 2021, performed a statistical analysis of “in-room-to-surgery-start time” in a retrospective cohort study of 44 patients affected by emergent perforated gastrojejunal ulcers. The comparison between RS and laparoscopic (24 and 20 cases respectively) showed encouraging results for RS (25 versus 31 min, p  = 0.01). Furthermore, no statistical differences were observed in terms of intra- and post-operative outcomes (operative time, complication rate, complication severity, hospital length of stay, discharge to home, and 30-day readmission). Despite RS showing higher surgical costs, authors concluded that emergency gastric perforation could be safely approached thanks to RS.

No complications were reported in all studies reported in our review [ 69 , 70 , 71 , 72 ].

Robotics in emergency colorectal surgery and appendectomies

Nowadays, emergency laparoscopy represents a safe and valid approach to colorectal disorders such as perforated diverticulitis with generalized peritonitis [ 73 ], iatrogenic colonoscopy perforations [ 74 ], bowel obstructions and anastomotic leaks management [ 75 , 76 , 77 ].

In 2012, Pedraza et al. [ 78 ] showed successful robotic colectomy due to iatrogenic colon perforation following colonoscopy.

Two years later, Felli et al. [ 79 ] described a case of an 86-year-old woman admitted to the emergency unit for massive intestinal bleeding due to ascending colon cancer. After patient resuscitation thanks to blood transfusions, surgeons carried out a robotic right colectomy. The postoperative period was uneventful.

Several series compared laparoscopic and robotic outcomes in patients who underwent elective colorectal surgery [ 80 , 81 , 82 ], suggesting the potential role of RS in this surgical field. Nevertheless, an interesting analysis was performed by Beltzer et al. [ 83 ] in 2019. 106 patients were treated for uncomplicated, complicated, or recurrent diverticulitis. The authors concluded that RS achieves better outcomes when compared to laparoscopic surgery in challenging cases (abscess or relapsing diverticulitis).

Three monocentric experiences reported by Kudsi et al. [ 84 , 85 , 86 ] showed the effectiveness of urgent RS for the treatment of obstructive transverse colon cancer, bleeding sigmoid diverticulosis and caecal volvulus.

However, RS could represent a crucial approach also in colorectal autoimmune diseases. Concerning this field, Anderson et al. [ 87 ] in 2020 reported a matched case-control study of 6 patients treated by urgent subtotal colectomy for ulcerative colitis using the robotic platform. In addition, authors compared patients who underwent RS to laparoscopic urgent procedures (6 versus 13 cases) concluding that no differences in perioperative outcomes were observed.

According to Yang et al. [ 88 ] estimation, more than 17 millions of patients were affected by appendicitis in 2019, making it the most common surgical emergency worldwide. Nevertheless, regarding urgent robotic appendectomies, only 5 literature experiences reported robotic approaches [ 89 , 90 , 91 , 92 , 93 ]. A total of 11 patients were collected and 3 (27.27%) required an appendix stump suture. No complications or conversions were reported. Moreover, Hüttenbrink et al. [ 94 ] described incidental appendicectomy during robotic prostatectomy.

Figure  3 shows our experience during RS for complicated sigmoid diverticulitis with sigmoid-bladder fistula.

Lunardi et al. [ 95 ] presented an interesting analysis of temporal trends in the use of minimally invasive surgery in Abdominal Emergency and Urgent Settings. The authors compared 89,098 emergency colectomies performed between 2013 and 2021. The increase per year for robotic colectomy was 0.9% (from 1.4% of total procedures in 2013 to 8.8% in 2021). As a result of this increase, a 0.7% decrease was registered for the open approach. Furthermore, patients who underwent RS were older, had more comorbidities and had higher BMI when compared to laparoscopic and open groups. Intraoperative outcomes were encouraging for RS: after Propensity Score Matching, a conversion rate of 25.5% (860/3,375 patients) was registered during laparoscopic surgery, while in 11.2% (379/3,375 patients) of RS cases, a conversion to open was required ( p  < 0.001). After Propensity Score Marching of patients underwent Emergency surgery only, RS demonstrated advantages in terms of conversion to open (27.5% vs. 12% in laparoscopic and robotic groups respectively, p  < 0.001) and post-operative LOS (7.12 vs. 6.85 days respectively, p  = 0.001).

In conclusion, conventional open surgery should be recommended for unstable and frail patients who require time-critical surgery. Nevertheless, it could be underlined that stable and frail patients may benefit from an enhanced recovery after surgery associated with RS in the acute setting when compared to open surgery.

Robotic approach during emergency setting for patients affected by complicated sigmoid diverticulitis with sigmoid-bladder fistula. We performed a fistula resection and bladder suture in double-layer barbed suture

Robotics in acute cholecystitis and biliary tree diseases

Another interesting field of application in AES may be the biliary tree and gallbladder diseases including cholecystitis, Mirizzi syndromes, biliary fistulas, iatrogenic diseases and common bile duct stones.

In 2016, Kubat et al. [ 96 ] published a retrospective series of 150 consecutive robotic single-site cholecystectomies (74 versus 76 cases treated in emergency scenarios and elective settings respectively). The mean operative time for ES cohort was significantly longer (95.0 ± 4.4 versus 71.9 ± 2.6 min; p  < 0.001). Both cohorts required 1 conversion to open (1.35% for the emergency group and 1.31% for the elective group). One bile duct injury (0.7%) was reported in patients treated in emergency conditions. The authors concluded that robotic single-site cholecystectomy can be performed safely and effectively in both elective and urgent scenarios with a learning curve of about 48 cases to reach acceptable perioperative outcomes.

Mirizzi syndrome represents one of the most challenging complications of cholelithiasis [ 97 , 98 , 99 ].

In 2014, Lee et al. [ 97 ] evaluated the outcomes of five patients treated by endoscopic biliary stent placement and subsequent robotic partial cholecystectomy due to Mirizzi syndrome. No conversion to open was reported and all patients experienced an uneventful postoperative course.

In 2017 Magge et al. [ 98 ] reported a 6-patient series. All cases were treated performing a combined endoscopic and robotic approaches. In 3 cases (50%) a Roux-en-Y hepatico-jejunostomy was carried out. In these challenging scenarios, RS showed relevant benefits when compared to laparoscopy, facilitating complex dissections, and reducing conversion to open rate.

The most representative cohort of patients was described by Gangemi et al. [ 100 ] in 2017. Authors compared a large series of 676 patients receiving a robotic cholecystectomy with 284 treated by conventional laparoscopy: data analysis showed a significantly lower conversion to open in RS group, especially in patients affected by acute or gangrenous cholecystitis.

A 3-patient experience was described by Milone et al. [ 101 ] in 2019, achieving good perioperative outcomes in acute cholecystitis treatment.

Major bile duct injuries after cholecystectomy require complex surgical repairs that are usually performed with a conventional open approach [ 102 ]. This field may represent an interesting application of RS to safety perform biliary anastomosis. Cubisino et al. presented a systematic review of 13 literature experiences on minimally invasive biliary anastomosis after iatrogenic bile duct injury [ 103 ]. 198 patients were included. 135 patients (63.1%) underwent laparoscopic biliary anastomosis, while 73 (36.1%) received an analogue robotic procedure. According to Strasberg’s classification [ 104 ], all Bile Duct Injuries were types D and E (E1–E5). No conversions occurred in the RS series, while 4 patients required conversion to open surgery among the laparoscopic ones. Postoperative complications were superimposable (18.7% and 19.7% in laparoscopic and robotic approaches, respectively). Nevertheless, the overall reoperation rate was 4.4%, 5.5% in laparoscopic and 2.6% in robotic repairs.

During the follow-up period (median 24.6 months), 9 patients developed an anastomotic stricture: 5 (3.70%) in laparoscopic and 4 (5.48%) in robotic series that required a redo-anastomosis in 60% and 25% respectively.

When compared to open and laparoscopic cholecystectomy in AES, RS showed an increase of 0.7% per year in Lunardi et al. cohort of 793’800 cholecystectomies [ 95 ], ranging from 2.5 to 8.8% between 2013 and 2021. It could be underlined that conversion rate and LOS were statistically lower in RS group ( p  < 0.001). Despite these findings, laparoscopic cholecystectomy yet represents the preferred approach in AES.

Robotics emergencies in hernia and abdominal wall surgery

Only a few studies analyzed urgent hernia operations treated using robotic surgery.

In 2020, Bou-Ayash et al. [ 105 ] published a retrospective series of 19 patients (including 23 surgical procedures) affected by inguinal hernia, treated from 2013 to 2020. The authors concluded that the robotic approach represents a safe procedure in selected patients, with a short length of stay and a low complication rate compared to open and laparoscopic surgery.

In 2021, Kudsi et al. [ 106 ] described perioperative outcomes of RS in a 34-patient cohort treated between 2013 and 2019. All patients experienced robotic ventral and incisional hernia repair in an emergency setting. 20% of patients were classified as Clavien-Dindo I or II, while about 11% Clavien-Dindo III and IV. Only 3% of the population experienced a recurrence.

Muysoms et al. [ 107 ] performed an extensive analysis of robotic cost. They retrospective evaluate laparoscopic (272 procedures of which 6 were emergency cases) and robotic (404 procedures of which 8 were emergency cases) inguinal hernia repairs. As reported in other literature experiences, authors concluded that Robotic inguinal hernia repair was significantly ( p  < 0.001) more expensive if compared to laparoscopic surgery (mean cost €2612 versus €1963, respectively). Nevertheless, in the robotic group, a larger number of patients were treated as outpatients with lower postoperative complications.

Regarding inguinal and ventral hernia repair, the analysis conducted by Lunardi et al. [ 95 ] showed encouraging data for RS approach: from 2013 to 2021 RS increased of 1.9% per year and 1.1% per year respectively. After propensity score matching, authors reported superimposable data in terms of CCI and BMI, comparing laparoscopic and robotic approaches. Nonetheless, RS showed benefits also in these fields: lower conversion rates were reported both in inguinal hernia repairs (18.1% vs. 3.8%, p  < 0.001) and in ventral hernia repair (16.2% vs. 4.8%, p  < 0.001). In addition, a statistically significant shorter postoperative LOS was registered in the RS group (the mean LOS in the inguinal hernia group was 3.34 vs. 3 days in laparoscopic and robotic approaches respectively, and the mean LOS in the ventral hernia group was 3.87 vs. 3.73 days, respectively).

Other abdominal emergency surgery and future perspectives

A rare indication for urgent RS was post-traumatic splenic bleeding reported by Giulianotti et al. [ 108 ].

Until now, no reports of RS in adhesive intestinal obstruction have been published.

A possible and useful application of RS is represented by telementoring and telesurgery [ 32 , 109 , 110 , 111 ]. The original aim of RS and the recent COVID-19 pandemic gave an important incentive in these directions. The advantage of telementoring and telepresence of an expert surgeon in a virtual way is nowadays possible and may be improved thanks to the modern and future highspeed internet connection (5G networks) as well as the telesurgery in ultra-remote countries, in low-volume centers and in an emergent civil or battlefield surgical scenarios [ 112 , 113 , 114 ].

The development of new modular robotic platforms may contribute to increase RS applications in emergency settings. Nowadays, several different robotic platforms are approved for human use, such as CMR Versius (Cambridge Medical Robotics, Cambridge, UK), Distalmotion Dexter (Distalmotion, Epalinges, Switzerland) and Medtronic Hugo (Medtronic Inc., Minneapolis, USA). Most of them share the opportunity of switching from a conventional laparoscopic setting to a robot-assisted one.

Limitations

The main bias of our study was represented by hospital organisations: RS devices are available in the same building as the General Surgery Unit at San Donato Hospital (Arezzo, Italy) facilitating emergency surgical procedures. On the other hand, Da Vinci Xi is situated in a separate building specifically dedicated to RS at the General and Robotic Surgery Unit of San Giovanni Battista Hospital (Foligno, Italy).

Furthermore, these findings represented a limit when the surgeons’ team wanted to perform a laparoscopic exploration to validate a minimally invasive robotic approach in emergency scenarios.

In our experience, it should also underline that the COVID-19 era has contributed to limiting RS adoption.

Future shreds of evidence from randomized clinical trials with long-term follow-up are required to define the potential role of RS in AES. Nevertheless, the unavailable data on the cost-effectiveness of RS in AES are linked to lower use of robotic devices if compared to laparoscopic approaches. Our experience suggested that RS costs are superimposable to laparoscopic surgery if we analyse LOS and conversion rate data. To optimize the delivery of robotic technology in AES, a well-coordinated effort among health systems, clinicians, payers, and policymakers and dedicated training program for robotic teams are imperative.

Our study demonstrates that RS may be an useful and reliable approach also to emergency surgical procedures, especially when performed in selected patients in very well trained robotic centers allowing a safe managing of surgical challenging procedures as main indications for this technology, reducing the conversion rate when compared to laparoscopy.

As for laparoscopy the patient selection for robotic approach need hemodinamically stable condition and require a sharing of the surgical strategy by all the team: surgeons, nurses and anaesthesiologists. All the staff need to be trained in laparoscopic and robotic elective surgery, including technology functioning. The hybrid use of robotic/laparoscopic technology may be taken into consideration (a laparoscopic exploration may be the first step) to decide the following approach. The robotic approach may be reserved to challenging steps of the operation (suture/microsuture/dissections).

The availability of the device is the sine qua non condition for emergent and of course urgent use. The current organization in which the platform is shared by different teams, represent for the diffusion of its use in the emergent setting.

The cost reduction of platforms and instruments, together with new robotic devices in the health market, may represent a future perspective for emergencies use of robotic technology. So, the robotic technology may be one of the tools available in every operating theatre, to use in selected cases according to patient condition and surgical team experience.

Data availability

No datasets were generated or analysed during the current study.

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G.C.: study conception and design, literature search, data acquisition, interpretation and analysis; drafting and critically revising the article for important intellectual content; final approval of the version to be published. P.A.: literature search, data acquisition, interpretation and analysis; drafting and critically revising the article for important intellectual content; final approval of the version to be published. F.C.: drafting and critically revising the article for important intellectual content; final approval of the version to be published. B.W.T.: data acquisition, interpretation and analysis; final approval of the version to be published. F.R.: study conception and design, literature search, data acquisition, interpretation and analysis; final approval of the version to be published. All authors: data acquisition, interpretation, and analysis; final approval of the version to be published. A.R.: supervision, drafted and critically revised the article for important intellectual content; final approval of the version to be published.

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Ceccarelli, G., Catena, F., Avella, P. et al. Emergency robotic surgery: the experience of a single center and review of the literature. World J Emerg Surg 19 , 28 (2024). https://doi.org/10.1186/s13017-024-00555-6

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Analysing near-miss incidents in construction: a systematic literature review.

1. Introduction

• Q 1 —Are near-miss events in construction industry the subject of scientific research?
• Q 2 —What methods have been employed thus far to obtain information on near misses and systems for recording incidents in construction companies?
• Q 3 —What methods have been used to analyse the information and figures obtained?
• Q 4 —What are the key aspects of near misses in the construction industry that have been of interest to the researchers?

2. Definition of Near-Miss Events

3. research methodology, 4.1. a statistical analysis of publications, 4.2. methods used to obtain information about near misses, 4.2.1. traditional methods.

• Traditional registration forms
• Computerized systems for the recording of events
• Surveys and interviews

4.2.2. Real-Time Monitoring Systems

• Employee-tracking systems
• Video surveillance systems
• Wearable technology
• Motion sensors

4.3. Methods Used to Analyse the Information and Figures That Have Been Obtained

4.3.1. quantitative and qualitative statistical methods, 4.3.2. analysis using artificial intelligence (ai), 4.3.3. building information modelling, 4.4. key aspects of near-miss investigations in the construction industry, 4.4.1. occupational risk assessment, 4.4.2. causes of hazards in construction, 4.4.3. time series of near misses, 4.4.4. material factors of construction processes, 4.5. a comprehensive overview of the research questions and references on near misses in the construction industry, 5. discussion, 5.1. interest of researchers in near misses in construction (question 1), 5.2. methods used to obtain near-miss information (question 2), 5.3. methods used to analyse the information and data sets (question 3), 5.4. key aspects of near-miss investigations in the construction industry (question 4), 6. conclusions.

• A quantitative analysis of the Q 1 question has revealed a positive trend, namely that there is a growing interest among researchers in studying near misses in construction. The greatest interest in NM topics is observed in the United States of America, China, the United Kingdom, Australia, Hong Kong, and Germany. Additionally, there has been a recent emergence of interest in Poland. The majority of articles are mainly published in journals such as Safety Science (10), Journal of Construction Engineering and Management (8), and Automation in Construction (5);
• The analysis of question Q 2 illustrates that traditional paper-based event registration systems are currently being superseded by advanced IT systems. However, both traditional and advanced systems are subject to the disadvantage of relying on employee-reported data, which introduces a significant degree of uncertainty regarding in the quality of the information provided. A substantial proportion of the data and findings presented in the studies was obtained through surveys and interviews. The implementation of real-time monitoring systems is becoming increasingly prevalent in construction sites. The objective of such systems is to provide immediate alerts in the event of potential hazards, thereby preventing a significant number of near misses. Real-time monitoring systems employ a range of technologies, including ultrasonic technology, radio frequency identification (RFID), inertial measurement units (IMUs), real-time location systems (RTLSs), industrial cameras, wearable technology, motion sensors, and advanced IT technologies, among others;
• The analysis of acquired near-miss data is primarily conducted through the utilisation of quantitative and qualitative statistical methods, as evidenced by the examination of the Q 3 question. In recent years, research utilising artificial intelligence (AI) has made significant advances. The most commonly employed artificial intelligence techniques include text mining, machine learning, and artificial neural networks. The growing deployment of Building Information Modelling (BIM) technology has precipitated a profound transformation in the safety management of construction sites, with the advent of sophisticated tools for the identification and management of hazardous occurrences;
• In response to question Q 4 , the study of near misses in the construction industry has identified several key aspects that have attracted the attention of researchers. These include the utilisation of both quantitative and qualitative methodologies for risk assessment, the analysis of the causes of hazards, the identification of accident precursors through the creation of time series, and the examination of material factors pertaining to construction processes. Researchers are focusing on the utilisation of both databases and advanced technologies, such as real-time location tracking, for the assessment and analysis of occupational risks. Techniques such as Analytic Hierarchy Process (AHP) and clustering facilitate a comprehensive assessment and categorisation of incidents, thereby enabling the identification of patterns and susceptibility to specific types of accidents. Moreover, the impact of a company’s safety climate and organisational culture on the frequency and characteristics of near misses represents a pivotal area of investigation. The findings of this research indicate that effective safety management requires a holistic approach that integrates technology, risk management and safety culture, with the objective of reducing accidents and enhancing overall working conditions on construction sites.

7. Gaps and Future Research Directions, Limitations

• Given the diversity and variability of construction sites and the changing conditions and circumstances of work, it is essential to create homogeneous clusters of near misses and to analyse the phenomena within these clusters. The formation of such clusters may be contingent upon the direct causes of the events in question;
• Given the inherently dynamic nature of construction, it is essential to analyse time series of events that indicate trends in development and safety levels. The numerical characteristics of these trends may be used to construct predictive models for future accidents and near misses;
• The authors have identified potential avenues for future research, which could involve the development of mathematical models using techniques such as linear regression, artificial intelligence, and machine learning. The objective of these models is to predict the probable timing of occupational accidents within defined incident categories, utilising data from near misses. Moreover, efforts are being made to gain access to the hazardous incident recording systems of different construction companies, with a view to facilitating comparison of the resulting data;
• One significant limitation of near-miss research is the lack of an integrated database that encompasses a diverse range of construction sites and construction work. A data resource of this nature would be of immense value for the purpose of conducting comprehensive analyses and formulating effective risk management strategies. This issue can be attributed to two factors: firstly, the reluctance of company managers to share their databases with researchers specialising in risk assessment, and secondly, the reluctance of employees to report near-miss incidents. Such actions may result in adverse consequences for employees, including disciplinary action or negative perceptions from managers. This consequently results in the recording of only a subset of incidents, thereby distorting the true picture of safety on the site.

Author Contributions

Institutional review board statement, informed consent statement, data availability statement, conflicts of interest.

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Woźniak, Z.; Hoła, B. Analysing Near-Miss Incidents in Construction: A Systematic Literature Review. Appl. Sci. 2024 , 14 , 7260. https://doi.org/10.3390/app14167260

Woźniak Z, Hoła B. Analysing Near-Miss Incidents in Construction: A Systematic Literature Review. Applied Sciences . 2024; 14(16):7260. https://doi.org/10.3390/app14167260

Woźniak, Zuzanna, and Bożena Hoła. 2024. "Analysing Near-Miss Incidents in Construction: A Systematic Literature Review" Applied Sciences 14, no. 16: 7260. https://doi.org/10.3390/app14167260

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