REVIEW OF LITERATURE

Xylanase is an industrial enzyme having applications in a wide range of processes. It is a glycoside hydrolase that catalyzes the

hydrolysis of xylan producing mainly xylobiose, xylotriose, and a small fraction of xylooligosacharides with higher degree of

polymerization. The present investigation aims at isolation of a potent xylanolytic bacterial strain and study of the production,

purification, characterization, immobilization and application of the xylanase secreted by this strain. Several reviews have been

published on xylanases (Gilbert and Hazlewood, 1993; Bajpai et al., 1994; Wong et al., 1988; Srinivasan and Rele, 1999; Bhat, 2000;

Bajpai, 2004; Beg et al., 2001; Subramaniyan and Prema, 2002; Sá-Pereira et al., 2003; Collins et al., 2005; Pollizeli et al., 2005;

Dhiman et al., 2008b; Juturu and Wu, 2011; Kuhad and Singh, 1993). This chapter presents a brief review of the literature pertaining

mainly to isolation, production, purification and characterization, immobilization, and application of bacterial xylanases.

2.1 Chemical structure and distribution of xylan (the substrate for xylanase)

Xylan is the substrate for xylanase. It a major component of hemicelluloses present in plant cell walls and is the second most

abundant polysaccharide next to cellulose in nature, accounting for approximately one-third of all renewable organic carbon on earth

(Biely, 1985; Prade, 1995). The term hemicellulose refers to a group of non-cellulosic polysaccharides which include xylan,

xyloglucan (a heteropolymer of D-xylose and D-glucose), glucomannan (a heteropolymer of D-mannose and D-glucose),

galactoglucomannan (a heteropolymer of D-galactose, D-glucose and D-mannose), arabinogalactans (a heteropolymer of D-galactose

and arabinose) (Shallom and Shoham, 2003). In plant cell wall, hemicelluloses occur in association with cellulose and lignin

constituting lignocellulose which on an average contains 35-45% cellulose, 20-30% hemicellulose and 8-15% lignin (a polymer of

phenylpropanoid residues). β-1,4-Xylans are mainly found in secondary walls, the major component of mature cell walls in woody

tissue. Although they also represent the major hemicellulose in the primary walls of monocots, xylans generally constitute a minor

component of the primary walls in dicots. Xylan functions as an adhesive by forming covalent and non-covalent bonds with lignin,

cellulose and other polymers essential to the integrity of the cell wall. Xylan is considered to form an interface between lignin and

cellulose (Kulkarni et al., 1999; Beg et al., 2001). The phenolic residues of lignin are linked to xylan by an ester linkage to 4-O-

methyl-D-glucuronic acid residues. Lignocellulosic materials in the form of agro-residues can serve as excellent substrate for inducing

xylanase production. Xylan is found in large quantities in hardwoods from angiosperms, softwoods from gymnosperms and annual

plants constituting 15–30%, 7–10% and

(A)

(B)

Fig. 2.1 Schematic representation of xylan structure: A) Softwood xylan and (B) Hardwood xylan (Woodward, 1984)

Xyl= xylopyranose residue; Me= Methyl group; GluA=Glucuronic acid residue; Arab= L-Arabinofuranosyl residue,

and Ac=Acetyl group

2.2 Xylanolytic enzymes

-------------------β(1 4)D-Xyl-β(1 4)D-Xyl-β(1 4)D-Xyl-β(1 4)D-Xyl--------------------- 2 3(2)

1 1 4-O-Me-α-D-GlcA- OAc

--------------------β(1 4)D-Xyl3-β(1 4)D-Xyl-β(1 4)D-Xyl7-β(1 4)D-Xyl----- 2 3

1 1 (4-O-Me-α-D-GlcA)3 α-L-Arabinose

A variety of xylanolytic enzymes are produced by microbes. Owing to the heterogeneity and complex chemical nature of

xylan, its complete hydrolysis to its constituent sugars requires the concerted action of several enzymes including endo-1,4-β-xylanase

(1,4-β-D-xylan xylanohydrolase; EC 3.2.1.8), β-D-xylosidase (1,4-β-D-xylan xylohydrolase; EC 3.2.1.37), α-L-arabinofuranosidase

(α-L-Arabinofuranosidase; EC 3.2.1.55), acetylxylan esterase (EC 3.2.1.6), α-D-glucuronidase (α-glucosiduronase; EC 3.2.1.139),

feruloyl esterase (EC 3.1.1.73) and p-coumaroyl esterase (Biely, 1985; Kuhad et al., 1997; Beg et al., 2001; Subramaniyan and Prema,

2002). Figure 2.2 shows a schematic structure of plant xylan along with the sites of action of various xylanolytic enzymes. Indeed,

complete xylanolytic enzyme systems, including all of the above activities have been found to be quite widespread among fungi

(Belancic et al., 1995; Sunna and Antranikian, 1997), actinomycetes (Elegir et al., 1994) and bacteria (Sunna and Antranikian, 1997;

Subramaniyan and Prema, 2002). Among the various xylanolytic enzymes, endoxylanase and β-xylosidase are the key enzymes

responsible for xylan hydrolysis. Xylan does not form tightly packed structures and is thus more accessible to hydrolytic enzymes.

Consequently, the specific activity of xylanases is 2-3 orders of magnitude greater than for cellulase hydrolysis of crystalline cellulose

(Gilbert and Hazlewood, 1993).

Endoxylanase (EC 3.2.1.8) catalyzes cleavage of the internal β-1,4-glycosidic bonds in the xylan backbone producing

xylobiobe, xylotriose and a small fraction of xylooligosaccharides with higher degree of polymerization. β-xylosidase acts on these

xylooligomers releasing xylose. Xylan is not attacked randomly, but the bonds selected for hydrolysis depend on the chain length,

degree of branching and the presence of substituents in the substrate molecule (Li et al., 2000). Although many xylanases are known

to release xylose during the hydrolysis of xylan or xylooligosaccharides, xylobiase activity has only been reported in β-xylosidases

(Wong et al., 1988).

β-D-Xylosidases (EC 3.2.1.37) catalyzes the hydrolysis of small xylooligosaccharides and xylobiose releasing xylose from the

non-reducing terminus. These can be classified according to their relative affinities for xylobiose and larger xylooligosaccharides.

Biely (1985) proposed that xylobiases and exo-1,4-β-xylanases can be recognized as distinct entities but are treated as xylosidases.

This enzyme is also able to cleave artificial substrates such as p-nitrophenyl- and o-nitrophenyl-β-D-xylopyranoside. The action of

xylanase may lead to the accumulation of short xylooligomers, which may inhibit the endoxylanase but β-xylosidase hydrolyzes these

products, removing the cause of inhibition, and thereby increasing the efficiency of xylan hydrolysis (Zanoelo et al., 2004). The

activity of β-xylosidase has been reported in Bacillus sp. (Panbangred et al., 1984) and fungi (Li et al., 2000).

α-D-Glucuronidase catalyzes the release of D-glucuronic acid or 4-O-methyl-D-glucuronic acid residues attached to the β-D-

xylopyranosyl residues of the glucuroxylan backbone via α-1,2-glycosidic bonds. The substrate specificity of glucuronidase varies

with the microbial source.

α-L-Arabinofuranosidase (EC 3.2.1.55) removes the α-L-arabinofuranosyl residues substituted at C-2 and C-3 of the β-D-

xylopyranosyl residues in the xylan backbone (Morales et al., 1995). These enzymes are of two types with distinct modes of action:

exo-α-L-arabinofuranosidase (EC 3.2.1.55) which degrades p-nitrophenyl-α-L-arabinofuranosides and branched arabinans whereas

endo-1,5-α-L-arabinase (EC 3.2.1.99) which hydrolyzes only linear arabinans. Most arabinofuranosidases investigated so far are of the

exo type.

Acetylxylan esterase (EC 3.1.1.6) catalyzes the removal of O-acetyl groups attached at either C-2 and/or C-3 of β-D-

xylopyranosyl residues of the acetylxylan backbone. Feruloyl esterase (EC 3.1.1.73) catalyzes cleavage of the bond between arabinose

side chain residue and ferulic acid whereas p-coumaroyl esterase (EC 3.1.1.-) catalyzes the cleavage of the bond between arabinose

side chain residue and p-coumaric acid (Christov and Prior, 1993).

Apparently, there is a considerable degree of synergy between xylanolytic enzymes (Poutanen et al., 1991). For example,

many xylanases will not cleave glycosidic bonds between xylose units which are substituted. Thus, side chains must be cleaved before

the xylan backbone can be completely hydrolyzed. Conversely, several accessory enzymes will only remove side chains from

xylooligosaccharides. These enzymes, therefore, require xylanases to partially hydrolyze the plant structural polysaccharide, before

side-chains can be cleaved (Poutanen et al., 1991). Synergistic and cooperative effects among the xylan-degrading enzymes enhance

the susceptibility of the heteropolymeric xylan to be attacked by endoxylanases (De Vries et al., 2000). Thus, the removal of acetyl

group from acetylxylan by acetyl xylan esterase increases the accessibility of the xylan backbone for endoxylanase attack. Conversely,

endoxylanase creates shorter acetylated polymers, which are preferred substrates for esterase activity (Biely et al. 1985, 1986). β-

Xylosidase enhances the hydrolysis of xylan by hydrolyzing the xylooligosaccharides generated by the action of endoxylanase and

hence relieving the end-product inhibition of endoxylanase. Addition of α-L-arabinofuranosidase to endoxylanase enhances the

saccharification of arabinoxylans.

Fig. 2.2 Schematic representation of chemical structure of plant xylan showing the sites of cleavage by different xylanolytic

enzymes (Source: Beg et al., 2001)

2.3 Occurrence of xylanases (EC 3.2.1.8)

Xylanases are produced mainly by microorganisms and take part in the breakdown of plant cell walls, along with other

enzymes that hydrolyze polysaccharides. Microbial sources of xylanases include bacteria (Akiba and Horikoshi, 1988; Sunna and

Antranikian, 1997; Beg et al., 2001; Anuradha et al., 2007; Ali et al., 2013; Anand et al., 2013; Mittal et al., 2013; Verma et al., 2013),

actinomycetes (Ninawe et al., 2007; Bajaj and Singh, 2010; Kumar et al., 2012; Thomas et al., 2013) and fungi (Sunna and

Antranikian, 1997; Polizeli et al., 2005). Xylanases also can be found in marine algae, protozoans, crustaceans, insects, snails and

seeds of land plants (Sunna and Antranikian, 1997; Kuhad et al., 1997). Bacterial xylanases mainly belong to the genus Bacillus,

Cellulosimicrobium, Chromohalobacter, Geobacillus, Paenibacillus, Flavobacterium and Arthrobacter. Among the bacteria, Bacilli

are the most important xylanolytic enzyme producers, which are known to produce elevated levels of xylanase exhibiting activity at

alkaline pH and high temperature (Akiba and Horikoshi, 1988; Sunna and Antrakian, 1997; Beg et al., 2001; Subramaniyan and

Prema, 2002; Lama et al., 2004; Mamo et al., 2006; Kumar and Satyanarayana, 2011; Nagar et al., 2012c; Kumar et al., 2013b).

Mesophilic xylanolytic fungi include Aspergillus sp., Trichoderma sp., Thermomyces sp., Fusarium sp. and Penicillium sp. among

others (Polizeli et al., 2005). Among actinomycetes, Streptomyces species are well known producers of xylanolytic enzymes (Beg et

al., 2001; Ninawe et al., 2006).

2.4 Multiplicity of xylanases

In addition to the production of a variety of xylanolytic enzymes, many micro-organisms produce multiple xylanases (Gilbert

and Hazlewood, 1993; Rizzatti et al., 2004). 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. For example, Aspergillus niger and Trichoderma viride secrete 15 and 13 isoenzymes of

xylanases (Biely et al., 1985). This multiplicity may be the result of genetic redundancy (Wong et al., 1988) and differential post-

translational processing (Biely, 1985). 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 (Collins et al., 2005).

2.5 Classification of xylanases

Wong et al. (1988) classified endoxylanases on the basis of their physicochemical properties, such as molecular weight (MW)

and isoelectric point (pI), into two groups: enzymes having basic pI and MW 30 kDa.

However, exceptions to this classification were found (Sunna and Antranikian, 1997) and about 30% xylanases, particularly fungal

enzymes, do not fit into this classification. Later, Henrissat et al. (1989) introduced a new classification system based on primary

structure comparisons of the catalytic domains only which classified not only xylanases, but glycosidases in general (EC 3.2.1.x), and

this system is now followed for the classification of these enzymes. The initial classification by Henrissat et al. (1989) grouped

cellulases and xylanases into 6 families (A–F) of related sequences, which continued to grow as new glycosidase sequences were

identified. The number of glycoside hydrolase families was 77 (1-77) in 1999 (Henrissat and Coutinho, 2001) and 96 in 2005

(carbohydrate–active enzyme CAZY server at http://afmb.cnrs-mrs.fr/~cazy/CAZY/) with approximately one-third of these families

containing enzymes with diverse substrate specificities (Collins et al., 2005). Enzymes within a particular family have a similar three-

dimensional structure (Henrissat and Coutinho, 2001) and similar molecular mechanism. In some families protein folds are more

conserved than their amino acid sequences, and these families are grouped into clans. Endo-xylanases fall into two main glycoside

hydrolase families 10 (formerly F) and 11 (formerly G) based on amino acid sequence similarities (Henrissat & Bairoch,

1996). The xylanases in these families differ both in their structure and catalytic properties (Biely et al., 1997). Family 10 (GH 10)

endoxylanases have high molecular mass, are more complex and diverse and hydrolyze both xylan and cellulose. They have a

(β/α)8 fold TIM barrel structure composed of a cellulose-binding domain and a catalytic domain connected by a linker peptide (Biely

et al., 1997). The GH 10 xylanases are capable of attacking the glycosidic linkages next to the branch points and towards the non-

reducing end. For cleavage, these enzymes require two unsubstituted xylopyranosyl residues between the branches. On the other hand,

family 11 (GH 11) xylanases have low molecular mass (

the production of endoxylanase. Xylanolytic activity was found in 10 of the 88 isolates. Two best endoxylanase producers (SB-9a and

TC-17d) belonged to the genus Bacillus.

Khandeparker and Bhosle (2007) isolated a xylanase producing bacterial strain from sediment sample collected from Mandovi

estuary, west coast of India using xylan in the isolation medium. The bacterial culture was identified as Arthrobacter sp. MTCC 5214.

Ko et al. (2007) isolated a xylanolytic bacterial strain from the samples of black liquor (released in kraft process) collected from

Hsinying paper mill of Taiwan. The isolated strain was identified as Paenibacillus sp. BL11 by using the 16S rRNA gene

amplification. Amplified 16S rDNA sequence (1545 bp) has been deposited in the GenBank data base under accession no. DQ232773.

Prakash et al. (2009) reported an extremely halophilic bacterium, Chromohalobacter sp. TPSV 101, which was able to produce

various exoenzymes which exhibited xylanase activity at 20% (w/v) NaCl. Roy and Rowshanul (2009) isolated Bacillus sp from soil

sample of Rajshahi university campus. The isolated bacterium was an aerobic, gram-positive, spore forming, rod-shaped organism and

was identified as Bacillus cereus on the basis of 16S rRNA gene amplification, morphological properties and taxonomic

characteristics.

Azeri et al. (2010) isolated four xylanolytic bacterial stains from Soda Lake (pH 8.2, salt concentration 9.9%) on media

containing xylan and characterized these on the basis of morphological, physiological and biochemical characters. These stains were

identified as belonging to the genus Bacillus. Inan et al. (2012) isolated a novel moderately thermophilic, Gram positive, endospore

forming, rod shaped, motile and alkaline active xylanase- producing bacterial strain D1021T from Kaynarca hot spring in the province

of Izmir, Turkey. The isolated bacterial strain was identified as Anoxybacillus kaynarcensis on the basis of phenotypic characteristics,

rpoB analysis and 16S rRNA gene amplification. Kaur et al. (2011b) isolated a xylanopectinolytic bacterial strain from the soil

contaminated with effluents of paper and pulp industry using cost-effective wheat bran in the isolation medium. The strain was

identified as B. pumilus by the Microbial Type Culture Collection Centre of IMTECH, Chandigarh, India. Khianngam et al. (2011)

isolated a xylan-degrading bacterium Paenibacillus xylanisolvens, which was Gram positive, facultative anaerobic and rod-shaped.

This strain was identified on the basis of its phenotypic characteristics and 16S rRNA gene amplification.

Kamble and Jadhav (2012b) reported the isolation of a cellulase-free xylanase producing bacterial strain and identified it as

Cellulosimicrobium sp. on the basis of morphological, physiological, biochemical characteristics as well as 16S rRNA sequencing by

Microbial Type Culture Collection (MTCC) and Gene Bank, Institute of Microbial Technology (IMTECH), Chandigarh, India. The

nucleotide length of the rRNA gene was 1408 bp and deposited in the genbank of NCBI with accession number FR729925.1. The

bacterial strain was deposited as Cellulosimicrobium sp. MTCC 10645. Xylanase activity of 4,962 U/gds was obtained after

optimization of xylanase production in SSF. Nagar et al. (2012) isolated 70 xylanolytic bacterial strains from 14 samples of decaying

wood, soil and compost on the basis of their zone of hydrolysis on wheat bran-agar medium, which is cost-effective as compared to

xylan-agar. Of all the isolates, five strains were selected as hyper producers of cellulase-free xylanase.

Abo-State et al. (2013) isolated 59 xylanolytic bacterial strains from different agricultural wastes viz. wheat bran, rice straw,

wheat straw, corn cob, sugarcane bagasse, potato peal and banana peel. These materials were also used as substrates for production of

xylanase. Ali et al. (2013) isolated 300 isolates from soil samples collected from different locations of Saudi Arabia and Egypt. The

best isolate was identified as Bacillus subtilis by the 16S rRNA gene amplification followed by similarity searching. The sequence of

the amplified 16S rRNA gene from this bacterial strain was deposited in GenBank under the accession number JF801740.

2.7 Production of bacterial xylanase

Microbial xylanase may be produced in submerged (SmF) or solid substrate fermentation (SSF). SmF involves growth of the

desired microorganism as a suspension in liquid medium in which various nutrients are either dissolved or suspended as particulate

solids. It is the preferred method for the production of most of the commercial enzymes, principally because sterilization and process

control are easier in these systems. About 90% of the total xylanase are produced worldwide by most of the xylanase manufacturers

using submerged fermentation techniques (Polizeli et al., 2005). On the other hand, SSF is the growth of microorganisms on moist

substrates in the absence or near absence of free-flowing water. The solid substrates act as source of carbon, nitrogen, minerals and

growth factors and have the capacity to absorb water in order to provide natural habitat and growth requirements of microbes. Enzyme

production in SSF is usually much higher than that of submerged fermentation (Haltrich et al., 1996). Enzyme production in SSF may

offer several economical and practical advantages over submerged cultivation such as need of simpler equipment, a simple

fermentation medium, requirement of less energy, higher product yield, reduced waste water output, lower capital and operational

costs, low catabolic repression, and does not require a rigorous control of fermentation parameters (Pandey et al., 1999; Krishna,

2005). Bacterial systems are being increasingly investigated for the production of enzymes and metabolites through SSF. Xylanase

production in SSF is of interest for countries like India with abundant biomass and agro-residues.

Culture conditions for xylanase production in SmF or SSF may differ for various bacterial strains/species. Each bacterial strain

may have a different set of optimal conditions for maximum enzyme yield. Therefore, optimization of culture conditions is essential

for obtaining maximum enzyme yield from the desired bacterial strain. Optimum concentration of inoculum is necessary for

maintaining balance between proliferating biomass and available nutrients to produce maximum enzyme level. An inoculum level

higher than the optimum could result in faster nutrient consumption and hence lower enzyme yield. Moreover, a large inoculum size is

undesirable during scale up of the fermentation process and is not preferred in industrial fermentation. Inoculum size required for SSF

is generally much larger than that required for SmF.

Cultivation period for the enzyme production is the time taken by the microorganism for utilizing the available nutrients from

the culture medium for the synthesis of desired product. The period of incubation depends upon the type of fermentation, growth of

microorganism and its growth pattern. Generally, the incubation period required by a microorganism for xylanase production in SSF is

longer as compared to SmF. Further, the duration of incubation period also varies according to the microorganism being smaller for

bacterial cultures as compared to fungi because the growth rate of the former is faster than the latter.

Incubation temperature and pH are the most important physical variables affecting the microbial growth and enzyme

production. Each bacterial strain results in maximum enzyme yield at its unique optimum growth temperature and pH of the

production medium. The pH of the production medium may change during fermentation due to metabolic activities of the

microorganism. Different types of microorganisms, species and genus require unique cultivation temperature and pH for enzyme

production. The optimum pH for xylan hydrolysis is around five for most fungal xylanases and they are normally stable between pH

values of two and nine. The pH optima of bacterial xylanases are generally slightly higher than the pH optima of fungal xylanases.

Alkalophilic Bacillus species and actinomycetes produce xylanases with high activity at alkaline pH value (Biely, 1985; Srinivasan

and Rele, 1999). Most of the fungal xylanases tolerate temperatures below 50°C (Subramaniyan and Prema, 2002). As compared to

fungi, bacteria are good source of alkaline and thermo stable xylanases (Kumar et al., 2013).

Carbon and nitrogen source are the main components of the production medium required for the growth of microorganism.

The choice of substrate is important for the enzyme production. Xylanases are usually inducible enzymes secreted in media containing

pure xylan or xylan-rich residues (Balakrishnan et al., 1997). However, constitutive production of xylanase has also been reported

(Khanna and Gauri, 1993; Khasin et al., 1993). 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 (Biely, 1985). It is believed that small amounts of constitutively produced enzymes liberate

xylo-oligomers which may be transported into the cell where they are degraded by β-xylosidases, or by intracellular xylanases and the

resulting products induce further xylanase synthesis. Various lignocellulosic materials, such as wheat bran, wheat straw, corn cobs,

sugarcane bagasse, cassava bagasse, barley bran and rice straw have been reported to act as good inducers of xylanase synthesis (Beg

et al., 2000a; Dhillon and Khanna, 2000; Sanghi et al., 2008; Parkash et al., 2009; Nagar et al., 2010b; Haddar et al., 2012; Sugumaran

et al., 2013). Agro-residues are the cheaper substitutes of pure xylan and could be exploited for xylanase production at industrial

scale. The level of enzyme production by various bacterial species/strains may be at variance with different agro-residues. Among the

various agro residues, wheat bran was reported to be the best substrate for inducing xylanase by Bacillus sp. in a number of research

papers (Sanghi et al., 2008; Nagar et al., 2010a&b; Mittal et al., 2013). Nitrogen source can be provided in either inorganic (KNO3,

ammonium chloride, ammonium dihydrogen phosphate etc) or organic form (peptone, yeast extract, beef extract etc.), the later being

more effective in stimulating xylanase production in most fermentation processes.

Agitation and aeration are generally used to meet the oxygen demand and uniform mixing of nutrients during fermentation

process. The lower enzyme production under stationary conditions might be due to reduction in oxygen level in the medium which in

turn adversely affected the enzyme yield. There are many reports related to the use of various metal ions, chelators, detergents and

surfactants for xylanase production. However, concentration of metal ion and mechanism of induction may vary from species to

species (Saxena et al., 1994).

In SSF, xylanase yield is affected by varying the moisture levels. Maximum enzyme yield is obtained at an optimum moisture

level. On increasing or decreasing the substrate to moisture ratio, xylanase yield declines. Hence, the moisture level should be

optimized, if enzyme is produced via SSF.

The above factors may be optimized for obtaining maximum enzyme yield by either one variable at a time approach or

statistical approach. Culture medium optimization by the traditional ‘one-factor-at-a-time’ technique requires a considerable amount

of work and time. Moreover, it is very difficult to determine the cumulative effect of more than two factors by using this approach.

However, optimization using statistical approach, such as Placket-Burman and Response surface methodology (RSM) can be used to

analyze the interactive effects of various factors and to optimize biotechnological processes (Bocchini et al., 2002; Nagar et al., 2011).

A brief account of the work done on xylanase production in SmF and SSF is given below:

2.7.1 Production of xylanase in SmF

Xylanase has been produced in SmF from different bacterial species/strains and the culture conditions employed by various

researchers are summarized in Table 2.1. Some of the reports pertaining to xylanase production in SmF are given below:

Ratto et al. (1992) reported the optimum xylanase activity of 400 IU/ml at neutral pH from B. circulans. It showed maximum

activity at pH 7.0 and 40% of activity was retained at pH 9.2. However, the culture supernatant also exhibited low levels of cellulose

activity. Balakrishnan et al. (2000) recorded 19.28 U/ml xylanase yield from B. circulans AB 16 when grown on rice straw medium.

Subramaniyan et al. (2001) observed 380 IU/ml xylanase activity when Bacillus SSP-34 was grown for a period of 96 h in the

production media containing (g/l) oat spelt xylan, K2HPO4, MgSO4 7H2O, yeast extract and peptone (pH 8.5). Fermentation was

carried out using 5% inoculum at 300 rpm at 35º ± 2ºC. Maximum enzyme yield was obtained in medium supplemented with 0.25%

each of yeast extract and peptone.

Sa-Pereira et al. (2002) isolated a xylanolytic B. subtilis from a hot-spring and the optimal production of cellulase free

xylanase (12 U/ml) was achieved at pH 6.0 and 50ºC, within 18h of fermentation. On increasing temperature to 55ºC, a higher

productivity was obtained in batch reactor (45000 U/l/h) as, compared to shake-flask fermentation (12000 U/l/h). The xylanase was

thermostable, presenting full stability at 60ºC during 3h.

Battan et al. (2007) produced xylanase from B. pumilus in SmF and obtained maximum enzyme yield in basal medium

supplemented with 2% wheat bran, peptone, yeast extract and potassium nitrate, pH 8.0 after incubation at 37ºC for 26 h. After

optimization, the enzyme yield increased by 13-fold. Sharma et al. (2007) reported that optimization of culture conditions resulted in

7.72-fold enhancement in cellulase-free and alkali-thermostable xylanase production in wheat bran-tryptone medium. The enzyme

yield was maximum (12.95 U/l) at pH 7.5, temperature 70ºC, agitation rate 250 rpm and incubation period of 72h. The enzyme

secretion was higher on supplementation of medium with 0.15% xylan and 0.1% Tween-80.

Kapoor et al. (2008) have optimized the culture conditions (pH 9.0, agitation 200 rpm, inoculum size 1.25% and inoculum age

2h) for maximum xylanase production by an alkaliphilic B. pumilus stain MK 001. Under these conditions, the bacterium secreted

1220 and 990 IU/ml on wheat bran and wheat straw, respectively. In contrast, the enzyme yield on birch wood xylan and oat spelt

xylan was 1190 and 1150 IU/ml, respectively. Prakash et al. (2009) observed maximum xylanase production from a halophilic and

alkali-tolerant Chromohalobacter sp. TPSV 101 on incubation at 40ºC for 140h in the medium containing 20% NaCl, pH 9.0

supplemented with 1% sugarcane bagasse and 0.5% feather hydrolyzate as carbon and nitrogen sources. Sugarcane bagasse (250

U/ml) and wheat bran (190 U/ml) were the best inducers of xylanase as compared to pure xylan (61 U/ml).

Sanghi et al. (2009) optimized the xylanase production from B. subtilis ASH in SmF using one variable approach. The use of

optimized conditions (pH 7.0, temperature 37ºC, 48 h incubation period, shaking at 200 rpm, 2% inoculum level, 2% wheat bran,

0.5% peptone and 0.5% yeast extract resulted in maximum xylanase production (410 IU/ml) which was 1.5-fold higher than under

unoptimized conditions.

Azeri et al. (2010) studied the effects of different factors such as pH (7.0-10.0), temperature (25-50°C) and inexpensive agro-

residues (wheat straw, wheat bran and corn cob) on xylanase production of four Bacillus stains under shake flask conditions.

Maximum enzyme activities (2.4-3.7 IU/ml) were obtained by cultivation in the medium containing birch wood xylan but high

enzyme production was also obtained on wheat straw and corn cob when cultivated at pH 8.5. The enzyme exhibited maximum

activity at pH 9.0 and temperature 60°C. The enzyme was stable at 60°C for more than 1h. Giridhar and Chandra (2010) isolated an

aerobic xylanolytic Gracilibacillus sp. TSCPVG capable of growing at extreme salinity (1–30%) and pH range 6.5–10.5. Maximum

xylanase (18.44 U/ml) and β-xylosidase (1.01 U/ml) activity was obtained after 60 h of incubation in the GSL-2 medium with

additions of (g/l) birchwood xylan (7.5), yeast extract (10.0), tryptone (8.0), proline (2.0), thiamine (2.0), Tween-40 (2.0) and NaCl

(35) at pH 7.5, 30 ºC and agitation rate 180 rpm.

Nagar et al. (2011) employed RSM to analyze the cumulative effect of peptone, yeast extract and KNO3 on xylanase

production by B. pumilus SV-85S. The use of optimal fermentation conditions i.e., 0.9% peptone, 0.9% yeast extract, 0.2% KNO3,

2.0% wheat bran, 0.1% KH2PO4, 0.01% MgSO4, 0.1% Tween 80, pH 6.0, temperature 37ºC, incubation time 36 h, agitation rate 150

rpm and 1.0% inoculums enhanced the enzyme production to 2995.20 IU/ml, which was 9.91-fold higher than that using basal

medium. Kamble and Jadhav (2011) studied the production of extracellular xylanase from an alkalo-thermophilic B. arseniciselenatis

DSM 15340 in SmF. The optimum xylanase production (749 U/ml) was found at pH 8.0 and temperature 65 ºC in basal salt yeast

extract medium supplemented with wheat bran after 48 h of incubation.

Nagar et al. (2012a) optimized the fermentation condition for hyper production of cellulase-free and alkali-stable xylanase

from B. pumilus SV-205. The bacterium secreted high levels of xylanase (7382.7 IU/ml) using wheat bran. The optimized condition

were found to be pH 10.0, incubation period 36h,ultivation temperature 37°C, agitation rate 150 rpm, wheat bran as carbon source,

and peptone and yeast extract as a nitrogen source. Haddar et al. (2012) optimized the xylanases production from B. mojavensis A21

using statistical methods: PB design was applied to find the key ingredients and Box–Behnken design was applied to optimize the

value of the four significant variables: barley bran, NaCl, agitation, and cultivation time. The optimal conditions (18.66 g/l barley

bran, 1.04 g/l NaCl, 176 rpm of agitation and 34.08 h cultivation time) resulted in 6.83-fold increase in xylanase production.

Mittal et al. (2013) optimized xylanase production from Bacillus sp. SV-34S by using the one variable approach in submerged

fermentation conditions. Cultivation using wheat bran as the carbon source and beef extract and (NH4) H2PO4 as the nitrogen source

resulted in highest xylanase titer. The optimal conditions for maximum xylanase production were: an incubation time of 48 h, pH 7.0,

temperature of 37°C, 1% inoculum of an 18-h-old culture, shaking at 200 rpm, 2% wheat bran, and 0.5% each of beef extract and

ammonium dihydrogen phosphate. A comparison of xylanase production measured under these optimized fermentation conditions

(3,454 IU/m) with that under non-optimal conditions (394.74 IU/ml) revealed an 8.74-fold increase in activity. Sugumaran et al.

(2013) studied the production of alkaline tolerant and thermostable xylanase by Bacillus subtilis in submerged fermentation using

various agricultural wastes such as wheat bran, wheat flour and cassava bagasse as a carbon source. Enzyme production and activity

was maximum using cassava bagasse as a carbon source and yeast extract as the nitrogen source.

Table 2.1 Xylanases produced by bacteria in submerged fermentation (SmF)

Bacterium Carbon

source

Nitrogen source Incubation

period

Temp pH Xylanase

activity (IU/ml)

Reference

B. cereus BSA-1 Xylan Yeast extract,

(NH4)SO4

84 35 6.0 6.02 Mandal et al., 2012

B. circulans Xylan Peptone, Yeast

extract

48 30 9.0 400 Ratto et al., 1992

Bacillus SPS-0 Wheat bran Yeast extract,

Tryptone

24 60°C 7.0 53nkat/ml Bataillon et al., 1998

B. circulans AB-1 Wheat straw Tryptone 96 72 6.0 50 Dhillon and Khanna,

2000

B. circulans D1 Xylan Beef extract,

Peptone

48 45 9.0 19 Bocchini et al., 2002

B. circulans Teri 42 Oat spelt xylan 24 37 1.10 Qureshy et al., 2002

B. licheniformis Wheat bran Corn steep liquor 50 7.0 756 Archana and

Satyanarayana, 1998

B. mojavensis A-21 Barley bran NH4Cl 34.08 30 8.0 7.5 Haddar et al., 2012

B. mojavensis

AG137

Oat bran Peptone, Yeast

extract

48 37 8.0 302.46 Sepahy et al., 2011

B. pumilus Oat spelt xylan

Wheat bran

Peptone, Yeast

extract

96

72

35 9.0 580

430

Poorna and Prema,

2006

B. pumilus ASH Wheat bran

Peptone, Yeast

extract, KNO3

26 37 8.0 5407 Battan et al., 2007

B. pumilus B20 Wheat bran,

Peptone, Yeast

extract

36 37 7.5 313 Geetha and

Gunasekaran, 2010

B. pumilus MK001 Wheat bran Peptone, Yeast

extract

48 37 9.0 1220 Kapoor et al., 2008

B. pumilus SV-85S Wheat bran Peptone, Yeast

extract, KNO3

48 37 6.0 2900 Nagar et al., 2010b

B. pumilus SV 34S Wheat bran Beef extract,

(NH4)2H2PO4

48 37 7.0 3454 Mittal et al., 2013

Bacillus sp. Wheat bran Peptone 48-60 50-55 8.0 4.0 Sharma et al., 2011

Bacillus sp. AG20 Wheat bran - 42-48 35 8.5 2.4-3.7 Azeri et al., 2010

Bacillus sp. SSP-34 Oat spelt xylan Yeast extract and

peptone

96 35 8.5 380 Subramaniyan et al.,

2001

Bacillus sp. V1-4 Birchwood

xylan

Corn steep liquid 48 37 10 49 Yang et al., 1995

B. subtilis Oat spelt xylan 18 50 6.0 12 Sa-Pereira et al.,

2002

B. subtilis Oat spelt xylan Peptone, Yeast

extract

36 55 9.0 128 Annamalai et al.,

2009

B. subtilis ASH Wheat bran Peptone, Yeast

extract, KNO3

48 37 7.0 410 Sanghi et al., 2009

B. subtilis 276NS Xylan Yeast extract 24 37 8.0 360 Ali et al., 2013

Chromohalobacter

sp. TPSV 101

Sugarcane

bagasse

Feather

hydrolyzate

140 40 9.0 250 Prakash et al., 2009

Flavobacterium sp. Larchwood

xylan

Yeast extract,

Peptone, Casein

hydrolyzate and

Nitrates

30 6.8 6.64 Bhatt et al., 1994

Geobacillus

thermoleovorans

Wheat bran Tryptone 72 70 7.5 12.95 Sharma et al., 2007

Gracilibacillus sp.

TSCPVG

Birchwood

xylan

Yeast extract and

Tryptone

60 30 7.5 18.44 Giridhar and

Chandra, 2010

Paenibacillus sp.

N1

Xylose (NH4)2HPO4 72 50 9.0 52.30 Pathania et al., 2012

2.7.2 Production of xylanase in SSF

Xylanase production in SSF has been reported from various bacterial species/strains as summarized in Table 2.1. Some of the

reports pertaining to xylanase production in SSF are given below:

Beg et al. (2000) reported the production of a thermostable and cellulase-free xylanase from Streptomyces sp. QG-11-3 in SSF

using wheat bran and eucalyptus kraft pulp as solid substrates. The substrates were uniformly mixed with mineral salts solution

containing (g/l): KH2PO4, 1; NaCl, 2.5; MgSO4.7H2O, 0.1; (NH4)2SO4, 1; CaCl2, 0.1; and soil extract, 2 ml (v/v) at pH 8.0. The

maximum xylanase yield obtained using these two substrates were 2360 U/g and 1200 U/g dry solid substrate at substrate: moisture

ratio of 1:3 and 1:2.5, respectively.

Sindhu et al. (2006) showed that optimization of SSF process parameters resulted in the highest xylanase activity at the

substrate to moistening agent ratio of 1: 1.5. Addition of 5% soya bean meal increased the xylanase production by 1.4-fold. An

inoculum level of 10% resulted in maximum enzyme yield after an incubation period of 96 h. Scale up of xylanase production by B.

megaterium in SSF to 500 g substrate level in enamel trays resulted in 846 U/g xylanase yield which was 12.14% lower than the

enzyme yield at 5 g level in shake flasks.

Poorna and Prema (2007) optimized the culture condition for the enhanced production of cellulase-free xylanase from Bacillus

pumilus in SSF. Batch studies were carried out to evaluate various agro-industrial residues such as rice bran, rice husk, rice straw,

sawdust, coconut pith, sugarcane bagasse and wheat bran for enzyme production. The enzyme production was highest on wheat bran

media (5582 U/gds), which was further enhanced by 3.8-fold through optimization of cultivation conditions.

Khandeparker and Bhosle (2007) optimized the xylanase production from Arthrobacter sp. MTCC 5214 in SSF using wheat

bran as a carbon source. Bacterial cultures were isolated by utilizing xylan from sediment sample collected from Mandovi estuary,

west coast of India. Among these isolates, Arthrobacter sp. MTCC 5112 produced the highest xylanase at 30 °C using wheat bran as a

substrate and moisture ratio 1:3 after incubation for 7 days.

Sanghi et al. (2008) optimized the xylanase production in SSF from an alkalophilic B. subtilis ASH using inexpensive

agricultural residues. Among these agro-residues, wheat bran was found to be the best substrate. Xylanase production was highest

(8,964 U/g) after 72 h of incubation at 37 ºC and a substrate to moisture ratio of 1:2 (w/v). An inoculum level of 15% resulted in

maximum production of xylanase. Addition of nutrients such as yeast extract, peptone and beef extract stimulated the enzyme

production. In contrast, addition of glucose and xylose repressed the production of xylanase.

Geetha and Gunasekaran (2010) studied the xylanase production from B. pumilus B20 by using DeMeo’s fractional factorial

design. Xylanase production increased up to 3.4-fold under the optimized culture medium consisting of K2HPO4, NaCl, peptone, yeast

extract and wheat bran. Among the different factors screened, wheat bran showed a positive effect in the first step of optimization and

MgSO4·7H2O and CaCl2·2H2O had a negative effect.

Kapilan and Arasaratnam (2011) optimized the SSF conditions for xylanase production from B. pumilus, using paddy husk

moistened with liquid fermentation medium (xylan, 20.0 g/L; peptone, 2.0 g/L; yeast extract, 2.5 g/L; K2HPO4, 2.5 g/L; KH2PO4, 1.0

g/L; NaCl, 0.1 g/L; (NH4)2SO4, 2.0 g/L, CaCl2·2H2O, 0.005 g/L; MgCl2·6H2O, 0.005 g/L; and FeCl3, 0.005 g/L) at pH 9.0. The

highest xylanase activity was obtained after six days of incubation at 30 °C using the paddy husk to liquid fermentation medium ratio

of 2: 9. Production of the xylanase was increased by sucrose, fructose, and arabinose but reduced by glucose, galactose, and lactose.

Nagar et al. (2011) optimized the xylanase production from B. pumilus SV-85S in SSF using wheat bran as substrate. The

optimization of fermentation conditions enhanced the enzyme production from 5300 IU/g to 73,000 IU/g. The enzyme titre was

highest after 48 h of incubation at 30 ºC with 1:3 ratio of substrate to moistening agent using the inoculum level of 15% and wheat

bran as a carbon source.

Bajaj et al. (2012) isolated a xylanolytic B. pumilus SS1 stain from soil and successfully produced considerable titer of

xylanase utilizing the wheat bran as the sole carbon source. Maximum enzyme production occurred at medium pH 8.0 and

temperature 45 ºC after an incubation period of 48 h. Yeast extract caused a substantial increase in xylanase production (14.2%) while

peptone did so moderately (7.1%) in comparison to the control.

Banu and Ingule (2012) optimized the production of xylanase from B. pumilus AB-1 under SSF. Maximum production of

xylanase was observed when bran moistened with mineral salt solution (MA III; pH 7.0) at a substrate to moistening agent ratio of

1:1.5 (w/v) was incubated at 30°C for 72 h. Yeast extract at 2.5% (w/v) concentration was found to enhance the xylanase production.

Kamble and Jadhav (2012b) optimized xylanase production from Cellulosimicrobium sp. MTCC 10645 in SSF. The optimized

fermentation conditions for growth and xylanase production were 72 h of incubation period at pH 7.0, cultivation temperature 40°C

inoculum size of 10% and substrate to moisture ratio of 1:1.8 (w/v). Among the different lignocellulosic substrates, wheat bran was

found to be best substrate. Xylanase production was stimulated by the addition of yeast extract and casein whereas addition of glucose,

xylose and carboxy methyl cellulose repressed the production of xylanase. Under optimized conditions, xylanase production in solid

state fermentation was 4,962 + 45.08 U/gds.

Walia et al. (2013) isolated Cellulosimicrobium sp. CKMX1 from mushroom compost and optimized the xylanase production

at 35 °C and pH 8.0 using the inexpensive apple pomace as the carbon source under SSF. The enzyme titre was increased to 535.6 U/g

after 72 h of incubation using 10% inoculum with wheat bran as a carbon source. Further, the optimization of enzyme production was

carried out using central composite design following RSM with four independent variables (yeast extract, urea, Tween 20 and CMC)

which resulted in very high levels of xylanase.

Table 2.2 Production of xylanases from bacteria in solid state fermentation (SSF)

Bacterial strain Solid

substrate

Nitrogen

source

Incubation

period

Temp pH Xylanase

Activity (IU/g)

References

Arthrobacter sp.

MTCC 6915

Saw dust Peptone and

beef extract

96 30°C 176.4 Murugan et al., 2011

Cellulosimicrobium

sp. CKMX1

Apple pomace Yeast extract 72 35°C 8.0 861.90 Walia et al., 2013

Cellulosimicrobium

sp. MTCC 10645

Wheat bran Yeast extract

and casein

72 40°C 7.0 4,962 Kamble and Jadhav, 2012b

B. licheniformis A99

Wheat bran Peptone 72 50°C

7.0 19.13 Archana and Satyanarayana,

1997 B. megaterium Wheat bran Soya bean meal 96 37°C 8.0 7,700 Sindhu et al., 2006

B. pumilus Paddy husk Soy meal

powder

144 30°C 9.0 142 Kapilan and Arasaratnam,

2011 B. pumilus ASH Wheat bran - 72 37 °C 8.0 5,407 Battan et al., 2007

B. pumilus SV-85 Wheat bran - 48 30°C 73,000

Nagar et al., 2011

Bacillus sp. AR-009

Wheat bran Yeast extract 72 37°C - 720 Gessesse and Mamo, 1999

Bacillus sp. GRE7 Wheat bran - 55 °C 7.0 3,950 Kiddinamoorthy et al., 2008

Bacillus sp. JB-99

Rice bran Yeast extract,

beef extract

72 50°C 10.0 3,644 Virupakshi et al., 2005

B. stearothermophilus

SDX

Wheat bran - 8.0 3,446 Dhiman et al., 2008a

B. subtilis ASH Wheat bran Yeast extract 72 37°C 7.0 8,964 Sanghi et al., 2008

2.8 Scale up of xylanase production

There are only a few reports pertaining to scale up of xylanase production from bacteria in SmF and SSF. These are as follows:

Sa-Pereira et al. (2002) investigated the production of cellulose- free xylanase from B. subtilis in shake-flasks and 2L batch

reactor capacity (working volume 1.5L) at pH 6.0 and 55ºC, within 18h of SmF and recorded a higher productivity (45000 U/l/h) as

compared to shake-flask fermentation (12000 U/l/h).

Gupta and Kar (2008) studied the scale up of xylanase production from B. licheniformis MTCC 9415 in SSF. Xylanase

production was higher in trays using bulk quantities of corn cob (80 g), as compared with Erlenmeyer flasks. Cultivation in large

enamel trays yielded 157.12 ±8.7 U/gds as compared to the yield 74.96 ±5.2 U/gds obtained in 250 ml flasks.

Kamble and Jadhav (2012b) carried out the scale up of xylanase production by Cellulosimicrobium sp. MTCC 10645 in SSF

by using enamel trays containing 80 g wheat bran. Cultivation in large enamel trays yielded 4,962 U/gds as compared to 965 U/gds

obtained in 250 ml flasks.

2.9 Purification and characterization of endoxylanase

Purification of xylanases to homogeneity is necessary for detailed biochemical and molecular studies, and for the successful

determination of their primary amino acid sequences and their three-dimensional structures. The isolation and purification methods

used for the recovery of microbial xylanases have been reviewed by Sa-Pereira et al. (2003). Owing to the presence of less number of

contaminating proteins in the culture filtrate, purification of an extracellular enzyme is easier than the intracellular one. Protein

purification varies from a simple one-step purification procedure to large scale purification processes. The key to obtaining successful

and efficient purification of an enzyme protein is the selection of appropriate techniques that maximize its yield and purity but

minimize the number of steps needed for its purification. Molecular cloning is being increasingly used to overproduce enzymes

including xylanases (Sa-Pereira et al., 2003; Juturu and Wu, 2011). The use of several purification techniques such as ammonium

sulfate precipitation, ion exchange chromatography, gel filtration and affinity chromatography has been reported for the purification of

xylanase from different microorganisms (Sa-Pereira et al., 2003). Table 2.3 summarizes the characteristics of purified bacterial

xylanases reported in various research papers. Some of the reports on purification of bacterial xylanase are as follows:

Table 2.3 Characteristics of purified bacterial xylanases

Organism MW pH Temp Km Vmax Fold

purificati

on

Yiel

d

(%)

Reference

Optimu

m

stabilit

y

Optimu

m

stability

Alicyclobacillus sp.

A4

oat spelt xylan

birchwood xylan

42.5 7.0 - 55 60

1.90c

1.56c

417.93d

335.8d

- - Bai et al., 2010

B. amyloliquefaciens 18.4 6.8-7.0 - 80 - - - 7.3 53.9 Breccia et al., 1998a

B. circulans AB 16 30

22

6.0-6.5

6.0-6.5

- 75-80

65-70

- 4.0c

25 c

2666.6f

20x102(

f)

38.5

125

29

10.6

Dhillon et al., 2000b

B. circulans 85

15

5.5-7.0

5.5-7.0

- - 4.5

9.1

- - - - Esteban et al., 1982

B. circulans Teri -42 71 7.0 - 50 - 2.86c 0.13d 2 40 Qureshy et al., 2002

B. halodurans 36 9.0 - 80 - - - 10.4 27.3 Kumar and

Satyanarayana, 2011

B. halodurans TSVP1 10.0 7-12 90 - - - - 26 Kumar et al., 2013b

B. halodurans TSEV-

1

40 9.0 7.0 -

12.0

80 40-90 2.05c 333.33d 13.5 30 Kumar and

Satyanaryana, 2013

B. licheniforms A99 14.4 38 Archana and

Satyanaryana, 2003

B. polymoxa CECT

153

61 6.5 - 50 - 17.7c 112d 30 0.5 Morales et al., 1995

B. pumilus 19 7.0 - 40 - 4.0c 0.068x

10-4 (J)

3.79 66 Monisha et al., 2009

B. pumilus GESF-1 39.6 8.0 7.0-8.0 40 - 5.3c 6593.4d 21.21 2.1 Menon et al., 2010

B. pumilus SS1 25 6–8 - 40–50 50 2.7c 36d 2.97 11.9

3

Bajaj et al., 2012

B. pumilus SSP-34 20 6.0 4.5-9.0 50 - 6.5 c 1233d 33.3 2.5 Subramaniyan, 2012

B. pumilus SV-85S 23.6 7.0 - 50 - 1.0c 333.3d 25.3 63.2 Nagar et al., 2012b

B. pumilus 276NS

xyl-1

xyl-2

xyl-3

14

35

60

7.0

6.0

6.0

-

50

50

55

20-40

4.0c

3.5 c

5000d

3448d

1.06

2.68

3.1

1.46

Asha Poorna, 2011

Bacillus sp. 23 8.0 - 70 - 11.8c 20.9d 13.5 13.6 Gessesse, 1998

48 9.0 75 2.6c 0.9d 9.5 0.6

Bacillus sp. 44 6.5 6.0-

10.0

50 - 0.02

5c

450m 54.87 56.2

7

Sapre et al., 2005

Bacillus sp. 48 7.0 5.4-

10.6

70 - 2.53

c

0.6j - - Zhang et al., 2010

Bacillus sp. AR 009 23

48

9.0

9.0-10

70

75

- 13.6

0.6

Gessesse and Mamo,

1999

B. arseniciselenatis

strain DSM-15340

29.8 8.0 6-7 50 - 5.26c 277.7d 3.06 41.5

9

Kamble and Jadhav,

2012a

Cellulosimicrobium

sp. MTCC 10645

78 7.0 7.0 50 30-40 4.76c 232.5d 16.2 2.71 Kamble and Jadhav,

2012b

Bacillus sp. AQ-1 15.7 7.0 - 60 - - - .68 7.11 Rahayu et al., 2008

Bacillus sp. BP-23. 32 5.5 9.5-10 50 55 - - 5 20.3 Blanco et al., 1995

Bacillus sp. BP-7 24 6.0 60 50/3h - - - Gallardo et al., 2004

Bacillus sp. C-125 6.0-10

6.0-7.0

70

70

- - - 21

31

3.7

26.5

Honda et al., 1985

Bacillus subtilis CCMI

966

140 6.0 - 60 - - - 2.7 3 Sa Pererira et al.,

2000

Bacillus sp. GRE 7 42 7.0 n.a. 70 n.a. 2.23c 296.8d 28.5 27 Kiddinamoorthy et

al., 2008

Bacillus sp. JB99 20 8.0 - 70 - 4.8c 218.6d 25.7 43.5 Shrinivas et al., 2010

Bacillus sp. K-8 24 7.0 n.a. n.a. n.a. n.a. n.a. 5.3 42.3 Tachaapaikoon et

al., 2006

Bacillus sp. MX47 26.4 8.0 40 3.24c 58.21d 36.7 16.1 Chi et al., 2012

Bacillus sp. NCIM 59 35

15.8

6.0 - 50-60 - - - 8

20

0.12 Dey et al., 1992

Bacillus sp. NTU-06 24 8.0 - 40 - 3.45c 387.3d 36.7 16.1 Wang et al., 2010

Bacillus sp SV 34s 27 6.5 - 50 - 3.7c 133.33d 12.94 13.4 Mittal et al., 2013

Bacillus sp. stain K-1 23

45

5.5 -. 60 30 - - - - Ratankhanokchchai

et al., 1999

Bacillus sp. SPS-0 99 6.0 - 75 -. 0.7c - 36 24.6 Bataillon et al., 2000

Bacillus sp. TAR-1 40 7.0 - 75 - 0.82c 280d 31.5 Nakamura et al.,

1995

Bacillus sp. YJ6 19 5.0 5.0-9.0 50 - - - 678 3.5 Yin et al., 2010

Bacillus sp. 41-M1 36 9.0 n.a. 50 n.a. 3.3c 1,100d 3.6 15.3 Nakamura et al.,

1993

B.

stearothermophilusT6

43 60 6.5-10 75 70 1.63c 288l 38.9 46 Khasin et al., 1993

B. subtilis - 8.0 -. 60 - 1.9g 0.1h - -. Sugumaran et al.,

2013

B. subtilis 36 9.0 - 55 - - - 1.69 34 Annamalai et al.,

2010

B. subtilis ASH 23 7.0 6.0-9.0 55 - 3.33c 100f 10.5 43 Sanghi et al., 2010

B. subtilis XP-10 23 8.5 - 40 - - -. - -. Tork et al., 2013

B. thermantarcticus 45 5.6 n.a. 80 n.a. 1.6c n.a. 119 19 Lama et al., 2004

G. thermodenitrificans 50 9.0 - 70 - 0.62c 555.5d - - Verma et al., 2013

G. thermodenitrificans

TSAAI

43 7.5 5.0-8.0 70 60 2.85c 45.45d 25 9 Anand et al., 2013

G. thermodenitrificans

JK11

47.4 6.0 - 70 - - - - - Gerasimova and

Kuisiene, 2012

Paenbaciillus sp.

HPL-002

38.4 8.0-9.0 - 50 40 0.06 55.3 4.7 - Park et al., 2012

Paenibacillus

macquariensis RC

1819

31 8.6 - 50 - 2.2c - - - Sharma et al., 2013

Staphylococcus spp.

SG-13

60 7.5

9.2

50 50 4.0c 90d 12 5 Gupta et al., 2000

c = mg/ml, d = µmol/min/mg, e = mol/mg, f = IU/ml, g = g/L, h = g/L/min, j = mmol/min/mg, k= mmol/mg U/ml=m

Gessesse (1998) purified two xylanases (Xyl-A and Xyl-B) from the alkaliphilic Bacillus sp. strain AR-009 using the DEAE-

Sepharose column chromatography. Their molecular mass by SDS-PAGE was found to be 23 and 48 kDa, respectively.

Ratanakhanokchai et al. (1999) purified an extracellular xylanases to homogeneity by affinity adsorption/desorption on insoluble

xylan from Bacillus sp. strain K-1. The molecular mass of purified xylanase was approximately 23 kDa. Metal ions such as Fe+2, Ca+2,

and Mg+2 increased the xylanase activity, whereas Mn+2 strongly inhibited it. Archana and Satyanarayana (2003) purified a xylanase

from B. licheniforms A99 by 14.4- fold with 38% yield using ammonium sulfate precipitation and ion-exchange chromatography

through DEAE-Sephadex A-50.

Wang et al. (2009) purified a xylanase by fast protein liquid chromatography (FPLC) and had a molecular mass of 24 kDa. The

enzyme was active over a concentration range of 0–20% sodium chloride in culture broth, although its activity was optimal in 5%

sodium chloride. Xylanase activity was maximal at pH 8.0 and 40°C. The xylanase had Km 3.45 mg/ml and Vmax 387 μmol/min/mg.

The deduced internal amino acid sequence of Bacillus sp. NTU-06 xylanase resembled the sequence of β-1,4-endoxylanase, which is a

member of glycoside hydrolase family 11. Monisa et al. (2009) partially purified a xylanase from B. pumilus by 3.79-fold with 66%

recovery using ammonium sulfate precipitation. Partially purified enzyme preparation exhibited a specific activity of 0.69

μM/min/mg, Km 4.0 mg/ml, Vmax 0.068 × 10-4 mM/min/mg and molecular mass 19 kDa.

Sanghi et al. (2010) purified an extracellular cellulase-free xylanase to homogeneity from B. subtilis ASH by 10.5-fold with

43% recovery in a single step using CM-Sephadex C-50 column chromatography. It showed an optimum pH at 7.0 and was stable

over the pH range 6.0-9.0. The optimum temperature of the enzyme was 55 °C. The purified enzyme revealed a single band on SDS-

PAGE gel with a molecular mass of 23 kDa. The Km and Vmax of the enzyme for birch wood xylan were found to be 3.33 mg/ml and

100 IU/ml, respectively. The purified enzyme was stable for six weeks at 4°C and lost 20% activity after 10 weeks. The enzyme

activity was strongly inhibited by Hg2+ and Cu2+ but stimulated by Co2+ and Mn2+.

Shrinivas et al. (2010) purified a highly thermostable xylanase from Bacillus sp. JB 99 to 25.7-fold and 43.5% recovery using

two step purification strategy involving chromatography through DEAE-Sepharose and Sephadex G-100. The purified enzyme

exhibited a molecular mass of 20 kDa, Km 4.8 mg/ml and Vmax 218.6 μM/min/mg for oat spelt xylan.

Menon et al. (2010) used (NH4)2SO4 fractionation, DEAE-Cellulose, and Sephadex-G-200 chromatography to purify xylanase

from B. pumilus strain, GESF1 resulting in 21.21-fold purification with a specific activity 112.42 U/mg protein, Km 5.3 mg/ml and

Vmax 0.42 μmol/min/ml. Prakash et al. (2009) reported the partial purification of a xylanase from Chromohalobacter sp. TPSV 101

using the protein concentrator. The partially purified enzyme had a molecular mass 15 kDa, optimum temperature 65°C and pH

optimum at 9.0.

Yin et al. (2010) reported the purification of xylanase from Bacillus sp. YJ6 using Sephacryl S-100 HR chromatography with

3.5% recovery and 678.1-fold purification. Xylanase had an optimal pH at 5.0 but was stable over the pH range 5.0-9.0. The enzyme

showed an optimum temperature of 50 ºC and was stable at temperatures less than 50 ºC. Xylanase activity was inhibited by Cu+2,

Fe+3, Hg+2, phenylmethyl sulfonyl fluoride (PMSF), N-tosyl-L-phenylalanine chloromethyl ketone (TPCK), N-ethylmaleimide, and

leupeptin, but activated by K+, Na+, Co+2, Mg+2, β-mercaptoethanol, and glutathione. The purified xylanase had high specificity for

beechwood, birchwood, and oat spelt xylans. The DNA fragment encoding this xylanase, corresponding to 213 amino acids, exhibited

about 95% homology with seven strains of Bacillus in the NCBI database. The purified enzyme had a molecular mass of 19 kDa and

specific activity 1436.0 U/mg protein.

Bai et al. (2010) cloned the gene (xynA4) encoding xylanase from Alicyclobacillus sp. A4 and expressed it in E. coli. It

encoded a 338-amino acid residue polypeptide with a calculated molecular mass of 42.5 kDa. Amino acid sequence was similar to

(53% identity) an endoxylanase from Geobacillus stearothermophilus belonging to family 10 of the glycoside hydrolases.

Recombinant XynA4 exhibited maximum activity at 55°C and pH 7.0, had broad pH stability (retaining 80% activity after incubation

at pH 2.6–12.0 for 1 h at 37°C), and was highly thermostable retaining 90% activity after incubation at 60°C for 1 h.

Zhang et al. (2010a) cloned the xylanase gene (xyn10) from alkaliphilic Bacillus sp. N16-5 and expressed it in Pichia pastoris.

The deduced amino acid sequence had 85% identity with xylanase xyn10A from B. halodurans and contained two potential N-

glycosylation sites. The glycosylated Xyn10 with MW 48 kDa could hydrolyze birchwood and oatspelt xylans. The enzyme had

optimum activity at pH 7.0 and 70°C, with the specific activity of 92.5U/mg. The Xyn10 retained over 90% residual activity at 60°C

for 30 min but lost all activity at 80°C over 15 min.

Chi et al. (2012) purified an extracellular xylanase from the culture broth of Bacillus sp. MX47 using two chromatographic

steps. The xylanase had an apparent molecular mass of 26.4 kDa with an amino terminal sequence (Gln-Gly-Gly-Asn-Phe) distinct

from that of reported proteins, implying that it was a novel enzyme. The optimum pH and temperature for xylanase activity were 8.0

and 40 °C, respectively. The enzyme activity was severely inhibited by many divalent metal ions and EDTA at 5 mM. The xylanase

was highly specific for beechwood and oat spelt xylans, however, not active on carboxymethyl cellulose (CMC), avicel, pectin, and

starch. Analysis of the xylan hydrolysis products by Bacillus sp. MX47 xylanase indicated that it was an endo-β-1,4-xylanase. The Km

and Vmax values toward beechwood xylan were 3.24 mg/ml and 58.21 μmol/min/mg protein, respectively.

Nagar et al. (2012a) purified an extracellular xylanase from B. pumilus 85S to apparent homogeneity by 25.3-fold with 63.2%

recovery using (NH4)2SO4 fractionation and CM-Sephadex column chromatography. The enzyme purity was tested by polyacrylamide

gel electrophoresis and HPLC. The purified enzyme revealed a molecular mass of 23.6 kDa estimated by SDS-PAGE. The Km and

Vmax values of the purified xylanase were 1.0 mg/mL and 333.3 IU/mL, respectively.

Mittal et al. (2013) reported the purification of Bacillus sp. SV-34S xylanase by 12.94-fold with a recovery of 13.4% and a

specific activity of 3417.2 IU/mg protein employing (NH4)2SO4 fractionation followed CM-Sephadex C-50 column chromatography.

The purified enzyme showed a molecular mass of 27 kDa, optimum temperature 50°C, optimum pH at 6.5, Km 3.7 mg/mL, and Vmax

133.33 IU/mL with birchwood xylan as the substrate.

Sharma et al. (2013) purified a xylanase secreted from Paenibacillus macquariensis RC 1819 using (NH4)2SO4 fractionation,

ion exchange chromatography using DEAE-cellulose, and gel filtration chromatography over Sephadex G-200 and Sephadex G-100.

The purified enzyme had a specific activity of 25.2 units/mg protein, molecular mass 31kDa, optimum pH at 8.6, optimum

temperature 50°C, and Km 2.2 mg/ml for birchwood xylan. Metal ions such as Co+2 and Mn+2 stimulated whereas Hg+2 inhibited the

enzyme activity.

Verma et al. (2013) cloned the xylanase encoding gene (1,224 bp) from Geobacillus thermodenitrificans in pET28a (+) vector

and successfully expressed in E. coli BL21 (DE3). Xylanase was purified by Ni2+-affinity chromatography. The eluted protein

appeared as a single band on 15% SDS-PAGE. The deduced amino acid sequence analysis revealed homology with that of glycosyl

hydrolase family 10 with a high molecular mass (50 kDa). The purified recombinant xylanase was optimally active at pH 9.0 and 70

°C with half-life of 10 min at 80 °C, and retained greater than 85% activity after exposure to 70°C for 180 min. The xylanase was

quite stable in the presence of the detergents tested. The recombinant xylanase showed Km 0.625 mgml−1 and Vmax 555.5

μmol/mg/min for birchwood xylan.

2.10 Immobilization of xylanase

Immobilization means restricting the mobility of biocatalysts by attaching them to an insoluble matrix, by cross-linking or

through encapsulation. The use of immobilized enzymes for catalyzing various biotransformations is now a widely used approach. For

practical applications, immobilization of microbial enzymes on solid materials may offer several advantages, including reusability of

enzymes, recovery of product with greater purity, easy separation of the enzyme catalyst from the reaction mixture, often

improvement of its chemical and thermal stability, and continuous operation of a process (Datta et al., 2013). Overall performance of

the immobilized enzyme preparations is based on immobilization yields, low mass transfer limitations, high operational stability and

reusability. The enhanced stability after immobilization may allow the application of immobilized enzymes under harsh conditions of

pH, temperature and non-aqueous media.

There are many methods available for immobilization which span from binding on prefabricated carrier materials to

incorporation into in situ prepared carriers. These methods include physical adsorption, ionic binding, covalent coupling, entrapment

and encapsulation (Datta et al., 2013). Each method has its own advantages and limitations. Immobilization of xylanase by entrapment

is not desirable as its substrate is large and hence its accessibility to the enzyme would be negligible. Covalent method involving the

formation of a covalent bond between the functional groups of amino acid residues on the surface of the enzyme and the functional

groups present on the surface of the carrier is likely to be more appropriate for immobilization of xylanase. The covalently

immobilized enzyme preparations are stable and enzyme leaching is minimal. Further, since enzyme molecules are on the surface,

contact with large substrates is possible as required in case of xylanase. A novel and versatile method for enzyme immobilization is

the production of cross-linked enzyme aggregates by extensive chemical cross-linking of enzymes dissolved in a solution. Such cross-

linked enzyme aggregates are stable and reusable (Sheldon et al., 2005; Dalal et al., 2007)

Immobilization of xylanase has been reported on various supports including polymethyl methacrylate nanofibers membrane

(Kumar et al., 2013a), Eudragit L-100 (Roy et al., 2003), silica (Kang et al., 2002; Sharma et al., 2012), chitin, HP-20 and gelatin

(Kapoor and Kuhad, 2007), glass beads (Kumar et al., 2009), chitosan beads (Jingmin et al., 2002), alginate beads (Pal and Khanum,

2011; Bhushan et al., 2013) and Eudragit S-100 (Gawande and Kamat, 1998, Edward, 2002). A multipurpose immobilized biocatalyst

with pectinase, cellulose and xylanase activities has been produced by cross-linking (Dalal et al., 2007). Some of the reports on

xylanase immobilization are as follows:

Dalal et al. (2007) designed a single multipurpose catalyst having pectinase, cellulose and xylanase activities. The preparation

was more stable at higher temperatures. The Vmax/Km values increased from 11, 75 and 16 to 14, 80 and 90 in case of pectinase,

xylanase and cellulose activities, respectively. Half life were improved from 17, 22 ad 32 minutes to 180, 82 and 91 minutes for

pectinase, xylanase and cellulose, respectively. All three of the enzyme in cross linked enzyme aggregate could be reused three times

without any lose of activity.

Kapoor and Kuhad (2007) immobilized B. pumilus strain MK001 xylanase on different matrices viz. gelatin (entrapment

method), chitin (physical adsorption), Q-sepharose (ionic binding), and HP-20 beads (covalent binding). Xylanase immobilized on

HP-20 beads showed the maximum efficiency and a shift of pH optimum by 1.0 unit. The immobilized xylanase exhibited 28.0%

higher pH stability as compared to free enzyme. Xylanase immobilized on HP-20, Q-S, CH, and GE retained 68.0, 64.0, 58.0, and

57.0% activity, respectively whereas free xylanase retained 50.0% activity after incubation for 3 h at 80.0 °C. The immobilized

xylanase retained up to 70.0% of its initial activity after seven cycles.

Anny et al. (2010) immobilized commercial xylanase NS50014 from Novozymes on glyoxyl-agarose, agarose-glutaraldehyde,

agarose-amino-epoxy support and on differently activated chitosan supports. Epoxy-chitosan-xylanase was found to be the best

chitosan derivative and presented 100% of immobilization yield and 64% of recovered activity. No significant increase on the thermal

stability was observed for all the chitosan-enzyme derivatives. The protein was then chemically modified with ethylenediamine and

immobilized on glyoxyl-agarose. The new enzyme derivatives were 40-fold more stable than the soluble, aminated, and dialyzed

enzyme (70°C, pH 7), with 100% of immobilization yield. Therefore, the increase of the number of amine groups in the enzyme

surface was confirmed to be a good strategy to improve the properties of immobilized xylanase.

Pal and Khanum (2011) reported the covalently immobilization of A. niger DFR-5xylanase on the glutaraldehyde- activated

alginate beads and the immobilized process was optimized by RSM. The assay of the immobilized enzyme was carried out at 37°C.

An increase in Km (from 0.9 to 1.49%), Vmax (from 7092 to 8000 IU/ml), optimum pH (from 5 to 5.5) and temperature (from 40 to 45

◦C) was realistic after immobilization. An improvement in thermostability of immobilized xylanase was also observed. Immobilized

xylanase could be reused 5 times while retaining more than 85% of its original activity.

Nagar et al. (2012d) covalent immobilized the xylanase from B. pumilus on the glutaraldehyde-activated aluminum oxide

pellets. Process parameters of immobilization were optimization by using the response surface methodology results in 83.65% yield.

The immobilized enzyme has shown an increase in temperature optima from 50 to 60 °C and Vmax from 3333.33 to 5000 IU/ml as

comparison with free enzyme. Similarly the pH and temperature stability of the immobilization xylanase were also enhanced.

Immobilized xylanase has retained 60% of its initial activity after 10 consecutive cycles.

Dhiman et al. (2012) covalently immobilized the Armillaria gemina xylanase onto functionalized SiO2 nanoparticles and

achieved 117 % immobilization efficiency. Immobilization caused a shift in both the pH optima and temperature, along with a 4-fold

improvement in the half-life of crude enzyme. Immobilized enzyme retained 92 % of the original activity after 17 cycles. The

production of xylo-oligosaccharides was 37.8% higher with immobilized enzyme as compared to its free counterpart.

Bhushan et al. (2013) immobilized A. flavous MTCC 9390 xylanase by encapsulation in alginate beads. The immobilized

enzyme had a pH optimum of 5.5 (as compared to 5.0 of free enzyme) and temperature optimum of 70 °C (10 °C higher than that of

soluble enzyme). The Vmax of the immobilized enzyme was 20 U/mL which is lower than that of free form, while Km value was 1.3

fold higher (1.5 % xylan for free and 2.08 % xylan for immobilized enzyme). The storage stability of immobilized enzyme was

appreciably higher as indicated by the presence of 80% residual activity after storage for one month at 4 °C. Samples of pineapple

juice showed relatively less viscosity, suspended solids and more clarity with immobilized enzyme treatment than its free counterpart.

Madakbas et al. (2013) immobilized xylanase on glutaraldehyde- activated polyaniline support. The immobilized enzyme

showed an increase in pH optimum by 1.0 unit and improved thermostability as compared to the free enzyme. Further, the

immobilized enzyme exhibited better reusability and storage stability than the free enzyme.

2.11 Applications of xylanase

Interest in microbial xylanases has increased markedly because of their wide range of potential biotechnological applications in

pulp and paper industry, production of xylo-oligosaccharides, texture improvement of bakery products, textile industry, nutritional

improvement of pig and poultry feed, fruit softening and clarification of juices and wines, bioconversion of lignocellulosic material

and agro-wastes to fuels and chemical feedstocks, production of pharmaceutically active polysaccharides for use as antimicrobial

agents or antioxidants, detergents, extraction of coffee, plant oils and pigments, and degumming of plant fibers such as flax, hemp and

jute (Kuhad and Singh, 1993; Bajpai, 2004; Wong et al., 1998; Beg et al., 2001; Subramaniyan and Prema, 2002; Polizeli et al., 2005;

Butt et al., 2008; Dhiman et al., 2008; Kulkarni et al., 2009; Harris and Ramalingam, 2010; Juturu and Wu, 2011; Sharma and Kumar,

2013). In the present study, application of xylanase produced by a newly isolated strain has been investigated in pulp biobleaching,

juice enrichment from fruit pulps and saccharification of wheat straw.

2.11.1 Application of xylanase in pulp biobleaching in pulp and paper industry

The main application of xylanases is in the bleaching of pulp. The conversion of wood into paper involves pulping (often kraft

pulping) and bleaching. Kraft process involves pretreatment of wood shavings with a combination of NaOH and sodium sulphide at

165°C in a digester. During this process, about 90-95% of the hemicellulose and lignin are dissolved and partially degraded. The

deposited lignin imparts a dark color to the pulp (Damiano et al., 2003). This is followed by washing and pre-bleaching of the brown

mass to remove minor impurities and a part of the remaining lignin. Subsequently, chemical bleaching is carried out which may use

ozone, chlorine, chlorine dioxide, hydrogen peroxide and sodium hydroxide. The main advantage of the Kraft process is the possibility

of recovering the chemical products from the black liquor. On the other hand, the disadvantages are the high initial costs, the strong

smell of gases emitted by the process, low yield (40–50%) and the high cost of bleaching. The use of chlorine based bleaching results

in the production of organochlorine compounds which are discharged in the effluent. These compounds are highly toxic, mutagenic,

persistent and harmful to biological systems (Bajpai and Bajpai, 1999). Environmental regulations have restricted the use of chlorine

compounds in bleaching processes in the paper and cellulose industries, especially in Western Europe and North America. Special

attention has been given to using xylanase in pre-bleaching, which would lower the amount of chlorine compounds used by up to

30%, so that a 15–20% reduction in organochlorines in the effluents could be achieved (Bajpai, 2004). The utilization of xylanases

could lead to the replacement of 5–7 kg of chlorine dioxide per ton of Kraft pulp thereby reducing environmental pollution (Polizeli et

al., 2005).

In many countries, enzymatic bleaching is employed in paper manufacture. The use of xylanase to enhance bleaching of the

pulp was first demonstrated by Viikari et al. (1986) and since then, several research groups have reported the use of xylanase in pulp

biobleaching. Xylanase-aided bleaching has been identified as a future technology (Marttila et al., 2000). The importance of xylanases

in pulp bleaching has been discussed in various reviews (Eriksson, 1990; Srinivasan and Rele, 1999; Bajpai, 1999; Techapun et al.,

2003; Bajpai, 2004; Polizeli et al., 2005; Kulkarni et al., 2009). Xylanases employed in paper industry do not need to be purified but

must be active at alkaline pH and high temperature and must be cellulose-free in order to preserve the cellulose fibers (Polizeli et al.,

2005). Xylanases from fungal sources have limited acceptability in pulp bleaching on a commercial scale due to the presence of

cellulase activity, low optimum pH and temperatures tolerance below 50°C (Subramaniyan and Prema, 2002). Therefore, cellulase-

free xylanases from other sources have been used in paper and pulp industry to ensure minimal damage to pulp fibres (Viikari, 1994;

Srinivasan and Rele, 1999; Buchert et al., 1992; Senthilkumar et al., 2005). A number of bacterial xylanases have been evaluated for

their potential in the pulp biobleaching process, which include Bacillus sp. NCIM 59 (Kulkarni and Rao, 1996); B. circulans (Dhillon

et al., 2000); B. pumilus (Bim and Franco, 2000; Kaur et al., 2011b), Bacillus sp. strain BP-23 (Torres et al., 2000); Staphylococcus sp.

SG-13 (Gupta et al., 2000); B. coagulans (Choudhury et al., 2006); B. megaterium (Sindhu et al., 2006); B pumilus ASH (Battan et al.,

2007; Garg et al., 2011); Bacillus sp. GRE7 (Kiddinamoorthy et al., 2008); B. subtilis ASH (Sanghi et al., 2009); B.

sterothermophilus (Dhiman et al.,2009), and Bacillus pumilus SV 85 (Nagar et al., 2013).

Xylanase may depolymerize the xylan reprecipitated on the surface of fibers during the kraft process resulting in increased

accessibility of bleaching chemicals to pulp fibers in the subsequent treatments (Viikari et al., 1994). Pretreatment with xylanase

might interrupt the lignin-carbohydrate bonds by depolymerizing xylan chains, thereby improving the accessibility of the bleaching

chemicals to the pulps and facilitating easier removal of solubilized lignin in bleaching (Paice et al., 1992; Wong et al., 1997; Lei et

al., 2008). Xylanase could remove the xylan derived chromophores such as hexenuronic acid, formed from 4-O-methylglucuronic acid

side groups of xylans during kraft cooking of wood, thereby increasing the brightness of pulp (Wong et al., 1997a). The work

published by some researchers on the use of xylanases in pulp bleaching is given below: -

Senior and Hamilton (1993) have shown that xylanase treatment and extraction change the reactivity of the pulp by enabling a

higher chlorine dioxide substitution to achieve target brightness and that they raise the brightness ceiling of fully bleached pulps. Garg

et al. (1996) optimized the enzyme pretreatment conditions of the pulp and found that enzyme treatment was to be effective at 65°C

and pH 6.0, at 5% consistency, for 3h on using the cellulase-free thermophilic xylanase from Streptomyces thermoviolaceus. Enzyme

treatment at 2% and 5% consistencies caused considerable decrease in long-span tensile strength and burst index, both of which are

predominantly a measure of the degree of interfiber bonding. Moreover, at 10% pulp consistency, the decrease in long-span tensile

strength and burst index was negligible.

Kulkarni and Rao (1996) investigated the biobleaching effect of cellulase-free xylanase produced from alkaliphilic

thermophilic Bacillus sp. NCIM 59 on the bagasse pulp at pH 7.0, temperature 50°C and an incubation period of 4 h. Xylanase

treatment of the bagasse pulp resulted in 2 unit decrease in the kappa number without altering the strength properties of the pulp.

Subsequently, the peroxide bleaching of the enzyme treated samples resulted in a decrease in the kappa number by 10 U and an

increase in brightness by 2.5%. The viscosity of xylanase treated samples remained unaltered.

Dhillon et al. (2000) pretreated eucalyptus kraft pulp with 7 IU/g of B. circulans AB 16 xylanase at 55°C for 3 h in a

multistage bleaching process using CEH bleaching sequence. Enzymatic prebleaching resulted in the reduction of 20% chlorine

consumption without any decrease in brightness. The viscosity of xylanase-treated pulp was 9.5–9.7 cp, whereas that of the pulp

treated exclusively with chlorine was 9.2 cp. Gupta et al. (2000) produced an alkalistable xylanase from Staphylococcus sp. SG-13 and

evaluated its bleach boosting on kraft pulps at an enzyme dose of 1.8U/g oven dried pulp at pH 9.5–10.0 and temperature 50 °C after 4

h of reaction time. Pretreatment of the pulp with xylanase and its subsequent treatment with 8% hypochlorite reduced the kappa

number by 30%, and enhanced the brightness and viscosity by 11% and 1.8%, respectively. They also observed an improvement in the

pulp properties such as tensile strength and burst factor by up to 10% and 17%, respectively as well as an increase in the CED

viscosity of pulp by 1.8%.

Torres et al. (2000) studied the application of xylanase from Bacillus sp. strain BP-23 in the bleaching of oxygen-delignified

eucalyptus kraft pulps. Treatment with the enzyme reduced the kappa number by 16% and chlorine dioxide consumption by 30% to

obtain similar kappa numbers after the Ep stage. Viscosity was not strongly affected by the use of xylanase treatment. At the same

chlorine dioxide consumption, enzyme treated pulps reached about 1% higher final brightness than pulps without xylanase

pretreatment, regardless of the pulp used.

Madlala et al. (2001) investigated the potential of xylanase P (a commercial xylanase from Sappi forest products, Southern

Africa) in bleaching of kraft pulp. The enzymatic pretreatment of pulp improved the brightness of kraft pulp by 5.6 brightness points

when used at 7 U/g of moisture-free pulp and caused an approximately 10% reduction in chlorine dioxide consumption.

Beg et al. (2005) used a thermostable and cellulase-free xylanase from Streptomyces sp. QG-11-3 for biobleaching of

eucalyptus kraft pulp at an enzyme dose 3.5 U/g oven dried pulp, pH 8.5, temperature 50°C and incubation period 2 h. A reduction in

chlorine consumption up to 8% could be achieved with biobleached pulp when subsequently subjected to chlorine treatment. Xylanase

treatment of the pulp reduced the kappa number by 25% and enhanced the brightness (%ISO) by 20%. In enzymatic bleaching, the

pulp properties such as tensile strength and burst factor showed an improved by up to 63% and 8%, respectively.

Choudhury et al. (2006) investigated the biobleaching potential of B. coagulans xylanase by treating eucalyptus kraft pulp with

enzyme dosage of 7 IU/g dry weight of pulp at pH 7.0–7.2, consistency 10% and temperature 50–55°C for 90 min. The enzymatic

bleaching of the kraft pulp increased the CED viscosity of pulp by 5.45%. Sindhu et al. (2006) observed that xylanase from B.

megaterium was the most effective in the enzymatic bleaching of the kraft pulp at 8 U/g odp, at 10% consistency and pH 8.0 on

incubation at 50 °C for 180 min. Enzymatic bleaching of the kraft pulp resulted in 8.12% (7 points) increase in brightness, 1.16%

increase in viscosity, 13.67% (2.26 points) decrease in kappa number, and 31% decrease in chlorine consumption at the CD stage. The

use of xylanase increased the CED viscosity of pulp by 13.67%.

Battan et al. (2007) studied the biobleaching of eucalyptus kraft pulp with B. pumilus xylanase at enzyme dose 5 IU/g odp, pH

7.0, temperature 60 °C and retention time 180 min. Treatment of kraft pulp with xylanase under the optimized conditions viz. xylanase

dose 5 IU/g pulp, consistency 10%, pH 7.0, incubation temperature 60 °C, and 180 min of treatment resulted in 20% reduction in

chlorine consumption without any change in brightness. An increase of 5% in brightness along with an increase of 21% in whiteness

and 28% in fluorescence was observed, whereas 18% decrease in the yellowness was observed. Enzyme treated pulp resulted in 16%

reduction in kappa number, and significant improvement in various pulp properties such as viscosity, tensile strength, breaking length,

burst factor, burstness, tear factor and tearness.

Khandeparker et al. (2007) produced thermoalkalophilic and cellulase-free xylanase from Arthrobacter sp. and evaluated it for

biobleaching of kraft pulp. Enzymatic prebleaching of kraft pulp showed 20% reduction in kappa number of the pulp without much

change in viscosity. Enzymatic treatment reduced the amount of chlorine by 29% without any decrease in brightness.

Kiddinamoorthy et al. (2008) documented 10 IU/g as the most effective dose of Bacillus sp. GRE7 xylanase for enzymatic

prebleaching of eucalyptus kraft pulp. The xylanase pretreatment of the pulp resulted in kappa number reduction and brightness gain

of 1.01 and 2.7 points, respectively and a savings of up to 30% chlorine dioxide consumption.

Dhiman et al. (2009) analyzed the biobleaching action of xylanase alone in the range of 0–12.5 U/g of oven dried pulp

for ‘single lay out’ (strategy I) and in combination with pectinase at a concentration of 2.5 U/g and 5 U/g of each enzyme for ‘mixed

lay out’ (strategy II) to investigate their bio-bleaching potentials. Strategy I was carried at 70 °C using 5 U/g of xylanase at pH 9.5 and

12.5 whereas strategy II was carried out at 70 °C using 5 U/g of each of the enzyme, respectively at pH 9.5. Bio-bleaching of the pulp

caused 15% and 20% less chlorine consumption though strategy I and II, respectively over chemical bleaching. Strategy II proved to

be 35.71% more efficient in ClO2 saving than the conventional method. Significant improvement in various pulp properties viz. tensile

strength 25.70%, breaking length 21.80%, burst factor 20.00%, burstness 13.86%, tear factor 6.61% and tearness 18.88%, was also

observed through ‘mixed lay out’ strategy.

Sanghi et al. (2009) pretreated the oven dried pulp at 10% consistency with 6 IU/g of B. subtilis ASH xylanase at pH 7.0 and

temperature of 60˚C for 2h. After enzymatic prebleaching, 4.6% reduction in kappa number (17.3 to 16.5), 11 % increase in ISO

brightness of kraft pulp, 4.9% increase in final brightness was observed. Xylanase pretreatment followed by chemical bleaching

according to the sequence CDED1D2 resulted in 28.6% reduction of chlorine consumption without any change in brightness. The

enzymatic pretreatment of the pulp resulted in an improved strength properties viz. gammage 6%, fiber thickness 6.1%, beating degree

35.5%, tensile index 1.34%, breaking length 13.3%, and tear index 18.7% and double fold 67.8% as compared to the control pulp.

Scanning electron microscopy revealed loosening and swelling of pulp fibers.

Kaur et al. (2011b) recorded 8.5% reduction in kappa number at prebleaching stage of kraft pulp and 25% reduction in active

chlorine consumption in subsequent bleaching stages without any decrease in brightness by the action of xylano-pectinolytic enzyme

from B. pumilus. A synergistic action of xylano-pectinolytic enzymes resulted in an increase of burst factor (9%), tear factor (4.6%),

breaking length (4.4%), double fold number (12.5%), gurley porosity (4%) and CED viscosity (11.8%) of enzyme- treated pulp

reflecting a significant improvement in pulp properties. Azeri et al. (2010) reported 1-3 % increase in ISO brightness of kraft pulp on

treatment with Bacillus sp. xylanase.

Garg et al. (2011) reported the optimum conditions for pretreatment of wheat straw pulp with B. pumilus ASH xylanase at 5

IU/g, pH 8.0, temperature 60°C and retention time 180 min. The pretreatment of wheat straw pulp resulted in final increase of 1.5%

ISO and 1.1% of brightness and whiteness and 6.0 % reduction in kappa number. Pretreatment